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Parasitic Diseases of Wild Birds
Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Parasitic Diseases of Wild Birds Edited by
Carter T. Atkinson Nancy J. Thomas D. Bruce Hunter
A John Wiley & Sons, Ltd., Publication
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Edition first published 2008 C 2008 Wiley-Blackwell Chapters 2, 3, 11, 20, and 25 are the work of the U.S. Government and is not subject to U.S. copyright. Blackwell Publishing was acquired by John Wiley & Sons in February 2007. Blackwell’s publishing program has been merged with Wiley’s global Scientific, Technical, and Medical business to form Wiley-Blackwell. Editorial Office 2121 State Avenue, Ames, Iowa 50014-8300, USA For details of our global editorial offices, for customer services, and for information about how to apply for permission to reuse the copyright material in this book, please see our website at www.wiley.com/wiley-blackwell. Authorization to photocopy items for internal or personal use, or the internal or personal use of specific clients, is granted by Blackwell Publishing, provided that the base fee is paid directly to the Copyright Clearance Center, 222 Rosewood Drive, Danvers, MA 01923. For those organizations that have been granted a photocopy license by CCC, a separate system of payments has been arranged. The fee codes for users of the Transactional Reporting Service are ISBN-13: 978-0-8138-2081-1/2008. Designations used by companies to distinguish their products are often claimed as trademarks. All brand names and product names used in this book are trade names, service marks, trademarks or registered trademarks of their respective owners. The publisher is not associated with any product or vendor mentioned in this book. This publication is designed to provide accurate and authoritative information in regard to the subject matter covered. It is sold on the understanding that the publisher is not engaged in rendering professional services. If professional advice or other expert assistance is required, the services of a competent professional should be sought. Library of Congress Cataloguing-in-Publication Data Parasitic diseases of wild birds / edited by Carter T. Atkinson, Nancy J. Thomas, D. Bruce Hunter. p. cm. Includes bibliographical references and index. ISBN-13: 978-0-8138-2081-1 (alk. paper) ISBN-10: 0-8138-2081-2 (alk. paper) 1. Birds–Parasites. 2. Birds–Diseases. I. Atkinson, Carter T. II. Thomas, Nancy J. (Nancy Jeanne), 1948– III. Hunter, D. Bruce. [DNLM: 1. Bird Diseases–parasitology. 2. Parasitic Diseases, Animal. SF 995.6.P35 P223 2008] SF995.6.P35P37 2008 636.5089 696–dc22
2008021325
A catalogue record for this book is available from the U.S. Library of Congress. R Set in 9.5/11.5pt Times by Aptara Inc., New Delhi, India Printed in Singapore by Markono Print Media Pte Ltd
The publisher and the author make no representations or warranties with respect to the accuracy or completeness of the contents of this work and specifically disclaim all warranties, including without limitation warranties of fitness for a particular purpose. No warranty may be created or extended by sales or promotional materials. The advice and strategies contained herein may not be suitable for every situation. This work is sold with the understanding that the publisher is not engaged in rendering legal, accounting, or other professional services. If professional assistance is required, the services of a competent professional person should be sought. Neither the publisher nor the author shall be liable for damages arising herefrom. The fact that an organization or Website is referred to in this work as a citation and/or a potential source of further information does not mean that the author or the publisher endorses the information the organization or Website may provide or recommendations it may make. Further, readers should be aware that Internet Websites listed in this work may have changed or disappeared between when this work was written and when it is read. 1
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Contents Preface Contributors Section I 1.
vii ix
Introduction
Parasitism: Costs and Effects Gary A. Wobeser
Section II
3
Protozoa
2.
Haemoproteus Carter T. Atkinson
13
3.
Avian Malaria Carter T. Atkinson
35
4.
Leucocytozoonosis Donald J. Forrester and Ellis C. Greiner
54
5.
Isospora, Atoxoplasma, and Sarcocystis Ellis C. Greiner
108
6.
Trichom*onosis Donald J. Forrester and Garry W. Foster
120
7.
Histomonas William R. Davidson
154
8.
Eimeria Michael J. Yabsley
162
9.
Disseminated Visceral Coccidiosis in Cranes Marilyn G. Spalding, James W. Carpenter, and Meliton N. Novilla
181
10.
Cryptosporidium David S. Lindsay and Byron L. Blagburn
195
11.
Toxoplasma J. P. Dubey
204
Section III
Helminths
12.
Trematodes Jane E. Huffman
225
13.
Schistosomes Jane E. Huffman and Bernard Fried
246
14.
Cestodes J. Daniel McLaughlin
261
15.
Acanthocephala Dennis J. Richardson and Brent B. Nickol
277
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Contents
16.
Eustrongylidosis Marilyn G. Spalding and Donald J. Forrester
289
17.
Trichostrongylus Daniel M. Tompkins
316
18.
Dispharynx, Echinuria, and Streptocara Ramon A. Carreno
326
19.
Tracheal Worms M. A. Fernando and John R. Barta
343
20.
Amidostomum and Epomidiostomum Alan M. Fedynich and Nancy J. Thomas
355
21.
Tetrameridosis John M. Kinsella and Donald J. Forrester
376
22.
Avioserpensosis John M. Kinsella
384
23.
Heterakis and Ascaridia Alan M. Fedynich
388
24.
Ascaridoid Nematodes: Contracaecum, Porrocaecum, and Baylisascaris Hans-Peter fa*gerholm and Robin M. Overstreet
413
25.
Diplotriaena, Serratospiculum, and Serratospiculoides Mauritz C. Sterner III and Rebecca A. Cole
434
26.
Filarioid Nematodes Cheryl M. Bartlett
439
27.
Capillarid Nematodes Michael J. Yabsley
463
Section IV 28.
Leeches
Leech Parasites of Birds Ronald W. Davies, Fredric R. Govedich, and William E. Moser
Section V
501
Arthropods
29.
Phthiraptera, the Chewing Lice Dale H. Clayton, Richard J. Adams, and Sarah E. Bush
515
30.
Acariasis Danny B. Pence
527
31.
Black Flies (Diptera: Simuliidae) Douglas C. Currie and D. Bruce Hunter
537
32.
Myiasis in Wild Birds Susan E. Little
546
Index
557
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Preface More that 30 years ago, John W. Davis, Roy C. Anderson, Lars Karstad, and Daniel O. Trainer edited the first edition of Infectious and Parasitic Diseases of Wild Birds. Since then there has been an explosion of new knowledge about parasitic diseases of wild birds, as wildlife disease specialists, ecologists, and evolutionary biologists have continued to unravel how parasitic protozoans, helminths, and ectoparasites affect wildlife populations. We continue in the footsteps of the first editors of this work by significantly expanding and updating the parasite portion of their original book. This work is a companion volume to Infectious Diseases of Wild Birds, which was published in 2007 by Blackwell Publishing, and complements Infectious Diseases of Wild Mammals, 3rd edition, edited by Elizabeth S. Williams and Ian K. Barker, and Parasitic Diseases of Wild Mammals, 2nd edition, edited by William M. Samuel, Margo J. Pybus, and A. Alan Kocan (Iowa State University Press). Taken together, these four volumes provide an important source of reference material for biologists and wildlife mangers, wildlife and veterinary students, professionals in the fields of animal health and wildlife disease, and evolutionary biologists with interests in disease ecology. We gratefully acknowledge our colleagues who established such excellent models for us to follow. This book focuses on the disease conditions produced by parasitic protozoans, helminths, leeches, and ectoparasitic arthropods, e.g. mites, and biting flies in free-living wild birds. Unlike most parasitology texts, this book emphasizes effects on the host rather than the parasites themselves, but still includes important information about their etiology, life cycles, transmission, and diagnosis. While no single work can cover the entire spectrum of wildlife parasites, we have attempted to assemble chapters that are both specific (e.g., Chapter 9, Disseminated Visceral Coccidiosis in Cranes) and general (e.g., Chapter 14, Cestodes) in their treatment of some of the diverse groups of organisms that use wild birds as intermediate or definitive hosts. In all cases, we have urged authors to avoid generalities and include specific examples of host–parasite
associations that can lead to clinical disease. We owe a great debt to the authors of these chapters both for their expertise in the material and for their willingness to endure the inevitable delays and revisions that are inherent in multiauthored works. Each chapter provides a classical description of the history, effects on the host, and causative agent, but the authors were also challenged to provide perspectives on the significance of the disease to wild birds and to document population impacts, an aspect that is particularly difficult to quantify in the wild. Unlike other volumes in this series, we elected to begin this book with an introductory chapter by Gary A. Wobeser who discusses some of the costs and effects of parasitism in wild avian populations. This chapter provides a succinct discussion of some of the difficulties in assessing impacts of parasitism on wild birds and provides a good framework for assimilating the detailed information in the sections that follow. We used The Clements Checklist of Birds of the World, 6th edition (Cornell University Press, 2007), as the authority for avian nomenclature and elected to allow authors to make individual decisions about whether to follow the proposed standardized nomenclature for parasitic diseases (SNOPAD; http://www. waavp.org/node/40). As a result, some chapters follow this terminology (e.g., Chapter 4, Leucocytozoonosis) while others retain the more traditional terminology (e.g., Chapter 7, Histomonas). Because many unpublished data on wild bird diseases have been compiled in laboratory and diagnostic files, citations of unpublished data were allowed for repositories of large, permanent, accessible institutions, such as the Canadian Cooperative Wildlife Health Centre, U.S. Geological Survey National Wildlife Health Center, and Southeastern Cooperative Wildlife Disease Study. Grateful acknowledgment goes to the Iowa State University Press, which guided this project through its initial stages, and to Blackwell Publishing, which took it over and shepherded it through to completion. We owe sincere debts of gratitude to Donald J. Forrester who was instrumental in the initial organization of the book and to Amy Miller for her significant
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Preface
contribution in the technical editing of the final manuscript. We acknowledge the support of the U.S. Geological Survey, Wildlife and Terrestrial Resources Program, and the University of Guelph. This book is dedicated to the Wildlife Disease Association, whose members initiated the revision of this book series and who continue to provide the backbone of growing
knowledge in the field of wildlife disease. Royalties that accrue from sales of this book will be provided to the Wildlife Disease Association. Carter T. Atkinson Nancy J. Thomas D. Bruce Hunter
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Contributors University of Utah Salt Lake City, Utah, U.S.A.
Richard J. Adams Department of Biology University of Utah Salt Lake City, Utah, U.S.A.
Rebecca A. Cole U.S. Geological Survey National Wildlife Health Center Madison, Wisconsin, U.S.A.
Carter T. Atkinson U.S. Geological Survey Pacific Island Ecosystems Research Center Hawaii National Park, Hawaii, U.S.A.
Douglas C. Currie Royal Ontario Museum Toronto, Ontario, Canada and Department of Ecology and Evolutionary Biology University of Toronto Toronto, Ontario, Canada
John R. Barta Department of Pathobiology Ontario Veterinary College University of Guelph Guelph, Ontario, Canada Cheryl M. Bartlett Department of Biology Cape Breton University Sydney, Nova Scotia, Canada
William R. Davidson D. B. Warnell School of Forestry and Natural Resources and Southeastern Cooperative Wildlife Disease Study College of Veterinary Medicine University of Georgia Athens, Georgia, U.S.A.
Byron L. Blagburn Department of Pathobiology College of Veterinary Medicine Auburn University, Alabama, U.S.A. Sarah E. Bush Natural History Museum and Biodiversity Research Center University of Kansas Lawrence, Kansas, U.S.A.
Ronald W. Davies Department of Biological Sciences University of Calgary Calgary, Alberta, Canada
James W. Carpenter Department of Clinical Sciences College of Veterinary Medicine Kansas State University Manhattan, Kansas, U.S.A.
J.P. Dubey Animal Parasitic Diseases Laboratory Animal and Natural Resources Institute Agricultural Research Service U.S. Department of Agriculture Beltsville, Maryland, U.S.A.
Ramon A. Carreno Department of Zoology Ohio Wesleyan University Delaware, Ohio, U.S.A.
Hans-Peter fa*gerholm Laboratory of Aquatic Pathobiology Department of Biology Abo Akademi University Åbo, Finland
Dale H. Clayton Department of Biology
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x Alan M. Fedynich Caesar Kleberg Wildlife Research Institute Texas A&M University-Kingsville Kingsville, Texas, U.S.A. M.A. Fernando Department of Pathobiology Ontario Veterinary College University of Guelph Guelph, Ontario, Canada Donald J. Forrester Department of Infectious Diseases and Pathology College of Veterinary Medicine University of Florida Gainesville, Florida, U.S.A. Garry W. Foster Department of Infectious Diseases and Pathology College of Veterinary Medicine University of Florida Gainesville, Florida, U.S.A. Bernard Fried Department of Biology Lafayette College Easton, Pennsylvania, U.S.A. Fredric R. Govedich Department of Biological Sciences Southern Utah University Cedar City, Utah, U.S.A. Ellis C. Greiner Department of Infectious Diseases and Pathology College of Veterinary Medicine University of Florida Gainesville, Florida, U.S.A. Jane E. Huffman Department of Biological Sciences Applied DNA Sciences Fish and Wildlife Microbiology Laboratory East Stroudsburg University East Stroudsburg, Pennsylvania, U.S.A. D. Bruce Hunter Department of Pathobiology Ontario Veterinary College University of Guelph Guelph, Ontario, Canada
Contributors John M. Kinsella Department of Infectious Diseases and Pathology College of Veterinary Medicine University of Florida Gainesville, Florida, U.S.A. David S. Lindsay Department of Biomedical Sciences and Pathobiology Virginia-Maryland Regional College of Veterinary Medicine Virginia Polytechnic Institute and State University Blacksburg, Virginia, U.S.A. Susan E. Little Department of Veterinary Pathobiology Center for Veterinary Health Sciences Oklahoma State University Stillwater, Oklahoma, U.S.A. J. Daniel McLaughlin Department of Biology Concordia University Montreal, Quebec, Canada William E. Moser Department of Invertebrate Zoology National Museum of Natural History Smithsonian Institution Washington, District of Columbia, U.S.A. Brent B. Nickol School of Biological Sciences University of Nebraska-Lincoln Lincoln, Nebraska, U.S.A. Meliton N. Novilla WIL Research Laboratories—Biotechnics LLC Greenfield, Indiana, U.S.A. Robin M. Overstreet The University of Southern Mississippi Gulf Coast Research Laboratory Ocean Springs, Mississippi, U.S.A. Danny B. Pence Department of Pathology Texas Tech University Health Sciences Center Lubbock, Texas, U.S.A.
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Contributors Dennis J. Richardson Department of Biological Sciences Quinnipiac University Hamden, Connecticut, U.S.A.
Daniel M. Tompkins Landcare Research Dunedin, New Zealand
Marilyn G. Spalding Department of Infectious Diseases and Pathology College of Veterinary Medicine University of Florida Gainesville, Florida, U.S.A.
Gary A. Wobeser Department of Veterinary Pathology Western College of Veterinary Medicine University of Saskatchewan Saskatoon, Saskatchewan, Canada
Mauritz C. Sterner III U.S. Geological Survey National Wildlife Health Center Madison, Wisconsin, U.S.A.
Michael J. Yabsley D. B. Warnell School of Forestry and Natural Resources and Southeastern Cooperative Wildlife Disease Study Department of Population Health College of Veterinary Medicine University of Georgia Athens, Georgia, U.S.A.
Nancy J. Thomas U.S. Geological Survey National Wildlife Health Center Madison, Wisconsin, U.S.A.
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Section I: Introduction
Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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1 Parasitism: Costs and Effects Gary A. Wobeser r difficulty in quantifying factors related to disease. It is impossible to assess the significance of a parasite for a population without the ability to calculate basic epidemiological proportions such as prevalence, incidence, morbidity, and mortality rates. The number of individuals affected by a parasite (the numerator for such calculations) is usually difficult to determine and the population at risk (the denominator) rarely can be measured adequately; r the need to consider the long-term effect of a parasite in wild birds. This may be very difficult, even when the number affected and the population at risk can be determined. If a disease, such as coccidiosis, occurs in a flock of chickens and 15% die, the significance of the disease is that 15% fewer chickens go to market. However, a similar 15% loss in a wild bird population might result in more resources per capita for the remaining birds, leading to reduced mortality from other factors and/or improved reproduction. The potential for compensation or other delayed effects may be very important in assessing the impact of a parasite on wild birds at the population level; r the sample of wild birds available for study is usually biased by the method of collection and may not represent the actual state of nature. Depending on the method of collection, affected birds may be under- or overrepresented, even in groups collected by mass-capture methods (Sulzbach and Cooke 1978); and r the anonymity of wild birds, except for the small number marked by the researcher. For instance, although age is an important disease determinant, the age of wild birds often cannot be determined except to differentiate hatch-year from after-hatch-year birds. Individuals seldom can be traced back in time to discover previous exposure to disease agents or forward in time to discover
Parasitism has been defined in many ways, but in terms of wildlife disease, it is usually taken to mean an obligatory trophic association between individuals of two species in which one (the parasite) derives its food from a living organism of the other species (the host). An individual host bird can be viewed as an island of habitat that provides resources for parasites, with the parasites deriving benefits while the host is harmed or bears some cost. Parasitism is common in nature; for example, Price (1980) estimated that half of all animal taxa are parasitic. Parasitism is ubiquitous in wild birds and individual birds are affected by many different parasites during their lifetime, but our understanding of the parasites that occur in wild birds is fragmentary. Moore and Clayton (1997) concluded that the majority of parasites of wild birds have yet to be described taxonomically. Some groups, such as blood-inhabiting protozoa (the hematozoa), have been studied widely, perhaps because of the ease with which blood can be collected from living birds, while little is known about other groups such as intestinal flagellates. But even within the hematozoa, species diversity has probably been greatly underestimated (Bensch et al. 2007). Similarly, more is known about the effects of arthropod ectoparasites than about the effect of protozoa and helminths on birds, and cavity nesting birds have been studied more extensively than most other species because of the relative ease in capturing, examining, and following these birds. Studying parasitism in wild birds is subject to a number of constraints that make working with disease in any free-ranging species more difficult than studying humans or domestic animals. These include:
r inadequate baseline information about the host species. Knowledge of avian life history traits is rudimentary (Zera and Harshman 2001), and so one often must extrapolate from other species and collect information about the basic biology of the host while trying to understand a host–parasite relationship; 3
Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Parasitic Diseases of Wild Birds their fate. Commonly used techniques such as retrospective and prospective case–control studies that are useful in human and veterinary epidemiology are impossible except in unusual circ*mstances, such as in birds with a high degree of nest site fidelity.
A fundamental feature of parasitism is that the presence of a parasite involves a cost to the host. The costs of parasitism may include:
r loss of resources extracted by the parasite directly from the host, for example, loss of blood to blood-feeding ectoparasites; r competition between the parasite and the host for resources, as occurs with cestodes that absorb nutrients from the host’s gut content; r costs to the host for defense against parasites. These may include foregoing resource-rich areas to avoid areas where parasites may be present, costs for grooming, moving away from parasites, or abandoning a nest, costs to develop and maintain innate and acquired resistance, and costs to activate these systems; r costs resulting from tissue injury related to the parasite. This may be direct damage caused by the parasite or, more often, injury from the inflammatory and immune response to the parasite. Some injuries may result in dysfunction, such as reduced mobility, reduced digestive efficiency, or increased loss of nutrients through intestinal or kidney injury, that interfere with obtaining or retaining resources; r costs related to improper development as a result of parasitism early in life (e.g., Spencer et al. 2005); and r costs to repair or replace damaged tissues. The diversity of parasites and the variety of ways that they interact with hosts make it difficult to measure the cost of a single parasite species; to compare the relative cost of different parasites such as the lice, intestinal coccidia, and tracheal worms, all of which might be infecting a single host; or to understand how these parasites may interact with each other and with other environmental factors to affect a host population. The costs described above are related to resources, and particularly to energy [“the single common denominator of life”; “something that is absolutely essential and involved in every action large or small” (Odum 1993)]. Energy is a measure of the ability to do work and is a “currency” that can be used to consider the costs of all types of parasitism, at least conceptually if not quantitatively at this time. Four basic features must be
considered when using energy as a currency to consider parasitism:
r The supply of energy is limited. Most birds are unable to increase their intake of energy readily, and so they must function within a finite budget. In other words, a bird cannot use more energy than it can assimilate or has in storage. r The amount of energy available and accessible is not constant or uniform. The energy available to a bird varies with the time of year, weather, habitat conditions, and the number of competitors for that energy. Not all individuals in a population have equal access to the resources that are available; thus, within a group or population some birds may have abundant resources while others do not. r Use of energy for one purpose reduces the amount available for other uses. Most of the energy assimilated by a bird is used for maintenance, that is, keeping the body functioning, repaired, maintaining a high core temperature, avoiding predators, and defending against disease. Energy that remains can be used for production (growth and reproduction) or stored as fat for future use. If extra energy is used to defend against parasites or to repair tissue injured by parasites, the energy available for reproduction or growth is reduced. For instance, the cost of producing antibody to a novel antigen is equivalent to that of producing half an egg in female House Sparrows (Passer domesticus) (Martin et al. 2003) and mounting an immune response resulted in asymmetry of flight feathers in nestling Mountain Chickadees (Poecile gambeli) (Whitaker and Fair 2002). Conversely, increased reproductive effort may result in reduced ability to mount a defense against parasites (Deerenberg et al. 1997). r The need for energy for various purposes is highly variable among individuals and at different times of year. Because an individual cannot maximize all life history traits simultaneously, life history theory suggests that a bird should adopt a strategy that optimizes energy use among resource-demanding activities, such as defense and reproduction, to maximize lifetime fitness. Ecologists use the term “trade-off” for this process of making physiological choices among competing needs for resources that should maximize the chances of an individual’s genes being passed on to the next generation. Individuals that make the wrong choices are less successful or “fit,” and this may provide a basis for genetic selection.
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Parasitism: Costs and Effects As a result of heterogeneity in both the supply of energy and the need for energy, the appropriate physiological trade-offs in relation to parasitism vary among individual birds and for different parasites, and the pattern of trade-offs is different seasonally and annually. For this reason, the reaction to parasites and the effects of parasitism must always be considered in terms of the context in which parasitism is occurring and of how the situation might influence resource trade-offs. For instance, during one season a bird may be in poor nutritional condition and need to direct all its available resources to simply staying alive, with little or no ability to mount an effective defense against parasites or to grow or reproduce. At another time of year the same bird may have ample resources to meet all needs, and so it can afford strong resistance to parasites and still be able to grow and reproduce effectively. Young birds may have different priorities than adults and the sexes may have different strategies and tradeoffs. For instance, Tschirren et al. (2003) suggested that a greater need for carotenoid-based coloration for signaling by male Great tit* (Parus major) might lead to a trade-off that results in reduced immunocompetence in males. Privileged individuals within the population, such as birds that possess a territory, may have a totally different context for trade-offs related to parasites than do the “have-nots” within the population. Changes in environmental conditions may change the context; for example, Blow Fly (Protocalliphora braueri) larvae had no effect on Sage Thrasher (Oreoscoptes montanus) nestling weight, size at fledging, or mean fledgling age, but in a year with cold wet weather, survival and fledging success were markedly reduced among parasitized birds compared to unparasitized birds (Howe 1992). Knowledge of how trade-offs occur in relation to parasitism is fragmentary at this time and general rules about which activity (reproduction, growth, defense against predators or parasites) should take precedence for resources are likely subject to many exceptions. For instance, hosts may be selected to develop acquired immunity to only some of the disease agents that they encounter (Boots and Bowers 2004). While mounting a strong defensive response to parasites is likely a “good” thing generally, in some situations it may be adaptive to suppress the defensive response. This may be the case in nesting Common Eiders (Somateria mollissima). Female eiders do not feed during breeding and face severe resource restrictions while incubating. Birds that do not begin with adequate resources abandon their nest in order to survive. Hanssen et al. (2004) immunized incubating female eiders with nonpathogenic antigens, including sheep red blood cells. Not surprisingly, the rate of successful immunization was not very good compared to what
5
would be expected at other times of year. Under these circ*mstances, it appears that the appropriate choice for many eiders is to use their limited resources to survive and reproduce rather than to mount an immune response. A second part of the same study compared survival of birds that mounted an immune response to that of birds that did not produce antibodies. Both responding and nonresponding eiders had sufficient resources to complete reproduction; however, only about 27% of birds that produced antibody to sheep red blood cells returned to the colony in subsequent years, compared with approximately 72% of birds that did not produce antibody. Under these conditions, females that invested in an immune response “experienced considerably impaired long-term survival” compared to females that did not respond. This example also serves to illustrate that the effect of a trade-off on fitness may be delayed. The cost to the host is not obvious for most parasites encountered in wild birds. It is only in a minority of situations, described elsewhere in this book, that parasitism is clearly associated with recognizable functional impairment of the host that we can characterize as disease. The apparently “benign” nature of many parasites could be because:
r the effect of the parasites actually is so trivial as to be undetectable; r the cost is not trivial but it is tolerable; that is, the bird has sufficient resources to cover the costs without significant negative effects on other functions under conditions at the time the effect was measured; r the cost of parasitism is obscured by other more proximate regulatory factors such as predation and competition. Predation is thought to be a major factor in shaping the life history of birds (Zera and Harshman 2001) and parasitized prey may be taken disproportionately by predators (Temple 1987). In some situations the parasite benefits if the infected host is eaten by an appropriate predator (parasite-induced trophic transmission; Lafferty 1999). But infections in which there is no apparent benefit to the parasite may make animals more susceptible to predators, perhaps because of the pathology induced by the parasite. Hudson et al. (1992a) found that Red Grouse (Lagopus lagopus scotica) killed by predators were more heavily parasitized by the cecal nematode (Trichostrongylus tenuis) than were hunter-killed birds and that birds with many worms may emit more scent and, hence, be more vulnerable to mammalian predators. In some situations, increased vulnerability to predators may be related to energy trade-offs and reduced resources for
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predator vigilance or avoidance. For instance, Common Redshanks (Tringa totanus) that are energetically stressed (as might result from parasitism) respond by taking risks that increase the probability of predation (Quinn and Cresswell 2004). The interaction between predation and parasitism is undoubtedly complex. Navarro et al. (2004) found that House Sparrows exposed to potential predators (cat or owl) had reduced T-cell-mediated immune response and a higher prevalence and intensity of infection with Haemoproteus spp. than did sparrows exposed to nonthreatening animals (rabbit or pigeon), suggesting that even the threat of predation may alter trade-offs that influence parasitism. Although little is known about the effect of parasitism on intraspecific competition, this may be an important factor. For instance, male Greater Sage-Grouse (Centrocercus urophasianus) infested with lice are discriminated against for breeding (Spurrier et al. 1991). Females appear to recognize infected males by the occurrence of petechial hemorrhages on the air sacs and males infested with lice are shunned, and so their reproductive input to the population is minimal; that is, their fitness is very low and there is likely negative selection against their genotype. In a similar manner, male Red Grouse infected with T. tenuis may have difficulty defending a territory (Delahay et al. 1995). Consideration of interactions between parasitism and competition must also include competition among species that share parasites, such as the Ring-necked Pheasant (Phasianus colchicus) and Gray Partridge (Perdix perdix) that share Heterakis gallinarum, with asymmetrically severe effects on the partridge (Tompkins et al. 2001b); and r the cost is not trivial but it goes undetected because of insensitivity of the methods used to look for effects. For instance, it would be very easy to dismiss the tiny hemorrhages caused by lice as inconsequential to male Greater Sage-Grouse, without even considering that they might have a profound effect on behavior, reproductive success, and natural selection. The costs of parasitism could also be overlooked because the wrong individuals within the population are examined, the interaction between parasite and host is examined in an inappropriate context (e.g., at the wrong time of year or in an experimental situation in which resources are not limited), inappropriate parameters are measured, or because the long-term (lifetime) consequences of parasitism are not measured. Møller (1994) suggested that the cost of parasitism
in nestling birds could be paid by the nestlings through reduced growth or survival or by the parents through reduced survival or future reproductive success as a result of having to provide additional resources to the parasitized young. Bize et al. (2003) found that nestling Alpine Swifts (Tachymarptis melba) can compensate for early growth retardation by rapid feather growth, so that if measured at fledging no effect might be obvious; however, rapid feather growth may result in poor feather quality with later effects (Dawson et al. 2000). Nutrient shortage in early development can have other serious long-term consequences including effects on adult dominance rank, morphology, and lifespan (Metcalfe and Monaghan 2001). Island Canaries (Serinus canaria) infected with plasmodia as nestlings have structural changes in their brain and reduced song repertoire as adults (Spencer et al. 2005). The effects of parasites are usually not distributed evenly or fairly among all members of a population, which complicates measuring their cost. Metazoa characteristically are distributed in an aggregated manner within the host population (Shaw et al. 1998). Most hosts have few or no parasites and a few individuals have many parasites (often referred to as the 20:80 rule: 20% of the population carries 80% of the parasites). Severe effects are likely to be confined to those individuals with many parasites. Measures of central tendency, such as average intensity of infection and average cost of parasitism, may not be helpful in understanding the significance of the parasite if effects are concentrated in a small group of heavily infected individuals. These animals at the extreme end of the distribution are also important as the major source of infection within the population, but samples drawn from the population are unlikely to contain these individuals unless the sample is very large. Much of the information available on the occurrence of parasites in wild birds comes from the study of birds that died of other causes, because it is inappropriate to kill large samples of birds simply to record their parasites. At one extreme, such a sample may primarily consist of the survivors of conditions that were severe, resulting in underestimation of the cost of parasitism. At the opposite extreme, the sample may contain the few significantly affected individuals in the population, and so the cost to the population is overestimated. “While the study of specific host–parasite relationships have proven insightful, they reflect only a small part of the wealth of parasites and pathogens in an
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Parasitism: Costs and Effects animal’s internal and external environment” (Lochmiller and Deerenberg 2000). Virtually all the information available about parasites of birds relates to the effects of individual parasite species, but individual birds are host to many different parasites, often simultaneously; for example, a single feather may be infested with 6 species of feather mite (P´erez and Atyeo 1984) and a group of 45 Lesser Scaup (Aythya affinis) were infected by almost 1 million individuals of 52 different helminth species (Bush and Holmes 1986). Examining the effect of parasitism as the interaction between two species fails to account for interactions among parasites that might be additive, synergistic, or antagonistic. Almost nothing is known about the effects or dynamics of parasite assemblages or communities in wild birds. The largest challenge for those interested in parasites of birds is to answer the question “Do parasites influence bird populations?” Most ecologists and wildlife managers have assumed that the answer is “No” (Tompkins et al. 2001a), but modeling suggests that parasites could regulate host populations if they reduce host survival and/or fecundity in a densitydependent manner (Anderson and May 1978; May and Anderson 1978). To understand the effect of a parasite on the host population, one needs to understand the effect of the parasite on the individual host, the prevalence and intensity of parasite infection within the host population, and the context within which the interaction is occurring. Parasites rarely result in obvious piles of dead birds but many studies have concentrated on the direct effect of parasites on mortality although “. . . highly pathogenic parasites tend not to have an impact at the population level. . . . ” (Hudson and Dobson 1997), because this type of parasite may kill the host rapidly, thus limiting transmission to other individuals. Sublethal effects of chronic infections that are mediated through reduced fecundity are more likely to have an effect at the population level. Much of the information available about parasites in birds is descriptive. More than 70 years ago, Aldo Leopold recognized that observational and correlational studies have limited ability to lead to an understanding of disease in wild species (Leopold 1933). Marzal et al. (2005) observed that knowledge of causal relationships of disease caused by parasites of birds “is still rudimentary due to a scarcity of experimental manipulation,” and Tompkins and Begon (2000) stated that “regulation by parasites can be established only by experimentally perturbing host/parasite systems away from their equilibrium levels and monitoring subsequent changes in both parasite and host densities relative to control.” Studies that include intervention through treatment of parasites in natural populations, such as by Hudson et al. (1992b, 1998) (T. tenuis and Red Grouse), Merino et al. (2000)
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(hematazoa in Eurasian Blue tit*, Cyanistes caeruleus), Hoodless et al. (2002) (ticks and Ring-necked Pheasants) and Marzal et al. (2005) (Haemoproteus prognei in House Martins, Delichon urbicum), and through experimental infection (e.g., Spencer et al. 2005), have provided insights into parasitism that would be unattainable with traditional observational study. As in all aspects of the study of parasitism, it is important to consider the long-term effects of such interventions. For instance, Hanssen et al. (2003) studied the effect of antiparasite treatment on nesting female eiders. There was no effect of treatment on nest success or on the survival to the next year of birds that nested successfully. However, among the females that were unsuccessful in nesting, 69% of treated birds survived compared with 18% of untreated birds. This suggests that birds that nested successfully were able to tolerate the effects of parasitism, while unsuccessful females were less able to bear the costs from parasites, resulting in a delayed effect on survival. In another example, McCutchan et al. (2004) found that a vaccine significantly protected canaries against natural infection with Plasmodium relictum in the year of vaccination. In the following year, survivors in the vaccinated group suffered much higher mortality than unvaccinated birds that had survived exposure in year 1, presumably because vaccine-induced immunity prevented acquisition of protective natural immunity. Wild birds have developed a suite of trade-offs that allow them to be successful under a particular set of conditions. Environmental cues, such as photoperiod, may guide the timing of these trade-offs. Our world is changing rapidly and dramatically, especially for many wild species. With rapid anthropogenic alterations, such as climate change and environmental contamination, cues that were reliable may no longer be associated with adaptive outcomes (Schlaepfer et al. 2002). If birds are trapped by their evolutionary response to cues, they may find themselves equipped with attributes that are no longer optimal. Schlaepfer et al. (2002) used the term “evolutionary trap” for decisions that are now maladaptive because of a sudden anthropogenic disruption. For instance, the optimal time for reproduction by seasonally breeding birds matches peak food supply with peak nestling demand. If birds schedule reproduction based on photoperiod while food supply is determined by temperature, a mismatch in timing may result in peak nestling demand occurring while food supplies are declining, with serious consequences for fitness (e.g., Thomas et al. 2001). The effect of this type of evolutionary trap on parasitism has not been explored, but mismatches between the phenology of parasites or disease vectors and birds, as well as range expansion by parasites as a result of
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climate change and interactions among parasites and contaminants, could result in parasites assuming different or greater significance in altered environments. In summary, although parasitism is a universal phenomenon in wild birds and many parasites have been observed and described, the information is still fragmentary and largely descriptive in nature. Little is known about the effect of most parasites on their hosts and almost nothing is known about interactions among the parasites that make up parasite assemblages or communities. The cost of parasites to their hosts is difficult to measure, but using energy as a currency may be a fruitful way to understand how costs are incurred, why birds must make trade-offs that influence both their exposure and resistance to parasites, and how being parasitized may affect basic life history traits including reproduction and susceptibility to predation. Parasitism can never be considered in isolation; it must always be considered in terms of the context in which it is occurring and this consideration must include the potential effects of anthropogenic changes. LITERATURE CITED Anderson, R. M., and R. M. May. 1978. Regulation and stability of host–parasite population interactions. I. Regulatory processes. Journal of Animal Ecology 47:219. Bensch, S., J. Waldenstr¨om, N. Jonz´en, H. Westerdahl, B. Hansson, D. Sejberg, and D. Hasselquist. 2007. Temporal dynamics and diversity of avian malarial parasites in a single host species. Journal of Animal Ecology 76:112. Bize, P., A. Roulin, L.-F. Bersier, D. Pfluger, and H. Richner. 2003. Parasitism and developmental plasticity in Alpine swift nestlings. Journal of Animal Ecology 72:633. Boots, M., and R. G. Bowers. 2004. The evolution of resistance through costly acquired immunity. Proceedings of the Royal Society of London, Series B 271:715. Bush, A. O., and J. C. Holmes. 1986. Intestinal helminths of lesser scaup ducks: An interactive community. Canadian Journal of Zoology 64:142. Dawson, A., S. A. Hinsley, P. N. Ferns, R. H. G. Bonser, and L. Eccleston. 2000. Rate of moult affects feather quality: A mechanism linking current reproductive effort to future survival. Proceedings of the Royal Society of London, Series B 267:2093. Deerenberg, C., V. Apanius, S. Daan, and N. Bos. 1997. Reproductive effort decreases antibody responsiveness. Proceedings of the Royal Society of London, Series B 264:1021. Delahay, R. J., J. R. Speakman, and R. Moss. 1995. The energetic consequences of parasitism: Effects of a developing infection of Trichostrongylus tenuis
(Nematoda) on red grouse (Lagopus lagopus scoticus) energy balance, body weight, and condition. Parasitology 110:473. Hanssen, S. A., I. Folstad, K. E. Erikstad, and A. Oksanen. 2003. Costs of parasites in common eiders: Effects of antiparasite treatment. Oikos 100:105. Hanssen, S. A., D. Hasselquist, I. Folstad, and K. E. Erikstad. 2004. Costs of immunity: Immune responsiveness reduces survival in a vertebrate. Proceedings of the Royal Society of London, Series B 271:925. Hoodless, A. N., K. Kurtenbach, P. A. Nuttall, and S. E. Randolph. 2002. The impacts of ticks on pheasant territoriality. Oikos 96:245. Howe, F. P. 1992. Effects of Protocalliphora braueri (Diptera: Calliphoridae) parasitism and inclement weather on nestling sage thrashers. Journal of Wildlife Diseases 28:141. Hudson, P. J., and A. P. Dobson. 1997. Host–parasite processes and demographic consequences. In Host–Parasite Evolution. General Principles and Avian Models, D. H. Clayton and J. Moore (eds). Oxford University Press, Oxford, pp. 128–154. Hudson, P. J., A. P. Dobson, and D. Newborn. 1992a. Do parasites make prey vulnerable to predation? Red grouse and parasites. Journal of Animal Ecology 61:681. Hudson, P. J., D. Newborn, and A. P. Dobson. 1992b. Regulation and stability of a free-living host–parasite system: Trichostrongylus tenuis in red grouse. I. Monitoring and parasite reduction experiments. Journal of Animal Ecology 61:477. Hudson, P. J., A. P. Dobson, and D. Newborn. 1998. Prevention of population cycles by parasite removal. Science 282:2256. Lafferty, K. D. 1999. The evolution of trophic transmission. Parasitology Today 15:111. Leopold, A. 1933. Game Management. Charles Scribner’s Sons, New York. Lochmiller, R., and C. Deerenberg. 2000. Trade-offs in evolutionary immunology: Just what is the cost? Oikos 88:87. Martin, L. B., II, A. Scheuerlein, and M. Wikelski. 2003. Immune activity elevates energy expenditure of house sparrows: A link between direct and indirect costs? Proceedings of the Royal Society of London, Series B 270:153. Marzal, A., F. de Lope, C. Navarro, and A. P. Møller. 2005. Malarial parasites decrease reproductive success: An experimental study in a passerine bird. Oecologia 142:541. May, R. M., and R. M. Anderson. 1978. Regulation and stability of host–parasite population interactions. II. Destabilizing processes. Journal of Animal Ecology 47:249.
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Parasitism: Costs and Effects McCutchan, T. F., K. C. Grim, J. Li, W. Weiss, D. Rathore, M. Sullivan, T. K. Graczyk, S. Kumar, and M. R. Cranfield. 2004. Measuring the effects of an ever-changing environment on malaria control. Infection and Immunity 72:2248. Merino, S, J. Moreno, J. J. Sanz, and E. Arriero. 2000. Are avian blood parasites pathogenic in the wild? A medication experiment in blue tit* (Parus caeruleus). Proceedings of the Royal Society of London, Series B 267:2507. Metcalfe, N. B., and P. Monaghan. 2001. Compensation for a bad start: Grow now, pay later? Trends in Ecology and Evolution 16:255. Møller, A. P. 1994. Parasites as an environmental component of reproduction in birds as exemplified by the swallow Hirundo rustica. Ardea 82:161. Moore, J., and D. H. Clayton. 1997. Conclusion: Evolution of host–parasite interactions. In Host–Parasite Evolution: General Principles and Avian Models, D. H. Clayton and J. Moore (eds). Oxford University Press, Oxford, pp. 370–376. Navarro, C., F. de Lope, A. Marzal, and A. P. Møller. 2004. Predation risk, host immune response, and parasitism. Behavioral Ecology 15:629. Odum, E. P. 1993. Ecology and Our Endangered Life-Support System, 2nd ed. Sinauer Associates Inc., Sunderland, MA. P´erez, T. M., and W. T. Atyeo. 1984. Site selection of feather and quill mites of Mexican parrots. In Acarology VI, D. A. Griffiths and C. E. Bowman (eds). Ellis Horwood, Chichester, UK, pp. 563–570. Price, P. W. 1980. Evolutionary biology of parasites. Princeton University Press, Princeton, NJ. Quinn, J. L., and W. Cresswell. 2004. Predator hunting behaviour and prey vulnerability. Journal of Animal Ecology 73:143. Schlaepfer, M. A., M. C. Runge, and P. W. Sherman. 2002. Ecological and evolutionary traps. Trends in Ecology and Evolution 17:474. Shaw, D. J., B. T. Grenfell, and A. P. Dobson. 1998. Patterns of macroparasite aggregation in wildlife host populations. Parasitology 117:597. Spencer, K. A., K. L. Buchanan, S. Leitner, A. R. Goldsmith, and C. K. Catchpole. 2005. Parasites
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affect song complexity and neural development in a songbird. Proceedings of the Royal Society of London, Series B 272:2037. Spurrier, M. F., M. S. Boyce, and B. F. J. Manly. 1991. Effects of parasites in mate choice by captive sage grouse. In Bird–Parasite Interactions. Ecology, Evolution, and Behaviour, J. E. Loye and M. Zuk (eds). Oxford University Press, Oxford, pp. 390–398. Sulzbach, D., and F. Cooke. 1978. Elements of non-randomness in mass-captured samples of snow geese. Journal of Wildlife Management 42:437. Temple, S. A. 1987. Do predators always capture substandard individuals disproportionately from prey populations? Ecology 68:669. Thomas, D. W., J. Blondel, P. Perret, M. M. Lambrechts, and J. R. Speakman. 2001. Energetic and fitness costs of mismatching resource supply and demand in seasonally breeding birds. Science 291:2598. Tompkins, D. M., and M. Begon. 2000. Parasites can regulate wildlife populations. Parasitology Today 15:311. Tompkins, D. M., A. P. Dobson, P. Arneberg, M. E. Begon, I. M. Cattadori, J. V. Greenman, J. A. P. Heesterbeek, P. J. Hudson, D. Newborn, A. Pugliese, A. P. Rizzoli, R. Ros`a, F. Rosso, and K. Wilson. 2001a. Parasites and host population dynamics. In The Ecology of Wildlife Diseases, P. J. Hudson, A. Rizzoli, B. T. Grenfell, H. Heesterbeek, and A. P. Dobson (eds). Oxford University Press, Oxford, pp. 45–62. Tompkins, D. M., J. V. Greenman, and P. J. Hudson. 2001b. Differential impact of a shared nematode parasite on two gamebird hosts: Implications for apparent competition. Parasitology 122:187. Tschirren, B., P. S. Fitze, and H. Richner. 2003. Sexual dimorphism in susceptibility to parasites and cell-mediated immunity in great tit nestlings. Journal of Animal Ecology 72:839. Whitaker, S., and J. Fair. 2002. The costs of immunological challenge to developing mountain chickadees, Poecile gambeli, in the wild. Oikos 99:161. Zera, A. J., and L. G. Harshman. 2001. The physiology of life history trade-offs in animals. Annual Review of Ecology and Systematics 32:95.
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Section II: Protozoa
Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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2 Haemoproteus Carter T. Atkinson INTRODUCTION The species of Haemoproteus that infect birds are vector-transmitted intraerythrocytic parasites closely allied to the true malarial parasites of vertebrates. Unlike their close relatives in the genus Plasmodium, they undergo asexual reproduction or merogony within tissues rather than circulating erythrocytes. They are some of the most common and widespread blood parasites of wild birds, yet their potential significance as disease agents in wild bird populations is largely unknown. They are receiving increasing attention by avian ecologists as models for testing evolutionary theories about effects of disease on host fitness and sexual selection, but these efforts have been hampered by lack of basic knowledge about their life cycles, vectors, and epizootiology. Some species of Haemoproteus can be highly pathogenic and cause severe myositis in avian hosts, but well-documented cases are still rare. These include reports of disease associated with developing tissue stages in Northern Bobwhite (Colinus virginianus) (Gardiner et al. 1984; Cardona et al. 2002), Luzon Bleeding-heart (Gallicolumba luzonica) (Earle et al. 1993), Rock Pigeons (Columba livia) (Farmer 1965), House Sparrows (Passer domesticus biblicus) (Paperna and Gil 2003), Blossom-headed Parakeets (Psittacula roseata) (Miltgen et al. 1981), and Wild Turkeys (Meleagris gallopavo) (Atkinson and Forrester 1987). Some of these reports reflect abnormal host–parasite associations, where susceptible hosts were moved outside of their natural ranges and exposed to haemoproteid parasites from closely related host species.
HISTORY The species of Haemoproteus that infect birds were first observed on unstained blood smears along with other intraerythrocytic hemosporidian parasites by the Russian zoologist V. Ya. Danilewsky as “. . . clear, colorless, transparent vacuoles, variable in shape and size, in which are present several refractile glossyblack granules” (cited in Hewitt 1940). With the advent of Giemsa staining to differentiate parasites from host cells (Garnham 1966), the diversity and broad host range of these parasites became evident, but their host specificity, life cycles, and vectors were not known. Considerable confusion existed as to what comprised a species, and the taxonomy of this group has been in a continual state of flux for over a hundred years. Major historical milestones over the past century include discovery that Haemoproteus columbae of pigeons and doves can be transmitted by the bite of ectoparasitic hippoboscid flies (Sergent and Sergent 1906) and the discovery that ceratopogonid flies in the genus Culicoides can transmit other species of Haemoproteus (Fallis and Wood 1957). Early recognition of the sexual stages of Haemoproteus (MacCallum 1898), the hippoboscid vectors (Sergent and Sergent 1906), and preerythrocytic tissue stages of H. colombae (Arag˜ao 1908a) led to a number of classic investigations of the sporogonic or asexual stages of the parasite within the invertebrate vector and the preerythrocytic development of H. columbae within the avian host (Acton and Knowles 1914; Adie 1915, 1924; Coatney 1933). These formed an important framework during the first two decades of the twentieth century for understanding the life cycles and development of closely related haemosporidia in the genera Plasmodium and Leucocytozoon. The vast bulk of published studies on avian species of Haemoproteus over the past 50 years have been surveys and taxonomic descriptions by parasitologists and disease workers. It is only in the past few years that there has been a renaissance in interest in these parasites by avian ecologists and evolutionary biologists
SYNONYMS Haemosporidiosis. Infection with avian species of Haemoproteus is sometimes referred to as avian malaria, particularly in the recent ecological literature, but distinctive life history characteristics clearly distinguish them from the true malarial parasites in the genus Plasmodium (Valki¯unas et al. 2005).
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because ease of sampling wild birds by noninvasive blood collection makes them potentially good models for testing evolutionary hypotheses. The role that these parasites may play as pathogens in wild birds has been speculated about since their discovery, but it is only in the past 20 years that clear evidence that they can have some measurable effects on host survival and reproduction has accumulated. DISTRIBUTION Avian haemoproteids have a worldwide distribution in temperate and tropical climates. This wide distribution is most likely a function of the diverse habitats occupied by their ceratopogonid and hippoboscid vectors (Greiner et al. 1975). Haemoproteids have been recorded from most parts of the globe where hippoboscid and ceratopogonid vectors occur, including remote islands in the central Pacific (Work and Raymeyer 1996; Padilla et al. 2004). The greatest diversity of species occurs in the Holarctic, Ethiopian, and Oriental zoogeographic regions, with fewer numbers of species recorded from both the Neotropical and Australian
regions (Valki¯unas 2005). In both North and South America, haemoproteids tend to have a relatively uniform distribution across the continent and are virtually absent in the high arctic tundra, most likely because of the absence of suitable vectors (Greiner et al. 1975; White et al. 1978; Bennett et al. 1992). HOST RANGE Over 130 species of Haemoproteus have been reported from 72 families of birds, depending on authority (Peirce 2005; Valki¯unas 2005). Diversity in terms of number of distinct morphological forms and species is highest among the Coraciiformes (kingfishers), Piciformes (woodpeckers), and Galliformes, but the highest number of species occurs within the Passeriformes (perching birds) (Bennett 1993). Of interest is the wide disparity in occurrence of haemoproteid infections among the avian orders (Bennett 1993; Valki¯unas 2005). Haemoproteus has not been reported in many of the more primitive orders of birds, but is very common among the Passeriformes (Table 2.1). Some of these differences are clearly related to vector distribution
Table 2.1. Host distribution of avian haemoproteids by avian order. Avian order Sphenisciformes Gaviiformes Podicipediformes Procellariiformes Pelecaniformes Tinamiformes Apterygiformes Struthioniformes Ciconiiformes Falconiformes Strigiformes Anseriformes Galliformes Gruiformes Charadriiformes Columbiformes Psittaciformes Cuculiformes Caprimulgiformes Apodiformes Piciformes Coliiformes Coraciiformes Trogoniformes Passeriformes
Host species 16 4 21 100 57 47 3 8 124 296 162 154 270 203 339 323 344 153 106 414 402 6 202 39 5,211
Number examined 16 3 7 30 44 12 1 4 89 168 66 113 133 87 154 135 143 84 51 75 201 3 118 20 2,409
Number infected 0 0 0 0 0 0 0 0 40 83 49 56 74 47 36 87 43 40 8 20 60 0 13 7 2,047
Percent infected 0 0 0 0 0 0 0 0 45 49 74 50 56 54 23 64 30 48 16 27 30 0 11 35 85
Note: Data are summarized from Table 1 in Bennett (1993) and represent number of reported host species for each avian order that are infected with one or more species of Haemoproteus.
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Haemoproteus and abundance, with correspondingly low prevalence in seabirds and shorebirds that have limited exposure to hippoboscid or ceratopogonid flies (Mendes et al. 2005), while others may be related to differences in host resistance and immune competence (Ricklefs 1992; Sol et al. 2003). ETIOLOGY Members of this genus are classified as members of the phylum Apicomplexa, class Aconoidasida, order Haemospororida, family Plasmodiidae, and are defined primarily by their intraerthrocytic development, production of prominent golden-brown or black pigment granules from digestion of host hemoglobin, and absence of asexual reproduction in the circulating blood cells (Peirce 2000). Virtually all species in this genus are distinguished by morphology of the circulating gametocytes, their presumed host specificity, and by distinctive changes in host erythrocyte morphology (Figures 2.1a, b, and 2.2). Five different morphological types of gametocytes are recognized that differ in shape (round or elongated) and
(a)
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how far they reach around the erythrocyte nucleus (Figure 2.2). Recent phylogenetic analyses based on mitochondrial gene sequences have placed Haemoproteus as a polyphyletic group within the same clade as Plasmodium (Perkins and Schall 2002). More recent analyses based on four genes find that the avian haemoproteids fall into two clades that are sister to Plasmodium: a basal group of columbiform parasites that uses hippoboscid flies as vectors and a second distinct group that is transmitted by ceratopogonid flies (Martinsen et al. 2008). These new analyses support the proposal by Bennett et al. (1965) to subdivide the genus, keeping columbiform parasites transmitted by hippoboscid flies in the genus Haemoproteus and moving the bulk of species that are likely transmitted by ceratopogonid flies into the genus Parahaemoproteus (Martinsen et al. 2008). While this distinction is currently made at the level of subgenus (Valki¯unas 2005), these recent phylogenetic studies suggest that the proposal by Bennett et al. (1965) should be revived. The most recent taxonomic revisions of this genus are by Peirce (2005) and Valki¯unas (2005). Peirce
(b)
Figure 2.1. Gametocytes of Haemoproteus meleagridis in erythrocytes of an experimentally infected domestic turkey. (a) After release from preerythrocytic meronts, merozoites (arrows) invade erythrocytes and develop into mature gametocytes. Intraerythrocytic merozoites have a large vacuole and small nucleus. As merozoites transform into young gametocytes (G), they become elongated and sausage-shaped, eventually encircling the erythrocyte nucleus. As many as seven gametocytes may be found within individual erythrocytes in intense infections. (b) Gametocytes (G) reach maturity within 7 days after invading erythrocytes. Pigment granules (arrows) become visible only during later stages of development. Giemsa stain, bar = 10 μm. Reproduced from Atkinson (1991a), with permission of the Journal of Vector Ecology.
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(a)
(b)
(c)
(d)
(f)
(e)
(g)
Figure 2.2. Five basic morphological forms of the mature gametocytes of avian species of Haemoproteus: (a) normal erythrocyte, (b) microhalteridial gametocyte, (c, d) halteridial gametocyte, (e) circumnuclear gametocyte, (f) rhabdosomal gametocyte, and (g) discosomal gametocyte. Reproduced from Bennett et al. (1988), with permission of the Journal of Natural History and Taylor & Francis Ltd. (http://www.informaworld.com).
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Haemoproteus (2005) lists 147 species in 72 avian families, while Valki¯unas synonymizes some species based on host range and lists 132 valid species. Early efforts by Gorden Bennett and coworkers to make some taxonomic sense out of the bewildering diversity of avian haemoproteids led to separation of morphologically similar forms by host family, based on limited experimental evidence indicating that species are specific to family (Bennett and Peirce 1988). As is the case with closely related parasites in the genus Leucocytozoon (Chapter 4), much of this evidence was based on small sample sizes and only a few attempts to actually infect members of different host families and orders (Valki¯unas 2005). Problems with this taxonomy have been summarized by Valki¯unas (2005), and he argues effectively that available evidence only supports specificity to level of host order. Our understanding of the relationships between traditional morphological species and parasite lineages defined by mitochondrial and nuclear gene sequences is rapidly evolving. Recent studies of diversity of mitochondrial and nuclear genes of avian haemoproteids suggest that the true number of species may be several orders of magnitude higher with multiple parasite lineages that can coexist within the same host and with host ranges that extend well beyond single avian families (Bensch et al. 2000, 2004; Ricklefs and Fallon 2002). As a result, many traditional species defined by morphological characteristics and host family may be composed of multiple cryptic species, while many others that are defined only by occurrence in different host families will need to be synonymized. It is likely that major strides in our understanding of the taxonomy, host specificity, and evolutionary relationships among these parasites and closely related species in the genus Plasmodium will occur in future years as molecular data are reconciled with life history characteristics of these organisms. EPIZOOTIOLOGY The complex life cycle of Haemoproteus involves both sexual (gametogenesis and fertilization) and asexual (sporogony) reproduction in the vector and asexual reproduction (merogony) in the avian host. Proven vectors of avian haemoproteids include both ceratopogonid flies in the genus Culicoides and ectoparasitic hippoboscid flies (Table 2.2). The sexual cycle begins when a blood meal containing mature sexual stages of the parasite, female macrogametocytes and male microgametocytes, is taken from an infected host. The sexual stages undergo gametogenesis and fertilization in the midgut of the vector and produce a motile zygote called the ookinete. Ookinetes subsequently penetrate the midgut wall and develop under the midgut basal
17
lamina as spherical oocysts during the asexual sporogonic cycle. Development of the parasite in both vectors is similar, but size of oocysts, number of sporozoites produced, and duration of sporogony differs. In ceratopognid flies, oocysts measure approximately 10 μm in diameter, while in hippoboscid flies oocysts are considerably larger and reach diameters of approximately 40 μm (Adie 1924; Fallis and Bennett 1960; Atkinson 1991b). Sporogony typically takes 4–6 days in ceratopogonid flies, eventually producing fewer than 100 sporozoites that bud from a single sporoblast. Sporogony in hippoboscid flies typically takes up to 10 days, eventually producing thousands of sporozoites that bud from multiple sporoblasts within the oocysts (Adie 1915, 1924). Oocysts subsequently rupture, releasing sporozoites into the haemocoel of the insect. These invade the salivary glands and pass through the salivary ducts during the next blood meal. The factors that affect the ability of particular species of Haemoproteus to develop in a particular species of arthropod vector are poorly understood. It is clear that individual species of avian Haemoproteus can be transmitted by a number of different hippoboscid or ceratopogonid vectors (Table 2.2), but successful development as measured by ability to complete sporogony and produce sporozoites that can reach the salivary glands varies in each species of Culicoides (Atkinson 1991a; Valki¯unas et al. 2002). It is not known whether blocks in development occur in the midgut, during passage through the peritrophic membrane that surrounds the blood meal during digestion, or within the midgut epithelium. It has never been demonstrated by experimental methods that haemoproteids transmitted by hippoboscid flies can also be transmitted by ceratopogonid flies, although complete development of Haemoproteus lophortyx from Northern Bobwhites in Culicoides bottimeri, Stilbometopa impressa, and Lynchia hirsuta suggests that this is possible (O’Roke 1930; Tarshis 1955; Mullens et al. 2006). However, the original experimental work on hippoboscid transmission of H. lophortyx (O’Roke 1930; Tarshis 1955) was done in facilities that were not adequately screened to prevent entry by ceratopogonid flies (Valki¯unas 2005; Table 2.2). Among other species of Haemoproteus, the rare occurrence of hippoboscid flies on Mourning Doves (Zenaida macroura) and high prevalence of Haemoproteus sacharovi strongly suggest that ceratopogonid flies may be involved in transmission of this parasite, but this possibility has not been investigated (Bennett and Peirce 1990). Given the results of recent phylogenetic studies (Martinsen et al. 2008), experimental tests of vector specificity of these two groups of haemoproteids should be pursued.
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Table 2.2. Known species of hippoboscid (Lynchia, Microlynchia, Ornithomyia, Pseudolynchia, Stilbometopa) and ceratopogonid (Culicoides) flies that can support complete asexual sporogonic development of Haemoproteus. Species
Host order
Vector
Haemoproteus nettionis Haemoproteus columbae
Anseriformes Columbiformes
Haemoproteus sacharovi* Haemoproteus maccallumi Haemoproteus turtur† Haemoproteus palumbis Haemoproteus lophortyx‡
Columbiformes Columbiformes Columbiformes Columbiformes Galliformes
Haemoproteus mansoni Haemoproteus meleagridis§
Galliformes Galliformes
Haemoproteus handai Haemoproteus velans
Psittaciformes Passeriformes
Haemoproteus fringillae
Passeriformes
Haemoproteus danilewskii
Passeriformes
Haemoproteus balmorali Haemoproteus dolniki Haemoproteus tartakovskyi Haemoproteus belopolskyi Haemoproteus lanii
Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes
Culicoides downesi Pseudolynchia canariensis Pseudolynchia brunnea Microlynchia pusilla Pseudolynchia canariensis Pseudolynchia canariensis Pseudolynchia canariensis Ornithomyia aviculria Stilbometopa impressa Lynchia hirsuta Culicoides bottimeri Culicoides sphagnumensis Culicoides edeni Culicoides hinmani Culicoides arboricola Culicoides haematopotus Culicoides knowltoni Culicoides nubeculosus Culicoides stilobezziodes Culicoides sphagnumensis Culicoides crepuscularis Culicoides stilobezziodes Culicoides sphagnumensis Culicoides impunctatus Culicoides crepuscularis Culicoides stilobezziodes Culicoides sphagnumensis Culicoides edeni Culicoides knowltoni Culicoides arboricola Culicoides impunctatus Culicoides impunctatus Culicoides impunctatus Culicoides impunctatus Culicoides impunctatus
Authors Fallis and Wood (1957) Sergent and Sergent (1906) Arag˜ao (1908b) Arag˜ao (1916) Huff (1932) Huff (1932) Rashdan (1998) Baker (1963, 1966) O’Roke (1930) Tarshis (1955) Mullens et al. (2006) Fallis and Bennett (1960) Atkinson et al. (1983) Atkinson et al. (1983) Atkinson et al. (1983) Atkinson (1988) Atkinson (1988) Miltgen et al. (1981) Khan and Fallis (1971) Khan and Fallis (1971) Fallis and Bennett (1961) Fallis and Bennett (1961) Fallis and Bennett (1961) Valki¯unas (1997) Bennett and Fallis (1960) Bennett and Fallis (1960) Fallis and Bennett (1961) Garvin and Greiner (2003a) Garvin and Greiner (2003a) Garvin and Greiner (2003a) Valki¯unas et al. (2002) Valki¯unas et al. (2002) Valki¯unas et al. (2002) Valki¯unas and Iezhova (2004) Valki¯unas and Iezhova (2004)
* Circ*mstantial evidence indicates that one or more species of Culicoides may also be involved in natural transmission of Haemoproteus sacharovi (Bennett and Peirce 1990). † Haemoproteus turtur is recognized as distinct from Haemoproteus columbae by Valki¯unas (2005), but considered a synonym of H. columbae by Peirce (2005). ‡ Tarshis (1955) was unable to demonstrate sporozoites of Haemoproteus lophortyx in Stilbometopa impressa and Lynchia hirsuta in spite of repeated attempts to infect them in the laboratory, but did successfully transmit H. lophortyx when flies were allowed to bite uninfected birds. Valki¯unas (2005) suggests that experimental cages may not have been impervious to ceratopogonid flies, based on unusually long prepatent periods for experimental infections and use of screened outdoor aviaries. O’Roke (1930), however, describes oocysts and sporozoites in L. hirsuta that fed on infected quail. Given the recent finding that Culicoides bottimeri is a likely natural vector, experiments with both S. impressa and L. hirsuta should be repeated. § Haemoproteus meleagridis is considered a junior synonym of Haemoproteus canachites by Valki¯unas (2005).
18
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Haemoproteus Complete life cycles are known for only a handful of avian haemoproteids and we still have only a rudimentary knowledge about the preerythrocytic development of these parasites. Haemoproteus columbae from pigeons and doves, Haemoproteus meleagridis from Wild Turkeys, and Haemoproteus danilewskii from Blue Jays (Cyanocitta cristata) have received the most detailed experimental study. Endogenous development of all three of these species begins when infective sporozoites are inoculated at the site where the vector takes a blood meal. These sporozoites develop within cells of the lymphoid–macrophage system, capillary endothelium, and/or myofibroblasts, undergoing one or more generations of asexual reproduction or merogony before penetrating circulating erythrocytes (Mohammed 1965; Atkinson et al. 1986). Here they develop as gametocytes, becoming infective to vectors within 7–10 days after invading the blood cells. At least two generations of preerythrocytic merogony occur in skeletal and cardiac muscle of domestic turkeys experimentally infected with H. meleagridis. The first begins when infective sporozoites invade capillary endothelial cells and myofibroblasts and develop into thin-walled round or oval meronts measuring 12– 20 μm in diameter. These produce long, slender merozoites between 5 and 8 days postinfection that subsequently invade new capillary endothelial cells in skeletal and cardiac muscle and develop as secondgeneration meronts. Early second-generation meronts are 5–8 μm in diameter and 28 μm in length. These grow rapidly to form large, fusiform, thick-walled megalomeronts measuring up to 500 μm in length (Figure 2.3). Megalomeronts reach maturity at 17 days postinfection and rupture to release small spherical merozoites that invade erythrocytes and develop into gametocytes (Figure 2.1a). Mature gametocytes that completely encircle the host erythrocyte nucleus develop within 7–10 days after red blood cells are invaded (Figure 2.1b). Parasitemias reach their peak intensity in the peripheral circulation at approximately 21 days postinfection and fall rapidly within 7 days to low intensities. A second, smaller peak in parasitemia may occur at approximately 35 days postinfection (Atkinson et al. 1986). The number of generations of preerythrocytic merogony has not been defined for H. columbae and H. danilewskii, but it is likely that they also undergo two or more cycles of asexual reproduction before invading erythrocytes. In these two species, the parasites invade capillary endothelial cells of the lungs where they undergo preerythrocytic development to form thin-walled, oval or branching meronts that radiate along pulmonary capillaries (Mohammed 1965; Garnham 1966; Garvin et al. 2003a; Valki¯unas 2005). Similar, thin-walled branching meronts have
19
Figure 2.3. Megalomeront of Haemoproteus meleagridis from the pectoral muscle of a naturally infected Wild Turkey (Meleagris gallopavo). The megalomeront is surrounded by a thick, hyaline wall (arrowheads) and is packed with spherical merozoites. Muscle fibers surrounding the megalomeront are swollen, pale, and hyaline and contain scattered basophilic granules (arrows). Note adjacent normal tissue (*). Hematoxylin and eosin, bar = 50 μm. Reproduced from Atkinson and Forrester (1987), with permission of the Journal of Wildlife Diseases.
been reported in a variety of other naturally infected avian hosts (Figure 2.4). Thick-walled megalomeronts have been reported in Luzon Bleeding-hearts (Gallicolumba luzonica) infected with H. columbae (Earle et al. 1993), but their relationship to the thin-walled, branching meronts of H. columbae described from Rock Pigeons is unclear and possibly related to presence of a mixed infection with another unidentified parasite (Peirce et al. 2004). Among haemoproteids transmitted by Culicoides, prepatent periods vary from 11 to 12 days for
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Parasitic Diseases of Wild Birds
Figure 2.4. Thin-walled, irregularly shaped meront in lung tissue from a White-throated Sparrow (Zonotrichia albicollis) infected naturally with Haemoproteus coatneyi. Reproduced from Khan and Fallis (1969), with permission of the Canadian Journal of Zoology.
Haemoproteus belopolskyi of Blackcaps (Sylvia atricapilla) (Valki¯unas and Iezhova 2004), from 11 to 14 days for Haemoproteus velans of woodpeckers (Khan and Fallis 1971), 14 days for Haemoproteus mansoni of Ruffed Grouse (Bonasa umbellus) (Fallis and Bennett 1960), approximately 16 days for Haemoproteus nettionis of ducks (Fallis and Wood 1957), 14 days for H. danilewskii of Blue Jays (Garvin et al. 2003a), and 17 days for H. meleagridis of Wild Turkeys (Atkinson et al. 1986). Among haemoproteids transmitted by hippoboscid flies, the prepatent period ranges from 17 to 37 days for H. columbae of Rock Pigeons and is about 14 days for Haemoproteus palumbis of Common Wood-Pigeons (Columba palumbus) (Baker 1966). Like the species of Haemoproteus that are transmitted by Culicoides, merozoites in circulating erythrocytes develop to mature microgametocytes and macrogametocytes that encircle the erythrocyte nucleus within approximately 5–10 days. Gametocyte numbers peak in the peripheral circulation approximately 10–20 days after first appearing in the circulation and then decline in numbers. Among species of Haemoproteus transmitted by ceratopogonid flies, transmission is seasonal and limited to the spring and summer months in more temperate parts of their range (Bennett and Fallis 1960), but can occur throughout the year in subtropical habitats in Florida and most likely other parts of the world where suitable vectors are present year round (Atkinson et al.
1988a). In temperate North America, by contrast, transmission of H. columbae by hippoboscid flies is seasonal and closely correlated with changes in vector populations, generally increasing in the fall and winter months and then declining as vector density decreases (Klei and DeGiusti 1975). More limited data from tropical and subtropical parts of the world where populations of hippoboscid flies remain more constant indicate that high rates of transmission and high prevalences of infection can be maintained throughout the year (Ayala et al. 1977; Sol et al. 2000). The role that host migratory behavior plays in cycles of transmission of avian haemoproteids is significant because of the potential of long distance migrants to disperse parasites both within and between continental landmasses (Laird 1960; Waldenstr¨om et al. 2002; Hasselquist et al. 2007). Limited information from the Nearctic and Palearctic indicates that some species of avian haemoproteids are transmitted on the breeding grounds, while others are transmitted in wintering areas in the tropics and subtropics, while others may be transmitted in both locations. This suggests that transmission may be linked in some cases to particular geographic locations or vector–parasite associations (Valki¯unas 1993; Valki¯unas and Iezhova 2001; Waldenstr¨om et al. 2002; Garvin et al. 2003b, 2004; Hasselquist et al. 2007; Hellgren et al. 2007b). Within individual hosts, intensity of infection varies after the initial acute phase and appears to be influenced by the complex interplay of host immunity, seasonal changes in photoperiod, and hormonal changes associated with reproduction. In temperate climates, a seasonal increase in intensity, termed the spring relapse, coincides with the breeding season when populations of blood-sucking insects typically increase and recently fledged susceptible birds are increasing in the population (Atkinson and van Riper 1991; Valki¯unas et al. 2004). Relapse of chronic Plasmodium infections can be triggered by corticosterone (Applegate and Beaudoin 1970) and other experimental evidence suggests that increases in photoperiod and subsequent physiological changes in levels of hormones such as melatonin that regulate circadian rhythms may also be important stimuli for initiating relapses among species of Haemoproteus (Valki¯unas et al. 2004). Other factors affecting intensity include stressmediated changes in the immune system that are associated with reproductive effort (Siikam¨aki et al. 1997), food availability (Appleby et al. 1999), concomitant infection with other parasites (Cox 1987), and exposure to predators (Navarro et al. 2004). Attempts to identify broad patterns and relationships in the prevalence of avian haemoproteids have met with variable success because of the diversity of this group of parasites. Much may depend on how prevalence data
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Haemoproteus are lumped, with positive relationships more evident where other species of hematozoans are included in the analyses. A wide variety of both intrinsic and extrinsic factors have been identified, including host specificity of the parasites (Bennett 1993), immune competency (Ricklefs 1992), host genotype (Bonneaud et al. 2006), host age and sex (Davidar and Morton 1993; Powers et al. 1994; McCurdy et al. 1998), geographic range of host species (Tella et al. 1999), whether or not host species are migratory (Bennett and Fallis 1960; Peirce and Mead 1978; Figuerola and Green 2000; Smith et al. 2004), plumage coloration (Yezerinac and Weatherhead 1995), and host foraging or nesting behavior (Greiner et al. 1975; Garvin and Remsen 1997). Extrinsic factors such as habitat, geographical region, and season are critically important because they can influence the distribution and abundance of vectors (Weatherhead and Bennett 1991, 1992; Sol et al. 2000; Mendes et al. 2005). Prevalence in the same host species can vary significantly across both large and small landscapes (Atkinson et al. 1988a; Sol et al. 2000; Wood et al. 2007), suggesting that vector distribution and abundance may be the most important determinant of prevalence. However, other factors may cause seasonal changes in parasite prevalence, including winter mortality in infected birds, and new infections associated with emergence of insect vectors and transmission to uninfected juvenile birds. CLINICAL SIGNS Clinical signs are usually not evident in low-intensity infections, but can become evident during acute phase infections when erythrocytic parasitemias and numbers of tissue meronts reach high intensities. Domestic turkey poults with experimental infections of H. meleagridis are lame in one or both legs and have lower weights and growth rates than do uninfected controls (Atkinson et al. 1988b). Similarly, Northern Bobwhites with natural infections of H. lophortyx are reluctant to move, have a ruffled, depressed appearance, and exhibit neurological signs such as loss of balance and difficulty walking (Cardona et al. 2002). Signs of infection in Rock Pigeons include weakness, anemia, and anorexia (Acton and Knowles 1914; Coatney 1933). Elevation in numbers of circulating lymphocytes, heterophils, basophils, eosinophils, and monocyte numbers has been observed in both natural and experimental infections with Haemoproteus, and it is likely that these increases represent a cell-mediated response to both erythrocytic and preerythrocytic stages of the parasite, particularly as the latter mature and rupture to release merozoites that invade erythrocytes (Ots and H˜orak 1998; Garvin et al. 2003a). No significant overall difference in plasma protein concentra-
21
tion, hemoglobin concentration, packed cell volume, or weight was observed between infected and uninfected Blue Jays (Garvin et al. 2003a). Other studies have also failed to report significant anemia in infections with Haemoproteus, including H. meleagridis in experimentally infected domestic turkeys (Atkinson et al. 1988b) and Haemoproteus spp. in Great tit* (Parus major) (Ots and H˜orak 1998). By contrast, O’Roke (1930) and Cardona et al. (2002) detected severe anemia in California Quail (Callipepla californica) and captive Northern Bobwhites with natural infections of H. lophortyx. Severe regenerative anemia with marked polychromasia has also been reported in Snowy Owls (Bubo scandiacus) infected with Haemoproteus noctuae (Evans and Otter 1998) and in Snowy Owls, Tawny Owls (Strix aluco), and Great Horned Owls (Bubo virginianus) infected with Haemoproteus syrnii (Mutlow and Forbes 1999). Mechanisms responsible for development of anemia in these host species are not known, although there may be a fine balance between removal of parasitized erythrocytes by the spleen and their replacement with immature red blood cells (Atkinson et al. 1988b). When an infected host lacks the physiological resources to replace infected blood cells because of stress associated with reproduction or limited food resources, anemia may result. PATHOGENESIS AND PATHOLOGY Virtually nothing is known about the pathogenesis of haemoproteid infections because so little is known about their development within natural and experimental hosts. Few host responses have been associated with development of thin-walled branching meronts that frequently occur in lung tissue (Mohammed 1965; Baker 1966; Garnham 1966) (Table 2.3). In one of the most detailed studies to date, no host responses were associated with preerythrocytic meronts at day 31 postinfection in Blue Jays infected experimentally with H. danilewskii. However by day 57 postinfection, juvenile jays had lesions in liver, spleen, and lung tissue. These included periportal and random individual cell necrosis in liver and lymphocytic infiltrates and epithelial hyperplasia around tertiary bronchi in lung tissue. Histological changes in splenic tissue included hyperplasia of white pulp arteriolar endothelium, random necrosis of lymphocytes, and increases in the number of macrophages, plasma cells, and Mott cells (Garvin et al. 2003a). The authors suggested that the lesions developed only after meronts matured and ruptured. Severe myositis has been reported in association with thick-walled megalomeronts in a variety of avian species (Table 2.3). These lesions are associated with intact and ruptured megalomeronts and are grossly visible as white flecks or dark hemorrhagic streaks
Host order
Haemoproteus balearicae
Lung
Lungs, heart, spleen
Tissue
22
W W E
Columbiformes Haemoproteus sacharovi Columbiformes Haemoproteus maccallumi Haemoproteus danilewskii Haemoproteus passeris Haemoproteus ptilotis* Haemoproteus ptilotis* Haemoproteus attenuatus Haemoproteus coatneyi
Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes
W
W
W
W
W
W
Lungs, heart, liver, spleen, cecum, kidneys
Lungs, spleen
Liver
Heart and spleen
Lungs, liver
Liver, spleen, lung
Lung
Lung
Lungs, heart
W, E, D Lungs, rarely liver and spleen
W
E
Status
Columbiformes Haemoproteus palumbis
Columbiformes Haemoproteus columbae
Gruiformes
Haemoproteus nettionis
Parasite
Peirce (1976)
Garvin et al. (2003a)
Greiner (1971)
Greiner (1971)
Baker (1966)
Arag˜ao (1908b), Mohammed (1965), and Peirce et al. (2004)
Sibley and Werner (1984) Peirce (1973)
Citations
No
No
Khan and Fallis (1969)
Valki¯unas (2005)
Tissue displacement, Peirce et al. (2004) inflammation No Peirce et al. (2004)
No
Minor inflammation
No
No
None reported or tissue displacement, blockage of vessels No
No
No
Pathology
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Common Wood-Pigeon (Columba palumbus) Mourning Dove (Zenaida macroura) Mourning Dove (Zenaida macroura) Blue Jay (Cyanocitta cristata) House Sparrow (Passer domesticus) Noisy Miner (Manorina melanocephala) Noisy Friarbird (Philemon corniculatus) European Robin (Erithacus rubecula) White-throated Sparrow (Zonotrichia albicollis)
Black Crowned-Crane (Balearica pavonina) Rock Pigeon (Columba livia)
Thin-walled oval or branching meronts Wood Duck (Aix sponsa) Anseriformes
Host species
Table 2.3. Preerythrocytic meronts and host responses reported from wild (W), domestic (D), captive (C), or experimentally infected (E) avian hosts.
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Domestic chicken, Red Jungle Fowl (Gallus gallus) Luzon Bleeding-heart (Gallicolumba luzonica) Mourning Dove (Zenaida macroura) Blossom-headed Parakeet (Psittacula roseata) Monk Parakeet (Myiopsitta monachus) Parakeet (species not reported) Arthrocystis galli* Haemoproteus columbae Haemoproteus sacharovi Haemoproteus handai Undetermined* Undetermined*
Galliformes Columbiformes Columbiformes Psittaciformes Psittaciformes Psittaciformes
Haemoproteus meleagridis W, E Cardiac and skeletal muscle
Galliformes
23 C
C
C
W
C
D
Heart, gizzard
Cardiac and skeletal muscle Skeletal muscle
Cardiac and skeletal muscle, gizzard, proventriculus Gizzard
Skeletal and cardiac muscle
Skeletal muscle
Hemorrhage
Myopathy
Myopathy
No
Myopathy
Myopathy
Myopathy
Myopathy
Hepatic necrosis, hemorrhage, inflammation Inflammation
Borst and Zwart (1972) Fowler and Forbes (1972) and Walker and Garnham (1972) (continues)
Miltgen et al. (1981)
Farmer (1965)
Atkinson and Forrester (1987) and Atkinson et al. (1988b) Levine et al. (1970) and Opitz et al. (1982) Earle et al. (1993)
Commichau and Jonas (1977) and Kˇucera et al. (1982) Cardona et al. (2002)
Ferrell et al. (2007)
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C
Haemoproteus lophortyx
Heart, lungs, liver, kidneys, spleen
Galliformes
D
Northern Bobwhite (Colinus virginianus) Domestic Turkey, Wild Turkey (Meleagris gallopavo)
Undetermined*
Liver
Anseriformes
C
Muscovy Duck (Cairina moschata)
Thick-walled meglomeronts Lesser Flamingo Phoenicopteriformes Haemoproteus sp.† (Phoenicopterus minor)
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24
W W C C
Passeriformes Haemoproteus halcyonis Passeriformes Undetermined* Passeriformes Haemoproteus sp.† Passeriformes Haemoproteus sp.† Liver
Heart, skeletal muscle, gizzard Liver
Skeletal muscle
Kidney
Liver, lungs, kidney
Skeletal muscle
Tissue
Hepatic necrosis, hemorrhage, inflammation Hepatic necrosis, hemorrhage, inflammation
Myopathy
No
No
Inflammation
No
Pathology
Ferrell et al. (2007)
Ferrell et al. (2007)
Lederer et al. (2002)
Peirce et al. (2004)
Mutlow and Forbes (1999) Garnham (1966) and Paperna and Gil (2003) Garnham (1966)
Citations
Note: Two primary types of preerythrocytic meronts have been reported: thin-walled, oval or branching forms that are associated with limited host reaction; and thick-walled, round or fusiform forms that occur in skeletal, gizzard, and cardiac muscle, as well as liver, spleen, and lung tissue. The most definitive associations are in birds with experimental infections with Haemoproteus nettionis, Haemoproteus columbae, Haemoproteus meleagridis, and Haemoproteus danilewskii. Remaining examples should be viewed with caution since hosts may have been infected with more than one parasite and often did not have circulating gametocytes. Table includes hosts infected with megalomeronts of undetermined or questionable taxonomic status that are suspected to belong to species of Haemoproteus. ∗ No parasitemia observed, possibly Leucocytozoon or other undetermined protozoan. † Identity determined by PCR amplification and sequencing of parasite cytochrome b gene.
W
Passeriformes Undetermined*
W
Status
W
Haemoproteus syrnii
Parasite
Passeriformes Haemoproteus passeris
Strigiformes
Host order
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Montezuma Oropendola (Gymnostinops montezuma)
Snowy Owl (Bubo scandiacus) Israeli House Sparrow (Passer domesticus biblicus) Java Sparrow (Padda oryzivora) Sacred Kingfisher (Todiramphus sanctus) Pied Currawong (Strepera graculina) Green Jay (Cyanocorax yncas)
Host species
Table 2.3. (Continued)
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Haemoproteus
25
in tissue macrophages of the liver and spleen and enlargement of these organs (Atkinson et al. 1986, 1988b; Atkinson and Forrester 1987). Megalomeronts with associated muscle pathology have been reported in a variety of other naturally infected avian hosts, but their role in life cycles of specific species of Haemoproteus is difficult to determine without experimental studies (Peirce et al. 2004; Table 2.3).
Figure 2.5. Formalin-fixed pectoral muscle from a domestic turkey with an experimental infection with Haemoproteus meleagridis. Note the scattered white streaks (arrowheads) and darkened hemorrhagic areas (arrows) that correspond to megalomeronts in histological sections. Hematoxylin and eosin, bar = 0.5 cm. Reproduced from Atkinson et al. (1988b), with permission of the Journal of Parasitology.
in skeletal and cardiac muscle. The lesions superficially resemble those from infections with Sarcocystis (Figure 2.5). Microscopically, megalomeronts are surrounded by mixed inflammatory infiltrates composed of macrophages, heterophils, giant cells, and red blood cells, and adjacent muscle fibers are often necrotic and calcified (Miltgen et al. 1981; Atkinson et al. 1988b; Cardona et al. 2002) (Figures 2.6 and 2.7). Other lesions include extensive deposition of parasite pigment
DIAGNOSIS The gold standard for diagnosis of Haemoproteus is a Giemsa-stained thin blood smear where it is possible to demonstrate the presence of erythrocytic gametocytes with prominent golden-brown or black pigment granules and absence of erythrocytic meronts that are diagnostic for Plasmodium spp. Individual species are traditionally defined by morphology of intraerythrocytic gametocytes (Figure 2.2) and host specificity, but this will likely undergo extensive revision in future years. Molecular methods are beginning to be applied to differentiation of genera and identification of unique parasite lineages. Their high sensitivity make them valuable for identifying birds with very low intensity infections, but these methods have not been refined to the point where they can be used to distinguish individual species. Recent studies, though, suggest that this may eventually be feasible (Hellgren et al. 2007a; Valki¯unas et al. 2007). Species of Haemoproteus may be difficult to distinguish from avian species of Plasmodium, particularly in chronic infections where number of circulating gametocytes is low and where it may be difficult to determine whether the intracellular meronts characteristic of Plasmodium are present or absent. Several recent sets of primers designed to amplify portions of parasite mitochondrial genome can distinguish Haemoproteus and Plasmodium from Leucocytozoon (Hellgren et al. 2004) or all three genera from each other following restriction digests of polymerase chain reaction (PCR) products (Beadell and Fleischer 2005). However, sequencing of PCR products is necessary for identifying individual parasite lineages and determining phylogenetic relationships. The morphology of tissue stages is difficult to use alone for making accurate diagnosis of infection with Haemoproteus. The thin-walled oval or branching meronts that are characteristic of some species of columbiform haemoproteids are similar in morphology to tissue stages of both Leucocytozoon and Plasmodium. Megalomeronts of Haemoproteus may be difficult to distinguish from those of Leucocytozoon. A variety of megalomeronts have been reported as aberrant Leucocytozoon infections (Levine et al. 1970; Borst and Zwart 1972; Fowler and Forbes 1972; Walker and
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Figure 2.6. Ruptured megalomeront from pectoral muscle of a domestic turkey with an experimental infection with Haemoproteus meleagridis. Hemorrhagic megalomeront (M) is surrounded and partially filled by red blood cells (RBCs). Thrombi (T) with embedded RBCs are adjacent to or within the megalomeront. Hematoxylin and eosin, bar = 100 μm. Reproduced from Atkinson et al. (1988b), with permission of the Journal of Parasitology. Garnham 1972; Hartley et al. 1981; Simpson 1991; Pennycott et al. 2006) or possible Besnoitia infections (Bennett et al. 1993; Peirce et al. 2004), but erythrocytic parasites were absent and it is possible that some of these reports may prove to be tissue stages of Haemoproteus. The difficulties in making diagnoses from wild birds that may be infected with multiple species of haemosporidians have been discussed by Lederer et al. (2002) who pointed out that accurate association between megalomeronts and infection with Haemoproteus or Leucocytozoon in wild birds requires experimental studies. The recent use of molecular methods for diagnosis may help resolve some of these problems. For example, hepatic megalomeronts in three species of captive birds in a zoo collection in Texas were recently shown to be associated with infection with an undetermined species of Haemoproteus by PCR amplification of a portion of the parasite mitochondrial cytochrome b gene (Ferrell et al. 2007). Haemoproteus appears to be antigenically distinct from Plasmodium and crude antigen extracts have been used to develop an ELISA test for H. columbae in Rock Pigeons (Graczyk et al. 1994). The specificity
and sensitivity of this serological test with other avian haemoproteids are not known, but they may prove useful for making genus level diagnoses in birds with lowintensity infections. IMMUNITY Virtually nothing is known about immune mechanisms in haemoproteid infections. Spontaneous recovery from infections with H. columbae has been reported in Rock Pigeons, with no immunity conferred to second infection (Sergent and B´equet 1914; Ahmed and Mohammed 1978). In most cases birds probably remain infected for long periods of time and have spontaneous relapses that may decrease in frequency, eventually leading to recovery (Coatney 1933; Ahmed and Mohammed 1978). Experimental evidence for this is very limited, however, and restricted to H. columbae of Rock Pigeons. Limited experimental data indicate that birds with chronic infections have concomitant immunity where a persistent chronic infection stimulates immunity to reinfection with hom*ologous parasites of the same species (Coatney 1933; Ahmed and Mohammed
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separation of most commercial poultry facilities from habitats where Wild Turkeys range. There are multiple reports of pathogenic infections of Haemoproteus in pigeons and doves. These are usually associated with high parasitemias (Coatney 1933) and the occurrence of megalomeronts (Farmer 1965; Earle et al. 1993), but most individuals appear to be able to tolerate very high parasitemias with no clinical signs of infection. Major outbreaks of infection with H. lophortyx have been reported in Northern Bobwhite raised in California where the natural reservoir host is California Quail. Outbreaks occur during warm weather when ceratopogonid populations increase (Cardona et al. 2002). Similarly, there have been a substantial number of reports of lethal Leucocytozoon-like infections affecting captive birds, particularly parakeets, that may actually be caused by species of Haemoproteus (Fowler and Forbes 1972; Smith 1972; Walker and Garnham 1972; Simpson 1991; Pennycott et al. 2006; Ferrell et al. 2007). In all these instances, captive birds were introduced to areas outside of their natural range.
Figure 2.7. Intact megalomeront from pectoral muscle of a domestic turkey with an experimental infection with Haemoproteus meleagridis. Megalomeront (M) is surrounded by giant cells (arrowheads) and hyaline and necrotic muscle fibers (arrows). Note thick hyaline wall (W) surrounding the megalomeront. Hematoxylin and eosin, bar = 50 μm. Reproduced from Atkinson et al. (1988b), with permission of the Journal of Parasitology.
1978). The relapses associated with chronic infections most likely originate from persistent tissue stages, but this has not been proven by experimental studies.
PUBLIC HEALTH CONCERNS Infected birds pose no health hazards to humans.
DOMESTICATED ANIMAL HEALTH CONCERNS Haemoproteus meleagridis of Wild Turkeys is a potential threat to domestic turkey production, but in practice this has never materialized—possibly because of
WILDLIFE POPULATION IMPACTS The effects of individual Haemoproteus infections are difficult to discern in wild hosts. The vast majority of studies are correlational and the avian hosts under investigation are frequently infected with other hematozoan parasites, including Leucocytozoon, Plasmodium, and Trypanosoma. In a thorough review of over 5,000 papers on avian blood parasites, Bennett et al. (1993) found that only about 4% reported mortality or pathogenicity in birds, with most dealing with domestic birds or birds in zoological collections. Mortality associated with Haemoproteus and other blood parasites in wild birds probably occurs more frequently than reported because sick individuals may be difficult to find for sampling or recover from the wild for necropsy. Epizootics are often hard to document for small passerines in areas where carcasses are rapidly scavenged (Bennett et al. 1993). Since the life cycle of Haemoproteus requires a vector, experimental manipulations of naturally acquired infections in the wild are difficult. One approach that has been successful is use of a single subcutaneous dose of primaquine to control H. majoris and Leucocytozoon majoris in naturally infected Eurasian Blue tit* (Cyanistes caeruleus) (Merino et al. 2000). The treated group had higher fledging success and lower nestling mortality, but the relative contributions of Haemoproteus and Leucocytozoon to decreased fledging success were not determined. Some studies have reported reduced survival in birds infected with Haemoproteus (Nordling et al. 1998;
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Dawson and Bortolotti 2000; H˜orak et al 2001; Sol et al. 2003) and negative effects on indices of immunity, condition, and reproductive success of their hosts (Allander and Bennett 1995; Ots and H˜orak 1998; Merino et al. 2000; Sanz et al. 2001). While some studies suggest that these changes may be reflected in plumage coloration (H˜orak et al. 2001), others have found limited association (Kirkpatrick et al. 1991). Effects of infection with Haemoproteus can also have indirect effects on host reproduction. Female Eurasian Kestrels (Falco tinnunculus) with Haemoproteus-infected mates laid smaller and later clutches than did females with unparasitized males (Korpim¨aki et al. 1995). Among American Kestrels (Falco sparverius) infected with Haemoproteus, pairs with lower intensity infections fledged more young than birds with higher intensities (Apanius 1991). There is growing evidence for a trade-off between reproductive effort and resistance to parasites that is thought to arise when limited resources must be partitioned between reproductive effort and disease resistance (Chapter 1). Parasite intensity (as measured by numbers of circulating gametocytes) increases with the degree of effort expended in reproduction (Norris et al. 1994; Ots and Horak 1996; Allander 1997; Siikam¨aki et al. 1997; Nordling et al. 1998) and may decrease when food resources are abundant (Wiehn and Korpimaki 1998). From other studies of the subclinical impacts of Haemoproteus infections on wild birds, results have often been conflicting and dependent on the particular host–parasite association under investigation, whether or not hosts had concurrent infections with other hematozoan parasites, whether stage of infection was acute or chronic, and age of the hosts. A number of studies have been unable to establish a relationship between infection with Haemoproteus and survivorship, mating success, reproductive success, host condition, or clinical chemistry (Bennett et al. 1988; Weatherhead and Bennett 1992; Davidar and Morton 1993; Powers et al. 1994; Korpim¨aki et al. 1995; Dale et al. 1996; H˜orak et al. 1998; Dawson and Bortolotti 2000; Schrader et al. 2003), yet others find subtle effects that are either difficult to detect or are equivocal (Dawson and Bortolotti 2000). For example, no association was detected between infection with Haemoproteus tinnunculi and return rates of American Kestrels when data from both sexes were combined, but there was a significant negative association between return rates and intensity of infection in females (Dawson and Bortolotti 2000). This suggests that acute or recrudescing infections may have more impact on host survivorship than chronic, lowintensity infections, but that effects may be subtle and easily masked when data for males and females are combined. By contrast, Purple Martins (Progne subis)
infected with Haemoproteus prognei returned to breeding sites earlier than uninfected birds, and infected females had higher numbers of fledged young than uninfected birds (Davidar and Morton 1993). These authors hypothesized that recovery from acute phases of infection of Haemoproteus was evidence of immunological superiority in surviving hosts and may actually be a measure of superior fitness.
TREATMENT AND CONTROL A number of antimalarial compounds are effective for reducing intensity of parasitemia in both wild and domestic birds with infections with Haemoproteus. These include atebrine, plasmochin, chloroquine sulfate, primaquine, and mefloquine (Coatney 1935; Evans and Otter 1998; Mutlow and Forbes 1999; Remple 2004) as well as the antitheilerial drug buparvaquone (El-Metenawy 1999). Other antimalarials may be effective including pyrimethamine, pyrimethamine–sulfadoxine combinations, and tetracyclines, but their effectiveness in birds is not wildly established (Mutlow and Forbes 1999). In captive situations, infections with Haemoproteus can be controlled by housing birds in screened, Culicoides-proof facilities and dusting birds to reduce or eliminate ectoparasitic hippoboscid flies.
MANAGEMENT IMPLICATIONS There are currently no broad-scale strategies for prevention or control of infections with Haemoproteus in wild birds. While reduction of vector populations will decrease transmission of species of Haemoproteus, this approach is currently not feasible for the many species of ceratopogonids that have larval habitats in damp soil and tree cavities (Blanton and Wirth 1979) or for ectoparasitic hippoboscid flies that occur on wild birds. It is likely that some species of Haemoproteus may become emerging disease threats in the event of global climate change as the range of hosts and vectors change, bringing previously isolated populations into contact with vectors and parasites to which they had no prior exposure. On a smaller scale, similar circ*mstances occur when avian species are transported or relocated outside of their normal range. Good examples are the recent epizootics of H. lophortyx in Northern Bobwhites that were relocated in California (Cardona et al. 2002), sporadic reports of myopathy from megalomeronts in captive psittacines (Pennycott et al. 2006), and periodic outbreaks in other captive birds and zoos where new exotic hosts are exposed to endemic vectors and parasites (Ferrell et al. 2007).
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Haemoproteus DISCLAIMER Any use of trade, product, or firm names in this publication is for descriptive purposes only and does not imply endorsem*nt by the U.S. government. ACKNOWLEDGMENTS I acknowledge financial support from the U.S. Geological Survey Wildlife and Invasive Species Programs and NSF biocomplexity grant DEB 0083944. LITERATURE CITED Acton, H. W., and R. Knowles. 1914. Studies on the Halteridium parasite of the pigeon Haemoproteus columbae, Celli & San Felice. Indian Journal of Medical Research 1:663–690. Adie, H. A. 1915. The sporogony of Haemoproteus columbae. Indian Journal of Medical Research 2:671–680. Adie, H. A. 1924. The sporogony of Haemoproteus columbae. Bulletin Societe Pathologie Exotique 1:605–613. Ahmed, F. E., and A. H. Mohammed. 1978. Haemoproteus columbae: Course of infection, relapse and immunity to reinfection in the pigeon. Zeitschrift f¨ur Parasitenkunde 57:229–236. Allander, K. 1997. Reproductive investment and parasite susceptibility in the Great tit. Functional Ecology 11:358–364. Allander, K., and G. F. Bennett. 1995. Prevalence and intensity of haematozoan infection in a population of Great tit* Parus major from Gotland, Sweden. Journal of Avian Biology 25:69–74. Apanius, V. 1991. Blood parasitism, immunity and reproduction in American Kestrels (Falco sparverius). In Biology and Conservation of Small Falcons, M. K. Nicholls and R. Clarke (eds). Proceedings of the 1991 Hawk and Owl Trust Conference. The Hawk and Owl Trust, London, pp. 117–124. Appleby, B. M., M. A. Anwar, and S. J. Petty. 1999. Short-term and long-term effects of food supply on parasite burdens in Tawny Owls, Strix aluco. Functional Ecology 13:315–321. Applegate, J. E., and R. L. Beaudoin. 1970. Mechanism of spring relapse in avian malaria: Effect of gonadotropin and corticosterone. Journal of Wildlife Diseases 6:443–447. ¨ Arag˜ao, H. B. 1908a. Uber den entwicklungsgang und die u¨ bertragung von Haemoproteus columbae. Archiv f¨ur Prostistenkunde 12:154–167. Arag˜ao, H. B. 1908b. Sobre o cyclo evloutivo e a transmiss˜ao do Haemoproteus columbae. Revista do Instituto de Medicina Tropical de S˜ao Paulo. 11:416–419.
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antibody ELISA. Journal of Parasitology 80:713– 718. Greiner, E. C. 1971. The Comparative Life Histories of Haemoproteus sacharovi and Haemoproteus maccallumi in the Mourning Dove (Zenaida macroura). Ph.D. Dissertation, University of Nebraska. Greiner, E. C., G. F. Bennett, E. M. White, and R. F. Coombs. 1975. Distribution of the avian hematozoa of North American. Canadian Journal of Zoology 53:162–1787. Hartley, W. J., G. L. Reddacliff, D. Finnie, and E. P. Turner. 1981. Suspected lethal Leucocytozoon infection in the pied currawong (Strepera graculina). In Wildlife Diseases of the Pacific Basin and Other Countries. Proceedings of the 4th International Conference of the Wildlife Disease Association, Sydney, Australia. Ames, Wildlife Disease Association, pp. 95–97. ¨ Ostman, ¨ Hasselquist, D., O. J. Waldenstr¨om, and S. Bensch. 2007. Temporal patterns of occurrence and transmission of the blood parasite Haemoproteus payevskyi in the great reed warbler Acrocephalus arundinaceus. Journal of Ornithology 148:401–409. Hellgren, O., J. Waldenstr¨om, and S. Bensch. 2004. A new PCR assay for simultaneous studies of Leucocytozoon, Plasmodium, and Haemoproteus from avian blood. Journal of Parasitology 90:797–802. Hellgren, O., A. Krizanauskiene, G. Valki¯unas, and S. Bensch. 2007a. Diversity and phylogeny of mitochondrial cytochrome b lineages from six morphospecies of avian Haemoproteus (Haemosporida, Haemoproteidae). Journal of Parasitology 93:899–896. Hellgren, O., J. Waldenstr¨om, J. Per´ez-Tris, E. Szoll Osi, D. Hasselquist, A. Krizanauskiene, U. Ottosson and S. Bensch. 2007b. Detecting shifts of transmission areas in avian blood parasites—a phylogenetic approach. Molecular Ecology 16:1281–1290. Hewitt, R. J. 1940. Bird Malaria. The American Journal of Hygiene Monographic Series, No. 15. The Johns Hopkins Press, Baltimore, MD. H˜orak, P., I. Ots, and A. Murum¨agi. 1998. Haematological health state indices of reproducing Great tit*: A response to brood size manipulation. Functional Ecology 12:750–756. H˜orak, P., I. Ots, H. Vellau, C. Spottiswoode, and A. P. Møller. 2001. Carotenoid-based plumage coloration reflects hemoparasite infection and local survival in breeding great tit*. Oecologia, 126:166–173. Huff, C. G. 1932. Studies on Haemoproteus of mourning doves. American Journal of Hygiene 16:618–623. Khan, R. A., and A. M. Fallis. 1969. Endogenous stages of Parahaemoproteus fringillae (Labb´e, 1894) and Leucocytozoon fringillinarum Woolco*ck 1910
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Merino, S., J. Moreno, J. J. Sanz, and E. Arriero. 2000. Are avian blood parasites pathogenic in the wild? A medication experiment in blue tit* (Parus caeruleus). Proceedings of the Royal Society of London, Series B 267:2507–2510. Miltgen, F., I. Landau, N. Ratanaworabhan, and S. Yenbutra. 1981. Parahaemoproteus desseri n. sp.; gam´etogonie et schizognie chz l’hˆote naturel: Psittacula roseate de Thailande, et sporogonie exp´erimentale chez Culicoides nubeculosus. Annales de Parasitologie Humaine et Compar´ee 56:123–130. Mohammed, A. H. H. 1965. Studies on the schizogony of Haemoproteus columbae Kruse 1890. Proceedings of the Egyptian Academy of Sciences 19:37–46. Mullens, B. A., C. J. Cardona, L. McClellan, C. E. Szijj, and J. P. Owen. 2006. Culicoides bottimeri as a vector of Haemoproteus lophortyx to quail in California, USA. Veterinary Parasitology 140:35–43. Mutlow, A., and N. Forbes. 1999. Haemoproteus in Raptors: Pathogenicity, Treatment and Control. Landsdown Veterinary Surgeons, Wallbridge, England. Navarro, C., F. de Lope, A. Marzal, and A. P. Møller. 2004. Predation risk, host immune response, and parasitism. Behavioral Ecology 15:629–635. Nordling, D., M. Andersson, S. Zohari, and L. Gustafsson. 1998. Reproductive effort reduces specific immune response and parasite resistance. Proceedings of the Royal Society of London, Series B 265:1291–1298. Norris, K., M. Anwar, and A. F. Read. 1994. Reproductive effort influences the prevalence of haematozoan parasites in Great tit*. Journal of Animal Ecology 63:601–610. Opitz, H. M., H. J. Jakob, E. Wiensenhuetter, and V. Vasandra Devi. 1982. A myopathy associated with protozoan schizonts in chickens in commercial farms in peninsular Malaysia. Avian Pathology 11:527–534. O’Roke, E. C. 1930. The morphology, transmission, and life history of Haemoproteus lophortyx O’Roke, a blood parasite of the California valley quail. University of California Publications in Zoology 36:1–50. Ots, I., and P. H˜orak. 1996. Great tit* Parus major trade health for reproduction. Proceedings of the Royal Society London, Series B 263:1443–1447. Ots, I., and P. H˜orak. 1998. Health impact of blood parasites on breeding great tit*. Oecologia 116:441–448. Padilla, L. R., D. Santiago-Alarcon, J. Merkel, R. E. Miller, and P. G. Parker. 2004. Survey for Haemoproteus spp. Trichom*onas gallinae, Chlamydophila psittaci, and Salmonella spp. in Galapagos Islands Columbiformes. Journal of Zoo and Wildlife Medicine 35:60–64.
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hemoproteids (Haemosporida: Haemoproteidae) of Sylviid birds, with a description of Haemoproteus parabelopolskyi sp. nov. Journal of Parasitology 93:680–687. Waldenstr¨om, J., S. Bensch, S. Kiboi, D. Hasselquist, and U. Ottosson. 2002. Cross-species infection of blood parasites between resident and migratory songbirds in Africa. Molecular Ecology 11:1545–1554. Walker, D., and P. C. C. Garnham. 1972. Aberrant Leucocytozoon infection in parakeets. Veterinary Record 91:70–72. Weatherhead, P. J., and G. F. Bennett. 1991. Ecology of red-winged blackbird parasitism by haematozoa. Canadian Journal of Zoology 69:2352–2359. Weatherhead, P. J., and G. F. Bennett. 1992. Ecology of parasitism of Brown-headed Cowbirds by haematozoa. Canadian Journal of Zoology 70:1–7. White, E. M., E. C. Greiner, G. F. Bennett, and C. M. Herman. 1978. Distribution of the hematozoa of Neotropical birds. Revista de Biologica Tropical 26:43–102. Wiehn, J., and E. Korpim¨aki. 1998. Resource levels, reproduction and resistance to haemoatozoan infections. Proceedings of the Royal Society London, Series B 265:1197–1201. Wood, M. J., C. L. Cosgrove, T. A. Wilkin, S. C. L. Knowles, K. P. Day, and B. C. Sheldon. 2007. Within-population variation in prevalence and lineage distribution of avian malaria in blue tit*, Cyanistes caeruleus. Molecular Ecology 16:3263–3273. Work, T. M., and R. A. Raymeyer. 1996. Haemoproteus iwa n. sp. in Great Frigatebirds (Fregata minor [gmelin]) from Hawaii: Parasite morphology and prevalence. Journal of Parasitology 82:489–491. Yezerinac, S., and P. J. Weatherhead. 1995. Plumage coloration, differential attraction of vectors and haematozoa infections in birds. Journal of Animal Ecology 64:528–537.
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3 Avian Malaria Carter T. Atkinson first recognized as common intraerythrocytic parasites of wild birds (Danilewsky 1889). The early years of this field have been reviewed in detail by Hewitt (1940), Garnham (1966), and Valki¯unas (2005), and it is clear that most of the major milestones in the field of human malariology were associated in one way or another with avian parasites. Highlights include the first descriptions of the characteristic pathological lesions of malaria in birds by Danilewsky (1889), the discovery of the mosquito transmission of P. relictum by Sir Ronald Ross (1898), discovery of the exoerythrocytic merogony of Plasmodium elongatum in reticuloendothelial cells in bone marrow and other organs in birds (Raffaele 1934), and the development of the theory of premunition or a resistance to reinfection that is conferred by a chronic malarial infection in avian hosts (Sergent and Sergent 1956). It was recognized relatively early that both wild and captive birds experience significant disease following infection with avian malaria, with reports as early as 1905 of die-offs from infection with Plasmodium in Gray Partridges (Perdix perdix) that were imported from Hungary and released in France (Garnham 1966). Despite this lengthy history, the number of reports of large-scale epizootics from avian malaria over the past 100 years are surprisingly limited, with most associated with wild Ciconiiformes in Venezuela (Gabaldon and Ulloa 1980), captive penguins (Fix et al. 1988), and native Hawaiian forest birds (Warner 1968). There has been a recent renaissance in the use of prevalence data on hematozoan infections in birds to investigate ecological and evolutionary hypotheses about sexual selection and the physiological costs of parasitism in wild bird populations (Hamilton and Zuk 1982; Kilpatrick et al. 2006; Gilman et al. 2007). Some of this work is based on the use of molecular methods to diagnose very low intensity infections and track host specificity and geographic distribution of mitochondrial lineages of these parasites (Ricklefs et al. 2005). These new tools are leading to fundamental revisions in how we define species of Plasmodium and will play
INTRODUCTION Avian malaria is a common mosquito-transmitted disease of wild birds that is caused by protozoan parasites in the genus Plasmodium. Infections are caused by a complex of more than 40 species that differ widely in host range, geographic distribution, vectors, and pathogenicity. The avian species of Plasmodium share morphological and developmental features with closely related haemosporidian parasites in the genera Haemoproteus and Leucocytozoon (Chapters 2 and 4), but are distinguished from both by the presence of asexual reproduction (merogony) in circulating erythrocytes. While there are numerous reports of individual birds with acute, pathogenic infections with Plasmodium, reports of epizootics are rare and mostly associated with captive birds in zoological collections and abnormal host–parasite associations following introductions of parasites or mosquito vectors to remote islands. Plasmodium relictum, one of the most widely distributed species of avian malaria (Beadell et al. 2006), continues to play an important role as a limiting factor in the current distribution and abundance of native Hawaiian forest birds (Warner 1968; Woodworth et al. 2005; Foster et al. 2007). SYNONYMS Avian malaria, haemoproteosis. Many reports in the recent ecological literature lump Plasmodium with Haemoproteus and refer to both genera as avian malaria, making it difficult to identify which genus is being discussed. Clear differences in life history characteristics of these two genera justify their continued separation (Valki¯unas et al. 2005) even though they are closely related (Martinsen et al. 2008). HISTORY The avian species of Plasmodium have played a seminal role as models for human malaria since they were
35 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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an important role in assessing their impact on wildlife populations. DISTRIBUTION The species of Plasmodium that infect birds have a cosmopolitan distribution and are found in all major zoogeographic regions of the world with the exception of Antarctica, where mosquito vectors responsible for their transmission do not occur. Reports of Plasmodium from the Australian region are notably fewer than others, but it is not clear whether this is because this region has not been adequately sampled or whether it reflects a true distributional anomaly (Bennett et al. 1993; Valki¯unas 2005). Seven species of Plasmodium have a cosmopolitan distribution and broad host range, with reports from as few as 67 species of avian hosts for P. elongatum to as many as 419 different species of birds for P. relictum (Bennett et al. 1993; Valki¯unas 2005). Plasmodium relictum and P. circumflexum have the broadest geographic distribution and are reported from the Nearctic, Palearctic, Oriental, Ethiopian, Neotropical, and Australian regions. Plasmodium vaughani, P. cathemerium, P. nucleophilum, P. rouxi, and P. elongatum have been reported from all regions with the exception of the Australian region (Bennett et al. 1993). HOST RANGE Infections with Plasmodium have been reported in birds from all avian orders with the exception of the Struthioniformes (ostriches), the Coliiformes (mousebirds), and the Trogoniformes (trogons and quetzals), but only about half of all avian species have been examined for these parasites. The greatest diversity of species of Plasmodium is recorded from the Galliformes, Columbiformes, and Passeriformes (Valki¯unas 2005). Important resources for locating host records and early literature on Plasmodium infections in wild birds have been prepared by the International Reference Centre for Avian Hematozoa (Herman et al. 1976; Bennett et al. 1981, 1982; Bishop and Bennett 1992). Plasmodium relictum has one of the widest host ranges of the avian plasmodia, occurring naturally in 70 different avian families. The relatively broad host range of most species of Plasmodium from birds is considered to be characteristic of the avian species of this genus, but exceptions are common. Based on identifications made by traditional morphological methods, some species appear to have very restricted host distributions in wild populations. For example, Plasmodium hermani and Plasmodium kempi have been described from domestic and Wild Turkeys (Meleagris
gallopavo) in North America, yet are different enough in morphological features to be described as separate species. Plasmodium hermani has also been found in Northern Bobwhite (Colinus virginianus) from the same habitats as Wild Turkeys in Florida, USA (Forrester et al. 1987), but prevalence in other species of wild birds from the same habitats is not known. While P. kempi is capable of infecting species of Galliformes and Anseriformes in the laboratory, Wild Turkeys are the only known natural host of this parasite (Christensen et al. 1983). Recent application of molecular methods to screen avian hosts has revealed a far greater complexity of genetic lineages of Plasmodium and the closely related genus Haemoproteus that are currently difficult to relate to more traditional morphological species (Bensch et al. 2004). Multiple lineages can occur in the same host individual, and their occurrence in species from a wide range of avian orders, families, and species is much broader than previously recognized (Fallon et al. 2005; Ricklefs et al. 2005; Szymanski and Lovette 2005). ETIOLOGY Members of this genus are classified as members of the phylum Apicomplexa, class Aconoidasida, order Haemospororida, family Plasmodiidae and are defined primarily by their intraerthrocytic development and asexual reproduction (merogony, also called schizogony) in the circulating blood cells (Peirce 2000). All members of this genus produce prominent goldenbrown or black pigment granules from digestion of host hemoglobin. The species of Plasmodium that infect birds are divided into five subgenera based on morphology of circulating gametocytes and meronts and on preference for mature or immature erythrocytes (Table 3.1; Figure 3.1; Valki¯unas 2005). Peirce and Bennett (1996) recognize a sixth subgenus among the avian parasites, Plasmodioides, that was erected for a single species, Fallisia neotropicalis, from pigeons and Ciconiiformes in Venezuela (Gabaldon et al. 1985). This unusual avian parasite lacks pigment granules in all stages of development and develops exclusively in circulating leukocytes and thrombocytes. Peirce and Bennett (1996) argue that similarities in life history characteristics justify including this parasite among the avian malarial parasites as a species of Plasmodium, but most workers now place this subgenus in the family Garniidae (genus Fallisia) with reptilian blood parasites that also undergo merogony in circulating leukocytes (Valki¯unas 2005). Species of Plasmodium are further distinguished by host range, vectors, and developmental characteristics of exoerythrocytic tissue stages. More than 40
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Table 3.1. Subgenera and species of avian Plasmodium and characteristics of erythrocytic stages of development. Subgenus
Characteristics
Species
Haemamoeba
Gametocytes round and exceed size of host cell nucleus Mature parasites displace host cell nucleus Meronts present in mature erythrocytes
Giovannolaia
Gametocytes elongate Mature parasites do not displace host cell nucleus Meronts present in mature erythrocytes Meronts larger than erythrocyte nucleus, with plentiful cytoplasm
Novyella
Gametocytes elongate Mature parasites do not displace host cell nucleus Meronts present in mature erythrocytes Meronts smaller than erythrocyte nucleus, without noticeable cytoplasm
Bennettinia
Gametocytes, round or oval, do not exceed size of host cell nucleus and stick to host nucleus Meronts present in mature erythrocytes Meronts round with scant cytoplasm and stick to host nucleus Gametocytes elongate Mature parasites do not displace host cell nucleus Meronts variable in form and size Meronts present in circulating erythrocyte precursors
Huffia
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Plasmodium relictum Plasmodium subpraecox Plasmodium cathemerium Plasmodium gallinaceum Plasmodium matutinum Plasmodium lutzi Plasmodium giovannolai Plasmodium griffithsi Plasmodium tejerai Plasmodium coturnixi Plasmodium parvulum Plasmodium fallax Plasmodium circumflexum Plasmodium polare Plasmodium lophurae Plasmodium durae Plasmodium pedioecetae Plasmodium pinottii Plasmodium formosanum Plasmodium gundersi Plasmodium anasum Plasmodium garnhami Plasmodium hegneri Plasmodium octamerium Plasmodium gabaldoni Plasmodium leanucleus Plasmodium vaughani Plasmodium columbae Plasmodium rouxi Plasmodium hexamerium Plasmodium nucleophilum Plasmodium dissanaikei Plasmodium paranucleophilum Plasmodium bertii Plasmodium kempi Plasmodium forresteri Plasmodium ashfordi Plasmodium juxtanucleare
Plasmodium elongatum Plasmodium huffi Plasmodium hermani
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(a)
(b)
(c)
(d)
(e)
(f)
(g)
(h)
Figure 3.1. Erythrocytic stages of Plasmodium circumflexum (subgenus Novyella) (a–d) and Plasmodium relictum (subgenus Haemamoeba) (e–h). Note elongated shape of meronts (b, c) and gametocyte that encircles the host erythrocyte nucleus (d). The latter is characteristic of species of Plasmodium in the subgenus Novyella. Note shift in host erythrocyte nuclei (e–h) and round shape of gametocyte (h) that is characteristic of species of Plasmodium in the subgenus Haemamoeba. (a) Trophozoite. Note cluster of pigment granules (arrow). (b) Mature meront. Individual merozoites (arrows) are evident. (c) Mature meront. Note individual merozoites (arrows). (d) Gametocyte. Gametocyte surrounds the host erythrocyte nucleus, filling the erythrocyte cytoplasm. Pigment granules (arrows) are scattered through the parasite cytoplasm. (e) A pair of trophozoites (arrows). (f) Mature meront. Developing merozoites surround a central mass of pigment. (g) Mature meront. Developing merozoites surround a central mass of pigment (arrow). (h) Gametocyte. Note round shape, displaced host erythrocyte nucleus, and scattered pigment granules (arrows).
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Avian Malaria species are currently recognized, but this number is in a continual state of flux as existing species are synonomized as more information is learned about their biological characteristics and new species are described (Table 3.1). The recent application of molecular methods to the taxonomy of this group has identified a bewildering array of lineages that are currently defined by sequence of mitochondrial genes (Bensch et al. 2004). In most cases, we know nothing about their erythrocytic morphology, natural vectors, or other life history characteristics and are only just beginning to link this information to the more traditional morphological and biological definition of individual species (Valki¯unas et al. 2007). Recent efforts to combine molecular data with life history information from members of five subgenera (Haemamaoba, Huffia, Bennettinia, Novyella, and Giovannolaia) indicate that the very distinctive characteristics of the subgenera Haemamoeba (large, round gametocytes and prominent host nucleus displacement), Huffia (predilection for immature erythrocytes), and Bennettinia (unusual morphology of the erythrocytic and sporogonic stages) are consistent with monophyletic origins for each of these subgenera. By contrast, Novyella and Giovannolaia form a clade composed of representatives from both subgenera, indicating that the less distinctive morphology of these parasites appears to be more plastic over evolutionary time (Martinsen et al. 2008). These findings suggest that some of the key morphological features used by parasitologists to distinguish these subgenera may not reflect true phylogenetic relationships (Martinsen et al. 2008). It is clear that our understanding of the taxonomy and phylogenetics of these parasites is rapidly evolving, and their classification will likely undergo further revision in future years. EPIZOOTIOLOGY Much of what we know about the detailed life cycle of species of Plasmodium from birds is based on a series of classic experiments by Clay Huff and coworkers with Plasmodium gallinaceum. In these studies, chickens and other birds were exposed to infective mosquito bites and examined at sequential time intervals to determine the location and morphology of the parasites (Huff and Coulston 1944; Huff 1951). These studies have provided us with specific details about how some of these parasites develop, but variations in the life cycle have been documented among other species of Plasmodium and further studies are needed. The life cycle of P. gallinaceum begins when infective sporozoites are inoculated by a mosquito vector into a susceptible host (Huff and Coulston 1944). Sporozoites invade macrophages and fibroblasts near
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the site of the mosquito bite and undergo an initial generation of asexual reproduction (merogony) as cryptozoites. These are relatively small in diameter and mature in approximately 36–48 h to release ovoid merozoites that invade cells of the lymphoid– macrophage system in brain, spleen, kidney, lung, and liver tissue to begin a second generation of merogony as metacryptozoites. Metacryptozoites mature and release merozoites that are capable of invading circulating erythrocytes and capillary endothelial cells of the major organs. The first two generations of merogony are referred to as the preerythrocytic stages of infection. Merozoites that continue with a third generation of merogony in stationary tissues of the host are called phanerozoites. Once they invade capillary endothelial cells and begin to reproduce by asexual merogony, they are referred to as exoerythrocytic meronts. Merozoites released from exoerythrocytic meronts can either invade circulating erythrocytes or reinvade endothelial cells to continue additional generations of merogony in stationary tissues. The exoerythrocytic meronts that occur in capillary endothelial cells are oval, elongate, or branching and similar in morphology to thin-walled meronts of Haemoproteus (Chapter 2). They are significantly larger than the preerythrocytic meronts and may contain hundreds of nuclei (Garnham 1966). Merozoites that invade the circulating erythrocytes undergo merogony and develop within 24–48 h into either mature meronts containing 8–32 ovoid merozoites or gametocytes that are infective to mosquito vectors (Table 3.1). Depending on the species of Plasmodium, meronts may be either round or elongate and produce numbers of merozoites that may be characteristic for particular species. Merozoites typically bud from a central residual mass and destroy their host erythrocyte when they are released. By contrast, gametocytes are elongate or round and have a single nucleus. The male gametocytes (microgametocytes) typically stain pink with Giemsa stain, while female gametocytes (macrogametocytes) stain pale blue. During growth in the erythrocyte, the parasites ingest host erythrocyte cytoplasm through a specialized structure known as a cytostome and digest host hemoglobin within one or more food vacuoles scattered throughout the cytoplasm of the parasite. Malarial pigment or hematozoin is produced as a by-product of the digestion of hemoglobin and may appear as golden-brown or black granules in the parasite cytoplasm. Clear food vacuoles with one or more pigment granules may be visible by light microscopy, depending on size of the vacuoles. Merogony may continue indefinitely in the circulating erythrocytes, and evidence suggests that merozoites from some erythrocytic meronts can reinvade stationary tissues and continue development as phanerozoites (Garnham 1966).
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Unlike P. gallinaceum (subgenus Haemamoeba), P. elongatum and other species in the subgenus Huffia do not develop in capillary endothelial cells of the major organs, but instead undergo exoerythrocytic merogony in hematopoietic tissues of the host (Garnham 1966). Specific details about preerythrocytic stages of development are not known. Gametocytes of all species of avian Plasmodium remain in the circulation and do not continue development until they are ingested by an arthropod vector. Once in the midgut of a suitable mosquito vector, they leave their host cells and undergo gametogenesis to form gametes. Male gametocytes undergo a process call exflagellation to produce up to eight, flagellated microgametes. One microgamete will fertilize a macrogamete and within 24 h a motile zygote develops, which is capable of penetrating the midgut wall and beginning development as an oocyst under the basal membrane of the mosquito midgut. These initial stages of gametogenesis and fertilization exhibit little or no host specificity for mosquito vectors and can be completed in vitro. It is only during invasion of the peritrophic membrane that surrounds the blood meal and subsequent penetration of the midgut epithelium that blocks in development of particular species of malaria, in particular mosquito hosts, can occur (Michel and Kafatos 2005). Oocysts undergo a type of asexual reproduction called sporogony and eventually produce thousands of sporozoites through a process of budding from multiple residual masses or sporoblasts. Oocysts mature within approximately 7 days after reaching a diameter of approximately 40 μm, depending on ambient temperature, and rupture to release sporozoites into the hemoceol of the mosquito. Sporozoites move via the hemocoel to the salivary glands, penetrate the glandular cells, and eventually gain access to the salivary ducts. When a mosquito takes a blood meal, these pass with the saliva into a new avian host to initiate a new infection. Birds typically undergo an acute phase of infection where parasitemia increases steadily to reach a peak in numbers, called the crisis, approximately 6–12 days after parasites first appear in the blood. This is followed by a rapid decline in intensity of infection to chronic levels as the host immune system begins to bring the infection under control. Chronic infections most likely persist for the lifetime of infected birds, and both circulating parasites and persistent exoerythrocytic meronts can serve as a source for recrudescing infections (Manwell 1934; Bishop et al. 1938; Garnham 1966). More than 60 different species of culicine and anopheline mosquitoes are capable of supporting experimental development of a variety of species of Plas-
modium from avian hosts (Huff 1965), but surprisingly, few natural mosquito vectors are known (Table 3.2). For example, more than 20 species of anopheline and culicine mosquitoes in four different genera (Culex, Aedes, Culiseta, and Anopheles) are capable of transmitting P. relictum in the laboratory, but only three—Culex quinquefasciatus, Culex tarsalis, and Culex stigmatasoma—are proven natural vectors of P. relictum in California and Hawaii (Reeves et al. 1954; LaPointe et al. 2005). After the initial acute phase of infection, intensity appears to be influenced by the complex interplay of host immunity, seasonal changes in photoperiod, and hormonal changes associated with reproduction. As has been described for other hematozoan parasites (Chapters 2 and 4), an increase in intensity of infection coincides with the breeding season when populations of blood-sucking insects typically increase, and recently fledged susceptible birds are increasing in the population (Atkinson and van Riper 1991; Valki¯unas et al. 2004). Termed the “spring relapse,” the increase in numbers of parasites in the peripheral circulation can be triggered by corticosterone (Applegate and Beaudoin 1970), increases in photoperiod, and subsequent physiological changes in levels of hormones such as melatonin that regulate circadian rhythms (Valki¯unas et al. 2004). Many of the same factors that affect intensity of infection with Haemoproteus (Chapter 2) probably affect intensity of infection with Plasmodium. These include stress-mediated changes in the immune system that are associated with reproductive effort (Siikam¨aki et al. 1997), food availability (Appleby et al. 1999), concomitant infection with other parasites (Wright et al. 2005), and exposure to predators (Navarro et al. 2004). While it is clear that most transmission of avian Plasmodium takes place during the spring and summer months in temperate climates, relatively little is known about dynamics of infection in tropical parts of the world. In Hawaii, transmission of P. relictum at lower elevations can take place throughout the year (Woodworth et al. 2005), but is more seasonal at higher elevations where both temperature and rainfall have significant effects on vector populations (Ahumada et al. 2004). By contrast, transmission of P. hermani in Wild Turkeys in subtropical Florida is limited primarily to late summer and early fall when populations of the primary vector, Culex nigripalpus, reach a peak. As is the case with Haemoproteus (Chapter 2), both the spatial and seasonal patterns of transmission depend on availability of suitable mosquito vectors and susceptible avian hosts. Among migratory species, recent evidence indicates that transmission of some species of Plasmodium and other haemosporidian parasites can occur on both the breeding and the wintering grounds,
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Avian Malaria
Table 3.2. Proven and suspected natural vectors of species of Plasmodium from birds, based on demonstration of oocysts or sporozoites from wild mosquitoes or transmission by wild-captured mosquitoes. Parasite species
Locality
Mosquito vector
Plasmodium relictum
California, USA Culex stimatosoma California, USA Culex tarsalis Hawaii, USA Culex quinquefasciatus
Plasmodium gallinaceum
Sri Lanka
Mansonia crassipes
Plasmodium circumflexum
Sri Lanka
Mansonia crassipes
Plasmodium rouxi
New Brunswick, Culiseta morsitans* Canada Algeria Culex pipiens
Plasmodium juxtanucleare
Malaysia Brazil
Plasmodium hermani Plasmodium elongatum
Florida, USA Maryland, USA
Plasmodium (Novyella) sp.
Venezuela
Plasmodium (Giovannolaia) sp. Venezuela
Reference Reeves et al. (1954) Reeves et al. (1954) LaPointe et al. (2005) and Woodworth et al. (2005) Niles et al. (1965) and Garnham (1966) Niles et al. (1965) and Garnham (1966) Meyer et al. (1974)
Sergent et al. (1928) and Garnham (1966) Culex sitiens Bennett et al. (1966) Culex annulus Bennett and Warren (1966) Culex saltanensis Lourenco-de-Oliveira and de Castro (1991) Culex nigripalpus Forrester et al. (1980) Culex pipiens* Beier and Trpis (1981) Culex restuans* Beier and Trpis (1981) Aedeomyia squamipennis Gabaldon et al. (1977) and Gabaldon and Ulloa (1980) Aedeomyia squamipennis Gabaldon et al. (1977) and Gabaldon and Ulloa (1980)
Note: Numerous other species of mosquitoes are capable of supporting development of avian species of Plasmodium under laboratory conditions (Huff 1965), but few studies have isolated Plasmodium from naturally infected vectors or linked these with demonstrated transmission in the wild. * Sporozoites or oocysts of undetermined species were demonstrated in wild mosquitoes; laboratory susceptibility was confirmed. leading to increases in parasite dispersal (P´erez-Tris and Bensch 2005; Hellgren et al. 2007). Both intrinsic and extrinsic factors affect the distribution and prevalence of the closely related genus Haemoproteus (Chapter 2). Many of these same factors also determine the prevalence of Plasmodium, but this has not been examined in as much detail. Based on surveys by microscopy, prevalence of Plasmodium is four to five times lower than either Haemoproteus or Leucocytozoon, with an overall prevalence of less than 4% in a sample of over 2,000 birds from North America (Greiner et al. 1975). Prevalence of Plasmodium differed in specific physiographic regions of the continent, ranging as high as almost 10% in the southeastern US to less than 1% in the arctic barrens (Greiner et al. 1975). Very low prevalences of Plasmodium relative to Haemoproteus and Leucocytozoon may largely be a sampling artifact because very low intensity chronic infections are extremely difficult
to detect by microscopy. Prevalence of Plasmodium is much higher when more sensitive diagnostic methods are used, such as those based on the polymerase chain reaction (PCR). For example, prevalence of Plasmodium in forest birds from American Samoa is 1% by microscopy, but approximately 60% by PCR amplification of parasite ribosomal genes (Jarvi et al. 2003; Atkinson et al. 2006). Given their higher sensitivity, molecular methods may be valuable for investigating the effects of host behavior and ecology on prevalence of infection. In a large study of host and parasite community relationships in southern Missouri, USA, prevalence was weakly correlated with host body mass, but not with foraging stratum, nest height, nest type, plumage brightness, sexual dichromatism, age, or sex (Ricklefs et al. 2005). Significant relationships may have been obscured, however, by analysis of multiple parasite lineages that may differ in specific life history
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characteristics. In more intensive studies of individual species of Plasmodium in defined host populations, prevalence may differ by both age and sex. For example, prevalence of infection with P. circumflexum and P. cathemerium is significantly higher in adults rather than juvenile Red-winged Blackbirds (Agelaius phoeniceus; Herman 1938), and differences in prevalence of Plasmodium by sex have been reported in other studies of this host species (Weatherhead and Bennett 1991). The potential confounding effects of simultaneous infection with other haemosporidian parasites may also influence prevalence of Plasmodium by maintaining infections at higher frequencies than might be expected. For example, specific Mhc alleles that seem to be associated with susceptibility to Plasmodium may be maintained in a population of House Sparrows (Passer domesticus) because they confer resistance to a coinfecting strain of Haemoproteus (Loiseau et al. 2008). CLINICAL SIGNS Infections with P. relictum (canaries, Hawaiian honeycreepers, penguins), P. gallinaceum (domestic chickens), P. juxtanucleare (domestic chickens), P. elongatum (penguins), and P. durae (domestic turkeys) can be extremely pathogenic during acute phases of infection in their respective hosts (Garnham 1966; Stoskopf and Beier 1979; Huchzermeyer 1993a; Yorinks and Atkinson 2000; Williams 2005). Infected birds are typically anemic, lethargic, anorexic, and have ruffled feathers. Hematocrits may fall by more than 50% (Figure 3.2). Domestic chickens infected with P. gallinaceum and P. juxtanucleare have been described as lethargic, having pale combs, green droppings, diarrhea, and partial or total paralysis (Garnham 1966). Young turkeys with infections of P. durae exhibit few clinical signs until immediately before death, when severe convulsions may occur (Garnham 1966). Adult turkeys typically become lethargic, anorexic, and often develop right pulmonary hypertension as a consequence of hypoxic pulmonary arterial hypertension (Huchzermeyer 1988). Adult birds may also develop edematous legs and gangrene of the wattles. Cerebral capillaries may be blocked by developing exoerythrocytic meronts, and infected birds may exhibit neurological signs and paralysis before death (Garnham 1966). During the crisis, when peripheral parasitemias reach their peak, chickens infected with P. gallinaceum have reduced plasma albumin and α2 -globulin as well as significant increases in γ1 - and γ2 -globulin (Williams 2005). These changes coincide with significant increases in plasma total protein and aspar-
Figure 3.2. Hematocrit for a Wild Apapane (Himatione sanguinea) (left) with an acute natural infection with Plasmodium relictum. The hematocrit from an uninfected control canary (right) illustrates the severity of the anemia. Birds with acute infections of this intensity are rarely captured with mist nets in the wild.
tate aminotransferase, glutamate dehydrogenase, and γ-glutamyltransferase and a decrease in creatinine that likely reflect tissue damage caused by developing both erythrocytic and exoerythrocytic parasites (Williams 2005). Increases in white blood cell counts, relative and absolute lymphocytosis, and total plasma solids have been documented in Hawaiian Crows (Corvus
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Avian Malaria hawaiiensis) and penguins with acute infections with P. relictum (Graczyk et al. 1994c; Massey et al. 1996). Hematological changes are much less evident in birds with chronic infections (Ricklefs and Sheldon 2007). PATHOLOGY AND PATHOGENESIS Avian malaria is primarily a disease of the blood and reticuloendothelial system, and the progress of the disease and clinical signs closely parallel increases in the number of parasites in the peripheral circulation (van Riper et al. 1994). In detailed studies of P. gallinaceum in experimentally infected chickens, clinical signs first become evident from 5 to 7 days after inoculation of infected blood (Williams 2005). These correspond to rapid increases in peripheral parasitemia and declines in hematocrit (Figure 3.3). Hemolysis of both infected and uninfected erythrocytes and catabolism of hemoglobin leads to production of excess biliverdin, which is excreted in the feces (Williams 1985). Infected birds begin to excrete green feces approximately
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4 days after infection. During phase I, lasting only a few hours, feces are normal in form with green pigment confined to the fecal portion of the dropping. Thin, mucoid, brilliant green diarrhea develops by day 5 (phase II), which persists about 2 days among birds that survive infection. During phase III, birds are recovering from infection, and green coloration of the droppings is intermediate in intensity between that observed during phase I and phase II. Droppings lose all green color by the time that parasitemia becomes undetectable (Williams 2005). It is not clear whether acute infections with Plasmodium cause the febrile paroxysms in birds that are so characteristic of human malarial infections. Increases in cloacal temperature have been measured during acute phases of infection with P. gallinaceum in chickens (Williams 2005). As is the case with human infections, the febrile period was relatively short-lived and closely paralleled increases in peripheral parasitemia. Following the crisis, cloacal temperatures fell and then remained below normal for several days (Figure 3.3).
Figure 3.3. Relative timing of clinical signs of Plasmodium gallinaceum in domestic chickens following a blood-induced infection. Lines represent deviations from baseline conditions in healthy birds. FCR, food conversion ratio. Reproduced from Williams (2005), with permission of Taylor & Francis Ltd. (http://www.informaworld.com).
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Figure 3.4. Livers and spleens from an uninfected control canary (right) and a canary with an experimental acute infection with Plasmodium relictum (left). Infected liver (bottom left) is enlarged, has rounded borders, is discolored from deposition of malarial pigment in tissue macrophages, and has multifocal areas of necrosis. Infected spleen (top left) is similarly enlarged and discolored from deposition of malarial pigment in tissue macrophages. Tissue has been fixed in 10% buffered formalin.
By contrast, canaries infected with P. relictum have significant declines in core body temperature during acute phases of infection and appear to lose the ability to thermoregulate (Hayworth et al. 1987). The hallmark gross lesions produced by acute infections with Plasmodium include thin, watery blood, and enlargement and discoloration of the liver and spleen by deposition of malarial pigment in tissue macrophages (Figure 3.4). Enlargement of these organs is due to hypercellularity and increased phagocytic activity of macrophages rather than edema (Al-Dabagh 1966). Development of gross lesions closely corresponds to a steady increase in peripheral parasitemia, intravascular hemolysis of infected erythrocytes as meronts mature, phagocytosis of parasitized erythrocytes, and increased fragility of unparasitized erythrocytes (Al-Dabagh 1966; Seed and Kreier 1972; van Riper et al. 1994; Williams 2005). Regenerative, hemolytic anemia is associated with a drop in erythrocyte counts, replacement with immature erythrocytes, and drops in hemoglobin concentration that peak during the crisis (Figure 3.5). Anoxia and intravascular agglutinations of erythrocytes (“sludging”
of blood) may lead to damage of endothelial cells lining the capillaries (Al-Dabagh 1966). Deposition of malarial pigment in macrophages of various organs, particularly liver and spleen, as infected cells are removed from the circulation can be extensive. In intense fatal infections, thrombi or emboli can form in some organs, particularly the spleen. Secondary shock may also occur during the terminal stages of some acute infections, resulting from destruction of large numbers of infected and uninfected erythrocytes. Capillaries and venules may be dilated and exhibit increased permeability, edema, and stasis of blood flow. Hemorrhage may be evident within the capillaries. Lowered blood pressure, lowered blood volume, disturbed fluid balance, increased coagulation times, and increased levels of potassium may also be evident in severe infections (Al-Dabagh 1966). Infections with P. cathemerium produce inflammatory myopathy in skeletal muscle of experimentally infected canaries. This is characterized by degeneration of capillaries and muscle fibers and presence of mononuclear cell infiltrates. Carmona et al. (1996) suggest that this may be related to obstruction of capillaries
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Figure 3.5. Blood smear from an Iiwi (Vestiaria coccinea) with an experimental infection with Plasmodium relictum. The normal cellular makeup of the blood is profoundly altered, with mature erythrocytes being replaced by erythroblasts (EB) and early polychromatic erythrocytes (PE). P, parasitized erythrocytes (Atkinson et al. 1995). by infected erythrocytes. Anemia may also lead to circulatory deficiency that is compensated in part by increased cardiac output and dilation and hypertrophy of heart muscle (Al-Dabagh 1966). While little or no host response is evident around preerythrocytic meronts of Plasmodium, the exoerythrocytic meronts of some species, for example, P. gallinaceum and P. durae, may partially or completely block capillaries, leading to leakage of plasma proteins, edema, and hemorrhage. These lesions may occur in the heart, lungs, renal glomeruli, and brain. When they occur in the brain, neurologic symptoms may appear and death can be sudden. There is a clear association between the severity of disease and dose. This has been demonstrated experimentally with both blood-induced infections (Permin and Juhl 2002) and sporozoite-induced infections (Atkinson et al. 1995). Birds exposed to higher numbers of infective sporozoites have higher parasitemias, more severe gross and microscopic lesions, and higher mortality (Atkinson et al. 1995, 2000).
DIAGNOSIS The gold standard for diagnosis of Plasmodium is a Giemsa-stained thin blood smear where it is possible to demonstrate the presence of erythrocytic meronts and gametocytes with prominent golden-brown or black pigment granules. Individual species are traditionally defined by size and shape of intraerythrocytic gametocytes and meronts (Table 3.1; Figure 3.1), number of merozoites produced by mature meronts, changes in morphology of the host erythrocyte, and other biological characteristics such as host range, susceptibility to species of mosquitoes, morphology, and location of exoerythrocytic meronts (Garnham 1966; Valki¯unas 2005). Since most identifications are made from blood smears, life history characteristics may be unknown, and it becomes essential to be able to find enough mature meronts and gametocytes on a smear to be able to make an accurate assessment of parasite morphology. Detailed keys and species descriptions have been recently revised by Valki¯unas (2005), and his monograph is currently the most
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up-to-date resource for identifying species of avian Plasmodium. Most infections of Plasmodium in wild birds are chronic, however, and intensity may be extremely low. In these cases, it may be impossible to identify parasites below level of subgenus. When erythrocytic meronts are not present, it may become difficult to distinguish gametocytes of Plasmodium from those of Haemoproteus, although gametocytes of Haemoproteus are often thicker and more robust than those of Plasmodium. The fact that species of Plasmodium have circulating meronts while species of Haemoproteus do not can be used to both isolate and identify an unknown species of Plasmodium if susceptible domestic or captive wild birds are available for experimental subinoculation of blood from the suspect bird. While it was common knowledge among early malariologists that Plasmodium can be passed to a new host by blood inoculation, Manwell and Herman (1935) and later Herman (1938) were the first to apply this method to diagnose infections with Plasmodium in wild birds. Blood from an infected host is passed by intravenous, intraperitoneal, or intramuscular inoculation into an uninfected host of the same species, and blood smears are prepared from the inoculated host for several weeks after injection. If the host is susceptible to the parasites, an acute phase infection will often result and meronts and gametocytes can be readily found for morphological analysis. When parasitemia is high, blood can be collected, treated with glycerin or dimethyl sulfoxide, aliquoted, and frozen in liquid nitrogen to create a frozen stabilite for further experimental studies (Garnham 1966). Given the importance of morphological characters to identify species of Plasmodium from birds, their consistency and stability between hosts of different species is critical for making accurate identifications. Surprisingly, few studies have looked at this issue in detail. In one of the most widely cited examples, when P. relictum from Silver Gulls (Larus novaehollandiae) was passed by sporozoites to sparrows and canaries, merozoite, and gametocyte morphology changed significantly (Lawrence and Bearup 1961). In gulls, gametocytes were elongate and mature meronts had 10 merozoites. In sparrows, morphology was more typical of P. relictum and gametocytes were round or oval, and mature meronts had on average 14 merozoites. Other reports have documented changes in morphology when parasites are inoculated into atypical hosts (Garnham 1966) or when parasitemias are extremely high in immature erythrocytes (Laird and van Riper 1981). By contrast, other reports have documented relatively constant morphology in hosts from multiple avian species and orders (van Riper et al. 1986; Iezhova et al. 2005; Valki¯unas et al. 2007; C. T. Atkinson, unpublished observations). This issue clearly needs further study, and
the relatively recent development of molecular methods to diagnose avian malaria with PCR primers to ribosomal and mitochondrial genes may help to resolve this problem. Despite their higher sensitivity, PCR methods may still miss infections that have extremely low parasitemias (Jarvi et al. 2002), although the recent application of real-time methods to malarial diagnostics may eventually solve these problems (Boonma et al. 2007). Several recent sets of primers designed to amplify portions of the parasite mitochondrial genome can distinguish Haemoproteus and Plasmodium from Leucocytozoon (Hellgren et al. 2004) or all three genera from each other following restriction digests of PCR products (Beadell and Fleischer 2005). However, sequencing of PCR products is necessary for identifying individual parasite lineages and determining phylogenetic relationships. Since so few isolates of avian Plasmodium of known identity have been sequenced and typed, it is often not known how to relate unknown mitochondrial lineages to traditional morphological species. Recent rapid progress in the molecular diagnosis of avian species of Plasmodium may eventually make it possible to identify species based on mitochondrial lineage (Valki¯unas et al. 2007). Plasmodium appears to be antigenically distinct from Haemoproteus, and crude antigen extracts have been used to develop an ELISA test for P. relictum in captive and wild penguins (Graczyk et al. 1994a, b). Standard immunoblotting techniques can also be used to identify antibodies to Plasmodium in wild and experimentally infected passerines (Atkinson et al. 2001). Although neither ELISA nor immunoblotting can distinguish species of Plasmodium, the techniques are useful for making diagnoses to level of genus in birds with low-intensity infections that may be missed by microscopy or PCR. IMMUNITY Birds infected with avian species of Plasmodium develop strong antibody and cell-mediated responses to erythrocytic parasites (van Riper et al. 1994), but appear to be unable to completely clear their infections. Limited evidence based on experimental studies in canaries (P. relictum), Hawaii Amakihi (Hemignathus virens) (P. relictum), and domestic turkeys (P. hermani) indicates that birds likely remain infected for life, but at chronic levels that stimulate immunity to reinfection with hom*ologous strains of the parasite (Bishop et al. 1938; Jarvi et al. 2002; Young et al. 2004). This phenomenon, termed premunition, was recognized in the early part of the twentieth century (Hewitt 1940; Sergent and Sergent 1956). When birds with blood or sporozoite-induced infections are rechallenged, they
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Avian Malaria may have only brief, low-intensity increases in peripheral parasitemia (Hewitt 1940; Atkinson et al. 2001; Paulman and Mcallister 2005). The persistence of subclinical infections may make birds vulnerable to the recrudescence of erythrocytic parasites if host immunity is compromised by stress or infection with other pathogens and provides an indirect measure of the cost of mounting an immune response. Experimental manipulation of clutch size led to increases in prevalence of Plasmodium in female Great tit* (Parus major) that laid more eggs, supporting the idea that there is a trade-off between the energetic costs of egg production and defense against parasites (Oppliger et al. 1996). Similarly, male Great tit* that expended extra energy to provision larger broods had a higher prevalence of malarial infection (Richner et al. 1995). Exposure to other infectious diseases that compromise the immune system may also lead to recrudescing infections. When Wild Turkeys are exposed simultaneously or sequentially to turkeypox virus and P. hermani, both parasitemia and mortality are higher in 1-week-old poults infected with both agents than those exposed to either malaria or pox alone (Wright et al. 2005). These effects are less evident in older poults, suggesting that the host age may also play a role in pathogenesis of concomitant infections.
PUBLIC HEALTH CONCERNS Avian species of Plasmodium do not infect humans, and infected birds pose no health risks to humans.
DOMESTICATED ANIMAL HEALTH CONCERNS Domestic poultry are susceptible to several species of avian malaria, but their most significant effects occur outside of North America and Europe and specifically where wild reservoir hosts serve as sources of infection for domestic birds. Plasmodium gallinaceum is highly pathogenic in domestic chickens, particularly when European breeds are introduced to endemic areas in southeastern Asia, Malaysia, India, and Sri Lanka where the natural host is the Red Junglefowl (Gallus gallus; Garnham 1966). The distribution of the parasite in domestic chickens coincides with the geographic range of the natural host and has not expanded with the movement of domestic poultry to other parts of the world. Plasmodium juxtanucleare is also a significant pathogen in domestic chickens in South America, southern Africa, and southeastern Asia. Proven wild reservoirs of this species are found in India, Malaysia, South Africa, and Taiwan and include
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Red Junglefowl, Gray-winged Francolins (Francolinus africanus), and Chinese Bamboo-Partridges (Bambusicola thoracicus), but natural hosts are not known for other parts of its range (Garnham 1966; Fernando and Dissanaike 1975; Manwell et al. 1976; Earle et al. 1991). Domestic turkeys are highly susceptible to P. durae in sub-Saharan Africa. This species is a parasite of wild francolins that infects domestic turkeys when wild reservoir hosts and vectors are present (Huchzermeyer 1993b). P. durae is highly pathogenic in domestic turkeys, and mortalities can be as high as 90% in young poults. Both P. kempi and P. hermani infect Wild Turkeys in North America, but have not reported to be a problem in domestic birds. WILDLIFE POPULATION IMPACTS There is relatively little evidence that species of avian Plasmodium are causes of major epizootic die-offs in their natural hosts. In a frequently cited example, high rates of transmission of species of Plasmodium from several subgenera have been documented in Venezuela among nesting Ciconiiformes, but clear evidence of malarial mortality in dead nestlings is not provided (Gabaldon and Ulloa 1980). In a thorough review of over 5,000 papers on avian blood parasites, Bennett et al. (1993) found that only about 4% reported mortality or pathogenicity in birds, with most dealing with domestic birds or birds in zoological collections. Evidence is beginning to accumulate, however, that both direct and indirect effects of acute and chronic infections can have measurable impacts on the lifetime reproductive success of their avian hosts. In a study of singing behavior in White-crowned Sparrows (Zonotrichia leucophrys oriantha), song consistency was influenced by infection with Plasmodium and Leucocytozoon. Birds infected with Plasmodium also sang fewer songs following experimental playback of recorded songs (Gilman et al. 2007). This could have a significant impact on mate choice and reproductive success of infected males. Similarly, the behavioral effects of acute infections may lead to increased predation of infected hosts (Yorinks and Atkinson 2000; Møller and Nielsen 2007). These questions are just beginning to be explored in detail in ecological studies of wild birds, and the careful integration of both field and laboratory studies may lead to significant progress in our understanding of the more subtle costs of infection with these parasites. The most significant reports of pathogenicity among species of Plasmodium that infect birds are in captive birds, zoological collections, and on isolated islands when new host–parasite associations become established. Avian malaria is particularly pathogenic in
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captive penguins whenever they are exposed to mosquito vectors outside of their natural range (Stoskopf and Beier 1979; Fix et al. 1988). Welldocumented cases of mortality from Plasmodium have not been reported in wild penguins (Jones and Shellam 1999; Sturrock and Tompkins 2007), but the introduction and spread of new mosquito vectors and the potential effects of global climate change may begin to place wild colonies at risk in future years (Miller et al. 2001; Tompkins and Gleeson 2006). The threat that introduced avian malaria poses to endemic birds on isolated islands is substantial. The accidental introduction of P. relictum and the southern house mosquito (C. quinquefasciatus) to the Hawaiian Islands has had a devastating impact on native Hawaiian forest birds (Warner 1968; van Riper et al. 1986) and continues to play a significant role in limiting the current geographic and altitudinal distribution of remaining species (Atkinson et al. 1995; Benning et al. 2002). Of more than 70 species and subspecies of endemic forest birds present at the end of the eighteenth century, at least 23 are now extinct and 30 of the remaining species and subspecies are listed as endangered by the U.S. Fish and Wildlife Service (Jacobi and Atkinson 1995). While numerous limiting factors have contributed to these extinctions, high susceptibility to malaria is believed to be one of the most important reasons why populations of native species have collapsed at low elevations in areas where suitable habitat still exists (van Riper et al. 1986; Atkinson et al. 1995). High rates of transmission are maintained by the extremely high susceptibility of native honeycreepers (Drepanidinae) to P. relictum (Atkinson et al. 1995, 2000), presence of high rates of malaria transmission in the lowlands (Woodworth et al. 2005), and presence of disease-free refugia on the highest mountaintops that provide a continual source of nonimmune birds for initiating epizootics at lower elevations (Atkinson and LaPointe 2009). While many of the more rare native species are continuing to decline, at least one, Hawaii Amakihi (Hemignathus virens), appears to be evolving some resistance to infection, and lowland populations in some parts of Hawaii have started to rebound in recent years (Woodworth et al. 2005; Foster et al. 2007). TREATMENT AND CONTROL Chloroquine phosphate, primaquine phosphate, pyrimethamine–sulfadoxine combinations, and mefloquine are effective in treating canaries, penguins, and raptors with avian malaria (Remple 2004). The anticoccidial drugs sulfamonomethoxine, sulfachloropyrazine, and halofuginone are somewhat effective in treating P. durae in domestic turkeys and may also be effective against P. gallinaceum.
Sulfamonomethoxine suppresses parasitemia, but does not provide full protection from mortality when given after the appearance of circulating parasites. Sulfachloropyrazine reduces mortality, but has no effect on parasitemia, suggesting that it has some efficacy against exoerythrocytic schizonts. Halofuginone delays parasitemia, but suppresses it to only a minor extent (Huchzermeyer 1993a). While birds were some of the first experimental models for development of vaccines against Plasmodium, practical methods for immunizing wild birds have not been developed and this probably presents the most significant challenge to controlling infection with this approach. A variety of different experimental vaccines have been used, including use of ultraviolet light-inactivated, formalin-inactivated, and irradiated sporozoites, merozoites, and gametes, and synthetic vaccines based on parasite surface molecules (van Riper et al. 1994). Two DNA vaccines based on the circ*msporozoite protein of P. gallinaceum and P. relictum have recently been evaluated in Jackass Penguins (African Black-footed Penguins, Spheniscus demersus; Grim et al. 2004), and canaries (McCutchan et al. 2004) exposed to natural transmission of P. relictum in a zoological park. Both provided protection to natural exposure to P. relictum, but immunity was short-lived in canaries, and birds were just as susceptible as unvaccinated controls when exposed to mosquito vectors 1 year later. As has been demonstrated with human malaria, reductions of populations of mosquito vectors can reduce transmission of Plasmodium, but this method has not been widely used to control infections in wild or captive birds. Efforts to control avian malaria in Hawaiian forest birds have focused on reducing larval habitat for the introduced mosquito, C. quinquefasciatus (Reiter and LaPointe 2007; LaPointe et al. in press). The most cost-effective measures for captive or domestic birds include housing cage birds in screened, mosquito-proof buildings, or locating birds in areas that are isolated from wild reservoir hosts. MANAGEMENT IMPLICATIONS The potential risk of exposure to avian malaria should be considered when threatened or endangered species are moved outside of their normal ranges and maintained in captive propagation facilities or zoological parks where they may be introduced to new vectors and locally transmitted strains of Plasmodium. This risk is well documented for penguins, but should also be considered for species of birds from remote and isolated island systems that may have no prior exposure to these parasites. Similarly, the unintentional introduction of both parasites and mosquito vectors to new
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Valki¯unas, G., P. Zehtindiiev, O. Hellgren, M. Ilieva, and T. A. Iezhova. 2007. Linkage between mitochondrial cytochrome b lineages and morphospecies of two avian malaria parasites, with a description of Plasmodium (Novyella) ashfordi sp. nov. Parasitology Research 100:1311–1322. van Riper, C., III, S. G. van Riper, M. L. Goff, and M. Laird. 1986. The epizootiology and ecological significance of malaria in Hawaiian land birds. Ecological Monographs 56:327–344. van Riper, C., III, C. T. Atkinson, and T. M. Seed. 1994. Plasmodia of birds. In Parasitic Protozoa, Vol. 7, J. P. Kreier (ed.). Academic Press, New York, pp. 73–140. Warner, R. E. 1968. The role of introduced diseases in the extinction of the endemic Hawaiian avifauna. Condor 70:101–120. Weatherhead, P. J., and G. F. Bennett. 1991. Ecology of red-winged blackbird parasitism by haematozoa. Canadian Journal of Zoology 69:2352–2359. Williams, R. B. 1985. Biliverdin production in chickens infected with the malarial parasite Plasmodium gallinaceum. Avian Pathology 14:409–419. Williams, R. B. 2005. Avian malaria: Clinical and chemical pathology of Plasmodium gallinaceum in the domesticated fowl Gallus gallus. Avian Pathology 34:29–47. Woodworth, B. L., C. T. Atkinson, D. A. LaPointe, P. J. Hart, C. S. Spiegel, E. J. Tweed, C. Henneman, J. LeBrun, T. Denette, R. DeMots, K. L. Kozar, D. Triglia, D. Lease, A. Gregor, T. Smith, and D. Duffy. 2005. Host population persistence in the face of introduced vector-borne diseases: Hawaii amakihi and avian malaria. Proceedings of the National Academy of Sciences of the United States of America 102:1531–1536. Wright, E. J., J. K. Nayar, and D. J. Forrester. 2005. Interactive effects of turkeypox virus and Plasmodium hermani on turkey poults. Journal of Wildlife Diseases 41:141–148. Yorinks, N., and C. T. Atkinson. 2000. Effects of malaria (Plasmodium relictum) on activity budgets of experimentally-infected juvenile Apapane (Himatione sanquinea). The Auk 117:731–738. Young, M. D., J. K. Nayar, and D. J. Forrester. 2004. Epizootiology of Plasmodium hermani in Florida: Chronicity of experimental infections in domestic turkeys and Northern Bobwhites. Journal of Parasitology 90:433–434.
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4 Leucocytozoonosis Donald J. Forrester and Ellis C. Greiner infection in domestic chickens in South and Southeast Asia).
INTRODUCTION Leucocytozoonosis is a vector-borne protozoan disease of birds caused by several species of Apicomplexa in the genus Leucocytozoon. There are many species of Leucocytozoon, but only a few are known to be pathogenic to their hosts. Avian groups at risk include waterfowl, pigeons, galliforms, raptors, and ostriches (Bennett et al. 1993b; Valki¯unas 2005). Several species cause significant mortality in domestic waterfowl and poultry, and one species (Leucocytozoon simondi) causes localized epizootics in wild ducks and geese (O’Roke 1931; Herman et al. 1975). Other species of Leucocytozoon cause disease on a smaller scale, but have not been studied extensively (Valki¯unas 2005). Undoubtedly, as more details on the life cycles and other biological aspects of the many relatively unstudied species are determined, others will be found to be pathogenic. All leucocytozoids are host specific at the avian order level and in some cases at the family level (e.g., Leucocytozoon simondi) and some even at the species level (e.g., Leucocytozoon caulleryi and Leucocytozoon smithi). They are closely related to species of the genera Plasmodium and Haemoproteus with similar life cycles, but are transmitted by black flies of the family Simuliidae, except for L. caulleryi, which is vectored by biting midges of the family Ceratopogonidae (Akiba 1960; Valki¯unas 2005). There is a considerable body of literature on the various species of Leucocytozoon, only a part of which is discussed here. Several monographs and reviews have been published (Sambon 1908; Hewitt 1940; Bennett et al. 1965; Garnham 1966; Cook 1971; Fallis et al. 1974; Kuˇcera 1981a, b; Atkinson and van Riper 1991; Greiner 1991; Desser and Bennett 1993; Valki¯unas 2005).
HISTORY The first publication on Leucocytozoon was by Sakharoff (1893) and was a morphological study of leucocytozoids of crows, magpies, and rooks in the Russian state of Georgia. This was followed by a paper written by Ziemann (1898) that included a description of leucocytozoids in the Little Owl (Athene noctua). He described the species as Leukocytozoon (sic) danilewskyi and was the first to stain blood films; his paper contained color illustrations of the gametocytes of this leucocytozoid (Ziemann 1898). The genus name Leucocytozoon was first used by Berestneff (1904) who described several species from owls, rooks, and crows, but Sambon (1908) was the first to define the genus Leucocytozoon. The family Leucocytozoidae was established later by Fallis and Bennett (1961). In 1965, the genus Akiba was established for one species of leucocytozoid (Akiba caulleryi) that is transmitted by biting midges rather than black flies (Bennett et al. 1965). This genus is currently considered by most as a subgenus of the genus Leucocytozoon (Valki¯unas 2005). It is interesting that the name Leucocytozoon was chosen originally for these blood protozoans because it was believed that they occupied only leukocytes. It was later discovered, however, that the gametocytes of this parasite developed in erythrocytes as well (Cook 1954; Desser 1967). Leucocytozoids were first identified as agents of disease by Tartakovsky (1913) who worked with domestic and wild anseriforms in northeastern Russia. Tartakovsky’s work was apparently overlooked by his contemporaries in Canada (Wickware 1915) and Germany (Knuth and Magdeburg 1922) who subsequently described the disease without reference to it (Valki¯unas 2005). Later work by O’Roke (1934), Karstad (1965), Fallis and Bennett (1966), Khan and Fallis (1968), Kocan (1968), Herman et al. (1975),
SYNONYMS Hematozoan disease, haemosporidian disease, blood parasite disease, Leucocytozoon disease, and Bangkok hemorrhagic disease (refers specifically to L . caulleryi
54 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Leucocytozoonosis Desser and Ryckman (1976), M¨orner and Wahlstr¨om (1983), and others defined L. simondi as an important disease agent. Leucocytozoon marchouxi was suspected to be pathogenic to columbiforms by Oosthuizen and Markus (1968) and Peirce (1984), but this was not confirmed until the 1990s by Peirce et al. (1997). Initial evidence of the pathogenicity of Leucocytozoon toddi to raptors was reported by Olsen and Gaunt (1985) and Korpim¨aki et al. (1995), while conclusive data were published later by Raidal and Jaensch (2000). The first discovery of arthropods as vectors of leucocytozoids occurred in the early 1930s when O’Roke (1930) and Skidmore (1931) simultaneously and independently showed that simuliid black flies transmitted L. simondi to ducks and L. smithi to turkeys. One species of Leucocytozoon (Leucocytozoon caulleryi) was found subsequently by Akiba (1960) to be transmitted by biting midges of the family Ceratopogonidae. It was not until the 1940s that megalomeronts (=megaloschizonts; exoerythrocytic stages that develop in macrophages and other cells of the reticuloendothelial system) were discovered (Huff 1942; Wingstrand 1947). These were significant contributions that eventually led to an understanding of the pathogenicity of several species of leucocytozoids (Valki¯unas 2005). During the 1960s and 1970s, considerable progress was made in understanding the morphology, development, and transmission of species of Leucocytozoon. Much of this was due to the efforts of Canadian researchers who were working with L . simondi in waterfowl, including A. M. Fallis, S. S. Desser, and R. A. Khan. From the early 1900s to the present, there have been a number of publications on the taxonomy and systematics of the Leucocytozoon fauna of birds (Valki¯unas 2005). Among these were papers by Mathis and L´eger (1909–1913), de Mello (1916–1937), Coatney (1937– 1938), Herman (1938–1976), Bennett and colleagues (1965–1995), Ashford (1971–1990), Nandi (1977– 1986), Peirce (1977–present), and Valki¯unas and colleagues (1983–present). The formation of the International Reference Centre for Avian Malaria Parasites in 1967 in St. John’s, Newfoundland, Canada, was an important milestone in the development of our knowledge of blood parasites, including species of Leucocytozoon (Bennett and Laird 1973). In 1975, the center was renamed the International Reference Centre for Avian Haematozoa, and in 1995, it was moved to the Queensland Museum in Brisbane, Australia. Laird and Bennett were active in initiating the formation of the center, which contains a large collection of the literature
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on blood parasites of birds and also a vast collection of over 64,000 preparations (mostly stained blood films) from throughout the world. Type material for many species of Leucocytozoon is contained in the collection of the center (Bennett et al. 1980). Bennett and many colleagues along with visiting scientists at the center produced a large number of publications describing new species and providing redescriptions of a number of known species of Leucocytozoon. In addition, they produced several publications containing bibliographies of pertinent literature (Herman et al. 1976; Bennett et al. 1981a; Bishop and Bennett 1992), a list of species names considered valid at the time (Bennett et al. 1994), and avian host–parasite checklists (Bennett et al. 1982b; Bishop and Bennett 1992). All this resulted in a new understanding, appreciation, and awareness of Leucocytozoon and other blood protozoans, and has provided a framework for subsequent work on this genus. This foundation of knowledge made a key theoretical paper possible on the influence of blood parasites, including leucocytozoids, on the development of bright plumage, peculiarities of song, choice of mating partners, and the resultant effects on populations (Hamilton and Zuk 1982). Publication of this paper led to a renaissance in the use of avian blood parasites to test a wide range of ecological hypotheses on the impacts of parasites on sexual selection and host fitness (Møller 1997), but not without some controversy. Among the criticisms that have been made of the research are that it has been conducted by nonparasitologists who do not fully understand and appreciate the complexity of the host–parasite systems in question and that the research has been based on identifications of the blood parasites only to the generic level (using morphological and molecular techniques) rather than identifying the parasites to species (Cox 1989; McLennan and Brooks 1991; Poulin and Vickery 1993; Poulin 1995; John 1997; Valki¯unas 2005). Regardless, publication of the paper by Hamilton and Zuk (1982) has helped to open this field up to interdisciplinary studies by a wide range of parasitologists, ecologists, and avian biologists. Recently, molecular techniques using polymerase chain reaction-based and restriction enzyme-based assays were developed to allow diagnosis of leucocytozoid infections in a time-efficient manner (Hellgren et al. 2004; Beadell and Fleischer 2005). These techniques should provide reliable and inexpensive methods for detecting Leucocytozoon infections, although not yet at the species level. At the present time, it seems wise to use both molecular and traditional morphometric (microscopic) techniques to diagnose leucocytozoid species and investigate their phylogenetic relationships such as has been done by Sehgal et al. (2006a) and Martinsen et al. (2006).
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Table 4.1. Prevalence of species of Leucocytozoon in birds in various zoogeographic regions throughout the world. Zoogeographic region Holarctic Ethiopian Oriental Neotropical Australian Antarctic
Number of birds examined
Number of birds infected
Prevalence of infection (%)
102,590 11,507 45,091 54,116 —* 0
16,619 529 1,327 79 —* 0
16.2 4.6 2.9 0.1 —* 0
Source: Modified from and includes data from Greiner et al. (1975), McClure et al. (1978), White et al. (1978), Peirce (1981), Valki¯unas (1987, 1996), Bennett et al. (1992a), and Forrester et al. (2001a). * There are no appropriate data on prevalence available for the Australian region. DISTRIBUTION Leucocytozoids are distributed worldwide except in the Antarctic (Valki¯unas 1996, 2005). All the currently known species are found in the Holarctic, Ethiopian, and Oriental zoogeographic regions with a few species being also found in the Neotropical and Australian regions (Table 4.1). The highest prevalence of leucocytozoids, the highest species diversity, and the greatest number of species specific to a particular zoogeographic region occur in the Holarctic. The fauna of the Neotropical region is the poorest (White et al. 1978), especially in comparison to the Nearctic region where in certain areas leucocytozoids are the dominant hematozoan (Greiner et al. 1975). The increase in the general prevalence of leucocytozoids on a gradient from the south (e.g., Neotropical region) to the north (e.g., Nearctic region) has been attributed to increased densities of host populations in the north and to changes in the dynamics of transmission (White et al. 1978; Valki¯unas 2005). Some species of Leucocytozoon are found even above the Arctic Circle and are transmitted there (Valki¯unas et al. 1990). These observations may be somewhat skewed because birds in the Holarctic region have been studied fairly extensively, whereas the avifaunas in other regions such as the Neotropic and Australian regions have received less attention (Valki¯unas 2005). The geographic distributions of eight species of Leucocytozoon that have been reported to cause leucocytozoonosis in domestic and wild birds are presented in Table 4.2. The three species of most concern in wild populations include L . simondi in waterfowl, L. marchouxi in pigeons and doves, and L. toddi in raptors. All three are found commonly in the Holarctic region, whereas L. marchouxi and L. toddi are common there as well as in the Oriental and Ethiopian regions. Leucocytozoon toddi probably has the most widespread geographic distribution of the three species; in addition to the Holarctic, Oriental, and Ethiopian regions, it is
found in the Neotropical region, although it seems to be less common in the latter three regions. The global distribution of L. simondi is given in Figure 4.1. There are no reports from Greenland, Iceland, South America, Australia, or Antarctica, and only one unconfirmed report from Africa. There are no reports of transmission of L. simondi in Africa, South Asia, and Mexico; the records from these regions were from overwintering Holarctic anatids (Valki¯unas 2007, Personal communications to D. J. Forrester, June 29, 2007, September 4, 2007, and October 22, 2007). On a smaller scale, the distribution of leucocytozoids within different zoogeographic regions is influenced by the presence or absence of flowing streams and rivers in which the vectors develop (Desser and Bennett 1993; Valki¯unas 2005).
HOST RANGE Species of Leucocytozoon have been reported from 22 of the 28 orders and 113 of the 204 avian families of birds recognized by Clements (2000) (Table 4.3). The highest numbers of species occur in Passeriformes (8 species), Galliformes (7), and Coraciiformes (4). Only 1, 2, or 3 species have been found in birds of the other avian orders. About 45% of the species of birds in the world have been investigated for blood parasites, and species of Leucocytozoon have been found in approximately 30% of these birds (Valki¯unas 2005). No leucocytozoids have been reported from five orders of birds, that is, Tinamiformes (tinnamous), Podicipediformes (grebes), Procellariiformes (albatrosses and petrels), Phoenicopteriformes (flamingos), and Pterocliformes (sandgrouse). In some cases, the records for some orders might reflect sporadic or accidental infections in an abnormal host. One example is the report of Leucocytozoon sp. in a gaviiform (the Common Loon, Gavia immer). The loon in question was in captivity in an outdoor pen for over a month
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Leucocytozoonosis Table 4.2. Geographic distribution of eight species of Leucocytozoon that are pathogenic to wild and domestic birds. Species of Leucocytozoon Leucocytozoon marchouxi Leucocytozoon simondi* Leucocytozoon toddi Leucocytozoon smithi
Main avian hosts
Common occurrence
Pathogenic to wild birds Pigeons and doves Holarctic and Ethiopian regions Ducks, geese, and Holarctic region swans Raptors
Holarctic and Ethiopian regions Pathogenic mainly to domestic birds Turkeys Nearctic region
Leucocytozoon macleani†
Chickens and pheasants
South and Southeast Asia
Leucocytozoon struthionis Leucocytozoon schoutedeni†
Ostriches Chickens
South Africa Ethiopian region
Leucocytozoon caulleryi
Chickens
South Asia and Southeast Asia
Occasional records Oriental region and Central America Mexico and other areas outside of Holarctic region Oriental and Neotropical regions Europe and South Africa (introduced) Central and southern Palearctic region and also Ethiopian and Oriental regions None Southeast Asia and USA (possibly introduced) None
Source: Prepared with data from Valki¯unas (2005, Personal communications to D. J. Forrester, June 29, 2007, September 4, 2007, and October 22, 2007). * Leucocytozoon simondi is also pathogenic to domestic waterfowl. † This species is mildly pathogenic. (Forrester and Spalding 2003). During that time, it was not protected from arthropod vectors and was debilitated due to aspergillosis. The immunocompromised condition of the bird may have made it susceptible to infection by a leucocytozoid from other birds in the rehabilitation facility or from free-ranging birds in the immediate area. Similarly, infections with Leucocytozoon struthionis have been found only in chicks of ostriches (Struthio camelus) up to 7 weeks of age and never in adult birds. Both the low number and morphologic condition of the gametocytes were suggestive of abnormal infections (Walker 1913; Bennett et al. 1992d) that were “remarkably similar” in form and dimensions to Leucocytozoon schoutedeni, a common parasite of chickens in the area. Leucocytozoonosis occurs primarily in members of the Anatidae (ducks, geese, and swans) (Table 4.4) and Columbidae (pigeons and doves) (Table 4.5), and less commonly in members of the Accipitridae (hawks, eagles, and kites) and Falconidae (falcons and caracaras) (Table 4.6). Leucocytozoon simondi, the most important pathogenic leucocytozoid in wild birds, particularly in the Holarctic, has been reported in 46 species of waterfowl from 17 countries (Table 4.4).
ETIOLOGY Species of Leucocytozoon are parasitic protozoans and are classified within the phylum Apicomplexa (Levine, 1970), class Coccidea (Leuckart, 1879), subclass Coccidia (Leuckart, 1879), order Haemosporida (Danilewsky, 1885), and family Leucocytozoidae Fallis and Bennett, 1961. In birds, there is one genus in the family (Leucocytozoon Berestneff, 1904) that is divided into two subgenera (Leucocytozoon Berestneff, 1904 and Akiba Bennett, Garnham and Fallis, 1965). Historically, the description of leucocytozoid species has been based mainly on the morphology of gametocytes in blood cells, although the examination of exoerythrocytic stages (i.e., meronts or schizonts) has been used to some extent. There are a number of problems associated with this practice (Valki¯unas 2005): (1) intensity of infection in wild birds is low, and prolonged microscopic searches of stained blood films are necessary to find adequate numbers of infected blood cells; (2) gametocytes are often deformed when blood films are prepared if appropriate precautions are not taken; (3) there are fewer morphologic characteristics available to use for leucocytozoids compared to other blood protozoa, and often the morphology of
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Figure 4.1. Distribution of Leucocytozoon simondi throughout the world. Solid circles indicate areas where infections were reported in either domestic, captive, or free-ranging wild waterfowl. It should be noted that no transmission has been reported from Africa, South Asia, and Mexico and that the records from these areas are from overwintering migratory Holarctic anatids. The data on which this figure is based are from Kucera ˇ (1981a), Valkiunas ¯ (Personal communications to D. J. Forrester, June 29, 2007, September 4, 2007, and October 22, 2007), and the references given in Table 4.4.
the host cell is more important than the morphology of the parasite; (4) most morphologic features overlap among various species and must be used carefully in species descriptions; and (5) the use of the morphology of meronts for taxonomic purposes is not always valid since these are unknown for most species, and some species (Leucocytozoon simondi, for example) produce meronts that are quite different depending on the avian host. Over time, several points of view on the host specificity of avian leucocytozoids have developed (Valki¯unas 2005). Early workers described new species on the basis of “a new host equals a new species,” and this resulted in the proliferation of a large number of named species, many of which had either minor or no morphometric differences. A limited number of experimental attempts to transmit leucocytozoids to
“abnormal” avian hosts (i.e., hosts of another avian family) via sporozoites from appropriate simuliid vectors eventually led to the idea that leucocytozoids are specific to host family. These attempts have been summarized by Bennett et al. (1991c). A more critical review of the literature indicates, however, that (1) three of the transmission attempts cited were not from published papers, but rather were “personal communications” and are of doubtful scientific value since details are not available; (2) one of the citations (Fallis and Bennett 1958) was incorrect; that is, it contained no information on the attempted transfer of Leucocytozoon bonasae from grouse to ducks and sparrows, rather they transferred L. bonasae from grouse to grouse; (3) the statement that Fallis et al. (1973) failed to transfer L . schoutedeni from chickens to guinea fowl and Leucocytozoon neavei from guinea fowl to chickens was
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Table 4.3. Distribution of species of Leucocytozoon by orders and families of birds. Avian order
Avian family
Struthioniformes Tinamiformes Sphenisciformes Gaviiformes Podicipediformes Procellariiformes Pelecaniformes
Struthionidae (Ostriches) — Spheniscidae (Penguins) Gaviidae (Loons) — — Anhingidae (Anhingas) Phalacrocoracidae (Cormorants) Ciconiiformes Ardeidae (Herons, Egrets, and Bitterns) Ciconiidae (Storks) Threskiornithidae (Ibises and Spoonbills) Ardeidae (Herons, Egrets, and Bitterns) Balaenicipitidae (Shoebills) Phoenicopteriformes — Anseriformes Anatidae (Ducks, Geese, and Swans) Falconiformes Accipitridae (Hawks, Eagles, and Kites) Cathartidae (New World Vultures) Falconidae (Falcons and Caracaras) Pandionidae (Ospreys) Sagittariidae (Secretary birds) Galliformes Meleagrididae (Turkeys) Cracidae (Guans, Chachalacas, and Curassows) Numididae (Guineafowl) Tetraonidae (Grouse, Ptarmigans, and Prairie Chickens) Phasianidae (Pheasants and Partridges)
Opisthocomiformes — Gruiformes Gruidae (Cranes) Otididae (Bustards) Rallidae (Rails, Gallinules, and Coots) Charadriiformes Charadriidae (Plovers and Lapwings) Scolopacidae (Sandpipers) Charadriidae (Plovers and Lapwings) Jacanidae (Jacanas) Recurvirostridae (Avocets and Stilts) Rostratulidae (Painted snipes) Sternidae (Terns) Pterocliformes — Columbiformes Columbidae (Pigeons and Doves) Psittaciformes Psittacidae (Parrots) Cacatuidae (co*ckatoos) Musophagiformes Musophagidae (Turacos) Cuculiformes Cuculidae (Cuckoos) Strigiformes Strigidae (Owls) Tytonidae (Barn Owls) Caprimulgiformes Caprimulgidae (Nightjars) Podargidae (Frogmouths) Apodiformes Trochilidae (Hummingbirds) Coliiformes Coliidae (Mousebirds) Trogoniformes Trogonidae (Trogons and Quetzals)
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Leucocytozoon species Leucocytozoon struthionis — Leucocytozoon tawaki Leucocytozoon sp* — — Leucocytozoon vandenbrandeni Leucocytozoon leboeufi Leucocytozoon nycticoraxi Leucocytozoon sp. — Leucocytozoon simondi Leucocytozoon toddi
Leucocytozoon smithi Leucocytozoon sp. Leucocytozoon neavei Leucocytozoon lovati Leucocytozoon cheissini Leucocytozoon macleani Leucocytozoon caulleryi Leucocytozoon schoutedeni — Leucocytozoon grusi Leucocytozoon sp. Leucocytozoon legeri Leucocytozoon sousadiasi Leucocytozoon sp.
— Leucocytozoon marchouxi Leucocytozoon sp. Leucocytozoon dizini Leucocytozoon centropi Leucocytozoon danilewskyi Leucocytozoon caprimulgi Leucocytozoon sp. Leucocytozoon colius Leucocytozoon sp. (continues)
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Table 4.3. (Continued ) Avian order Coraciiformes
Avian family
Leucocytozoon species
Coraciidae (Rollers) Bucerotidae (Hornbills) Upupidae (Hoopoes)† Coraciidae (Rollers) Momotidae (Motmots) Alcedinidae (Kingfishers)
Leucocytozoon bennetti Leucocytozoon communis
Meropidae (Bee-eaters) Piciformes Passeriformes
Capitonidae (Barbets) Picidae (Woodpeckers) Laniidae (Shrikes) Malaconotidae (Bushshrikes) Corvidae (Crows, Jays, and Magpies) Nectariniidae (Sunbirds and Spiderhunters) Zosteropidae (White-eyes) Certhiidae (Creepers) Emberizidae (Buntings and Sparrows) Estrildidae (Waxbills) Hirundinidae (Swallows) Icteridae (Troupials) Parulidae (New World Warblers) Ploceidae (Weavers) Prionopidae (Helmetshrikes) Prunellidae (Accentors) Thraupidae (Tanagers) Tyrannidae (Tyrant Flycatchers) Viduidae (Indigobirds) Oriolidae (Old World Orioles) Paradoxornithidae (Parrotbills) Passeridae (Old World Sparrows) Pittidae (Pittas) Alaudidae (Larks) Bombycillidae (Waxwings) Cardinalidae (Saltators and Cardinals) Chloropseidae (Leafbirds) Fringillidae (Siskins and Crossbills) Mimidae (Mockingbirds and Thrashers) Motacillidae (Wagtails and Pipits) Muscicapidae (Old World Flycatchers)
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Leucocytozoon eurystomi Leucocytozoon sp. Leucocytozoon communis Leucocytozoon eurystomi Leucocytozoon nyctyornis Leucocytozoon eurystomi Leucocytozoon nyctyornis Leucocytozoon squamatus Leucocytozoon balmorali Leucocytozoon berestneffi Leucocytozoon sakharoffi Leucocytozoon dubreuili Leucocytozoon fringillinarum
Leucocytozoon majoris
Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon dubreuili Leucocytozoon fringillinarum Leucocytozoon dubreuili Leucocytozoon fringillinarum Leucocytozoon dubreuili Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon dubreuili Leucocytozoon majoris (continues)
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Table 4.3. (Continued ) Avian order
Avian family
Leucocytozoon species
Paridae (Chickadees and tit*) Pynonotidae (Bulbuls) Sturnidae (Starlings) Sylviidae (Old World Warblers)
Timaliidae (Babblers) Turdidae (Thrushes)
Vireonidae (Vireos) Aegithinidae (Ioras) Bucconidae (Puffbirds) Campephagidae (Cuckoo-shrikes) Cinclidae (Dippers) Climactreridae (Australian Treecreepers) Corcoracidae (Choughs and Apostlebirds) Cracticidae (Bellmagpies) Dicaeidae (Flowerpickers) Dicruridae (Drongos) Eurylaimidae (Broadbills) Furnariidae (Ovenbirds) Grallinidae (Mudnest Builders) Indicatoridae (Honeyguides) Irenidae (Fairy-bluebirds) Meliphagidae (Honeyeaters) Paradisaeidae (birds of paradise) Philepittidae (Asities) Picathartidae (Rockfowl) Pipridae (Manakins) Ptilogonatidae (Silky-flycatchers) Ptilonorhynchidae (Bowerbirds) Regulidae (Kinglets) Sittidae (Nuthatches) Troglodytidae (Wrens) Vangidae (Vangas)
Leucocytozoon hamiltoni Leucocytozoon majoris Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon balmorali Leucocytozoon dubreuili Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon dubreuili Leucocytozoon fringillinarum Leucocytozoon maccluri Leucocytozoon majoris Leucocytozoon dubreuili Leucocytozoon fringillinarum Leucocytozoon majoris Leucocytozoon sp.
Source: Prepared from data presented by Bennett and Campbell (1975), Greiner and Kocan (1977), Bennett et al. (1982, 1992a), Bishop and Bennett (1992), Forrester et al. (1994), Super and van Riper (1995), Rintam¨aki et al. (1999), Adlard et al. (2002, 2004), Valki¯unas et al. (2002), Savage (2004), Savage et al. (2004, 2006a, b), Jones et al. (2005), Peirce et al. (2005), and Valki¯unas (2005). * Identity of species uncertain. † Some ornithologists consider the family Upupidae to be in another order (Upupiformes), but we follow Clements (2000) for the purposes of this analysis.
61
Mallard
Common name
Anas platyrhynchos
Species Belarus Canada (Alberta) Canada (Alberta and Saskatchewan) Canada (Labrador) Canada (Manitoba and Saskatchewan) Canada (Maritime Provinces) Canada (Newfoundland) Canada (Northwest Territories) Canada (Nova Scotia) Canada (Ontario) Canada (Quebec) Czechoslovakia Germany Lithuania Lithuania Mexico (Coahuila) Norway (Rendalen) Portugal Kazakhstan Russia (Salechard) Russia (Tomsk) Russia (Volga River Delta) USA (California) USA (California) USA (Colorado) USA (Maryland) USA (Massachusetts) USA (Michigan) USA (Minnesota) USA (Ohio) USA (Oklahoma) USA (South Dakota) USA (Washington) USA (Wisconsin) USA (Wisconsin)
Location
Table 4.4. Reports of Leucocytozoon simondi in wild waterfowl.
U 51 2,667 33 85 23 —* — — — — — ∼50 19 46 10 — U 27 — 37 17 15 368 110 59 624 220 — — 402 169 837 174 208
Number of examinations U 27 19 18 <1 13 —* — — — — — NG 26 15 20 — U 4 — 57 41 13 <1 10 2 15 4 — — 9 28 24 62 1
Prevalence (%) Dyl’ko (1966) Williams et al. (1977) Bennett et al. (1982a) Bennett et al. (1991b) Burgess (1957) Bennett et al. (1975) IRCAH records Williams et al. (1977) IRCAH records Karstad (1965) IRCAH records IRCAH records B¨oing (1925) Valki¯unas (1985) Valki¯unas et al. (1990) Bennett et al. (1991a) Eide et al. (1969) Fran¸ca (1912) Yakunin and Zhazyltaev (1977) Valki¯unas et al. (1990) Valki¯unas† Valki¯unas† Wood and Herman (1943) Herman (1951) Stabler et al. (1975) Wetmore (1941) Bennett et al. (1974a) DeJong and Muzzall (2000) Green et al. (1938) Al-Dabagh (1964) Kocan et al. (1979) Polcyn and Johnson (1968) Clark (1980) Trainer et al. (1962) Bradshaw and Trainer (1966)
Literature source
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62
63
Anas querquedula
Anas clypeata
Northern Shoveler
Anas acuta
Canada (Alberta) Canada (Alberta & Saskatchewan) Canada (Labrador) Canada (Manitoba & Saskatchewan) Canada (Maritime Provinces) Canada (New Brunswick) Canada (Northwest Territories) Canada (Nova Scotia) Canada (Prince Edward Island) Canada (Quebec) India (Rajasthan) Russia (Chaun River Delta) Russia (Salechard) Russia (Kliuchi, Kamchatka) Russia (Ust-Kara) USA (California) USA (California) USA (Colorado) USA (Louisiana) USA (Maryland) USA (Wisconsin) Germany India (Rajasthan) Iran (Dashti Arjan) Italy Kazakhstan Russia (Volga River Delta) Russia (Salechard) Canada (Alberta and Saskatchewan) Russia (Kliuchi, Kamchatka) Russia (Salechard) USA (California) USA (Colorado) USA (Florida)
49 505 14 138 228 — 18 — — — 66 26 48 22 — 24 263 68 — — — NG 34 — — 10 — 60 10 — — 55 — —
18 11 35 <1 18 — 4 — — — 6 58 83 45 — 29 1 16 — — — — 3 — — 20 — 87 10 — — 11 — — (continues)
Williams et al. (1977) Bennett et al. (1982a) Bennett et al. (1991b) Burgess (1957) Bennett et al. (1975) IRCAH records Williams et al. (1977) IRCAH records IRCAH records Laird and Bennett (1970) McClure et al. (1978) Valki¯unas et al. (1990) Valki¯unas et al. (1990) Valki¯unas et al. (1990) Valki¯unas et al. (1990) Wood and Herman (1943) Herman (1951) Stabler et al. (1975) O’Roke (1934) Wetmore (1941) Trainer et al. (1962) B¨oing (1925) McClure et al. (1978) IRCAH records Peirce (1981) Yakunin and Zhazyltaev (1977) Valki¯unas† Valki¯unas et al. (1990) Bennett et al. (1982a) Valki¯unas et al. (1990) Valki¯unas et al. (1990) Herman (1951) Stabler et al. (1975) Forrester and Spalding (2003)
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Garganey
Northern Pintail
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Species Anas discors
Anas crecca
Common name
Blue-winged Teal
Eurasian Teal
Table 4.4. (Continued )
Canada (Alberta) Canada (Labrador) Canada (Manitoba) Canada (Maritime Provinces) Canada (New Brunswick) Canada (Nova Scotia) Canada (Prince Edward Island) Canada (Alberta and Saskatchewan) USA (Colorado) USA (Massachusetts) USA (Oklahoma) USA (Texas) Canada (Alberta and Saskatchewan) Canada (Labrador) Canada (Manitoba and Saskatchewan) Canada (Maritime Provinces) Canada (New Brunswick) Canada (Newfoundland) Canada (Nova Scotia) Canada (Quebec) India (Rajasthan) Iran Norway (Rendalen) Russia (Chaun River Delta) Russia (Klyuchi, Kamchatka) Russia (Salechard) Russia (Volga River Delta) USA (Colorado) USA (Maine) USA (Massachusetts) USA (Oklahoma) USA (Texas)
Location — — — 1,286 — — — 446 39 87 58 314 119 73 25 387 — — — — 75 13 — — 34 — — 35 — 87 49 89
Number of examinations — — — 4 — — — 6 18 14 5 4 38 82 8 17 — — — — 5 8 — — 47 — — 54 — 23 18 7
Prevalence (%) IRCAH records Bennett et al. (1991b) IRCAH records Bennett et al. (1975) IRCAH records IRCAH records IRCAH records Bennett et al. (1982a) Stabler et al. (1975) Bennett et al. (1974) Kocan et al. (1979) Loven et al. (1980) Bennett et al. (1982a) Bennett et al. (1991b) Burgess (1957) Bennett et al. (1975) IRCAH records IRCAH records IRCAH records Laird and Bennett (1970) McClure et al. (1978) McClure et al. (1978) Eide et al. (1969) Valki¯unas et al. (1990) Valki¯unas et al. (1990) Valki¯unas et al. (1990) Valki¯unas† Stabler et al. (1975) Nelson and Gashwiler (1941) Bennett et al. (1974a) Kocan et al. (1979) Fedynich et al. (1993)
Literature source
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64
Anas rubripes
American Black Duck
Anas falcata Anas formosa Anas penelope
Anas strepera
Gadwall
Canada (Alberta and Saskatchewan) Canada (Maritime Provinces) Canada (New Brunswick) USA (California) USA (Colorado) USA (Maryland) USA (Oklahoma) USA (Texas) Canada (Alberta and Saskatchewan) USA (Colorado) USA (Florida) Canada (Labrador) Canada (Labrador) Canada (Maritime Provinces) Canada (New Brunswick) Canada (Newfoundland) Canada (Nova Scotia) Canada (Ontario) Canada (Prince Edward Island) Canada (Quebec) USA (Maine) USA (Maine) USA (Maryland) USA (Massachusetts) USA (Massachusetts) USA (Michigan) USA (New York) USA (Nebraska) USA (Ohio) USA (Washington, DC) USA (Wisconsin) Russia (Kliuchi, Kamchatka) Russia (Kliuchi, Kamchatka) India (Rajasthan and Tamil Nadu)
28 180 — 43 40 — 104 64 24 16 — 20 382 1,750 — — — — — — 408 29 89 85 203 — — — 13 — — 23 — 44
18 2 — 5 23 — 11 5 4 25 — 25 71 23 — — — — — — 75 89 7 13 28 — — — 38 — — 48 — 11
65
(continues)
Bennett et al. (1982a) Bennett et al. (1975) IRCAH records Herman (1951) Stabler et al. (1975) Wetmore (1941) Kocan et al. (1979) Fedynich et al. (1993) Bennett et al. (1982a) Stabler et al. (1975) Forrester and Spalding (2003) Bennett (1972) Bennett et al. (1991b) Bennett et al. (1975) IRCAH records IRCAH records IRCAH records Clarke (1946) IRCAH records Laird and Bennett (1970) O’Meara (1956) Nelson and Gashwiler (1941) Williams and Bennett (1978) Herman (1938) Bennett et al. (1974a) DeJong and Muzzall (2000) Reilly (1956) IRCAH records Al-Dabagh (1964) IRCAH records Trainer et al. (1962) Valki¯unas et al. (1990) Valki¯unas et al. (1990) McClure et al. (1978)
September 29, 2008
Falcated Duck Baikal Teal Eurasian Wigeon
Anas americana
American Wigeon
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Netta rufina Clangula hyemalis Mergus merganser
Red-crested Pochard Long-tailed Duck
Common Merganser
Aythya valisineria
Canvasback
Aythya ferina Aythya fuligula
Aythya americana
Redhead
Common Pochard Tufted Duck
Aythya affinis
Lesser Scaup
Aythya collaris
Aythya marila
Greater Scaup
Number of examinations — 51 — 63 — — — 39 180 23 17 13 — 88 — 178 — — 283 — — 26 — — 40 — — 15 — — 21 — 55
Location Russia (Kliuchi, Kamchatka) Russia (Salechard) Canada (Labrador) Russia (Kliuchi, Kamchatka) Russia (Salechard) Russia (Ust-Kara) Canada (Northwest Territories) USA (Colorado) USA (Texas) Canada (Alberta and Saskatchewan) USA (California) USA (Colorado) USA (Louisiana) USA (Maryland) USA (Washington, DC) Canada (Maritime Provinces) Canada (New Brunswick) Canada (Prince Edward Island) USA (Florida) USA (Louisiana) Kazakhstan India (Rajasthan) Macedonia Russia (Kliuchi, Kamchatka) Russia (Salechard) Kazakhstan North America Russia (Chaun River Delta) Canada (Labrador) Canada (Ontario) USA (Colorado) USA (Maine) USA (Michigan)
— 92 — 35 — — — 8 36 7 12 23 — 6 — 4 — — 9 — — 8 — — 88 — — 53 — — 5 — 47
Prevalence (%) Valki¯unas et al. (1990) Valki¯unas et al. (1990) Bennett et al. (1991b) Valki¯unas et al. (1990) Valki¯unas et al. (1990) Valki¯unas et al. (1990) IRCAH records Stabler et al. (1975) Loven et al. (1980) Bennett et al. (1982a) Wood and Herman (1943) Stabler et al. (1975) O’Roke (1934) Kocan and Knisley (1970) IRCAH records Bennett et al. (1975) IRCAH records IRCAH records Forrester et al. (2001b) O’Roke (1934) Yakunin and Zhazyltaev (1977) McClure et al. (1978) W¨ulker (1919) Valki¯unas et al. (1990) Valki¯unas et al. (1990) Yakunin and Zhazyltaev (1977) Herman (1963) Valki¯unas et al. (1990) Bennett et al. (1991b) Fallis et al. (1954) Stabler et al. (1975) Nelson and Gashwiler (1941) DeJong et al. (2001)
Literature source
September 29, 2008
Ring-necked Duck
Species
Common name
Table 4.4. (Continued )
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66
Mergus serrator Mergellus albellus Lophodytes cucullatus
Melanitta perspicillata Melanitta nigra Bucephala clangula Somateria fischeri Somateria mollissima Aix sponsa
Red-breasted Merganser
Smew Hooded Merganser
Surf Scoter Black Scoter
Common Goldeneye
Spectacled Eider Common Eider
Wood Duck
Canada (Quebec) USA (California) USA (Massachusetts) Russia (Salechard) Canada (Quebec) USA (Maine) USA (Massachusetts) USA (Texas) Canada (Labrador) Canada (Labrador) USA (North Carolina) Canada (Maritime Provinces) Canada (New Brunswick) USA (Maine) Russia (Chaun River Delta) Canada (Labrador) Canada (Newfoundland) Russia (Ust-Kara) Canada (Maritime Provinces) Canada (New Brunswick) Canada (Nova Scotia) Canada (Ontario) USA (Connecticut) USA (Florida) USA (Georgia) USA (Maine) USA (Maine) USA (Maine) USA (Maine) USA (Maine) USA (Maryland) USA (Maryland) USA (Missouri)
— — — 10 — — 11 20 — — — — — — 16 18 126 — 51 — — 66 79 2,143 729 77 322 13 24 944 11 93 371
— — — 20 — — 9 10 — — — — — — 44 <1 <1 — 14 — — 16 4 2 2 59 51 69 83 70 9 1 1
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67
(continues)
Laird and Bennett (1970) Wood and Herman (1943) IRCAH records Valki¯unas et al. (1990) Laird and Bennett (1970) Nelson and Gashwiler (1941) Bennett et al. (1974a) Loven et al. (1980) Bennett (1972) IRCAH records NWHC records Bennett et al. (1975) IRCAH records Nelson and Gashwiler (1941) Valki¯unas et al. (1990) Bennett (1972) Bennett and Inder (1972) Valki¯unas et al. (1990) Bennett et al. (1975) IRCAH records IRCAH records Thul and O’Brien (1990) Thul and O’Brien (1990) Thul and O’Brien (1990) Thul and O’Brien (1990) Nelson and Gashwiler (1941) O’Meara (1956) Herman et al. (1971) Thul et al. (1980) Thul and O’Brien (1990) Thul et al. (1980) Thul and O’Brien (1990) Odell and Robbins (1994)
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68
Aix galericulata Oxyura jamaicensis Cygnus buccinator
Cygnus columbianus
Cygnus olor
Branta canadensis
Tundra Swan
Mute Swan
Canada Goose
Species
Number of examinations 230 730 1,066 46 721 912 35 10 840 26 608 225 1,418 217 40 U — 75 38 — — — — 62 — — 66 —
Location USA (Massachusetts) USA (Massachusetts) USA (Massachusetts) USA (New Hampshire) USA (New York) USA (North Carolina) USA (Ohio) USA (Pennsylvania) USA (Pennsylvania) USA (Rhode Island) USA (South Carolina) USA (Vermont) USA (Virginia) USA (West Virginia) USA (Wisconsin) Northeastern Asia USA (Maryland) Canada (Alberta) Canada (Northwest Territories) Canada (Northwest Territories) USA (Alaska) USA (Michigan) USA (Utah) Sweden USA (New Hampshire) Canada (Labrador) Canada (Maritime Provinces) Canada (New Brunswick)
<1 33 5 26 4 2 3 10 1 8 3 5 2 5 8 U — 1 21 — — — — 16 — — 2 —
Prevalence (%)
Herman et al. (1971) Bennett et al. (1974a) Thul and O’Brien (1990) Thul and O’Brien (1990) Thul and O’Brien (1990) Thul and O’Brien (1990) Herman et al. (1971) Thul et al. (1980) Thul and O’Brien (1990) Thul and O’Brien (1990) Thul and O’Brien (1990) Thul and O’Brien (1990) Thul and O’Brien (1990) Thul and O’Brien (1990) Trainer et al. (1962) Valki¯unas (2005) Williams and Bennett (1978) Bennett et al. (1981b) Bennett et al. (1992b) IRCAH records IRCAH records IRCAH records IRCAH records M¨orner and Wahlstr¨om (1983) IRCAH records Bennett (1972) Bennett et al. (1975) IRCAH records
Literature source
September 29, 2008
Mandarin Duck Ruddy Duck Trumpeter Swan
Common name
Table 4.4. (Continued )
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69
Chen canagica Anser albifrons Anser anser
Anser cygnoides Anser fabalis
Emperor Goose Greater White-fronted Goose
Graylag Goose
Swan Goose Taiga Bean-Goose
— 9 — 3 4 1 — 2 — — — 4 — 1 — — 4 — — NG 7 U U —
Laird and Bennett (1970) Levine and Hanson (1953) IRCAH records Herman (1938) DeJong and Muzzall (2000) Kocan et al. (1979) Trainer et al. (1962) Bradshaw and Trainer (1966) Wood and Herman (1943) IRCAH records Minchin (1910) Bennett and MacInnes (1972) Bennett et al. (1982a) Hollmen et al. (1998) IRCAH records Wood and Herman (1943) Kloss et al. (2003) IRCAH records IRCAH records B¨oing (1925) Yakunin and Zhazyltaev (1977) Fran¸ca (1912) Valki¯unas (2005) Valki¯unas et al. (1990)
IRCAH, International Reference Centre for Avian Haematozoa, Queensland Museum, South Brisbane, Queensland, Australia; NWHC, National Wildlife Health Center, U.S. Geological Survey, Madison, WI, USA; NG, not given by author; U, values not known: either not given by author or we were unable to obtain the reference. * Number of birds examined was less than 10 and therefore the prevalence was not calculated. † Personal communications to D. J. Forrester, June 29, 2007, September 4, 2007, and October 22, 2007.
Branta hutchinsii Branta bernicla Alopochen aegyptiaca Chen caerulescens
— 353 — 31 77 98 — 175 — — — 570 — 134 — — 46 — — ∼ 200 42 U U —
September 29, 2008
Cackling Goose Brant Egyptian Goose Snow Goose
Canada (Quebec) USA (Illinois) USA (Maine) USA (Massachusetts) USA (Michigan) USA (Oklahoma) USA (Wisconsin) USA (Wisconsin) USA (California) Canada (Northwest Territories) Uganda Canada (Northwest Territories) Canada (Alberta and Saskatchewan) USA (Alaska) Canada (Northwest Territories) USA (California) USA (Texas) Canada (New Brunswick) Canada (Quebec) Germany Kazakhstan Portugal North Central Asia Russia (Salechard)
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Columba palumbus
Columba arquatrix Columba oenas Columba eversmanni Columba larvata Columba vitiensis Nesoenas mayeri Geophaps lophotes
Common Wood-Pigeon
Rameron Pigeon Stock Dove Pale-backed Pigeon
Lemon Dove Metallic Pigeon Pink Pigeon
Crested Pigeon
Columba guinea
Speckled Pigeon
Patagioenas fasciata
Columba livia
Rock Pigeon
70 Australia
Australia Azerbaijan Belarus England Georgia Iraq Kazakhstan South Africa Tajikistan Turkey Turkmenistan USA (Colorado) Nigeria South Africa South Africa USA (California) USA (Colorado) USA (Colorado) Mexico England England Germany Kazakhstan Morocco Poland Uganda Kazakhstan Kazakhstan Kazakhstan (Tyan-Shan) Ethiopia NG Mauritius
Location 27 U U — U 12 —* 33 75 16 — 86 — 11 50 105 109 364 — 109 22 128 19 10 — — 337 301 — 18 U 313 328 127
Number of examinations 4 U U — U 8 —* 3 1 31 — 1 — 9 2 30 36 65 — 15 5 30 32 30 — — 12 5 — 11 U 29 18 1
Prevalence (%) Adlard et al. (2004) Zeiniev (1975) Dyl’ko (1966) Coles (1914) Burtikashvili (1978) Shamsuddin and Mohammad (1980) Yakunin and Zhazyltaev (1977) Earl´e and Little (1993) Shakhmatov et al. (1972) Ozmen et al. (2005) Berdyev (1979) Stabler and Holt (1963) Cowper (1969) Thomas and Dobson (1975) Earl´e and Little (1993) Stabler et al. (1977) Stabler and Holt (1963) Stabler et al. (1977) Stabler et al. (1977) Baker (1974) Peirce (1980) B¨oing (1925) Yakunin and Zhazyltaev (1977) Gaud and Petitot (1945) Ramisz (1962) Valki¯unas et al. (2005) Yakunin and Zhazyltaev (1977) Yakunin and Zhazyltaev (1977) Kairullaev and Yakunin (1982) Ashford et al. (1976) Berson (1964) Swinnerton et al. (2005) Bunbury et al. (2007) Adlard et al. (2004)
Literature source
September 29, 2008
Band-tailed Pigeon
Species
Common name
Table 4.5. Reports of Leucocytozoon marchouxi in wild pigeons and doves.
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71
Spotted Dove
Philippines India Japan Nigeria Philippines NG Philippines USA (Arizona) USA (District of Columbia) USA (California) USA (Colorado) USA (Florida) USA (Georgia) USA (Illinois) USA (Iowa) USA (Maryland) USA (New Jersey) USA (New Mexico) USA (Ohio) USA (Vermont) Zenaida asiatica Mexico Columbina talpacoti NG Streptopelia semitorquata Ethiopia Kenya Mauritius Uganda Streptopelia chinensis Japan Mauritius Philippines USA (California)† Phapitreron leucotis Treron sphenurus Treron sieboldii Treron australis Treron vernans Treron sphenurus Ducula carola Zenaida macroura
(continues)
79 1 McClure et al. (1978) — — Ray (1952) — — Murata (2002) — — Cowper (1969) 15 7 McClure et al. (1978) U U Berson (1964) — — McClure et al. (1978) — — Wood and Herman (1943) — — Wetmore (1941) — — Wood and Herman (1943) 269 14 Stabler and Holt (1963) 918 <1 Shamis and Forrester (1977) — — Thompson (1943) 464 2 Hanson et al. (1957) 41 15 Farmer (1960) 227 1 Williams and Bennett (1978) 119 4 Huffman and Cali (1983) 339 15 Guti´errez (1973) 102 3 Al-Dabagh (1964) 27 19 Barnard and Bair (1986) 72 3 Saunders (1959) U U Berson (1964) 23 9 Ashford et al. (1976) — — Bennett and Herman (1976) 60 8 Swinnerton et al. (2005) — — Minchin (1910) 111 5 Ogawa (1912) — — Swinnerton et al. (2005) 30 7 McClure et al. (1978) 25 4 Wood and Herman (1943)
September 29, 2008
White-winged Dove Ruddy Ground-Dove Red-eyed Dove
White-eared Dove Wedge-tailed Pigeon White-bellied Pigeon Madagascar Green-Pigeon Pink-necked Pigeon Wedge-tailed Pigeon Spotted Imperial-Pigeon Mourning Dove
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72 U 458 22 — U —
Streptopelia orientalis
U 14 5 — U —
Ashford et al. (1976) McClure et al. (1978) Peirce et al. (1977a) Leger and Leger (1914) Earl´e et al. (1991) Leger (1913) B¨oing (1925) Burtikashvili (1978) Shamsuddin and Mohammad (1980) Franchini (1924) Yakunin and Zhazyltaev (1977) Kairullaev and Yakunin (1982) Mohammed and Al-Taqi (1975) Gaud and Petitot (1945) Covaleda Ortega and G´allego Berenguer (1946) Ulugzadaev and Abidzhanov (1975) Yakunin and Zhazyltaev (1977) Kairullaev and Yakunin (1982) McClure et al. (1978) Ogawa (1912) McClure et al. (1978)
September 29, 2008
Oriental Turtle-Dove
5 — — — 38 — — U — U 9 19 11 12 U
Number of Prevalence examinations (%) Literature source
Uzbekistan Kazakhstan Kazakhstan (Tyan-Shan) India Japan Korea
Location 22 — — — 40 — — U — U 194 33 19 25 U
Species
Streptopelia senegalensis Ethiopia India Kenya Senegal South Africa Eurasian Turtle-Dove Streptopelia turtur Corsica Germany Georgia Iraq Italy Kazakhstan Kazakhstan (Tyan-Shan) Kuwait Morocco Spain
Laughing Dove
Common name
Table 4.5. (Continued )
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73 Turtur tympanistria Turtur afer Turtur abyssinicus
Tambourine Dove
Blue-spotted Wood-Dove Black-billed Wood-Dove
South Africa Ethiopia Kenya Vietnam Philippines India Malaysia South Africa Australia Mauritius Mauritius Philippines Kenya Zambia Ethiopia Tanzania Uganda Ethiopia Ethiopia
— 51 — U 235 15 — — — 13 17 178 — — 22 — 15 44 —
— 4 — U <1 7 — — — 8 12 <1 — — 5 — 7 2 —
Jansen (1952) Ashford et al. (1976) Bennett and Herman (1976) Mathis and Leger (1910) McClure et al. (1978) McClure et al. (1978) McClure et al. (1978) Jansen (1952) Reece et al. (1992) Swinnerton et al. (2005) Peirce et al. (1977b) McClure et al. (1978) Peirce et al. (1977a) Peirce (1984) Ashford et al. (1976) Bennett and Herman (1976) Bennett et al. (1974b) Ashford et al. (1976) Ashford et al. (1976)
NG, not given by author; U, values not known: either not given by author or we were unable to obtain the reference. * Number of birds examined was less than 10 and therefore the prevalence was not calculated. † Nonendemic; introduced into California where it now breeds.
Turtur chalcospilos
Streptopelia capicola Streptopelia decipiens Streptopelia lugens Streptopelia tranquebarica Streptopelia bitorquata Streptopelia decaocto Macropygia ruficeps Oena capensis Geopelia humeralis Geopelia striata
September 29, 2008
Emerald-spotted Wood-Dove
Ring-necked Dove African Mourning Dove Dusky Turtle-Dove Red Collared-Dove Island Collared-Dove Eurasian Collared-Dove Little Cuckoo-Dove Namaqua Dove Bar-shouldered Dove Zebra Dove
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Species
Location
Hooded Vulture White-backed Vulture Necrosyrtes monachus Gyps africanus
Accipitridae Gambia South Africa Zimbabwe Lappet-faced Vulture Torgos tracheliotus Zimbabwe Short-toed Eagle Circaetus gallicus Algeria Black-shouldered Kite Elanus caeruleus Ethiopia Philippines India South Africa Eurasian Buzzard Buteo buteo Czech Republic England France Germany Israel Italy Kazakhstan (Tyan-Shan) Kazakhstan Scotland Spain Sweden South Africa Ukraine Madagascar Buzzard Buteo brachypterus Madagascar Jackal Buzzard Buteo rufofuscus Kenya Long-legged Buzzard Buteo rufinus Iraq Kazakhstan Turkmenistan Black Eagle Ictinaetus malayensis Bhutan Lizard Buzzard Kaupifalco monogrammicus Sub-Saharan Africa DR of the Congo “French Congo” Guinea-Bissau Nigeria
Common name
Table 4.6. Reports of Leucocytozoon toddi in wild falconiforms.
34 35 330 —* — — — — U 99 — 26 189 — 39 19 13 — — — 18 — — — — U — — 11 — U U —
6 3 <1 —* — — — — U 38 — 46 31 — 23 84 23 — — — 33 — — — — U — — 64 — U U —
Number of Prevalence examinations (%) Todd and Wolbach (1912) Greiner and Mundy (1979) Greiner and Mundy (1979) Greiner and Mundy (1979) Sergent and Fabiani (1922) Ashford et al. (1976) McClure et al. (1978) Nandi and Mandal (1984) Enigk (1942) Svobodov´a and Vot´ypka (1998) Simpson (1991) Mikaelian and Bayol (1991) Krone et al. (2001) Bishop and Bennett (1992) Sacchi and Prigioni (1984) Kairullaev and Yakunin (1982) Yakunin and Zhazyltaev (1977) Peirce and Marquiss (1983) Peirce et al. (1983) Wingstrand (1947) Bennett et al. (1992c) Glushchenko (1962) Bennett and Blancou (1974) Peirce and Cooper (1977a) Shamsuddin and Mohammad (1980) Yakunin (1972) Berdyev (1979) McClure et al. (1978) Bennett et al. (1992c) Todd (1907) Aubert and Heckenroth (1911) Tendeiro (1947) Cox and Vickerman (1965)
Literature source
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74
Circaetus cinereus Buteo jamaicensis
DR of the Congo — USA (California) 458 USA (Colorado) 10 USA (Florida) 14 USA (Louisiana) — USA (Maryland) 16 USA (Minnesota) — USA (Nebraska) — USA (New Jersey) 10 USA (Oklahoma) 34 USA (Washington) — USA (Wyoming) — Red-shouldered Hawk Buteo lineatus USA (California) 40 USA (Florida) 22 USA (Oklahoma) — Rough-legged Hawk Buteo lagopus Germany — Kazakhstan — USA (Colorado) — USA (Oklahoma) — Ukraine U Ferruginous Hawk Buteo regalis USA (California) — USA (Colorado) 11 USA (Oklahoma) — Broad-winged Hawk Buteo platypterus Canada (Quebec) — USA (Minnesota) 10 Swainson’s Hawk Buteo swainsoni USA (Colorado) 14 USA (Montana) — Bald Eagle Haliaeetus leucocephalus Canada (British Columbia) U USA (Florida) 23 USA (Michigan) 12 USA (Minnesota) — African Fish-Eagle Haliaeetus vocifer DR of the Congo — White-tailed Eagle Haliaeetus albicilla Germany 15
Brown Snake-Eagle Red-tailed Hawk
— 26† 20 7 — 19 — — 30 35 — — 38† 5 — — — — — U —† 55 — — 80 43 — U 17 100 — — <1
Rodhain et al. (1913) Sehgal et al. (2006b) Stabler and Holt (1965) Forrester et al. (1994) Olsen and Gaunt (1985) Williams and Bennett (1978) Taft et al. (1996) Coatney and Roudabush (1937) Kirkpatrick and Lauer (1985) Kocan et al. (1977) Clark et al. (1968) Smith et al.(1998) Sehgal et al. (2006b) Forrester et al. (1994) Kocan et al. (1977) Krone et al. (2001) Valki¯unas (1989) Stabler and Holt (1965) Kocan et al. (1977) Glushchenko (1963) Sehgal et al. (2006b) Stabler and Holt (1965) Kocan et al. (1977) CCWHC records Taft et al. (1996) Stabler and Holt (1965) Coatney and Jellison (1940) Tucker and Stewart (1988) Forrester et al. (1994) Stuht et al. (1999) Stuht et al. (1999) Laveran and Nattan-Larrier (1911) Krone et al. (2001) (continues)
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76 Accipiter gentilis
Milvus migrans
Black Kite
Northern Goshawk
Aquila wahlbergi Aquila clanga Lophaetus occipitalis Spizaetus cirrhatus Milvus milvus
Wahlberg’s Eagle Greater Spotted Eagle Long-crested Eagle Changeable Hawk-Eagle Red Kite
Accipiter cooperii
Aquila pomarina Aquila rapax
Lesser Spotted Eagle Tawny Eagle
USA (Colorado) USA (Mississippi) USA (Ohio) Germany Russia (Volgograd) South Africa South Africa Kazakhstan DR of the Congo India England Germany DR of the Congo Sub-Saharan Africa Kazakhstan Russia (Volgograd) DR of the Congo USA (Arizona) USA (California) USA (Colorado) USA (Florida) USA (Michigan) USA (Minnesota) USA (New Jersey) USA (Oklahoma) USA (Wisconsin) Canada (Ontario) England Germany Spain USA (Colorado) USA (Minnesota) Wales
Location — — — 20 — 12 — — — — — 24 — 69 — — — 62 82 11 — — 27 — — 80 — 10 227 — 29 48 48
Number of examinations — — — 5 — 8 — — — — — 8 — 3 — — — 8 48† 64 — — 78 — — 59 — 30 9 — 62 50 19
Prevalence (%) Stabler and Holt (1965) NWHC records Al-Dabagh (1964) Krone et al. (2001) Kobyshev and Chashchina (1972) Bennett et al. (1992c) Bennett et al. (1992c) Valki¯unas (1989) Schwetz (1935) Nandi and Mandal (1984) Bishop and Bennett (1992) Krone et al. (2001) Rodhain et al. (1913) Bennett et al. (1992c) Yakunin and Zhazyltaev (1977) Kobyshev and Chashchina (1972) Neave (1909) Boal et al. (1998) Sehgal et al. (2006b) Stabler and Holt (1965) Forrester et al. (1994) Hartman (1927) Taft et al. (1996) Kirkpatrick and Lauer (1985) Kocan et al. (1977) Taft et al. (1994) Bennett and Fallis (1960) Peirce and Cooper (1977b) Krone et al. (2001) Peirce et al. (1983) Stabler and Holt (1965) Taft et al. (1996) Toyne and Ashford (1997)
Literature source
September 29, 2008
Cooper’s Hawk
Aquila chrysaetos
Species
Golden Eagle
Common name
Table 4.6. (Continued )
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Accipiter badius
Accipiter nisus
Eurasian Sparrowhawk
Canada (Ontario) USA (California) USA (Colorado) USA (Florida) USA (Louisiana) USA (Minnesota) USA (New Jersey) USA (New Mexico) USA (Pennsylvania) Azerbaijan Czech Republic England Germany India Iraq Italy Kazakhstan Kazakhstan (Tyan-Shan) Kazakhstan Kazakhstan Lithuania Portugal Russia Scotland Sweden Switzerland Ethiopia Guinea-Bissau India Kazakhstan Mali Sub-Saharan Africa Zambia
— — — 11 — 55 166 75 83 U 308 307 132 — — — 146 — 536 11 21 — U 195 — — — U 12 11 — 15 —
— — — 9 — 73 60 24 17 U 29 23 5 — — — 17 — 94 45† 48 — U 67 — — — U 33 27 — 80 —
Clarke (1946) Wood and Herman (1943) Stabler and Holt (1965) Forrester et al. (1994) Olsen and Gaunt (1985) Taft et al. (1996) Kirkpatrick and Lauer (1985) Smith et al. (2004) Powers et al. (1994) Zeiniev (1975) Svobodov´a and Vot´ypka (1998) Ashford et al. (1991) Krone et al. (2001) McClure et al. (1978) Shamsuddin and Mohammad (1980) Sacchi and Prigioni (1984) Yakunin and Zhazyltaev (1977) Kairullaev and Yakunin (1982) Valki¯unas (1989) Sehgal et al. (2006b) Valki¯unas (1985) Fran¸ca (1912) Valki¯unas (1985) Peirce and Marquiss (1983) Wingstrand (1947) Geigy et al. (1962) Ashford et al. (1976) Tendeiro (1947) McClure et al. (1978) Valki¯unas (1989) Commes (1918) Bennett et al. (1992c) Peirce (1984) (continues)
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Shikra
Accipiter striatus
Sharp-shinned Hawk
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Accipiter minullus Accipiter gularis Accipiter tachiro Accipiter soloensis Accipiter virgatus Accipiter brevipes Melierax metabates Circus cyaneus
Circus macrourus
Circus pygargus Kaupifalco monogrammicus
Japanese Sparrowhawk
African Goshawk
Chinese Goshawk Besra
Levant Sparrowhawk
Dark Chanting Goshawk
Northern Harrier
Pallid Harrier
Montagu’s Harrier
Lizard Buzzard
Species
Little Sparrowhawk
Common name
Table 4.6. (Continued )
— — 10 — — — — — — — 10 — — — — — 20 — — 32 27 — U —
— — — —
Number of examinations
— — 10 — — —† — — — — 30 —† — — — — 20 — — 22 78 — U —
— — — —
Prevalence (%)
Ashford et al. (1976) Bishop and Bennett (1992) McClure et al. (1978) McClure et al. (1978) McClure et al. (1978) Sehgal et al. (2006b) McClure et al. (1978) Rousselot (1953) Bennett et al. (1992c) Sacchi and Prigioni (1984) Yakunin and Zhazyltaev (1977) Sehgal et al. (2006b) Stabler and Holt (1965) Taft et al. (1996) Kirkpatrick and Lauer (1985) Franchini (1923) Yakunin and Zhazyltaev (1977) Kairullaev and Yakunin (1982) Abidzhanov (1967) Yakunin and Zhazyltaev (1977) Valki¯unas (1989) Sacchi and Prigioni (1984) Tendeiro (1947) Bishop and Bennett (1992)
Ashford et al. (1976) Earl´e et al. (1991) McClure et al. (1978) McClure et al. (1978)
Literature source
September 29, 2008
Ethiopia South Africa Philippines India Malaysia Kazakhstan Thailand Mali South Africa Italy Kazakhstan USA (California) USA (Colorado) USA (Minnesota) USA (New Jersey) Italy Kazakhstan Kazakhstan (Tyan-Shan) Uzbekistan Kazakhstan Kazakhstan Italy Guinea-Bissau South Africa
Ethiopia South Africa Malaysia Philippines
Location
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Circus aeruginosus
Micronisus gabar Pernis apivorus
Coragyps atratus Cathartes aura Milvago chimango Falco peregrinus
Falco columbarius Falco tinnunculus
Western Marsh-Harrier
Gabar Goshawk European Honey-Buzzard
Black Vulture Turkey Vulture
Chimango Caracara Peregrine Falcon
Merlin
Eurasian Kestrel
Germany Iraq Italy Kazakhstan Lithuania Russia (Volgograd) Tadzhikistan Ethiopia Egypt Kazakhstan (Tyan-Shan) Spain Cathartidae USA (Florida) USA (Maryland) Falconidae Chile Australia England Japan Kuwait Malaysia England Kazakhstan Russia (Volgograd) Finland Germany Italy Kazakhstan Kazakhstan South Africa
79 15 — — — — — — — — 227 136 20 26 16 —
Forrester et al. (2001a) Raidal et al. (1999) Peirce and Cooper (1977b) Ogawa (1912) Tarello (2006) McClure et al. (1978) Peirce (1980) Yakunin and Zhazyltaev (1977) Kobyshev et al. (1975) Korpim¨aki et al. (1995) Krone et al. (2001) Sacchi and Prigioni (1984) Yakunin and Zhazyltaev (1977) Valki¯unas (1989) Bennett et al. (1992c) (continues)
Forrester and Spalding (2003) Wetmore (1941)
<1 3 87 — — — — — — — — <1 <1 5 27 6 —
B¨oing (1925) Shamsuddin and Mohammad (1980) Franchini (1924) Valki¯unas (1989) Valki¯unas (1985) Kobyshev and Chashchina (1972) Subkhanov (1980) Ashford et al. (1976) Mohammed (1958) Kairullaev and Yakunin (1982) Peirce et al. (1983)
— U U 67 — — — — U — —
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211 79
— U U 12 — — — — U — —
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Falco cherrug Falco eleonorae Falco rupicoloides Falco naumanni Falco cenchroides Falco subbuteo Pandion haliaetus Sagittarius serpentarius
Saker Falcon Eleonora’s Falcon Greater Kestrel Lesser Kestrel Australian Kestrel Eurasian Hobby
Osprey
Secretary-bird
Canada (Saskatchewan) Mexico USA (Colorado) USA (Oklahoma) Kuwait ¨ ais) Greece (Ag¨ South Africa Kazakhstan Australia Kazakhstan Kazakhstan Pandionidae Georgia Sagittariidae Mozambique
Location
80 —
U
442 U 58 — — 16 19 13 — 11 36
Number of examinations
—
U
<1 U 22 — — 13 5 38 — 9 6
Prevalence (%)
Travassos Santos Dias (1954)
Burtikashvili (1978)
Dawson and Bortolotti (1999) Beltr´an and Pardinas (1953) Stabler and Holt (1965) Kocan et al. (1977) Tarello (2006) Wink et al. (1979) Bennett et al. (1992c) Yakunin and Zhazyltaev (1977) Raidal and Jaensch (2000) Yakunin and Zhazyltaev (1977) Valki¯unas (1989)
Literature source
September 29, 2008
Note: Unless otherwise indicated prevalences were determined by blood film analysis. CCWHC, Canadian Cooperative Wildlife Health Centre, University of Saskatchewan, Saskatoon, Saskatchewan, Canada; NWHC, National Wildlife Health Center, U.S. Geological Survey, Madison, WI, USA; U, values not known: either not given by author or we were unable to obtain the reference. * Number of birds examined was less than 10 and therefore the prevalence was not calculated. † Prevalence determined by polymerase chain reaction technique.
Falco sparverius
Species
American Kestrel
Common name
Table 4.6. (Continued )
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Leucocytozoonosis incorrect—they did not attempt those cross family transfers, but instead successfully transferred the guinea fowl leucocytozoid to other guinea fowl and the chicken leucocytozoid to other chickens; (4) the statement that Khan and Fallis (1970a) were unable to transmit Leucocytozoon dubreuili from thrushes (Turdidae) to emberizids, icterids, and ducks was incorrect—they did not report attempts to make such transfers, but only transferred L. dubreuili from thrushes to thrushes, and (5) the claim that Skidmore (1931) was unable to transfer L. smithi from turkeys to other galliforms was incorrect—he experimentally transferred L. smithi from turkey to turkey, but did not attempt to transfer it to other galliforms. Regarding this last point, Skidmore (1931) did mention, however, that there were chickens, ducks, and geese on a farm where infected turkeys existed and that these other birds remained free of L. smithi. It should be noted here that in another paper (Solis 1973) that was not mentioned by Bennett et al. (1991c), unsuccessful attempts to experimentally transfer L. smithi from turkeys to quail, partridges, domestic ducks, and pheasants were reported. Also, not mentioned in Bennett et al. (1991c) were unsuccessful attempts by Fallis and Bennett (1966) and Fallis et al. (1954) to transmit Leucocytozoon lovati from grouse to geese and a sparrow, Leucocytozoon danilewskyi from an owl to ducks and a sparrow, L. simondi from ducks to grouse, chickens, turkeys, and pheasants as well as reported failures to transmit L. caulleryi from chickens to nine other galliform species (Morii and Kitaoka 1971). In conclusion, there is only limited experimental evidence to support the hypothesis that leucocytozoids are all specific at the avian family or subfamily level. Rather, there is substantial information that would lead us to conclude that they are specific at least to the ordinal level. Some leucocytozoids such as L. smithi of turkeys and L. caulleryi of chickens are host species specific, some are host genera specific such as Leucocytozoon sakharoffi of crows and ravens and Leucocytozoon berestneffi of jays, while some like L. simondi are family specific, and others such as L. toddi, Leucocytozoon fringillinarum, Leucocytozoon majoris, and L. dubreuili are found in numerous families, but all within the same order (Table 4.3). Valki¯unas (2005) reviewed the citations mentioned above and other literature on experimental infections and concluded that “. . . there are no scientific facts available which confirm the possibility that the same species of Leucocytozoon infect birds belonging to different orders.” There are possible exceptions to this as illustrated by the leucocytozoid infections described earlier in a Common Loon and in young Ostriches. However, these infections might have originated from birds of other orders, although this was not proven. This is unlikely to be a common phenomenon.
81
Using the family host specificity approach, the number of described species of Leucocytozoon is around 143, whereas the ordinal host specificity and distinct morphological approach results in numerous synonymies and a list of only 36 valid species (Table 4.3). There is some evidence that cryptic species of Leucocytozoon exist (Sehgal et al. 2006b) and future research combining techniques of traditional parasitology and molecular biology will most certainly result in the determination of additional synonyms and rethinking of the systematics of the leucocytozoids. Leucocytozoids have a number of stages that occur in the simuliid vector and in the blood and other tissues of the avian host. Two morphological forms of gametocytes, either round or elongate, occur in the blood of the avian host. These various stages will be discussed further in the next section in relation to the life cycle. The ultrastructure of the various stages of L. simondi has been studied by S. S. Desser and associates (Desser 1970a–c, 1972, 1973; Desser et al. 1970;), and for meronts of L. toddi by Raidal and Jaensch (2000). Strains with varying degrees of pathogenicity have been recognized for L. caulleryi in chickens and L. simondi in waterfowl (Desser et al. 1978; Morii et al. 1986) and may exist for other species. In the case of L. simondi, several strains were recognized in Canada Geese (Branta canadensis) from Michigan, USA, and Ontario, Canada. One strain from geese in the Seney National Wildlife Refuge (NWR) in the upper peninsula of Michigan underwent complete development with primary development in the liver, resulting in round gametocytes in the blood and secondary development in reticuloendothelial cells, resulting in the production of many elongate gametocytes and was associated with mortality of goslings. Other strains from geese in Cusino Wildlife Research Station (40 km west of Seney NWR), White Pine Copper Company (278 km west of Seney NWR in Michigan), and Algonquin Park in Ontario underwent development only in the liver, producing only round gametocytes, and did not result in mortality (Desser et al. 1978). A pathogenic Norwegian strain has also been recognized, in which merogony is completed more rapidly than is the case with strains in North America; other differences were recognized in the location and size of megalomeronts (Eide and Fallis 1972). Recently, partial sequences of the cytochrome b gene have been used to characterize species and perhaps subspecies of L . fringillinarum from House Sparrows (Passer domesticus) in Israel (Martinsen et al. 2006), L. schoutedeni from domestic chickens in Uganda (Sehgal et al. 2006a), and L. toddi in diurnal raptors in California, Kazakhstan, and Lithuania (Sehgal et al. 2006b). This approach will eventually help to define
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intraspecific genetic diversity and the taxonomic and phylogenetic relationships of the leucocytozoids. EPIZOOTIOLOGY Leucocytozoids have an indirect life cycle that involves biting flies of the order Diptera as vectors. All are species of Simuliidae (black flies), with the exception of L. caulleryi, which uses biting midges of the genus Culicoides. The best-studied life cycle is that of L. simondi (Figure 4.2). The works of O’Roke (1934), Huff (1942), Chernin (1952a), Desser (1967), Desser et al. (1968), Khan et al. (1969), Aikawa et al. (1970), Yang et al. (1971), and Eide and Fallis (1972) were foundational to understanding the life cycle of this species and of leucocytozoids in general. The following is a summary of the life cycle of L. simondi based on the works listed above as presented by Valki¯unas (2005). Sporozoites (stages infective to avian hosts) are injected into the blood stream by biting flies while they are taking a blood meal. These sporozoites penetrate hepatic cells where they develop into first-generation meronts (Figure 4.2, 1–3). Over a 4- or 5-day period these meronts increase in size, undergo multiple nuclear divisions, and form a number of separate sections called cytomeres. These further develop into uninuclear merozoites and syncytia containing several nuclei. Some of these merozoites and syncytia enter the blood stream. These merozoites then invade erythrocytes and develop into gametocytes (round forms) (Figure 4.2, 4–6). Syncytia are carried by the blood to many organs (spleen, lymph nodes, liver, brain, etc.), where they are engulfed by macrophages and form megalomeronts (Figure 4.2, 7–8). These megalomeronts contain thousands of merozoites that rupture from the megalomeront and in turn penetrate lymphocytes and other leukocytes and develop into gametocytes (fusiform or elongate forms) (Figure 4.2, 9–11; Figure 4.3). The dynamics of parasitemia is illustrated in Figure 4.4. The round or fusiform gametocytes (male gametocytes or microgametocytes and female gametocytes or macrogametocytes) are infective for the dipteran vector. Once the gametocytes are ingested by a blood-feeding vector, they undergo sexual reproduction and form a zygote that becomes an ookinete (Figure 4.2, 12–14). The ookinete penetrates the midgut of the vector, undergoes sporogony, and produces sporozoites (Figure 4.2, 15–18). These sporozoites then migrate to the salivary glands of the insect vector (Figure 4.2, 19) and then can be injected into the next bird when the vector takes a blood meal. While development of all leucocytozoid species that have been studied in vectors is similar, it varies de-
Figure 4.2. Diagrammatic illustration of the life cycle of Leucocytozoon simondi. Upper section of the illustration represents events that occur in the vector and lower section represents events that occur in the bird. 1, sporozoite or merozoite in hepatocyte; 2–4, hepatic meronts; 5. merozoites in erythrocytes; 6, gametocytes in round host cells; 7, syncytia of merozoites in reticuloendothelial cells; 8 and 9, megalomeronts; 10, merozoites in mononuclear leukocytes; 11, gametocytes in fusiform host cells; 12, macrogamete; 13, microgamete that is exflagellating; 14, fertilization of macrogamete; 15, ookinete penetrating the peritrophic membrane of the vector’s gut wall; 16, young oocyst; 17 and 18, sprogony; 19, sporozoites in the salivary glands of the vector. From Valkiunas ¯ (2005), with permission of the author and CRC Press.
pending on species and avian host. First-generation meronts develop in the parenchymal cells of the liver in all leucocytozoids except L. caulleryi, which develops in the endothelial cells of the capillaries of many organs (Valki¯unas 2005). First-generation meronts of L. dubreuili develop in liver cells and also in endothelial cells of the kidney (Khan and Fallis 1970a), but in L. smithi, they develop only in hepatocytes (Steele and Noblet 1992). Little is known about details of the
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Leucocytozoonosis
Figure 4.3. Illustrations of the round and elongate forms of the gametocytes of Leucocytozoon simondi based on a blood smear from a Eurasian Wigeon (Anas penelope). 1–4 and 6, macrogametocytes; and 5 and 7, microgametocytes. Uninfected erythrocyte is between 3 and 4. Chc, cytoplasm of host cell; Nc, nucleolus; Nhc, nucleus of host cell; Np, nucleus of parasite; Vc, vacuole; Vg, valutin granule. From Valkiunas ¯ et al. (1990), with permission of the author and Parazitologiya (St. Petersburg).
development and morphology of meronts or megalomeronts in the avian hosts of L. marchouxi and L. toddi, although they are known to occur. Peirce et al. (1997) described and provided photomicrographs of megalomeronts of L. marchouxi in a Pink Pigeon (Nesoenas mayeri), as did Simpson (1991) for L. toddi in a Eurasian Buzzard (Buteo buteo). Vectors are known for 14 of the 36 leucocytozoid species (Table 4.7). Studies on transmission of species of Leucocytozoon include Skidmore (1932), Fallis et al. (1956), Anderson et al. (1962), Barrow et al. (1968), Baker (1970), Noblet et al. (1975), Allison et al. (1978), Greiner and Forrester (1979), Pinkovsky et al. (1981), and Kiszewski and Cupp (1986).
83
Some species of Leucocytozoon can be transmitted by more than one species of black fly, and some species of black fly can transmit more than one species of Leucocytozoon. The geographic range of the parasite being transmitted is restricted to the range of the susceptible vector(s) as well as other ecological and behavioral factors. For example, L. simondi is absent in nonmigratory Mottled Ducks (Anas fulvigula), Fulvous Whistling-Ducks (Dendrocygna bicolor), Canada Geese, and Wood Ducks (Aix sponsa) in Florida (Thul and O’Brien 1990; Forrester and Spalding 2003) because of a number of biotic and abiotic factors, including behavioral and physiological chacteristics of both hosts and vectors. Cnephia ornithophilia is a capable vector of L. simondi (Tarshis 1972) and has been found in 14 counties in northern Florida from October through May (Pinkovsky 1976; Pinkovsky and Butler 1978), yet there is no evidence that transmission of L. simondi between infected migratory birds and uninfected resident species takes place. Cnephia ornithophilia may be either spatially separated from nonmigratory ducks and geese in Florida, or does not feed on these birds. However, this does not preclude migratory species from carrying the parasite with them when they fly south to their wintering grounds from the northern breeding range where transmission occurs. Patent infections have been reported for migratory Wood Ducks and Ring-necked Ducks (Aythya collaris; Thul and O’Brien 1990; Forrester et al. 2001b); but as these birds recover from breeding and migrate, their parasitemias may be reduced to a point where they may be too low to infect vectors. As with other vector-borne diseases and parasites, transmission of Leucocytozoon is dependent on availability of appropriate vectors and presence of a sufficient number of gametocytes in the peripheral circulation of the avian host to infect those vectors. In areas with temperate climates, this is achieved through “spring relapse” (Desser et al. 1968; Khan and Fallis 1970b). While the avian host is preparing either for spring migration to the breeding ground or for breeding, it undergoes hormonal changes that induce an increase in the number of circulating blood parasites. Day length and interspecific stress also play a role (Chernin 1952b; Barrow 1963). This raises the parasitemia to a level that facilitates infection of vectors just prior to the production of na¨ıve young of the year. Parasitemias during spring relapse vary among different species of waterfowl. In one study in Ontario, Canada, parasitemias were higher in American Black Ducks (Anas rubripes) than in Mallards (Anas platyrhynchos) or domestic ducks (Khan and Fallis 1968). Parasitemias typically decrease after the breeding season and are maintained at a low level. This undoubtedly conserves parasite resources by not having
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Parasites / 1000 red blood cells
Parasitic Diseases of Wild Birds
90%
50% 50%
10%
100%
90%
16
10%
14 12 10 8 6 4 2
rupture of hepatic schizonts 4
6
8
rupture of megalo schizonts 10
12
14
Days postinfection Figure 4.4. Dynamics of parasitemia of Leucocytozoon simondi in experimentally infected ducklings and the ratio of round and fusiform host cells during the development and rupture of hepatic meronts and megalomeronts. (a) Period of rupture of hepatic meronts; (b) period of rupture of megalomeronts. Ordinate indicates mean parasitemia from eight ducklings expressed as number of parasites per 1,000 erythrocytes; abscissa indicates days post inoculation of sporozoites. Adapted from Desser 1967, with permission of the author and the Journal of Protozoology.
parasites being produced when there are no vectors available for transmission and causes less damage to the avian host. Relapses have been observed in birds infected with L. danilewskyi, L. dubreuili, L. lovati, L. simondi, L. smithi, L. toddi, and some other leucocytozoids (Ashford et al. 1990; Valki¯unas 2005). The behavior of avian hosts is an important feature of the epizootiology of leucocytozoid infections. This includes nesting and roosting habits (Figure 4.5) as well as migratory behavior. Some colonial-nesting birds have a higher diversity of haemosporidian parasites, including leucocytozoids, and higher prevalances of infection than do solitary-nesting birds (Tella 2002). This is most likely related to greater efficiency of transmission where host density is high. There are exceptions to this, however; some colonial nesting birds have very low prevalences of blood parasites and some have none at all, probably due to factors that may either hin-
der or limit numbers of vectors (Valki¯unas 2005). In England, L . toddi is transmitted to adult and nestling Eurasian Sparrowhawks (Accipiter nisus) primarily at the nest site prior to dispersal of nestlings at about 2 months of age, resulting in a 33% prevalence of infection (Ashford et al. 1990, 1991). Migratory waterfowl (and other avian species as well) are exposed to a more diverse community of parasites than are nonmigratory species, and therefore have a higher risk of infection (Figuerola and Green 2000). One example of this is L . simondi infections in Wood Ducks in the Atlantic Flyway in North America. Wood Ducks were sampled from 82 sites in 19 states and provinces throughout the flyway from Florida, USA, to New Brunswick, Canada, during the nesting season and before migration began (Thul et al. 1980; Thul and O’Brien 1990). Infections with L. simondi were found in ducks from Ontario, Canada, and several northern
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Table 4.7. Species of dipterans known to serve as vectors of species of Leucocytozoon. Species of Leucocytozoon 1 2 3
Leucocytozoon balmorali Leucocytozoon bennetti Leucocytozoon berestneffi
4 5
Leucocytozoon caprimulgi Leucocytozoon caulleryi
6 7 8 9 10
Leucocytozoon centropi Leucocytozoon cheissini Leucocytozoon colius Leucocytozoon communis Leucocytozoon danilewskyi
11 12
Leucocytozoon dizini Leucocytozoon dubreuili
13 14
15 16 17 18 19
Leucocytozoon eurystomi Leucocytozoon fringillinarum
Leucocytozoon grusi Leucocytozoon hamiltoni Leucocytozoon leboeufi Leucocytozoon legeri Leucocytozoon lovati
20 21
Leucocytozoon maccluri Leucocytozoon macleani
22 23 24
Leucocytozoon majoris Leucocytozoon marchouxi Leucocytozoon neavei
Table 4.7. (Continued ). Species of Leucocytozoon
Species of vector Unknown Unknown Prosimulium decemarticulatum Simulium aureum Unknown Culicoides arakawae Culicoides circ*mscriptus Culicoides odibilis Culicoides schultzei Unknown Unknown Unknown Unknown Prosimulium decemarticulatum Simulium aureum Simulium latipes Unknown Cnephia ornithophilia Prosimulium decemarticulatum Simulium aureum Simulium croxtoni Simulium latipes Simulium quebecense Unknown Cnephia ornithophilia Prosimulium decemarticulatum Simulium aureum Simulium croxtoni Simulium latipes Simulium quebecense Unknown Unknown Unknown Unknown Simulium aureum Simulium croxtoni Simulium latipes Simulium minus Simulium quebecense Unknown Eusimulium geneculare Simulium metatarsale Unknown Unknown Simulium adersi Simulium impukane Simulium nyasalandicum
25 26 27
Leucocytozoon nycticoraxi Leucocytozoon nyctyornis Leucocytozoon sakharoffi
28
Leucocytozoon schoutedeni
29
Leucocytozoon simondi
30
Leucocytozoon smithi
31 32 33 34
Leucocytozoon sousadiasi Leucocytozoon squamatus Leucocytozoon struthionis Leucocytozoon tawaki
35
Leucocytozoon toddi
36
Leucocytozoon vandenbrandeni
Species of vector Unknown Unknown Prosimulium decemarticulatum Simulium angustitarse Simulium aureum Simulium latipes Simulium quebecense Simulium adersi Simulium impukane* Simulium nyasalandicum Simulium vorax Cnephia ornithophilia Simulium anatinum Simulium fallisi Simulium innocens Simulium latipes Simulium parnassum Simulium rendalense Simulium rugglesi Simulium venustum Simulium vittatum Prosimulium hirtipes Simulium aureum Simulium congareenarum Simulium jenningsi Simulium meridionale Simulium pictipes Simulium slossanae Simulium venustum Simulium vittatum Unknown Unknown Unknown Austrosimulium australense Austrosimulium dumbletoni Austrosimulium ungulatum Prosimulium decemarticulatum† Simulium aureum† Simulium quebecense† Unknown
Sources: Chang (1975), Greiner (1991), and Valki¯unas (2005). * Identification uncertain (Fallis et al. 1973). † Sporogony occurs in these vectors, but transmission to birds has not been proven (Bennett et al. 1993a).
85
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Figure 4.5. Transmission of Leucocytozoon smithi to sentinel domestic turkeys maintained in cages in Wild Turkey (Meleagris gallopavo) habitat at Fisheating Creek Wildlife Management Area in southern Florida during 1976 and 1977. Tree cages were located in the canopy where turkeys roost at night; ground cages were in ground-level habitat where they feed, rest, and nest. Broken lines indicate missing data due to deaths of sentinel birds unrelated to disease. From Forrester and Spalding (2003).
states (Maine, Massachusetts, New York, Maryland, and Pennsylvania), but not in any of the states south of Maryland. After migration began in the fall, infected ducks were found in southern states; they had acquired their infections in the north prior to flying south for the winter. Another example is the presence of leucocytozoids in several species of passeriforms in the Curonian Spit, which is located partly in Lithuania and partly in Russia and projects into the Baltic Sea (Valki¯unas 1993). Through a long-term banding study, it was determined that leucocytozoids were not transmitted to the migratory population of passeriforms that were hatched on the spit, but were present in the same species of passeriforms that had migrated south and had overwintered in southern Europe and Africa where they acquired the parasites and then returned to the Curonian Spit. It was determined that there were no simuliid vectors on the Curonian Spit due to the lack of flowing, well-oxygenated fresh water needed for the flies to breed.
Transmission of leucocytozoids is also dependent on a number of abiotic factors including favorable environmental conditions, particularly temperature, rainfall, humidity, and the presence or absence of running water. Running water is necessary for black fly vectors to reproduce (Adler et al. 2004), and this requirement in turn influences the transmission of leucocytozoids. For example, the prevalence of L. simondi in waterfowl is lower in years of drought than it is in normal years in western Canada (Bennett et al. 1982a). In southern Florida, there is a significant correlation between the prevalence of L. smithi in Wild Turkeys (Meleagris gallopavo) and the depth of nearby creeks. Prevalence of infection is higher during periods when water levels in streams and adjacent cypress swamps increase and available habitat for black flies expands (Figure 4.6). As a result of variation in abiotic factors, transmission of leucocytozoids occurs during restricted periods of time in northern climates or throughout the year in warmer climates. Sentinel birds have been used
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87
Figure 4.6. Comparison of prevalences of Leucocytozoon smithi in Wild Turkeys (Meleagris gallopavo) at Fisheating Creek Wildlife Management Area in southern Florida during the months of July, August, September, and October, with the depth of water in Fisheating Creek during the preceding March and April over a 15-year period, 1969–1983. From Forrester and Spalding (2003). to study the dynamics of transmission of a number of leucocytozoids including L. simondi in waterfowl (Herman and Bennett 1976) and L. smithi in Wild Turkeys (Forrester and Spalding 2003). In eastern Canada, transmission of L. simondi to sentinel ducks occurs in June and July. Transmission of L. smithi to sentinel turkeys takes place over a more extended period of time in the subtropical climate of southern Florida (Figure 4.5), but occurs only during March, April, and May in northern Florida, possibly because of weather or the absence of the primary vector (Simulium slossonae) during the rest of the year (Atkinson and van Riper 1991). CLINICAL SIGNS Clinical signs of leucocytozoonosis are usually nonspecific and may not be apparent (Wobeser 1997). Young ducks and geese are most susceptible to leucocytozoonosis and may die within a short time after infection. Ducklings may be active and normal in the morning, ill and with no interest in eating by midafternoon, and dead by the following morning (O’Roke 1934). Older birds may be listless and lose their wariness of humans, but rarely die of the disease.
Anemia is the most important clinical sign (Maley and Desser 1977) and packed cell volumes may be only 20% of normal (Fallis et al. 1951). Other signs are anorexia, lethargy, labored breathing, and diarrhea (Wobeser 1997). Some birds exhibit nervous signs such as marked excitement (O’Roke 1934) and convulsions (Khan and Fallis 1968). Doves infected with L. marchouxi have been reported to exhibit listlessness, ruffled feathers, anemia, and below average body weights (Oosthuizen and Markus 1968; Peirce 1984). Clinical signs in raptors infected with L. toddi may range from erratic flight, reduced flight speeds, lack of coordination, depression, blindness, spontaneous erratic vocalization, and seizures to anorexia, weight loss, vomiting, weakness, labored breathing, and ruffled feathers (Raidal and Jaensch 2000; Tarello 2006).
PATHOGENESIS Three species of Leucocytozoon are reported to be pathogenic to wild birds, L. simondi, L. marchouxi, and L. toddi (Table 4.2). There is some evidence that L. danilewskyi may be pathogenic to owls, but definitive
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Figure 4.7. Mean percentage hematocrits and numbers of gametocytes for ducklings infected with Leucocytozoon simondi compared with uninfected controls. Bars surrounding data points indicate standard error of the mean. From Maley and Desser (1977), with permission of the authors and the Canadian Journal of Zoology. data are lacking. There is one report that L. danilewskyi infection may have caused a reduction in egg production (Korpim¨aki et al. 1993), but necropsies as well as clinical and histopathologic studies were not performed to verify this. In another study, mortality of fledgling owls was attributed to severe black fly feeding in concert with L. danilewskyi infections (Hunter et al. 1997), but not to leucocytozoonosis alone. The beststudied species is L. simondi (see reviews in Wobeser (1997) and Valki¯unas (2005)). The pathogenesis of leucocytozoonosis in waterfowl due to L. simondi can best be understood with reference to the life cycle and development of gametocytes and exoerythrocytic stages in the tissues of infected birds over time (see Figure 4.2). It begins with the injection of sporozoites into the blood stream and their subsequent invasion of hepatic cells. Here, they undergo further development into meronts over a period of 5 days (Desser 1967). Beginning on days 4– 6 postinfection (PI), erythrocytes become fragile and
birds become anemic as numbers of erythrocytes begin to drop (Figure 4.7). Anemia is associated with the rupture of meronts and the release of merozoites and syncytia into the circulation. Merozoites invade erythrocytes and develop into round gametocytes; syncytia are carried via blood to various organs including spleen, lymph nodes, liver, and brain, and are engulfed by macrophages and form numerous megalomeronts containing thousands of merozoites. Megalomeronts are quite large, some reaching 60–189 μm in diameter (Desser 1967). It has been estimated that in some infections megalomeronts can make up to three-fourths of the mass of spleen and lymphoid tissue by day 9 PI (Desser 1967). Rupture of megalomeronts and release of merozoites occurs 9–12 days PI and coincides with the peak of erythrocyte fragility and anemia. This anemia is thought to be due to an “anti-erythrocyte factor” released from the meronts or their host cells rather than destruction of the erythrocytes by gametocytes, since the peak of anemia precedes the peak of
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Leucocytozoonosis parasitemia (Kocan and Clark 1966; Desser and Ryckman 1976). The highest mortality occurs in ducklings at day 12 PI when anemia reaches its peak and most of the megalomeronts have ruptured (Kocan and Clark 1966; Maley and Desser 1977; Valki¯unas 2005). No detailed studies have been conducted on the pathogenesis of L. marchouxi in pigeons and doves and L. toddi in raptors, although megalomeronts have been described in numerous internal organs in Pink Pigeons (Peirce et al. 1997) and in Eurasian Buzzards (Simpson 1991). Pathogenic and nonpathogenic strains of L. simondi have been recognized (Eide and Fallis 1972; Desser et al. 1978). The pathogenic strains (i.e., the Norway strain and the Seney strain) undergo primary merogony in the liver and secondary merogony and formation of megalomeronts in various additional organs. Nonpathogenic strains such as Cusino, White Pine, and Algonquin undergo only primary merogony in the liver and do not produce megalomeronts. Pathogenicity of some leucocytozoids seems to be related to the development of megalomeronts (Valki¯unas 2005). However, of the eight species of Leucocytozoon that are pathogenic to domestic and wild birds (Table 4.2), three (Leucocytozoon simondi, L. marchouxi, and L. caulleryi) produce megalomeronts, whereas two other species (Leucocytozoon macleani and L. smithi) do not. It is not known if the other two species (Leucocytozoon struthionis and L. schoutedeni) produce megalomeronts. Megalomeronts of L. toddi have been described (Simpson 1991), but lack cytomeres that are characteristic of megalomeronts in other species and may actually be large primary meronts (Peirce et al. 1997). PATHOLOGY Gross lesions in waterfowl with fatal leucocytozoonosis include enlargement of the spleen (Figure 4.8) and liver, paleness of tissues, and thin watery blood (Wobeser 1997). Histological studies of infections of L. simondi have been reported by several authors (see Wobeser 1997), but the most complete and detailed description was that of Newberne (1957). Capillaries of lungs, liver, and spleen were distended by the presence of many gametocytes, but local host tissue reaction was not evident. Megalomeronts in the brain had moderate to marked cellular reactions (Figure 4.9a), but meronts elicited moderate, slight, or no such reaction. Megalomeronts were located in close association with small blood vessels, and the host reaction was characterized by the proliferation of large mononuclear cells. Megalomeronts that had ruptured and contained no merozoites were filled with an eosinophilic coagulum and large mononuclear cells. In some cases, there were scattered
89
Figure 4.8. Gross view of a spleen from a duck infected with Leucocytozoon simondi (a) compared with one from an uninfected control (b). From Newberne (1957), with permission of the American Journal of Veterinary Research. scars that were believed to be the remnants of depleted megalomeronts. Some megalomeronts in the lung had marked host reaction consisting of several layers of lymphoid cells, plasma cells, large mononuclear cells, and fibroblasts (Figure 4.9b), whereas others had less severe reactions or none at all. Megalomeronts in other organs such as spleen (Figure 4.9c) and cardiac muscle (Figure 4.9d) had no local host tissue reactions. A variety of microscopic changes were noted in various organs. In liver, severe central necrosis was so widespread in some cases that the necrotic areas were confluent. In these areas, there were also marked periportal and diffuse lymphocytic infiltration, prominent Kupffer cells that contained hemosiderin, and macrophages containing pigment. Enlarged spleens were congested and contained many macrophages that were swollen and contained large amounts of pigment and cellular debris. The normal splenic architecture was almost completely obliterated in most birds. Some birds had pulmonary congestion and some had infiltrates of histiocyte-type cells containing cellular
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(a)
(b)
(c)
(d)
Figure 4.9. Megalomeronts of Leucocytozoon simondi in various tissues of an infected duck. Hematoxylin and eosin. From Newberne (1957), with permission of the American Journal of Veterinary Research. (a) Megalomeront in the brain, showing cellular reaction (A) and round cytomeres (B). ×233. (b) Megalomeront (A) in the lung, showing a prominent cellular reaction (bottom arrow). ×233. (c) Megalomeront in the spleen, showing islands of cytoplasmic masses (arrows). ×300. (d) Elongated megalomeront (arrow) in the cardiac muscle. ×300.
90
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91
Figure 4.10. Megalomeront of Leucocytozoon marchouxi in the spleen of an infected Pink Pigeon (Nesoenas mayeri) from Mauritius. Hematoxylin and eosin. ×1140. From Peirce et al. (1997), with permission of Veterinary Record.
Figure 4.11. Structures identified as megalomeronts of Leucocytozoon toddi in the spleen of a Eurasian Buzzard (Buteo buteo) from England. Hematoxylin and eosin. From Simpson (1991), with permission of Veterinary Record.
debris and traces of pigment in septa of air spaces. Lymphocytic infiltration of the myocardium was seen in some birds. There was moderate to marked hyperplasia and replacement of fat by proliferating cells in bone marrow. Little is known about gross and histologic lesions in doves and pigeons infected with L. marchouxi. A 7week-old Pink Pigeon squab from Mauritius that died of leucocytozoonosis had megalomeronts in various stages of development in the liver, pancreas, heart, kidney, intestine, and spleen (Peirce et al. 1997). Megalomeronts measuring up to 210 μm in diameter were very numerous in the spleen (Figure 4.10). There was also liver and renal tubular necrosis and hemorrhage in the myocardium. Splenomegaly has been reported as a gross lesion of leucocytozoonosis in a Eurasian Buzzard infected with L. toddi (Simpson 1991). Histopathological features have been described in connection with central nervous disease and blindness in Peregrine Falcons (Falco peregrinus) and Australian Kestrels (Falco cenchroides) in Australia (Raidal et al. 1999; Raidal and Jaensch
2000). Lesions included severe endarteritis, pectenitis, and meningoencephalomyelitis. The arterioles of the meninges, brain, optic papillae, optic nerve, and spinal cord had marked proliferation of endothelial cells and numerous meronts measuring from 40 to 60 μm in diameter. Meronts were also present in smaller numbers in lung, liver, heart, and intestines. Megalomeronts were not reported in the falcons and kestrels, but Simpson (1991) reported these stages in the spleen, pectoral muscle, and heart of a Eurasian Buzzard in England (Figure 4.11). No hemorrhage or myodegeneration was associated with the megalomeronts. However, these may not be megalomeronts, but actually very large meronts, since cytomeres were not present (Peirce et al. 1997).
DIAGNOSIS Diagnosis of Leucocytozoon infections (i.e., leucocytozooniasis) and diagnosis of the disease caused by Leucocytozoon spp. (i.e., leucocytozoonosis) must be
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considered separately. Infections by Leucocytozoon spp. can be diagnosed readily by examining stained thin films made from peripheral blood and finding the characteristic gametocytes. Valki¯unas (2005, pp. 213–216) has provided an excellent description of the methods of making and staining thin blood smears. By noting the morphologic and metric characteristics of the gametocytes, the host involved, and by using appropriate descriptive literature, the species can be determined. The recent monograph by Valki¯unas (2005) contains pertinent information gathered from the world literature on blood protozoans, including measurements, illustrations, and keys, and is a significant resource for identifying these organisms. Diagnosis of leucocytozoonosis should include observation of appropriate clinical signs (especially anemia), the presence of typical gross and histologic lesions, and the identification of gametocytes of Leucocytozoon spp. in the blood (Wobeser 1997). It must be remembered, however, that mortality of young birds may occur in the absence of parasitemia, and birds may be parasitemic without having leucocytozoonosis (Herman et al. 1975; Wobeser 1997). Since the early 1970s, a number of serological tests have been developed to detect antibodies against L. caulleryi in chickens. These include agar gel precipitation (Morii 1972), counter-immunoelectrophoresis (Fujisaki et al. 1980), immunofluorescence (Fujisaki et al. 1981; Isobe and Akiba 1982), an enzyme-linked immunosorbent assay (ELISA) (Isobe and Suzuki 1986, 1987a, b), immunoblot analysis (Isobe et al. 1998), and a latex agglutination test using recombinant R7 antigen (Ito and Gotanda 2005). Similar tests have not been developed for other species of Leucocytozoon. Over the past 12 years, a number of molecular genetic tests have been developed for screening birds for the presence of blood parasites. Most of the tests are polymerase chain reaction-based assays that target fragments of small unit (18S) ribosomal RNA (Feldman et al. 1995; Jarvi et al. 2002) or mitochondrial cytochrome b genes (Bensch et al. 2000; Fallon et al. 2003; Waldenstr¨om et al. 2004) to identify Plasmodium and Haemoproteus at the generic level. Several recent modifications of these assays allow Leucocytozoon to be distinguished from Plasmodium and Haemoproteus (Hellgren et al. 2004; Beadell and Fleischer 2005; Cosgrove et al. 2006), but none of these tests allow identification of leucocytozoids below the level of genus. Recent efforts to link molecular tests with traditional morphological species that are recognized as valid will lead to development of important diagnostic tools for investigating the ecology of these organisms and for recognizing cryptic species
or subspecies (Martinsen et al. 2006; Sehgal et al. 2006a).
NATURAL RESISTANCE AND IMMUNITY Information on natural or innate resistance to leucocytozoids is sparse, although some data are available on experimental infections of L . simondi in domestic ducks, American Black Ducks, and Mallards (Khan and Fallis 1968). Primary infections using ducklings and adults of all three species resulted in higher mortality in domestic ducks (white Pekins) than in American Black Ducks and Mallards given the same doses of sporozoites. Relapse parasitemias were higher in American Black Ducks than in either domestic ducks or Mallards. Overall, domestic ducks were more susceptible to L. simondi than the endemic wild species. These observations are similar to those made by several earlier investigators (Anderson et al. 1962; Trainer et al. 1962; Fallis and Bennett 1966). With the exception of L. caulleryi infections in chickens and L. simondi infections in waterfowl, very little is known about acquired immunity to leucocytozoids. Chickens that have recovered from primary infections of L. caulleryi are resistant to reinfection (Morii et al. 1986, 1989), although young chickens are less resistant than older ones (Morii and Kitaoka 1970). IgM and IgG antibodies are involved in immunity (Isobe and Suzuki 1987b) as well as cell-mediated responses (Nakata et al. 2003; Ito and Gotanda 2005). Complete protection against L. caulleryi is achieved by immunization with a recombinant R7 vaccine which is expressed against second-generation meronts (Ito and Gotanda 2005). It is not known if the immune responses in chickens infected with L. caulleryi also occur in primary infections of wild birds with other species of Leucocytozoon. In waterfowl infected with L. simondi, this does not seem to be the case. Domestic ducks exposed to a single infection and not challenged until 6 weeks later are not resistant to reinfection (Fallis et al. 1951). However, ducks exposed to primary infections of L. simondi and then repeatedly exposed to infected vectors over a 3-week period have persistently lower parasitemias than do uninfected control ducks that are exposed to infected vectors at the same time (Fallis et al. 1951). Fallis et al. (1974) referred to this as a state of premunition. However, chronically infected birds that are exposed to infection a year later develop high parasitemias and die. Immunological factors such as concentrations of white blood cells (Ortego and Espada 2007) and immunoglobulins (Tom´as et al. 2007) have been used to
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Leucocytozoonosis
PUBLIC HEALTH CONCERNS There are no public health concerns since leucocytozoids infect only birds.
100 90 80 70 % Mortality
assess disease risk and the impact of blood parasites (including Leucocytozoon spp.) on breeding populations of wild birds. Unfortunately, these studies have involved birds infected by multiple species of parasites (protozoans and arthropods), and the role of the species of Leucocytozoon in the process is not clear.
60 50 40 30 20 10 0 1960
1962
1964
1966
1968
1970
1972
Year
DOMESTIC ANIMAL HEALTH CONCERNS Six species of Leucocytozoon cause significant disease in domestic birds (Table 4.2). These include L. simondi in domestic waterfowl in the US, Canada, and Europe; L. smithi in domestic turkeys in the US and Canada; L. macleani in chickens in Southeast Asia; L. struthionis in captive ostriches in South Africa; L. schoutedeni in chickens in sub-Saharan Africa and Southeast and Southern Asia; and L. caulleryi in chickens of many Southeast and Southern Asian countries (Fallis et al. 1974; Springer 1978; Valki¯unas 2005). In an analysis of 237 published reports of mortality or pathogenicity cause by avian blood protozoans, 95% were reports on leucocytozoonosis caused by L. caulleryi (n = 90), L. simondi (n = 72), and L. smithi (n = 37) in domestic chickens, ducks, and turkeys (Bennett et al. 1993b). L. simondi and L. smithi are also found in wild waterfowl and Wild Turkeys, respectively, and these hosts may serve as reservoirs of the parasite for their domestic counterparts. Leucocytozoon smithi and L . caulleryi cause significant economic losses in poultry in certain areas of the US and Asia (Fallis et al. 1974; Morii 1992; Bennett et al. 1993b), whereas the other species are of lesser concern. Leucocytozoon smithi causes mortality of poults and adult turkeys (Stoddard et al. 1952), but also causes a decrease in production and hatchability of eggs in those that survive (Jones et al. 1972). Leucocytozoon caulleryi also causes mortality and retarded growth in young chicks and reduction of egg production in adult chickens (Morii 1992).
WILDLIFE POPULATION IMPACTS The impact of leucocytozoonosis on populations of wild birds is not clear. As mentioned earlier, three species (Leucocytozoon simondi in waterfowl, L. marchouxi in doves and pigeons, and L. toddi in raptors) are of concern, and there may be other pathogenic species that are not recognized as such at this time.
Figure 4.12. Annual mortality of Canada Geese (Branta canadensis) goslings at Seney National Wildlife Refuge in northern Michigan from 1960 to 1972. Figure was prepared with data published by Herman et al. (1975).
Leucocytozoonosis caused by L. simondi mainly occurs in the northern Holarctic and has caused localized mortality in wild waterfowl in the northern US (O’Roke 1931, 1934; Herman et al. 1975), Canada (Karstad 1965; Leighton and Riddell 1979), and Sweden (M¨orner and Wahlstr¨om 1983). The best example is the documented annual mortality of Canada Goose goslings at Seney NWR in northern Michigan (Herman et al. 1975). Mortality of goslings was noted in the refuge since its inception in 1935. The best records were from 1960 to 1972 when mortality of goslings reached over 70% every 4 years (Figure 4.12). Correlations between these cyclic fluctuations of mortality and weather, hunting, predation, and other parasites and diseases have not been found. Other reports of mortality are more anecdotal in nature. Mortality as high as 90% was observed in Mallards and American Black Ducks in some areas of Michigan in the early 1930s (O’Roke 1931, 1934). Death of a juvenile Mallard in 1963 due to leucocytozoonosis in Ontario, Canada, was described by Karstad (1965). No information was given on the number of Mallards at risk in the population from which the duck came or if there was additional mortality of Mallards in the area in question. There is another similar report of a wild duckling (species not identified) dying of leucocytozoonosis in Saskatchewan, Canada (Leighton and Riddell 1979). In southern Sweden, 10 of 62 young Mute Swans (Cygnus olor) examined at necropsy over a 10-year period were found to have died of leucocytozoonosis (M¨orner and Wahlstr¨om 1983). In a 1993 analysis of the
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literature on mortality caused by avian blood parasites, 199 reports on Leucocytozoon spp. were found and of those 79 were concerned with L. simondi. However, of those 79 reports, only 7 dealt with wild waterfowl, the other 72 were records of mortality in domestic ducks and geese (Bennett et al. 1993b). Since this analysis was published, two studies designed to test the effect of L. simondi on mortality (Shutler et al. 1996) and growth rates (Shutler et al. 1999) of Mallard and American Black Duck ducklings under conditions of natural exposure to L. simondi in Ontario, Canada, have been completed. No adverse effects were observed in either study, but the authors may have been dealing with nonpathogenic strains of the parasite. It is clear that although pathogenic strains of L. simondi cause some mortality among wild anseriforms, there is little or no evidence, with the possible exception of Canada Geese at Seney NWR, that the parasite is controlling population densities of waterfowl. In 1993–1994, mortality of two Pink Pigeon squabs from Mauritius was attributed to leucocytozoonosis caused by L. marchouxi (Peirce et al. 1997). A subsequent study of the same population of Pink Pigeons was conducted in 2003 (Bunbury et al. 2007). Pigeons less than 1 year of age had higher prevalences of infection with L. marchouxi (∼45%) compared to older birds (∼10–20%). There was no measurable effect on body condition based on measurements of body mass, tarsus length, culmen-gape, culmen-skull, culmen-feathers, and wing length. However, mortality during the 12month period after sampling was 21% for infected birds compared to 12% for uninfected birds. There was a statistically reduced likelihood of infected birds surviving to 90 days postsampling compared to uninfected birds. Although L . toddi has been shown to cause leucocytozoonosis and mortality in falcons and kestrels in Australia (Raidal et al. 1999; Raidal and Jaensch 2000), observations on the effects on raptor populations are limited and indicate that there is no impact. Long-term studies have been conducted on Eurasian Sparrowhawks in England (Ashford et al. 1990, 1991; Ashford 1994). No statistical differences were found between the survival rate of infected and uninfected nestlings and adults. Reduction of the growth rate of nestlings, increased mortality in young fledged birds, and reduction in the fecundity of infected adults were only temporary and not statistically verified because of small sample sizes. Similarly, no difference was found in the survival of infected versus uninfected nestlings and fledglings of Northern Goshawks (Accipiter gentilis) in a 1994 study of 48 nestlings from 23 nests in Wales (Toyne and Ashford 1997). On the other hand, it must be remembered that chronic infections may have harmful sublethal influ-
ences on avian populations that are not obvious or readily measured. Wild birds also commonly have concurrent infections with a wide variety of other infectious and noninfectious disease agents. Infections with Leucocytozoon might have additive effects or interact with these agents in a synergistic fashion and lead to compromised behavior or health. Leucocytozoid infections might not be the direct cause of death, but may elevate host susceptibility to predation or other disease agents or compromise host fitness for reproduction or migration. Little is known about the physiological and ecological costs of leucocytozoid infections and more research is needed. PREVENTION, TREATMENT, AND CONTROL A variety of techniques have been used to prevent and treat clinical disease caused by L. caulleryi and L. smithi in domestic poultry and have met with some success. Two vaccines have been developed against the megalomeronts of L. caulleryi to protect chickens against leucocytozoonosis. One is a formalin-killed vaccine containing second-generation megalomeronts (Morii et al. 1990), while the other is a recombinant vaccine based on a second-generation megalomeront protein (Ito and Gotanda 2004). Pyrimethamine and a combination of sufamonomethoxine and pyrimethamine are effective when administered in food (Akiba et al. 1963, 1964; Akiba 1970). Repellents such as DA14-7 have been used effectively inside chicken houses and directly on feathers to decrease biting by the vectors of L. caulleryi (Hori et al. 1964; Kitaoka et al. 1965). Among domestic turkeys, clopidol is effective in reducing numbers of gametocytes of L. smithi in the blood, but does not eliminate infections completely (Siccardi et al. 1974). Vector control in areas where domestic turkey populations are at risk from infection with L. smithi has been tested by applying Bacillus thuringiensis israelensis (Bti) to flowing streams, the source of black fly vectors. In one study, all streams within 7 km of a turkey farm were treated with a wettable formulation of Bti, leading to a reduction in transmission, reduced parasitemias, and prevention of morbidity or mortality among domestic turkeys (Horosko and Noblet 1968). Temephos, an organophosphate larvicide, was also effective against the larvae of black flies when applied by air to running streams in an area that was endemic for L. smithi. No harmful effects to nontarget stream biota were detected (Kissam et al. 1973, 1975). When possible, vector-proof screening of pens may be of use, although this is not always feasible. Prevention, treatment, and control of leucocytozoonosis in free-living populations of wild birds
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Leucocytozoonosis are difficult. While leucocytozoonosis in captive waterfowl and raptors can sometimes be treated satisfactorily with quinine derivatives (O’Roke 1934), atebrine (Coatney and West 1937), trimethoprim and sulfamethoxazole (Remple 2004), and melarsomine (Tarello 2006), currently there are no effective treatments for L. simondi or L. toddi in wild birds (Bennett 1987; Bermudez 2003). In one field study of L. majoris in a small sample of Eurasian Blue tit* (Cyanistes caeruleus), prevalence was significantly lower (20%) in females captured in nest boxes and treated with primaquine than in untreated controls (62%) (Tom´as et al. 2005).
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ACKNOWLEDGMENTS The following people are acknowledged for their kind assistance in the translation of literature: Gabriele Forrester (German), Eiji Sato (Japanese), Victor Shille (Russian), and Gediminas Valki¯unas (Russian). We also thank Robert Adlard, Jennifer Baker, Jacqui Brown, Nancy Bunbury, Frederick Leighton, Antoinette MacIntosh, Jaimie Miller, Krysten Schuler, Brittany Sears, Karl Skirnisson, and Gediminas Valki¯unas for assistance in the acquisition of published and unpublished information on leucocytozoonosis.
LITERATURE CITED MANAGEMENT IMPLICATIONS For most free-ranging avian populations, Leucocytozoon infections are probably of little concern, although our knowledge of this topic is limited. However, as previously discussed, pathogenic strains of L. simondi, L. marchouxi, L. toddi, and perhaps other species exist and therefore some avian populations may be at risk. Little can be done for most wild populations to mitigate the impact of leucocytozoonosis, especially on a large scale. In the cases of small populations, subpopulations of endangered species, or in rehabilitation settings, it may be advisable to try to manage infections. Since injured raptors with blood parasite infections (including L. toddi) have significantly longer rehabilitation times and higher mortality rates than do uninfected raptors (Olsen and Gaunt 1985), treatment of hematozoan infections may be beneficial and increase survival when birds are released back into the wild. Treating (Tom´as et al. 2005) or vaccinating (Plumb et al. 2007) a free-ranging population might be done in situations where significant numbers of individuals can be easily captured. Examples include cavity-nesting species, birds such as geese that are flightless for periods of time during molt, and social or colonial species that can be readily captured through use of baits or nets. This approach may be applicable to small populations of endangered or threatened birds. Judicious water management or the use of chemicals or biological control agents to treat streams to eliminate or reduce the black fly vectors may also be effective. If certain subpopulations of birds harbor a pathogenic strain of Leucocytozoon (L. simondi in waterfowl, for example), it might be advisable to selectively reduce or eliminate the subpopulation in an attempt to prevent its spread to other populations. The possible impact of leucocytozoonosis on avian populations, especially waterfowl, should be considered when changes in water flow patterns associated with hydroelectric dams or other types of river and stream management are planned.
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Peirce, M. A., A. G. Greenwood, and J. E. Cooper. 1983. Haematozoa of raptors and other birds from Britain, Spain and the United Arab Emirates. Avian Pathology 12:443–446. Peirce, M. A., A. G. Greenwood, and K. Swinnerton. 1997. Pathogenicity of Leucocytozoon marchouxi in the pink pigeon (Columba mayeri) in Mauritius. Veterinary Record 140:155–156. Peirce, M. A., R. D. Adlard, and R. Lederer. 2005. A new species of Leucocytozoon Berestneff, 1904 (Apicomplexa: Leucocytozoidae) from the avian family Artamidae. Systematic Parasitology 60:151–154. Pinkovsky, D. D. 1976. The Black Flies (Diptera: Simuliidae) of Florida and their Involvement in the Transmission of Leucocytozoon smithi to turkeys. Ph.D. dissertation, University of Florida, Gainesville, FL. Pinkovsky, D. D., and J. F. Butler. 1978. Black flies of Florida I. Geographic and seasonal distribution. The Florida Entomologist 61:257–267. Pinkovsky, D. D., D. J. Forrester, and J. F. Butler. 1981. Investigations on black fly vectors (Diptera: Simuliidae) of Leucocytozoon smithi (Sporozoa: Leucocytozoidae) in Florida. Journal of Medical Entomology 8:153–157. Plumb, G., L. Babiuk, J. Mazet, S. Olsen, P. P. Pastoret, C. Rupprecht, and D. Slate. 2007. Vaccination in conservation medicine. Revue Scientifique et Technique Office International des Epizooties 26:229–241. Polcyn, G. M., and A. D. Johnson. 1968. Hematozoa of the Mallard Duck Anas platyrhynchos L. in South Dakota. Bulletin of the Wildlife Disease Association 4:11. Poulin, R. 1995. Evolutionary and ecological parasitology: a changing of the guard? International Journal for Parasitology 25:861–862. Poulin, R., and W. L. Vickery. 1993. Parasite distribution and virulence: implications for parasite-mediated sexual selection. Behavioural Ecology and Sociobiology 33:429–436. Powers, L. V., M. Pokras, K. Rio, C. Viverette, and L. Goodrich. 1994. Hematology and occurrence of hemoparasites in migrating Sharp-shinned Hawks (Accipiter striatus) during fall migration. Journal of Raptor Research 28:178–185. Raidal, S. R., and S. M. Jaensch. 2000. Central nervous disease and blindness in Nankeen Kestrels (Falco cenchroides) due to a novel Leucocytozoon-like infection. Avian Pathology 29:51–56. Raidal, S. R., S. M. Jaensch, and J. Ende. 1999. Preliminary report of a parasitic infection of the brain and eyes of a peregrine falcon Falco peregrinus and Nankeen kestrels Falco cenchroides in western Australia. Emu 99:291–292.
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5 Isospora, Atoxoplasma, and Sarcocystis Ellis C. Greiner been Atoxoplasma. In fact, Levine (1982) felt the genus Atoxoplasma was valid and placed 19 species into this genus, including some species originally named in these other genera. We have still not addressed questions posed by Baker et al. (1972) in their review of avian blood coccidians: (1) “Are all atoxoplasms really Isospora?” (2) “If they are, are they one widespread species?” and (3) “If there are two different groups referred as atoxoplasms, are the two groups monospecific or not?” In this chapter, the recent synonomy of Atoxoplasma with the genus Isospora is followed, but the disease is continued to be referred to as atoxoplasmosis to distinguish it from disease caused by enteric species of Isospora. Species of Isospora are monoxenous, with single host life cycles. There are numerous species of Isospora for which their entire life cycle is restricted to the intestinal epithelium of their avian hosts. Most species of Isospora are considered host species specific. Little is known about most of them other than the morphology of the oocysts. Thus, if they do have an impact on wild avian populations, it is not recognized.
INTRODUCTION The genera Isospora, Atoxoplasma, and Sarcocystis are coccidian parasites closely related to Toxoplasma and Eimeria (Chapters 8, 9, and 11). They produce oocysts with two sporocysts containing four sporozoites each. The life cycles are different for each of these genera, and location of the endogenous or tissue stages of the parasites in the avian host determines which genus is present. Confusion can occur as to which genus is involved depending on how much of the life cycle has been detected. There are species that are pathogenic to the avian host; there are species that are not; and for most species, we do not know their impact on birds. Host specificity varies from being species specific to being able to infect a variety of birds. Most information on these genera is from examination of captive birds.
ATOXOPLASMA AND ISOSPORA Atoxoplasmosis is a disease of birds that is caused by unusual coccidian parasites in the genus Atoxoplasma. While this genus has recently been synonymized with Isospora (Barta et al. 2005), a great deal of confusion still surrounds it. Most avian coccidia with direct life cycles complete their development in the epithelial cells of the gut and occasionally in the bile ducts of the liver and the collecting tubules of the kidney. The species of Isospora that cause atoxoplasmosis have a phase that is extraintestinal in monocytes. The avian taxa that appear to be most at risk are the members of the avian families Fringillidae and Sturnidae. Atoxoplasmosis is a disease of the recticuloendothelial system as well as the intestines, and a wide range of birds are infected with these organisms. Whereas most infections do not cause disease, they are fatal in some avian hosts. The stages of this parasite that inhabit the blood have been confused with species of Lankesterella, and undoubtedly also Hepatozoon and Haemogregarina. This is not to imply that these latter genera are synonymous with Atoxoplasma, but some of the reports of these other genera may actually have
SYNONYMS Atoxoplasmosis is sometimes called “going light” as infected birds may stop eating and lose weight. When associated with disease, infections with Isospora spp. will be referred to as coccidiosis, but this condition may be caused by other genera such as Eimeria and Caryospora. Infection with Isospora spp. in the absence of clinical disease is called coccidiasis.
HISTORY The genus Atoxoplasma was defined by Garnham (1950) as those “parasites which inhabit the monocytes of birds from many parts of the world, are strictly host specific, non-pathogenic and possess a delicately granular cytoplasm not enclosed by a periplast, and a large diffuse nucleus with a tiny karyosome.” He
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Isospora, Atoxoplasma, and Sarcocystis stated that the genus was created for the “sake of convenience” to assist in naming the parasites with this morphology. Parts of this description are not correct as some species are pathogenic to their hosts, and we have no proof of the host specificity of these parasites. There are 19 species listed as valid. Levine (1982) listed these and their presumed synonyms (species of Lankesterella, Hepatozoon, and Haemogregarina) and added a species name to the parasite responsible for an epizootic in Evening Grosbeaks (Coccothraustes vespertinus) that occurred in Algonquin Park, Canada (Khan and Desser 1971). Atoxoplasma is a synonym of Lankesterella, and the red mite (Dermanyssus gallinae) is the vector of the parasite in the House Sparrow (Passer domesticus) (Lainson 1959). A closely related parasite in canaries (Serinus canaria) was transmitted by fecal contamination with oocysts that were structurally similar to species of Isospora (Box 1970). Red mites were not present on the canaries, thus demonstrating that this was truly a coccidian parasite. DISTRIBUTION Infections of Atoxoplasma have been reported from all continents except Antarctica. Many of the reports are based on examinations of blood smears of freeranging birds. Cases where the stages of the parasite that inhabit circulating monocytes have been clearly associated with fecal oocysts are usually in captive birds in aviaries or under laboratory conditions. Therefore, the true geographic distribution of atoxoplasmosis is probably best indicated by the presence of stages seen on smears of whole blood or white blood cells from the “buffy coat” from centrifuged whole blood, rather than in feces where the infections cannot be distinguished from species of Isospora that are restricted to the gut. Related to this point, atoxoplasmosis is a major problem in captive propagation of the Bali Myna (Leucopsar rothschildi; Partington et al. 1989) and is apparently caused by the only species of coccidian that has been reported from this host, Isospora rothschildi (Upton et al. 2001). While many species of avian Isospora are recognized that are restricted to the gut, the geographic distribution of these has not been determined. HOST RANGE The species of Isospora that cause atoxoplasmosis have been reported as blood parasites in at least 58 avian families (Bennett et al. 1982; Bishop and Bennett 1992). Unfortunately, it is not clear whether species of Lankesterella, Hepatozoon, and Haemogregarina were correctly differentiated from Isospora when these identifications were made. Thus, many orders of birds
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from a variety of habitats are infected with these parasites. This does not imply that disease occurs in all these hosts, but that disease is possible if conditions are correct. The level of host specificity for species of Isospora that cause atoxoplasmosis is unknown. Khan and Desser (1971) inoculated blood and tissue hom*ogenates (not oocysts) from infected Evening Grosbeaks (Coccothraustes vespertinus) into ducks and four species of passeriforms. All but the ducks became positive within 8–14 days. This suggests that there is some host specificity, but at a high taxonomic level. Box (1970) was able to transmit Isospora from House Sparrow to House Sparrow, but not to canary. In a study conducted at the San Diego Zoological Gardens (McAloose et al. 2001), a variety of passerines were found to be infected with a species of Isospora that causes atoxoplasmosis. The birds ranged in age from several days to nearly 18 years. Most appeared to be infected with one species based on polymerase chain reaction (PCR) amplification of a portion of the small subunit rRNA, but a second potential species was also present with a more restricted host distribution (Schrenzel et al. 2001). Therefore, there is evidence for some host specificity and some indication that at least some of these parasites are transmitted among unrelated host species. Approximately 140 species of enteric Isospora have been reported from a wide variety of avian families (Duszynski et al. 2000 ).
ETIOLOGY Oocyst morphology is distinct among species and these can be distinguished by differences in size in some mixed infections (Figures 5.1–5.4). There is no way
Figure 5.1. Oocysts of Isospora rothschildi from Bali Myna (Leucopsar rothschildi) (1,000×; 20 × 20 μm).
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Figure 5.2. Oocysts of Isospora canaria (larger arrow; 22 × 21 μm) and Isospora serini (smaller arrow; 16 × 16 μm) from Island Canary (Serinus canaria) (400×).
to distinguish oocysts of enteric species of Isospora from those that cause atoxoplasmosis. The mononuclear cell stages or merozoites of species of Isospora that cause atoxoplasmosis have few distinctive morphological features (Figures 5.5– 5.8). Thus, the named species of Atoxoplasma were distinguished primarily by host species.
EPIZOOTIOLOGY Coccidia have a direct life cycle and are transmitted by a fecal–oral route that involves ingestion of infective oocysts. The oocysts of species of Isospora need to undergo asexual reproduction (=sporogony) in the abiotic environment before they become infective. When ingested by a suitable avian host, the oocysts excyst in the intestines and release sporozoites. Sporozoites invade epithelial cells that line the mucosa. These then
Figure 5.3. Oocyst of Isospora sp. from Fish Crow (Corvus ossifragus) (1,000×; 20 μm).
Figure 5.4. Oocyst of Isospora sp. from Red-billed Leiothrix (Leiothrix lutea) (1,000×; 26 × 21 μm).
undergo asexual reproduction (=merogony) and produce progeny called merozoites. These escape and kill their host cells and invade other host cells where they are genetically programmed to undergo a set number of additional generations of merogony. This process greatly increases the number of parasites in the host. Merozoites from the last generation of merogony invade epithelial cells and initiate the sexual phase of the life cycle (=gametogony). Gametes are produced that fuse to form zygotes. An oocyst wall forms around the zygote and unsporulated oocysts are released, killing their host cells in the process. Oocysts are shed in the feces and undergo sporogony within a few days to become infectious to the next avian host. Those species of Isospora that cause atoxoplasmosis undergo early merogony in mononuclear phagocytes in the gut mucosa. Some of these infected monocytes leave the gut and are found in the circulation, but final stages of
Figure 5.5. Merozoite in monocyte of Bali Myna (Leucopsar rothschildi).
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Figure 5.6. Merozoite in monocytes in Superb Starling (Lamprotornis superbus).
Figure 5.8. Merozoites in monocytes in Golden-crested Myna (Ampeliceps coronatus).
merogony and gametogony occur in intestinal epithelial cells like most enteric coccidia (Box 1977). Regardless of whether infections develop only in epithelial cells of the intestines or also in monocytes, oocysts are the infective stage. Coccidiasis caused by the enteric species of Isospora develops only if hosts have no previous exposure to the parasites and if the dose of oocysts is sufficiently high. Most infections do not cause disease. Normally, when coccidia cause disease there are factors that allow a large buildup of infectious oocysts that can be acquired by a susceptible host within a short period of time. In some birds, the number of oocysts shed by adults may increase during nesting. Transmission to nestlings may occur when these are shed during egg laying, brooding, or while the parents are feeding the young in the nest. The type of nest construction might also be important for collection of and buildup of oocysts.
Prevalence of atoxoplasmosis in free-ranging birds is poorly understood. In a 4-year study in Ontario, Khan and Desser (1971) estimated that the annual prevalence in Evening Grosbeaks ranged from 29 to 68%. Prevalence in four species of birds from Hawaii ranged from 0.1% in Japanese White-eyes (Zosterops japonicus) to 2.9% in House Finches (Carpodacus mexicanus), to 8.6% in House Sparrows, and to 17.4% in Nutmeg Mannikins (Lonchura punctulata) (van Riper et al. 1987). Prevalence was 100% in 90 House Sparrows and Eurasian Tree Sparrows (Passer montanus) in Poland (Kruszewicz 1991). Ball et al. (1998) detected oocysts that they were able to correlate with monocyte infections in the feces of 136 of 922 (14.8%) European Greenfinches (Carduelis chloris) in Great Britain. Atoxoplasmosis was the primary cause of death in the loss of 95 of 98 Black Siskins (Carduelis atrata) when captive birds were imported from South America to Italy (Giacomo et al. 1997). PCR has recently been used to identify infections associated with atoxoplasmosis in feces, blood, and tissues of captive birds. Tissues from 19 of 32 dead tanagers, representing 15 species, were positive for Isospora by PCR in a zoo in the northern United States based on amplification of a fragment of 18s rRNA (Adkesson et al. 2005). These included Purple Honeycreeper (Cyanerpes caeruleus), Red-legged Honeycreeper (Cyanerpes cyaneus), Blue Dacnis (Dacnis cayana), Violaceous Euphonia (Euphonia violacea), Hawaii Amakihi (Hemignathus virens), Apapane (Himatione sanguinea), Silverbeaked Tanager (Ramphocelus carbo), Passerini’s Tangara (Ramphocelus passerinii), White-lined Tanager (Tachyphonus rufus), Burnished-buff Tanager (Tangara cayana), Paradise Tanager (Tangara chilensis), Turquoise Tanager (Tangara mexicana), Green-andgold Tanager (Tangara schrankii), Blue-gray Tanager (Thraupis episcopus), and Iiwi (Vestiaria coccinea).
Figure 5.7. Merozoite in monocyte in Wattled Starling (Creatophora cinerea).
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Prevalence of infection has not been reported for other host species infected with Isospora spp. that cause atoxoplasmosis. These include American Goldfinches (Carduelis tristis) in Canada (Middleton and Julian 1983), Nashville Warblers (Vermivora ruficapilla) in Michigan, USA (Swayne et al. 1991), Eurasian Bullfinch (Pyrrhula pyrrhula) in Great Britain (McNamee et al. 1995), and Northern Cardinal (Cardinalis cardinalis) in Arizona, USA (Baker et al. 1996). CLINICAL SIGNS Clinical signs associated with atoxoplasmosis include loss of appetite, weight loss, diarrhea, lethargy, and ruffled feathers (Norton et al. 1993). Clinical signs are usually not evident in birds that pass oocysts of enteric species of Isospora, but clinical signs might mimic those of atoxoplasmosis when disease is present. PATHOGENESIS AND PATHOLOGY Much of the available information on pathology of atoxoplasmosis is from captive birds because they are readily available for observation before death and can be necropsied soon afterward. It is only in the most fortuitous occasions that free-ranging birds are observed in the early stages of disease and even more rarely that one can observe the disease as it progresses. Gross lesions associated with atoxoplasmosis include enlargement of the liver and spleen and presence of tiny white foci of necrosis through the parenchyma of both organs and sometimes on the surface of the heart. The pancreas may be hemorrhagic and edematous, the intestines will be filled with fluid, and the air sacs and the pericardium may be filled with a yellowish clear fluid. Enlargement of the liver and spleen is, in part, caused by an infiltrate of mononuclear cells including macrophages, lymphocytes, and plasma cells (Partington et al. 1989; Norton et al. 1993; S. Terrell, personal communication). Host cells are eventually killed by developing parasites in infections with Isospora, but like many coccidia, there is a balance between loss of these cells and their replacement with new ones. When rate of loss is equivalent to rate of replacement, the parasites may be in harmony with their hosts with no evidence of clinical disease. Coccidiosis can develop in na¨ıve individuals when large numbers of oocysts are ingested within a short period of time. DIAGNOSIS Based on personal observations of atoxoplasmosis in a large number of captive Bali Mynas, there is rarely any correlation between presence of fecal oocysts and
presence of mononuclear merozoites in the same bird at the same time. Standard fecal flotation using centrifugation with Sheather’s sugar is the best way to concentrate and cleanse the oocysts for visualization and measurement. This flotation medium is more viscous than saturated salt solutions and makes it possible to use oil immersion lenses and higher magnification to observe finer points of oocyst morphology without having the oocysts move out of the field of view when adjusting focus. Key morphological features of oocysts that are used to distinguish species include length and width of the oocyst and sporocyst, their shapes, presence or absence of a nipple-like Stieda body at one end of the sporocyst, presence of granular material in both the oocyst and the sporocyst called a residuum, presence of a thinning at one end of the oocyst called the micropyle, presence of a cap over the micropyle, presence of large refractile granules in the oocysts called polar bodies, and number of layers in the oocyst wall. Oocysts of Isospora need to be aerated for about a week to allow sporulation to occur and to produce infectious oocysts. Fully sporulated oocysts have all the morphological features that are used to identify species. Oocysts can be sporulated in a 3% potassium dichromate solution or a 1% sulfuric acid solution by gently bubbling air through the solution. This process will prevent bacterial overgrowth and reduce adverse odors. Mature oocysts of Isospora will contain two sporocysts and each will contain four sporozoites (Figures 5.1–5.4). To confirm a diagnosis of atoxoplasmosis, both fecal oocysts and merozoites in the monocytes must be present. This may require collection of multiple blood samples over the course of 5–7 days to find merozoites in the monocytes. Mononuclear merozoites are most easily found in impression smears of spleen or liver, but smears prepared from the buffy coat will provide an adequate sample of monocytes when the host is still alive. This procedure will provide a higher number of monocytes for review than in a normal blood smear and thus enhance the potential of detecting the parasite. These are prepared by centrifuging whole blood in a microhematocrit centrifuge tube, breaking the tube at the top of the erythrocyte pellet where white blood cells are concentrated into a thin white band, smearing the top portion of the pellet containing the buffy coat onto a glass microscope slide, and rapidly drying. Both smears of the buffy coat and impression smears should be fixed in absolute methanol and then stained with Giemsa or Wrights/Giemsa. In the vast majority of cases, a single merozoite will be present in the monocyte causing an indentation in the host cell nucleus (Figures 5.5–5.8). It is possible to find meronts in these monocytes as well as single parasites. If one does not find infected monocytes, then it is possible that the bird is infected with an enteric species of
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Isospora, Atoxoplasma, and Sarcocystis Isospora and thus will have only meronts and gametocytes in the gut epithelium and oocysts in the feces. Fecal collections should ideally be spread over a 5day interval and collected from the host at least three times during this period. At necropsy, gross signs of atoxoplasmosis include hepatomegaly, splenomegaly, white pinpoint foci on the surface and cut surfaces of spleen, liver, and occasionally heart, presence of an edematous pancreas, and presence of fluid in the intestinal tract. It is useful to collect the normal range of tissues for histopathology and to make impression smears of at least the spleen and liver. IMMUNITY No studies have been conducted on the immunology of avian species of Isospora. Atoxoplasmosis is usually a disease of young birds, particularly fledglings, and adults are usually not affected.
SARCOCYSTIS In contrast to Isospora, species of Sarcocystis have indirect life cycles. Intermediate hosts containing the large, distinctive intramuscular tissue cysts need to be eaten by the definitive host to transmit the infection. The intermediate host is, in turn, infected by the fecal– oral route by ingesting sporocysts that are excreted by the definitive host. When disease occurs in the intermediate host, it is caused by meronts during early phases of infection and not the large obvious Sarcocystis. SYNONYMS Infections with Sarcocystis spp. are called sarcocystosis. HOST RANGE AND DISTRIBUTION Few studies of host range have been conducted for the species of Sarcocystis that infect birds. Twelve species of this genus use birds as definitive hosts and twentytwo species use birds as intermediate hosts. Two additional species can use birds for both definitive and intermediate hosts (Table 5.1; Odening 1998). The most detailed studies of host range in avian hosts have been conducted with Sarcocystis falcatula. When sarcocysts from Brown-headed Cowbirds (Molothrus ater) and Boat-tailed Grackles (Quiscalus major) are fed to opossums, the opossums produce oocysts and sporocysts that are infective to canaries and House Sparrows (Box and Duszynski 1978). Budgerigars (Melopsittacus undulatus), Zebra Finches (Taeniopygia guttata), and Rock Pigeons (Columba livia) are susceptible to infection with sporocysts from opossums, but domestic chickens (Gallus gallus) and
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Helmeted Guineafowl (Numida meleagris) are not (Box and Smith 1982). Thus, four different orders of birds can serve as hosts for this species. Species of Sarcocystis that infect birds are widely distributed, with reports from all continents with the exception of Antarctica (Table 5.1). ETIOLOGY Like their relatives in the genus Isospora, species of Sarcocystis are coccidian parasites and have both intestinal and extraintestinal tissue stages and produce infective oocysts that are passed in the feces of their definitive hosts. Meronts of S. falcatula occur in endothelial cells of the intermediate host and may be visualized with immunoperoxidase staining or hematoxylin and eosin (Figure 5.9). Sarcocysts are large spindle-shaped structures that occur in the muscle fibers of the intermediate host. They are round when seen in cross section and narrow and elongate when viewed in longitudinal sections (Figure 5.10). Sporocysts (Figure 5.11) typically rupture from their thin-walled oocysts when shed in the feces of the definitive host and are fully sporulated and infectious to the intermediate host as soon as they reach the external environment. EPIZOOTIOLOGY The life cycle of Sarcocystis is similar to Isospora, but requires two hosts. When intermediate hosts consume infective sporocysts, sporozoites are liberated in the intestine. These move through the intestinal wall into the arterial endothelium of the mesenteric lymph nodes where the initial round of asexual reproduction (merogony) occurs (Dubey et al. 1989). Subsequent cycles of merogony occur in endothelial cells in other organs and it is during this phase of development that most pathology occurs. At completion of merogony, merozoites are released that enter muscle cells and develop into septate cysts, which will undergo another form of asexual reproduction called endopolygony. This process leads to the formation of countless infective forms called bradyzoites. Mature sarcocysts will persist in muscle tissue until consumed by a carnivore. After ingestion by a suitable definitive host, bradyzoites will be released in the intestine, enter gut epithelial cells, and undergo sexual reproduction or gametogony. This process forms gametocytes, which mature to produce male (microgametes) and female (macrogametes) that unite to form fertile zygotes. A thin oocyst wall then forms around each zygote and these undergo a final round of asexual reproduction (sporogony) to generate two sporocysts that each contains four sporozoites. Unlike the oocysts of enteric species of Isospora that undergo
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Table 5.1. Species of Sarcocystis that parasitize birds. Sarcocystis spp.
Intermediate host
Definitive host
Geographic region
Sarcocystis accipitris Sarcocystis alectoributeonis Sarcocystis alectorivulpes Sarcocystis ammodrami Sarcocystis aramidis Sarcocystis buteonis
Island Canary (Serinus canaria)
Northern Goshawk (Accipiter gentilis) Eurasian Buzzard (Buteo buteo) Mammal (Canidae)
Europe
Kazakhstan
Unknown
South America
Unknown
South America
Eurasian Buzzard, Red-tailed Hawk (Buteo jamaicensis) Eurasian Kestrel (Falco tinnunculus) Black Kite (Milvus migrans) Eurasian Buzzard
Holarctic
Kazakhstan
Unknown
Africa Europe
Passeriformes Cuculiformes Columbiformes Psittaciformes Little Egret (Egretta garzetta)
Northern Long-eared Owl (Asio otus), Barn Owl (Tyto alba) Northern Saw-whet Owl (Aegolius acadicus) Opossums (Didelphis spp.) Unknown
South Africa
Mammal (Cricetidae)
Eurasian Buzzard
Western Palearctic
Domestic chicken (Gallus gallus)
Unknown
Europe
Blue-black Grassquit (Volatinia jacarina) Laughing Dove (Streptopelia senegalensis) Siamese Fireback (Lophura diardi), Common Hill Myna (Gracula religiosa) Eurasian Buzzard
Unknown
South America
Unknown
South Africa
Unknown
Southeast Asia Central America
Unknown
Europe
Unknown
America
Mammal (Canidae)
Unknown
Unknown
Africa
Unknown
Unknown†
Chukar (Alectoris chukar) Chukar
Sarcocystis cernae
Grassland Sparrow (Ammodramus humeralis) Slaty-breasted Wood-Rail (Aramides saracura) Mammal (Cricetidae, Muridae, Chinchillidae, Erethizotidae, Leporidae) Mammal (Cricetidae)
Sarcocystis cheeli
Unknown
Sarcocystis citellibuteonis Sarcocystis colii
Mammal (Sciuridae)
Sarcocystis dispersa Sarcocystis espinosai Sarcocystis falcatula Sarcocystis garzettae Sarcocystis glareoli Sarcocystis horvathi Sarcocystis jacarinae* Sarcocystis kaiserae Sarcocystis kirmsei Sarcocystis nontenella Sarcocystis oliverioi* Sarcocystis peckai Sarcocystis phoeniconaii Sarcocystis ramphastosi
Red-faced Mouse Bird (Colius erythromelon) Mammal (Muridae)
Mammal (Cricetidae)
Green-rumped Parrotlet (Forpus passerinus) Ring-necked Pheasant (Phasianus colchicus) Lesser Flamingo (Phoenicopterus minor) Keel-billed Toucan (Ramphastos sulfuratus)
Kazakhstan
Europe India
North America America
(continues)
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Isospora, Atoxoplasma, and Sarcocystis Table 5.1. (Continued ) Sarcocystis spp.
Intermediate host
Definitive host
Geographic region
Sarcocystis rauschorum Sarcocystis rileyi
Mammal (Cricetidae)
Snowy Owl (Bubo scandiacus) Mammal (Mustelidae, Marsupialia)
North America
Tawny Owl (Strix aluco) Tawny Owl
Europe Europe
Unknown
North America
Sarcocystis scotti Sarcocystis sebeki Sarcocystis setophagae* Sarcocystis spaldingae
Sarcocystis sulfuratusi Sarcocystis wenzeli
Dabbling ducks (Anas spp.), Goldeneye (Bucephala spp.), American Wigeon (Anas americana), White-winged Scoter (Melanitta fusca), Blue-winged Teal (Anas discors), Northern Shoveler (Anas clypeata) Mammal (Muridae) Mammal (Muridae, Leporidae, Mustelidae) American Redstart (Setophaga ruticilla) Great Blue Heron (Ardea herodias), Striated Heron (Butorides striata), Great Egret (Ardea alba), Little Blue Heron (Egretta caerulea), White Ibis (Eudocimus albus), Yellow-crowned Night-Heron (Nyctanassa violacea) Keel-billed Toucan
North America
Unknown
North America
Unknown
Unknown
Domestic Chicken
Mammal (Canidae)
Europe
Note: Information is summarized from Odening (1998) and Dubey et al. (2004). * Considered by Odening (1998) to be possible synonyms of Sarcocystis falcatula. † Described from captive bird; natural range unknown. sporogony outside of the host, these complete sporogony prior to passing out of the gastrointestinal tract and are immediately infectious to intermediate hosts. A classic problem in the distribution of avian sarcocystosis in North America developed when old world psittacines were brought into contact with Virginia
Figure 5.9. Elongate meront of Sarcocystis falcatula (arrows) in endothelial cells of a pulmonary capillary from a Lory (Lorinus sp.). Hematoxylin and eosin (1000×).
opossums (Didelphis virginianus) and exposed to infection with thesporocysts of S. falcatula—a parasite of Brown-headed Cowbirds. Infections are often fatal
Figure 5.10. Sarcocysts of Sarcocystis falcatula in Brown-headed Cowbird (Molothrus ater). Muscles are cut in cross and longitudinal sections. Hematoxylin and eosin (100×; maximum width of sarcocysts is 92 μm).
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Figure 5.11. Sporocysts of Sarcocystis falcatula from feces of Virginia Opossum (Didelphis virginiana) (1000×; 11 × 7 μm). (Hillyer et al. 1991) and may influence the success of captive propagation programs aimed at reintroduction of rare parrots into endemic sites. Infections in captivity result when opossums roam around aviaries and deposit their feces where food or water can be contaminated (Hillyer et al. 1991). Exposure may also occur when co*ckroaches feed on opossum feces and then enter cages and are eaten by the birds. The sporocysts are not destroyed in the gut of the roach and remain viable and infectious (Clubb and Frenkel 1992). This might occur in the wild where introduced species of parrots have become established in regions where this parasite is cycling. No one seems to know what happened to the massive population of budgerigars that became established in southeastern Florida, but the role that S. falcatula may have played in their disappearance is not known. One introduced psittacine pest species that is from the Neotropics, the Monk Parakeet (Myiopsitta monachus), is not susceptible to this parasite (E. Greiner, personal observations). CLINICAL SIGNS Many species of birds contain sarcocysts that often reach very high intensities, yet health problems are usually not apparent. Sarcocystosis in birds is best known from work on S. falcatula in old world psittacines. Susceptible parrots may be fine and active one day and dead the next without showing any outward signs of infection. Some will become anorexic, weak, have difficulty in breathing, have blood in the oral cavity and trachea, and exhibit neurologic signs (Hillyer et al. 1991). PATHOGENESIS AND PATHOLOGY As indicated earlier, developing meronts rather than sarcocysts are the primary cause of pathology in inter-
mediate hosts. Sarcocystis falcatula is one of the beststudied species in birds, and detailed knowledge about pulmonary and hepatic pathology is derived from experimental infections using budgerigars. Early meronts develop in pulmonary endothelial cells and cause these cells to enlarge and obstruct the capillaries. Inflammatory infiltrates develop in response to tissue damage, and blockage of the blood vessels can lead to interstitial, air space edema, and pulmonary congestion (Smith et al. 1987). Most meronts develop in the lungs with a much smaller proportion in the liver and kidney. The earliest merogony begins about 12 h after ingestion of sporocysts and infection of the lamina propria of the intestines. Meronts are found by day 2 in the lungs and liver. The first sarcocysts develop in cardiac muscle by day 7, but these cysts degenerate by 30–40 days postinfection. Sarcocysts in skeletal muscle appear by day 8 in the pectoral and major leg muscles, but those in the pectoral muscles degenerate (Smith et al. 1989). This does not always happen as massive numbers of sarcocysts can sometimes be seen grossly at necropsy of psittacines (Bolon et al. 1989). While most deaths are attributed to pneumonitis, inflammation of the liver, muscles, kidneys, and brain may also be evident (Smith et al. 1989). Sarcocystosis in captive old world psittacines can be very acute. The most frequent sign of infection is pulmonary edema that may sometimes be associated with hemorrhage (Hillyer et al. 1991). Parrots may also develop an enlarged spleen and liver with marked inflammation in a number of internal organs (Page et al. 1992). By contrast, infections with Sarcocystis are usually nonpathogenic in other avian hosts.
DIAGNOSIS Sporulated oocysts of species of Sarcocystis usually rupture in the feces when they pass out of host, releasing infective sporocysts into the environment (Figure 5.11). The sporocysts may be concentrated from the feces of definitive hosts by the same fecal flotation methods with saturated salt or sugar solutions that are used to recover oocysts from other genera. In the intermediate host, one must inspect the muscles of freshly dead birds for grossly visible, elongate white, thread-like sarcocysts or find the sarcocysts in histologic sections (Figure 5.10). Meronts are less obvious and will require careful scrutiny of sections of host tissues such as the lung (Figure 5.9). Since nothing is shed from the intermediate hosts, necropsy or biopsy is necessary to detect infections with Sarcocystis in these hosts.
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Isospora, Atoxoplasma, and Sarcocystis IMMUNITY No studies have been done on immunity to Sarcocystis in birds. Species of Sarcocystis may cause problems in birds of any age, for example S. falcatula, but these are usually abnormal hosts for the parasite.
ATOXOPLASMA, ISOSPORA, AND SARCOCYSTIS PUBLIC HEALTH CONCERNS Species of Isospora will infect only birds. The avian species of Sarcocystis may use other vertebrate classes as either intermediate or definitive hosts. None are known to be infectious to humans. DOMESTIC ANIMAL HEALTH CONCERNS There are no records of infections of Isospora in poultry. There are species of Isospora that infect domestic mammals, but none of them infect birds. There are no known cases where dogs or cats serve as definitive hosts of species of Sarcocystis that infect birds. Captive wild birds may be at risk from other vertebrates since the complete life cycle and definitive hosts of many species Sarcocystis are unknown (Table 5.1). WILDLIFE POPULATION IMPACTS Khan and Desser (1971) reported the first outbreak of atoxoplasmosis (=Lankesterella) in wild populations, but there have been relatively few reports of atoxoplasmosis in free-ranging birds since then. These include reports of atoxoplasmosis in American Goldfinches and Nashville Warblers that were brought into captivity (Middleton and Julian 1983; Swayne et al. 1991) and a case of pneumonia in a Northern Cardinal that was attributed to this disease (Baker et al. 1996). Most reports of atoxoplasmosis originate from aviaries where this disease may have significant impacts on efforts to use captive propagation to reestablish threatened or endangered species in the wild. This disease has had a substantial impact on captive propagation of the Bali Myna (Partington et al. 1989; Norton et al. 1993), and possibly the largest influence these genera will have on wildlife populations will be their effects on captive populations of threatened or endangered species. TREATMENT AND CONTROL Species of Isospora are normally not treated in freeranging birds, but in captive situations both hygiene and anticoccidial drugs have been used successfully to control atoxoplasmosis. Compounds that are effective in drinking water include sulfachlorpyrazine
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(ESB3) and toltrazuril (Baycox). Sulfachlorpyridazine (Vetisulid) may be substituted for sulfachlorpyrazine, but a vitamin B12 supplement should be used during the treatment (Norton et al. 2002). The suggested treatment for infections with enteric species of Isospora is use of trimethoprimsulphamethoxazole (Clyde and Patton 1996). Combinations of trimethoprim-sulphamethoxazole and pyrimethamine or trimethoprim-sulfadiazine have been used successfully to control Sarcocystis (Page et al. 1992; Clyde and Patton 1996). Most problems with these genera will be in captivity rather than in the wild. Some of the main problems with atoxoplasmosis will be with captive propagation where the intention is to release individuals back into the wild such as is documented for the Species Survival Plan for the Bali Myna (Norton et al. 2002). Similar concerns might occur with species of Sarcocystis. In captivity, a means of reducing contact between psittacines and opossums is to use a hot wire about 10 cm above the ground and about 1 m from the aviary barrier (Susan Clubb, personal communication). ACKNOWLEDGMENTS I thank the many keepers and veterinarians in zoos and aviaries across the country for getting me involved with atoxoplasmosis and sarcocystosis. In particular, I thank Dr Terry Norton for his patience and continued prodding and Dr Carter Atkinson for his energetic, professional, and highly useful editorial assistance with this chapter. LITERATURE CITED Adkesson, M. J., J. M. Zdziarski, and S. E. Little. 2005. Atoxoplasmosis in tanagers. Journal of Zoo and Wildlife Medicine 36:265–272. Baker, D. G., S. A. Speer, A. Yamaguchi, S. M. Griffey, and J. P. Dubey. 1996. An unusual coccidian parasite causing pneumonia in a northern cardinal (Carduelis carduelis). Journal of Wildlife Disease 32:130–132. Baker, J. R., G. F. Bennett, G. W. Clark, and M. Laird. 1972. Avian blood coccidians. Advances in Parasitology 10:1–30. Ball, S. J., M. A. Brown, P. Daszak, and R. M. Pittilo. 1998. Atoxoplasma (Apicomplexa: Eimeriorina: Atoxoplasmatidae) in the greenfinch (Carduelis chloris). Journal of Parasitology 84:813–817. Barta, J. R., M. D. Schrenzel, R. Carreno, and B. A. Rideout. 2005. The genus Atoxoplasma (Garnham 1950) as a junior objective synonym of the genus Isospora (Schneider 1881) species infecting birds and resurrection of Cystoisospora (Frenkel 1977) as the correct genus for Isospora species infecting mammals. Journal of Parasitology 91:726–727.
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Bennett, G. F., M. Whiteway, and C. Woodworth-Lynas. 1982. Host–parasite catalogue of the avian haematozoa. Occasional Papers in Biology, Memorial University of Newfoundland 5:243. Bishop, M. A., and G. F. Bennett. 1992. Host–parasite catalogue of the avian haematozoa, Supplement 1, and bibliography of the avian blood-inhabiting haematozoa, Supplement 2. Occasional Papers in Biology, Memorial University of Newfoundland 15:1–244. Bolon, B., E. C. Greiner, and M. Calderwood Mays. 1989. Microscopic features of Sarcocystis falcatula in skeletal muscle from a Patogonian Conure. Veterinary Pathology 26:282–284. Box, E. D. 1970. Atoxoplasma associated with an isosporan oocyst in canaries. Journal of Protozoology 17:391–396. Box, E. D. 1977. Life cycles of two Isospora species in the Canary, Serinus canarius Linnaeus. Journal of Protozoology 24:57–67. Box, E. D., and D. W. Duszynski. 1978. Experimental transmission of Sarcocystis from icterid birds to sparrows and canaries by sporocysts from the opossum. Journal of Parasitology 64:682–688. Box, E. D., and J. H. Smith. 1982. The intermediate host spectrum in a Sarcocystis species of birds. Journal of Parasitology 68:668–673. Clubb, S. L., and J. K. Frenkel. 1992. Sarcocystis falcatula of opossums: transmission by co*ckroaches with fatal pulmonary disease in psittacine birds. Journal of Parasitology 78:116–124. Clyde, V. L., and S. Patton. 1996. Diagnosis, treatment, and control of common parasites in companion and aviary birds. Seminars in Avian and Exotic Pet Medicine 5:75–84. Dubey, J. P., E. Lane, and E. Van Wilpe. 2004. Sarcocystis ramphastosi sp. nov. and Sarcocystis sulfuratusi sp. nov. are described from natural infected keel-billed toucan (Ramphastos sulfuratus). Acta Parasitologia 49:93–101. Dubey, J. P., C. E. Speer, and R. Fayer. 1989. Sarcocystosis of Animal and Man. CRC Press, Boca Raton, FL. Duszynski, D. W., S. J. Upton, and L. Couch. 2000. The Coccidia of the World. Department of Biology, University of New Mexico. Available at http://biology.unm.edu/biology/coccidia/home.html. Garnham, P. C. C. 1950. Blood parasites of East African vertebrates, with a brief description of exo-erythrocytic schizogony in Plasmodium pitmani. Parasitology 40:328–329. Giacomo, R., P. Stefania, T. Ennio, V. C. Giorgina, B. Giovanni, and R. Giacomo. 1997. Mortality in Black Siskins (Carduelis atrata) with systemic coccidiosis. Journal of Wildlife Diseases 33:152–157.
Hillyer, E. V., M. P. Anderson, E. C. Greiner, C. T. Atkinson, and J. K. Frenkel. 1991. An outbreak of Sarcocystis in a collection of psittacines. Journal of Zoo Wildlife Medicine 22:434–445. Khan, R. A., and S. S. Desser. 1971. Avian Lankesterella infections in Algonquin Park, Ontario. Canadian Journal of Zoology 49:1105–1110. Kruszewicz, A. 1991. Lesions in sparrows. Veterinary Record 128:167. Lainson, R. 1959. Atoxoplasma Garnham, 1950, as a synonym for Lankesterella Labbe, 1899. Its life cycle in the English sparrow (Passer domesticus, Linn). Journal of Protozoology 6:361–371. Levine, N.D. 1982. The genus Atoxoplasma (Protozoa, Apicomplexa). Journal Parasitology 68:719–723. McAloose, D., L. Keener, M. Schrenzel, and B. Rideout. 2001. Atoxoplasmosis: beyond Bali Mynahs. Proceedings of the American Association of Zoo Veterinarians, American Association of Wildlife Veterinarians, Association of Reptile Amphibian Veterinarians, and National Association of Zoo Wildlife Veterinarians, Joint Conference, pp. 64– 67. McNamee P., T. Pennycott, and S. McConnell. 1995. Clinical and pathological changes associates with Atoxoplasma in a captive bullfinch (Pyrrhula pyrrhula). Veterinary Record 136:221–222. Middleton, A. L. A., and R. J. Julian. 1983. Lymphoproliferative disease in the American goldfinch, Carduelis tristis. Journal of Wildlife Disease 19:280–285. Norton, T. M., E. C. Greiner, K. S. Latimer, and E. Dierenfeld. 1993. Medical aspects of Bali Mynahs (Leucopsar rothschildi). Proceedings of the American Association of Zoo Veterinarians, pp. 29–37. Norton, T. M., E. C. Greiner, K. Latimer, and S. E. Little. 2002. Medical protocols recommended by the US Bali Mynah SSP. Riverbanks Zoo and Garden, Columbia, SC. Available at http://www.riverbanks. org/AIG/new.htm. Odening, K. 1998. The present state of speciessystematics in Sarcocystis Lankester, 1882 (Protista, Sporozoa, Coccidia). Systematic Parasitology 41:209–233. Page, C. D., R. E. Schmidt, J. H. English, C. H. Gardiner, G. B. Hubbard, and G. C. Smith. 1992. Antemortem diagnosis and treatment of sarcocystosis in two species of psittacines. Journal of Zoo Wildlife Medicine 23:77–85. Partington, C. J., C. H. Gardiner, D. Fritz, L. G. Phillips, Jr., and R. J. Montali. 1989. Atoxoplasmosis in Bali mynahs (Leucopsar rothschildi). Journal of Zoo Wildlife Medicine 20:328–335. Schrenzel, M, L. Keener, D. McAloose, I. Stalis, R. S. Papendick, R. Klieforth, G. Maalouf, and
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Isospora, Atoxoplasma, and Sarcocystis B. Rideout. 2001. Diagnosis and molecular characterization of Atoxoplasma in passerine birds. Proceedings of the American Association of Zoo Veterinarians, American Association of Wildlife Veterinarians, Association of Reptile Amphibian Veterinarians, and National Association of Zoo Wildlife Veterinarians, Joint Conference, pp. 214. Smith, J. H., J. L. Meier, P. J. G. Neill, and E. D. Box. 1987. Pathogenesis of Sarcocystis falcatula in the Budgerigar. II. Pulmonary pathology. Laboratory Investigation 56:72–84. Smith, J. H., P. J. G. Neill, and E. D. Box. 1989. Pathogenesis of Sarcocystis falcatula (Apicomplexa: Sarocystidae) in the Budgerigar (Melopsittacus undulatus). III. Pathologic quantitative parasitologic
119
analysis of extrapulmonary disease. Journal of Parasitology 75:270–287. Swayne, D. E., D. Getzy, R. D. Siemons, C. Bocetti, and L. Kramer. 1991. Coccidiosis as a cause of transmural lymphocytic enteritis and mortality in captive Nashville warblers (Vermivora ruficapilla). Journal of Wildlife Disease 27:615–620. Upton, S. J., S. C. Wilson, T. M. Norton, and E. C. Greiner. 2001. A new species of Isospora (Apicomplexa: Eimeriidae) from the Bali (Rothschild’s) Mynah (Leucopsar rothschildi) (Passeriformes: Sturnidae). Systematic Parasitology 48:47–53. Van Riper, C., III, S. Van Riper, and M. Laird. 1987. Discovery of Atoxoplasma in Hawaii. Journal of Parasitology 73:1071–1073.
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6 Trichom*onosis Donald J. Forrester and Garry W. Foster 1500s which contained a description of the disease (Turbervile 1575). It was not until 300 years later that T. gallinae was identified as the etiologic agent (Rivolta 1878). Rivolta described the organism from the upper digestive tract and liver of a Rock Pigeon (Columba livia). Following Rivolta’s work, there was a period of approximately 100 years when numerous papers were published concerning the nomenclature for this parasite as well as its morphology and its host and geographic distribution throughout the world (Stabler 1954). At that time, there was also considerable research conducted on the cultivation and nutritional requirements of the organism, particularly during the 1930s and 1940s by R. Cailleau, A. Bos, and others. Most of this work was done in Europe and North America. These studies on the cultivation of the parasite were foundational to research that followed on improved methods of diagnosis, treatment, and control and led to studies on virulence, pathogenicity, and immunity in the 1950s, 1960s, and 1970s. The contributions of R. M. Stabler, R. M. Kocan, B. M. Honigberg, and their colleagues were especially noteworthy during that period. From 1980 to the 2000s there were a number of significant findings related to immunity, pathology, ultrastructural morphology of the agent, and the use of molecular techniques to improve diagnosis and understand the phylogeny of trichom*onads in general, including T. gallinae. Additional details on the history of T. gallinae and trichom*onosis can be found in the reviews by Stabler (1954), Kocan and Herman (1971), BonDurant and Honigberg (1994), Knispel (2005), and Bunbury (2006).
INTRODUCTION Trichom*onosis is a protozoan disease caused by the flagellate Trichom*onas gallinae (Rivolta 1878). It is primarily a disease of the upper digestive and respiratory tracts of columbiforms, raptors, psittaciforms, and a few other birds. Effects vary from subclinical infections (i.e., trichom*oniasis) to significant disease (i.e., trichom*onosis) that leads to severe organ necrosis, caseation, tissue invasion, and death (Kocan and Herman 1971). Although many instances of this disease relate to individual birds or siblings in the nest, epizootics of sizeable proportions are known, especially among free-ranging columbiforms (Haugen and Keeler 1952). There is a sizeable body of literature on trichom*onosis, only a part of which will be discussed here. Several general reviews have been published (Florent 1938; Stabler and Herman 1951; Stabler 1954; Kocan and Herman 1971; Conti 1993; BonDurant and Honigberg 1994; and Cole 1999). Pokras et al. (1993) summarized publications on the disease in owls. SYNONYMS Trichom*oniasis, canker, roup (columbiforms, psittaciforms, and other birds), frounce (raptors). (Note: The authors recognize that the term trichom*oniasis has been used commonly in the historical literature to refer to the disease caused by T. gallinae as well as infection without apparent disease, but we are following the convention of referring to the disease as trichom*onosis and the infection without clinical signs as trichom*oniasis. See Kassai (2006) for a discussion of this practice.)
DISTRIBUTION Trichom*onas gallinae is cosmopolitan and has been reported from every major land mass except Antarctica, Greenland, and the northern parts of North America, Europe, and Asia (Figure 6.1). Its distribution is correlated closely with that of the Rock Pigeon, one of its most important hosts.
HISTORY Trichom*onosis is probably the oldest known wildlife disease for which there are written records. Many years before the cause of the disease was discovered, there were reports of the lesions in birds of prey used for falconry. For example, a book was published in the
120 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Trichom*onosis
121
Figure 6.1. Distribution of Trichom*onas gallinae throughout the world. Solid circles indicate areas where infections were reported either in captive or free-ranging wild birds. This figure is based on information from the references listed in Tables 6.1 and 6.2 and locality records from Babes and Puscariu (1890), Ratz (1913), Volkmar (1930), Bos (1932), Callender and Simmons (1937), Hees (1938), Bushnell and Twiehaus (1940) Russell (1951), Ahmed et al. (1970), Ballouh and Eisa (1980), Minowa et al. (1982), Zhang et al. (1982), Garner and Sturtevant (1992), Rosskopf and Woerpel (1996), and Silvanose et al. (1998). HOST RANGE Trichom*onas gallinae is common especially in columbiforms, and one of these, the Rock Pigeon, is considered to be its primary host (Stabler 1954). Infections have been reported in Rock Pigeons from 31 countries representing every continent except Antarctica (Table 6.1). In addition to the Rock Pigeon, infections are known to occur in 18 other species of columbiforms (Table 6.1), 26 species of falconiforms (Table 6.2), and 9 species of strigiforms (Table 6.2). Other captive and experimental hosts include psittaciforms, passeriforms, galliforms, gruiforms, and anseriforms. There is also one report of trichom*onosis (lesions and trichom*onad identification) from a charadriiform (an unidentified species of gull) from the Shetland Isles off the coast of Scotland (Hees 1938). Although there are several reports of successful experimental infections, some accompanied by lesions, in several species
of passeriforms (Callender and Simmons 1937; Levine et al. 1941; Stabler 1953), natural infections in these birds are not common. However, in 2002 there was an outbreak of trichom*onosis in Kentucky that involved approximately 200 wild House Finches (Carpodacus mexicanus) and House Sparrows (Passer domesticus) (NWHC 2002). In addition, widespread mortality attributed to a trichom*onosis-like disease was reported in several areas in England during 2005 and 2006 and involved large numbers of European Greenfinches (Carduelis chloris) and Chaffinches (Fringilla coelebs) (Pennycott et al. 2005; Lawson et al. 2006).
ETIOLOGY Rivolta (1878) discovered the etiologic agent of trichom*onosis in 1878 and named it Cercomonas gallinae. Rivolta also described another flagellate from
Rock Pigeon (Columba livia)
Host
122 W C* C* C*
Hungary India (Hyderabad) India (Calcutta) Iran
Greece
Germany
England Ecuador (Galapagos Islands)
Egypt
Canada (Alberta) Canada (Ontario) Canada (Quebec) Chile (Santiago) Croatia
— 2 2 NG
1 22 NG 1 18 104 NG 6 2 11 148 232 60 35 23 8 10 6 8 NG 32 41 14
Infected
— — — —
— 71 — — 27 62 — — — 11 52 70 57 40 24 — 100 38 44 60–100 — — 13
%
Githkopoulos and Liakos (1987) Ratz (1913) Mohteda (1956) Bhattacharya et al. (1997) Bozorgmehri-Fard and Moeinvaziri (1985)
Padilla et al. (2004) Friedhoff (1982) Knispel (2005)
DEFRA (2003) Harmon et al. (1987)
Hart (1941) McKeon et al. (1997) Reece et al. (1985) Krenn (1935) Tasca and DeCarli (1999) De Carli et al. (1979) Pybus and Onderka (2001) CCWHC (2007) CCWHC (2007) Toro et al. (1999) Greguric et al. (1986) Boˇsnjak and Greguric (1989) Abd-El-Motelib et al. (1994)
Literature source
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— 2 2 ∼720
1 31 NG 3 68 167 NG NG NG 100 285 332 106A 87J 95N NG 10 16 18 NG NG NG 110
C* C C* C* W W W* W* W* W W W W C* C* W* W W* W C* C* C W*
Australia (Glenfield) Australia (Perth) Austria (Vienna) Brazil
Examined
Status
Location
Number of birds
Table 6.1. Reports of Trichom*onas gallinae in free-ranging and captive columbiforms.
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123 USA (Colorado) USA (Connecticut)
USA (Alabama) USA (Arkansas) USA (California)
South Africa (Cape Town) South Africa (Wellington) Spain Sudan Switzerland Trinidad Turkey United Arab Emirates
W* NG NG C* W* W* C* W W C* W W C W W W, C C* C* W NG* C W W W* C* W W C* W W* C* 262 85 109 NG NG NG NG NG NG NG — 57 96 293 139 399 2 10 101 NG NG 44 80 60 150 1 10 >100 11 100 50
41 6 35 NG NG NG NG NG NG NG — NG NG 109 11 304 2 5 80 NG NG 9 60 21 102 1 8 NG 11 69 30
16 7 32 — — — — — — — — — — 37 8 76 — 50 79 — — 20 75 35 68 — — 30 100 69 60 (continues)
Haugen and Keeler (1952) Barrows (1975) Niemeyer (1939) Stabler and Herman (1951) Stabler (1951) Stabler and Herman (1951)
Dobeic (2003) Dovc et al. (2004) Zadravec et al. (2006) Jowett (1907) Pepler and Oettl´e (1992) Mart´ınez-Moreno et al. (1989) Ballouh and Eisa (1980) Sporri (1938) Kaminjolo et al. (1988) Gulegen et al. (2005) Bailey et al. (2000)
Delogu et al. (1997) Tacconi et al. (1993) Catelli et al. (1999) Oguma (1931) Minowa et al. (1982) Pepler and Oettl´e (1992) Bos (1932) Oyedapo et al. (2004) Samour et al. (1995) Tongson et al. (1969) Babes and Puscariu (1890) Kuliˇsic et al. (1996)
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Slovenia (Ljubljana)
Japan (Nakano) Japan (Tokyo) Namibia Netherlands Nigeria “Persian Gulf States” Philippines Romania Serbia (Belgrade)
Italy
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124 Netherlands Scotland Spain
USA (Louisiana) USA (Maryland, New Jersey, and Pennsylvania) USA (Maryland) USA (Minnesota) USA (Nebraska) USA (New Jersey) USA (New York) USA (North Carolina) USA (South Carolina) USA (Pennsylvania) USA (Virginia) USA (Texas) USA (Washington, DC) England
USA (Hawaii) USA (Illinois) USA (Iowa)
USA (Florida)
Location 13 27 7 2 50 NG 1 515 55 187 148 16 4 NG 62 1 10 20 20 153 148 NG 1,026 — NG 6A 91
W* W* W* W* W
Examined
W W C C* W W C* C* C W W* C* W W* C* W* W C W W W* W*
Status 9 21 6 2 50 NG 1 102 54 102 109 16 2 NG 17 1 1 20 12 19 109 “mass mortality” 79 — 1 2 31
Infected
Number of birds
Locke and Herman (1961) Waller (1934) Greiner and Baxter (1974) Cauthen (1934) Cauthen (1936) McCulloch (1950) Barrows (1975) Stabler and Shelanski (1936) Barrows (1975) Panigrahy et al. (1982) Stabler and Herman (1951) Duff (2003) Cousquer (2003) Jansen (1944) SACVS (2006) H¨ofle et al. (2004) Villanua et al. (2006)
<1 — — — 34
Shamis (1977) Forrester and Spalding (2003) Forrester and Foster (2001) Yager and Gleiser (1946) Jaskoski and Plank (1967) Stiles (1939) Kietzmann (1993) Rosenwald (1944) Stabler (1941b)
Literature source
69 78 — — 100 — — 20 98 55 74 100 — — 27 — — 100 65 12 74 —
%
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Common Wood-Pigeon (Columba palumbus)
Host
Table 6.1. (Continued )
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125
Red-eyed Dove (Streptopelia semitorquata)
Ring Turtle-Dove (Streptopelia risoria)§
NG 665 NG 125 NG NG 6 11 14 36 288 NG
W* W* NG W* W W* W* C* E* E* C* W*
England England Germany Italy Northern Ireland Scotland USA (Florida) USA (California) USA (Iowa) USA (Iowa) USA (New York) South Africa (Constantia)
Seychelles
USA (Colorado) Mauritius
156 NG NG NG NG NG 109 41 2,991 3
W W* W* W* W* W* W W W* W
USA (Arizona) USA (California)
41† 12
W W*
USA (Florida Keys)
2 25 NG 66 6 “several” 6 11 14 16 136 NG
8 NG ∼2,000 ∼16,000 ∼2,000 ∼300 21 9 1,504 1
36‡ 12
— <1 — 53 — — — 100 100 44 47 —
5 — — — — — 19 22 50¶ —
88 100
(continues)
Sileo and Fitzhugh (1969) Stabler and Braun (1979) NWHC (1995) Cole (1999) NWHC (2004) NWHC (2006) Stabler (1951) Swinnerton et al. (2005) Bunbury (2006) Bunbury (personal communication, September 16, 2007) Cornelius (1972) Cousquer (2003) Knispel (2005) Delogu et al. (1997) Beggs and Kennedy (2005) SACVS (1994) Forrester and Spalding (2003) Stabler and Herman (1951) Kietzmann (1993) Powell and Hollander (1982) Cauthen (1936) Pepler and Oettl´e (1992)
Kocan and Sprunt (1971)
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Pink Pigeon (Nesoenas mayeri) Seychelles Blue-Pigeon (Alectroenas pulcherrima) Eurasian Collared-Dove (Streptopelia decaocto)
White-crowned Pigeon (Patagioenas leucocephala) Band-tailed Pigeon (Patagioenas fasciata)
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126
Mourning Dove (Zenaida macroura)
USA (Colorado) USA (Connecticut) USA (Florida)
USA (California)
USA (Arkansas)
USA (Arizona)
USA (Hawaii) Canada (Ontario) USA (Alabama)
Australia (Perth) Mauritius South Africa (Constantia) Mauritius
Australia (Sydney) Malaysia Mauritius
Mauritius
Location W W W* C* W W W W W* W W W* W* W* W* W* W* W* W W* W W* W W* W* W* W
Status 109 247 1 8 3 32 76 4 NG 9 17 2 NG 204 NG NG NG NG NG 60 10 450 55 NG 100 40 142
Examined 17 115 1 3 1 6 35 2 NG 3 10 2 60 5 ∼50 3 1 108 NG 1 8 200 21 ∼1,400 23 28 10
Infected
Number of birds
16 47 — — — 19 46 — — — 59 — — 2 — — — — 16 2 80 45 38 — 23 70 7
%
Swinnerton et al. (2005) Bunbury et al. (2007) Hart (1940) Amin-Babjee et al. (1986) Swinnerton et al. (2005) Bunbury et al. (2007) McKeon et al. (1997) Bunbury et al. (2007) Pepler and Oettl´e (1992) Swinnerton et al. (2005) Bunbury et al. (2007) Kocan and Banko (1974) CCWHC (2007) Haugen and Keeler (1952) NWHC (2003) NWHC (1993) NWHC (1995) NWHC (2003) Hedlund (1998) Stabler and Herman (1951) Barrows (1975) Stabler and Herman (1951) Rupiper and Harmon (1988) NWHC (2004) Stabler (1951) Stabler and Herman (1951) Conti and Forrester (1981)
Literature source
September 11, 2008
Laughing Dove (Streptopelia senegalensis) Zebra Dove (Geopelia striata)
Madagascar Turtle-Dove (Streptopelia picturata) Spotted Dove (Streptopelia chinensis)
Host
Table 6.1. (Continued )
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127 W* W* W* C* W* W* W* W* W*
USA (Nevada) USA (New Mexico)
USA (North Carolina) USA (Ohio)
USA (New York)
USA (Nebraska)
W W* W* W W* W*
USA (Massachusetts) USA (Michigan) USA (Missouri)
W* W W* W* W* C* W* W* C* W* W* W W* W* NG 10 NG NG NG 4 NG NG NG NG NG NG 32 14N 520J,A 80 NG NG 4,052 1 121A 17J 7N NG NG NG 5 NG NG 2N NG NG
5 1 5 ∼18 ∼40 4 3 34 “several” ∼15 13 3 1 2 13 2 ∼22 ∼10 226 1 57 7 2 6 ∼800 ∼300 4 143 ∼500 1 NG ∼195
— 10 — — — — — — — — — — 3 14 3 3 — — 6 — 47 41 29 — — — — — — — 11 —
(continues)
NWHC (2003) Cole (1999) NWHC (1995) Cauthen (1936) NWHC (1998) Cole (1999) Harwood (1946) Stabler and Herman (1951) NWHC (1983)
Stabler and Herman (1951) NWHC (1991) NWHC (1998) Schulz et al. (2005) Padilla et al. (2004) Greiner and Baxter (1974)
NWHC (1989) Barrows (1975) NWHC (1998) NWHC (2001) NWHC (2002) Stabler and Herman (1951) NWHC (1995) Barnes (1951) Stabler and Herman (1951) NWHC (1993) NWHC (2003) Locke and Herman (1961) Stabler and Herman (1951) Locke and Herman (1961)
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USA (Kentucky) USA (Maryland)
USA (Indiana) USA (Iowa) USA (Kansas)
USA (Illinois)
USA (Georgia)
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128 W W W
USA (Wisconsin) Ecuador (Galapagos Islands) USA (Arizona)
USA (Florida)
USA (West Virginia)
USA (Virginia)
USA (Texas) USA (Utah)
USA (Tennessee)
W* W* W*
USA (Oregon) USA (Pennsylvania) USA (South Carolina) W* W* W W* W W W* W* W* W* W W* W* W* W
W
Status
USA (Oklahoma)
Location
19A 23N NG 25
163A,J 20N NG NG 56A 252J NG NG NG NG 101N 155 230 1 6N NG 20 NG NG 2 27
Examined
11 21 NG 25
21 6 ∼90 ∼24 9 5 5 ∼12 1 ∼34 53 27 1 1 5 ∼15 13 10 ∼15 2 3
Infected
Number of birds
58 91 98 100
13 30 — — 16 2 — — — — 52 17 <1 — — — 65 — — — 11
%
Hedlund (1998) Conti et al. (1985)
Toepfer et al. (1966)
Stabler and Herman (1951) Sprunt (1957) NWHC (2000) Barrows (1975) NWHC (1998) NWHC (2000) Stabler and Herman (1951) Harmon et al. (1987)
NWHC (2001) NWHC (2002) Locke and Herman (1961) NWHC (2001) Locke and Herman (1961) Ostrand et al. (1995)
NWHC (1991) NWHC (2003) Kocan and Amend (1972)
Carpenter et al. (1972)
Literature source
September 11, 2008
Galapagos Dove (Zenaida galapagoensis) White-winged Dove (Zenaida asiatica)
Host
Table 6.1. (Continued )
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129 W
USA (Texas)
2
2
65 6 51 97 73 3 3 NG 3 NG —
97 35 100 100 99 — — 52 — —
Hayse and James (1964)
Stabler and Holt (1962) Locke et al. (1961) Hedlund (1998) Locke and James (1962) Callender and Simmons (1937)
Conti and Forrester (1981) Locke and Kiel (1960) Stabler (1961) Glass et al. (2001)
A, adults; W, wild bird infected naturally; C, captive bird infected naturally; E, bird experimentally infected; J, juveniles; N, nestlings; NG, not given by authors. * Lesions reported. † Number of nests examined that contained squabs. The total number of squabs examined was not given. ‡ Number of nests examined that contained at least one infected squab. The total number of infected squabs was not given. § Clements (2000) did not list this as a valid species. Stevenson and Anderson (1994) stated that “According to Goodwin (1967) and other authorities, the turtle-dove, long kept in captivity, was derived from the African Collared-Dove (Streptopelia roseogrisea), native to North Africa and Arabia. There is some doubt as to whether it is now specifically distinct from that species.” ¶ This prevalence value is derived from multiple examinations of 426 Pink Pigeons over a 20-month period. The numbers examined here and the numbers positive for Trichom*onas gallinae are actually the number of examinations rather than the numbers of birds examined. Many birds were examined multiple times.
White-tipped Dove (Leptotila verreauxi)
USA (Florida) USA (Texas) USA (Arizona) USA (Texas) Panama
67 17 51 97A 74J 4 4 NG 3 NG
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Common Ground-Dove (Columbina passerina) Inca Dove (Columbina inca)
W W W W W W W W W* E*
USA (Texas)
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C* W*
England South Africa W*
W* E* W* W W* W* W* W*
South Africa
USA (Arizona) USA (Pennsylvania)
Canada (British Columbia) Spain England Germany Poland
England
W* W* W* C*
Status
South Africa USA (New York)
Location
130 36 269N 39N
144N 2 89A 233N NG NG
1
1 2
1 1 2 1
Examined
14 175 61
115 1 1 140 3N 1J
1
1 2
1 1 2 1
Infected
39 65 64
79 — <1 60 — —
—
— —
— — — —
%
Cooper and Petty (1988) Krone et al. (2005) Wieliczko et al. (2003)
Rosenfield et al. (2002) Knispel (2005)
Boal and Mannan (1999) Stabler (1941b) Boal et al. (1998)
Keymer (1972) Oettl´e (1990); Pepler and Oettl´e (1992) Oettl´e (1990); Pepler and Oettl´e (1992)
Pepler and Oettl´e (1992) Rettig (1978) Stone and Nye (1981) Keymer (1972)
Literature source
September 11, 2008
Western Marsh-Harrier (Circus aeruginosus) Northern Goshawk (Accipiter gentilis)
Falconiforms Black Kite (Milvus migrans) Bald Eagle (Haliaeetus leucocephalus) Egyptian Vulture (Neophron percnopterus) Shikra (Accipiter badius) African Goshawk (Accipiter tachiro) Rufous-chested Sparrowhawk (Accipiter rufiventris) Cooper’s Hawk (Accipiter cooperii)
Host
Number of birds
Table 6.2. Reports of Trichom*onas gallinae from free-ranging and captive falconiforms and strigiforms.
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Booted Eagle (Aquila pennata) Secretary-Bird (Sagittarius serpentarius) Lesser Kestrel (Falco naumanni) Eurasian Kestrel (Falco tinnunculus) American Kestrel (Falco sparverius)
131 USA (Pennsylvania)
USA (New York)
NG 1 1
NG NG NG
C* W* C* W* W* C*
6
C?*
Japan/Republic of Botswana† Bahrain Spain Bahrain
Spain
10 1 12N 39 NG NG
374 NG 1
W* E* W* W* W W*
W* W W*
England Germany South Africa
2 1 1 1
USA (Idaho/Oregon) USA (Pennsylvania) Portugal Spain
W* W* E* W*
USA (Arizona) USA (Florida) USA (Pennsylvania) USA (Florida)
1 1 1
5 1J 7
3
4 1 6 14 3N 1A
9 1 1
2 1 1 1
— — —
— — —
—
(continues)
Tangredi (1978) Stone and Janes (1969) Stabler and Shelanski (1936)
Samour et al. (1995) Knispel (2005) Samour et al. (1995)
Koyama et al. (1971)
Cousquer (2003) Knispel (2005) Oettl´e (1990); Pepler and Oettl´e (1992) Beecham and Kochert (1975) Stabler (1941b) H¨ofle et al. (2000) Real et al. (2000) Knispel (2005) Knispel (2005)
<1 — — 40 — 50 36 — —
Stensrude (1965) Forrester and Spalding (2003) Stabler and Shelanski (1936) Forrester and Spalding (2003)
— — — —
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Golden Eagle (Aquila chrysaetos) Bonelli’s Eagle (Aquila fasciata)
Gray Hawk (Buteo nitidus) Red-shouldered Hawk (Buteo lineatus) Red-tailed Hawk (Buteo jamaicensis) Eurasian Buzzard (Buteo buteo)
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Strigiforms Barn Owl (Tyto alba)
Peregrine Falcon (Falco peregrinus)
C* C* C* C* C C* W* W* NG C* C* W* W
Saudi Arabia
Italy South Africa Spain
England
W* W* W* W* W*
C C* C* C*
USA (Pennsylvania) Bahrain Saudi Arabia Bahrain
USA (Colorado) USA (Pennsylvania) Saudi Arabia Canada (Saskatchewan) England Germany Saudi Arabia South Africa Spain USA (Pennsylvania)
C*
Status
England
Location
180 98 20 1 NG
2 12 NG 1 1 NG NG NG NG NG 1 NG 10
1 NG NG NG
1
Examined
1 1 10 1 1A
2 12 346 1 1 5 1 1 NG 30 1 1N 2
1 310 8 1,345
1
Infected
Number of birds
<1 <1 50 — —
— 100 — — — — — — — — — — 20
— — — —
—
%
Hardy et al. (1981) Cousquer (2003) Delogu et al. (1997) Pepler and Oettl´e (1992) Knispel (2005)
Samour (2000a) Samour (2000b) Samour and Naldo (2003) Hamilton and Stabler (1953) Stabler (1969) Samour and Naldo (2003) CCWHC (2007) DEFRA (2003) Knispel (2005) Samour and Naldo (2003) Pepler and Oettl´e (1992) Knispel (2005) Stabler (1941b)
Stabler (1969) Samour et al. (1995) Samour and Naldo (2003) Samour et al. (1995)
Keymer (1972)
Literature source
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Gyrfalcon (Falco rusticolus)
Red-necked Falcon (Falco chicquera) Merlin (Falco columbarius) Lanner Falcon (Falco biarmicus) Saker Falcon (Falco cherrug)
Host
Table 6.2. (Continued )
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W* W* W* C
Canada (Ontario) USA (California) USA (Florida) Germany W* W* W* W* W W W*
South Africa England Italy USA (Florida) USA (Louisiana) USA (Massachusetts) Italy
338 40 NG NG 1 118
1
1 3 NG NG
3
10 11 3 2 1 1
1
1 3 1 1
1
40 8 20 1 1
<1 28 — — — 1
—
— — — —
—
2 — 25 — 10
Oettl´e (1990); Pepler and Oettl´e (1992) Cousquer (2003) Delogu et al. (1997) Forrester and Spalding (2003) Pokras et al. (1993) Pokras et al. (1993) Delogu et al. (1997)
CCWHC (2007) Jessup (1980) Forrester and Spalding (2003) Knispel (2005)
Forrester and Spalding (2003)
Schulz (1986) Pokras et al. (1993) Work and Hale (1996) Pokras et al. (1993) Delogu et al. (1997)
NG, not given; A, adults; J, juveniles; N, nestlings; W, wild bird infected naturally; C, captive bird infected naturally; E, bird experimentally infected. * Lesions reported. † These birds were captured in Botswana and subsequently brought to Japan. Eleven to thirteen days after being put in a zoo, three of the birds died with lesions compatible with trichom*onosis. Therefore, it is not known whether the birds were infected in Botswana or in Japan.
Little Owl (Athene noctua)
Barred Owl (Strix varia)
W*
USA (Florida)
1,638 8 81 NG 10
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Eurasian Eagle-Owl (Bubo bubo) Spotted Eagle-Owl (Bubo africanus) Tawny Owl (Strix aluco)
European Scops-Owl (Otus scops) Eastern Screech-Owl (Megascops asio) Great Horned-Owl (Bubo virginianus)
USA (Hawaii) USA (Louisiana) Italy
W* W* W* W W*
USA (California)
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the liver of a pigeon and called it Cercomonas hepaticum. Later, both these species of Cercomonas were recognized as T. gallinae and were considered synonyms (Stabler 1938). Taxonomically, T. gallinae is in the family Trichom*onadidae (phylum Parabasalia, order Trichom*onadida) and is closely related to several other parasitic flagellates of veterinary and medical importance, including Tritrichom*onas foetus in cattle, Trichom*onas phasiani in game-farm pheasants, and Trichom*onas vagin*lis in humans. The molecular phylogeny of T. gallinae and other trichom*onads has been studied extensively during the past 10 years (see Kleina et al. 2004; Cepicka et al. 2005, 2006; Gaspar da Silva et al. 2007). The living organism varies from pear-shaped to round and measures from 12.5 to 20 μm in length, has four anterior flagella, no free posterior flagellum, an axostyle that protrudes posteriorly, and a well-developed undulating membrane (BonDurant and Honigberg 1994) (Figures 6.2a–6.2c). Recently, it has been discovered that T. gallinae has pseudocyst stages (Tasca and DeCarli 2003). These are spherical forms that internalize the flagella, do not have a true cyst wall (Figures 6.2d–6.2f), and behave as resistant forms under stressful environmental conditions. Their formation is reversible (Pereira-Neves et al. 2003). Their role in the epidemiology of trichom*onosis is not known, but pseudocysts of a related species, T. foetus, have been found to adhere to vagin*l epithelial cells at a higher ratio than the trophozoite forms and may have some role in host cell infectivity (Mariante et al. 2004). For additional details on morphology, the reader is referred to Stabler (1941a), Abraham and Honigberg (1964), and Tasca and DeCarli (2003). Trichom*onas gallinae reproduces by binary fission (Stabler 1941a) and grows readily in agnotobiotic and axenic cultures in a number of media, both liquid and semiliquid (BonDurant and Honigberg 1994). As a result, there is an extensive amount known about carbohydrate, nitrogen, and lipid metabolism and various nutritional requirements of this organism (see reviews by Stabler (1954) and BonDurant and Honigberg (1994)). It has been known for some time that there are variations in the strains of T. gallinae (Stabler 1954; BonDurant and Honigberg 1994). Most strains are either nonpathogenic or moderately pathogenic, but there are also virulent strains. One of the most virulent is the Jones’ Barn (JB) strain that kills nonimmune pigeons at approximately 8 days postinfection (PI). The pathogenicity of the JB strain decreases when the organism is grown in nonliving media and is restored subsequently when the strain is passed serially in nonimmune pigeons (Stabler et al. 1964). Another strain, the avirulent Amherst (AG) strain, loses its infectivity to pigeons after prolonged growth in culture (Honigberg
1979). Although strains can lose their pathogenicity and infectivity while being cultured, no changes in virulence or antigenic properties have been reported after 12 years of cryopreservation (Bosch and Frank 1972). DNA and RNA from a virulent stain can increase the virulence of a nonpathogenic strain grown in culture (Honigberg et al. 1971). The pathogenicity of strains of T. gallinae has been determined by several techniques. One is the use of a subcutaneous mouse assay developed and evaluated by Honigberg (1961) and Frost and Honigberg (1962). This assay consists of injection of quantified numbers of axenically cultured trophozoites of T. gallinae under the skin of purebred mice and measurement of the mean volumes of any subsequent lesions at 6 days after injection. There is a close correlation between the lesion volumes and pathogenicity, larger lesions resulting from highly pathogenic strains. Two virulent strains (JB and Eiberg or IBERG) were distinguished from the avirulent Stabler-gallinae (SG) strain by the use of isoenzyme electrophoresis (Nadler and Honigberg 1988). Mattos et al. (1997) were also able to distinguish between three strains of T. gallinae by the use of isoenzyme electrophoresis, but pathogenicity of the three strains was not reported. Restriction enzyme analyses have been used to distinguish between strains of T. gallinae from several avian species (pigeons, raptors, canaries, and parakeets), but the technique has not been applied to comparisons of pathogenic versus nonpathogenic strains (Knispel 2005). Hemolysis of erythrocytes has been correlated with pathogenicity of strains of T. vagin*lis (Dailey et al. 1990), but this has not been determined to be useful for the differentiation of pathogenic and nonpathogenic strains of T. gallinae. Comparative sequence analyses of 5.8S rRNA genes and internal transcribed spacer regions have been used to identify genera and species of trichom*onads, including T. gallinae. This technique was used to study 24 isolates of T. gallinae—19 from Pink Pigeons (Nesoenas mayeri) and 5 from Madagascar Turtle Doves (Streptopelia picturata) from 6 different sites on the Indian Ocean island of Mauritius (Gaspar da Silva et al. 2007). All isolates had identical sequences both to each other and to an unrelated but previously sequenced isolate of T. gallinae, indicating that the locus (ITS1/5.8S/ITS2) can be used as a species marker. Random amplified polymorphic DNA analyses of these same isolates allowed the authors to identify geographic and host species differences, indicating that these were different strains of T. gallinae. These techniques have not been applied to the differentiation of pathogenic and nonpathogenic strains of T. gallinae (Felleisen 1997; Knispel 2005; Gaspar da Silva et al. 2007). Further studies on these techniques are needed so that assays can be developed to readily identify
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Figure 6.2. Scanning electron micrographs of trophozoites (a–c) and the formation of pseudocysts (d–f) of a strain of Trichom*onas gallinae obtained from a Rock Pigeon (Columba livia) and cultured axenically in vitro. Note the invagin*tion of the flagellae and disappearance of the undulating membrane in the organisms shown in (d) and (e). Part (f) represents a pseudocyst. AF, anterior flagella; UM, undulating membrane; AX, axostyle; PC, periflagellar canal. Reprinted from Tasca and DeCarli (2003), with permission of Veterinary Parasitology. pathogenic strains of T. gallinae. Such assays would revolutionize our understanding of many epizootiological aspects of trichom*onosis and the impact of this disease on populations of wild birds.
Some strains exhibit sites of tissue/organ predilection. For example, the JB strain and the Eiberg strains are predominantly hepatotrophic while the Mirza strain primarily infects the head (sinuses, orbital regions,
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brain, and neck tissues) and the mucosa of the upper digestive tract (Narcisi et al. 1991; BonDurant and Honigberg 1994). When the JB strain was given to Mourning Doves (Zenaida macroura), however, the lesions occurred predominately in the lungs rather than in the liver (Kocan 1969b). This is usually the case with infections with the JB strain in Rock Pigeons. EPIZOOTIOLOGY Among columbiforms, the primary source of infection by T. gallinae is the Rock Pigeon (Stabler 1954; BonDurant and Honigberg 1994). Some pigeons and doves develop immunity to the harmful effects of virulent strains of T. gallinae because of previous infection with an avirulent strain and act as carriers. These birds can have concomitant infections of both virulent and avirulent strains that can be transmitted to other birds, a virulent strain on one occasion and an avirulent strain during another (Kocan and Herman 1971). The life cycle of T. gallinae is direct, the organism being passed from one host to another without involvement of intermediate or paratenic hosts. There are no resistant cyst stages (although there are pseudocysts; see Etiology section) and the trichom*onads are very sensitive to desiccation (Kocan and Herman 1971). Transmission from host to host occurs by several methods. In the case of columbiforms, the organism is transferred directly from the upper digestive tract and mouth cavity of infected adults to squabs via regurgitation of pigeon milk produced in the crop of the adult bird (Stabler 1947; Kietzmann 1990). Thus, newly hatched squabs become infected during their first feeding. Transmission also occurs via direct contact between infected and uninfected columbiforms while cross-feeding or billing during courtship (Kocan and Herman 1971). Infected birds with oral or throat lesions have difficulty swallowing large pieces of grain and will pick them up, contaminate them with the organism, and subsequently drop them (Kocan and Herman 1971). These contaminated pieces of grain can then be ingested by an uninfected bird. Under normal circ*mstances, both uninfected doves and pigeons and those infected with avirulent strains of T. gallinae will also drop seeds that they pick up while feeding. These contaminated seeds can be ingested by another susceptible uninfected bird. A third method is by the ingestion of contaminated water. Trichom*onas gallinae can live for 20 min to several hours in water depending on the salinity and for at least 5 days in moist grains (Kocan 1969a). Transmission of T. gallinae via contaminated drinking water was demonstrated experimentally by Kietzmann (1990) using caged Ring Turtle Doves (Streptopelia risoria). Using InPouch culture kits (see Diagnosis section), Bunbury et al. (2007) cultured water from
sources utilized by infected columbids in Mauritius and found that 2 of 15 samples were positive for T. gallinae. Galliforms, psittaciforms, and passeriforms are infected by using the same feeding and watering areas as infected columbiforms. Raptors, however, are infected by feeding on infected prey, especially columbiforms (Stabler 1941b). Trichom*onas gallinae has been shown to survive in dove carcasses for up to 48 h after death of the host (Erwin et al. 2000). Pseudocysts might have a role in this survival, which in turn could influence the transmission of the disease to raptors that feed on these carcasses, but this has not been investigated. Trichom*onosis can result from the transfer of only one trichom*onad. This was shown by Stabler and Kihara (1954), who infected five Rock Pigeons each with one trichom*onad of the highly pathogenic JB strain; all five birds died with typical lesions and signs of trichom*onosis within 8–14 days after infection. The prevalences of T. gallinae in various hosts from a variety of geographical locations are presented in Table 6.1 for columbiforms and Table 6.2 for raptors. The Rock Pigeon is the most commonly and widely reported host, and infected birds have been recorded from 31 countries. The prevalence, based on examinations of 4,778 Rock Pigeons from 17 countries, ranged from 7 to 100%, while the mean was 47%. Next to the Rock Pigeon, the Mourning Dove in the US has been studied the most extensively. Prevalences, based on examinations of 6,932 doves from 20 states, ranged from 2 to 100%, with a mean prevalence of 11%. These values for both Rock Pigeons and Mourning Doves are probably underestimates since most of the earlier reports were based on standard wet-mount microscopy which has been shown to be considerably less sensitive than culture or polymerase chain reaction (PCR) techniques that have been applied more recently (Bunbury et al. 2005). Bunbury (2006) found no differences in prevalence between sexes in endangered Pink Pigeons in Mauritius and an increasing probability of infection with increasing age. She also found that although higher temperatures and lower rainfall were associated with higher prevalences in Madagascar Turtle-Doves in Mauritius (Bunbury et al. 2007), temperature had the most significant influence. Although a determination of the prevalence of the etiologic agent T. gallinae in a population of birds by swab, culture, and PCR techniques is useful information, these values provide only minimal epizootiological data. This is because the number of birds that had been infected at one time, but are free of infection and thereby immune at the time of sampling, is unknown. Kocan and Knisley (1970) used a challenge technique as a means to determine the prevalence of immune birds in a population. They livetrapped Rock Pigeons
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Trichom*onosis and Mourning Doves in the Maryland–Washington DC area and determined by swab and culture techniques that the prevalence of infection by T. gallinae was 52% for the Rock Pigeons and 0% for the Mourning Doves. When the negative birds were challenged subsequently with the JB strain of T. gallinae, all became positive for the organism (trichom*oniasis), but 88% of the Rock Pigeons and 82% of the Mourning Doves were immune and did not develop the disease (trichom*onosis). As discussed in the Immunity section, the duration and loss of infections and the possibility of premunition could both be important factors in the epidemiology of trichom*onosis. When the same individuals were tested every 2 months over a 20-month period in Mauritius, Pink Pigeons were more likely to remain either positive or negative than they were to acquire or lose infections of T. gallinae (Bunbury 2006). However, a number of birds also gained and lost infections several times over the screening period. More information is needed on this topic. Information on the relative importance of trichom*onosis as a mortality factor in avian populations is sparse. Although studies of carcasses submitted for necropsy are sometimes limited by collection biases, they can provide some insights into causes of death when interpreted with caution. In Mauritius, 54% of 35 free-living Pink Pigeons found dead over a 4-year period died of trichom*onosis, the leading cause of death (Bunbury 2006). Similarly, Gerhold et al. (2007) reported that trichom*onosis accounted for 40% of the cases and was the leading cause of death of 135 Mourning Doves submitted for diagnostic determination from 8 southeastern states in the US over a 35-year period. The factors that trigger an epizootic of trichom*onosis are unknown, although a number of suggestions have been made, particularly in the case of Rock Pigeons living in a pigeon loft. These include improper food, crowding, poor ventilation, wet and dirty litter, and lack of sunlight (Stabler 1947). It is uncertain if such factors apply to other columbiforms. A massive outbreak of trichom*onosis in Common Wood-Pigeons (Columba palumbus) on wintering roosts in southern Spain during 2001 was attributed to concentrations of Wood-Pigeons at birdfeeders that were numerous and set up for pheasants and partridges at a nearby hunting estate (H¨ofle et al. 2004). It was felt that transmission was enhanced by contamination of grain at the feeders as described earlier (see also Kocan 1969a). CLINICAL SIGNS Most of the clinical signs of infected columbiforms are related to the oral lesions which prevent or impair feeding. These include weight loss, listlessness, and ruffled feathers. Yellowish caseous lesions can be seen
137
around the beak or eyes of infected birds and their faces look swollen (Cole 1999). Also, there can be an excess of watery saliva and a foul cheese-like smell (Bunbury 2006). Some of these same signs can be seen in infected raptors along with dyspnea and nasal and oral exudation (Pepler and Oettl´e 1992). Signs in captive psittaciforms such as Budgerigars (Melopsittacus undulatus) include wasting, matted feathers, diarrhea, and repeated vomiting (Baker 1986; Murphy 1992). PATHOGENESIS AND PATHOLOGY As mentioned previously, infections by some strains of T. gallinae occur in the absence of apparent disease (i.e., trichom*oniasis), while others vary from being mildly pathogenic to very pathogenic (i.e., trichom*onosis) (Stabler 1948a; Kocan and Herman 1971). This variation is thought to be related to a greater antigenic diversity in avirulent strains that may stimulate a stronger host immune response than occurs during infection with more virulent strains with lower antigenic diversity (Stepkowski and Honigberg 1972). Infection with mild strains results in excessive salivation and some inflammation of the oral cavity and throat, whereas infections with more virulent strains result in caseous lesions in the mouth (Figure 6.3), throat, and crop and even invasion of the sinuses, skull, and skin of the neck. Some highly virulent strains cause lesions only in the head, neck, and crop, but other strains invade internal organs such as liver, lungs, pericardium, air sacs, and pancreas via the bloodstream (Stabler and Engley 1946; Jaquette 1950; Kocan and Herman 1971). In severe cases, death can result as early as 4 days after infection. Kocan and Herman (1971) described the progression of the disease. Oral lesions are wellcirc*mscribed, yellow masses located on the floor or roof of the mouth or in the pharyngeal region. Early lesions may be small and flush with the surface of the epithelium, but later they often develop small spur-like projections in their centers and may coalesce to form large, caseous masses in the mouth and throat. These can completely block the passage of food, so that the bird becomes emaciated and dies of starvation. Death may also occur as a result of respiratory failure if the lesion blocks the trachea (Kocan and Herman 1971) or hepatic dysfunction if organisms invade the liver (Narcisi et al. 1991). Lungs and other organs may be involved in infections by highly pathogenic strains. Host factors rather than characteristics of the etiologic agent may be more important in determining which organs are invaded. For example, the highly pathogenic JB strain of T. gallinae invades the liver of Rock Pigeons, whereas the same strain invades the lungs of Mourning Doves (Kocan 1969b).
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Figure 6.3. Gross lesion (arrow) of Trichom*onas gallinae in the oral cavity of a Mourning Dove (Zenaida macroura). Reproduced from Cole (1999), with permission of the author.
Histopathological changes associated with infections of pathogenic strains of T. gallinae in Rock Pigeons have been studied experimentally by PerezMesa et al. (1961) for the JB strain and by Narcisi et al. (1991) for the Eiberg strain. Times required for the trichom*onads to reach the liver and cause death were 3 and 7–10 days PI for the JB strain and 7 and 14– 17 days for the Eiberg strain, but other histopathological findings were similar. Highlights of the histopathological findings of Perez-Mesa et al. (1961) follow. On day 2, trichom*onads were arranged side by side, perpendicular to the surface of the epithelium of the pharynx and formed a layer that resembled columnar epithelium (Figure 6.4a). There was no inflammatory reaction in this area except for a mild mononuclear reaction near the gland openings. On day 3, there were occasional shallow pharyngeal ulcers with an infiltration of leukocytes in the submucosa around the glands. In one bird, there were lung abscesses with necrotic centers surrounded by lymphocytes, mononuclear cells, and rare giant cells. Trichom*onads were seen between the necrotic centers and the peripheral normal lung tissues. They were also seen in the sinusoidal capillaries and in Disse’s spaces in the liver.
There were abscesses in the liver described as focal necrosis with infiltrations of mononuclear cells and heterophils. On day 4, pharyngeal ulcers had massive inflammatory reactions. The liver contained large advanced abscesses with necrotic areas surrounded by heterophils, mononuclear cells, and trichom*onads. On days 5 and 7, the pharyngeal ulcers became deeper (Figure 6.4b) and the liver abscesses were larger than on day 4. On day 7, trichom*onads were seen close to the vessels in the pharynx and were very numerous near the periphery of hepatic abscesses. During the course of this study, birds died between days 5 and 10 PI. PerezMesa et al. (1961) described the basic pathological response in Rock Pigeons infected with the JB strain as purulent inflammation. They further concluded that the pigeons died of massive hepatic destruction. Further insights into the pathogenesis of trichom*onosis were reported by Kietzmann (1993) in a study using scanning electron microscopy. He infected juvenile Ring Turtle-Doves with a pathogenic strain of T. gallinae obtained from a Rock Pigeon and followed the progression of infection for 240 h PI, with special emphasis on the events prior to canker formation. Between 6 and 19 h PI, small numbers of amoeboid
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(a)
139
(b)
Figure 6.4. Histopathological changes related to experimental infection of a Rock Pigeon (Columba livia) with the Jones’ Barn strain of Trichom*onas gallinae. (a) Trichom*onads arranged perpendicularly to the surface of intact squamous epithelium in the pharynx on day 2 postinfection. 200×. (b) Ulcer in the pharynx on day 5 postinfection. The center of the ulcer consists of necrotic purulent exudate surrounded by mononuclear cells. 80×. Reproduced from Perez-Mesa et al. (1961), with permission of Avian Diseases.
trophozoites of T. gallinae attached to microfolds and cell borders of squamous epithelium of the palatal– esophageal junction (Figure 6.5a). Kietzmann (1993) postulated that some unknown parasite-secreted factor initiated squamous cell damage, separation, and removal. This was followed by invasion of areas beneath the squamous cells by trichom*onads (Figure 6.5b) and accelerated desquamation, invasion of the mucosa, and the development of cankers between 19 and 240 h PI (Figures 6.6 and 6.7). There are no comparable experimental studies of the pathogenesis of trichom*onosis in birds of prey, psittaciforms, and other birds. However, a number of authors have described the lesions of trichom*onosis in raptors (Jessup 1980; Cooper and Petty 1988; Pokras et al. 1993; Samour et al. 1995; Heidenreich 1997; Samour 2000a). Infected areas include oral and nasal cavities, esophagus, crop, and sometimes soft tissues, the skull, and various internal organs such as the heart. Stomatitis due to bacterial infection by Pseudomonas aeruginosa has been reported as a sequel to trichom*onosis in captive Saker Falcons (Falco cherrug) in Saudi Arabia (Samour 2000b). In captive psittaciforms, the disease has been reported to involve the oral cavity, crop, esophagus, pharynx, inner nares, sinuses, and other parts of the respiratory tract (Ruhl et al. 1982; Baker 1986; Ramsay et al. 1990; Garner and Sturtevant 1992; Murphy 1992). Intracellular viruses or virus-like particles have been found in some strains of T. vagin*lis and T. foetus
(Benchimol et al. 2002; Vancini and Benchimol 2005). The role of these viruses in the pathogenesis of the diseases caused by these trichom*onads is not understood. Such viruses or virus-like particles have not been identified in T . gallinae. DIAGNOSIS The presence of trichom*onads can be determined by microscopic examination of wet smears prepared with sterile cotton-tipped swabs from the mucus of the mouth and oropharyngeal area for the presence of motile, flagellated protozoans (BonDurant and Honigberg 1994). In situations where the numbers of organisms are low, it is helpful to inoculate scrapings into a suitable growth medium and examine samples after the organisms have had time to multiply. Trichom*onas gallinae grows readily in a variety of liquid and semisolid media. Diamond’s medium or a modification of it has been used by a number of authors (Diamond 1954; Kocan and Amend 1972) and these were discussed by Stabler (1954) and BonDurant and Honigberg (1994). Cover et al. (1994) used a commercial product originally designed to culture infections of T. foetus in cattle (InPouch TF, BioMed Diagnostics, White City, Oregon, USA). These pouches had the same sensitivity as Diamond’s medium and were convenient and effective for use in the field. This InPouch system has been used successfully in a number of studies involving columbiforms (Glass et al. 2001; Schulz
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Figure 6.5. Scanning electron photomicrographs of the palatal–esophageal junction of a Ring Turtle-Dove (Streptopelia risoria) infected experimentally with a pathogenic strain of Trichom*onas gallinae. (a) Nineteen hours postinfection (PI). One trichom*onad (arrow) is located at a squamous cell border and another is involved at a cell border separation (B). (b) Nineteen hours PI. A trichom*onad can be seen under a loosened squamous cell within the intercellular space. Border remnants (arrows) can be seen where the cell was once attached to other cells. Scale bars = 5 μm. Reproduced from Kietzmann (1993), with permission of The Journal of Parasitology.
et al. 2005; Bunbury et al. 2007) and raptors (Boal et al. 1998). Bunbury et al. (2005) found the sensitivity of the InPouch system to be more than twice that of conventional wet-mount microscopy. Definitive identification of trichom*onads is accomplished by the amplification of the 5.8S rRNA region by PCR. Since the assay was first developed (Felleisen 1997), it has been used for studies on T. vagin*lis in humans (e.g., Mayta et al. 2000), T. foetus in cattle (e.g., Grahn et al. 2005), as well as T. gallinae in birds (e.g., H¨ofle et al. 2004; Villanua et al. 2006; Gaspar da Silva et al. 2007). At least two commercial companies in North America offer diagnostic testing of samples for T. gallinae by PCR (Zoologix Inc., Chatsworth, California and HealthGene Corporation, Toronto, Ontario, Canada). Liebhart et al. (2006) developed an in situ hybridization procedure for detecting Histomonas meleagridis in paraffin-embedded tissue samples and also evaluated probes for T. gallinae and
other organisms. Their probes were specific for H . meleagridis, but could not differentiate between T. gallinae and Tetratrichom*onas gallinarum. Refinement of this technique might be possible and would provide a useful diagnostic tool for retrospective studies of trichom*onosis in birds. Lesions in the throat or oral cavity of a living or dead bird can also be used to obtain a diagnosis of trichom*onosis. A definitive diagnosis of the disease is made by demonstrating the presence of the organism by PCR assay and by observing the typical lesions. Lesions caused by fungi (Aspergillus sp., Candida sp.), poxvirus, nematode infections (Capillaria sp.), or vitamin A deficiency can be similar superficially to those of trichom*onosis and this should be considered when making a diagnosis (Kocan and Herman 1971; Cole 1999). Birds that recover from trichom*onosis often lack pharyngeal folds (located in the back of the throat) as a result of the necrotizing process that occurs during
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Figure 6.6. Scanning electron photomicrograph of the palatal–esophageal junction of a Ring Turtle-Dove (Streptopelia risoria) 48 h after being infected experimentally with a pathogenic strain of Trichom*onas gallinae. Note the extensive desquamation of squamous cell sheets and palisading of trichom*onads (arrows). A palatal papilla is labeled (P). Scale bar = 50 μm. Reproduced from Kietzmann (1993), with permission of The Journal of Parasitology. infection. This observation can be helpful in identifying previous cases of disease, at least in some species (Kocan and Herman 1971). IMMUNITY Several authors have reviewed immunologic aspects of infection with T. gallinae in columbiforms (Stabler 1954; Kocan and Herman 1971; Honigberg and Lindmark 1987; BonDurant and Honigberg 1994). Kocan and Amend (1972) challenged Mourning Doves from an epizootic area in South Carolina and from an area in Maryland where no epizootic had occurred for at least 3 years. Eighty-five percent of the doves from the epizootic area were immune to trichom*onosis, whereas only 69% of the doves from the nonepizootic area were immune. Birds infected with a moderately virulent or avirulent strain of T. gallinae have strong protection against the pathogenic effects of a subsequent infection by a virulent strain (Stabler 1948b). This immunity has both cellular and humoral components. Phagocytosis of trichom*onads by leukocytes appears to be sufficient to arrest the disease in primary infections that involve avir-
ulent or mildly virulent strains, but this is not true for infections with highly virulent strains. The exact role of phagocytosis in immune birds is not known (Kocan and Herman 1971). Humoral antibodies may be more important and have been shown to provide protection. This protection can be passively transferred from immune to nonimmune hosts via plasma or serum (Kocan 1970; Kocan and Herman 1970). The exact mechanism is not clear, but Kocan and Herman (1971) suggested that the trichom*onads might be inhibited from penetrating the epithelium of the upper digestive tract or that they might be lysed after penetration. Goudswaard et al. (1979) demonstrated that IgA is found in pigeon milk and is also transferred into the bloodstream of squabs, probably by pinocytosis. Secretory IgA may play a role in transfer of immunity to T. gallinae, but this has not been investigated. Experimental studies using mice have resulted in additional evidence that protective immunity occurs in infections with T. gallinae. Warren et al. (1961) used a mouse model that included subcutaneous injections of antigens from T. gallinae and found that complete protection against infection was observed in 50% of the animals tested. Honigberg (1978) was unable to
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Figure 6.7. Scanning electron photomicrograph of the palatal–esophageal junction of a Ring Turtle-Dove (Streptopelia risoria) 216 h after being infected experimentally with a pathogenic strain of Trichom*onas gallinae. Note erosion of palatal papilla (P) where several layers of epithelium have been removed. Scale bar = 50 μm. Reproduced from Kietzmann (1993), with permission of The Journal of Parasitology.
confirm these findings, but did observe protection of mice via intraperitoneal injections of a living strain of T. gallinae of mild pathogenicity. There is some evidence that premunition may be important (Jaquette 1948; Stabler 1954). Although some pigeons are positive for T. gallinae for up to 620 days after infection, others lose their infections with time. Stabler (1954) stated that immunity gradually diminishes after infections are lost, but gave no data to back up this assertion. This should be further investigated. Little new information on immunity has been published since the review of BonDurant and Honigberg (1994). However, studies on immunology of related organisms such as T. foetus in cattle and T. vagin*lis in humans are numerous and may provide some clues to immunologic aspects of trichom*onosis and T. gallinae infections in birds. In the case of immunity to T. foetus infections, antibody on the mucosal surface is critical for protection and vaccination to stimulate production of IgA and IgG1 pathogen-specific antibodies has proven successful (Corbeil et al. 2003). There is no clear proof of the protective characteristics of antibodies in immunity to T. vagin*lis infections and it
has been concluded that in addition to antibody, innate immune and acquired cellular responses are likely as important (Schwebke and Burgess 2004). Similar patterns may hold true for avian trichom*onosis and should be investigated in order to understand more about the host response to T . gallinae. PUBLIC HEALTH CONCERNS There are no reports of infections of T. gallinae in humans (Cole 1999). DOMESTIC ANIMAL HEALTH CONCERNS There are reports of trichom*onosis and infections with T. gallinae in domestic poultry. However, such infections are seen only occasionally in turkeys, and are even more rare in chickens (Willoughby et al. 1995). Feral Rock Pigeons are the source of infection in domestic pigeons and poultry (Kocan and Herman 1971), although other free-ranging doves might also be involved. There are also a number of records of trichom*onosis in captive psittaciforms kept as pets (Garner and Sturtevant
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Trichom*onosis 1992; Murphy 1992; Rosskopf and Woerpel 1996). These infections may have originated from Rock Pigeons or other columbiforms as well. WILDLIFE POPULATION IMPACTS Trichom*onosis has a negative effect on its avian host, but as with most wildlife diseases, the impact at the population level is difficult to measure. Although trichom*onosis has caused mortality in many different species of free-ranging columbiforms and raptors in various parts of the world (Tables 6.1 and 6.2), these have usually involved small or moderate numbers of birds. The most significant recorded outbreaks have occurred in Mourning Doves in the US, although many of these were local and involved only 100–800 birds. Such outbreaks, for example, have been reported in California (Stabler and Herman 1951; Cole 1999), Nebraska (Greiner and Baxter 1974), New Mexico (Cole 1999), North Carolina (Cole 1999), and South Carolina (Kocan and Amend 1972). The largest epizootic on record occurred in the southeastern US over a 2-year period (1950–1951) when an estimated 50,000–100,000 Mourning Doves died (Haugen 1952; Haugen and Keeler 1952). The die-off was centered in Alabama, but other neighboring states were also involved. Within Alabama, mortality was widespread, with losses being reported in 43 of the state’s 67 counties. The overall negative impact of this die-off on dove populations was reflected by poor hunting success during the 2 years following the outbreak (Haugen 1952). Another major epizootic occurred in California during 1988 in which at least 16,000 Band-tailed Pigeons (Patagioenas fasciata) died (Cole 1999). During the spring and summer of 2001 an epizootic involving at least 1,000 Eurasian Collared-Doves (Streptopelia decaocto) occurred in Florida (Forrester and Spalding 2003). Trichom*onad lesions were primarily in the liver, although there were a few birds with cankers in the oral cavity, throat, and crop. This outbreak was complicated by the presence of concurrent infections with pigeon paramyxovirus, which made interpretation of the cause of the mortality difficult. In 2001 and 2002, large numbers of Common Wood-Pigeons died of trichom*onosis in southwestern Spain (H¨ofle et al. 2004) and southern England (Duff 2002). The number of birds dying in Spain was estimated at 2,600, which represented about 15% of the wintering population. Even so, the disease did not appear to have a serious effect on the population as judged by the numbers of birds counted in the following winter (H¨ofle et al. 2004). The impact on the Common Wood-Pigeon population in England is not known. Several authors have suggested that trichom*onosis may have played an important role in the final extinc-
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tion of the Passenger Pigeon (Ectopistes migratorius) (Stabler 1954; Hanson 1969). There are no definitive data to support this idea, but it has been noted that Rock Pigeons were introduced into North America by the earliest colonists and may have been infected with virulent strains of T. gallinae. These Rock Pigeons may have been the source of infections in native species of columbiforms including the Passenger Pigeon (Haugen 1952). The role of trichom*onosis in the downward trend in populations of Mourning Doves in Utah was investigated (Ostrand et al. 1995). In a 2-year study, the prevalence of T. gallinae was 17%, but only 1 of 230 doves examined had lesions. Because of the low prevalence of birds with lesions, it was concluded that trichom*onosis was not a factor contributing to the decline in Mourning Doves. However, the use of data on the prevalence of lesions might lead to an underestimation of the impact of trichom*onosis since many, if not most, infected birds might die and thereby would not be included in the analysis. In a recovery program in Mauritius for the endangered endemic Pink Pigeon, survival of squabs to 30 days of age increased from 27 to 62% when the birds were treated with carnidazole (Swinnerton et al. 2005). However, treatment did not significantly increase juvenile (postfledging) survival to 150 days. In the same program, a negative effect of infection with T . gallinae on adult survival, reproductive success, and fledgling survival of Pink Pigeons was documented (Bunbury 2006; Bunbury et al. 2008). Birds that were not infected with T. gallinae had a significantly higher probability of surviving for 2 years after examination than those that were (Figure 6.8). Common Wood-Pigeons infected with a nonpathogenic strain of T. gallinae were lesion-free, but had lower body masses and fat reserves (Villanua et al. 2006). It was concluded that, although not fatal in and of themselves, these effects could lead to increased susceptibility of these birds to predation or other diseases and thereby exert a negative impact on the population. Additionally, the authors stated that this increased susceptibility of infected birds to predation would put birds of prey at a higher risk of exposure to T. gallinae after ingestion of these infected pigeons. The effect of trichom*onosis on populations of Peregrine Falcons (Falco peregrinus), which commonly feed on columbiforms, has been addressed (Stabler 1969). It was concluded that even though there is evidence that some Peregrine Falcons and other columbiform-eating raptors are infected with T. gallinae and contract the disease, the population impact was negligible. Trichom*onosis was diagnosed as the cause of death in 14 nestling Northern Goshawks (Accipiter gentilis)
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Figure 6.8. Kaplan–Meier survivorship curves for Pink Pigeons (Nesoenas mayeri) in Mauritius that were tested for Trichom*onas gallinae in three consecutive 2-month periods (n = 233). Pigeons were either not infected, intermediately infected, or always infected. Reproduced from Bunbury et al. (2008), with permission of Biological Conservation.
during a study conducted in Scotland over a 13-year period (Cooper and Petty 1988). It was concluded that the disease slowed the expansion of the reintroduced population of Northern Goshawks by about 15%. During the course of a comparative study of the breeding ecology of Cooper’s Hawks (Accipiter cooperii) in urban and exurban (undeveloped, natural) areas in southeastern Arizona, no nestling mortality of exurban birds due to trichom*onosis was found (Boal and Mannan 1999). By contrast, 80% of the 73 nestlings found dead in the urban study area died of the disease. The food habits of Cooper’s Hawks in the two areas were very different. Only a small proportion of the diet of exurban Cooper’s Hawks consisted of columbiforms, whereas in the urban area 84% of the diet was Mourning Doves and Inca Doves (Columbina inca), which were known
to have prevalences of T. gallinae infections of 16 and 52%, respectively. TREATMENT AND CONTROL Treating free-ranging wild birds is problematic. While mortality during an outbreak of trichom*onosis in Common Wood-Pigeons in Spain ceased after dimetridazole was used to treat grain at game bird feeders, harmful effects were documented in nontarget species (H¨ofle et al. 2004). Dimetridazole can be toxic to birds (Reece et al. 1985), and in the area where the 2001 outbreak occurred, reductions in numbers of chicks and lower than normal populations of adult Red-legged Partridges (Alectoris rufa) were noted in the following autumn. By contrast, carnidazole, ronidazole, and
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Trichom*onosis dimetridozole have been used with limited success to treat trichom*onosis in one subpopulation of Pink Pigeons on Mauritius, successfully increasing survival rates of squabs, juveniles, and adults (Swinnerton et al. 2005). Drugs that have been used to successfully treat infections in captive pigeons, raptors, and psittaciforms include some of the nitroimidazoles such as metronidazole, dimetridazole, ronidazol, and carnidazole (Emanuelson 1983; Ramsay et al. 1990; Pokras et al. 1993). Dimetridazole has been used successfully in drinking water to treat captive Rock Pigeons (Inghelbrecht et al. 1996), while metronidazole and carnidazole have been used effectively in raptors (Redig 2003). However, therapeutic failures due to drug resistance to several nitroimidazoles have been documented (Franssen and Lumeij 1992; Munoz et al. 1998). Currently, dimetridazole and metronidazole are not approved for use in birds in the US (Janzen 2006). Several synthetic compounds (chalcones) show evidence of potent activity against T. gallinae along with low toxicity to the host (Oyedapo et al. 2004) and may be good alternatives to nitroimidazoles when drug resistance is a problem. Appropriate measures to control trichom*onosis among wild and captive columbiforms and other birds should include actions to reduce sources of infection (Swinnerton et al. 2005). These include measures to minimize the use of contaminated communal food and water sources and measures to reduce stress from factors such as other pathogens and food shortages which can lead to reduction in resistance to trichom*onosis. Backyard bird feeders and artificial watering areas should be kept clean. Food should be changed regularly and the feeders and other types of food platforms should be disinfected with a 10% bleach solution. To prevent disease transmission, attention should also be given to prevention of flocks of doves and pigeons coming to feed at grain storage facilities and feedlots for livestock (Cole 1999). The control of trichom*onosis in wild raptors is very difficult, if not impossible, but in captive birds it can be prevented by avoiding the use of infected columbiforms as food sources (Halliwell 1979). When outbreaks of trichom*onosis occur in birds other than Rock Pigeons, the Rock Pigeons in the immediate area should be checked to determine if they contain lethal strains of T. gallinae. Until an assay is developed that can distinguish pathogenic from nonpathogenic stains, this must be done by culturing the parasite and then conducting transmission tests using Rock Pigeon squabs that have never been exposed to T. gallinae (Conti et al. 1985). Flocks of wild Rock Pigeons that are identified as being infected with lethal strains can be captured and treated or humanely elim-
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inated. This approach may not be acceptable in many areas or countries, but might be worthy of consideration under appropriate circ*mstances. MANAGEMENT IMPLICATIONS Since there are pathogenic and nonpathogenic strains of T. gallinae, management of trichom*onosis requires knowledge about the distribution of lethal strains before actions to minimize or even eliminate their impact on wild and captive populations of columbiforms, falconiforms, strigiforms, domestic poultry, and other susceptible birds of concern can be taken. This can be accomplished by conducting surveillance of freeranging doves and pigeons, especially Rock Pigeons that are the primary source of lethal strains (Stabler 1954; BonDurant and Honigberg 1994). This could be followed by treatment or eradication of infected birds. The translocation of nonendemic columbiforms into new areas should be avoided or done with great caution to eliminate the possibility of introducing pathogenic strains of T. gallinae into new areas. This is particularly true for islands where endemic species may be highly susceptible (see review by Wikelski et al. 2004). For example, pathogenic strains of T. gallinae are believed to have been introduced to Mauritius after release of Rock Pigeons and several species of exotic doves. This disease is now a serious threat to the survival of the endemic Pink Pigeon (Swinnerton et al. 2005; Bunbury 2006). ACKNOWLEDGMENTS The following people are acknowledged for their kind assistance in the translation of references: Claus Buergelt, Gabriele Forrester, Sandra Seng (German), Teresa DeLaFuente (French), Maarten Drost (Dutch), Mayuko Omori (Japanese), and QiYun Zeng (Chinese). We also thank Thomas Bailey, Robert BonDurant, Jacqui Brown, Nancy Bunbury, Kathy Converse, Glen Cousquer, Paul Duff, Glenn Kietzmann, Krysten Schuler, and Kevin Tyler for their assistance with acquisition of published and unpublished information on trichom*onosis. Nancy Bunbury was especially helpful in that regard and also read and critiqued an early draft of the manuscript. LITERATURE CITED Abd-El-Motelib, T. Y., G. Galal B El, and B. El Gamal Galal. 1994. Some studies on Trichom*onas gallinae infections in pigeons. Assiut Veterinary Medical Journal 30:59, 277–288. Abraham, R., and B. M. Honigberg. 1964. Structure of Trichom*onas gallinae (Rivolta). Journal of Parasitology 50:608–619.
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Dailey, D. C., T. H. Chang, and J. F. Alderete. 1990. Characterization of Trichom*onas vagin*lis haemolysis. Parasitology 101:171–175. De Carli, G. A., M. C. G. Pansera, and J. Guerrero. 1979. Trichom*onas gallinae (Rivolta, 1878) Stabler, 1938, no trato digestivo superior de pombos dom´esticos, Columbia livia, no Rio Grande do Sul—Primeiro Regestro. Acta Biologia Leopoldensia 1:85–95. DEFRA. 2003. Wildlife diseases in the UK. Cases reported in 2003. Department of Environment, Food, and Rural Affairs. Available at Defra.gov.uk/ corporate/vla/science/documents/science-end.oie03. pdf. Accessed February 9, 2007. Delogu, M., M. A. de Marco, V. Guberti, and S. Govoni. 1997. Epidemiologia della tricomoniasi in popolazioni di falconiformi, accipitriformi e strigiformi. Selezione Veterinaria 8–9:819–825. Diamond, L. S. 1954. A comparative study of 28 culture media for Trichom*onas gallinae. Experimental Parasitology 3:251–258. Dobeic, M. 2003. Regulation of population size of street pigeons in Ljubljana, Slovenia. Acta Veterinaria Beograd 53:171–182. Dovc, A., O. Zorman-Rojs, A. Vergles-Rataj, V. Bole-Hribovsek, U. Krapez, and M. Dobeic. 2004. Health status of free-living pigeons (Columba livia domestica) in the city of Ljubljana. Acta Veterinaria Hungarica 52:219–226. Duff, J. P. 2002. Wildlife diseases in the UK 2002. Report to the DEFRA and OIE, pp. 12. Duff, J. P. 2003. Surveillance report—wildlife. Veterinary Laboratories Agency, Regional Laboratories, Monitoring of Diseases of Wildlife Project (DEFRA), Penrith, United Kingdom, Quarterly Report 4:1–4. Emanuelson, S. 1983. Avian trichom*oniasis. In Current Veterinary Therapy VIII, R. W. Kirk (ed.). W. B. Saunders Company, Philadelphia, PA, pp. 619–621. Erwin, K. G., C. Kloss, J. Lyles, J. Felderhoff, A. M. Fedynich, S. E. Henke, and J. A. Robinson. 2000. Survival of Trichom*onas gallinae in white-winged dove carcasses. Journal of Wildlife Diseases 36:551–554. Felleisen, R. S. J. 1997. Comparative sequence analysis of 5·8S rRNA genes and internal transcribed spacer (ITS) regions of trichom*onadid protozoa. Parasitology 115:111–119. Florent, A. 1938. La trichom*oniase de pigeon. Annales de Medecine Veterinaire 83:401–428. Forrester, D. J., and G. W. Foster. 2001. Unpublished data. College of Veterinary Medicine, University of Florida, Gainesville, FL. Forrester, D. J., and M. G. Spalding. 2003. Parasites and Diseases of Wild Birds in Florida. University Press of Florida, Gainesville, FL.
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Trichom*onosis Swinnerton, K. J., A. G. Greenwood, R. E. Chapman, and C. G. Jones. 2005. The incidence of the parasitic disease trichom*oniasis and its treatment in reintroduced and wild pink pigeons Columba mayeri. Ibis 147:772–782. Tacconi, G., A. Moretti, D. Piergili-Floretti, and M. Latini. 1993. Endoparasitoses of pigeons (Columba livia, Gmelin 1789): Epidemiological survery in the city of Terni. Zootecnica International 4:83–85. Tangredi, B. P. 1978. Occurrence of trichom*oniasis on Long Island. New York Fish and Game Journal 25:89–90. Tasca, T., and G. A. DeCarli. 1999. Prevalence of Trichom*onas gallinae from the upper digestive tract of the common pigeon, Columba livia in the southern Brazilian state, Rio Grande do Sul. Parasitologia al dia 23:42–43. Tasca, T., and G. A. DeCarli. 2003. Scanning electron microscopy study of Trichom*onas gallinae. Veterinary Parasitology 118:37–42. Toepfer, E. W., L. N. Locke, and L. H. Blankenship. 1966. The occurrence of Trichom*onas gallinae in white-winged doves in Arizona. Bulletin of the Wildlife Disease Association 2:13. Tongson, M. S., M. N. Novilla, S. Loningkit, and E. Balediata. 1969. An outbreak of avian trichom*oniasis in the Philippines. Philippine Journal of Veterinary Medicine 8:141–145. Toro, H., C. Saucedo, C. Borie, R. E. Gough, and H. Alca´ıno. 1999. Health status of free-living pigeons in the city of Santiago. Avian Pathology 28:619–623. Turbervile, G. 1575. The Book of Faulconrie or Hawking. Christopher Barker, London. Vancini, R. G., and M. Benchimol. 2005. Appearance of virus-like particles in Tritrichom*onas foetus after drug treatment. Tissue and Cell 37:317–323. Villanua, D., U. H¨ofle, L. Perez-Rodriguez, and C. Gortazar. 2006. Trichom*onas gallinae in wintering common wood pigeons Columba palumbus in Spain. Ibis 148:641–648.
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Volkmar, F. 1930. Trichom*onas diversa n. sp. and its association with a disease of turkeys. Journal of Parasitology 17:85–89. Waller, E. F. 1934. A preliminary report on trichom*oniasis of pigeons. Journal of the American Veterinary Medical Association 84:596–602. Warren, L. G., W. B. Kitzman, and E. Hake. 1961. Induced resistance of mice to subcutaneous infection with Trichom*onas gallinae (Rivolta, 1878). Journal of Parasitology 47:533–537. Wieliczko, A., T. Piasecki, G. M. Dorrestein, A. Adamski, and M. Mazurkiewicz. 2003. Evaluation of the health status of goshawk chicks (Accipiter gentilis) nesting in Wroclaw vicinity. Bulletin of the Veterinary Institute in Pulawy 47:247–257. Wikelski, M., J. Foufopoulos, H. Vargas, and H. Snell. 2004. Galapagos birds and diseases: Invasive pathogens as threats for island species. Ecology and Society 9:5 Available at http://www. ecologyandsociety.org/vol9.iss1/art5 Willoughby, D. H., A. A. Bickford, B. R. Charlton, and G. L. Cooper. 1995. Esophageal trichom*oniasis in chickens. Avian Diseases 39:919–924. Work, T. M., and J. Hale. 1996. Causes of owl mortality in Hawaii, 1992–1994. Journal of Wildlife Diseases 32:266–273. Yager, R. H., and C. A. Gleiser. 1946. Trichom*onas and Haemoproteus infections and the experimental use of DDT in the control of ectoparasites in a flock of Signal Corps pigeons in the Territory of Hawaii. Journal of the American Veterinary Medical Association 109:204–207. Zadravec, M., O. Z. Rojs, J. Racnik, A. V. Rataj, K. Vlahovic, and A. Dovc. 2006. The prevalence of trichom*oniasis in ornamental and free-living pigeons (Columba livia) in Slovenia. Veterinarske Novice 32:5–10. Zhang, T. L., J. W. Jiang, Z. Z. Qu, and Y. Zhao. 1982. Trichom*oniasis in fowls. Chinese Journal of Veterinary Medicine Zhongguo Shouyi Zashi 8:15–16.
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7 Histomonas William R. Davidson disease in domestic poultry was correlated with areas with climate and soils suitable for transmission of the parasite. Most reports of histomoniasis among wild birds have been from North America, but whether this reflects true prevalence of the disease is unclear.
INTRODUCTION Histomoniasis is a disease of galliform birds (order Galliformes) caused by the protozoan Histomonas meleagridis. For many years following its recognition in the late 1800s, this disease was a major problem in the production of domestic poultry, especially turkeys. Histomoniasis is also recognized as a severe disease of Wild Turkeys (Meleagris gallopavo) and has been reported on occasion among other species of wild galliform birds.
HOST RANGE Histomoniasis is a disease almost exclusively of birds in the order Galliformes. At least 12 species of nondomestic galliform birds are susceptible to infection (Table 7.1), although there is wide variation in susceptibility and clinical response to infection among species. Among wild populations, histomoniasis has been reported most frequently as a disease of Wild Turkeys and much less often in other species. However, histomoniasis is common in several species of galliform game birds raised in captivity. Reports in nongalliform birds, such as captive Ostriches (Struthio camelus) (Borst and Lambers 1985), are rare. Mallards (Anas platyrhynchos) and domestic geese were essentially refractory to experimental infection (Lund et al. 1974).
SYNONYMS Blackhead disease, infectious enterohepatitis, typhlohepatitis. HISTORY Excellent historical reviews of histomoniasis, including descriptions of earlier controversies regarding its etiology and epizootiology, have been published by Reid (1967) and Lund (1977). On the basis of a series of experimental infections in various species of galliform birds, Lund and Chute (1974) theorized that H. meleagridis evolved in Asia, probably as a parasite of Ring-necked Pheasants (Phasianus colchicus) or related Phasianus spp. Recognition that domestic chickens were also reservoir hosts for H. meleagridis led to the poultry industry axiom of not raising chickens and turkeys together (Reid 1967).
ETIOLOGY Histomonas meleagridis is the only member within the genus and is a pleomorphic flagellate in the family Monocercomonadidae, order Trichom*onadida, phylum Parabasalia (Brugerolle and Lee 2000). Histomonads are 4–30 m in diameter, rounded to elongate, have a single nucleus, exhibit active ameboid movement, and may have a single flagellum. The morphologic form is dependent on the stage of the infection and location of the parasite within the avian host. Both ameboid and flagellate trophozoites exist in the cecal lumen of infected birds. Organisms within lesions in the cecal wall or in the liver lack a flagellum. Reproduction is by binary fission. There is no cyst or environmentally resistant stage, and trophozoites shed in feces do not survive outside the avian host.
DISTRIBUTION Histomoniasis has been reported throughout the world in regions where chickens, turkeys, or other domesticated galliform birds are raised. The disease is more prevalent in warmer regions of the globe, but has occurred with some frequency near the limits of both northern and southern temperate zones (Lund 1972). Prior to implementation of effective prevention and control practices, the frequency of occurrence of the
154 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Table 7.1. Species of the order Galliformes reported to be susceptible to infection by Histomonas meleagridis. Species
Host status
Disease severity
Status as reservoir
Reference
Wild Turkey (Meleagris gallopavo)
Wild, captive, experimental
Severe
Poor
Northern Bobwhite (Colinus virginianus)
Wild, captive, experimental
Moderate
Marginal
Ruffed Grouse (Bonasa umbellus) Greater Prairie-Chicken (Tympanuchus cupido) Ring-necked Pheasant (Phasianus colchicus) Chukar (Alectoris chukar)
Captive
Severe
Poor
Stoddard (1935, 1936), Mosby and Handley (1943), Kozicky (1948), Snyder (1953), Roberts (1956), Thomas (1964), Bailey and Rinell (1968), Prestwood et al. (1973), Lund et al. (1975), Hurst (1980), Davidson et al. (1985), Schorr et al. (1988), Ley et al. (1989), Davidson and Wentworth (1992), and Forrester (1992) Kellogg and Reid (1970), Lund and Chute (1971b), Davidson et al. (1978), Zeakes et al. (1981), and Davidson et al. (1982) Bump et al. (1947)
NR
NR
NR
Braun and Willers (1967)
Captive, experimental
Mild
Superior reservoir
O’Roke (1933), and Lund and Chute (1972b–d, 1974)
Captive, experimental
Severe
Poor
Indian Peafowl (Pavo cristatus) Gray Partridge (Perdix perdix) Black Francolin (Francolinus francolinus) Red Junglefowl (Gallus gallus)
Captive, experimental Experimental
Severe
Poor
Mild
Poor
Wild
NR
NR
Chaddock (1948), Sims (1960), and Lund and Chute (1971a, 1972b, 1974) Lund and Chute (1972b, c, 1974) Lund and Chute (1972b, 1974) Bump and Bump (1964)
Wild (released), Captive Experimental
Mild
Important
Kellogg et al. (1971, 1978)
Mild
Poor
Mild to severe
Suitable
Lund and Ellis (1967), and Lund and Chute (1972b, 1974) Chute and Lund (1972, 1974), and Lund and Chute (1972e)
Japanese Quail (Coturnix japonica) Helmeted Guineafowl (Numida meleagris)
Experimental
NR, not reported.
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EPIZOOTIOLOGY The epizootiology of H. meleagridis is unusual in that under natural conditions transmission is dependent on the cecal nematode Heterakis gallinarum, which also infects many species of galliform birds. This discovery by Graybill and Smith (1920) is considered a milestone in parasitology (Lund 1977). Histomonads, in addition to infecting the ceca of the bird, also infect the cecal worms. Within the ovaries of female cecal worms, the histomonads become incorporated in the eggs of H. gallinarum (Lee 1969; Lund and Chute 1973). The protective covering of the cecal worm egg shields the delicate histomonads from deleterious environmental factors which otherwise rapidly kill the protozoans. When infective (larvated) histomonad-bearing heterakid eggs are ingested by a suitable host, both parasites are released in the ceca. Although some histomonads are released when the heterakid eggs hatch, the majority is liberated from cecal worm larvae that die and decompose within the ceca (Lund and Chute 1972a, 1974). This is important because infection of H. gallinarum by H. meleagridis occurs principally when the heterakid larvae are 10–20 days old, and histomonads from simultaneously acquired heterakid larvae that have died are most numerous in the cecal lumen during this time period (Lund 1968, 1971; Lund and Chute 1974). In addition to direct transmission via heterakid eggs, earthworms serve as important paratenic hosts of histomonad-infected H. gallinarum larvae. Under field conditions, earthworms can accumulate and store large numbers of heterakid larvae in somatic tissues. Both parasites are transmitted together, especially following periods of rain, when earthworms come to the soil surface and are consumed easily by susceptible birds (Lund et al. 1966; Kemp and Franson 1975). Grasshoppers can also serve as paratenic hosts for Heterakis and Histomonas (DeVolt and Davis 1936; Frank 1953) but their role is less important than earthworms (Reid 1967). It was recently demonstrated that histomonads can be effectively transmitted laterally by the fecal–oral route (Hu and McDougald 2003). However, this means of transmission appears to be restricted only to the crowded conditions of confinement-reared domestic poultry production and is not important among wild bird populations. Different members of the order Galliformes exhibit wide variation in their susceptibility to clinical histomoniasis, spanning the spectrum from an essential tolerance of the protozoan with minimal lesions to severe disease with a high case-fatality rate (Lund and Chute 1972b). Species such as Ring-necked Pheasants, domestic chickens, or junglefowl that develop minimal disease harbor and readily transmit both H.
meleagridis and H. gallinarum. Such species function as critical reservoir hosts for both parasites. Apart from the potential reduction in transmission caused by host deaths from histomoniasis, the clinical response to H. meleagridis infection has a major influence on transmission of both parasites. In ceca with lesions caused by Histomonas, the survival of the heterakid nematode is extremely poor, and heterakid infrapopulations in diseased hosts are often completely eliminated by altered conditions within the ceca. Thus, individual birds or species in which severe cecal lesions develop are poor reservoir hosts (Lund and Chute 1972b, 1974). Lund and Chute (1974) presented a well-supported hypothesis that both H. meleagridis and H. gallinarum evolved in Asia with the Ring-necked Pheasant or a close relative of the pheasant. This concept was based on experimental studies in various galliform species in which the number of histomonad-infected heterakid eggs produced per heterakid egg ingested were measured. There were great differences in the egg output/input ratios among the host species, ranging from less than 1 to 1 for Japanese Quail (Coturnix japonica), Gray Partridge (Perdix perdix), Peafowl, and Northern Bobwhites (Colinus virginianus) to greater than 5 to 1 for pheasants, chickens, and guineafowl (Lund and Chute 1974). These experiments delineated reservoir hosts, in which disease is rare or mild, from vulnerable hosts, in which disease is more severe. Junglefowl also are efficient reservoir hosts (Kellogg et al. 1978). Worldwide, domestic chickens currently are the most widely distributed and abundant reservoir host for H. meleagridis; however, in areas where they occur, wild populations of pheasants and junglefowl can serve as important reservoirs. In addition, H. meleagridis and H. gallinarum are both common in captivereared game birds, and if they are released in the wild, these birds can serve as sources of infection for vulnerable species. Captive-reared game birds can also differ from their wild counterparts in the risk they pose for transmitting H. meleagridis. For example, captive-reared Northern Bobwhites are often infected with H. gallinarum, whereas wild Northern Bobwhites rarely harbor H. gallinarum but are commonly infected with another cecal worm, Heterakis isolonche (or Heterakis bonasae). This distinction is important because Heterakis isolonche apparently does not transmit H. meleagridis, and thus, captive-reared and wild bobwhites differ in their epizootiologic risks (Davidson et al. 1978). CLINICAL SIGNS Clinical signs of histomoniasis are well described for domestic poultry, especially turkeys, and typically appear from 1 to 3 weeks following infection. Alterations
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of serum enzymes, serum proteins, and histochemical changes in the ceca and liver have been described in infected domestic poultry (Clarkson 1966; McDougald and Hansen 1970; Wilkins and Lee 1976). Affected birds are inactive, depressed, develop inappetence, and stand with drooped wings, closed eyes, retracted head, and ruffled feathers. Feces may be sulfur colored or contain flecks of blood and mucus. Birds that survive several weeks often become emaciated. Similar signs have been reported in wild and captive galliform game birds; however, because of the difficulties of monitoring wild populations, most affected individuals are reported only because they are so ill that they are easily captured or are found dead.
mal intestinal bacterial flora and gnotobiotic poults with monospecific intestinal bacterial flora consisting of Escherichia coli, Escherichia intermedia, Clostridium perfringens, Streptococcus fecalis, or Bacillus subtilis developed mild to severe disease. Monospecific infection with certain other bacteria allowed colonization of ceca by H. meleagridis but did not result in lesions (Reid 1967). Other host compromising factors may also influence outcome of infection. For example, exposure to the insecticide Sevin (1-napthyl N -methyl carbamate) increased the susceptibility of bobwhites to histomoniasis and markedly increased mortality (Zeakes et al. 1981).
PATHOGENESIS AND PATHOLOGY Histomonas meleagridis organisms are liberated from H. gallinarum larvae in the ceca where they begin to reproduce within the lumen. Within 4–7 days, organisms can be detected within small ulcerations in the cecal mucosa where they elicit a mixed inflammatory response that is predominantly composed of heterophils. Necrosis ensues and focal mucosal and submucosal lesions expand to become confluent, eventually extending into the muscularis layer. The lumen of the ceca fills with a mixture of cecal contents, serous and hemorrhagic exudates, inflammatory cells, and necrotic debris. The concentric layers of this admixture typically form a caseous cecal core. Histomonads continue to reproduce within lesions in the cecal wall and may gain entry into the venous blood vessels. They then are carried to the liver where they continue to reproduce, setting up discrete foci of hepatic necrosis that are grossly visible at about 10 days. Initially, foci of hepatic necrosis appear as small white spots that progress to gray depressed areas surrounded by a narrow rim of hemorrhage. As they enlarge, these lesions may coalesce and become more firm and white or yellow as fibrosis occurs. In severely affected birds, death often occurs between 14 and 21 days (Clarkson 1962). Although H. meleagridis is the critical etiologic agent of histomoniasis, experiments with gnotobiotic turkeys have demonstrated clearly that intestinal bacteria are essential for development of lesions. During the 1960s, separate laboratories independently confirmed that intestinal bacteria play two essential roles in the pathogenesis of histomoniasis (Doll and Franker 1963; Franker and Doll 1964; Bradley and Reid 1966). One role is enabling H. meleagridis to colonize the ceca of the bird; bacteria-free domestic turkey poults rarely could be infected with H. meleagridis and did not develop histomoniasis. The second role is enhancing the development of lesions. In marked contrast to bacteriafree poults, both conventional turkey poults with nor-
DIAGNOSIS The combination of necrotic cecal cores and multifocal hepatic necrosis in a galliform bird (Figure 7.1) is strong presumptive evidence of histomoniasis. Confirmation of infection in diseased hosts can be accomplished by histologic demonstration of histomonads (Figure 7.2) in ceca or liver using various stains such as periodic acid Schiff or silver stains (Kemp and Reid 1966), demonstration of live histomonads in saline mount preparations (75–80◦ F slide warming device required), in vitro cultivation of histomonads from ceca
Figure 7.1. Gross lesions of Histomonas meleagridis-infected liver (upper) and ceca (lower) from an experimentally infected turkey poult illustrating (arrows) multifocal hepatic necrosis, thickened and hemorrhagic cecal walls, and caseous cecal core. Reprinted with permission from Waters (1992).
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Parasitic Diseases of Wild Birds tion of commercial chicken and turkey flocks, and improved husbandry practices in range turkeys have greatly reduced losses due to histomoniasis. Appropriate husbandry and biosecurity practices are effective in preventing transmission of histomoniasis from wild or captive game birds and noncommercial chickens to commercial turkey flocks. Histomoniasis has no known public health implications.
Figure 7.2. Photomicrograph of submucosal region of cecum demonstrating Histomonas meleagridis (arrows) within lesions. Hematoxylin and eosin stain (891×). Reprinted with permission from Waters (1992).
or liver (McDougald and Galloway 1973), or by rectal inoculation of turkey poults with saline suspensions of cecal or liver lesions. Confirmation of infection in unaffected reservoir hosts is best accomplished by in vitro cultivation of cecal contents, by rectal inoculation of turkey poults with cecal suspensions, or by feeding embryonated heterakid eggs from the suspected host to turkey poults. Fresh specimens are required for in vitro cultivation, direct visualization in wet mounts, and bioassay procedures using turkey poults. Freezing may impair confirmation by histologic means. Differential diagnoses should include coccidiosis, coligranuloma, salmonellosis, and neoplasia. IMMUNITY Several species of galliform birds exhibit an age resistance to infection by H. meleagridis, with younger hosts being more susceptible to infection and developing more severe disease (Lund and Chute 1970; Lund 1972; Levine 1985). However, virulent strains of H. meleagridis can cause disease in hosts of any age. Birds that recover from histomoniasis develop immunity to reinfection (Lund 1972; Levine 1985). DOMESTIC ANIMAL AND PUBLIC HEALTH CONCERNS Historically, histomoniasis was a major disease among domestic turkeys; however, recognition of the reservoir role played by domestic chickens, the separa-
WILDLIFE POPULATION IMPACTS Although histomoniasis is frequently mentioned in scientific, semitechnical, and popular literature as a significant disease risk for wild galliform birds, especially Wild Turkeys, there are relatively few primary accounts of the disease in wild populations (Davidson and Wentworth 1992). Despite the few reports from Wild Turkeys, histomoniasis is believed to be one of the more important diseases of this species in the southeastern US (Hurst 1980; Davidson et al. 1985). Histomoniasis was the second most common infectious disease among sick or dead Wild Turkeys from eight southeastern states diagnosed and accounted for 14% of nontrauma diagnoses (Davidson et al. 1985). In sick or dead Wild Turkeys from Florida, histomoniasis was less frequent but accounted for 5% of nontrauma diagnoses (Forrester 1992). Histomoniasis appears to be rare among populations of other wild galliform birds, although information for many species is sparse.
PREVENTION, CONTROL, AND MANAGEMENT IMPLICATIONS Prevention and control of histomoniasis is predicated on separating reservoir hosts from vulnerable species and on breaking the cycle of the cecal worm vector. These objectives can be accomplished among captive flocks by not commingling reservoir and vulnerable species (e.g., chickens and turkeys), by housing flocks in deep stone or wire floored pens that reduce ingestion of eggs and earthworm paratenic hosts, or by removal of H. gallinarum with appropriate anthelmintics. Preventive actions applicable for wild galliform populations that achieve similar objectives include not introducing reservoir hosts (e.g., Ring-necked Pheasants or junglefowl) in habitats occupied by wild vulnerable species (e.g., Wild Turkey) and not using untreated manure from domestic chickens as fertilizer on areas frequented by vulnerable species. Although manure from commercial broiler chickens grown under modern husbandry practices poses little risk of histomoniasis, commercial breeder and layer flocks and noncommercial chickens still have high prevalences of infection (Waters et al. 1994). Risk of introducing
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Histomonas histomoniasis from junglefowl into native wild turkey populations was one factor in the abandonment of an earlier foreign game bird introduction program in the US (Kellogg et al. 1978).
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Davidson, W. R., and E. J. Wentworth. 1992. Population influences: Diseases and parasites. In The Wild Turkey: Biology and Management, J. G. Dickson (ed.). Stackpole Books, Harrisburg, PA, pp. 101–118. Davidson, W. R., G. L. Doster, and M. B. McGhee. 1978. Failure of Heterakis bonasae to transmit Histomonas meleagridis. Avian Diseases 22:627–632. Davidson, W. R., F. E. Kellogg, and G. L. Doster. 1982. An overview of disease and parasitism in southeastern bobwhite quail. In Proceedings of the Second National Bobwhite Quail Symposium, Series Number 6, F. sh*toskey, Jr., E. C. sh*toskey, and L. G. Talent (eds). Oklahoma State University Environmental, Stillwater, OK. Davidson, W. R., V. F. Nettles, C. E. Couvillion, and E. W. Howerth. 1985. Diseases diagnosed in wild turkeys (Meleagris gallopavo) of the southeastern United States. Journal of Wildlife Diseases 21:386–390. DeVolt, H. M., and C. R. Davis. 1936. Blackhead (infectious enterohepatitis) in turkeys, with notes on other intestinal protozoa. University of Maryland Agricultural Experiment Station Bulletin 392:493–567. Doll, J. P., and C. K. Franker. 1963. Experimental histomoniasis in gnotobiotic turkeys. I. Infection and histopathology of the bacteria-free host. Journal of Parasitology 49:411–414. Forrester, D. J. 1992. A synopsis of disease conditions found in wild turkeys (Meleagris gallopavo L.) from Florida, 1969–1990. Florida Field Naturalist 20(2):29–56. Frank, J. F. 1953. A note on the experimental transmission of enterohepatitis of turkeys by arthropods. Canadian Journal of Comparative Medicine 17:230–232. Franker, C. K., and J. P. Doll. 1964. Experimental histomoniasis in gnotobiotic turkeys. II. Effects of some cecal bacteria on pathogenesis. Journal of Parasitology 50:636–640. Graybill, H. W., and T. Smith. 1920. Production of fatal blackhead in turkeys by feeding embryonated eggs of Heterakis papillosa. Journal of Experimental Medicine 31:647–655. Hu, J., and L. R. McDougald. 2003. Direct lateral transmission of Histomonas meleagridis in turkeys. Avian Diseases 47:489–492. Hurst, G. A. 1980. Histomoniasis in wild turkeys in Mississippi. Journal of Wildlife Diseases 16:357–358. Kellogg, F. E., and W. M. Reid. 1970. Bobwhites as possible reservoir hosts for blackhead in wild turkeys. Journal of Wildlife Management 34:155–159. Kellogg, F. E., G. L. Doster, and J. K. Johnson. 1971. Diseases and parasites encountered in pen-raised Indian red junglefowl. Journal of Wildlife Diseases 7:186–187.
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Kellogg, F. E., T. H. Eleazer, and T. R. Colvin. 1978. Transmission of blackhead from junglefowl to turkey. In Proceedings of the Annual Conference of the Southeastern Association of Fish and Wildlife Agencies 32:378–379. Kemp, R. L., and J. C. Franson. 1975. Transmission of Histomonas meleagridis to domestic fowl by means of earthworms recovered from pheasant yard soil. Avian Diseases 19:741–744. Kemp, R. L., and W. M. Reid. 1966. Staining techniques for differential diagnosis of histomoniasis and mycosis in domestic poultry. Avian Diseases 10:357–363. Kozicky, E. L. 1948. Some protozoan parasites of the eastern wild turkey in Pennsylvania. Journal of Wildlife Management 12:263–266. Lee, D. L. 1969. The structure and development of Histomonas meleagridis (Mastigamoebidae: Protozoa) in the female reproductive tract of its intermediate host, Heterakis gallinarum (Nematoda). Parasitology 59:877–884. Levine, N. D. 1985. Veterinary Protozoology. Iowa State University Press, Ames, IA. Ley, D. H., M. D. Ficken, D. T. Cobb, and R. N. Witter. 1989. Histomoniasis and reticuloendotheliosis in a wild turkey (Meleagris gallopavo) in North Carolina. Journal of Wildlife Diseases 25:262–265. Lund, E. E. 1968. Acquisition and liberation of Histomonas wenrichi by Heterakis gallinarum. Experimental Parasitology 22:62–67. Lund, E. E. 1971. Histomonas meleagridis and H. wenrichi: Time of acquisition by Heterakis gallinarum. Experimental Parasitology 29: 59–65. Lund, E. E. 1972. Histomoniasis. In Diseases of Poultry, 6th ed., M. S. Hofstad, B. W. Calnek, C. F. Helmboldt, W. M. Reid, and H. W. Yoder, Jr. (eds). Iowa State University Press, Ames, IA, pp. 990–1006. Lund, E. E. 1977. The history of avian medicine in the United States IV. Some milestones in American research on poultry parasites. Avian Diseases 21:459–480. Lund, E. E., and A. M. Chute. 1970. Relative importance of young and mature turkeys and chickens in contaminating soil with Histomonas-bearing Heterakis eggs. Avian Diseases 14:342–348. Lund, E. E., and A. M. Chute. 1971a. Histomoniasis in the chukar partridge. Journal of Wildlife Management 35:307–315. Lund, E. E., and A. M. Chute. 1971b. Bobwhite, Colinus virginianus, as a host for Heterakis and Histomonas. Journal of Wildlife Diseases 7:70–75. Lund, E. E., and A. M. Chute. 1972a. Transfer of ten-day Heterakis gallinarum larvae: Effect on retention and development of the heterakids, and
liberation of Histomonas and Parahistomonas. Experimental Parasitology 31:361–369. Lund, E. E., and A. M. Chute. 1972b. Reciprocal responses of eight species of galliform birds and three parasites: Heterakis gallinarum, Histomonas meleagridis, and Parahistomonas wenrichi. Journal of Parasitology 58:940–945. Lund, E. E., and A. M. Chute. 1972c. Heterakis and Histomonas infections in young peafowl, compared to such infections in pheasants, chickens, and turkeys. Journal of Wildlife Diseases 8:352–358. Lund, E. E., and A. M. Chute. 1972d. The ring-necked pheasant (Phasianus colchicus torquatus) as a host for Heterakis gallinarum and Histomonas meleagridis. The American Midland Naturalist 87:1–7. Lund, E. E., and A. M. Chute. 1972e. Potential of young and mature guinea fowl in contaminating soil with Histomonas-bearing heterakid eggs. Avian Diseases 16:1079–1086. Lund, E. E., and A. M. Chute. 1973. Means of acquisition of Histomonas meleagridis by eggs of Heterakis gallinarum. Parasitology 66:335–342. Lund, E. E., and A. M. Chute. 1974. The reproductive potential of Heterakis gallinarum in various species of galliform birds: Implications for survival of H. gallinarum and Histomonas meleagridis to recent times. International Journal for Parasitology 4:455–461. Lund, E. E., and D. J. Ellis. 1967. The Japanese quail, Coturnix coturnix japonica, as a host for Heterakis and Histomonas. Laboratory Animal Care 17:110–113. Lund, E. E., E. E. Wehr, and D. J. Ellis. 1966. Earthworm transmission of Heterakis and Histomonas to turkeys and chickens. Journal of Parasitology 52:899–902. Lund, E. E., A. M. Chute, and M. E. L. Vernon. 1974. Experimental infections with Histomonas meleagridis and Heterakis gallinarum in ducks and geese. Journal of Parasitology 60:683–686. Lund, E. E., A. M. Chute, and G. C. Wilkins. 1975. The wild turkey as a host for Heterakis gallinarum and Histomonas meleagridis. Journal of Wildlife Diseases 11:376–381. McDougald, L. R., and R. B. Galloway. 1973. Blackhead disease: In vitro isolation of Histomonas meleagridis as a potentially useful diagnostic aid. Avian Diseases 17:847–850. McDougald, L. R., and M. F. Hansen. 1970. Histomonas meleagridis: Effect on plasma enzymes in chickens and turkeys. Experimental Parasitology 27:229–235. Mosby, H. S., and C. O. Handley. 1943. Diseases and pathological conditions. In The Wild Turkey in Virginia: Its Status, Life History and Management. Virginia Commission of Game and Inland Fisheries, Richmond, VA, pp. 138–146.
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Histomonas O’Roke, E. C. 1933. Some important problems in game bird pathology. Transactions of the American Game Conference 19:424–431. Prestwood, A. K., F. E. Kellogg, and G. L. Doster. 1973. Parasitism and disease among southeastern wild turkeys. In Wild Turkey Management: Current Problems and Programs, G. C. Sanderson and H. C. Schultz (eds). University of Missouri Press, Columbia, MO, pp. 159–167. Reid, W. M. 1967. Etiology and dissemination of the blackhead disease syndrome in turkeys and chickens. Experimental Parasitology 21:249–275. Roberts, H. A. 1956. Investigations of the frequency and kinds of disease and parasites found among wild turkeys in Pennsylvania. Semiannual Progress Report, Pennsylvania Game Commission, Harrisburg, PA. Schorr, L. F., W. R. Davidson, V. F. Nettles, J. E. Kennamer, P. Villegas, and H. W. Yoder, Jr. 1988. A survey of parasites and diseases of pen-raised wild turkeys. In Proceedings of the Annual Conference of the Southeastern Association of Fish and Wildlife Agencies 42:315–328. Sims, H. M. 1960. A preventive disease program for the chukar partridge. Modern Game Breeding 30(9):6–7. Snyder, R. L. 1953. Ability of spring released wild turkeys to survive and adapt to a natural environment. Quarterly Progress Report, Pennsylvania Game Commission, Harrisburg, PA, pp. 23–24.
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Stoddard, H. L. 1935. Wild turkey management. Transactions of the American Game Conference 21:326–333. Stoddard, H. L. 1936. Management of wild turkey. In Proceedings of the North American Wildlife Conference 1:352–356. Thomas, J. W. 1964. Diagnosed diseases and parasitism in Rio Grande wild turkeys. The Wilson Bulletin 76:292. Waters, C. V., III. 1992. An evaluation of the risk of commercial poultry litter as a source of histomoniasis for wild turkeys and other susceptible galliform species. M.S. Thesis, The University of Georgia, Athens, GA. Waters, C. V., L. D. Hall, W. R. Davidson, E. A. Rollor, and K. A. Lee. 1994. Status of commercial and noncommercial chickens as potential sources of histomoniasis among wild turkeys. The Wildlife Society Bulletin 22:43–49. Wilkins, D., and D. L. Lee. 1976. Qualitative and quantitative histochemical changes in the caecum and liver of turkeys infected with Histomonas meleagridis. Parasitology 72:51–63. Zeakes, S.J., M.F. Hansen, and R.J. Robel. 1981. Increased susceptibility of bobwhites (Colinus virginianus) to Histomonas meleagridis after exposure to Sevin insecticide. Avian Diseases 25:981–987.
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8 Eimeria Michael J. Yabsley normally do not cause disease can produce pathogenic effects and cause coccidiosis. In general, however, species of Eimeria rarely cause disease in free-ranging birds. Young birds or adults that are stressed or unhealthy are more likely to develop clinical coccidiosis.
INTRODUCTION The coccidia infect all classes of vertebrates and are a large and complex group of obligate intracellular parasites in the phylum Apicomplexa. Many of the coccidia are important medical and veterinary pathogens, but in this chapter, only the species of Eimeria that infect the intestines, kidneys, and liver of wild birds are discussed. Classification of the coccidia is based on morphologic characteristics, especially those of the environmentally-resistant sporulated oocyst. This stage is the only one that is passed in feces of the host and is therefore the stage most often observed. The vast majority of coccidia are described solely on the morphology of voided oocysts, and in many cases nothing more about their development in the host is known. The majority of avian species of Eimeria infect and develop within intestinal epithelial cells; however, some species of Eimeria develop in extraintestinal locations. Asexual stages, sexual stages, and the oocysts all develop within the cytoplasm or nucleus of infected cells. Compared with intestinal species of Eimeria, little is known regarding the host specificity and endogenous development of renal coccidia due to the difficulty in getting oocysts to sporulate to the infective stage. To date, virtually all extraintestinal species of Eimeria have been detected within infected renal epithelial cells. Exceptions are two species of Eimeria—Eimeria gruis and Eimeria reichenowi—that infect multiple organs in cranes (Chapter 9) and a single case of hepatic coccidiosis in a Magpie-lark (Grallina cyanoleuca). An important consideration when discussing coccidia is to differentiate infection and disease. Coccidian infections are frequently asymptomatic; thus, the correct terminology for infection is “infection with” or “coccidiasis.” Coccidiosis refers to infections resulting in clinical disease, but the term is often used inappropriately to indicate infection. Under certain varying circ*mstances, including age of host, high inoculation dose, stress, lack of previous infection, concurrent disease, or immunosuppression, species of Eimeria that
HISTORY Coccidiosis of domestic birds was recognized as early as the late 1800s (Railliet and Lucet 1890; Salmon 1899). Coccidium truncatum (=Eimeria truncata) from domestic geese was the first species of renal Eimeria to be named and described (Railliet and Lucet 1890). The life cycle of the first intestinal species of avian Eimeria was described in 1910 (Fantham 1910a). This species, detected in grouse in the UK, was named Eimeria avium and initially was thought to cause coccidiosis in numerous avian species (Fantham 1910b, 1911). It is now known that members of this genus are generally host specific and almost 200 species of avian Eimeria have been formally described. Numerous more have been reported but not described as distinct species.
INTESTINAL EIMERIA DISTRIBUTION AND HOST RANGE Intestinal species of avian Eimeria have been reported throughout the world. Intestinal coccidiosis is a common and economically important disease of domesticated fowl such as chickens, turkeys, and geese, whereas only sporadic cases of intestinal coccidiosis in wild birds have been reported. Infection with intestinal coccidia is probably ubiquitous among avian species; however, prevalence of infection with Eimeria varies among the avian orders (Table 8.1). To date, approximately 196 species of Eimeria have been formally described from 17 avian orders. However, uncharacterized species of Eimeria have been reported from numerous other wild avian
162 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Eimeria
Table 8.1. The approximate number of species of Eimeria that have been reported from avian hosts.
Host order Struthioniformes Casuariiformes Rheiformes Tinamiformes Sphenisciformes Gaviiformes Podicipediformes Procellariiformes Pelecaniformes Ciconiiformes Phoenicopteriformes Anseriformes Falconiformes Galliformes Gruiformes Charadriiformes Columbiformes Psittaciformes Cuculiformes Strigiformes Caprimulgiformes Apodiformes Coliiformes Trogoniformes Coraciiformes Piciformes Passeriformes Total
Number of avian species
Number of avian species with Eimeria (%)
Number of Eimeria spp.
Undescribed Coccidia reported
1 7 2 46 17 5 19 107 62 124 5 157 296 287 183 360 298 352 161 194 115 425 6 39 208 396 5,593 9,465
0 0 0 2 (4.3) 0 1 (20) 0 2 (1.9) 4 (6.5) 4 (3.2) 0 36 (22.9) 2 (0.7) 39 (13.6) 10 (5.5) 18 (5) 12 (4) 5 (1.4) 1 (0.6) 9 (4.6) 0 0 0 0 3 (1.4) 5 (1.3) 9 (0.16) 162 (1.7)
0 0 0 2 0 1 0 2 5 4 0 30 2 79 13 19 9 4 2 8 0 0 0 0 4 5 8 196
Yes Yes Yes No Yes No No Yes Yes Yes No Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes No Yes No Yes Yes Yes
Note: Host orders and species follow Dickinson (2003). hosts. The majority of species of Eimeria found in freeranging wild birds are reported from the orders Anseriformes and Galliformes. Likewise, coccidiosis is also reported most often from these two orders. Species of birds from some orders, such as Passeriformes, are primarily infected with Isospora and/or Atoxoplasma (Chapter 5). Although undescribed species of Eimeria have been detected in many orders of birds, there are many other orders where Eimeria likely occurs but has not been observed because few hosts have been examined. In other orders, for example Passeriformes, a considerable number of hosts have been examined for coccidia, but few Eimeria spp. have been detected. In general, Eimeria are highly host specific, but some species of Eimeria do infect multiple, often closely related, hosts. For example, Eimeria dispersa infects turkeys, chickens, Chukar (Alectoris chukar), Ring-
necked Pheasants (Phasianus colchicus), and Northern Bobwhite (Colinus virginianus) (Doran 1978), and Eimeria mulardi infects domestic ducks (Anas platyrhynchos), Muscovy Ducks (Cairina moschata), and their hybrid mule duck offspring (Sercy et al. 1996). Controlled experimental infections and/or genetic studies are needed to prove or disprove the occurrence of a single Eimeria species in multiple hosts, especially those hosts in distinct genera. ETIOLOGY Several genera of coccidia infect the intestinal epithelium of avian hosts, including Eimeria, Isospora, Atoxoplasma, Tyzzeria, Caryospora, Cryptosporidum, and Sarcocystis. The genus Eimeria is in the family Eimeriidae, order Eucoccidiorida, phylum
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Parasitic Diseases of Wild Birds
Figure 8.1. Major structural characteristics of the sporulated oocyst of a typical species of Eimeria. Drawing by S. E. J. Gibbs, CSIRO. Reproduced from Yabsley and Gibbs (2006), with permission of the Journal of Parasitology.
Apicomplexa and is one of the most common coccidia reported from birds. Eimeria can be differentiated from other genera by their characteristic oocysts— each contains four sporocysts, each of which contains two sporozoites (Figure 8.1). Other genera of avian coccidia have oocysts that differ in morphology. Members of the genera Isospora and Atoxoplasma (Chapter 5) have two sporocysts, each with four sporozoites. Members of the genera Cryptosporidium (Chapter 10) and Tyzzeria do not have sporocysts and contain four and eight sporozoites, respectively, within the oocyst. Members of the genus Caryospora have oocysts that possess a single sporocyst with eight sporozoites. Oocysts of species of Sarcocystis are extremely thin and free sporocysts containing four sporozoites are often the only forms detected in feces (Chapter 5). Molecular characterization of avian coccidia has shown that many of these early classifications have created polyphyletic genera and that morphologic characters are not sufficient to de-
termine relationships (e.g., avian Isospora are more closely related to Eimeria than to species of mammalian Isospora) (Carreno and Barta 1999; Yabsley and Gibbs 2006). EPIZOOTIOLOGY Coccidia in the genus Eimeria have direct life cycles (i.e., they are transmitted from one host to another without the aid of vectors or intermediate hosts). Development within the host includes both asexual and sexual stages that reside in epithelial cells. Unsporulated, noninfective oocysts passed in the feces of the host undergo sporulation in the environment to become infective. The first step of sporulation is the asexual process of sporogony by which sporocysts and sporozoites (infective stage) are produced from the germ ball within the environmentally resistant oocyst (Figure 8.1). This process is regulated by various environmental variables (oxygen, light, temperature, etc.)
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Eimeria that are parasite species dependent. In general, oocysts are extremely resistant and can even tolerate desiccation and freezing (Sathyanarayanan and Ortega 2006), although hard freezes or extreme heat can kill them (Parker and Jones 1990). Once ingested by an appropriate host, various chemical and physical cues cause the oocysts to rupture, releasing sporocysts which then rupture and release sporozoites. The sporozoites invade intestinal epithelial cells and transform into trophozoites. Trophozoites replicate asexually to form meronts, which later transform into merozoites by a process called merogony. These merozoites break out of the cell and enter other epithelial cells either to undergo additional rounds of merogony or to begin gametogony. The number of cycles of merogony and number of merozoites produced during each cycle differs among species of Eimeria. During gametogony, merozoites transform into macrogametocytes (female cell) or microgametocytes (male cell) (Figure 8.2). A macrogametocyte develops into a single macrogamete while a microgametocyte buds to form many flagellated microgametes. These microgametes exit the host cell and enter cells containing macrogametes, where fertilization occurs. A fertilized macrogamete develops an outer wall to become an oocyst (Figure 8.2), which is passed in the feces of the host. Coccidiosis is rare in free-ranging birds and is usually related to captive rearing, crowding, or stress. For
Figure 8.2. Developmental stages of Eimeria. Macrogametocytes (short arrow), microgametocytes (arrow head), and oocyst (long arrow) within villous epithelial cells. Flattened host cell nuclei are seen in some cells. Hematoxylin and eosin stain. Bar = 20 μm. Courtesy of A. E. Ellis, University of Georgia.
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several species of Eimeria, disease severity increases with ingestion of increasing numbers of oocysts in a dose-dependent fashion (Ruff and Wilkins 1987; Williams 2001). Young or na¨ıve birds exposed to high numbers of infective oocysts are most likely to exhibit disease. Anseriformes Species of Eimeria have been reported from only 22% of avian species in the order Anseriformes, but this group accounts for the second highest number of species of Eimeria reported from birds (Table 8.1). Several of these species of Eimeria have been reported to infect multiple avian hosts, but this needs to be confirmed by both experimental studies and application of molecular methods to identify morphologically similar cryptic species. An excellent review of coccidia of Anseriformes, including full morphologic descriptions and life history information, has been published (Gajadhar et al. 1983b). Several epizootics of intestinal coccidiosis caused by Eimeria aythyae have been reported from freeranging Lesser Scaup (Aythya affinis) in the US (Table 8.2; Bump 1937; Farr 1965; Windingstad et al. 1980; Southeastern Cooperative Wildlife Disease Study, unpublished data; US Geological Survey, National Wildlife Health Center, unpublished data). All outbreaks occurred during the spring. In Nebraska, at least 29% of Lesser Scaup died in each of three consecutive outbreaks during the springs of 1976–1978 (Windingstad et al. 1980). These outbreaks were associated with low water levels that could have crowded and stressed the birds. Attempts to transmit the infection to the Tufted Duck (Aythya fuligula) failed. An outbreak of intestinal and renal coccidiosis in 1993 (caused by an undescribed Eimeria sp. and Eimeria somateriae, respectively) resulted in the death of hundreds of Common Eider ducklings (Somateria mollissima) in Iceland (Table 8.2; Skirnisson 1997). This outbreak could have been caused by the washing of large amounts of mud into the sea near the nesting sites, which led to decreased foraging success and undernourishment and stress of ducklings (Skirnisson 1997). Galliformes Coccidiosis is a worldwide economically important disease of domestically raised fowl, primarily chickens and turkeys. In general, species of Galliformes harbor multiple species of coccidia (e.g., at least eight in chickens and seven in turkeys). Because of the economic importance of coccidiosis to domestic chickens, many excellent reviews of domestic chicken coccidia have been published (Allen and Fetterer 2002; Shirley
Lesser Scaup
Anseriformes
Aythya affinis
Host scientific name
166 Branta canadensis Eimeria fulva
Canada Goose
Eimeria bucephalae
Bucephala clangula
Common Goldeneye
Eimeria sp.
Eimeria aythyae
Eimeria spp.
Location of tissue stages
Clinical signs and/or lesions Mortality
Citations
Cytoplasm of small Sloughing of intestinal Extensive outbreaks Bump (1937), Farr intestine mucosa with (1965), epithelial cells extensive hemorrhage Windingstad et al. (1980), and U.S. Geological Survey, National Wildlife Health Center and Southeastern Cooperative Wildlife Disease Study, unpublished data Skirnisson (1997) Small localized Iceland Cytoplasm of small Focal necrosis of outbreaks in infected intestinal intestine epithelial cells cells malnourished ducklings; contributed to outbreak of renal coccidiosis Denmark Cytoplasm of small Thickening of intestinal Outbreaks in young Christiansen and birds Madsen (1948) wall with intestine epithelial cells hemorrhagic lesions and grayish-white foci; necrosis of subepithelial tissues Small localized Farr (1953) USA Cytoplasm of small Thickening of the outbreaks intestine intestinal wall and epithelial cells accumulation of greenish mucus in the intestinal lumen (continues)
USA
Locality
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Common Eider Somateria mollissima
Host common name
Host order
Table 8.2. Common pathogenic intestinal species of Eimeria.
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Galliformes
167 Eimeria angusta
Eimeria lettyae
Colinus virginianus
Northern Bobwhite
Greater Sage- Centrocercus urophasianus Grouse and and Bonasa Ruffed umbellus Grouse
Eimeria meleagrimitis
Meleagris gallopavo
USA
Cytoplasm of lower Dilated intestine and thickened wall, thick ileum, ceca, and creamy material, or rectum epithelial caseous casts cells USA Cytoplasm of upper Necrotic enteritis and mid-small intestine epithelial cells, lamina propria, or deeper tissues Listlessness, droopiness, USA Duodenum with and anorexia but no rare parasites in gross or the ileum and histopathologic cecum lesions noted Western USA Cytoplasm of Thickening of mucosa, cecum epithelial hemorrhage; diarrhea, cells depression, and weight loss
USA
Eimeria Meleagris dispersa gallopavo and Colinus virginianus Eimeria galMeleagris gallopavo lopavonis
Gajadhar et al. (1986)
Simon (1940), Historically, Honess and Post significant losses (1968), Barker of young sage et al. (1984), and grouse, no cases Connelly et al. documented since (2000) 1960s; deaths of ruffed grouse only in captivity (continues)
Not associated with Ruff (1985), and Ruff and Wilkins disease in wild birds (1987)
Not associated with Blakey (1932), and disease in wild Kozicky (1948) birds
Not associated with Blakey (1932), and Kozicky (1948) disease in wild birds
Not associated with Blakey (1932), and disease in wild Kozicky (1948) birds
Not associated with Blakey (1932), and disease in wild Kozicky (1948) birds
Small localized outbreaks
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Wild Turkey
Wild Turkey and Northern Bobwhite Wild Turkey
Reddening of mucosal surface and focal enteritis
Cytoplasm of lower Dilated intestine and thickened wall, thick ileum, ceca, and creamy material, or rectum epithelial caseous casts cells Cytoplasm of small Creamy, mucoid intestine and ceca enteritis epithelial cells
USA, Canada Nucleus of ileum and colon epithelial cells
USA
Eimeria stigmosa
Eimeria adenoeides
Canada Goose Branta canadensis and Lesser and Chen Snow caerulescens Goose caerulescens Wild Turkey Meleagris gallopavo
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Phasianus colchicus
Phasianus colchicus
Perdix spp.
Ring-necked Pheasant
Ring-necked Pheasant
Partridge species
Helmeted Numida Guineafowl meleagris
Phasianus colchicus
Host scientific name
Ring-necked Pheasant
Host common name
Psittaciformes
Budgerigar
Melopsittacus undulatus
Worldwide
Locality
Location of tissue stages
Clinical signs and/or lesions
Cytoplasm of Petechial hemorrhages cecum epithelial in heavy infections of cells captive birds Eimeria Worldwide Cytoplasm of small Ruffled feathers, phasiani intestine incoordination, epithelial cells mucoid diarrhea, and decreased weight gain Eimeria Worldwide Cytoplasm of small Anorexia and slight duodenalis intestine depression but no epithelial cells mortality in captive birds Eimeria Europe Cytoplasm of White cecal cores with procera cecum epithelial clotted blood, cells thickened intestinal walls Eimeria sp. Nigeria, Niger Cytoplasm of Intestines thickened and congested, intestine epithelial cells edematous, and/or hemorrhagic Worldwide Cytoplasm of small Intestines distended, Eimeria inflamed, and contain intestine labbeana petechial epithelial cells and Eimeria hemorrhages columbarum Yellow, pasty feces, Eimeria Worldwide Cytoplasm of dusingi duodenum enlarged duodenum epithelial cells
Eimeria colchici
Eimeria spp.
Citations
Not associated with Farr (1960), and Panigrahy et al. disease in wild birds (1981)
Ayeni et al. (1983) Not typically associated with disease in wild birds Not associated with Hunt and O’Grady disease in wild (1976) birds
Not associated with Goldov´a et al. disease in wild (2000) birds
Not associated with Jones (1966) disease in wild birds
Not associated with Jones (1966) disease in wild birds Not associated with Jones (1966), and disease in wild Trigg (1967) birds
Mortality
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Columba and Columbiformes Various pigeons and Streptopelia spp. doves
Host order
Table 8.2. (Continued)
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Eimeria et al. 2005). The eight species of Eimeria described from domestic chickens (Eimeria acervulina, Eimeria brunetti, Eimeria maxima, Eimeria mitis, Eimeria mivati, Eimeria necatrix, Eimeria praecox, and Eimeria tenella) each infect different regions of the lower gastrointestinal tract and cause variable presentations of disease (McDougald 2003). Not surprisingly, similar species of Eimeria have been found in the free-living ancestors of domestic chickens from Asia, the wild Red Junglefowl (Gallus gallus) and Ceylon Junglefowl (Gallus lafayetii); however, reports of disease in these hosts are lacking (Fernando and Remmler 1973a, b; Long et al. 1974). Numerous species of Eimeria—E. dispersa, Eimeria meleagrimitis, Eimeria gallopavonis, Eimeria meleagridis, Eimeria innocua, Eimeria subrotunda, Eimeria adenoeides, and an undescribed Eimeria sp.— have been reported from Wild Turkeys (Meleagris gallopavo). None of these species of Eimeria have been associated with disease in free-ranging birds, but four species have been associated with disease in captive birds (Table 8.2). In the southeastern US, Prestwood et al. (1971) reported that 50% of poults and 17% of juvenile and adult Wild Turkeys were positive for species of Eimeria. Despite the high prevalence, no lesions or clinical disease was observed in any birds. Similar findings were found for pen-raised Wild Turkeys; 66% were positive for Eimeria and mixed infections were common (Ruff et al. 1988a). At least 12 species of Eimeria have been reported from various species of quail. E. dispersa, the first coccidian described from a quail and also a common parasite of turkeys and pheasants, may cause disease in young Northern Bobwhite (Table 8.2). Eimeria lettyae from Northern Bobwhite has been reported from Pennsylvania and Florida (Ruff 1985) but probably occurs throughout the range of the Northern Bobwhite. E. lettyae appears to be host specific, as attempts to experimentally infect Japanese Quail (Coturnix japonica), Chukar, Ring-necked Pheasants, domestic turkeys, and chickens have failed. Although E. lettyae can be pathogenic for young Northern Bobwhite (Table 8.2) (Ruff and Wilkins 1987), coccidiosis does not appear to be a significant disease problem of wild free-ranging species of quail. Captive Japanese Quail are also susceptible to coccidiosis caused by Eimeria uzura, Eimeria tsunodai, and Eimeria taldykurganica (Ruff et al. 1984), although they have not been reported from wild Japanese Quail. Three-day-old quail were more susceptible to disease (100% mortality) than seventeen-day-old quail (8% mortality) when birds were inoculated with 105 oocysts of a mixture of these three species of Eimeria. Experimental inoculations of Northern Bobwhite, Chukar, Ring-necked Pheasants, domestic
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chickens, and domestic turkeys failed to produce infections (Ruff et al. 1984). Ten species of Eimeria have been described from pheasants, of which three—Eimeria colchici, Eimeria duodenalis, and Eimeria phasiani—are associated with severe disease in captive-bred Ring-necked Pheasants (Table 8.2) (Jones 1966). E. colchici is considered to be the most pathogenic. Chickens are experimentally susceptible when inoculated with large numbers of oocysts, but development in the cecal epithelial cells is limited and no clinical signs have been observed (Looszova et al. 2001). Eimeria angusta has been associated with cecal coccidiosis in both captive and free-ranging species of grouse (Table 8.2). Mortality has not been observed in free-ranging Greater Sage-Grouse (Centrocercus urophasianus) since the 1960s, presumably because numbers of grouse have declined, leading to decreased crowding and/or stress and reduced transmission of the parasite (Honess and Post 1968; Connelly et al. 2000). Gruiformes A total of 13 species of Eimeria have been reported from the order Gruiformes, 7 from cranes and 6 from coots and their relatives. All these species infect and develop in intestinal epithelial cells, but 2 species— Eimeria gruis and Eimeria reichenowi—can disseminate, develop extraintestinally, and cause severe disease (Chapter 9). Columbiformes Several species of Eimeria have been reported from free-ranging pigeons and doves (Table 8.1). None have been associated with disease among free-ranging birds; however, two species—Eimeria labbeana and Eimeria columbarum—have caused significant losses of captive pigeons and doves (Table 8.2) (Wages 1987; McDougald 2003). Psittaciformes Four species of Eimeria have been described from the Psittaciformes, but only Eimeria dunsingi has been associated with clinical disease in captive Budgerigars (Melopsittacus undulatus) (Farr 1960; Panigrahy et al. 1981). No disease was noted in naturally infected freeranging Budgerigars or Musk Lorikeets (Glossopsitta concinna) from Australia (Gartrell et al. 2000). Piciformes There is a single report of clinical coccidiosis and high mortality in captive Toco Toucan (Ramphastos toco) infected with an Eimeria sp., but clinical or
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epizootiological details of the infections were not reported (Martins et al. 2006). Eimeria forresteri was described from feces of captive Toco Toucans that did not exhibit clinical disease (Upton et al. 1984). Passeriformes Perching birds in the order Passeriformes are rarely infected with species of Eimeria. Some reports may be erroneous and could be pseudoparasites ingested with food items. There is one report of hemorrhagic enteritis associated with an Eimeria sp. in a Common Hill Mynah (Gracula religiosa) (Korbel and Kosters 1998). CLINICAL SIGNS Most birds infected with species of intestinal Eimeria do not exhibit any clinical signs because low-intensity infections destroy a limited number of epithelial cells that can be quickly replaced. Large numbers of cells are destroyed in infections of moderate to high intensity, leading to reduced food and water consumption, decreased intestinal absorption, and hemorrhage. Affected birds sometimes exhibit diarrhea tinged with blood or mucus, lack of appetite, emaciation, droopiness, loss of coordination, ruffled feathers, and decreased egg production (Hunt and O’Grady 1976; Wages 1987). Development of clinical signs depends on many factors including intensity of infection, species of parasite, and host factors such as age and health. For example, aberrant hosts may become infected, but replication and oocyst shedding by the parasite may be limited, resulting in no disease (Looszova et al. 2001; Revajov´a et al. 2006). Dosage can be an important factor that leads to development of disease. Ringnecked Pheasants experimentally infected with low numbers (10,000 oocysts) of E. phasiani developed only diarrhea, but when pheasants were inoculated with 100,000 oocysts, birds exhibited ruffled feathers, incoordination, mucoid diarrhea, and decreased weight gain (Trigg 1967). Similarly, experimental infections of Ring-necked Pheasants with variable numbers of E. colchici resulted in a dose-dependent disease (Norton 1967). Eight of ten birds exposed to low numbers (20,000 oocysts) survived infections, but none survived exposure to 80,000 or 320,000 oocysts. PATHOLOGY Pathologic changes vary widely, depending on the host and parasite species and severity of infection. Grossly, intestines may be ballooned, filled with mucus, hemorrhagic, and discolored (Figure 8.3). Sloughing of
Figure 8.3. Intestine, Common Eider (Somateria mollissima). Distinct light-colored areas (arrows) within the wall of the intestine. Courtesy of J. C. Franson, U.S. Geological Survey.
the intestinal mucosa is often observed in severe infections. Some species of Eimeria cause formation of white caseous cores in the ceca. Among Lesser Scaup with chronic infections, dry crusts have been reported on the mucosal surface of the intestine (Cole 1999). Among species of Galliformes, infections with Eimeria cause intestinal damage and changes in intestinal motility that may predispose the gut to infections with other pathogens such as Clostridium perfringens or Salmonella typhimurium. Cecal coccidiosis can exacerbate infections with Histomonas meleagridis (blackhead) (McDougald 2003). Developing meronts, gamonts, and oocysts of Eimeria can easily be observed within intestinal epithelial cells by microscopy (Figure 8.2). Most species of Eimeria develop in the cytoplasm of infected cells, but several species from geese develop within the epithelial cell nuclei. In clinically ill birds, extensive histopathologic lesions should be evident including host cell destruction and lymphocytic infiltration (Figure 8.4). However, lymphocytic infiltration may or may not be present depending on species of parasite and severity of infection.
DIAGNOSIS A diagnosis of coccidiosis is based on detection and identification of oocysts in feces along with clinical signs of the disease in live birds or characteristic lesions at necropsy. Feces collected from live birds or at necropsy can be examined directly for oocysts or by first concentrating oocysts by flotation using standard zinc sulfate or Sheather’s sugar. Diurnal periodicity in the shedding of oocysts has not been reported for
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Eimeria
Figure 8.4. Intestine, Lesser Scaup (Aythya affinis). Crypts are dilated and contain multiple coccidian stages with small amounts of necrotic cell debris. Crypt epithelium is attenuated or lost (arrow). Small numbers of inflammatory cells, primarily lymphocytes, are present in the surrounding lamina propria. Hematoxylin and eosin stain. Bar = 50 μm. Courtesy of R. W. Gerhold and A. E. Ellis, University of Georgia.
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IMMUNITY The bulk of knowledge related to development of host immunity to intestinal species of Eimeria is derived from studies of the domestic fowl. Immunity in chickens against coccidiosis is primarily T-cell mediated (reviewed by Lillehoj and Lillehoj 2000 and Yun et al. 2000). Anti-Eimeria IgM, IgY, and IgA antibodies are produced, but these antibodies are not effective at eliminating the parasite. Some level of protection develops in young birds that survive infection; however, this protection is specific to species of Eimeria; that is, it does not confer cross-protection against other species or strains of Eimeria. In some cases, birds may not develop complete immunity and the host becomes infected, but the infection will be less severe and result in development of fewer numbers of infective stages. For example, experimental infection of Northern Bobwhite with E. lettyae did not prevent reinfection (Ruff 1985). Lack of disease in wild birds is probably related to repeated exposures to low numbers of oocysts, which causes limited pathology and allows development of immunity.
PUBLIC HEALTH CONCERNS There are no known public health concerns regarding avian species of intestinal Eimeria. species of poultry Eimeria (Long 1982), but has been reported for an Eimeria sp. from the Red-legged Partridge (Alectoris rufa), which more commonly sheds oocysts in the late afternoon (Villan´ua et al. 2006). This phenomenon is also commonly observed among species of Isospora that occur in passerines (Barre and Troncy 1974; Brawner and Hill 1999; Misof 2004; M. J. Yabsley, unpublished data). It is unknown whether species of Eimeria from wild birds exhibit diurnal periodicity in oocyst shedding, but this should be considered when surveys of wild hosts are done. For identification of species, oocysts must be allowed to sporulate by placing feces in 1–3% (w/v) potassium dichromate, stored at room temperature, and examined daily for evidence of sporulation. Sporulation is facilitated by placing the fecal/potassium dichromate solution in a covered petri dish. Enough potassium dichromate solution must be used to prevent desiccation. Once sporulated, feces should be stored at 4◦ C to maintain morphologic characteristics. Because most infections are nonclinical, the finding of oocysts in a fecal sample does not indicate that a species of Eimeria is the cause of disease; significant pathologic lesions must be present at necropsy or other potential causes of the illness must be ruled out in live birds.
DOMESTIC ANIMAL CONCERNS Intestinal coccidiosis is a serious disease of many species of domesticated and captive wild birds and is associated with how birds are managed in captivity. Because of the presumed strict host specificity of the avian species of Eimeria, wild avian coccidia pose little threat to unrelated domesticated birds, but mixing of wild birds of different species is still discouraged. Wild birds and their domesticated counterparts (e.g., Wild Turkey and domesticated turkey) can be infected by the same species of coccidia; however, rarely would these wild birds pose any unusual threat to domesticated birds, as domesticated birds are commonly infected with the same repertoire of coccidia. For example, four species of Eimeria of Wild Turkeys— E. adenoeides, E. dispersa, E. gallopavonis, and E. meleagrimitis—are significant pathogens of domestic turkeys (McDougald 2003); however, these infections circulate within domestic turkeys without exposure to Wild Turkeys. Interestingly, several species of Eimeria have been associated with coccidiosis in domestic ducks (Eimeria saitamae) and domestic geese (Eimeria anseris, Eimeria kotlani, and Eimeria nocens) but have not been found to cause disease in
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free-ranging waterfowl (Levine 1953; Inoue 1967; Gajadhar et al. 1983a). Wild birds that are kept in zoological parks or other captive facilities are more likely to develop coccidiosis than their free-ranging counterparts (Panigrahy et al. 1981; Barker et al. 1984; Swayne et al. 1991; Giacomo et al. 1997; Novilla and Carpenter 2004). This problem can be compounded in facilities where large numbers of very closely related host species are housed together because closely related hosts may be susceptible to the same coccidian species. WILDLIFE POPULATION IMPACTS Outbreaks of intestinal coccidiosis are occasionally reported and cause mortality of free-ranging birds, but this condition does not appear to have a significant impact on wild populations. Reduced egg production and fertility have been reported in experimental studies of coccidiosis and could also occur in free-ranging birds. For example, adult Northern Bobwhite and Japanese Quail experimentally infected with species of Eimeria did not die, but egg production and fertility were reduced and maturation of males was delayed (Ruff et al. 1984, 1988b; Ruff and Wilkins 1987). Reductions in weight gain have not been reported in young wild birds with eimerian infections; however, this phenomenon is common in both domestic fowl and experimental studies and could be unrecognized in wild birds (Ruff et al. 1984). Field studies of the subclinical effects of coccidian infections are needed. TREATMENT AND CONTROL Much of what is known about treatment or control of avian coccidiosis is derived from studies concerning Eimeria of domestic fowl and birds in zoological collections. Historically, the use of anticoccidial feed or water supplements (e.g., amprolium and monensin) has been the primary method for controlling coccidiosis for poultry producers. In recent years, resistance has been documented against many of the common anticoccidial drugs (Martin et al. 1997). Poultry coccidia induce a strong immunity; therefore, vaccination has been investigated as an alternative to drugs for controlling disease. Early vaccines were made of live, wild-type, or attenuated parasites, but these vaccines were specific to species of Eimeria and, in some cases, specific to particular parasite strains. Wild-type vaccines work by providing a low-level of exposure, so uniform exposure among all birds is essential to preventing development of disease and for the development of protective immunity against future infections with large numbers of parasites (Shirley et al. 2005). Attenuated strains (those that have a reduced re-
productive capacity) are as immunogenic as wild-type strains but reduce the risk of clinical disease. New vaccines based on recombinant protective antigens are under development and may further increase the ability to vaccinate poultry safely and cheaply (Shirley et al. 2007). Currently, vaccines are parasite species/strain specific, difficult to produce, costly, and would not be feasible for use in wild birds. Captive wild birds that develop coccidiosis may respond to commercially available anticoccidial drugs, but studies on their effectiveness and safety are limited. In addition, each species of Eimeria may vary in susceptibility to the most commonly used drugs. For example, cecal coccidiosis (Eimeria colchici) in Ring-necked Pheasants is easily controlled with medicated feed containing zoalene or amprolium, but sulfaquinoxaline is ineffective (Norton 1967). Treatment of captive Rock Pigeons (Columba livia) infected with E. labbeana was successful with amprolium and sulfaquinoxaline (Hunt and O’Grady 1976). Sulfamethazine has also been used successfully to treat coccidiosis in captive Rock Pigeons and Budgerigars when added to drinking water (Panigrahy et al. 1981; McDougald 2003). Maintaining clean housing or raising birds on wire prevents buildup of infective oocysts and can decrease the risk of coccidiosis. Outbreaks of coccidiosis in free-ranging birds are difficult to treat because neither dosage nor regular dosing intervals can be easily controlled. Preventing crowding or stress may be more effective approaches to reducing or preventing outbreaks of coccidiosis in free-ranging birds.
MANAGEMENT IMPLICATIONS Infections of free-ranging birds with species of Eimeria are common, but coccidiosis among these birds in undisturbed habitat is rarely a significant problem. Outbreaks can occur when factors conspire to crowd or stress birds (e.g., breeding and loss of habitat). Most importantly, keeping wild birds in captivity can result in significant disease from coccidiosis.
RENAL EIMERIA ETIOLOGY Species of Eimeria are the primary cause of renal coccidiosis in birds. Although many host species harbor both renal and intestinal coccidia, the species of Eimeria that infect the kidneys are distinct and different from those that occur in intestinal tissues (Gajadhar et al. 1983a, b; Yabsley and Gibbs 2006).
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Eimeria EPIZOOTIOLOGY Similar to intestinal Eimeria, species of renal Eimeria have direct life cycles. Instead of developing in intestinal epithelial cells, sporozoites of renal species invade and develop in renal epithelial cells. Oocysts are passed unsporulated into the ureters and out of the cloaca. Oocysts shed in the feces of an infected host sporulate in the environment to become infective. As with the intestinal coccidia, sporulation is dependent on several factors including temperature, moisture, and levels of oxygen. Transmission of renal coccidia probably occurs in the fall as the prevalence in ducks is significantly higher in the fall than in spring in Saskatchewan, Canada, and Sweden (Walden 1963; Gajadhar et al. 1983a). Likewise, more geese were found infected in the fall than spring in Saskatchewan and Manitoba, Canada, and all reported outbreaks of renal coccidiosis of Double-crested Cormorants (Phalacrocorax auritus) have occurred between November and January (Gajadhar et al. 1983a; Clinchy and Barker 1994; Yabsley et al. 2002). As with intestinal coccidia, juvenile birds are more likely to be infected (Walden 1963; Nation and Wobeser 1977; Yabsley and Gibbs 2006). This may explain why prevalence increases during the fall when large numbers of na¨ıve young birds enter the population. HOST RANGE AND PREVALENCE Renal coccidia have been reported from numerous families of birds, but the greatest diversity of infected hosts is among species of Anseriformes. Pathogenic species of renal Eimeria and their associated traits are listed in Table 8.3. Procellariiformes In a study of the potential causative agents of “limey disease” (soiling of vent feathers by whitish excrement) in Short-tailed Shearwaters (Puffinus tenuirostris) from Tasmania, Munday et al. (1971) discovered renal coccidia (later described as Eimeria serventyi) in underweight chicks (Table 8.3; Pellerdy 1974). Renal coccidia have also been reported from Cory’s Shearwater (Calonectris diomedea), but no morphologic or pathologic information is available (Munday et al. 1971). Pelecaniformes From 1984 to present, more than 1,300 Double-crested Cormorants were reported to have died of renal coccidiosis during 11 mortality events (Table 8.3) (Yabsley et al. 2002; US Geological Survey, National Wildlife
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Health Center, unpublished data). Eimeria auritusi was described during an outbreak of coccidiosis in Georgia (Yabsley et al. 2002). While several substantial outbreaks of renal coccidiosis have been reported, a recent study in Georgia showed that 18 of 80 (23%) healthy double-crested cormorants were positive for renal coccidia identified as E. auritusi (Yabsley and Gibbs 2006). In general, low numbers of oocysts (<10) were detected in positive kidney samples and no gross lesions were noted, indicating that E. auritusi is not always pathogenic for double-crested cormorants. Anseriiformes Two species of renal Eimeria have been described from ducks: Eimeria somateriae from the common eider and long-tailed duck (Clangula hyemalis) and Eimeria boschadis from the mallard (Anas platyrhynchos) (Table 8.3) (Christiansen 1952; Walden 1963). However, the description of E. boschadis was based on unsporulated oocysts, making it an invalid description (Gajadhar et al. 1983a, b). Uncharacterized renal coccidia have been reported from virtually all species of ducks where substantial numbers of individuals have been examined. Many of these unidentified species of Eimeria differed morphologically from recognized species, indicating that many new species still need to be described. In a survey of 336 ducks of 12 species from Saskatchewan, Canada, 151 (45%) were infected with renal coccidia, most of which are undescribed species (Gajadhar et al. 1983a). Renal coccidia were detected in 11 of the species of ducks that were examined including the American Widgeon (Anas americana), Blue-winged Teal (Anas discors), Eurasian Teal (Anas crecca), Gadwall (Anas strepera), Mallard, Northern Pintail (Anas acuta), Northern Shoveler (Anas clypeata), Canvasback (Aythya valisineria), Common Goldeneye (Bucephala clangula), Lesser Scaup, and Redhead (Aythya americana). Eleven Bufflehead (Bucephala albeola) were sampled and were found to be negative. Renal coccidia were detected in one of six Red-breasted Mergansers (Mergus serrator) from Florida (Forrester and Spalding 2003). Renal coccidia are common in several species of geese and have been reported worldwide from Greater and Lesser Snow Geese (Chen caerulescens atlantica and Chen caerulescens caerulescens), Ross’ Geese (Chen rossii), Graylag Geese (Anser anser), domestic geese, and multiple subspecies of Canada Geese (Branta canadensis) (Gajadhar et al. 1983a, b). To date, E. truncata is the only species formally described from geese (Table 8.3). It is likely that multiple species of renal Eimeria infect geese, but controlled experimental infections and/or molecular studies are
USA, Canada
Finland
Eimeria auritusi Eimeria somateriae Eimeria truncata Eimeria truncata Eimeria truncata
Phalacrocorax auritus Somateria mollissima Branta canadensis Chen caerulescens caerulescens Anser anser
Double-crested Cormorant Common Eider
Canada Goose
Lesser Snow Goose
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Graylag Goose
Denmark, Iceland, Sweden, and Scotland USA
USA
Tasmania
Small outbreaks; many infections are subclinical
Few numbers of birds; many infections are subclinical Occasionally in malnourished geese
Outbreaks associated with concurrent parasitic and fungal infections Loss of young underweight birds; morbidity as high as 1–5%; can cause high mortality Small- to medium-sized outbreaks Occasional deaths; a few ep*rnitics
Mortality
Oksanen (1994)
Gomis et al. (1996)
Christiansen (1952), Persson et al. (1974), Mendenhall (1976), and Skirnisson (1997) Farr (1954), and Tuggle and Crites (1984)
Yabsley et al. (2002)
Munday et al. (1971), and Pellerdy (1974)
Obendorf and McColl (1980)
Citations
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Anseriformes
Pelecaniformes
Eimeria serventyi
Puffinus tenuirostris
Short-tailed Shearwater
Procellariiformes
Victoria, Australia
Eimeria sp.
Little Penguin
Sphenisciformes
Eudyptula minor
Locality
Eimeria spp.
Host common name
Host order
Host scientific name
Table 8.3. Common pathogenic species of renal Eimeria.
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needed to establish the true number. Because of the difficulty in getting oocysts of renal coccidia to sporulate under typical conditions that are successful for intestinal Eimeria, experimental infections are difficult, but Gajadhar et al. (1982) have successfully studied the life cycle of a renal Eimeria in Lesser Snow Geese. Mortality events caused by E. truncata have been reported in free-ranging Canada Geese, Lesser Snow Geese, and Graylag Geese (Table 8.3) (Farr 1954; Tuggle and Crites 1984; Oksanen 1994; Gomis et al. 1996). The prevalence of renal coccidia from 309 Canada Geese along the Mississippi flyway was almost 7% (Tuggle and Crites 1984). The majority of infections were regarded as subclinical, but renal coccidiosis was determined as the cause of death for one goose found dead during the study.
tralia (Table 8.3) (Obendorf and McColl 1980). Threequarters of the 48 penguins submitted for necropsy between 1977 and 1978 were in poor body condition and 23% had gross and histopathologic lesions associated with renal coccidiosis.
Charadriiformes Eimeria wobeseri and Eimeria goelandi were described from the European Herring Gull (Larus argentatus) by Gajadhar and Leighton (1988). Neither species was associated with disease. Interestingly, E. goelandi sporulated endogenously, which is unusual for members of the genus Eimeria. Morphologically, E. goelandi is correctly described as a species of Eimeria as it has four sporocysts, each containing two sporozoites. A single coccidian species, Eimeria fraterculae, has been described from the Atlantic Puffin (Fratercula arctica) from Newfoundland, Canada (Leighton and Gajadhar 1986). In the original description, 7 of 50 nestling puffins were infected, but none exhibited any clinical signs. Renal coccidia have been reported from both captive-raised and free-ranging American Woodco*cks (Scolopax minor) from numerous states in the eastern US and Ontario, Canada (Locke et al. 1965; Pursglove 1973). Prevalence in free-ranging and captive American Woodco*cks was 28 and 19% (72/265 and 6/31), respectively (Pursglove 1973). Clinical disease was not apparent and kidneys of many infected birds appeared grossly normal with the exception of occasional whitish streaks. Developmental stages of the parasites were observed in collecting tubules in histological sections. These tubules were markedly enlarged with cell nuclei displaced toward the basem*nt membrane. No fresh material was available in either study to attempt sporulation and description of oocysts.
CLINICAL SIGNS AND PATHOLOGY As with intestinal Eimeria species, most reports of species of renal Eimeria come from surveys of presumably healthy birds and clinical signs are rarely observed. If birds develop disease due to renal coccidia, they are often found dead. Clinical signs observed in experimentally infected or domestic birds include diarrhea, weakness, ataxia, difficulty in flying, depression, lack of appetite, and emaciation (Gajadhar et al. 1983b; McDougald 2003). The majority of infections with renal coccidia result in few or no gross lesions. Birds may be emaciated, but if death occurred quickly, they may still be in good physiological condition (Obendorf and McColl 1980; Yabsley et al. 2002). In low-intensity infections, kidneys may be slightly enlarged and mottled with rare white streaks or nodules. In severe, very intense infections, kidneys are pale, grossly enlarged, friable, and mottled with white streaks and nodules (Figure 8.5). Microscopic lesions consist primarily of dilation of infected tubules with distortion of normal architecture and associated inflammation (Leighton and Gajadhar 1986; Yabsley et al. 2002). Infected cells are swollen and contain numerous developing intracellular parasites (Figure 8.6). Large numbers of oocysts can obstruct tubules (Figure 8.6) and may be found in ureters. Rarely, oocysts may enter the bloodstream and become lodged in organs such as the lungs (Yabsley et al. 2002). Affected tubules are often surrounded by infiltrates of macrophages, lymphocytes, plasma cells, and heterophils. Necrosis of infected cells and tubules is often evident (Thompson and Wright 1978; Gajadhar et al. 1983b; Yabsley et al. 2002). In mild infections, lesions are limited to few infected tubules and are surrounded by normal kidney tissue, but in severe cases, most tubules are infected with parasites, resulting in little normal kidney tissue being present.
Sphenisciformes Renal coccidiosis, together with extreme gastric parasitism and aspergillosis, was associated with mortality of Little Penguins (Eudyptula minor) in Victoria, Aus-
Apterygiformes An unclassified coccidian species has been reported to cause renal coccidiosis in a captive-bred, 1-monthold North Island Brown Kiwi (Apteryx mantelli) (Thompson and Wright 1978). The bird was depressed and at necropsy had pale kidneys. Three other clinically normal kiwis at the same location were found to be passing oocysts in their feces. None sporulated, so specific identification was not possible.
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Figure 8.5. Kidney, Double-crested Cormorants (Phalacrocorax auritus). (a) Normal size and color; (b) enlarged kidneys with pale areas from a bird infected with renal coccidiosis. Courtesy of J. C. Franson, U.S. Geological Survey.
DIAGNOSIS In the cases of suspected fatal renal coccidiosis, gross pathology is highly suggestive; however, lesions in fatal cases must be sufficient to impede kidney function, and other causes of death must be ruled out. For confirmation, oocysts can be observed in direct smears of kidney tissue or parasites can be observed in histological sections of kidney tissue. Many hosts infected with renal coccidia have only developing parasites in a limited number of renal tubules; therefore, morbidity and mortality in most infected hosts is low. For detection of oocysts in lowintensity infections from asymptomatic birds, kidney tissues should be placed in 2% (w/v) potassium dichromate and disrupted with a blender or tissue macerator. Care must be taken during dissection to avoid contamination of samples with intestinal contents that may contain unrelated species of coccidia. The tissue can be filtered through a layer of cheese cloth and the filtrate, containing oocysts and small pieces of kidney tissue, centrifuged so that the resulting pellet can be examined for oocysts by direct examination or standard zinc sulfate or Sheather’s sugar flotation. Asymptomatic infections can also be detected during histopathologic examination of kidney tissue; developing meronts, gamonts, and oocysts are easily observed in the cytoplasm of tubular epithelial cells.
Figure 8.6. Kidney, Double-crested Cormorant (Phalacrocorax auritus). (a) Renal tubular epithelial cells distended by oocysts of Eimeria auritusi in their cytoplasm. (b) Multiple developing gamonts per infected epithelial cell. Hematoxylin and eosin stain. Bar = 25 μm. Reproduced from Yabsley et al. (2002), with permission of the Journal of Parasitology.
For specific diagnosis, oocysts must be sporulated in potassium dichromate as described for intestinal Eimeria. To date, researchers have had limited success in sporulation of many samples of renal coccidia. This has hampered detailed studies of oocyst morphology and made experimental studies difficult. Other renal coccidia reported from avian species include disseminated toxoplasmosis (Toxoplasma gondii) in Wild Turkeys (Chapter 11) (Quist et al. 1995), renal cryptosporidiosis in many avian species (Chapter 10) (Gardiner and Imes 1984; Randall 1986; Trampel et al. 2000), and a probable case of Klossiella from a Great Horned Owl (Bubo virginianus) (Helmboldt 1967).
IMMUNITY Domestic geese develop immunity to reinfection with E. truncata (McDougald 2003). Nothing is known about the development of immunity to renal Eimeria in naturally infected free-ranging birds. Similar to intestinal coccidia, infection of young birds with low levels of renal coccidia probably results in mild infections that may protect birds from future heavy infections and subsequent clinical disease.
PUBLIC HEALTH CONCERNS There are no known public health concerns associated with renal coccidia.
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Eimeria DOMESTIC ANIMAL HEALTH CONCERNS Eimeria truncata is a significant pathogen of domestic geese throughout the world (Gajadhar et al. 1983b). There is evidence to suggest that E. truncata occurs in multiple species of wild geese, but controlled experimental infections and/or molecular studies are needed to confirm this finding. Wild geese probably do not pose a threat to domestic geese because the pathogen is frequently present in domestic geese populations that do not have exposure to wild geese.
WILDLIFE POPULATION IMPACTS Small- to medium-sized outbreaks of renal coccidiosis involving up to several hundred birds have been reported in several species of free-ranging birds, but these outbreaks do not appear to have had a significant impact on wild populations.
TREATMENT AND CONTROL The efficacy and safety of common anticoccidial drugs for treating renal coccidiosis in wild birds is unknown. Numerous coccidiostats and anticoccidial drugs commonly used in domestic chickens (e.g., amprolium, dulfaquinoxaline, clopidol, zoalene, narasin, nicarbazin, robenidin, and salinomycin) are tolerated by domestic geese and are used to control both intestinal and renal coccidiosis. However, current therapy in domestic birds is aimed at reducing clinical signs and numbers and transmission of parasites rather than complete elimination of the parasites. Control of transmission in the wild is not feasible, based on difficulties in administering drugs and the widespread occurrence of asymptomatic infections in wild birds.
MANAGEMENT IMPLICATIONS Coccidian infections in free-ranging birds in undisturbed habitats are rarely a significant problem because birds harbor asymptomatic infections that rarely cause mortality. Any activity that concentrates birds (e.g., extreme weather conditions or habitat degradation) can lead to outbreaks of coccidiosis. Renal coccidiosis may become a problem when susceptible wild avian species are kept in captivity.
HEPATIC EIMERIA One species of Eimeria has been reported to develop in the cytoplasm of bile duct epithelial cells of the Magpie-lark from Australia (Reece 1989). The single infected bird was extremely emaciated and the liver was enlarged with many white foci. Oocysts of this
177
species of Eimeria sporulated within the liver tissue and were passed in the feces as fully developed oocysts. Endogenous sporulation also occurs in species of renal Eimeria from gulls. While Eimeria grallinida was proposed as the name for the parasite, a detailed description was not given. LITERATURE CITED Allen, P. C., and R. H. Fetterer. 2002. Recent advances in biology and immunobiology of Eimeria species and in diagnosis and control of infection with these coccidian parasites of poultry. Clinical Microbiology Reviews 15:58–65. Ayeni, J. S., O. O. Dipeolu, and A. N. Okaeme. 1983. Parasitic infections of the grey-breasted helmet guinea-fowl (Numida meleagris galeata) in Nigeria. Veterinary Parasitology 12:59–63. Barker, I. K., A. Garbutt, and A. L. Middleton. 1984. Endogenous development and pathogenicity of Eimeria angusta in the ruffed grouse, Bonasa umbellus. Journal of Wildlife Diseases 20:100–107. Barre, N., and P. M. Troncy. 1974. Note on a coccidia of some Ploceidae in Chad: Isospora xerophila n. sp. Zeitschrift f¨ur Parasitenkunde 44:139–147. Blakey, H. L. 1932. Biological problems confronting the artificial propagation of wild turkeys in Missouri. Transactions of the 19th North American Game Conference 19:337–343. Brawner, W. R., III, and G. E. Hill. 1999. Temporal variation in shedding of coccidial oocysts: Implications for sexual-selection studies. Canadian Journal of Zoology 77:347–350. Bump, G. 1937. Annual Report to the Legislature. New York Conservation Department 1936:288–335. Carreno, R. A., and J. R. Barta. 1999. An Eimeriid origin of isosporoid coccidia with Stieda bodies as shown by phylogenetic analysis of small subunit ribosomal RNA gene sequences. Journal of Parasitology 85:77–83. Christiansen, M. 1952. Nyrecoccidiose hos vildtlevend andefugle (Anseriformes). Nordisk Veterinaermedicin 4:1173–1191. Christiansen, M., and H. Madsen. 1948. Eimeria bucephalae n. sp. pathogenic in goldeneye in Denmark. Danish Review Game Biology 1:62–73. Clinchy, M., and I. K. Barker. 1994. Dynamics of parasitic infections at four sites within lesser snow geese (Chen caerulescens caerulescens) from the breeding colony at La Perouse Bay, Manitoba, Canada. Journal of Parasitology 80:663–666. Cole, R. A. 1999. Intestinal coccidiosis. In Field Manual of Wildlife Disease General Field Procedures and Diseases of Birds, M. Friend, and J. C. Franson (eds). U.S. Geological Survey Information and Technology Report 1999–2001, pp. 207–213.
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Connelly, J. W., M. A. Schroeder, A. R. Sands, and C. E. Braun. 2000. Guidelines to manage sage grouse populations and their habitats. Wildlife Society Bulletin 28:967–985. Dickinson, D. C. (ed.). 2003. The Howard and Moore Complete checklist of the Birds of the World, 3rd ed. Princeton University Press, Princeton, NJ. Doran, D. J. 1978. The life cycle of Eimeria dispersa Tyzzer 1929 from the turkey in gallinaceous birds. Journal of Parasitology 64:882–885. Fantham, H. B. 1910a. The morphology and life history of Eimeria (coccidium) avium: A sporozoon causing a fatal disease among young grouse. Proceedings of the Zoological Society of London 3:672–691. Fantham, H. B. 1910b. Experimental studies on avian coccidiosis, especially in relation to young grouse, fowls, and pigeons. Proceedings of the Zoological Society of London 3:708–721. Fantham, H. B. 1911. Coccidiosis in British game birds and poultry. Journal of Economic Biology 6:75–96. Farr, M. M. 1953. Three new species of coccidia from the Canada goose (Branta canadensis). Journal of the Washington Academy of Sciences 43:336–340. Farr, M. M. 1954. Renal coccidiosis of Canada geese. Journal of Parasitology 40:46. Farr, M. M. 1960. Eimeria dunsingi n. sp. (Protozoa: Eimeriidae) from the intestine of a parakeet, Melopsittacus undulatus (Shaw). Libro Homenaje al Dr. Eduardo Caballerol y lal. Jubileo 1930-1960. pp. 31–35. Farr, M. M. 1965. Coccidiosis of the lesser scaup duck, Aythya affinis (Eyton, 1938) with a description of a new species, Eimeria aythyae. Proceedings of the Helminthological Society of Washington 32:236–238. Fernando, M. A., and O. Remmler. 1973a. Eimeria diminuta sp. n. from the Ceylon jungle fowl, Gallus lafayettei. Journal of Protozoology 20:357. Fernando, M. A., and O. Remmler. 1973b. Four new species of Eimeria and one of Tyzzeria from the Ceylon jungle fowl Gallus lafayettei. Journal of Protozoology 20:43–45. Forrester, D. J., and M. G. Spalding. 2003. Parasites and diseases of wild birds in Florida. University Press of Florida, Gainesville, FL. 1152 pp. Gajadhar, A. A., and F. A. Leighton. 1988. Eimeria wobeseri n. sp. and Eimeria goelandi n. sp. (Protozoa: Apicomplexa) in the kidneys of herring gulls (Larus argentatus). Journal of Wildlife Diseases 24:538–546. Gajadhar, A. A., R. J. Cawthorn, and D. J. Rainnie. 1982. Experimental studies on the life cycle of a renal coccidium of lesser snow geese (Anser c. caerulescens). Canadian Journal of Zoology 60:2085–2092. Gajadhar, A. A., R. J. Cawthorn, G. A. Wobeser, and P. H. G. Stockdale. 1983a. Prevalence of renal coccidia
in wild waterfowl in Saskatchewan. Canadian Journal of Zoology 61:2631–2633. Gajadhar, A. A., G. Wobeser, and P. H. G. Stockdale. 1983b. Coccidia of domestic and wild waterfowl. Canadian Journal of Zoology 61:1–24. Gajadhar, A. A., D. J. Rainnie, and R. J. Cawthorn. 1986. Description of the goose coccidium Eimeria stigmosa (Klimes, 1963), with evidence of intranuclear development. Journal of Parasitology 72:588–594. Gardiner, C. H., and G. D. Imes, Jr. 1984. Cryptosporidium sp. in the kidneys of a black-throated finch. Journal of American Veterinary Medical Association 185:1401–1402. Gartrell, B. D., P. O’Donoghue, and S. R. Raidal. 2000. Eimeria dunsingi in free living musk lorikeets (Glossopsitta concinna). Australian Veterinary Journal 78:717–718. Giacomo, R., P. Stefania, T. Ennio, V. C. Giorgina, B. Giovanni, and R. Giacomo. 1997. Mortality in black siskins (Carduelis atrata) with systemic coccidiosis. Journal of Wildlife Diseases 33:152–157. Goldov´a, M., V. Letkov´a, and G. Csizsm´arov´a. 2000. Life cycle of Eimeria procerca in experimentally infected grey partridges (Perdix perdix). Veterinary Parasitology 90:255–263. Gomis, S., A. B. Didiuk, J. Neufeld, and G. Wobeser. 1996. Renal coccidiosis and other parasitologic conditions in lesser snow goose goslings at Tha-anne River, west coast Hudson Bay. Journal of Wildlife Diseases 32:498–504. Helmboldt, C. F. 1967. An unidentified protozoan parasite in the kidney of the great-horned owl (Bubo virginianus). Bulletin of the Wildlife Disease Association 3:23–25. Honess, R. F., and G. Post. 1968. History of an epizootic in sage grouse. Part I. Sage grouse coccidiosis. Science Monograph No. 14, Agricultural Experimental Station, University of Wyoming, Laramie, WI, 28 pp. Hunt, S., and J. O’Grady. 1976. Coccidiosis in pigeons due to Eimeria labbeana. Australian Veterinary Journal 52:390. Inoue, I. 1967. Eimeria saitamae n. sp.: A new cause of coccidiosis in domestic ducks (Anas platlyrhyncha var. domestica). Japanese Journal of Veterinary Science 29:209–205. Jones, M. B. 1966. Survey of game bird diseases. Report of the Game Research Association 5:34. Korbel, R., and J Kosters. 1998. Beos. In Krankheiten der Heimtiere, 4th ed., K. Gabrish and P. Zwart (eds). Schlutersche, Hannover, Germany, pp. 397–428. Kozicky, E. L. 1948. Some protozoan parasites of the eastern wild turkey in Pennsylvania. The Journal of Wildlife Management 12:263–266.
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Eimeria Leighton, F. A., and A. A. Gajadhar. 1986. Eimeria fraterculae n. sp. in the kidneys of Atlantic puffins (Fratercula arctica) from Newfoundland, Canada: Species description and lesions. Journal of Wildlife Diseases 22:520–526. Levine, N. D. 1953. A review of coccidian from the avian orders Galliformes, Anseriformes, and Charadriiformes, with descriptions of three new species. American Midland Naturalist 49:696–719. Lillehoj, H. S., and E. P. Lillehoj. 2000. Avian coccidiosis. A review of acquired intestinal immunity and vaccination strategies. Avian Diseases 44:408–425. Locke, L. N., W. H. Stickel, and S. A. Geis. 1965. Some diseases and parasites of captive woodco*cks. Journal of Wildlife Management 29:156–161. Long, P. L. 1982. The Biology of the Coccidia. University Park Press, Baltimore, MD, 502 pp. Long, P. L., M. A. Fernando, and O. Remmler. 1974. Experimental infections of the domestic fowl with a variant of Eimeria praecox from the Ceylon jungle fowl. Parasitology 69:1–9. Looszova, A., V. Revajova, M. Levkut, and J. Pistl. 2001. Pathogenesis of Eimeria colchici in the intestine of chickens and the related immune response. Acta Veterinaria Brno 70:191–196. Martin, A. G., H. D. Danforth, J. R. Barta, and M. A. Fernando. 1997. Analysis of immunological cross-protection and sensitivities to anticoccidial drugs among five geographical and temporal strains of Eimeria maxima. International Journal for Parasitology 27:527–533. Martins, N. R. S., A. C. Horta, A. M. Siqueira, S. Q. Lopes, J. S. Resende, M. A. Jorge, R. A. Assis, N. E. Martins, A. A. Fernandes, P. R. Barrios, T. J. R. Costa, and L. M. C. Guimar˜aes. 2006. Macrorhabdus ornithogaster in ostrich, rhea, canary, zebra finch, free range chicken, turkey, guinea-fowl, columbina pigeon, toucan, chuckar partridge and experimental infection in chicken, Japanese quail and mice. Arquivo Brasileiro de Medicina Veterin´aria e Zootecnia 58:291–298. McDougald, L. R. 2003. Coccidiosis. In Diseases of Poultry, 11th ed., Y. M. Saif, H. G. Barnes, J. R. Glisson, A. M. Fadly, L. R. McDonald, and D. E. Swayne (eds). Iowa State Press, Ames, IA, pp. 974–990. Mendenhall, V. 1976. Survival and causes of mortality in eider ducklings on the Ythan Estuary, Aberdeenshire, Scotland. Wildfowl 27:160. Misof, K. 2004. Diurnal cycle of Isospora spp. oocyst shedding in Eurasian blackbirds (Turdus merula). Canadian Journal of Zoology 82:764–768. Munday, B. L., R. W. Mason, R. J. H. Wells, and J. H. Arundel. 1971. Further studies on “Limey-disease” of
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Tasmanian mutton birds (Puffinus tenuirostris). Journal of Wildlife Diseases 7:126–129. Nation, P. N., and G. Wobeser. 1977. Renal coccidiosis in wild ducks in Saskatchewan. Journal of Wildlife Diseases 13:370–375. Norton, C. C. 1967. Eimeria colchici sp. nov. (Protozoa: Eimeriidae), the cause of cecal coccidiosis in English covert pheasants. Journal of Protozoology 14:772. Novilla, M. N., and J. W. Carpenter. 2004. Pathology and pathogenesis of disseminated visceral coccidiosis in cranes. Avian Pathology 33:275–280. Obendorf, D. L., and K. McColl. 1980. Mortality in little penguins (Eudyptula minor) along the coast of Victoria, Australia. Journal of Wildlife Diseases 16:251–259. Oksanen, A. 1994. Mortality associated with renal coccidiosis in juvenile wild greylag geese (Anser anser anser). Journal of Wildlife Diseases 30:554–556. Panigrahy, B., J. J. Mathewson, C. F. Hall, and L. C. Grumbles. 1981. Unusual disease conditions in pet and aviary birds. Journal of American Veterinary Medical Association 178:394–395. Parker, R. J., and G. W. Jones. 1990. Destruction of bovine coccidial oocysts in simulated cattle yards by dry tropical winter weather. Veterinary Parasitology 35:269–272. Pellerdy, L. 1974. Coccidia and Coccidiosis. Verlag Paul Parey, Berlin and Hamburg, Germany, 959 pp. Persson, L., K. Borg, and H. Falt. 1974. On the occurrence of endoparasites in eider ducks in Sweden. Viltrevy 9:1–24. Prestwood, A. K., F. E. Kellog, and G. L. Doster. 1971. Coccidia in eastern wild turkeys of the southeastern United States. Journal of Parasitology 57:189–190. Pursglove, S. R., Jr. 1973. Some Parasites and Diseases of the American Woodco*ck, Philohela minor (Gmelin). Ph.D. Dissertation, The University of Georgia, 221 pp. Quist, C. F., J. P. Dubey, M. P. Luttrell, and W. R. Davidson. 1995. Toxoplasmosis in wild turkeys: A case report and serologic survey. Journal of Wildlife Diseases 31:255–258. Railliet, A., and A. Lucet. 1890. Une nouvelle maladie parasitaire de l’oie domestique, determine par des coccidies. Comptes rendus des s´eances et m´emoires do la Soci´et´e de biologie. Paris 42:292–294. Randall, C. J. 1986. Renal and nasal cryptosporidiosis in a junglefowl (Gallus sonneratii). The Veterinary Record 119:130–131. Reece, R. L. 1989. Hepatic coccidiosis (Eimeria sp.) in a wild magpie-lark (Grallina cyanoleuca). Avian Pathology 18:357–362. Revajov´a, V., A. Lo´oszov´a, M. Goldov´a, M. Zibr´ın, R. Herich, and M. Levkut. 2006. Morphological study of partridge Eimeria procera development in the foreign
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host—Leghorn chicks (Gallus gallus). Journal of Protozoology Research 16:26–32. Ruff, M. D. 1985. Life cycle and biology of Eimeria lettyae sp. n. (Protozoa: Eimeriidae) from the northern bobwhite, Colinus virginianus (L.). Journal of Wildlife Diseases 21:361–370. Ruff, M. D., and G. C. Wilkins. 1987. Pathogenicity of Eimeria lettyae Ruff, 1985 in the northern bobwhite (Colinus virginianus L.). Journal of Wildlife Diseases 23:121–126. Ruff, M. D., J. M. fa*gan, and J. W. Dick. 1984. Pathogenicity of coccidia in Japanese quail (Coturnix coturnix japonica). Poultry Science 63:55–60. Ruff, M. D., L. Schorr, W. R. Davidson, and V. F. Nettles. 1988a. Prevalence and identity of coccidia in pen-raised wild turkeys. Journal of Wildlife Diseases 24:711–714. Ruff, M. D., M. A. Abdel Nabi, R. N. Clarke, M. Mobarak, and M. A. Ottinger. 1988b. Effect of coccidiosis of reproductive maturation of male Japanese quail. Avian Diseases 32:41–45. Salmon, D. E. 1899. The Diseases of Poultry. The Feather Library, George E. Howard and Co., Washington, DC. Sathyanarayanan, L., and Y. Ortega. 2006. Effects of temperature and different food matrices on Cyclospora cayetanensis oocyst sporulation. Journal of Parasitology 92:218–222. Sercy, O., K. Nie, A. Pascalon, G. Fort, and P. Yvore. 1996. Receptivity and susceptibility of the domestic duck (Anas platyrhynchos), the Muscovy duck (Cairina moschata), and their hybrid, the mule duck, to an experimental infection by Eimeria mulardi. Avian Diseases 40:23–27. Shirley, M. W., A. L. Smith, and F. M. Tomley. 2005. The biology of avian Eimeria with an emphasis on their control by vaccination. Advances in Parasitology 60:285–330. Shirley, M. W., A. L. Smith, and D. P. Blake. 2007. Challenges in the successful control of avian coccidia. Vaccine 25:5540–5547. Simon, F. 1940. The parasites of sage grouse, Centrocercus urophasianus. University of Wyoming Publications 7:77–100. Skirnisson, K. 1997. Mortality associated with renal and intestinal coccidiosis in juvenile eiders in Iceland. Parassitologia 39:325–330. Swayne, D. E., D. Getzy, R. D. Slemons, C. Bocetti, and L. Kramer. 1991. Coccidiosis as a cause of transmural lymphocytic enteritis and mortality in captive Nashville warblers (Vermivora ruficapilla). Journal of Wildlife Diseases 27:615–620. Thompson, E. J., and I. G. A. Wright. 1978. Coccidiosis in kiwis. New Zealand Veterinary Journal 26:167.
Trampel, D. W., T. M. Pepper, and B. L. Blagburn. 2000. Urinary tract cryptosporidiosis in commercial laying hens. Avian Diseases 44:479–484. Trigg, P. I. 1967. Eimeria phasiani Tyzzer, 1929—A coccidium from the pheasant (Phasianus colchicus). II. Pathogenicity and drug action. Parasitology 57:147. Tuggle, B. N., and J. L. Crites. 1984. Renal coccidiosis in interior Canada geese, Branta canadensis interior Todd, of the Mississippi Valley population. Journal of Wildlife Diseases 20:272–278. Upton, S. J., J. V. Ernst, S. L. Clubb, and W. L. Current. 1984. Eimeria forresteri n. sp. (Apicomplexa: Eimeriidae) from Ramphastos toco and a redescription of Isospora graculai from Gracula religiosa. Systematic Parasitology 6:237–240. Villan´ua, D., L. P´erez-Rodr´ıguez, C. Gort´azar, U. H¨ofle, and J. Vi˜nuela. 2006. Avoiding bias in parasite excretion estimates: The effect of sampling time and type of faeces. Parasitology 133:251– 259. Wages, D. P. 1987. Diseases of pigeons. Veterinary Clinics of North America: Small Animal Practice 17:1089–1107. Walden, H. W. 1963. Observations on renal coccidia in Swedish anseriform birds, with notes concerning two new species, Eimeria boschadis and Eimeria christianseni (Sporozoa, Telosporidia). Arkiv for Zoologi 15:97–104. Williams, R. B. 2001. Quantification of the crowding effect during infections with the seven Eimeria species of the domesticated fowl: its importance for experimental designs and the production of oocyst stocks. International Journal for Parasitology 31:1056–1069. Windingstad, R. M., M. E. McDonald, L. N. Locke, S. M. Kerr, and J. A. Sinn. 1980. Epizootic of coccidiosis in free-flying lesser scaup. Avian Diseases 24:1044–1049. Yabsley, M. J., and S. E. J. Gibbs. 2006. Description and phylogeny of a new species of Eimeria from double-crested cormorants (Phalacrocorax auritus) near Fort Gaines, Georgia. Journal of Parasitology 92:385–388. Yabsley, M. J., N. L. Gottdenker, and J. R. Fischer. 2002. Description of a new Eimeria species and associated lesions in the kidneys of double-crested cormorants (Phalacrocorax auritus). Journal of Parasitology 88:1230–1233. Yun, C. H., H. S. Lillehoj, and E. P. Lillehoj. 2000. Intestinal immune responses to coccidiosis. Developmental and Comparative Immunology 24:303–324.
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9 Disseminated Visceral Coccidiosis in Cranes Marilyn G. Spalding, James W. Carpenter, and Meliton N. Novilla venile (13–18 days old) and one adult (9 years old) Whooping Cranes died (Carpenter et al. 1980). At the time of the outbreak, the Whooping Cranes were highly endangered with fewer than 100 birds remaining in both the captive and wild population (Doughty 1989). Documentation of DVC in wild cranes based on oral granulomas and a few mortalities seems to be limited to North America, Korea, and Japan (Carpenter et al. 1979; Forrester and Spalding 2003; Watanabe et al. 2003). The pathogenesis of DVC was later characterized experimentally in Sandhill Cranes and reviewed by Novilla and Carpenter (2004).
INTRODUCTION Disseminated visceral coccidiosis (DVC) is a widely distributed intestinal and extraintestinal granulomatous disease of cranes caused by infection with intracellular apicomplexan protozoan parasites from the genus Eimeria. Two species of Eimeria are associated with the disease: Eimeria reichenowi and, to a lesser extent, Eimeria gruis. Disseminated visceral coccidiosis has caused morbidity in various species of cranes. The most significant role of this disease has been in the captive rearing of Whooping Cranes (Grus americana) for reintroduction (Carpenter et al. 1980). Although the prevalence of DVC in wild Sandhill Cranes (Grus canadensis) and recently released Whooping Cranes is high, there have been no records of mortality of wild birds without other contributing factors.
HOST RANGE Five species of Eimeria have been described in 8 of the 15 species of cranes in the world, including cranes from North America, Europe, Asia, and Africa, but only two, E. reichenowi and E. gruis, can be considered common, and are implicated as causes of DVC. It is these two species that are further discussed in this chapter (Table 9.1). The only documentation of DVC-infected wild birds comes from North American Whooping and Sandhill Cranes, and Asian White-naped (Grus vipio) and Red-crowned Cranes (Grus japonensis). Information from other crane species comes from captive birds in zoos and is not necessarily representative of wild populations. Table 9.1 lists host species for which data are available, their natural distribution, and evidence of infection with E. reichenowi and E. gruis. We found no reports of E. reichenowi and E. gruis in cranes of the subspecies Balearica, a separate subfamily that includes the crowned cranes of Africa, in spite of their common occurrence in zoos.
SYNONYMS Systemic coccidiosis, extraintestinal coccidiosis. HISTORY The agents of DVC were discovered prior to the recognition of the disseminated form of the disease. Both species, E. reichenowi and E. gruis, were originally described by Yakimoff and Matschoulsky (1935) from a captive Demoiselle Crane (Anthropoides virgo) in a zoo in Russia. Pande et al. (1970) then described Eimeria grusi n. sp. (note different spelling, considered a junior synonym of E. reichenowi) from a Sarus Crane (Grus antigone) in a zoo in India. Nodular granulomatous oral lesions containing protozoa, later to be ascribed to DVC, were first observed in captive sandhill cranes at the Patuxent Wildlife Research Center (PWRC) in Laurel, Maryland (Carpenter et al. 1979). The disease was first recognized and named after a die-off event took place at PWRC in 1978. Three ju-
ETIOLOGY Two species of Eimeria are implicated as causes of DVC. This is somewhat surprising since species of
181 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
+*
Demoiselle Crane (Anthropoides virgo): Asia, Africa Russia zoo, captive
182 15/16 (94%) — —
16/16 (100%) — —
Maryland, Texas, USA, captive Maryland, USA, experimental Maryland, USA, experimental
—
—
61/161 (38%) 139/212 (66%)
+
—
Maryland, USA, captive
New Mexico, USA
116/226 (51%) 160/212 (75%)
—
Blue Crane (Anthropoides paradiseus): southern Africa Georgia, USA
Eimeria gruis
—
16/16 (100%) —
—
68/164 (41%) 118/212 (56%)
+
—
Eimeria reichenowi
—
—
31/95 (33%) —
192/423 (45%) 42/64 (67%)
—
—
Oral granulomas
0/10 (0%) —
—
64/64 (100%† ) 24/58 (41%) GI —
—
—
1
Postmortem lesions
3/11 (27%)
—
—
6
3/226 (1%) —
—
1
Mortality
Novilla et al. (1989)
Carpenter et al. (1992)
Forrester et al. (1978)
Carpenter et al. (1979)
M. G. Spalding (unpublished data) Parker and Duszynski (1986)
Yakimoff and Matschoulsky (1935)
S. E. Little (unpublished data) and S. P. Terrell (unpublished data)
References
September 12, 2008
Sandhill Crane (Grus canadensis): North America Florida, USA
Eimeria sp.
Host and distribution
Fecal oocysts
Table 9.1. Distribution and prevalence of oocyst shedding, oral granulomas, postmortem lesions, and mortality of Eimeria spp. infections in both captive and wild cranes.
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183
Texas, Nebraska, Oklahoma, Alaska, USA, and Saskatchewan, Canada
Saskatchewan, Canada
New Mexico, USA
Indiana, USA
Florida, USA
Florida, USA
Greater Sandhill Crane (Grus canadensis tabida): North America Arizona, USA
—
40/50 (80%) —
2/73 (3%) 32/51 (63%) +
+
3/3 (100%)
60/90 (67%) +
—
36/50 (72%) —
5/14 (36%) 62/72 (86%) 11/29 (38%) —
3/3 (100%)
52/90 (58%) 4/16 (25%)
—
32/50 (64%) —
4/14 (29%) 66/72 (92%) 9/29 (31%) +
3/3 (100%)
42/90 (47%) 5/16 (31%)
—
—
—
14/51 (27%) —
—
—
—
—
—
1/1 (100%) 91/382 (24%)
4/4 (100%) —
—
—
—
—
—
—
—
—
—
—
—
—
—
—
(continues)
Carpenter et al. (1984)
Carpenter et al. (1984)
Parker and Duszynski (1986)
M. G. Spalding (unpublished data) Carpenter et al. (1984)
Courtney et al. (1975)
Courtney et al. (1975)
Forrester et al. (1978)
Courtney et al. (1975)
Parker and Duszynski (1986)
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Canadian Sandhill Crane (Grus canadensis rowani): western North America Texas, USA
Texas, USA
Lesser Sandhill Crane (Grus canadensis canadensis): East Siberia, western North America New Mexico, USA
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184 +
Sarus Crane (Grus antigone): India, Asia, Australia India zoo, captive
+§
— — —
— — —
— —
—
1
—
1
—
12/14 (86%) 55/122 (45%) 5/5 (100%)
1
+
11/14 (79%) 46/119 (39%) 5/5 (100%)
3/14 (21%) 72/144 (50%) 5/5 (100%)
4/4 (100%) —
Eimeria reichenowi
Maryland, USA, captive White-naped Crane (Grus vipio): East Asia Chulown, Korea Korea zoo, captive the Netherlands, captive
Mississippi Sandhill Crane (Grus canadensis pulla): Mississippi, USA Mississippi, USA‡
Maryland, USA, captive
3/4 (75%) —
Eimeria gruis
4/4 (100%) —
Eimeria sp.
Fecal oocysts
—
— — —
—
—
173/423 (41%) —
—
0/12 (0%)
—
Oral granulomas
—
1 1 1
—
2
—
—
—
8/12 (67%)
—
Postmortem lesions
—
1 1 1
—
2
—
—
—
1/12 (8%)
—
Mortality
Pande et al. (1970)
Kwon et al. (2006) Kim et al. (2005) Dorrestein and Van Den Brand (2006)
N. J. Thomas (unpublished data) and J. C. Franson (unpublished data) Forrester et al. (1978)
M. G. Spalding (unpublished data) Forrester et al. (1978)
Courtney et al. (1975)
Carpenter et al. (1984)
Forrester et al. (1978)
References
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Florida, USA
Maryland, USA, experimental Florida Sandhill Crane (Grus canadensis pratensis): Florida, USA Florida, USA
Maryland, USA, captive
Host and distribution
Table 9.1. (Continued)
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185 2/16 (13%)
—
2/16 (13%)
—
—
305/656 (46%) 62/146 (42%) —
—
—
—
—
4/4 (100%) —
79/121 (65%) 38/66 (58%) —
—
—
4
1
Note: Hosts are wild unless otherwise indicated. Dashes indicate that no data are available. * Indicates samples were positive but exact numbers were not given. † Each bird had a lesion in at least one tissue type; GI, gastrointestinal tract. ‡ Released captive raised birds in wild for >5 months. § Described as Eimeria grusi, synonym with Eimeria reichenowi. ¶ Reported as Hepatozoon-like protozoal disease lacking sexual reproductive stages, probably DVC.
Maryland, USA, captive
Maryland, USA, captive
Florida, USA, >1 year reintroduced Texas, USA
33/656 (5%) 9/139 (6%) 2/19 11% —
+
+ 22/656 (3%) 4/139 (3%) 4/19 21% —
+
+
91/377 (24%) + 56/686 (8%) 12/143 (8%) 6/19 32% —
—
—
—
—
—
—
4/21 (19%) —
0/121 (0%) 0/66 (0%) —
—
—
4
1
Carpenter et al. (1980) and Novilla et al. (1989) Forrester et al. (1978)
M. G. Spalding (unpublished data) M. G. Spalding (unpublished data) Forrester et al. (1978)
Watanabe et al. (2003)
Watanabe et al. (2003)
T. McNamara and E. C. Greiner, unpublished data Shimizu et al. (1987)
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Hokkaido, Japan, captive Whooping Crane (Grus americana): North America Florida, USA, reintroduced
Kagoshima, Japan, captive¶ Red-crowned Crane (Grus japonensis): East Asia Hokkaido, Japan
Hooded Crane (Grus monacha): East Asia New York, USA, captive
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(a)
(b)
Figure 9.1. Photomicrographs and line drawings of oocysts of Eimeria reichenowi (a) and Eimeria gruis (b) from the feces of a Sandhill Crane (Grus canadensis). Line drawing reprinted from Courtney et al. (1975), with permission of the Journal of Parasitology.
Eimeria are generally very host specific (Marquardt 1973). Even more interesting is the fact that both E. reichenowi and E. gruis frequently coinfect most of the host populations that have been examined. Phylogenetically, E. reichenowi and E. gruis collected from Redcrowned, Hooded (Grus monacha), and White-naped Cranes are similar but distinct from each other, forming their own cluster when compared with species of Eimeria from chickens, ruminants, and rodents that infect the intestine only. This indicates that these two eimerians may have evolved independently from the species of Eimeria found only in the intestine (Matsubayashi et al. 2005; Honma et al. 2007). In experimental studies, DVC has been reproduced using inocula-containing mixtures of oocysts of both
E. reichenowi and E. gruis and inocula-containing oocysts of either species alone (Novilla et al. 1981, 1989; Augustine et al. 1998, 2001). However, the predominance of data from experimental studies and mortality in captive birds indicates that E. reichenowi is more often associated with the acute pathogenic form of DVC. In Whooping Cranes that died at PWRC, oocysts recovered from the feces and intestinal stages of the parasites morphologically resembled E. reichenowi (Carpenter et al. 1980). Oocysts of E. reichenowi and E. gruis are distinctly different in morphology (Figure 9.1). Oocysts of E. gruis are ellipsoid to pyriform and measure on average 18 × 11.4 μm, whereas oocysts of E. reichenowi are round to ovoid/ellipsoid measuring on average
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17.8 × 15.3 μm (Courtney et al. 1975). Comparative measurements with other species are presented elsewhere (Courtney et al. 1975; Novilla et al. 1981; Parker and Duszynski 1986). Three other species of Eimeria have been described in single reports from cranes, but no evidence of their involvement in DVC has been demonstrated. They include Eimeria tropicalis and E. grusi from a captive Sarus Crane (Pande et al. 1970), and Eimeria bosquei from Lesser Sandhill Cranes (Grus canadensis canadensis) in New Mexico (Parker and Duszynski 1986). EPIZOOTIOLOGY While most eimerian protozoa confine their parasitic life cycle to the intestinal tract, the infectious agents of DVC can be found in almost any tissue in cranes. The sexual cycle can be completed in both the respiratory and the digestive tracts. A probable life history is illustrated in Figure 9.2 (Novilla et al. 1981, 1989; Novilla and Carpenter 2004). The intestinal infection in cranes follows a typical eimerian life cycle and consists of repeated cycles of asexual reproduction (merogony resulting in merozoites) followed by sexual reproduction (gametogony resulting in oocysts). This phase of the life cycle can be completed in 12 days with a noninfectious unsporulated oocyst passing in the feces by 12 days postinfection (PI). Orally ingested sporulated oocysts rupture when digested, releasing sporozoites that invade the intestinal epithelium. The site of invasion for E. gruis and E. reichenowi in experimentally infected Sandhill Cranes is predominantly the distal jejunum and ileum. By 6 h PI they can be found in the lamina propria (Augustine et al. 1998, 2001). It is not known whether these sporozoites actually invade or are engulfed by the epithelium. As with other species of Eimeria, these then undergo one or more cycles of merogony (asexual reproduction) before they become gamonts that produce macrogametocytes (female) or microgametocytes (male), which unite to form a zygote (gametogony, sexual reproduction) which matures to produce oocysts (Novilla and Carpenter 2004). The eimerians of cranes appear to uniquely differ from other eimerians in their ability to complete their life cycle in extraintestinal tissues. Unlike the asexual stages of more typical intestinal coccidians, the asexual stages of E. reichenowi and E. gruis are taken up by mononuclear cells, including large lymphocytes or macrophages, and are transported to other tissues by way of blood or lymph where further cycles of merogony can occur and where gametogony can also occur in the lung (Figure 9.3)
Figure 9.2. Probable life cycle of Eimeria sp. in cranes. (1) A sporulated oocyst is consumed and ruptures in the gut, releasing sporocysts. Sporocysts rupture to release sporozoites, which penetrate the mucosal epithelium of the distal jejunum and (2) undergo asexual merogony (m) for one or more generations. The merozoites reinfect intestinal cells or are taken up by mononuclear phagocytes and move to other organs and the lungs. (3) Merozoites initiate sexual reproduction or gametogony (G) and develop into microgametocytes (male) and macrogametocytes (female). These join to form a zygote that matures to form an oocyst. Oocysts produced in the lung are coughed up, swallowed, and pass out in feces. Oocysts produced in the intestine pass out in the feces. (4) Oocysts passed out in feces are exposed to the environment and sporulate under favorable conditions to become infective. (5) In chronic DVC, merozoites survive in granulomas in various tissues. Adapted from Novilla et al. (1981).
(Novilla et al. 1981). Why this happens in some hosts and not others is not known. Intracellular sporozoites or merozoites initiate additional cycles of merogony in a variety of tissues and form granulomas. If development occurs in the lungs, they undergo gametogony to produce oocysts. These oocysts are coughed up, swallowed, and are passed in the feces along with oocysts that are produced in the intestines. Oocysts can occur in the lung as early as 14 days PI (Novilla et al. 1989). Grossly visible nodular granulomas do not appear until 28 days PI and represent the chronic and more commonly observed phase of the disease (Figure 9.4).
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Figure 9.3. Section of pulmonary bronchus from a Sandhill Crane chick (Grus canadensis) killed 24 days after exposure to a pen contaminated with oocysts of Eimeria spp. Note gamonts (white arrows) and merozoites (black arrow) in the submucosa. Periodic acid–Schiff stain. Bar = 50 μm. Reproduced from Novilla et al. (1989), with permission of the Journal of Wildlife Diseases.
Figure 9.4. Photograph of a granuloma (arrow) in the oral cavity (top) of a wild Whooping Crane (Grus americana) and multiple granulomas (arrow) in the liver (bottom) of an experimentally infected Sandhill Crane (Grus canadensis) that died from disseminated visceral coccidiosis.
Figure 9.5. Peripheral mononuclear blood cells infected with Eimeria sp. (arrow) from a captive Hooded Crane (Grus monacha). Courtesy of Ellis Greiner, Department of Pathobiology, College of Veterinary Medicine, University of Florida, Gainesville, Florida.
In an experimental study using E. reichenowi, sporozoites were also noted in capillaries, possibly indicating the route of extraintestinal infection (Augustine et al. 1998, 2001). Extracellular merozoites may also be transported to other tissues in this way. Infected phagocytes and free merozoites have been found in peripheral blood 9 days PI in experimentally infected Sandhill Cranes, at death in a captive White-naped Crane (Dorrestein and Van Den Brand 2006), and 4 days prior to death and on the day of death in a captive Hooded Crane (Novilla et al. 1989; T. McNamara and E. C. Greiner, unpublished data) (Figure 9.5). These parasites have not been observed in the blood of wild cranes, possibly because of the low level and transient nature of the parasitemia (Box 1977). Gametogony occurs in the epithelium of the digestive and respiratory tracts with gamonts and oocysts visible by 14 days PI (Novilla et al. 1981). There have been rare observations of macrogamonts in the liver of experimentally infected Sandhill Cranes (Augustine et al. 2001) and in the oral mucosa of a Wild Sandhill Crane (Parker and Duszynski 1986). Once oocysts reach the external environment after being passed in the feces, they sporulate and become infective. Transmission occurs when a crane ingests sporulated oocysts. Cranes forage on the ground and frequently probe the soil for subterranean food. In situations where birds are crowded, particularly in
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Disseminated Visceral Coccidiosis in Cranes captivity, cranes may feed on numerous food items that are contaminated with feces. Cranes that are more widely distributed are less likely to be exposed to large numbers of sporulated oocysts. Oocysts can also be mechanically transmitted in or on insects. Since this parasite is specific to cranes, it appears that the only reservoir is an infected crane. Little information is available about environmental tolerances for the oocysts of E. gruis and E. reichenowi. Eimerian oocysts of other species can generally survive in the environment for several weeks, especially under cool, moist conditions (McDougald and Reid 1997). Oocysts do not survive freezing or high temperatures (55◦ C). The oocysts of E. reichenowi are able to survive prolonged refrigeration, longer than those of E. gruis (Novilla et al. 1989). Infection with species of Eimeria is very common in North American cranes and both E. reichenowi and E. gruis are commonly found (Table 9.1). Coinfection with both species has been reported in up to 72% of Sandhill Cranes in New Mexico (Parker and Duszynski 1986; M. G. Spalding, unpublished data). There is some association between the presence of visceral nodules and oocyst shedding, with the highest correlation being 84% (Parker and Duszynski 1986). Age has an effect on both prevalence and intensity of infection. Juvenile cranes more commonly shed oocysts and have oral granulomas (40%) than adults (20%). Intensity of infections in juveniles when measured as number of oral granulomas was approximately twice as high (4.0 granulomas per bird) as that in adults (1.7 granulomas per bird). No difference in prevalence was noted between males and females (Carpenter et al. 1984; Parker and Duszynski 1986; M. G. Spalding, unpublished data). Factors important in the epizootiology of the coccidia are reviewed by Fayer (1980). Species of Eimeria have only a single host and the oocyst is the only mechanism for parasites to infect new hosts. As a result, magnitude of oocyst production, duration of shedding of oocysts, prevalence of infected hosts, environmental conditions that affect sporulation and survival of oocysts, distribution of oocysts in the environment, and density of hosts are all factors that influence transmission. Novilla et al. (1989) proposed that low mortality of infected cranes reflects tolerance to the parasites and may represent a mechanism for the host to act as a carrier of the organism. The disseminated form of coccidiosis may make hosts better carriers by prolonging the shedding of oocysts or facilitating reemergence of the disease. In cranes, prolonged shedding may ensure contamination of both wintering and summering grounds. Since most cranes migrate out of cold weather during the winter when oocysts would be killed by cold temperatures, maintenance of
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the parasite in the population would require that birds either act as carriers or become reinfected on the summering grounds. Most cranes that migrate gather on staging grounds during migration, and fly together in large flocks. This behavior can also increase transmission when infected birds mix with uninfected birds while they wait for favorable conditions to migrate. Additionally, for some subspecies of migratory cranes such as the Greater Sandhill Cranes (Grus canadensis tabida), they comingle with resident Florida Sandhill Cranes (Grus canadensis pratensis) during the winter. Severe fulminant, clinically significant, or fatal DVC has not been reported in wild cranes. Two captivereared Mississippi Sandhill Cranes (Grus canadensis pulla) that were released in the field for about 5 months died from DVC, but also had other contributing concurrent diseases (N. Thomas and J. C. Franson, personal communication). In a few cases, young birds that were shedding oocysts were treated successfully in veterinary clinics, but it is not known if they had extraintestinal lesions. The relatively rare occurrence of disease in wild birds may be because they are less likely to ingest more than a few food items that are contaminated with sporulated oocysts. Alternatively, sick cranes may be killed by predators before the disease runs it course. Fatal wild infections, particularly in highly susceptible young chicks, may be underestimated because predators or scavengers may consume carcasses before they can be recovered for necropsy. In one of the most intensive studies of the epizootiology of DVC, captive-reared and released Whooping Cranes were monitored in Florida for the presence of DVC by fecal analysis, oral examination, and necropsy. These birds had access to a coccidiostat during the time of the release and for the time that they still remained near the release pen. This treatment reduced the number of Whooping Cranes that were shedding fecal oocysts by an order of magnitude relative to wild Sandhill Cranes at the release site (Table 9.1). Treatment with coccidiostats had no effect on the prevalence of oral granulomas in the Whooping Cranes, indicating that merogony was still taking place in spite of exposure to these drugs. When Whooping Cranes older than 1 year and no longer on coccidiostat therapy were examined, there was no significant change in the prevalence of fecal oocysts, indicating that the lack of therapy may be balanced by the decrease in exposure as birds moved away from a confinement situation. At necropsy, 32 of 74 (43%) Whooping Crane carcasses that were of sufficient quality for histological examination had characteristic lesions of DVC. Oocysts and gamonts were seen only in the lungs of two birds, while most of the remaining birds had hepatic lymphohistiocytic phlebitis. Asexual stages were frequently not visible, possibly due to exposure to coccidiostats, but
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tissues with grossly visible nodules were positive for Eimeria by polymerase chain reaction (PCR) . Lesions in two Whooping Cranes were severe enough to predispose the cranes to predation. Primary acute DVC has not been observed in wild Whooping Cranes (Forrester and Spalding 2003; M. G. Spalding, unpublished data). CLINICAL SIGNS Sandhill crane chicks with experimental infections of both E. reichenowi and E. gruis exhibit progressive weakness, emaciation, greenish diarrhea, and recumbency (Novilla et al. 1989). Little information is available about clinical signs in Whooping Cranes other than observations of lethargy and severe diarrhea in an adult bird (Carpenter et al. 1980). Although there are anecdotal reports of serum enzyme changes in birds infected with species of Eimeria (Carpenter et al. 1979; M. G. Spalding, unpublished data), there are no good clinical markers of acute disseminated coccidiosis. Birds may die prior to shedding oocysts. Infected cells in the peripheral circulation are rare and observed occasionally in only the most severe cases. Oral granulomas and fecal oocyst shedding can demonstrate infection but tell little about the stage of disease or prognosis. Such information, however, can be very useful for monitoring and managing captive populations. Clinical signs in wild birds are limited to the presence of oral granulomas and the shedding of oocysts. Oral granulomas are relatively common in cranes and can also be caused by several less common conditions (see Diagnosis section). PATHOGENESIS AND PATHOLOGY Clinical signs and mortality in Sandhill Crane chicks with experimental infections of E. reichenowi appear to be associated with widespread merogony, with mortality occurring at 10–11 days PI (Novilla et al. 1989). Sporozoites are first observed in the intestines by 6 h PI and subsequently appear in the liver, spleen, and lungs (Augustine et al. 1998). Merozoites in mononuclear cells are seen by 9 days PI and oocysts appear in feces by 12–14 days PI. A granulomatous inflammatory reaction to the presence of infected or ruptured cells or to necrosis of parenchymal cells results in the formation of granulomas in various tissues. The exact cause of the necrosis that presumably initiates the granulomatous response is not known. In acute cases, cranes die with mild to severe bronchointerstitial pneumonia, granulomatous inflammation of the trachea, esophagus, gastrointestinal tract, liver, heart, kidney, spleen, thymus, bursa, and many other extraintestinal tissues. In these cases, mortality often precedes gametogony.
Figure 9.6. Well-circ*mscribed oral granuloma from a wild Sandhill Crane (Grus canadensis) from Florida. Note the chronic nature of the granulomatous nodule in the submucosa and the meronts (arrows) within parasitophorous vacuoles (inset).
The more chronic form of DVC, and the form seen most often in wild birds, is characterized by widely disseminated lymphohistiocytic nodules. These nodules tend to lack the more active inflammation and necrosis found in the more acute phases of the disease (Figure 9.6). In severe acute cases of DVC in juvenile Whooping Cranes and experimentally infected Sandhill Cranes, birds may have turgid intestines containing fluid and greenish white mucoid material, hyperemic mucosa, congested consolidated lungs with airways that contain frothy fluid, enlarged mottled liver and spleen, and scattered orange-white nodules in many organs and tissues. Mild necrosis of the intestinal epithelium with epithelial cells containing coccidial oocysts and gametocytes is present. Asexual stages are present in the lamina propria and developing meronts are present within the cytoplasm of macrophages (Carpenter et al. 1980; Novilla et al. 1989). Less severe and more chronic cases are characterized by white raised, 0.5–4.0-mm nodules scattered through a variety of tissues including oral mucosa, esophagus, feathered skin, eyelid, lung, air sac, liver, kidney, heart, spleen, adventitia of vessels, submucosa, and serosa of the intestinal tract (Figure 9.6). The most common locations for nodules are oral and esophageal mucosa, liver, and heart. In active cases, nodules in the liver are surrounded by a red rim of hemorrhage. Microscopic lesions are well illustrated in the published literature (Carpenter et al. 1980, 1984; Novilla et al. 1981, 1989; Gardiner et al. 1988; Forrester and
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Figure 9.7. Photomicrograph of liver with mononuclear cells containing Eimeria sp. meronts (curved black arrow) infiltrating a vein and parenchyma (area delineated by white arrows) in a wild Sandhill Crane (Grus canadensis) from Florida. The white star is over a bile duct. The inset illustrates meronts within mononuclear cells in the liver at a higher magnification.
Spalding 2003) as is the ultrastructure of Eimeria sp. (Carpenter et al. 1979; Novilla et al. 1981, 1989; Parker and Duszynski 1986; Augustine et al. 2001; Dorrestein and Van Den Brand 2006). Granulomatous nodules are most commonly associated with veins, especially in the liver, resulting in phlebitis that may protrude into the lumen of the vein (Figure 9.7). The granulomas consist predominantly of lymphocytes and macrophages that displace the normal tissue architecture. In liver, heart, and blood vessels, the granulomas may invade into surrounding parenchyma. In liver, the periphery of the lesion frequently consists of an area of hemorrhage, hepatocellular necrosis, and disruption of hepatic cords. Basophilic cytoplasmic inclusions displace the nuclei of mononuclear cells (Figure 9.7, inset). Depending on the age of the lesion, variable amounts of necrotic cellular debris can be present at the center. Nodular granulomas within other tissues, such as the oral mucosa, esophagus, lungs, and skin, are usually more discrete and circ*mscribed with little evidence of necrosis and inflammatory reaction. At the light microscope level, it is sometimes difficult to discern protozoan parasites within mononuclear cells, possibly due to low numbers of parasites. In a survey of Greater (Grus canadensis tabida) and Lesser Sandhill Cranes from New Mexico, granuloma-
191
tous nodules were most prevalent in the oral mucosa (67%), followed by liver (41%), small intestine (12%), heart (10%), esophagus (3%), peritoneum (2%), and mesenteries (2%). Although protozoa were not always seen within granulomas, they were present in 67% of oral granulomas, 38% of nodules in the liver, and 14% of nodules in the small intestine. The authors reported macrogamonts in the lung of only one crane (Parker and Duszynski 1986). In experimentally infected Sandhill Cranes and captive Whooping Cranes, more generalized changes were noted such as bronchointerstitial pneumonia and parasitemia. Gamonts and oocysts were seen in bronchial epithelium (Figure 9.4). Oocysts were present in airways and the esophagus of chicks of both experimentally infected Sandhill Cranes and naturally infected captive Whooping Cranes (Carpenter et al. 1984). Captive-reared Whooping Cranes released into the wild in Florida had lesions that differed from the typical chronic granulomas, possibly because of treatment with coccidiostats during the release process. Protozoa were frequently difficult to see at the light microscope level and almost every bird had hepatic phlebitis. It was later demonstrated by PCR that Eimeria was present in these lesions (see Diagnosis section). Intestinal lesions were rare in wild Sandhill and Whooping Cranes that were a part of this study (M. G. Spalding, unpublished data). When present, lesions were very mild. DIAGNOSIS A presumptive diagnosis of DVC is made based on finding extraintestinal nodules that contain protozoal meronts or oocysts. The disease can be confirmed if eimerian oocysts are present in fecal flotation tests and there are gamonts and oocysts in the intestinal tract and/or lungs. In the cases of fulminant disease, birds may die before oocysts are produced, making identification of the protozoan more difficult. Demonstration of Eimeria within the granulomas by PCR has proved useful in the cases when merozoites are not seen by light microscopy (Terrell et al. 1999). Indirect immunofluorescence microscopy with monoclonal antibodies has also been used to study the sporozoite stage (Augustine et al. 1998, 2001). Electron microscopy may also be useful for documenting infection. Shedding of Eimeria oocysts in feces alone is not necessarily diagnostic for the disseminated form of the disease. Differential diagnoses for DVC based on gross lesions include avian tuberculosis and neoplasia, especially cholangiocarcinoma. Oral lesions can also be caused by bacteria, Candida sp., Capillaria sp., avian pox, and vitamin A deficiency. At the light microscope level, intracellular protozoa must be differentiated from other protozoans such as Leucocytozoon,
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Sarcocystis, Toxoplasma, and Isospora. This is especially true when birds die prior to development of oocysts. Clusters of parasitized cells in tissues such as liver and spleen can superficially resemble lymphosarcoma.
IMMUNITY Very little is known about how parasites interact with the host immune system in DVC. Among other species of Eimeria, infection with one species greatly reduces the severity of disease when birds are reinfected by the same species (Augustine 1999). Generally, reinfection with a second species of Eimeria results in a diminution of the reproductive potential of the second species. In chickens, T-cell-mediated immunity by intestinal lymphocytes is the predominant mechanism of protection against Eimeria (Lillehoj and Lillehoj 2000). There are both synergistic and immunosuppressive interactions between coccidiosis and other infectious diseases (reviewed by McDougald and Reid 1997). It is likely that na¨ıve, young chicks are more susceptible to the severe consequences of exposure than are older birds. Environmental contamination with oocysts and the possibility of increased transmission may explain the severity of disease in crowded conditions. However, immunosuppression related to poor diet, stress, loss of genetic diversity, and drug therapy may also be an important factor that affects severity of infection.
PUBLIC HEALTH AND DOMESTIC ANIMAL CONCERNS Infections with species of Eimeria are generally species specific, although the causative agents of DVC appear to have a much wider host range among multiple species of cranes. There have been no reports of this disease in humans or domestic animals. Broiler chicks, ducks, and dogs inoculated with pooled oocysts of E. reichenowi and E. gruis are refractory to infection (Novilla et al. 1981; M. N. Novilla and J. W. Carpenter, unpublished data).
WILDLIFE POPULATION IMPACTS The impact of DVC on wild crane populations is probably best understood for the Florida Sandhill Crane and the introduced Whooping Crane. The disease is common, but seldomly fatal in juvenile and adult cranes of both species. The role of DVC as a cause of mortality in young chicks is less well known. Mortality of wild, young crane chicks is relatively high during the first few weeks after hatching and poorly documented because of the difficulty in finding carcasses.
Mortality from DVC in wild Whooping Cranes has never been documented; however, severe lesions in birds killed by predators suggest that ill birds may be predisposed to predation. At the time of release, cranes congregate around feeders and receive pellets that contain a coccidiostat to counteract the increased risk of DVC associated with crowding. Thus, our understanding of the importance of DVC to the Florida Whooping Crane population is confounded by these unnatural factors. Fledged chicks in Florida have oral granulomas and the few that have died have disseminated granulomas, but they have not been severe enough to be considered detrimental to health. TREATMENT AND CONTROL Although treatment of wild cranes is not feasible, it is recommended for cranes in captive collections, breeding facilities, or for those being released into pens. Because DVC is a significant clinical problem in young cranes, a coccidiostat should be used in the food or water. Among coccidiostats that have been tested, monensin is the only one that provides protection against experimentally induced DVC in Sandhill Cranes (Carpenter et al. 1992, 2005). Coccidian infections in individual birds have also been treated with some success with trimethoprim-sulfamethoxazole, ormetoprim-sulfa, sulfadimethoxine, or amprolium. There is increasing evidence that coccidia can develop resistance to coccidiostats. Therefore, alternating treatment between monensin and amprolium or newer generation coccidiostats may be advisable. Captive cranes should also be monitored for oocysts, and treated as appropriate to reduce contamination of pens and potential exposure of chicks. In addition to the use of coccidiostats, reducing crowding in pens, rotation of pens, facility hygiene, quarantine, prophylactic or therapeutic treatment of new birds, and separating birds by age are integral components for controlling DVC in captivity. When birds are being released into the wild, control may still be possible because they continue to forage at the release site. Inclusion of a coccidiostat in feed is recommended. Failure to do this may lead to development of significant lesions in cranes that are being released. For example, a Whooping Crane that escaped from a pen earlier than intended developed significant lesions of DVC, most likely because it did not have access to treated feed (M. G. Spalding, unpublished data). Use of a coccidiostat during the vulnerable period when birds are young and crowded together may aid in development of immunity to infection, by suppressing multiplication of the parasite while allowing exposure to the parasite. Gradual withdrawal from the
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Disseminated Visceral Coccidiosis in Cranes coccidiostat, as birds learn to forage on natural foods, may also allow birds to develop immunity while they are still partly protected. MANAGEMENT IMPLICATIONS Because concentrations of the oocysts of Eimeria spp. in the soil may increase substantially where cranes use feeding stations, DVC is an important consideration for the management of cranes in captivity and during reintroduction into the wild. All cranes kept in close quarters and allowed to feed from soil that is contaminated with feces should receive feed treated with a coccidiostat. The significance of DVC to wild populations is probably low based on the high morbidity and low mortality observed in subadult and adult Sandhill Cranes and reintroduced Whooping Cranes in North America. Dead chicks, however, are rarely found and since they are probably more susceptible to infection, the impact of DVC on young wild birds may be greater than realized. Many endangered species of cranes are being intensively propagated around the world to help prevent their extinction. Disseminated visceral coccidiosis is an excellent example of how important disease can be when management activities that involve population manipulation in the wild or in captivity are undertaken. LITERATURE CITED Augustine, P. C. 1999. Prior or concurrent exposure to different species of avian Eimeria: Effect on sporozoite invasion and chick growth performance. Avian Diseases 43:461–468. Augustine, P. C., P. N. Klein, and H. D. Danforth. 1998. Use of monoclonal antibodies against chicken coccidia to study invasion and early development of Eimeria gruis in the Florida Sandhill Crane (Grus canadensis). Journal of Zoo and Wildlife Medicine 29:21–24. Augustine, P., G. Olsen, H. Danforth, G. Gee, and M. N. Novilla. 2001. Use of monoclonal antibodies developed against chicken coccidia (Eimeria) to study invasion and development of Eimeria reichenowi in Florida Sandhill Cranes. Journal of Zoo and Wildlife Medicine 32:65–70. Box, E. D. 1977. Life cycles of two Isospora species in the canary, Serinus canaria Linnaeus. Journal of Protozoology 24:57–67. Carpenter, J. W., T. R. Spraker, C. H. Gardiner, and M. N. Novilla. 1979. Disseminated granulomas caused by an unidentified protozoan in Sandhill Cranes. Journal of the American Veterinary Medical Association 175:948–951.
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Carpenter, J. W., T. R. Spraker, and M. N. Novilla. 1980. Disseminated visceral coccidiosis in Whooping Cranes. Journal of the American Veterinary Medical Association 177:845–848. Carpenter, J. W., M. N. Novilla, R. Fayer, and G. C. Iverson. 1984. Disseminated visceral coccidiosis in sandhill cranes. Journal of the American Veterinary Medical Association 185:1342–1346. Carpenter, J. W., M. N. Novilla, and J. S. Hatfield. 1992. The safety and physiologic effects of the anticoccidial drugs monensin and clazuril in Sandhill Cranes (Grus canadensis). Journal of Zoo and Wildlife Medicine 23:214–221. Carpenter, J. W., M. N. Novilla, and J. S. Hatfield. 2005. Efficacy of selected coccidiostats in Sandhill Cranes (Grus canadensis) following challenge. Journal of Zoo and Wildlife Medicine 36:391–400. Courtney, C. H., D. J. Forrester, J. V. Ernst, and S. A. Nesbitt. 1975. Coccidia of Sandhill Cranes, Grus canadensis. Journal of Parasitology 61:695–699. Dorrestein, G. M., and J. M. A. Van Den Brand. 2006. Disseminated visceral coccidiosis in a White-naped Crane (Grus vipio). In European Association of Zoo and Wildlife Veterinarians, 6th Scientific Meeting, Budapest, Hungary, May 24–28, 2006. Doughty, R. W. 1989. Return of the Whooping Crane. University of Texas Press, Austin, TX, 182 pp. Fayer, R. 1980. Epidemiology of protozoan infections: The coccidia. Veterinary Parasitology 6:75–103. Forrester, D. J., J. W. Carpenter, and D. R. Blankinship. 1978. Coccidia of Whooping Cranes. Journal of Wildlife Diseases 14:24–27. Forrester, D. J., and M. G. Spalding. 2003. Parasites and Diseases of Wild Birds in Florida. University Press of Florida, Gainesville, FL, 1132 pp. Gardiner, C. H., R. Fayer, and J. P. Dubey. 1988. An atlas of protozoan parasites in animal tissues. In Agriculture Handbook No. 651. U.S. Department of Agriculture, Washington, DC. Honma, H., T. Yokoyama, M. Inoue, A. Uebayashi, F. Matsumoto, Y. Watanabe, and Y. Nakai. 2007. Genetical identification of coccidian in Red-crowned Crane, Grus japonensis. Parasitology Research 100:637–640. Kim Y., E. W. Howerth, N.-S. Shin, S.-W. Kwon, S. P. Terrell, and D.-Y. Kim. 2005. Disseminated visceral coccidiosis and cloacal cryptosporidiosis in a Japanese White-naped Crane (Grus vipio). Journal of Parasitology 91:199–201. Kwon, Y.-K., W.-J. Jeon, M.-I. Kang, J.-H. Kim, and G. H. Olsen. 2006. Disseminated visceral coccidiosis in a wild White-naped Crane (Grus vipio). Journal of Wildlife Diseases 42:712–714. Lillehoj, H. S., and E. P. Lillehoj. 2000. Avian coccidiosis: A review of acquired intestinal immunity
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and vaccination strategies. Avian Diseases 44:408–425. Marquardt, W. C. 1973. Host and site specificity in the coccidian. In The Coccidia, Hammond, D. M. (ed.). University Park Press, Baltimore, MD, Chapter 2. Matsubayashi, M., K. Takami, N. Abe, I. Kamata, H. Tani, K. Sasai, and E. Baba. 2005. Molecular characterization of crane coccidian, Eimeria gruis and E. reichenowi, found in feces of migratory cranes. Parasitological Research 97:80–83. McDougald, L. R., and W. M. Reid. 1997. Coccidiosis. In Diseases of Poultry, 10th ed., B. W. Calnek (ed.). Iowa State University Press, Ames, IA, pp. 865–883. Novilla, M. N., and J. W. Carpenter. 2004. Pathology and pathogenesis of disseminated visceral coccidiosis in cranes. Avian Pathology 33:275–280. Novilla, M. N., J. W. Carpenter, T. R. Spraker, and T. K. Jeffers. 1981. Parenteral development of eimerian coccidia in sandhill and whooping cranes. Journal of Protozoology 28:248–255. Novilla, M. N., J. W. Carpenter, T. K. Jeffers, and S. L. White. 1989. Pulmonary lesions in disseminated visceral coccidiosis of Sandhill and Whooping Cranes. Journal of Wildlife Diseases 25:527–533. Pande, B. P., B. B. Bhatia, P. P. S. Chauhan, and R. K. Garg. 1970. Species composition of coccidia of some of the mammals and birds at the Zoological Gardens,
Lucknow (Uttar Pradesh). Indian Journal of Animal Science 40:154–166. Parker, B. B., and D. W. Duszynski. 1986. Coccidiosis of Sandhill Cranes (Grus canadensis) wintering in New Mexico. Journal of Wildlife Diseases 22:25–35. Shimizu, T., N. Yamada, I. Kono, and T. Koyama. 1987. Fatal infection of Hepatozoon-like organisms in the young captive cranes (Grus monacha). Memoirs of the Faculty of Agriculture, Kagoshima University 23:99–107. Terrell, S. P., S. E. Little, M. G. Spalding, and C. M. Johnson. 1999. Detection of the causative agent of disseminated visceral coccidiosis (Eimeria sp.) in Sandhill Cranes (Grus canadensis) and Whooping Cranes (Grus americana) by polymerase chain reaction amplification of 18S rDNA. In Proceedings of the Annual Conference of the Wildlife Disease Association, Athens, GA, p. 33. Watanabe, Y., F. Matsumoto, and K. Koga. 2003. A survey of the coccidian infection of wild Japanese Cranes Grus japonensis in Hokkaido, Japan (Japanese with English abstract). Journal of the Yamashina Institute of Ornithology 35:55–60. Yakimoff, W. L., and S. N. Matschoulsky. 1935. Die Kokzidiose der Kraniche. Zeitschrift fur Parasitenkunde 8:239–240.
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10 Cryptosporidium David S. Lindsay and Byron L. Blagburn baileyi, Cryptosporidium galli, and Cryptosporidium tyzzeri from chickens and turkeys, and Cryptosporidium anserinum from domestic geese. Cryptosporidium baileyi is considered to be distinct from C. meleagridis from turkeys because it differs in oocyst structure, is only moderately infectious for turkeys, and differs in site of endogenous development in the intestine. Both molecular and morphological data have confirmed that C. galli Pavlasek, 1999, from chickens is a valid species (Ryan et al. 2003). This species develops in the proventriculus and has oocysts that are larger than those of C. baileyi. The name Cryptosporidium blagburni was given to oocysts from three types of finches, the Gouldian Finch (Chloebia gouldiae), the Red-faced Pytilia (Pytilia hypogrammica), and the Plum-headed Finch (Neochmia modesta) (Morgan et al. 2001), but it is no longer considered to be a valid species (Ryan et al. 2003). The two remaining species, C. anserinum from domestic geese and C. tyzzeri from domestic chickens, were not adequately described and are considered nomina nuda. Neither of the original reports gave adequate descriptions of the oocysts or provided other useful information that would support their status as new species. Ng et al. (2006) examined the genetic diversity of oocysts collected from the feces of birds from Western Australia and the Czech Republic. They found four nondescribed genotypes plus Cryptosporidium andersoni, a species reported from adult cattle (Lindsay et al. 2000), Cryptosporidium muris, C. baileyi, C. galli, and C. meleagridis, suggesting that the diversity of species of Cryptosporidium in avian hosts may be even higher. The status of a species of Cryptosporidium that infects Northern Bobwhite (Colinus virginianus) is currently being investigated. The oocysts are structurally distinct from those of C. baileyi and resemble those of C. meleagridis (Lindsay et al. 1989). Unlike C. meleagridis, parasites from the Northern Bobwhite produce a generalized small intestinal infection that is associated with extreme morbidity and mortality in captive quail. Studies of the molecular genetics and host specificity of this parasite are needed before its status as a distinct
INTRODUCTION Members of the Genus Cryptosporidium belong to the protozoan Phylum Apicomplexa. They are coccidiallike parasites that develop in the microvillus border of epithelial cells in the digestive, respiratory, and urinary tracts of vertebrates. They have asexual and sexual reproductive stages in their life cycle and are excreted as fully sporulated oocysts. Molecular studies indicate that they are more closely related to the gregarine parasites of invertebrates than to the true coccidial parasites of vertebrates (Carreno et al. 1999). Cryptosporidium has been recognized as an increasingly important disease of commercial poultry, but has only been identified on an individual basis in wild birds, usually from fecal evaluation. There have been no recognized dieoffs in wild bird populations from cryptosporidiosis. SYNONYMS Cryptosporidiosis. HISTORY The Genus Cryptosporidium was first described by Dr E. E. Tyzzer. He was also the first to report cryptosporidial infection in an avian species and described a species in the ceca of domestic chickens (Gallus gallus) (Tyzzer 1929) that was structurally similar to Cryptosporidium parvum from mice (Mus musculus). However, he did not name or describe this parasite. A species of Cryptosporidium was subsequently reported in the ileum of turkey poults (Meleagris gallopavo) suffering from enteritis (Slavin 1955). Slavin (1955) named the parasite Cryptosporidium meleagridis and partially described its endogenous life cycle. The first complete life cycle for an avian species was described for Cryptosporidium baileyi, a species that was isolated from broiler chickens (Current et al. 1986). ETIOLOGY AND HOST RANGE Five different species of Cryptosporidium have been described from birds: Cryptosporidium meleagridis, C.
195 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Figure 10.1. Generalized life cycle of a typical species of Cryptosporidium that infects birds. Adapted from Fayer et al. (1990). species can be determined. The status of a species of Cryptosporidium from ostriches (Struthio camelus) is not fully understood. This undescribed species is not infectious for suckling mice, chickens, turkeys, or Japanese Quail (Coturnix japonica) (Gajadhar 1994), and molecular studies indicate that it is closely related, but distinct from, C. baileyi (Meireles et al. 2006). It is clear that many genotypes and species of Cryptosporidium can be found in the feces of wild birds. Studies of life cycles, molecular genetics, and host specificity are needed before we can accurately determine their true number. EPIZOOTIOLOGY The life cycles of C. baileyi (Current et al. 1986) and C. meleagridis (Slavin 1955; Pavlasek 1994) are
known in detail (Figure 10.1). Sporulated oocysts are ingested in contaminated food or water. Oocysts excyst in the digestive tract and sporozoites penetrate the microvilli of epithelial cells at locations in the intestinal tract that are specific for different species of Cryptosporidium. The sporozoite rounds up and becomes a trophozoite and undergoes asexual reproduction or merogony to become a multinucleated type I meront. These produce eight type I merozoites (Figure 10.2) that invade additional epithelilial cells and either initiate an additional cycle of type I merogony or develop into type II meronts. Type II meronts produce four type II merozoites that penetrate microvilli and become sexual stages. The male stages are microgamonts and they produce nonflagellated microgametes. The female stage is the macrogamont (Figure 10.3). Fertilization occurs and oocysts are produced. Two types of
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Figure 10.2. Transmission electron micrographs of Cryptosporidium baileyi in the respiratory tract of domestic chickens. Type II meront. Note the meront residuum (MR) and parasitophorous vacuole (PV). Bar = 1 μm. Courtesy of M. A. Cheadle. oocysts are produced and both types sporulate endogenously (Cheadle et al. 1999). Both types contain four sporozoites, an oocyst residuum, and no sporocysts. Thin-walled oocysts are autoinfective (Figure 10.4).
Figure 10.3. Transmission electron micrographs of Cryptosporidium baileyi in the respiratory tract of domestic chickens. Macrogamont. Bar = 1 μm. Courtesy of M. A. Cheadle.
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Figure 10.4. Transmission electron micrographs of Cryptosporidium baileyi in the respiratory tract of domestic chickens. Thin-walled oocyst. Note the oocyst residuum (OR) and the thin oocyst wall (arrowhead). Bar = 1 μm. Courtesy of M. A. Cheadle.
The thick-walled oocysts (Figure 10.5) are excreted in the feces (Lindsay et al. 1986b). If sporozoites or merozoites reach epithelial cells in the respiratory, conjunctival, or urinary tracts, then development can occur in these locations. Infections of these nonintestinal sites are not due to transport of infective stages by the blood (Lindsay et al. 1987b). Oocysts from the environment or oocysts currently being excreted probably come in direct contact with respiratory or conjunctival tissues, excyst, and initiate new infections on these surfaces.
CLINICAL SIGNS Cryptosporidiosis in birds manifests itself as enteritis, respiratory disease, or renal disease. Usually only one condition is present in an outbreak, but combinations of the three forms have been observed. Clinical signs of intestinal cryptosporidiosis include nonbloody diarrhea. In respiratory infections, birds may suffer from rales, coughing, sneezing, and dyspnea (ho*rr et al. 1978; Ranck and ho*rr 1987; Goodwin et al. 1988a). Prolapses of the phallus and cloaca have been reported in Ostrich chicks (Penrith et al. 1994).
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Parasitic Diseases of Wild Birds Respiratory Cryptosporidiosis Excess mucous may be present in the trachea and nasal cavities of birds with respiratory cryptosporidiosis. Infection of nasal tissues is associated with “swollen head syndrome” (Goodwin and Waltman 1994). Air sacculitis may be present. Microscopic lesions generally consist of hypertrophy and hyperplasia of infected epithelial surfaces (Mason and Hartley 1980). Infiltrates of macrophages, heterophils, lymphocytes, and plasma cells usually are present. Cilia generally are reduced or are absent on ciliated epithelial surfaces.
Figure 10.5. Transmission electron micrographs of Cryptosporidium baileyi in the respiratory tract of domestic chickens. Thick-walled oocyst. Note the oocyst residuum (OR), presence of the oocyst in a parasitophorous vacuole (PV), and thick oocyst wall (arrowhead). Bar = 1 μm. Courtesy of M. A. Cheadle. Because cryptosporidiosis in birds manifests itself as enteritis, respiratory disease, or renal disease, it is difficult to diagnose. Many other agents can cause clinical signs that are similar to those of birds with cryptosporidiosis. There is no one clinical sign that indicates cryptosporidiosis. PATHOLOGY Intestinal Cryptosporidiosis Nonbloody diarrhea is associated with intestinal cryptosporidiosis. Gross lesions are confined to the intestinal tract (ho*rr et al. 1986; Goodwin et al. 1988b). The small intestine may be distended with mucoid intestinal contents and gas. Similar lesions may be seen in the ceca. Microscopic lesions consist of villous atrophy, villous fusion, and crypt hyperplasia (ho*rr et al. 1986; Goodwin et al. 1988b). Infiltrates of lymphocytes, heterophils, macrophages, and plasma cells may be present. Cryptosporidia generally are on the distal 2/3 of the villi and are usually not seen in the ceca. Nonbloody diarrhea is also associated with cryptosporidial infection confined to the proventriculus (Blagburn et al. 1990). Lesions in the proventriculus consist of focal cuboidal metaplasia of glandular epithelial cells and deposition of amyloid in the perivascular interstitial tissues at the base of the glands (Blagburn et al. 1990)
Renal Cryptosporidiosis Infected kidneys are often pale and grossly enlarged (Abbassi et al. 1999). Urate distention may be seen in surface tubules. The epithelial cells of the collecting ducts, collecting tubules, distal convoluted tubules, and ureters are hypertrophic and hyperplastic in response to the infection (Abbassi et al. 1999; Trampel et al. 2000). Interstitial tissues may be infiltrated by lymphocytes, macrophages, heterophils, and plasma cells. Fibrotic areas may be present (Abbassi et al. 1999; Trampel et al. 2000).
DIAGNOSIS Examination of feces for oocysts or of intestinal tissues for developmental stages is the method used for documenting Cryptosporidium infections in birds and other animals. The small size of the parasite makes it difficult to detect. Oocysts of Caryospora, Isospora, and Eimeria, and sporocysts of Sarcocystis can frequently be found in the feces of wild birds. The oocysts of Caryospora, Isospora, and Eimeria are usually excreted nonsporulated. Oocysts of Sarcocystis spp. sporulate in the intestinal mucosa and are excreted as sporocysts that may be confused with oocysts of Cryptosporidium. The sporocysts of Sarcocystis contain four sporozoites, are surrounded by a sporocyst wall, and are identical in structure to the oocysts of Cryptosporidium oocyst. They can be distinguished by size and morphology of residual material. Sporocysts of Sarcocystis are 10–12 by 4–7 μm and contain several granular residual granules. Oocysts of Cryptosporidium are 4–8 by 5–6 μm and contain a compact residual body. Microscopic examination of fresh preparations from birds with respiratory signs is useful in obtaining a diagnosis (Ranck and ho*rr 1987; Goodwin et al. 1988a). Cryptosporidium oocysts can be observed in standard fecal flotations. Sheather’s sugar solution is the best flotation medium. Oocysts are difficult to see
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Cryptosporidium because of their small size. They will float in a plane higher than helminth ova and other protozoan cysts. Oocysts are observable using the 40× objective of a light microscope. They are often light pink to greenish in color with brightfield microscopy depending on the microscope objective lens. A central residual body is usually visible in Cryptosporidium oocysts in fecal flotations. More than 10 different types of staining methods have been developed to detect Cryptosporidium oocysts in fecal smears with standard light microscopy (Arrowood 1997). The Ziehl-Neelsen acidfast-staining technique is used most often and produces red-stained oocysts against a blue-green background of fecal material. Immunodetection of Cryptosporidium oocysts in feces was pioneered by the human medical community. Fluorescently labeled monoclonal antibodies that bind to the oocysts of C. parvum are used in a direct immunofluorescent antibody test. Most species of avian Cryptosporidium will cross-react with the reagents in commercial immunofluorescent antibody tests that were developed for C. parvum (Graczyk et al. 1996a). Serum from birds infected with avian species of Cryptosporidium also cross-reacts with C. parvum in enzyme-linked immunosorbent assays, making these tests useful in detecting the presence of cryptosporidial antibodies in infected birds. Detection of cryptosporidial stages in avian tissues is readily done using routine histological procedures and hematoxylin and eosin staining of tissues. The wide variety of locations within the avian host where Cryptosporidium can develop makes it important that tissues from many different anatomical sites be collected from birds at necropsy. For example, C. baileyi is usually found in the bursa of Fabricius and cloaca, C. meleagridis is usually found in the small intestine, and C. blagburni is usually found in the proventriculus. Failure to collect these tissues could result in false negative findings. Tissue should be taken from the head, trachea, lungs, kidneys, proventriculus, duodenum, jejunum, ileum, bursa of Fabricius, and cloaca to insure that all potential sites of development are examined. IMMUNITY Most studies on immunity to avian species of Cryptosporidium have been done with experimental infections of C. baileyi in chickens. Some partial immunity, as measured by oocyst excretion, may be passed in the eggs of hens infected with oocysts of C. baileyi (Hornok et al. 1998). It is not known if this occurs with other species of Cryptosporidium that infect birds. Younger chickens are more susceptible to
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infection with C. baileyi (Lindsay et al. 1988) and will produce more oocysts over a longer period of time than older birds. Severity of clinical signs, respiratory disease, and magnitude of oocyst production are also greater in younger chickens when they are inoculated with oocysts in the respiratory tract. Birds develop serum antibodies following exposure to oocysts of C. baileyi (Current and Snyder 1988). Studies using chemical bursectomy and treatment with cyclosporin A indicate that cell-mediated immunity (CMI) is more important in resistance to C. baileyi than circulating antibody. Hatkin et al. (1993) found that bursectomy altered serum antibody production, but not CMI, as measured by the delayed-type hypersensitivity skin reaction. They also reported that chickens treated with cyclosporin were more susceptible to respiratory disease than untreated controls. Sr´eter et al. (1996) found that immunity to reinfection was inhibited in thymectomized chickens infected with oocysts of C. baileyi, further indicating the role of CMI in resistance to Cryptosporidium. Infection with Eimeria species does not induce immunity to Cryptosporidium in chickens; however, experimental infection with C. parvum does induce some cross-resistance (Sr´eter et al. 1997). Infection with C. baileyi may inhibit development of antibodies to other infectious agents. Rhee et al. (1998a, 1998b) have demonstrated decreased antibody production to Brucella abortus vaccine and sheep red blood cells in chickens infected with C. baileyi. Secondary infections with bacteria and viruses may also be present in natural cases and add to the severity of disease. PUBLIC HEALTH CONCERNS Birds as a Source of C. parvum and Cryptosporidium hominis In the early 1980s, C. parvum was identified as a major cause of intestinal disease in AIDS patients and it was later found in immunocompetent humans. It is now well recognized as a public health problem. Viable oocysts of C. parvum can pass undamaged through the digestive system of several avian hosts (Graczyk et al. 1996b, 1997) and the oocysts of C. parvum have been detected in feces of wild Canada Geese (Branta canadensis) (Graczyk et al. 1998). These oocysts are infectious for mice, indicating that migratory waterfowl can disseminate infectious oocysts. Oocysts comprising a minimum of five different genotypes of Cryptosporidium can be found in the feces of Canada Geese (Jellison et al. 2004), including oocysts of C. hominis (Zhou et al. 2004).
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Wild birds can contaminate water with fecal droppings containing oocysts of Cryptosporidium. Water is a major source of oocysts and outbreaks can occur in developed countries including North America, the United Kingdom, and Japan (Fayer 2004). Surveys of surface water, groundwater, estuaries, and seawater indicate that contamination of water with Cryptosporidium is common and not isolated to specific geographical regions (Fayer 2004). Migratory birds are the likely source of oocysts of Cryptosporidium because environmental samples are more likely to be positive when birds are present (Jellison et al. 2007). C. meleagridis Infections in Humans C. meleagridis is reported frequently from humans. In a large survey in England of human cases with diarrhea and oocysts, 0.9% of 2,414 cases were due to C. meleagridis (Leoni et al. 2006). Oocysts of C. meleagridis have been reported from humans (Xiao et al. 2001). These findings have all been based on molecular-based diagnostic tests for oocysts and not on actual recovery of oocysts from fecal material. These reports indicate that C. meleagridis may pose a public health risk. Most human infections with C. meleagridis have been in children or immunocompromised individuals, although diarrhea can also occur in individuals that have no identifiable immune deficiency (PedrazaD´ıaz et al. 2001). Oocysts of C. meleagridis that were isolated from turkeys were infectious to immunosuppressed mice (Sr´eter et al. 2000), lending evidence to the potential zoonotic threat posed by this species. C. baileyi Infections in Humans C. baileyi is not infectious for laboratory mammals under experimental conditions (Lindsay et al. 1986a), although C. baileyi-like parasites have been found in human feces on a few occasions (Ditrich et al. 1991, 1993). There is one report of transmission from humans to chickens (Ditrich et al. 1993). DOMESTIC ANIMAL HEALTH CONCERNS Since migratory birds can carry infectious oocysts of C. parvum in their feces (Graczyk et al. 1998), they may serve as a source of infection for domestic livestock. Because of the confinement of domestic birds, wild birds do not appear to serve as a significant source of cryptosporidial oocysts. Wild birds may serve as a source of oocysts for free range poultry because they can contaminate the environment with oocysts. WILDLIFE POPULATION IMPACTS No clinical outbreaks of avian cryptosporidiosis have been reported in wild flocks of birds. European
Herring Gulls (Larus argentatus) and Black-headed Gulls (Larus ridibundus) have been reported to be naturally infected with Cryptosporidium based on the presence of oocysts in the feces (Smith et al. 1993; Pavlasek 1993), but infections did not cause morbidity or mortality. TREATMENT AND CONTROL There is presently no proven treatment for avian cryptosporidiosis. Commonly used ionophorous anticoccidials are not effective (Lindsay et al. 1987a; Varga et al. 1995). The addition of the antioxidant, duokvin, to feed that contains ionophors increases their efficacy, but the combination is toxic (Varga et al. 1995). Diclazuril and toltrazuril are also not effective (Sr´eter et al. 1999). Enrofloxacin is marginally effective and paromomycin causes a reduction in oocyst output by 67 to 82% (Sr´eter et al. 2002). Paromomycin has a positive effect on weight gain. Control methods may be useful in limiting or preventing avian cryptosporidiosis in zoos or rehabilitation centers. Several commonly used disinfectants were evaluated for the ability to kill oocysts of C. baileyi in an excystation assay (Sundermann et al. 1987). None of the disinfectants was effective at concentrations recommended by the manufacturers. Commercially available ammonia compounds were effective when used at 50% (v/v) concentration, with less than 5% of oocysts remaining viable. A similar concentration of commercial bleach (5.25% sodium hypochlorite) was somewhat effective with less than 15% of oocysts remaining viable. Treatment of metal brooders, feeders, and waterers with a bleach solution followed by exposure to direct sunlight for 3 days, cleaning concrete floors, and replacing wood shavings, were effective methods for controlling an outbreak of cryptosporidiosis in young Northern Bobwhite (ho*rr et al. 1986). Like most coccidial infections, cryptosporidiosis is a disease of confined birds or birds that are present in an area in large numbers. Any management program that decreases the numbers of birds in an area will decrease the probability of transmission of the parasite. LITERATURE CITED Abbassi, H., F. Coudert, Y. Ch´erel, G. Dambrine, J. Brug`ere-Picoux, and M. Naciri. 1999. Renal Cryptosporidiosis (Cryptosporidium baileyi) in specific-pathogen-free chickens experimentally coinfected with Marek’s disease virus. Avian Diseases 43:738–744. Arrowood, M. J. 1997. Diagnosis. In Cryptosporidium and cryptosporidiosis, R. Fayer (ed.). CRC Press, Boca Raton, FL, pp. 43–64.
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Cryptosporidium Blagburn, B. L., D. S. Lindsay, F. J. ho*rr, A. L. Atlas, and M. Toivio-Kinnucan. 1990. Cryptosporidium sp. infection in the proventriculus of an Australian diamond firetail finch (Stagonopleura bella: Passeriformes, Estrildidae). Avian Diseases 34:1027– 1030. Carreno, R. A., D. S. Martin, and J. R. Barta. 1999. Cryptosporidium is more closely related to the gregarines than to coccidia as shown by phylogenetic analysis of apicomplexan parasites inferred using small-subunit ribosomal RNA gene sequences. Parasitology Research 85:899–904. Cheadle, M. A., M. Toivio-Kinnucan, and B. L. Blagburn. 1999. The ultrastructure of gametogenesis of Cryptosporidium baileyi (Eimeriorina; cryptosporidiidae) in the respiratory tract of broiler chickens (Gallus domesticus). Journal of Parasitology 85:609–615. Current, W. L., and D. B. Snyder. 1988. Development of and serologic evaluation of acquired immunity to Cryptosporidium baileyi by broiler chickens. Poultry Science 67:720–729. Current, W. L., S. J. Upton, and T. B. Haynes. 1986. The life cycle of Cryptosporidium baileyi n. sp. (Apicomplexa, Cryptosporidiidae) infecting chickens. Journal of Protozoology 33:289–296. Ditrich, O., P. Kopacek, and Z. Kucerova. 1993. Antigenic characterization of human isolates of cryptosporidia. Folia Parasitologica (Praha) 40:301– 305. Ditrich, O., L. Palkovic, J. Sterba, J. Prokopic, J. Loudova, and M. Giboda. 1991. The first finding of Cryptosporidium baileyi in man. Parasitology Research 77:44–47. Fayer, R. 2004. Cryptosporidium: a water-borne zoonotic parasite. Veterinary Parasitology 126:37– 56. Fayer, R., C. A. Speer, and J. P. Dubey. 1990. General biology of Cryptosporidium. In Cryptosporidiosis of Man and Animals. J. P. Dubey, C. A. Speer, and R. Fayer (eds.). CRC Press, Boca Raton, FL, 1–29. Gajadhar, A. A. 1994. Host specificity studies and oocyst description of a Cryptosporidium sp. isolated from ostriches. Parasitology Research 80:316–319. Goodwin, M. A., and W. D. Waltman. 1994. Clinical and pathological findings in young Georgia broiler chickens with oculofacial respiratory disease (“so-called swollen heads”). Avian Diseases 38:376– 378. Goodwin, M. A., K. S. Latimer, J. Brown, W. L. Steffens, P. W. Martin, R. S. Resurreccion, M. A. Smeltzer, and T. G. Dickson. 1988a. Respiratory cryptosporidiosis in chickens. Poultry Science 67: 1684–1693.
201
Goodwin, M. A., W. L. Steffens, I. D. Russell, and J. Brown. 1988b. Diarrhea associated with intestinal cryptosporidiosis in turkeys. Avian Diseases 32:63– 67. Graczyk, T. K., M. R. Cranfield, and R. Fayer. 1996a. Evaluation of commercial enzyme immunoassay (EIA) and immunofluorescent antibody (FA) test kits for detection of Cryptosporidium oocysts of species other than Cryptosporidium parvum. American Journal of Tropical Medicine and Hygiene 54:274– 279. Graczyk, T. K., M. R. Cranfield, R. Fayer, and M. S. Anderson. 1996b. Viability and infectivity of Cryptosporidium parvum oocysts are retained upon intestinal passage through a refractory avian host. Applied and Environmental Microbiology 62:3234– 3237. Graczyk, T. K., M. R. Cranfield, R. Fayer, J. Trout, and H. J. Goodale. 1997. Infectivity of Cryptosporidium parvum oocysts is retained upon intestinal passage through a migratory water-fowl species (Canada goose, Branta canadensis). Tropical Medicine and International Health 2:341–347. Graczyk, T. K., R. Fayer, J. M. Trout, E. J. Lewis, C. A. Farley, I. Sulaiman, and A. A. Lal. 1998. Giardia sp. cysts and infectious Cryptosporidium parvum oocysts in the feces of migratory Canada geese (Branta canadensis). Applied and Environmental Microbiology 64:2736–2738. Hatkin, J., J. J. Giambrone, and B. L. Blagburn. 1993. Correlation of circulating antibody and cellular immunity with resistance against Cryptosporidium baileyi in broiler chickens. Avian Diseases 37:800– 804. ho*rr, F. J., F. M. Ranck, and T. F. Hastings. 1978. Respiratory cryptosporidiosis in turkeys. Journal of the American Veterinary Medical Association 173: 1591–1593. ho*rr F. J., W. L. Current and T. B. Haynes. 1986. Fatal cryptosporidiosis in quail. Avian Diseases 30:421–425. Hornok, S., Z. Bitay, Z. Szell, and I. Varga. 1998. Assessment of maternal immunity to Cryptosporidium baileyi in chickens. Veterinary Parasitology 79:203–212. Jellison, K. L., D. L. Distel, H. F. Hemond, and D. B. Schauer. 2004. Phylogenetic analysis of the hypervariable region of the 18S rRNA gene of Cryptosporidium oocysts in feces of Canada geese (Branta canadensis): evidence for five novel genotypes. Applied Environmental Microbiology 70:452–458. Jellison, K. L., D. L. Distel, H. F. Hemond, and D. B. Schauer. 2007. Phylogenetic analysis implicates birds as a source of Cryptosporidium spp. oocysts in
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agricultural watersheds. Environmental Science and Technology 41:3620–3625. Leoni, F., C. Amar, G. Nichols, S. Pedraza-D´ıaz, and J. McLauchlin. 2006. Genetic analysis of Cryptosporidium from 2414 humans with diarrhea in England between 1985 and 2000. Journal of Medical Microbiology 55:703–707. Lindsay, D. S., B. L. Blagburn, and C. A. Sundermann. 1986a. Host specificity of Cryptosporidium sp. isolated from chickens. Journal of Parasitology 72: 565–568. Lindsay, D. S., B. L. Blagburn, C. A. Sundermann, J. F. ho*rr, and J. A. Ernest. 1986b. Experimental Cryptosporidium infections in chickens; oocyst structure and tissue specificity. American Journal of Veterinary Research 47:876–879. Lindsay, D. S., B. L. Blagburn, C. A. Sundermann, and J. A. Ernest. 1987a. Chemoprophylaxis of cryptosporidiosis in chickens, using halofuginone, salinomycin, lasalocid, or monensin. American Journal of Veterinary Research 48:354–355. Lindsay, D. S., B. L. Blagburn, C. A. Sundermann, F. J. ho*rr, and J. J. Giambrone. 1987b. Cryptosporidium baileyi: effects of intravenous and intra-abdominal inoculation of oocysts on infectivity and site of development in broiler chickens. Avian Diseases 31:841–843. Lindsay, D. S., B. L. Blagburn, C. A. Sundermann, and J. J. Giambrone. 1988. Effect of broiler chicken age on susceptibility to experimentally induced Cryptosporidium baileyi infection. American Journal of Veterinary Research 49:1412–1414. Lindsay, D. S., B. L. Blagburn, and C. A. Sundermann. 1989. Morphometric comparison of the oocysts of Cryptosporidium meleagridis and C. baileyi from birds. Proceedings of the Helminthological Society of Washington 56:91–92. Lindsay, D. S., S. J. Upton, D. S. Owens, U. M. Morgan, J. R. Mead, and B. L. Blagburn. 2000. Cryptosporidium andersoni n. sp. (Apicomplexa: Cryptosporiidae) from cattle, Bos taurus. Journal of Eukaryotic Microbiology 47:91–95. Mason, R. W., and W. J. Hartley. 1980. Respiratory cryptosporidiosis in a peaco*ck chick. Avian Diseases 24:771–776. Meireles, M. V., R. M. Soares, M. M. dos Santos, and S. M. Gennari. 2006. Biological studies and molecular characterization of a Cryptosporidium isolate from ostriches (Struthio camelus). Journal of Parasitology 92:623–626. Morgan, U. M., P. T. Monis, L. Xiao, J. Limor, I. Sulaiman, S. Raidal, P. O’Donoghue, R. Gasser, A. Murray, R. Fayer, B. L. Blagburn, Altaf A. Lal, and R. C. A. Thompson. 2001. Molecular and phylogenetic characterization of Cryptosporidium
from birds. International Journal for Parasitology 31:289–296. Ng, J., I. Pavlasek, and U. Ryan. 2006. Identification of novel Cryptosporidium genotypes from avian hosts. Applied Environmental Microbiology 72:7548– 7553. Pavlasek, I. 1993. The black-headed gull (Larus ridibundus L.), a new host for Cryptosporidium baileyi (Apicomplexa: Cryptosporidiidae). Veterinary Medicine (Praha) 38:629–638. Pavlasek, I. 1994. Localization of endogenous developmental stages of Cryptosporidium meleagridis Slavin, 1955 (Apicomplexa: Cryptosporidiidae) in birds. Veterinary Medicine (Praha) 39:733–742. Pedraza-D´ıaz, S., C. F. Amar, J. McLauchlin, G. L. Nichols, K. M. Cotton, P. Godwin, A. M. Iversen, L. Milne, J. R. Mulla, K. Nye, H. Panigrahl, S. R. Venn, R. Wiggins, M. Williams, and E. R. Youngs. 2001. Cryptosporidium meleagridis from humans: molecular analysis and description of affected patients. Journal of Infection 42:243–250. Penrith, M. L., A. J. Bezuidenhout, W. P. Burger, and J. F. Putterill. 1994. Evidence for cryptosporidial infection as a cause of prolapse of the phallus and cloaca in ostrich chicks (Struthio camelus). Onderstepoort Journal of Veterinary Research 61: 283–289. Ranck F. M., and F. J. ho*rr. 1987. Cryptosporidia in the respiratory tract of turkeys. Avian Diseases 31:389–391. Rhee, J. K., H. C. Kim, and B. K. Park. 1998a. Effect of Cryptosporidium baileyi infection on antibody response to sRBC in chickens. Korean Journal of Parasitology 36:33–36. Rhee, J. K., H. J. Yang, and H. C. Kim. 1998b. Verification of immunosuppression in chicks caused by Cryptosporidium baileyi infection using Brucella abortus strain 1119-3. Korean Journal of Parasitology 36:281–4. Ryan, U. M., L. Xiao, C. Read, I. M. Sulaiman, P. Monis, A. A. Lal, R. Fayer, and I. Pavlasek. 2003. A redescription of Cryptosporidium galli Pavlasek, 1999 (Apicomplexa: Cryptosporidiidae) from birds. Journal of Parasitology 89: 809–913. Slavin, D. 1955. Cryptosporidium meleagridis (sp. nov.). Journal of Comparative Pathology 65:262– 266. Smith, H. V., J. Brown, J. C. Coulson, G. P. Morris, and R. W. Girdwood. 1993. Occurrence of oocysts of Cryptosporidium sp. in Larus spp. gulls. Epidemiology and Infection 110:135–143. Sr´eter, T., I. Varga, and L. Bekesi. 1996. Effects of bursectomy and thymectomy on the development of resistance to Cryptosporidium baileyi in chickens. Parasitology 82:174–177.
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Cryptosporidium Sr´eter, T., S. Hornok, I. Varga, L. Bekesi, and Z. Szell. 1997. Attempts to immunize chickens against Cryptosporidium baileyi with C. parvum oocysts and Paracox vaccine. Folia Parasitologica (Praha) 44:77– 80. Sr´eter, T., Z. Sz´ell, and I. Varga I. 1999. Attempted chemoprophylaxis of cryptosporidiosis in chickens, using diclazuril, toltrazuril, or garlic extract. Journal of Parasitology 85:989–991. Sr´eter, T., G. Kovacs, A. J. da Silva, N. J. Pieniazek, Z. Szell, M. Dobos-Kovacs, K. Marialigeti, and I. Varga. 2000. Morphologic, host specificity, and molecular characterization of a Hungarian Cryptosporidium meleagridis isolate. Applied and Environmental Microbiology 66:735–738. Sr´eter, T., Z. Sz´ell, and I. Varga. 2002. Anticryptosporidial prophylactic efficacy of enrofloxacin and paromomycin in chickens. Journal of Parasitology 88:209–211. Sundermann, C. A., D. S. Lindsay, and B. L. Blagburn. 1987. Evaluation of disinfectants for ability to kill
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avian Cryptosporidium oocysts. Companion Animal Practice 1:36–39. Trampel, D. W., T. M. Pepper, and B. L. Blagburn. 2000. Urinary tract cryptosporidiosis in commercial laying hens. Avian Diseases 44:479–484. Tyzzer, E. E. 1929. Coccidiosis in gallinaceous birds. American Journal of Hygiene 10:269–383. Varga, I., T. Sr´eter, and L. Bekesi. 1995. Potentiation of ionophorous anticoccidials with duokvin: battery trials against Cryptosporidium baileyi in chickens. Journal of Parasitology 81:777– 780. Xiao, L., C. Bern, J. Limor, I. Sulaiman, J. Roberts, W. Checkley, L. Cabrera, R. H. Gilman, and A. A. Lal. 2001. Identification of 5 types of Cryptosporidium parasites in children in Lima, Peru. Journal of Infectious Diseases 183:492–497. Zhou, L., H. Kassa, M. L. Tischler, and L. Xiao. 2004. Host-adapted Cryptosporidium spp. in Canada geese (Branta canadensis). Applied Environmental Microbiology 70:4211–4215.
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11 Toxoplasma J. P. Dubey INTRODUCTION Toxoplasma gondii is a protozoan parasite of worldwide distribution. It infects virtually all warm-blooded animals, including birds and humans (Dubey and Beattie 1988; Dubey 1993; Remington et al. 1995; Tenter et al. 2000; Dubey and Odening 2001). It can cause serious disease in many hosts, especially Australasian marsupials, new world monkeys, and those with immunodeficiencies. Severe toxoplasmosis has been reported in endangered Hawaiian Crows (Corvus hawaiiensis), canaries (Serinus spp.), and finches (Fringillidae).
DISTRIBUTION AND HOST RANGE Toxoplasma gondii has a worldwide distribution. Because there are several T. gondii-like parasites in birds (Atoxoplasma, Isospora, Sarcocystis) (Dubey 2002; Chapter 5), true hosts of T. gondii are only those where T. gondii has been demonstrated by bioassays. Reports of isolation of viable T. gondii from tissues of various avian species without clinical signs are summarized in Table 11.1. For bioassays, tissue hom*ogenates from naturally exposed animals are inoculated into laboratory animals or cell cultures to observe the development of T. gondii; outbred Swiss albino mice are the animals most commonly used. Detection of T. gondii DNA is not sufficient to prove the presence of the parasite; rather, development of the organisms in cell culture or subinoculated mice is the most definitive assay. Demonstration of antibodies to T . gondii only indicates exposure to the organism but does not provide any information whether the host harbors live parasites. Serologic tests, in most cases, do not provide information about whether infections are recent or whether they are latent (Table 11.2).
HISTORY Toxoplasma gondii was first discovered in a Tunisian rodent, Ctenodactylus gundi, by Nicolle and Manceux (1908, 1909). At about the same time, Splendore (1909) independently described a similar parasite in a laboratory rabbit in S˜ao Paulo, Brazil. The complete life cycle was not discovered until 1970 when cats were found to be the only definitive hosts (reviewed by Dubey and Beattie 1988 and Dubey 2007). Toxoplasma-like parasites were first observed in birds by Carini (1911) in smears prepared from the liver and spleen of a Rock Pigeon (Columba livia) in S˜ao Paulo, Brazil. Previously, there were reports of Toxoplasma-like parasites in sparrows and other birds, but they were believed to be hemoprotozoans. Toxoplasma was subsequently reported from several species of birds (Dubey 2002), but these identifications may not have been accurate because there were no T. gondiispecific serologic or immunohistochemical techniques available prior to 1950. The development of the dye test by Sabin and Feldman (1948) provided a reliable serological method for evaluating and comparing presumed infections with T. gondii among various animal species. In the 1950s and 1960s, it became clear that there were no morphologic or serologic differences among various isolates of T. gondii from avian or mammalian hosts.
ETIOLOGY Toxoplasma gondii is a coccidian parasite with cats as the definitive host and warm-blooded animals as intermediate hosts. Current classifications place it in the phylum Apicomplexa Levine, 1970; class Sporozoasida Leukart, 1879; subclass Coccidiasina Leukart, 1879; order Eimeriorina Leger, 1911; and family Toxoplasmatidae Biocca, 1956. There is only one species, T. gondii, but genetic differences exist among isolates of T. gondii, even within a given host. For example, both pandemic strains and strains specific to different continents were recently identified among over 200 isolates of T. gondii from free-range chickens (Lehmann et al. 2006). Isolates of T. gondii have been classified by biological characteristics as mouse virulent or avirulent and by molecular methods into three main lineages (Types I, II, and III). Type I lineages are
204 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
205 Larus ridibundus Sterna hirundo
Black-headed Gull Common Tern
Charadriiformes
Columbiformes Columba palumbus Columba livia
Meleagris gallopavo Fulica atra
Wild Turkey Eurasian Coot
Gruiformes
Galliformes
Common Wood-Pigeon Rock Pigeon
Falco tinnunculus Falco sparverius Circus macrourus Aegypius monachus Buteo jamaicensis Buteo lineatus Perdix perdix Phasianus colchicus
Eurasian Kestrel American Kestrel Pallid Harrier Cinereous Vulture Red-tailed Hawk Red-shouldered Hawk Gray Partridge Ring-necked Pheasant
Falconiformes
Streptopelia decaocto
Aythya ferina Aythya fuligula Anas acuta Anas strepera Anser anser Branta canadensis Accipiter gentilis Accipiter cooperi Buteo buteo
Common Pochard Tufted Duck Northern Pintail Gadwall domestic goose Canada Goose Northern Goshawk Cooper’s Hawk Eurasian Buzzard
184 19 8 25 57 93 1 1 10 4 123 12 1 3 3 4 27 12 16 590 1 16 43 29 61 84 14 3 60 12 12 606 3 16 1 50
N 12.0 5 12.5 28.0 1.8 1.1 100 100 10.0 25 8.1 8.3 100 33.3 33.3 25.0 41.1 66.7 18.7 2.4 100 50 4.6 3.4 16.4 1.2 7.1 33.3 5.0 50 8.3 1.0 100 12.5 100 2
Infected (%)
(continues)
Liter´ak et al. (1992) Dubey et al. (2003) Liter´ak et al. (1992) Liter´ak et al. (1992) Pak (1976) Pak (1976) Dubey et al. (2007) Dubey et al. (2004) Liter´ak et al. (1992) Lindsay et al. (1993) Liter´ak et al. (1992) Pak (1976) Liter´ak et al. (1992) Lindsay et al. (1993) Pak (1976) Pak (1976) Lindsay et al. (1993) Lindsay et al. (1993) Liter´ak et al. (1992) Liter´ak et al. (1992) ˇ ar (1974) Cat´ Lindsay et al. (1994) Liter´ak et al. (1992) Pak (1976) Liter´ak et al. (1992) Pak (1976) Pak (1976) Pak (1970) Liter´ak et al. (1992) ˇ ar (1974) Cat´ Liter´ak et al. (1992) Liter´ak et al. (1992) Siim et al. (1963) ˇ ar (1974) Cat´ Feldman and Sabin (1949) Manwell and Drobeck (1951)
Reference
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Eurasian Collared-Dove
Czech Republic Egypt Czech Republic Czech Republic Kazakhstan Kazakhstan USA USA Czech Republic USA Czech Republic Kazakhstan Czech Republic USA Kazakhstan Kazakhstan USA USA Czech Republic Czech Republic Slovakia USA Czech Republic Kazakhstan Czech Republic Kazakhstan Kazakhstan USSR Czech Republic Slovakia Czech Republic Czech Republic Denmark Slovakia USA USA
Anas platyrhynchos
Mallard
Anseriformes
Country
Species
Scientific name
Order
Table 11.1. Isolation of Toxoplasma gondii from tissues of naturally infected wild birds.
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Scientific name
Streptopelia senegalensis Columbina talpacoti Melopsittacus undulatus Glaucidium brasilianum Athene noctua Bubo virginianus Strix varia Strix aluco Lanius excubitor Emberiza citrinella Fringilla coelebs Passer domesticus
Passer montanus Garrulus glandarius Sturnus vulgaris Thraupis palmarum Turdus merula
Species
Laughing Dove Ruddy Ground-Dove Budgerigar Ferruginous Pygmy-Owl Little Owl Great Horned Owl Barred Owl Tawny Owl Northern Shrike Yellowhammer
Chaffinch
House Sparrow
Eurasian Tree Sparrow
Eurasian Jay European Starling
Palm Tanager Eurasian Blackbird
N 80 16 20 79 2 1 15 5 15 1 1 5 185 133 152 106 (1,907) 40 177 5 412 4 316 178 43 69 430 3 54 4
Country USA USA Kazakhstan Panama Switzerland Costa Rica Kazakhstan USA USA France Czech Republic Czech Republic Czech Republic Czech Republic Czech Republic Costa Rica Czech Republic Czech Republic Kazakhstan Slovakia USSR Czech Republic Czech Republic Kazakhstan Czech Republic Czech Republic Kazakhstan Panama Czech Republic Slovakia
5 6 5.0 3 100 Not given 6.7 20 26.7 100 100 20 0.5 0.7 0.7 16 0.5 17.5 1.7 40 0.5 25 0.6 0.6 2.3 1.4 0.5 33.3 1.9 25
Infected (%) Jacobs et al. (1952) Gibson and Eyles (1957) Pak (1976) Frenkel et al. (1995) Galli-Valerio (1939) Holst and Chinchilla (1990) Pak (1976) Lindsay et al. (1993) Lindsay et al. (1993) Aubert et al. (2008) Liter´ak et al. (1992) Hejl´ıcˇ ek et al. 1981 Liter´ak et al. (1992) Liter´ak et al. (1992) Liter´ak et al. (1992) Ruiz and Frenkel (1980) Liter´ak et al. (1992) Hejl´ıcˇ ek et al. 1981 Pak (1976) ˇ ar (1974) Cat´ Pak (1972) Hejl´ıcˇ ek et al. 1981 Liter´ak et al. (1992) Pak (1976) Liter´ak et al. (1992) Liter´ak et al. (1992) Pak (1976) Frenkel et al. (1995) Liter´ak et al. (1992) ˇ ar (1974) Cat´ (continues)
Reference
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Passeriformes
Psittaciformes Strigiformes
Order
Table 11.1. (Continued)
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206
207
Sitta europaea Certhia familiaris Carduelis chloris Corvus brachyrhynchos Corvus corone Corvus monedula Corvus frugilegus
Eurasian Nuthatch Eurasian Treecreeper European Greenfinch American Crow Carrion Crow
Eurasian Jackdaw Rook
Source: Modified from Dubey (2002).
Turdus viscivorus Turdus philomelos Erithacus rubecula Parus major Slovakia Slovakia Slovakia Czech Republic Slovakia Slovakia Slovakia Slovakia USA Kazakhstan Slovakia Czech Republic Czech Republic
1 7 8 215 5 6 1 1 82 58 4 5 495
100 71.4 37.5 1.4 40 33 100 100 1.2 1.7 50 20.0 18.0
ˇ ar (1974) Cat´ ˇ ar (1974) Cat´ ˇ ar (1974) Cat´ Liter´ak et al. (1992) ˇ ar (1974) Cat´ ˇ ar (1974) Cat´ ˇ ar (1974) Cat´ ˇ ar (1974) Cat´ Finlay and Manwell (1956) Pak (1976) ˇ ar (1974) Cat´ Liter´ak et al. (1992) Liter´ak et al. (1992)
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Mistle Thrush Song Thrush European Robin Great Tit
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208
Passeriformes
Strigiformes
Strix aluco Thryothorus modestus Mimus polyglottos Turdus migratorius Turdus grayi
Tawny Owl Plain Wren Northern Mockingbird American Robin
Clay-colored Robin
Fulica americana Larus delawarensis Larus atricilla Columba livia
Gruiformes American Coot Charadriiformes Ring-billed Gull Laughing Gull Columbiformes Rock Pigeon
Streptopelia chinensis Columbina talpacoti Tyto alba
Canada USA USA USA Norway Nigeria USA France USA USA USA USA USA Belgium Germany Italy South Africa Taiwan USA USA USA USA USA USA Panama USA USA USA France France Panama USA USA USA Panama
Struthio camelus Bubulcus ibis Aix sponsa Anseranas semipalmata Branta leucopsis Gyps africanus Cathartes aura Buteo buteo Meleagris gallopavo
Struthioniformes Ostrich Ciconiiformes Cattle Egret Anseriformes Wood Duck Magpie Goose Barnacle Goose Falconiformes White-backed Vulture Turkey Vulture Eurasian Buzzard Galliformes Wild Turkey
973 40 16 11 149 240 2 14 130 16 38 13 33 220 49 108 16 665 20 15 80 34 322 134 79 38 80 28 18 12 1 133 23 20 12
2.9 2.5 6 10.8 7 64.8 50 79 10 71 3 15.3 6 3.18 2 3 100 4.7 10 6 8.7 5.9 8.6 8.2 12.6 27.3 22.5 10.7 11 50 100 0.75 8.6 5 16.6
MAT IHAT IHAT MAT MAT MAT IHAT MAT MAT MAT IHAT IHAT IHAT MAT DT DT IHAT LAT DT DT DT MAT IHAT DT MAT MAT MAT MAT MAT MAT MAT IHAT IHAT IHAT MAT
1:25 1:64 1:64 1:25 1:40 1:25 1:64 1:25 1:25 1:25 1:64 1:64 1:64 1:64 1:16 1:50 1:64 1:32 1:16 1:16 1:16 1:40 1:16 1:16 1:5 1:40 1:40 1:40 1:25 1:25 1:5 1:64 1:64 1:64 1:5
N Positive (%) Test Cutoff Dubey et al. (2000) Burridge et al. (1979) Burridge et al. (1979) Dubey et al. (2001) Prestrud et al. (2007) Arene (1999) Franti et al. (1975) Aubert et al. (2008) Quist et al. (1995) Lindsay et al. (1994) Franti et al. (1976) Burridge et al. (1979) Burridge et al. (1979) Cotteleer and Famer´ee (1978) Niederehe (1964) Mandelli and Persiani (1966) Mushi et al. (2001) Tsai et al. (2006) Feldman and Sabin (1949) Gibson and Eyles (1957) Jacobs et al. (1952) Kirkpatrick et al. (1990) Pendergraph (1972) Wallace (1973) Frenkel et al. (1995) Kirkpatrick et al. (1990) Kirkpatrick et al. (1990) Kirkpatrick et al. (1990) Aubert et al. (2008) Aubert el al. (2008) Frenkel et al. (1995) Burridge et al. (1979) Franti et al. (1975) Franti et al. (1976) Frenkel et al. (1995) (continues)
Reference
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Spotted Dove Ruddy Ground-Dove Barn Owl
Country
Species
Scientific name
Order
Table 11.2. Serologic prevalence of antibodies to Toxoplasma gondii in wild birds.
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Ramphocelus dimidiatus Thraupis episcopus Thraupis palmarum Quiscalus quiscula Quiscalus mexicanus Agelaius phoeniceus Euphagus cyanocephalus Passer domesticus Passer montanus Sturnus vulgaris Corvus brachyrhynchos
Panama 8 12.5 MAT 1:5 Frenkel et al. (1995) Panama 15 33 MAT 1:5 Frenkel et al. (1995) Panama 3 33 MAT 1:5 Frenkel et al. (1995) USA 27 37 IHAT 1:64 Burridge et al. (1979) Panama 33 33 MAT 1:5 Frenkel et al. (1995) USA 31 6.4 IHAT 1:64 Franti et al. (1975) USA 4 25 IHAT 1:64 Franti et al. (1975) Czech Republic 227 12.3 IFAT 1:10 Liter´ak et al. (1997) Czech Republic 41 4.9 IFAT 1:10 Liter´ak et al. (1997) USA 563 4.8 IHAT 1:64 Haslett and Schneider (1978) USA 74 14 IHAT 1:64 Franti et al. (1976)
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Source: Modified from Dubey (2002). Note: It is not accurate to compare distribution, prevalence, and host distribution of Toxoplasma gondii based on information in this table because different serological tests were used, different cutoff values were employed, and different numbers of birds were tested. None of the serological tests have been validated for use in wild birds using isolation of T. gondii from naturally infected animals as the standard. Most of the information gathered in these reports is from opportunistic samples rather than planned surveys. DT, dye test; IFAT, indirect fluorescent antibody test; IHAT, indirect hemagglutination test; LAT, latex agglutination test; MAT, modified agglutination test.
Crimson-backed Tanager Blue-gray Tanager Palm Tanager Common Grackle Great-tailed Grackle Red-winged Blackbird Brewer’s Blackbird House Sparrow Eurasian Tree Sparrow European Starling American Crow
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considered more virulent in mice than are Types II and III. However, there are no firm data indicating whether pathogenicity in mice reflects pathogenicity in other hosts, including birds (Dubey 2006). In the only outbreak of acute toxoplasmosis in avian hosts where associated T. gondii strains were genotyped, isolates from five Blacked-winged Lories (Eos cyanogenia) were identified as Type III (Dubey et al. 2004). It is likely that all T. gondii isolates in nature are capable of causing illness under appropriate conditions. The name Toxoplasma (toxon = arc and plasma = form) is derived from the crescent shape of the tachyzoite stage (Figure 11.1a). There are three infectious stages of T. gondii that are linked in a cycle: tachy-
zoites (in groups), bradyzoites (in tissue cysts), and sporozoites (in oocysts) (Figure 11.2). The tachyzoite is often crescent-shaped with a pointed anterior end and a rounded posterior end and measures approximately 2 × 6 μm in size in smears (Figure 11.1a). It has a pellicle (outer covering) and several organelles including subpellicular microtubules, a mitochondrion, endoplasmic reticulum, a Golgi apparatus, an apicoplast, ribosomes, a micropore, and a well-defined nucleus (Dubey and Beattie 1988). The nucleus is usually situated toward the central area of the cell. Tachyzoites vary in shape and size, depending on the stage of division or plane of section. Dividing tachyzoites are often globular in shape
Figure 11.1. Developmental stages of Toxoplasma gondii from experimentally infected Budgerigars (Melopsittacus undulatus). (a) Tachyzoites in impression smear of intestine. Note crescent-shaped individual tachyzoites (arrowheads) and dividing tachyzoite (arrow). Giemsa stain. (b) Section of small intestine showing tachyzoites (arrowheads) and necrosis of the lamina propria cells and enterocytes. Note faint staining of tachyzoites. Hematoxylin and eosin stain. (c) Section of small intestine after immunohistochemical labeling with antibodies to T. gondii. Numerous tachyzoites (arrowhead) are darkly stained. Note differences in sizes of tachyzoites in parts (a)–(c). (d) Section of cerebrum showing granulomatous inflammation surrounding a tissue cyst. Note the thin cyst wall (arrow) and terminal nuclei in bradyzoites (arrowhead). From Dubey and Hamir (2002).
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Figure 11.2. Life cycle of Toxoplasma gondii. From Dubey and Beattie (1988). and bigger in size than undividing ones (Figure 11.1a). In histological sections stained with hematoxylin and eosin, tachyzoites are often globular and only about 2 μm in diameter. They are difficult to distinguish from degenerating host cells (Figure 11.1b). Tachyzoites in tissue sections labeled with T. gondii-specific antibodies are often larger than those in hematoxylin and eosinstained sections (Figure 11.1c). Bradyzoites are a slowly growing developmental stage of T. gondii and are enclosed in an elastic membrane. This entire structure is called a tissue cyst. Tissue cysts vary in size from 5 to 70 μm (Figure 11.1d). Although tissue cysts may develop in visceral organs, including lungs, liver, and kidneys, they are more prevalent in muscular and neural tissues, including the brain (Figure 11.1d) eye, skeletal, and cardiac muscle. Intact tissue cysts probably persist for the life of the host. A tissue cyst may enclose hundreds of the bradyzoites that are approximately 7 × 1.5 μm in size. Bradyzoites differ structurally only slightly from tachyzoites. They have a nucleus that is situated to-
ward the posterior end of the cell, whereas the nucleus in tachyzoites is more centrally located. The rhoptries in well-developed bradyzoites are electron-dense, whereas in tachyzoites they are electron-lucent (Dubey et al. 1998a). EPIZOOTIOLOGY Birds may acquire infections with T. gondii by ingestion of oocysts from the environment or by ingestion of tissue-inhabiting stages of the parasite in their prey. The oocyst is the environmentally resistant stage of the parasite and is excreted only by cats. In freshly passed feline feces, oocysts are unsporulated and noninfective. Unsporulated oocysts are subspherical to spherical and measure 10 × 12 μm in diameter. They sporulate and become infectious outside the cat within 1–5 days depending on aeration and temperature. Sporulated oocysts contain two ellipsoidal sporocysts. Each sporocyst contains four sporozoites that measure 2 × 6–8 μm in size. Carnivorous birds are more likely to become infected by ingesting infected tissues from their prey and
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are expected to have a high prevalence of T. gondii (Table 11.1). By contrast, ground-feeding avian species are most likely to become infected by ingesting oocysts from contaminated soil. Contamination of the environment by oocysts is widespread as oocysts are shed by all members of the Felidae. Domestic cats are probably the major source of environmental contamination as oocyst formation is greatest in these hosts and they are extremely common. While as few as 1% of cats may be shedding T. gondii oocysts at any given time, a cat may excrete millions of oocysts after ingesting only one bradyzoite or one tissue cyst. Since many tissue cysts may be present in one infected mouse or bird (Dubey 2001), numbers of excreted oocysts may be even higher. Congenital infection can also occur in cats, and congenitally infected kittens can excrete oocysts, thus providing another common source for contamination. Sporulated oocysts survive for long periods under most environmental conditions. They can survive in moist soil, for example, for months and even years (Dubey and Beattie 1988; Dubey 2004) and can be mechanically spread by flies, co*ckroaches, dung beetles, and earthworms. Environmental resistance of oocysts and the enormous numbers that are shed in the feces of domestic cats assure widespread contamination (Dubey 2004). Cats are thought to become infected by eating both birds and rodents. Hence, prevalence of infection in cats is determined by prevalence of infection in the local avian and rodent populations (Ruiz and Frenkel 1980). As environmental contamination with oocysts increases, prey animals are more likely to be infected, leading to higher rates of infection in cats. CLINICAL SIGNS Clinical signs of avian toxoplasmosis are nonspecific and cannot be used to make a definitive diagnosis. These signs include anorexia, depression, dull ruffled feathers, diarrhea, and dyspnoea. Unusual clinical signs of toxoplasmosis have been observed in Island Canaries (Serinus canaria), including cataracts and blindness (Vickers et al. 1992; Lindsay et al. 1995; Gibbens et al. 1997; Williams et al. 2001). In ocular cases, the eyes became dull, sightless, closed, and sunken into the head, but birds were otherwise alert and continued to feed (Vickers et al. 1992; Lindsay et al. 1995; Gibbens et al. 1997; Williams et al. 2001). In one outbreak, half of the affected canaries had evidence of central nervous system involvement, including head twitch and disoriented walking in circles. PATHOGENESIS AND PATHOLOGY After ingestion, sporozoites from oocysts penetrate intestinal epithelial cells and multiply as tachyzoites in
cells of the lamina propria. Toxoplasma gondii may spread to distant organs via lymphatics or blood. A host may die of acute toxoplasmosis because of necrosis of the intestine and associated lymphoid tissues before other organs are severely damaged (Figure 11.3), but more often recovers. Focal areas of necrosis may develop in many organs. By about the third week after infection, tachyzoites begin to disappear from visceral tissues and may localize as tissue cysts in neural and muscular tissues during recovery. Inflammatory lesions may persist in the central nervous system (Figure 11.1d). Toxoplasma gondii causes tissue necrosis by active destruction of host cells; it does not produce a toxin. Tachyzoites can be found in lesions (Figure 11.3), often at the periphery of necrotic foci. In chronic lesions, tissue cysts may be found. The presence of tissue cysts in the absence of lesions indicates only persistent infection and not the clinical disease. In naturally infected animals, lesions predominate in the liver, lungs, spleen, brain, and eyes (Figures 11.4–11.6) (Table 11.3), and occasionally in adrenal glands and bursa of Fabricius. Necrosis of hepatocytes and mononuclear cell infiltrations in periportal areas and sinusoids are characteristic of hepatic lesions (Figure 11.7). Many tachyzoites and occasionally tissue cysts are present among lesions. In lung tissue, pneumonitis is characterized by necrosis of pulmonary parenchyma and infiltrations of mononuclear cells. Tachyzoites are often present in pulmonary macrophages. Necrosis of splenic parenchyma is the primary lesion in spleen tissue. Neural lesions consist of necrosis of neuropils, gliosis, and perivascular infiltrations of mononuclear cells. Ocular lesions in canaries are characterized by acute, severe diffuse choroiditis and retinal necrosis (Figure 11.6), panophthalmitis, optic neuritis, cataracts, and osseous replacement of the globe. Tachyzoites have been found in the choroid, retina, vitreous, and even in the lens (Vickers et al. 1992). Many factors may determine the outcome of toxoplasmosis, including stage of the parasite ingested, dose, and host species. Oocyst-acquired infections are generally thought to be more clinically severe in birds than infection from the ingestion of infected tissues. The number of organisms ingested may not be the determining factor because hundreds of bradyzoites are contained in a tissue cyst versus only eight sporozoites in an oocyst. Currently, there is no test that can determine whether a host was infected with oocysts or tissue cysts. The following information is derived from avian species with experimental oral infections with oocysts or tissue cysts; data derived from birds infected by parenteral routes are not comparable. Oral infection of birds with tissue cysts has been reported in only one
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Toxoplasma
(a)
(b)
(c)
Figure 11.3. Sections of small intestines from Rock Partridges (Alectoris graeca) that were fed Toxoplasma gondii oocysts. Hematoxylin and eosin stain. (a) Desquamation (arrow) of intestinal contents into the lumen at 10 days postinfection. (b) Necrosis of lamina propria (arrow) with intact epithelium (arrowheads) at 7 days postinfection. (c) Higher magnification of part (b). Note tachyzoites (arrows). From Dubey et al. (1995).
study. Great Horned Owls (Bubo virginianus), Barred Owls, (Strix varia), and Eastern Screech-Owls (Megascops asio) fed tissues of rats containing many tissue cysts of the GT1 and CT1 (Type I, mouse virulent strains of T. gondii) became infected but did not develop clinical signs (Dubey et al. 1992). Rock Partridges (Alectoris graeca) fed 10,000 oocysts of the GT1 strain (the strain fed to owls) died of peracute toxoplasmosis 4–6 days postinoculation, whereas clinical signs were less severe in Rock Partridges fed the Me-49 (Type II, mouse avirulent) strain of T. gondii. However, even 10 oocysts of the Me-49 strain killed 2 of 6 Rock Partridges 12 and 13 days postinoculation (Dubey et al. 1995). It is interesting that Red-legged Partridges (Alectoris rufa) are less susceptible than Rock Partridges. Red-legged Partridges fed 10,000 oocysts of the OV-51/95 (genotype unknown) became infected but did not develop clinical signs (Mart´ınez-Carrasco et al. 2004, 2005, 2006). Japanese Quail (Coturnix japonica), Northern Bobwhite (Colinus virginianus), and domestic turkeys were more susceptible to toxoplasmosis than Ringnecked Pheasants (Phasianus colchicus) (Dubey et al. 1993a, b, 1994a, b). Although severe toxoplasmosis
has been reported in some psittacine species (Table 11.3), Budgerigars (Melopsittacus undulatus) are relatively resistant to clinical toxoplasmosis (Dubey and Hamir 2002; Kajerov´a et al. 2003).
DIAGNOSIS Serologic, histopathologic, immunohistochemical, and molecular methods can aid diagnosis, and this subject has been discussed in detail elsewhere (Dubey and Beattie 1988; Dubey 1993; Dubey and Odening 2001). Although the dye test is the most specific test for the detection of antibodies to T. gondii in humans, it does not work with sera from most avian species (Frenkel 1981; Dubey 2002). The latex agglutination test (LAT), indirect hemagglutination test (IHAT), and the modified agglutination test (MAT) have been evaluated in experimentally infected, domestic Wild Turkeys (Meleagris gallopavo), Ring-necked Pheasants, Rock Partridges, Red-legged Partridges, Northern Bobwhites, Japanese Quail, Great Horned Owls, Barred Owls, and Eastern Screech-Owls. The MAT is the most specific and sensitive test (Dubey et al. 1992, 1993a, b, 1994a, b, 1995; Martinez-Carrasco et al. 2004) and is simple,
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Figure 11.4. Sections of brain of birds fed Toxoplasma gondii oocysts. Hematoxylin and eosin stain. (a) Cerebrum of a Japanese Quail (Coturnix japonica) at 16 days postinfection. Note mononuclear cell infiltrate and necrosis of neutrophils associated with tachyzoites (arrows). From Dubey et al. (1994b). (b) Cerebrum of a Budgerigar (Melopsittacus undulatus) at 35 days postinfection. Tissue cyst (arrow) is located at the periphery of the lesion in a glial nodule. From Dubey and Hamir (2002). reliable, does not require specific reagents, and works well with plasma (Dubey et al. 1992). A titer of 1:25 is considered indicative of infection, but titer intensity does not reflect clinical status. The LAT also works well with avian sera but titers can decline to undetectable levels (Kajerov´a et al. 2003). Hematologic values are unaffected and of little use for diagnosing infection with Toxoplasma (Kajerova et al. 2003), although enzymes indicative of tissue necrosis (e.g., lactic dehydrogenase) may help indicate involvement of specific organ systems. Toxoplasma gondii DNA can be detected by polymerase chain reaction methodology in fresh, frozen,
and sometimes in fixed tissue by using T. gondiispecific primers. Fixation in formalin for a long period may denature T. gondii DNA. In most cases, however, diagnosis is made by histologic examination of tissues submitted for necropsy that may have already been fixed in buffered neutral 10% formalin. A preliminary diagnosis can be made by examining Giemsa-stained impression smears of affected tissues (Figure 11.1a). T. gondii tachyzoites are crescentic to globular in smears, depending on the stage of division. However, in histologic sections, tachyzoites are globular to oval and about half the size of those in smears (Figure 11.1b). Toxoplasma gondii tissue cysts are often globular, have a thin cyst wall (<0.5 μm), and enclose small (5 μm), slender bradyzoites (Figure 11.1d). The bradyzoites are periodic acid Schiff positive and there are no intracystic septa (Dubey and Beattie 1988). Presence of tissue cysts in the absence of lesions generally indicates latent infection. Immunohistochemical staining with T. gondii-specific antibodies can aid diagnosis (Figure 11.1c). Polyclonal antibodies raised against whole parasites are often superior to monoclonal antibodies for immunohistochemical diagnosis of toxoplasmosis in tissue sections. Preservation of tissues in 10% formalin does not interfere with the immunohistochemical reaction. Atoxoplasma and Sarcocystis are two parasite genera that should be considered in the differential diagnosis of avian toxoplasmosis. Proliferative stages (merozoites) of Atoxoplasma sp. are smaller than T. gondii tachyzoites, both in smears and in histologic sections (Figure 11.8). Other parasites related to Atoxoplasma may also be found in birds (Baker et al. 1996; Speer et al. 1997). Sarcocystis falcatula and S. falcatula-like infections can cause generalized disease in birds, especially in passerines and psittacines (Smith et al. 1989; Hillyer et al. 1991). Some unidentified species of Sarcocystis can cause neural and myocardial sarcocystosis in association with development of merents in affected tissues (Gustafsson et al. 1997; Dubey et al. 2001). Neural sarcocystosis can simulate toxoplasmosis and has been reported in a Northern Goshawk (Accipiter gentilis atricapillus), Wild Turkeys, Eurasian Capercaillie (Tetrao urogallus), a Straw-necked Ibis (Threskiornis spinicollis), Golden Eagle (Aquila chrysaetos), and a Bald Eagle (Haliaeetus leucocephalus) (Aguilar et al. 1991; Dubey et al. 1991, 1998b, 2000, 2001; Teglas et al. 1998; Olson et al. 2007). PUBLIC HEALTH AND DOMESTIC ANIMAL CONCERNS Toxoplasma gondii infection is widespread among humans and its prevalence varies widely from place to
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(a)
215
(b)
Figure 11.5. Spleen of a Rock Partridge (Alectoris graeca) 10 days after the bird was fed Toxoplasma gondii oocysts. (a) Note pale foci (arrows) on the surface and in the parenchyma. Unstained. (b) Spleen section stained with hematoxylin and eosin. Note coagulative necrosis (arrow) and tachyzoites (arrowheads). From Dubey et al. (1995). place. Most infections in humans are asymptomatic, but at times the parasite can produce devastating disease. Toxoplasma gondii is capable of causing severe disease in animals other than humans (Dubey and Beattie 1988; Tenter et al. 2000). Toxoplasmosis causes great losses in sheep and goats and may cause embryonic death and resorption, fetal death and mummification, abortion, stillbirth, and neonatal death in these animals. Wild birds probably play a minor role as a source of infections for both humans and domestic animals. Most cases originate from exposure to oocysts and consumption of raw or undercooked meat. WILDLIFE POPULATION IMPACTS Virtually any species of bird can be an intermediate host for T. gondii and a source of infection for cats. There are no firm data on the impact of T. gondii on
decline or mortality of birds in the wild, but this organism can pose a significant threat to small populations of critically endangered species. Approximately 20% of the wild population of the Hawaiian Crow died from toxoplasmosis in the late 1990s during attempts to restore this species in former habitat on the island of Hawaii (Work et al. 2000).
TREATMENT, CONTROL, AND PREVENTION Sulfadiazine and pyrimethamine (Daraprim) are two drugs widely used for therapy of toxoplasmosis. These drugs are effective during acute stages of the disease when there is active multiplication of the parasite, but will not usually eradicate infection. Sulfadiazine and pyrimethamine have little effect on subclinical
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Figure 11.7. Sections of liver from a Barred Owl (Strix varia) naturally infected with Toxoplasma gondii and labeled by immunohistochemical staining with anti-T. gondii antibodies. Note foci of necrosis (arrows) and numerous black tachyzoites (arrowheads). Inset: Higher magnification view of tachyzoites (arrow). From Mikaelian et al. (1997).
Figure 11.6. Chorioretinitis in a naturally infected Island Canary (Serinus canaria) with fatal toxoplasmosis. Hematoxylin and eosin stain. (a) Fragmentation (large arrow) and detachment (small arrow) of retina in vitreous humor. (b) Higher magnification of a portion of retina in the vitreous humor. Note tachyzoites (arrows). From Vickers et al. (1992).
infections, but the growth of tissue cysts in mice has been restrained with sulfonamides (Beverley 1958). Lindsay et al. (1995) successfully treated T. gondiiassociated blindness in canaries with sulfadiazine and trimethoprim. Diclazuril, a benzene acetonitride, was effective when used to treat toxoplasmosis in Hawaiian Crows (Work et al. 2000). To prevent infection with T. gondii, aviaries should be made cat-proof. Feed should be stored in covered containers to prevent contamination with cat feces, and meat fed to birds should be cooked thoroughly to 67◦ C. If cooking is not practical, meat should be frozen at −12◦ C for at least 24 h. Freezing meat in a household freezer kills most, if not all, T. gondii. There is no direct transmission of T. gondii from birds to humans or other birds other than by eating infected meat.
Figure 11.8. Atoxoplasma sp. merozoites (a) and Toxoplasma gondii tachyzoites (b) in impression smears of avian tissues stained with Giemsa. (a) Individual (arrow) and dividing (arrowheads) Atoxoplasma in spleen of a Bali Mynah (Leucopsar rothschildi). (b) Two T. gondii tachyzoites (arrow) within a mononuclear cell in the lung of a Rock Partridge (Alectoris graeca). From Dubey (2002).
Species
217
Strigiformes Psittaciformes
Columbiformes
Spheniscus humboldti Spheniscus magellanicus Spheniscus demersus Eudyptula minor Sula sula Anseranus semipalmata Branta sandvicensis Anas platyrhynchos Meleagris gallopavo
Scientific name USA USA USA Australia USA
Country 4 2 1 1 1 2 2 Many 1 1 3 1 1 1 3 1 4 2 1 4 3 1 1 1 1 1 1 5
N
C C C W C
C C C C C C C C W W C W C C C C C C C C C C
Died
Reference
Ratcliffe and Worth (1951) Ratcliffe and Worth (1951) Ratcliffe and Worth (1951) H, IHC Mason et al. (1991) H, IHC Work et al. (2002) H, IHC Dubey et al. (2001) H, IHC Work et al. (2002) H Boehringer et al. (1962) H, TEM Howerth and Rodenroth (1985) H, IHC Quist et al. (1995) H Pokorny (1955) H, IHC Work et al. (2002) H, I Kageruka and Willaert (1971) H Tackaert-Henry and Kageruka (1977) H, I Poelma and Zwart (1972) H Ratcliffe and Worth (1951) H Tackaert-Henry and Kageruka (1977) H, I Poelma and Zwart (1972) H, I Poelma and Zwart (1972) H, I Poelma and Zwart (1972) H, IHC Hartley and Dubey (1991) H, IHC Hartley and Dubey (1991) H, I Poelma and Zwart (1972) H, IHC Mikaelian et al. (1997) H, IHC Hartley and Dubey (1991) H, IHC Hartley and Dubey (1991) H, IHC, TEM Howerth et al. (1991) H, I Howerth et al. (1991) (continues)
H
Method
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USA Argentina USA USA Gray Partridge Perdix perdix Czech Republic Erckel’s Francolin Francolinus erckelii USA Western Crowned-Pigeon Goura cristata Belgium Belgium The Netherlands Victoria Crowned-Pigeon Goura victoria USA Belgium The Netherlands Pied Imperial-Pigeon Ducula bicolor The Netherlands Southern Crowned-Pigeon Goura scheepmakeri The Netherlands Torresian Imperial-Pigeon Ducula spilorrhoa Australia Wonga Pigeon Leucosarcia melanoleuca Australia Luzon Bleeding-heart Gallicolumba luzonica The Netherlands Barred Owl Strix varia Canada Regent Parrot Polytelis anthopeplus Australia Superb Parrot Polytelis swainsonii Australia Red Lorry Eos bornea USA USA
Sphenisciformes Humboldt Penguin Magellanic Penguin Jackass Penguin Little Penguin Pelecaniformes Red-footed Booby Anseriformes Magpie Goose Hawaiian Goose Mallard Galliformes Wild Turkey
Order
Table 11.3. Reports of toxoplasmosis at necropsy from birds that died in captivity (C) or in the wild (W).
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Species
Scientific name
1 3
Australia Australia Australia 3 USA 1 USA 5 Australia 1 Australia 1 Australia 1 Australia 12/24* Australia 2/40* United Kingdom 2/44* United States 2/9*
1
N
The Netherlands
Country
C C C C C C C
C C
C C
C
Died
H, IHC H, IHC A, H, I, IHC H, IHC H, IHC H, IHC H, I, IHC H H H
H, IHC H, IHC
H, I
Method
Hartley et al. (2008) Gerhold and Yabsley (2007) Work et al. (2000) Hartley and Dubey (1991) Hartley and Dubey (1991) Hartley and Dubey (1991) Vickers et al. (1992) Lindsay et al. (1995) Gibbens et al. (1997) Williams et al. (2001)
Hartley and Dubey (1991) Hartley et al. (2008)
Dubey et al. (2004) Poelma and Zwart (1972)
Reference
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218
Note: Methods of diagnosis include antibodies to Toxoplasma gondii in serum (A), histology (H), immunohistochemical staining with anti-T. gondii antibodies (IHC), isolation in mice (I), and transmission electron microscopy (TEM). In most cases, birds died from pneumonia, hepatitis, splenitis, encephalitis, and/or ophthalmitis. In addition to data compiled in this table, clinical toxoplasmosis has been reported in unspecified species of pigeons as well as Rock Pigeons (Columba livia) from Brazil (Carini, 1911; Pires and Dos Santos 1934; Reis and N´obrega 1936; Springer 1942), Democratic Republic of Congo (Wiktor 1950), Ecuador (Rodriguez 1954), Italy (de Mello 1915; Alosi and Iannuzzi 1966), Mexico (Paasch 1983), Panama (Johnson 1943), Scandinavia (Siim et al. 1963), Venezuela (Vogelsang and Gallo 1954), Uruguay (Cassamagnaghi et al. 1952, 1977), and from an unspecified source (Hubbard et al. 1986). Toxoplasmosis has also been reported from canaries and finches from Uruguay (Cassamagnaghi et al. 1952, 1977) and Italy (Parenti et al. 1986) and from Budgerigars (Melopsittacus undulatus) from Switzerland (Galli-Valerio 1939). The number of birds affected and the methods of diagnosis were not detailed in several of these reports. * Fraction of total that were examined by histopathology.
Black-winged Lorry Rainbow Lorikeet
Eos cyanogenia Trichoglossus haematodus moluccanus Crimson Rosella Platycercus elegans Red-fronted Parakeet Cyanoramphus novaezelandiae Yellow-fronted Parakeet Cyanoramphus auriceps Piciformes Red-bellied Woodpecker Melanerpes carolinus Passeriformes Hawaiian Crow Corvus hawaiiensis Satin Bowerbird Ptilonorhynchus violaceus Regent Bowerbird Sericulus chrysocephalus Red-whiskered Bulbul Pycnonotus jocosus Island Canary Serinus canaria
Order
Table 11.3. (Continued)
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Toxoplasma LITERATURE CITED Aguilar, R. F., D. P. Shaw, J. P. Dubey, and P. Redig. 1991. Sarcocystis-associated encephalitis in an immature northern goshawk (Accipiter gentilis atricapillus). Journal of Zoo and Wildlife Medicine 22:466–469. Alosi, C., and L. Iannuzzi. 1966. Toxoplasmosi acuta spontanea dei colombi. Segnalazione di un focolaio a Messina. Acta Medicina Veterinaria 12:265–273. Arene, F. O. I. 1999. Seroprevalence of Toxoplasma gondii in vultures (Pseudogyps africanus) from eastern Nigeria. Acta Parasitologica 44:79–80. Aubert, D., M. E. Terrier, A. Dum`etre, J. Barrat, and I. Villena. 2008. Prevalence of Toxoplasma gondii in raptors from France. Journal of Wildlife Distribution 44, 172–173. Baker, D. G., C. A. Speer, A. Yamaguchi, S. M. Griffey, and J. P. Dubey. 1996. An unusual coccidian parasite causing pneumonia in a northern cardinal (Cardinalis cardinalis). Journal of Wildlife Diseases 32:130–132. Beverley, J. K. A. 1958. A rational approach to the treatment of toxoplasmic uveitis. Transactions of the Ophthalmological Societies of the United Kingdom 78:109–121. Burridge, M. J., W. J. Bigler, D. J. Forrester, and J. M. Hennemann. 1979. Serologic survey for Toxoplasma gondii in wild animals in Florida. Journal of the American Veterinary Medical Association 175:964–967. Carini, A. 1911. Infection spontan´ee du pigeon et du chien due au Toxoplasma cuniculi. Bulletin de la Soci´et´e de Pathologie Exotique 4:518–519. Cassamagnaghi, A., A. Bianchi Bazerque, R. Scelza, and H. Ferrando. 1952. La toxoplasmosis. Su incorporacion en la patologia Uruguaya. Reconocimento de dos cepas en nuestras aves dom´esticas. Su trasmisi´on y car´acter infeccioso para los mam´ıferos. Bollettin Ministerio de Ganader´ıa y Agricultura 33:34–38. Cassamagnaghi, A., A. Bianchi Bazerque, R. Scelza, and H. Ferrando. 1977. La toxoplasmosis. Su incorporacion en la patologia Uruguaya. Annali della Facolta di Medicina Veterinaria di Uruquay 14:27–46. Cat´ar, G. 1974. Toxoplazm´oza v ekologick´ych podmienkach na slovensku. Biologick´e Pr´ace (Bratislava) 20:1–138. Cotteleer, C., and L. Famer´ee. 1978. Parasites intestinaux et anticorps antitoxoplasmiques chez les colombins en Belgique. Schweizer Archiv fur Tierheilkunde 120:181–187. de Mello, F. 1915. Preliminary note on a new haemogregarine found in the pigeon’s blood. Indian Journal of Medical Research 3:93–94. Dubey, J. P. 1993. Toxoplasma, Neospora, Sarcocystis, and other tissue cyst-forming coccidia of humans and animals. In Parasitic Protozoa, Vol. VI, J. P. Kreier (ed.). Academic Press, New York, pp. 1–158.
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Dubey, J. P. 2001. Oocyst shedding by cats fed isolated bradyzoites and comparision of infectivity of bradyzoites of the VEG strain Toxoplasma gondii to cats and mice. Journal of Parasitology 87:215–219. Dubey, J. P. 2002. A review of toxoplasmosis in wild birds. Veterinary Parasitology, 106:121–153. Dubey, J. P. 2004. Toxoplasmosis—A waterborne zoonosis. Veterinary Parasitology 126:57–72. Dubey, J. P. 2006. Comparative infectivity of oocysts and bradyzoites of Toxoplasma gondii for intermediate (mice) and definitive (cats) hosts. Veterinary Parasitology 140:69–75. Dubey, J. P. 2007. The history and life cycle of Toxoplasma gondii. In Toxoplasma gondii. The Model Apicomplexan: Perspectives and Methods, L. Weiss and K. Kim (eds). Academic Press, San Diego, CA, pp. 1–17. Dubey, J. P., and C. P. Beattie. 1988. Toxoplasmosis of Animals and Man. CRC Press, Boca Raton, FL, 220 pp. Dubey, J. P., and K. Odening. 2001. Toxoplasmosis and related infections. In Parasitic Diseases of Wild Mammals, B. Samuel, M. Pybur and A. M. Kocan (eds). Iowa State University Press, Ames, IA, pp. 478–519. Dubey, J. P., and A. N. Hamir. 2002. Experimental toxoplasmosis in budgerigars (Melopsittacus undulatus). Journal of Parasitology 88:514–519. Dubey, J. P., S. L. Porter, A. L. Hattel, D. C. Kradel, M. J. Topper, and L. Johnson. 1991. Sarcocystosis-associated clinical encephalitis in a golden eagle (Aquila chrysaetos). Journal of Zoo and Wildlife Medicine 22:233–236. Dubey, J. P., S. L. Porter, F. Tseng, S. K. Shen, and P. Thulliez. 1992. Induced toxoplasmosis in owls. Journal of Zoo and Wildlife Medicine 23:98–102. Dubey, J. P., M. D. Ruff, O. C. H. Kwok, S. K. Shen, G. C. Wilkins, and P. Thulliez. 1993a. Experimental toxoplasmosis in bobwhite quail (Colinus virginianus). The Journal of Parasitology 79:935–939. Dubey, J. P., M. E. Camargo, M. D. Ruff, G. C. Wilkins, S. K. Shen, O. C. H. Kwok, and P. Thulliez. 1993b. Experimental toxoplasmosis in turkeys. The Journal of Parasitology 79:949–952. Dubey, J. P., M. D. Ruff, G. C. Wilkins, S. K. Shen, and O. C. H. Kwok. 1994a. Experimental toxoplasmosis in pheasants (Phasianus colchicus). Journal of Wildlife Diseases 30:40–45. Dubey, J. P., M. A. Goodwin, M. D. Ruff, O. C. H. Kwok, S. K. Shen, G. C. Wilkins, and P. Thulliez. 1994b. Experimental toxoplasmosis in Japanese quail. Journal of Veterinary Diagnostic Investigation 6:216–221. Dubey, J. P., M. A. Goodwin, M. D. Ruff, S. K. Shen, O. C. H. Kwok, G. L. Wilkins, and P. Thulliez. 1995. Experimental toxoplasmosis in chukar partridges (Alecotoris graeca). Avian Pathology 24:95–107.
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Dubey, J. P., D. S. Lindsay, and C. A. Speer. 1998a. Structure of Toxoplasma gondii tachyzoites, bradyzoites and sporozoites, and biology and development of tissue cysts. Clinical Microbiology Reviews 11:267–299. Dubey, J. P., E. Rudb¨ack, and M. J. Topper. 1998b. Sarcocystosis in capercaillie (Tetrao urogallus) in Finland: Description of the parasite and lesions. Journal of Parasitology 84:104–108. Dubey, J. P., W. B. Scandrett, O. C. H. Kwok, and A. A. Gajadhar. 2000. Prevalence of antibodies to Toxoplasma gondii in ostriches (Struthio camelus). The Journal of Parasitology 86:623–624. Dubey, J. P., M. M. Garner, M. M. Willette, K. L. Batey, and C. H. Gardiner. 2001. Disseminated toxoplasmosis in magpie geese (Anseranas semipalmata) with large numbers of tissue cysts in livers. The Journal of Parasitology 87:219–223. Dubey, J. P., D. H. Graham, E. Dahl, M. Hilali, A. El-Ghaysh, C. Sreekumar, O. C. H. Kwok, S. K. Shen, and T. Lehmann. 2003. Isolation and molecular characterization of Toxoplasma gondii from chickens and ducks from Egypt. Journal of Parasitology 114, 89–95. Dubey, J. P., P. G. Parnell, C. Sreekumar, M. C. B. Vianna, R. W. de Young, E. Dahl, and T. Lehmann. 2004. Biologic and molecular characteristics of Toxoplasma gondii isolates from striped skunk (Mephitis mephitis), Canada goose (Branta canadensis), blacked-winged lory (Eos cyanogenia), and cats (Felis catus). Journal of Parasitology 90:1171–1174. Dubey, J. P., D. M. Webb, N. Sundar, G. V. Velmurugan, L. A. Bandini, O. C. H. Kwok, and C. Su. 2007. Endemic avian toxoplasmosis on a farm in Illinois: clinical disease, diagnosis, biologic and genetic characteristics of Toxoplasma gondii isolates from chickens (Gallus domesticus), and a goose (Anser anser). Veterinary Parasitology 148, 207–212. Feldman, H. A., and A. B. Sabin. 1949. Skin reactions to toxoplasmic antigen in people of different ages without known history of infection. Pediatrics 4:798–804. Finlay, P., and R. D. Manwell. 1956. Toxoplasma from the crow, a new natural host. Experimental Parasitology 5:149–153. Franti, C. E., G. E. Connolly, H. P. Riemann, D. E. Behymer, R. Ruppanner, C. M. Willadsen, and W. Longhurst. 1975. A survey for Toxoplasma gondii antibodies in deer and other wildlife on a sheep range. Journal of the American Veterinary Medical Association 167:565–568. Franti, C. E., H. P. Riemann, D. E. Behymer, D. Suther, J. A. Howarth, and R. Ruppanner. 1976. Prevalence of Toxoplasma gondii antibodies in wild and domestic animals in northern California. Journal of the
American Veterinary Medical Association 169:901–906. Frenkel, J. K. 1981. False-negative serologic tests for Toxoplasma in birds. The Journal of Parasitology 67:952–953. Frenkel, J. K., K. M. Hassanein, R. S. Hassanein, E. Brown, P. Thulliez, and R. Quintero-Nunez. 1995. Transmission of Toxoplasma gondii in Panama City, Panama: A five-year prospective cohort study of children, cats, rodents, birds, and soil. American Journal of Tropical Medicine and Hygiene 53:458–468. Galli-Valerio, B. 1939. Sur une toxoplasmiase du Melopsittacus undulatus Shaw. Schweizer Archiv fur Tierheilkunde 81:458–460. Gerhold, R. W., and M. J. Yabsley. 2007. Toxoplasmosis in a red-bellied woodpecker (Melanerpes carolinus). Avian Diseases 51:992–994. Gibbens, J. C., E. J. Abraham, and G. MacKenzie. 1997. Toxoplasmosis in canaries in Great Britain. The Veterinary Record 140:370–371. Gibson, C. L., and D. E. Eyles. 1957. Toxoplasma infections in animals associated with a case of human congenital toxoplasmosis. American Journal of Tropical Medicine and Hygiene 6:990–1000. Gustafsson, K., M. Book, J. P. Dubey, and A. Uggla. 1997. Meningoencephalitis in capercaillie (Tetrao urogallus L.) caused by a Sarcocytis-like organism. Journal of Zoo and Wildlife Medicine 28:280–284. Hartley, W. J., and J. P. Dubey. 1991. Fatal toxoplasmosis in some native Australian birds. Journal of Veterinary Diagnostic Investigation 3:167–169. Hartley, W. J., R. Booth, R. Slocombe, and J. P. Dubey. 2008. Lethal toxoplasmosis in an aviary of kakarikis (Cyanoramphus supp.) in Australia. Journal of Parasitology. In press. Haslett, T. M., and W. J. Schneider. 1978. Occurrence and attempted transmission of Toxoplasma gondii in European starlings (Sturnus vulgaris). Journal of Wildlife Diseases 14:173–175. Hejl´ıcˇ ek, K., F. Prosek, and F. Treml. 1981. Isolation of Toxoplasma gondii in free-living small mammals and birds. Acta Veterinaria Brno 50:233–236. Hillyer, E. V., M. P. Anderson, E. C. Greiner, C. T. Atkinson, and J. K. Frenkel. 1991. An outbreak of Sarcocystis in a collection of psittacines. Journal of Zoo and Wildlife Medicine 22:434–445. Holst, I., and M. Chinchilla. 1990. Development and distribution of cysts of an avirulent strain of Toxoplasma and the humoral immune response in mice. Revista de Biologia Tropical 38:189–193. Howerth, E. W., and N. Rodenroth. 1985. Fatal systemic toxoplasmosis in a wild turkey. Journal of Wildlife Diseases 21:446–449. Howerth, E. W., G. Rich, J. P. Dubey, and K. Yogasundram. 1991. Fatal toxoplasmosis in a red lory (Eos bornea). Avian Diseases 35:642–646.
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Toxoplasma Hubbard, G., W. Witt, M. Healy, and R. Schmidt. 1986. An outbreak of toxoplasmosis in zoo birds. Veterinary Pathology 23:639–641. Jacobs, L., M. L. Melton, and F. E. Jones. 1952. The prevalence of toxoplasmosis in wild pigeons. The Journal of Parasitology 38:457–461. Johnson, C. M. 1943. Immunological and epidemiological investigation under the direction of C. M. Johnson, protozoologist. Annual Report of the Gorgas Memorial Laboratory 15–16. Kageruka, P., and E. Willaert. 1971. Toxoplasma gondii (Nicolle et Manceaux 1908) isole chez Goura cristata pallas et Manis crassicaudata Geoffroy. Acta Zoologica et Pathologica Antverpiensia 52:3–10. Kajerov´a, V., I. Literak, E. Bartova, and K. Sedlak. 2003. Experimental infection of budgerigars (Melopsittacus undulatus) with a low virulent K21 strain of Toxoplasma gondii. Veterinary Parasitology 116:297–304. Kirkpatrick, C. E., B. A. Colvin, and J. P. Dubey. 1990. Toxoplasma gondii antibodies in common barn-owls (Tyto alba) and pigeons (Columba livia) in New Jersey. Veterinary Parasitology 36:177–180. Lehmann, T., P. L. Marcet, D. H. Graham, E. R. Dahl, and J. P. Dubey. 2006. Globalization and the population structure of Toxoplasma gondii. In Proceedings of the National Academy of Sciences 103:11423–11428. Lindsay, D. S., P. C. Smith, F. J. ho*rr, and B. L. Blagburn. 1993. Prevalence of encysted Toxoplasma gondii in raptors from Alabama. The Journal of Parasitology 79:870–873. Lindsay, D. S., P. C. Smith, and B. L. Blagburn. 1994. Prevalence and isolation of Toxoplasma gondii from wild turkeys in Alabama. Journal of the Helminthological Society of Washington 61:115–117. Lindsay, D. S., R. B. Gasser, K. E. Harrigan, D. N. Madill, and B. L. Blagburn. 1995. Central nervous system toxoplasmosis in roller canaries. Avian Diseases 39:204–207. Liter´ak, I., K. Hejlicek, J. Nezval, and C. Folk. 1992. Incidence of Toxoplasma gondii in populations of wild birds in the Czech Republic. Avian Pathology 21:659–665. Liter´ak, I., J. Pinowski, M. Anger, Z. Juricova, H. Kyu-Hwang, and J. Romanowski. 1997. Toxoplasma gondii antibodies in house sparrows (Passer domesticus) and tree sparrows (P. montanus). Avian Pathology 26:823–827. Mandelli, G., and G. Persiani. 1966. Ricerche sierologiche sulla presenza e diffusione della toxoplasmosi new piccioni torraioli (Columba livia). Clinica Veterinaria 89:161–166. Manwell, R. D., and H. P. Drobeck. 1951. Mammalian toxoplasmosis in birds. Experimental Parasitology 1:83–93.
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Martin´ez-Carrasco, C., J. M. Ortiz, A. Bernab´e, M. R. Ruiz deYb´an˜ ez, M. Garijo, and F. D. Alonso. 2004. Serologic response of red-legged partridges (Alectoris rufa) after oral inoculation with Toxoplasma gondii oocysts. Veterinary Parasitology 121:143–149. Mart´ınez-Carrasco, C., A. Bernab´e, J. M. Ortiz, and F. D. Alonso. 2005. Experimental toxoplasmosis in red-legged partridges (Alectoris rufa) fed Toxoplasma gondii oocysts. Veterinary Parasitology 130, 55–60. Mart´ınez-Carrasco, C., A. Bernab´e, J. M. Ortiz, and F. D. Alonso. 2006. Lesiones asociadas a una toxoplasmosis aguda en perdices rojas (Alectoris rufa) infectadas experimentalmente. Anales de Veterinaria de Murcia 22, 93–98. Mason, R. W., W. J. Hartley, and J. P. Dubey. 1991. Lethal toxoplasmosis in a little penguin (Eudyptula minor) from Tasmania. The Journal of Parasitology 77:328 Mikaelian, I., J. P. Dubey, and D. Martineau. 1997. Severe hepatitis resulting from toxoplasmosis in a barred owl (Strix varia) from Qu´ebec, Canada. Avian Diseases 41:738–740. Mushi, E. Z., M. G. Binta, R. G. Chabo, R. Ndebele, and R. Panzirah. 2001. Seroprevalence of Toxoplasma gondii and Chlamydia psittaci in domestic pigeons (Columba livia domestica) at Sebele, Gaborone, Botswana. Onderstepoort Journal of Veterinary Research 68:159–161. Nicolle, C., and L. Manceaux. 1909. Sur un protozoaire nouveau du gondi. Comptes Rendus des Seances de l’Academie des Sciences 148:369–372. Nicolle, M. M. C., and L. Manceaux. 1908. Sur une infection a corps de Leishman (ou organismes voisins) du gondi. Comptes Rendus des Seances de l’Academie des Sciences 147:763–766. Niederehe, H. 1964. Toxoplasma-Infektion bei verwilderten Tauben. Tier¨arztliche Umschau 19:256–257. Olson, E. J., A. W¨unschmann, and J. P. Dubey. 2007. Sarcocystis-associated meningoencephalitis in a bald eagle (Haliaeetus leucocephalus). Journal of Veterinary Diagnostic Investigation 19:564–568. Paasch, M. L. 1983. Toxoplasmosis en palomas. Veterinaria M´exico 14:39–41. Pak, S. M. 1970. A strain of Toxoplasma gondii isolated from the common tern. Contributions on the Natural Nidality of Diseases 3:49. Pak, S. M. 1972. Toxoplasmosis of sparrows. Contributions to the Natural Nidality of Diseases 5:116–125. Pak, S. M. 1976. Toxoplasmosis of birds in Kazakhstan (in Russian). Nauka Publishing, Alma Ata, Kazakhstan, 115 pp. Parenti, E., S. Cerruti Sola, C. Turilli, and S. Corazzola. 1986. Spontaneous toxoplasmosis in canaries (Serinus
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parasitologic analysis of extrapulmonary disease. Journal of Parasitology 75:270–287. Speer, C. A., D. G. Baker, A. Yamaguchi, and J. P. Dubey. 1997. Ultrastructural characteristics of a Lankesterella-like coccidian causing pneumonia in a northern cardinal (Cardinalis cardinalis). Acta Protozoologica 36:39–47. Splendore, A. 1909. Sopra un nuovo protozoo parassita de’ conigli. Revista da Sociedade Scientifica de Sao Paulo 4:75–79. Springer, L. 1942. Toxoplasmose epizootica entre pombos. Arquivos de Biologia 26:74–76. Tackaert-Henry, M. C., and P. Kageruka. 1977. Une e´ pizootie de toxoplasmose parmi les pigeons couronn´es, Goura cristata Pallas et Goura victoria Frazer, du Zoo d’Anvers. Acta Zoologica et Pathologica Antverpiensia 69:163–168. Teglas, M. B., S. E. Little, K. S. Latimer, and J. P. Dubey. 1998. Sarcocystis-associated encephalitis and myocarditis in a wild turkey (Meleagris gallopavo). Journal of Parasitology 84:661–663. Tenter, A. M., A. R. Heckeroth, and L. M. Weiss. 2000. Toxoplasma gondii: From animals to humans. International Journal for Parasitology 30:1217–1258. Tsai, Y. J., W. C. Chung, H. H. Lei, and Y. L. Wu. 2006. Prevalence of antibodies to Toxoplasma gondii in pigeons (Columba livia) in Taiwan. Journal of Parasitology 92:871. Vickers, M. C., W. J. Hartley, R. W. Mason, J. P. Dubey, and L. Schollam. 1992. Blindness associated with toxoplasmosis in canaries. Journal of the American Veterinary Medical Association 200:1723–1725. Vogelsang, E. G., and P. Gallo. 1954. Toxoplasmosis en aves de Venesuela. Revista De Medicina Veterinaria y Parasitologia 13:59–61. Wallace, G. D. 1973. Intermediate and transport hosts in the natural history of Toxoplasma gondii. American Journal of Tropical Medicine and Hygiene 22:456–464. Wiktor, T. J. 1950. Toxoplasmose animale. Sur une e´ pid´emie des lapins et des pigeons a` Stanleyville (Congo Belge). Annales de la Societe Belge de Medecine Tropicale 30:97–107. Williams, S. M., R. M. Fulton, J. A. Render, L. Mansfield, and M. Bouldin. 2001. Ocular and encephalic toxoplasmosis in canaries. Avian Diseases 45:262–267. Work, T. M., J. G. Massey, B. A. Rideout, C. H. Gardiner, D. B. Ledig, O. C. H. Kwok, and J. P. Dubey. 2000. Fatal toxoplasmosis in free-ranging endangered ’alala from Hawaii. Journal of Wildlife Diseases 36:205–212. Work, T. M., J. F. Massey, D. Lindsay, and J. P. Dubey. 2002. Toxoplasmosis in three species of native and introduced Hawaiian birds. Journal of Parasitology 88: 1040–1042.
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12 Trematodes Jane E. Huffman INTRODUCTION Trematodes are flat, leaflike parasitic helminthes that are classified in the phylum Platyhelminthes—a diverse group of free-living and parasitic worms that also includes the cestodes (Chapter 14). Current taxonomy places trematodes into two classes—the monogenetic trematodes that are primarily ectoparasites of fish and possess a posterior attachment organ or haptor armed with hooks and the digenetic trematodes that are common endoparasites of a variety of vertebrate hosts and have muscular oral and/or ventral suckers. The digenetic trematodes have life cycles with both sexual and asexual phases of reproduction—the former in their vertebrate definitive hosts and the latter in molluscan intermediate hosts. With the exception of the schistosomes (Chapter 13), trematodes are hermaphroditic and generally have one or two large, sometimes branched, testes, a comparatively small ovary, an often long and looping uterus, and a single common genital pore. They typically have a bifurcated intestine and blind ceca that exit the body through an anus or via an excretory vesicle. Unlike nematodes, and similar to both cestodes and acanthocephalans, the tegument lacks a cuticle (Marquardt et al. 2000). A wide range of digenetic trematodes occur in wild birds. However, the majority of these species are not associated with significant disease. Trematodes that cause large-scale epizootics in birds include Sphaeridiotrema globulus (Psilostomatidae), Cyathocotyle bushiensis (Cyclocoelidae), and Leyogonimus polyoon (Lecithodenriidae). Epizootics with these three parasites usually occur in the spring and fall, often when large numbers of birds stop over at particular areas during migration. The success of these parasites depends on the presence of first, second, and definitive hosts in the environment. Large late-summer mortalities of waterfowl attributable to these trematodes have been reported since the 1960s, when thousands of ducks were apparently killed by mixed infections of the digeneans C. bushiensis and S. globulus along the St. Lawrence River south of Quebec, Canada.
SYNONYMS Flukes, flatworms. HISTORY Much of the early history of the digenetic trematodes of birds was summarized by Dawes (1946). It seems likely that the larger flukes that are parasitic in mammals, for example, Fasciola, have been recognized since at least the fourteenth century. Their occurrence in birds was noted by Goeze (1782, cited in Dawes 1946) who published a work on the natural history of parasitic worms. Zeder in 1800 developed a systematic classification of parasitic worms and described Cyclocoelum mutabile, a trematode of the Black Scoter (Melanitta nigra) and Common Moorhen (Gallinula chloropus). Rudolphi broadened the foundations of our knowledge of parasitic worms with two works in 1810 and 1819, and coined the term “trematode” to replace the term “sucking worm.” Rudolphi described a number of trematode species from British birds. Jagerskiold published a series of papers between 1896 and 1908 on the trematode parasites of birds (Dawes 1946). In the US, the first report of an epizootic in waterfowl was reported by Price (1934) in Lesser Scaup (Aythya affinis) in the Potomac River in Washington, DC. The causative agent was S. globulus. ETIOLOGY McDonald (1981) listed 536 species of digenetic trematodes from 125 genera and 27 families of birds. A detailed treatment of this diverse group of parasites is well beyond the scope of this chapter, although they all have a number of similarities in life cycles, morphology, and development within the gut and other tissues of their definitive hosts. The most significant trematodes of birds occur in 6 of the 10 orders of digenetic trematodes listed by Brooks and McLennan (1993). These trematodes are distinguished by morphological features of both adult and immature forms. Details about their morphology and taxonomy can be found
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in Dawes (1946), Yamaguti (1971), and McDonald (1981). Ventral suckers are generally about the same size as the oral suckers, and located centrally on the anterior, ventral surface. Often, the ventral suckers are quite close behind the oral sucker. The location, number, and morphology of the suckers are used to separate members of this group. A distome is a digenetic trematode with an anterior oral sucker and the posterior sucker located on the ventral surface and a monostome possesses a single sucker, oral or ventral, rather than both. A holostome is a type of adult digenetic trematode having a portion of the ventral surface modified as a complex adhesive organ.
(Fulica atra) and Common Moorhens; in the US, the only known host currently is the American Coot. S. globulus and C. bushiensis share similar waterfowl hosts in both Europe and the US. Coinfections with S. globulus and C. bushiensis have been reported (Hoeve and Scott 1988). All three parasites can produce mortality in the avian host within 3–8 days after infection. Annual migrations can disseminate avian trematodes and of special importance is the cross migration and movement of birds following the breeding season and preceding the beginning of fall migration. This can allow for widespread dissemination of these parasites over the breeding grounds as long as the intermediate hosts are present.
HOST RANGE AND DISTRIBUTION The geographical distribution of trematodes is influenced by environmental conditions that affect the distribution of their intermediate hosts. These conditions include biotic variables such as vegetation cover and abiotic variables of the lentic environment such as size, average depth, salinity, and characteristics of the sediments. For example, a trematode that uses a particular species of mollusk as an intermediate host may only occur where that mollusk is found. The hosts of Uvulifer ambloplitis, a parasite of kingfishers (Megaceryle spp.), include snails, fish, and birds. The distribution of this trematode is a combination of the ranges of all three hosts. Nonetheless, the distribution of a particular trematode can span large areas, particularly if definitive and intermediate host species live in a broad range of habitats or if the definitive host migrates over vast regions. Some species of trematodes occur with great frequency in a large variety of avian hosts, others seem to be rarer, even specific for one or more hosts. Monostomes such as Notocotylus attenuatus and Catatropis verrucosa occur in a large variety of hosts. Holostomes such as Strigea spp. are common parasites of birds of prey, ducks, and gulls. Prosthogonimus ovatus likewise occurs in numerous birds, more than half of which are passerines of wading birds. The species P. cuneatus shows an even greater preference for passerines. At the other extreme, the two most common echinostomes—Echinostoma revolutum and Hypoderaeum conoideum—are almost entirely confined to ducks and their close relatives. Table 12.1 summarizes the distribution and host range of L. polyoon, C. bushiensis, and S. globulus, the cause of current epizootics in the US. The exotic Faucet Snail (Bithynia tentaculata), originally native to Europe, can serve as the intermediate host, and the American Coot (Fulica americana) can serve as the definitive host for all three parasites. In Europe, the only definitive hosts for L. polyoon are Eurasian Coots
EPIZOOTIOLOGY Digenetic trematodes produce eggs in their definitive hosts which pass with feces either into water or onto land. The egg of most forms is oval and has a lidlike hatch on one end called an operculum. When eggs reach freshwater, the operculum opens and a ciliated free-swimming larva called a miracidium swims out. The miracidium will then use chemotactic cues to find a suitable intermediate host, which is usually a snail. The miracidium penetrates the snail, loses its cilia, and develops into a sporocyst. Sporocycsts reproduce asexually to form either more sporocysts or a number of rediae. Rediae reproduce asexually to form more rediae or tailed forms called cercariae. The cercariae emerge from the snail and penetrate a second intermediate host (either a mollusk, amphibian, or fish), the final host, or encyst on vegetation where they transform into metacercariae. Adult worms develop from metacercariae when they are ingested by a definitive host (Figure 12.1) (Marquardt et al. 2000). Trematodes of birds generally develop in specific locations in the body of the host. The most common site is the intestine. Both holostomes and echinostomes show a preference for the lower end of the intestine. Other sites include the bursa of Fabricii (Prosthogonimus) and the cloaca (Leucochloridium). Most monostomes inhabit the air sacs and Collyriculum faba forms cysts under the skin. Species of Opisthorchiidae and Dicrocoeliidae infect the liver (Table 12.2). A number of trematodes develop in very specific and unusual sites. Both Lyperosomum longicauda, a common fluke of Carrion Crows (Corvus corone) in Europe, and Athesmia heterolecithodes from the Ruffed Grouse (Bonasa umbellus) develop specifically in the liver. Clinostomum complanatum is found in the buccal cavity and the upper ends of the esophagus and trachea. Renicola pinguis (Troglotrematidae) and Eucotyle nephritica (Eucotylidae) inhabit the kidney. Some birds are exceptional in harboring two or more species of trematodes that
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227
Trematodes
Table 12.1. Summary of host range and distribution information for Sphaeridiotrema globulus (family Psilostomatidae), Cyathocotyle bushiensis (family Cyathocotylidae), and Leyogonimus polyoon (family Lecithodendriidae). Trematode*
Intermediate hosts
Sphaeridiotrema Bithynia globulus tentaculata† Elimia virginica Fluminicola virens Oxytrema silicula
Cyathocotyle bushiensis
Leyogonimus polyoon
Avian hosts (USA)
Location
American Coot (Fulica Potomac River americana) Mute Swan (Cygnus olor) NJ, NY, WI, OR, USA Tundra Swan (Cygnus columbianus) Lesser Scaup (Aythya affinis) Canvasback (Aythya valisineria) Common Goldeneye (Bucephala clangula) Ruddy Duck (Oxyura jamaicensis) Long-tailed Duck (Clangula hyemalis)
Avian hosts (Europe) Tufted Duck (Aythya fuligula) Greater Scaup (Aythya marila) Northern Pintail (Anas acuta) Long-tailed Duck (Clangula hyemalis) Razorbill (Alca torda) Common Merganser (Mergus merganser)
Whooper Swan (Cygnus cygnus) Red-breasted Merganser (Mergus serrator) St. Lawrence Long-tailed Duck River, Canada (Clangula hyemalis) Great Lakes European Shag Basin, WI, USA (Phalacrocorax aristotelis)
Bithynia tentaculata American Coot (Fulica americana) American Black Duck (Anas rubripes) Blue-winged Teal (Anas discors) Green-winged Teal (Anas carolinensis) WI Bithynia tentaculata American Coot (Fulica americana)
Eurasian Coot (Fulica atra) Common Moorhen (Gallinula chloropus chloropus)
Note: All three trematodes were introduced from Europe. Epizootics caused by Sphaeridiotrema were documented in 1928, while those caused by Cyathocotyle bushiensis occurred in the 1960s. Leyogonimus polyoon was identified as a cause of eipizootics in Wisconsin in 1996. These trematodes occur in the lower intestine (Sphaeridiotrema, Cyathocotyle) or upper and middle intestines (Leyogonimus) of their hosts. * References for avian hosts can be found in Dawes (1946), Gower (1939), and Yamaguti (1971). † Bithynia tentaculata was introduced from Europe into Lake Michigan in the 1970s. belong to the same genus. The Black Scoter has been reported to be parasitized by 7 species of Gymnophallus (Microphallidae) (Dawes 1946). Skrjabin (1926) described an example of a bird parasitized by 17 species of helminths. The larval stages of trematodes are influenced by density-independent factors such as temperature and moisture. Their populations may fluctuate dramatically over time as a result of environmental changes, possibly leading to local extinctions (Bush et al. 2001).
Trematodes that use hosts that are also strongly influenced by density-independent factors and parasites living at the edges of their geographic ranges may also be strongly influenced by density-independent factors (Bush et al. 2001). In contrast, density-dependent effects primarily occur within vertebrate and intermediate hosts and can reduce trematode survival or fecundity and ultimately regulate parasite abundance. Two of the factors most likely to be important in exerting density-dependent effects are host immune responses
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8:57
Parasitic Diseases of Wild Birds that gradual debilitation of a wild bird from a specific disease may predispose it to several other potential pathogens within its environment. A statistically significant association between poor body condition and high numbers of intestinal trematodes has been reported in Common Loons (Gavia immer) from maritime Canada (Daoust et al. 1998). Factors involved in susceptibility of a host population include density of infective stages of the parasite in the environment, rate of exposure between host and infective stage, and host susceptibility. Density and nutritional status of avian populations and the interaction with avian trematodes may have consequences that result in epizootics.
Figure 12.1. Typical life cycle of a digenetic trematode. Eggs are released by adult worms and pass with feces of their avian hosts either into water or onto land. When eggs reach freshwater, a ciliated free-swimming miracidium is released which penetrates a snail, loses its cilia, and develops into a sporocyst. Sporocycsts reproduce asexually to form either more sporocysts or a number of rediae. Rediae reproduce asexually to form more rediae or tailed forms called cercariae. The cercariae emerge from the snail and penetrate a second intermediate host (either a mollusk, amphibian, or fish), the final host, or encyst on vegetation where they transform into metacercariae. Adult worms develop from metacercariae when they are ingested by a suitable definitive host. and competitive interactions either within or between trematode species. Both these factors may lead to host mortality as trematode density increases, with ultimate effects on overall abundance of the parasites (Bush et al. 2001). Pathogenicity of a trematode may differ among species of birds as well as different populations of the same species. The effects of trematodes on individual birds are well documented in the literature. The identification of multiple concurrent disease problems in individual birds in poor body condition suggests
CLINICAL SIGNS Clinical signs of trematode infections vary and depend on the number of parasites, species of trematode involved, and the organs and organ systems affected. Signs seen in gastrointestinal infections include watery blood-stained diarrhea and pericloacal feathers stained with blood (Roscoe and Huffman 1982), weakness (i.e., wing droop) (Huffman and Roscoe 1989), leg weakness (van Haitsma 1931), inability to fly (Kocan and Kocan 1972), unsteady gait, disorientation, and a weak raspy call (Huffman and Roscoe 1989). Emaciation (Poonswad et al. 1992) and diarrhea (Dedrick 1965; Patnaik et al. 1970; Graczyk and Schiff 1993) also occur in wild birds. A high intensity of infection with Paratanaisia bragai in the kidney can cause apathy, loss of weight, diarrhea, and death in pigeons (Portugal et al. 1972; Arnizaut et al. 1992). Cloacal discharges have been reported in waterfowl infected with gastrointestinal trematodes (Annereaux 1940; Biester and Schwarte 1959). Increases in cloacal temperature have also been noted (Gagnon et al. 1993). Anemia can be pronounced in infected birds (Kocan and Kocan 1972; Huffman and Roscoe 1989; Luppi et al. 2007). Increases in hemoglobin and packed cell volume have been reported (Gagnon et al. 1993). Mallards (Anas platyrhynchos) experimentally infected with S. globulus develop increased prothrombin time. Excysted metacercariae were shown to produce beta hemolysis on blood agar (Tabery et al. 1988). Approximately 25 proteins have been isolated from the excretory/secretory products of S. globulus (Babu 2000). These have an effect on the coagulation factors Xa and IIa (Isopi 2000)—causing a 54% inhibition of factor Xa and a 17% inhibition of IIa. How the excretory/secretory products inhibit the factors has not been determined. PATHOLOGY Trematodes can cause lesions in their hosts by a number of different mechanisms. The most pathogenic species of trematodes are described in this section.
Brachylaemidae Leucochloridiomorpha Cathaemasiidae Cathaemasia Clinostomidae Clinostomum Cortrematidae Cortrema Cyathocotylidae Cyathocotyle Cyclocoelidae Bothrigaster Cyclocoelum Typhlocoelum Hyptiasmus Ophthalmophagus Wardianum Dicrocoeliidae Athesmia Lyperosomum Oswaldoia Platynosomum
Trematode
+ +
Ac B
+
+
Bd BF Bv C
c
Ca
E
+
+
Es
+
I
+ +
IO
K
229 +
+ +
L
+
M
+ + +
N
N
Od PV Sc
+
+
Tb
(continues)
T
+
Tc
October 16, 2008
+
+ +
As
Table 12.2. Some trematodes and their anatomical location in avian hosts. As, air sacs; Ac, abdominal cavity; B, gall bladder; Bd, bile ducts; BF, bursa Fabricii; Bv, blood vessels; C, cloaca; c, conjunctiva; Ca, ceca, E, eye; Es, esophagus; I, intestine; IO, infra-orbital sinus; K, kidney, L, liver; M, buccal cavity; N, nasal cavity; n, nictitating membrane; Od, oviduct; PV, proventriculus; s, subcutaneous cysts; T, trachea; Tb, trachea and bronchi; Tc, thoracic cavity (Dawes 1946; Yamaguti 1971).
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Diplostomatidae Diplostomum Neodiplostomum Posthodiplostomum Uvulifer Echinostomatidae Chaunocephalus Echinostoma Echinoparyphium Echinochasmus Himasthla Hypoderaeum Parorchis Stephanoprora Eucotylidae Eucotyle Paratanaisia Eumegacetidae Eumegacetes Heterophyidae Ascocotyle Cryptocotyle Phagicola Lecithodendriidae Leyogonimus Macyella Leucochloridiidae Leucochloridium Urotocus
Trematode
Table 12.2. (Continued) As
Ac B
230 +
+
+ +
c
Ca
E
Es
+ +
+ + +
+
+ + + +
+ + + +
I
IO
+ +
K
L
M
N
N
Od PV Sc
T
Tb
Tc
October 16, 2008
+ +
+
+
Bd BF Bv C
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Microphallidae Gymnophallus Maritrema Spelotrema Levinseniella Microscaphidiidae Polyangium Notocotylidae Notocotylus Catatropis Paramonostomum Opisthorchiidae Amphimerus Metorchis Opisthorchis Pachytrema Pseudamphimerus Orchipedidae Orchipedum Paramphistomidae Zygocotyle Philophthalmidae Philophthalmus Cloacitrema Parorchis Plagiorchiidae Plagiorchis Pronocephalidae Parapronocephalum Prosthogonimidae Prostogonimus Schistogonimus + + +
+ +
231 + +
+
+
+
+
+ +
+
+
+
+ +
+
+
+
+ + +
+ + + + +
+
+ +
+ (continues)
+
October 16, 2008
+
+
BLBS014-Atkinson 8:57
Psilostomatidae Psilostomum Psilochasmus Ribeiroia Sphaeridiotrema Schistosomatidae Austrobilharzia Gigantobilharzia Bilharziella Stomylotrematidae Laterotrema Stomylotrema Strigeidae Cotylurus Apatemon Parastrigea Strigea Thapariellidae Thapariella Troglotrematidae Collyriclum Renicolla
Trematode
Table 12.2. (Continued) As
Ac B
+
c
Ca
E
Es
232 +
+ + + +
+
+ + + +
I
IO
K
L
M
N
N
+
+ +
Od PV Sc
T
Tb
Tc
October 16, 2008
+
+
+ + +
Bd BF Bv C
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Trematodes Pathology may be due to a direct host–parasite interaction, mechanical insult resulting in tissue damage, or by the ingestion of host tissue. Host immune responses cause inflammation and immune-mediated pathology. The lesions are closely related to the anatomical location of the parasite. Liver, Bile Ducts, and Gall Bladder Gross lesions caused by trematodes that develop within the bile ducts and gall bladder of their avian hosts include irritation and inflammation of the bile duct epithelium, cholangiectasis, enlargement of the bile duct lumen, blockage of the duct, and back flow of bile into the liver, leading to dystrophic and atrophic changes in the liver. Lesions have been reported from infections of Metorchis bilis in White-bellied Sea-Eagles (Haliaeetus leucogaster) (Krone et al. 2006), and infections of Opisthorchis sp. in Western Marsh Harriers (Circus aeruginosus) and Northern Harriers (Circus cyaneus) (Averikhin et al. 1984). Intensity of infection likely plays a role in severity of lesions. For example, over 200 individuals of an Opisthorchis sp. were recovered from the bile ducts of infected Western Marsh-Harriers, causing inflammation of the duct walls, enlargement of the lumen, and stasis of the contents, leading to dystrophic and atrophic changes in the liver. Fatal hepatic trematodiasis caused by Amphimerus elongatus has been diagnosed in Double-crested Cormorants (Phalacrocorax auritus), Common Loons, and Bald Eagles (Haliaeetus leucocephalus). Most birds had enlarged livers with irregular capsular surfaces and numerous white, dark green, or black foci and tracts. Microscopic lesions consisted of granulomas composed of multinucleated giant cells with fibrous connective tissue and other inflammatory cells surrounding necrotic debris, trematode eggs, trematode pigment, and, occasionally, bacterial colonies. Gravid trematodes were associated with compression of adjacent hepatocytes in portal areas. Amphimerus heterolecithodes infects the bile ducts of the liver of Wild Turkeys (Meleagris gallopavo). The ducts can be occluded and there can be hyperplasia or complete desquamation of the epithelium of the duct walls. In areas that contain numerous parasites, there is extensive fibrosis. The parasite has been reported free in the liver parenchyma. Trematode eggs have been found in the kidney and pancreas of Double-crested Cormorants, suggesting that the parasite migrates via the bile duct, pancreatic duct, and ureter to reach these organs (Kuiken et al. 1999). Kidneys Gross lesions caused by trematodes that develop within the kidney of birds include distention of the collecting tubules and a thickening of their walls and exten-
233
sive cellular infiltration of the parenchyma (dos Santos 1934). At necropsy, Blue- and Yellow-Macaws (Ara ararauna), Blue-winged Macaws (Primolius maracana), White-eared Parakeets (Pyrrhura leucotis), and Ringnecked Pheasants (Phasianus colchicus) infected with P. bragai had enlarged kidneys with brown-yellow discoloration and irregular cortical surfaces. Microscopic lesions consisted of granulomatous nephritis and included an interstitial, multifocal to coalescent, lymphoplasmacytic infiltrate with some epithelioid macrophages and a few heterophils. Adult worms and eggs were observed within dilated tubules and in the renal pelvis. In one bird, some parasite eggs were located interstitially and associated with an intense adjacent granulomatous reaction (Luppi et al. 2007).
Air Sacs Bothrigaster variolaris (Cyclocoelidae) infects the air sacs of Snail Kites (Rostrhamus sociabilis) (Cole et al. 1995). Grossly, the air sacs were opaque and tan granular deposits had accumulated in the folds and angles of the tissues. Primary microscopic lesions included moderate pyogranulomatous bronchitis and peribronchitis, with mild squamous metaplasia of the epithelium near intrabronchial trematodes. Mild granulomatous airsaculitis composed exclusively of large, pigment-laden macrophages was also noted.
Gastrointestinal Tract Lesions associated with gastrointestinal trematodes can be mild to severe depending on the number of parasites and species. The character of the lesions also depends on the species of trematode. Lesions are generally confined to the gastrointestinal tract and can range from mild enteritis to severe ulcerative hemorrhagic enteritis. Sphaeridiotrema globulus, C. bushiensis, and L . polyoon are three gastrointestinal parasites that cause epizootics in waterfowl in the US. Infections with S. globulus cause ballooning of the jejunum and ileum and the affected intestine may have a generalized cyanotic appearance. Foci of hemorrhage circ*mscribe trematodes and are visible through the serosa. Ulcers in the jejunum and ileum may penetrate the mucosa to the circular muscle layer. The intensity of fatal infections appears to be host species dependent. American Coots and Mute Swans (Cygnus olor) can die from an infection of S. globulus with as few as 20 parasites, whereas the fatal worm burden for Muscovy Ducks (Cairina moschata), Mallards, and Canada geese (Branta canadensis) ranges from 100 to 3,300 (Trainer and Fischer 1963; Campbell and Jackson 1977; Roscoe and Huffman 1982).
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Parasitic Diseases of Wild Birds
Cyathocotyle bushiensis infects the lower intestine and most commonly the cecae of ducks (Table 12.3). Both ceca can be affected and may externally appear to be dark, elongated, and regularly distended. Internally, the ceca have numerous hemorrhagic areas and whitish caseous plaques. Ulceration, generalized mucosal necrosis, and firm, irregular cores may be present (Gibson et al. 1972). Intensity of infection in Bluewinged Teals (Anas discors) ranged from 1 to 649 worms per individual, with an average of 260 worms. Intensity of infection in American Black Ducks (Anas rubripes) was 180 worms per individual (Gibson et al. 1972). Hoeve and Scott (1988) reported that as few as seven parasites could cause mortality in experimentally infected ducks. Lesions from low-intensity infections may heal rapidly once the worms complete their life span. Mortality is probably attributable to the effects of severe infections, including hemorrhage and fluid loss. Leyogonimus polyoon infects primarily the upper and middle areas of the small intestine. Gross lesions include severe enteritis characterized by thickening of the intestinal wall and a fibrinous to caseous core of necrotic debris that blocks the lumen of the intestine (Cole and Friend 1999). Echinostoma spp. can cause mild to severe enteritis in birds (Griffiths et al. 1976; Hossain et al. 1980). However, a slight abrasion of the mucosal surface at the site of attachment was the only damage noted in Mallards infected with E. trivolvis (Mucha et al. 1990). Enlargement of the proventriculus and reddening around the orifices of the glands has been observed with infection with Ribeiroia. In heavy infections, grayish exudates on the surface and superficial ulceration have been reported. Histologically, the mucosal surface is covered with a fibrinous exudate and the outer portion is necrotic with a polymorphonuclear leukocytic infiltration (Kocan and Locke 1974). Eyes Noticeable irritation of the eyes and a retracted nictitating membrane can be observed in chickens experimentally infected with Philophthalmus gralli (West 1961). Waterfowl infected with P. gralli have swollen and hyperemic nictitating membranes (Schmidt and Toft 1981). Erosion and ulceration of the conjunctival membrane and an intense inflammatory response were evident in histological sections of infected areas of the eye. Diffuse conjunctivitis was present adjacent to the attachment site of P. gralli in a Swan Goose (Anser cygnoides) (Schmidt and Hubbard 1987). Oviduct Infection of the oviduct in White-throated Sparrows (Zonotrichia albicollis) and Wild Turkeys with P. mar-
crochis results in distention and the accumulation of considerable amounts of exudates and egg material. The oviduct may have varying degrees of inflammation, depending on the number of parasites. A catarrhal to a fibrinous exudate or a caseous mass may be present in the oviduct lumen, where broken yolks and frequently large concentrations of yolk and albumen will also be found. If the oviduct ruptures, albumen and yolk material will be present in the body cavity and peritonitis with possible organ adhesions will result (Biester and Schwarte 1959). DIAGNOSIS Anatomical location of adult trematodes is an important clue for their identification (Table 12.2). Birds should be closely examined for the presence of conjunctival discharges that may indicate infection with eye flukes such as Philophthalmus sp. or the presence of cloacal prolapse or soiling around the vent that may indicate infection with intracloacal or intestinal flukes. The diagnosis of a trematode infection may be based on the microscopic identification of eggs in the stool. Trematode eggs are relatively small, typically have an operculum, and contain either an embryo or, in mature eggs, a ciliated miracidium. By contrast, nematode eggs usually have thin shells and contain either a morula in unembryonated eggs or a recognizable larval worm. Nematode eggs do not have an operculum, but some species may have unusual but well-defined structural modifications. Cestode eggs have thickened walls and contain a larva called an onchosphere, which possesses six hooklets. Acanthocephalan eggs contain a partially developed embryo or acanthor. The eggs of schistosomes (blood-dwelling trematodes) do not have an operculum, but do possess terminal or lateral spines (Chapter 13). If fecal samples are examined within less than 72 h, no preservatives are needed. However, some eggs may embryonate or hatch during this time unless air is excluded from the container. To maintain fecal samples longer than 72 h, the fecal sample should be fixed in 10–15 volumes of 10% formalin. Determination of the genus and species of the trematode can be done after fixing and staining adult worms by using traditional morphological methods (Pritchard and Kruse 1982). Trematodes that are recovered at necropsy or passed in the feces require relaxation before fixation. Chilling the worms, either in saline or in tap water overnight in the refrigerator, relaxes them with the least handling. Another method is to place them into 5–10% ethyl alcohol at room temperature. The relaxed worms can be fixed in 10% formalin or preferably alcohol–formalin–acetic acid fixative (Pritchard and Kruse 1982).
Host species
Parasite family
Parasite
235
American Black Duck (Anas rubripes) Blue-winged Teal (Anas discors) Green-winged Teal (Anas carolinensis) Northern Pintail (Anas acuta) Northern Shoveler (Anas clypeata) Canvasback (Aythya valisineria) Mallard (Anas platyrhynchos) Green-winged Teal (Anas carolinensis) Blue-winged Teal (Anas discors) Greater Scaup (Aythya marila) Lesser Scaup (Aythya affinis) Mottled Duck (Anas fulvigula)
Cattle Egret (Bubulcus ibis)
Asian Openbill (Anastomus oscitans)
Great Blue Heron (Ardea herodias) White Stork (Ciconia ciconia) Black Stork (Ciconia nigra) Clinostomum attentuatum Cathaemasia hians
Cyathocotyle bushiensis
Typhlocoelum cucumerium
Cyathocotylidae
Cyclocoelidae
Echinostomatidae Pegosomum sp.
Echinostomatidae Chaunocephalus ferox
Cathaemasiidae
Clinostominae
Cosmopolitan
North America
Japan
Thailand
Europe
North America
Tracheal obstruction
Mutifocal hepatitis Kuiken et al. (1999) Bile duct hyperplasia Esophageal Forrester and Spalding obstruction (2003) Esophageal Stoskopf et al. (1982), obstruction and Merino et al. (2001) Catarrhal enteritis Patnaik et al. (1970), Poonswad et al. (1992), and Hofle et al. (2003) Cholangitis and Murata et al. (1998) cholecystitis Typhlitis Gibson et al. (1972)
Canada
(continues)
Gower (1937), Town (1960), Cornwell and Cowan (1963), Taft (1971), Kinsella and Forrester (1972), Broderson et al. (1977), Mahoney and Threlfall (1978), and Scott et al. (1980)
Greve et al. (1986)
Villus atrophy
Florida, USA
Harrigan (1992)
References
Liver necrosis
Lesion
Australia
Geographic local
October 16, 2008
Anseriformes
Ciconiiformes
Sphenisciformes Little Penguin (Eudyptula minor) Prosthogonimidae Mawsonotrema eudyptulae Pelecaniformes Brown Pelican (Pelecanus Heterophyidae Phagicola longa occidentalis) Cyathocotylidae Mesostephanus appendiculatoides Double-crested Cormorant Opisthorchiidae Amphimerus (Phalacrocorax auritus) elongatus
Host order
Table 12.3. The major parasite families and species of trematodes that can cause disease in wild birds.
BLBS014-Atkinson 8:57
Echinostomatidae
Parasite family
Parasite
236 Cyclocoelidae
Psilostomatidae
Renicolidae
Philophthalmidae
Prairie Falcon (Falco mexicanus) Strigeidae
Florida Snail Kite (Rostrhamus sociabilis plumbeus)
Osprey (Pandion haliaetus)
Wood Duck (Aix sponsa)
Ruddy Duck (Oxyura jamaicensis) Blue-winged Teal (Anas discors) Mallards (Anas platyrhynchos) Dabbling Ducks (Anas spp.)
Bothrigaster variolaris
Ribeiroia ondatrae
Sphaeridiotrema globulus Philophthalmus sp. Renicola lari
Sphaeridiotrema globulus
Echinoparyphium recurvatum Echinosotma sp. Echinosotma trivolvis Notocotylidae Notocotylus attenuatus Anas spp. Paramphistomatidae Zygocotyle lunata American Black Duck (Anas Microphallidae Maritrema rubripes) acadiae Blue-winged Teal (Anas discors) Maritrema sp. Mute Swan (Cygnus olor) Psilostomatidae Sphaeridiotrema Tundra Swan (Cygnus globulus columbianus) Whooper Swan (Cygnus cygnus) Lesser Scaup (Aythya affinis) Sphaeridiotrema globulus Greater Scaup (Aythya marila) Sphaeridiotrema globulus
Mallard (Anas platyrhynchos) Northern Pintail (Anas acuta) Mallard (Anas platyrhynchos) Canada Goose (Branta canadensis) Anas spp.
Host species
Enteritis Typhlitis Intestinal ulceration Intestinal enteritis UHE*
UHE
USA North America Nova Scotia
Washington, DC, USA North Central Minnesota, USA Wisconsin, USA
North America
Florida, USA
North America
North America
North America
Canada
Kennedy and Frelier 1984 Kocan and Locke (1974); Kinsella et al. (1996) Cole et al. (1995)
Schmidt and Toft (1981)
Hoeve and Scott (1988)
Minnesota Department of Natural Resources (2007) United States Geological Survey (1997)
Price (1934)
Hoeve and Scott (1988) Roscoe and Huffman (1982, 1983)
Mettrick (1959) Swales (1933)
Griffiths et al. (1976)
Huffman (2000)
Soulsby (1965)
References
Air sacculitis Pyogranulomatous bronchitis Diarrhea Dedrick (1965)
Hyperplasia of the proventriculus
Nephritis
Conjunctivitis
UHE
UHE
UHE
Enteritis
Cosmopolitan
Canada North America
Enteritis
Lesion
Cosmopolitan
Geographic local
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Falconiformes
Host order
Table 12.3. (Continued)
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Galliformes
Dicrocoeliidae
Psilostomatidae
Strigeidae Parastrigea tulipoides Ribeiroia ondatrae Athesmia jolliei
Opisthorchis sp.
Cryptocotyle lingua Metorchis bilis
Heterophyidae Opisthorchiidae
Strigea falconis
Strigeidae
Cholangiectasis
North America Granuloma
North America Dystrophic and atrophic liver damage North America None described
Finland
North America Hyperplasia of the Proventriculus North America Emaciation
Miller and Harkema (1965) Beaver (1939)
Averikhin et al. (1984)
Krone et al. (2006)
Smith (1978)
Kinsella et al. (1998)
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North America Fibrosis of the bile Schell (1957) ducts Plagiorchiidae Plagiorchis North America Enteritis Clausen and elegans Gudmundsson (1981) Wild Turkey (Meleagris Brachylaemidae Postharmostomum North America Typhlitis Soulsby (1965) gallopavo) gallinum Opisthorchiidae Amphimerus het- North America Obstruction and Kingston (1984), and erolecithodes fibrosis of bile Davidson and ducts Wentworth (1992) Prosthogonimidae Prosthogonimus North America Oviduct Davidson and macrorchis inflammation Wentworth (1992) White-throated Sparrow Prosthogonimidae Prosthogonimus Canada None described Brooks et al. (1993) (Zonotrichia albicollis) macrorchis Paratanaisia South America Nephritis Travassos et al. (1969), Spot-winged Wood Quail Eucotylidae bragai Costa et al. (1975), (Odontophorus capueira) Silva et al. (1990), Wild Turkey (Meleagris Menezes et al. (2001), gallopavo) and Pinto et al. (2004) Ring-necked Pheasant (Phasianus Paratanaisia Brazil Nephritis Gomes et al. (2005) colchicus) bragai (continues)
White-tailed Eagle (Haliaeetus albicilla) Western Marsh-Harrier (Circus aeruginosus) Northern Harrier (Circus cyaneus) Red-shouldered Hawk (Buteo lineatus) Cooper’s Hawk (Accipiter cooperii) American Kestrel (Falco sparverius) Gyrfalcon (Falco rusticolus)
Bald Eagle (Haliaeetus leucocephalus)
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American Coot (Fulica americana)
Gruiformes
Cyclocoelidae
238
∗
Dicrocoeliidae
American Robin (Turdus migratorius)
Brachylecithum mosquensis
Amphimerus elongates Cathaemasiidae Pulchrosoma pulchrosoma Troglotrematidae Collyriclum faba
Opisthorchiidae
Belted Kingfisher (Megaceryle alcyon) Ringed Kingfisher (Megaceryle torquatus) Wood Thrush (Hylocichla mustelina)
UHE, Ulcerative hemorrhagic enteritis.
Passeriformes
Coraciiformes
Philophthalmus gralli Paratanaisia bragai
Philophthalmus hegeneri
Cyclocoelum mutabile
Renal medullary collecting ducts and ureters Bile duct hyperplasia Lung granulomas
South America
North America
North and Central America
Peru
Merino et al. (2003)
Boyd and Fry (1971)
Pinto et al. (2004)
Schmidt and Toft (1981)
Farner and Morgan Wasting and (1944), and Kirmse anemia, obstruction of the (1987) cloaca Obstruction of bile Schell (1957) ducts
Conjunctivitis
North America
References
Cole and Friend (1999), and Cole and Franson (2006) Enteritis Trainer and Fischer (1963) Hemopericardium, McLaughlin (1976, blood-filled air 1977, 1983) sacs, biliary congestion Retardation of Underhill et al. (1994), moult and Branton et al. (1985) Conjunctivitis Nollen and Kanev (1995)
Enteritis
Lesion
North America
North and South America North America
North America
North America
Sphaeridiotrema globulus Cyclocoelum mutabile
Psilostomatidae
Geographic local North America Europe
Parasite
Lecithodendriidae Leyogonimus polyoon
Parasite family
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Charadriiformes Red Knot (Calidris canutus) Cyclocoelidae Greater Yellowlegs (Tringa melanoleuca) Royal Tern (Thalasseus maximus) Philophthalmidae Laughing Gull (Larus atricilla) Yellow-crowned Night-Heron (Nyctanassa violacea) European Herring Gull (Larus argentatus) Columbiformes Ruddy Ground-Dove (Columbina Eucotylidae talpacoti)
Host species
Host order
Table 12.3. (Continued)
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Trematodes In studies designed to clarify relationships between morphologically similar species, molecular techniques are being developed to identify trematodes (Galazzo et al. 2002). Adult specimens of the opisthorchiid liver flukes Opisthorchis felineus from the Western MarshHarrier and M. bilis found in White-tailed Eagles (Haliaeetus albicilla) can be identified by using speciesspecific primers based on a part of the mitochondrial cytochrome c oxidase I gene (Pauly et al. 2003). To better understand the systematics and biogeography of Ribeiroia sp. from Great Blue Herons (Ardea herodias), the intertranscribed spacer region 2 of the ribosomal gene complex has been sequenced to determine differences between species (Wilson et al. 2005). IMMUNITY There is experimental evidence of acquired and agerelated immunity in wild birds with trematode infections. Acquired resistance to infection with S. globulus has been reported in experimentally infected Mallards (Huffman and Roscoe 1986). When exposed to a moderate dose of metacercariae of S. globulus, Mallards can develop resistance to subsequent reinfection. Host cell-mediated immunity and wound healing in Mallards experimentally infected with S. globulus has been evaluated (Mucha and Huffman 1991). An increase in mast cells and eosinophils occurred in intestinal tissue of infected ducks, but not in controls. Antibodies that were reactive with antigens of S. globulus have been demonstrated in Mallards (Jones 1993). Immunity to reinfection with Zygocotyle lunata has also been reported (Willey 1941). The age of the host at the time of infection may be a factor in the number and size of eye flukes (Philophthalmus sp.) recovered from laboratory infected chickens or geese (Nollen 1971). No protection to a challenge infection was provided by a 10-day initial infection with Philophthalmus megalurus. An initial infection with Philophthalmus hegeneri failed to protect chickens against a hom*ologous challenge 12–14 days later (Fried 1963). It appears from these studies that there is little host immunity after infection with Philophthalmus, although higher antibody titers were reported in infections with P. megalurus infection than those from P. gralli (Snyder 1991). There appears to be age-specific immunity to reinfection with Crytocotyle lingua. Ducks have been reported to be refractory to reinfection with this species and older terns and gulls harbor relatively few mature worms and pass recently excysted metacercariae in their feces. In contrast, young birds are usually heavily parasitized (Willey and Stunkard 1942). In a comparison of pairs of closely related species of birds that differ with respect to whether they are mi-
239
gratory or residents, the size of two immune defense organs (the bursa of Fabricius and the spleen) was consistently larger in the migratory species (Møller and Erritzøe 1998). Since the bursa is found only in juvenile, sexually immature birds, adaptations for immune defense appear to exist before the start of the first migration. PUBLIC HEALTH CONCERNS Nollen and Kanev (1995) documented several cases of human infections with preadult (prepatent) eye flukes. A human eye infection with Philophthalmus sp. was reported from Japan from a 67-year-old farmer (Mimori et al. 1982). However, most avian trematodes pose no threats to humans. DOMESTIC ANIMAL HEALTH CONCERNS Echinostomiasis (Echinostoma spp.) is a significant cause of mortality in commercial duck farms in Europe and Asia (Kishore and Sinha 1982). The parasite can be maintained within the domestic flock or brought in by wild birds. Psilochasmus oxyurus has been reported from domestic geese in Brazil where flocks are generally maintained under poor sanitary conditions (Fernandes et al. 2007). Three types of integrated fish-cum-duck farming practices have been developed in China that can allow exposure of domestic waterfowl to trematode infections: (1) raising large groups of ducks in open rivers, lakes, and reservoirs during the day and confining the birds in pens at night, (2) raising ducks on the edge of ponds where a large duck pen is constructed on flat areas of the shore with appropriate cemented areas for dry and wet runs, and (3) embankment and fencing of ponds to form both dry and wet runs (Bao-tong and Hua-zhu 1984). WILDLIFE POPULATION IMPACTS Birds are hosts to a wide variety of trematodes, but with few exceptions the significance of these infections on wild populations is unknown. Mixed trematode infections are common and the effect of any one parasite species depends on other parasites, diseases, or stressors that may be present. When trematodes do not directly kill the host they may, however, affect behavior, reproduction, the assimilation of nutrients, and in other ways contribute to the ill health of birds (Threlfall 1986). Severe and repeated epizootics in wild waterfowl have been caused by S. globulus, C. bushiensis, and L. polyoon. An epizootic in wild ducks was attributed to Maritrema acadiae by Swales (1933). This species has
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only recently been reported again from Mar Chiquita coastal lagoon, Buenos Aires province, Argentina, when adults of Maritrema bonaerensis n. sp. were collected from the intestine of Brown-hooded Gulls (Larus maculipennis) and Olrog’s Gulls (Larus atlanticus) (Etchegoin and Martorelli 1997). TREATMENT AND CONTROL Treatment and control measures for avian trematodes are few, especially for free-ranging waterfowl. The only practical solution is to remove birds from the source of infection. This can be done if the intermediate hosts are known. Control measures could involve reduction of snail intermediate hosts through the use of molluscicides or by draining snail habitat. Good waste management practices help prevent infection in captive situations. Oxyclozanide has been used on duck farms in Poland for treatment of the trematode N. attenuatus. Treatment was successful in eliminating the worm and preventing contamination of the pasture (Robertson and Courtney 1995). Birds infected naturally with P. gralli at the San Antonio, Texas, zoo were treated successfully with creoline (Nollen and Murray 1978), and the eyes were immediately flushed with sterile distilled water to remove the worms. Greve and Harrison (1980) reported that young Ostriches (Struthio camelus) raised in captivity were found to harbor large numbers of adult P. gralli in the orbital cavity between the nictitating membrane and outer eyelid. Persistent treatment with carbamate powder and antibiotics finally eliminated the worms. In raptors, trematode infections are usually regarded as being of little clinical significance. If diagnosed and considered to be significant, they can be treated with praziquantel (Kollias et al. 1987). MANAGEMENT IMPLICATIONS Massive late-summer mortalities of American Black Ducks, Blue-winged Teal, and Mallards have been attributed to mixed infections of C. bushiensis and S. globulus in the St. Lawrence River south of Quebec. Infections were linked to ingestion of the invasive European gastropod Bithynia tentaculata (Hoeve and Scott 1988). In 1997, tens of thousands of American Coots were killed in Wisconsin’s Shawano Lake by a third digenean trematode (L. polyoon) that was known formerly only from Europe. Once again, introduced Bithynia played a key role in transmission of the trematodes (USGS-NWHC Fact Sheet). Since 2002, all three worms have been implicated in massive waterfowl mortalities in Wisconsin ( United States Geological Survey 2007). Waterfowl mortality attributable to Cyathocotyle and Sphaeridiotrema in Minnesota’s
Lake Winnibigoshish has been linked to Viviparus georgianus (Minnesota Department of Natural Resources 2007). This snail is a native of the American southeast and is much more widely distributed throughout the US than Bithynia. If these trematodes can infect Viviparus, they would also seem likely to be able to exploit other indigenous snails and in so doing expand their potential ranges nationwide. As stopover sites for waterfowl become fewer, remaining refuges become more important in sustaining populations of migratory birds. When migrants become concentrated within refuges, the probability that epizootics may occur increases. The introduction of waterfowl into new continents may have led to the transfer and establishment of their parasites into these new locations. For example, S. globulus was first reported in 1927 in the US, most likely as a result of the importation and release of Mute Swans. During migration, wild birds may carry trematodes over long distances to new areas and the resulting introduction of their helminth parasites can put na¨ıve, native hosts at risk. Enhancing productivity of an aquatic habitat, through the impoundment of flowing water, eutrophication, and increased thermal inputs, can increase mollusk populations and other intermediate hosts of trematodes. This can increase the prevalence of trematode parasitism in birds using the area. Both migration and pollutants may stress avian hosts, suppress immune responses, and enhance vulnerability to parasitic disease. To better manage healthy populations of wild birds, continued research and surveillance are critical. Bird population monitoring programs should focus on identifying the foci, pathways, and intermediate hosts for trematodes; continue to develop methods for detecting new populations of intermediate hosts and parasites; and develop strategies and methods to control and manage populations of intermediate hosts. Hazing, or chasing waterfowl elsewhere, would not be effective at reducing losses and may aid in spread of the diseases to other lakes or wetlands.
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Trematodes Kollias, G. V., E. C. Greiner, and D. Heard. 1987. The efficacy of ivermectin and praziquantel against gastrointestinal nematodes and trematodes in raptors. In Proceedings of the First International Conference of Zoo Avian Medicine 1:6–11. Krone, O., T. Stjernberg, N. Kenntner, F. Tataruch, J. Koivusaari, and I. Nuuja. 2006. Mortality factors, helminth burden, and contaminant residues in white-tailed sea eagles (Haliaeetus albicilla) from Finland. Ambio 35:98–104. Kuiken, T., F. A. Leighton, G. Wobeser, and B. Wagner. 1999. Causes of morbidity and mortality and their effect on reproductive success in double-crested cormorants from Saskatchewan. Journal of Wildlife Diseases 35:331–346. Luppi, M. M., A. L. de Melo, R. O. C. Motta, M. C. C. Malta, C. H. Gardiner, and R. L. Santos. 2007. Granulomatous nephritis in psittacines associated with parasitism by the trematode Paratanaisia spp. Veterinary Parasitology 146:363–366. Mahoney, S. P., and W. Threlfall. 1978. Digenean, nematoda, and acanthocephala of two species of ducks from Ontario and eastern Canada. Canadian Journal of Zoology 56:436–439. Marquardt, W. C., R. S. Demaree, and R. B. Grieve. 2000. Parasitology & Vector Biology, 2nd ed., Academic Press, San Diego, CA, 720 pp. McDonald, M. E. 1981. Key to Trematodes Reported in Waterfowl. Resource publication 142, United States Department of the Interior, U.S. Fish and Wildlife Service, Washington DC, 156 pp. McLaughlin, J. D. 1976. Experimental studies on the life cycle of Cyclocoelum mutabile (Zeder) (Trematoda: Cyclocoelidae). Canadian Journal of Zoology 54:48–54. McLaughlin, J. D. 1977. The migratory route of Cyclocoelum mutabile (Zeder) (Trematoda: Cyclocoelidae) in the American coot, Fulica americana. Canadian Journal of Zoology 55:274–279. McLaughlin, J. D. 1983. Growth and development of Cyclocoelum mutabile (Cyclocoelidae) in coots, Fulica americana (Gm.). Journal of Parasitology 69:617–620. Menezes, R. C., D. G. Mattos-J´unior, R. Tortelly, L. C. Muniz-Pereira, R. M. Pinto, and D. C. Gomes. 2001. Trematodes of free range reared guinea fowls (Numida meleagris Linnaeus, 1758) in the state of Rio de Janeiro, Brazil: Morphology and pathology. Avian Pathology 30:209–214. Merino, S., J. Martinez, G. Alcantara, M. Navarro, S. Mas-Coma, and F. Rodriguez-Caabeiro. 2003. Pulchrosoma pulchrosoma (Trematoda: Cathaemasiidae) in ringed kingfishers (Megaceryle torquata torquata) from Iquitos, Peru: With inferences
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on life-cycle features. Avian Pathology 32:351– 354. Merino, S., J. Martonez, P. Lanzarot, L. S. Cano, M. Fernandez-Garco, and F. Rodroguez-Caabeiro. 2001. Cathaemasia hians (Trematoda: Cathaemasiidae) infecting black stork nestlings (Ciconia nigra) from central Spain. Avian Pathology 30:559–561. Mettrick, D. F. 1959. Zygocotyle lunata. A redescription of Zygocotyle lunata (Diesing,1836) Stunkard, from Anas platyrhynchos domesticus in southern Rhodesia. Rhodesia Agricultural Journal 56:197– 198. Miller, G. C., and R. Harkema. 1965. Studies on helminths of North Carolina vertebrates. IV. Parastrigea tulipoides sp. n., a trematode (Strigeida: Strigeidae) from the Red-Shouldered Hawk. Journal of Parasitology 51:21–23. Mimori, T., H. Hirai, T. Kifune, and K. Inada. 1982. Philophthalmus sp. in a human eye. American Society of Tropical Medicine and Hygiene 31:859–861. Minnesota Department of Natural Resources. 2007. Parasite Likely Cause of Scaup, Coot Deaths at Lake Winnibigoshish. Minnesota Department of Natural Resources, St. Paul, MN. Møller A. P., and J. Erritzøe. 1998. Host immune defense and migration in birds. Evolutionary Ecology 12:945–953. Mucha, K. H., J. E. Huffman, and B. Fried. 1990. Mallard ducklings (Anas platyrhynchos) experimentally infected with Echinostoma trivolvis (Digenea). Journal of Parasitology 76:590–592. Mucha, K. H., and J. E. Huffman. 1991. Inflammatory cell stimulation and wound healing in Sphaeridiotrema globulus experimentally infected mallard ducks (Anas platyrhynchos). Journal of Wildlife Disease 27:428–434. Murata, K., A. Noda, T. Yanai, T. Masegi, and S. Kamegai. 1998. A fatal Pegosomum sp. (Trematoda: Echinostomatidae) infection in a wild cattle egret (Bubulcus ibis) from Japan. Journal of Zoo and Wildlife Medicine 29:78–80. Nollen, P. M. 1971. Studies on growth and infection of Philophthalmus megalurus (Cort, 1914) (Trematoda) in chicks. Journal of Parasitology 57:261–266. Nollen, P. M., and I. Kanev. 1995. The taxonomy and biology of Philophthalmid eyeflukes. Advances in Parasitology 36:205–269. Nollen, P. M., and H. D. Murray. 1978. Philopthalmus gralli: Identification, growth characteristics and treatment of an oriental eyefluke of birds introduced into the continental United States. Journal of Parasitology 64:178–180. Patnaik, M. M., A. T. Rao, L. N. Acharjyo, and D. N. Mohanty. 1970. Notes on a nodular disease of the intestine of the open-billed stork—Anastomus
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oscitans caused by Chaunocephalus ferox. Journal of Wildlife Diseases 6:64–66. Pauly A., R. Schuster, and S. Steuber. 2003. Molecular characterization and differentiation of opisthorchiid trematodes of the species Opisthorchis felineus (Rivolta, 1884) and Metorchis bilis (Braun, 1790) using polymerase chain reaction. Parasitology Research 90:409–414. Pinto, R. M., R. C. Menezes, and R. Tortelly. 2004. Systematic and pathologic study of Paratanaisia bragai (Santos, 1934) Freitas, 1959 (Digenea, Eucotylidae) in ruddy ground dove, Columbina talpacoti (Temminck, 1811). Arquivo Brasileiro de Medicina Veterinaria e Zootecnia 56:472– 479. Poonswad, P., P. Chatikavanij, and W. Thamavit. 1992. Chaunocephalosis in a wild population of Asian open-billed storks in Thailand. Journal of Wildlife Diseases 28:460–466. Portugal, M. A. S. C., G. F. Oliveira, F. L. Fenerich, C. E. M. P. M. Cappellaro, and V. Chiarelli. 1972. Ocorrˆencia de Paratanaisia bragai (Santos, 1934) Freitas, 1959 (Trematoda, Euco-tylidae), em pomba dom´estica (Columba livia domestica). Arquivos Do Instituto Biologico 39:189–194. Price, E. W. 1934. Losses among wild ducks due to infestation with Sphaeridiotrema globulus (Rudolphi) (Trematoda: Psilostomidae). Proceedings of the Helminthological Society of Washington 1:32–34. Pritchard, M. H., and G. O. W. Kruse. 1982. The Collection and Preservation of Animal Parasites. University of Nebraska Press, Lincoln, NE. Robertson, E. L., and C. H. Courtney. 1995. Veterinary Pharmacology and Therapeutics, H. R. Adams (ed.). Iowa State University Press, Ames, IA. Roscoe, D. E., and J. E. Huffman. 1982. Case report—Trematode (Sphaeridiotrema globulus)—Induced ulcerative hemorrhagic enteritis in wild mute swans (Cygnus olor). Avian Diseases 26:214–224. Roscoe, D. E., and J. E. Huffman. 1983. A lethal Sphaeridiotrema globulus infection of a whistling swan. Journal of Wildlife Diseases 19:370–371. Schell, S. C. 1957. Dicrocoeliidae from birds in the northwest. Transactions of the American Microscopical Society 76:184–188. Schmidt, R. E., and G. P. Hubbard. 1987. Atlas of Zoo Animal Pathology, Vol II. CRC Press, Boca Raton, FL, 192 pp. Schmidt, R. E., and J. D. Toft, III. 1981. Ophthalmic lesions in animals from a zoologic collection. Journal of Wildlife Diseases 17:267–275. Scott, M. E., M. E. Rau, and J. D. McLaughlin. 1980. Prevalence and intensity of Typhlocoelum
cucumerinum (Digenea) in wild anatids of Quebec, Canada. Journal of Wildlife Diseases 16:71– 75. Silva C. C., D. G. Mattos-J´unior, and P. M. Ramirez. 1990. Helmintos parasitos de Columba livia (Gm.) no munic´ıpio de S˜ao Gon¸calo, Rio de Janeiro. Arquivo Brasileiro de Medicina Veterinaria e Zootecnia 42:391–394. Skrjabin, K. I. 1926. Infestation simultanee d’un oiseau de Transbaikalie. Annales de Parasitologie Humaine et Comparee 6:80–87. Smith, H. J. 1978. Cryptocotyle lingua infection in a bald eagle (Haliaeetus leucocephalus). Journal of Wildlife Diseases 14:163–164. Snyder, P. E. 1991. A Study of the Humoral Immune Response in Chickens to the Eyeflukes, Philophthalmus gralli, and Philophthalmus megalurus. Masters Thesis, Western Illinois University, MaComb, IL. Soulsby, E. J. L. 1965. Textbook of Veterinary Clinical Parasitology. F.A. Davis, Co. Philadelphia, PA, 824 pp. Stoskopf, M. K., S. Patton, and E. Bueding. 1982. Treatment of two marabou storks (Leptoptilos crumeniferus) infected with the esophageal fluke (Cathaemasia spectabilis). Journal of Zoo Animal Medicine 13:51–55. Swales, W. E. 1933. On Streptovitella acadiae (gen et spec. nov.), a trematode of the family Heterophyidae from the black duck (Anas rubripes). Journal of Helminthology 11:115–118. Tabery, P., J. E., Huffman, and B. Fried. 1988. Hemolytic and coagulation properties of Sphaeridiotrema globulus (Trematoda). Journal of Parasitology 74:730–731. Taft, S. J. 1971. Incidence of the trematode family Cyclocoelidae in some North American birds. Journal of Parasitology 57:831. Threlfall, W. 1986. Parasites: An ignored factor in the study of the energetics and food of seabirds. Pacific Seabird Group 13:110–111. Town, R. H. 1960. A Survey of Helminth Parasites in Diving Ducks Found Dead in the Lower Detroit River. M.S. Thesis, University of Michigan, 61 pp. Trainer, D. O., and G. W. Fischer. 1963. Fatal trematodiasis of coots. Journal of Wildlife Management 27:483–486. Travassos, L., J. F. T. Freitas, and A. Kohn. 1969. Tremat´odeos do Brasil. Memorias do Instituto Oswaldo Cruz 67:1–886. Underhill, L. G., R. A. Earle, T. Piersma, I. Tulp, and A. Veister. 1994. Knots (Calidris canutus) from Germany and South Africa parasitized by trematode Cyclocoelum mutabile. Journal f¨ur Ornithologie 135:236–239.
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Trematodes United States Geological Survey 1997. National Wildlife Health Center, Quarterly Wildlife Mortality Report October–December 1997. Available at http://www.nwhc.usgs.gov United States Geological Survey 2007. Exotic Parasite of American Coot Discovered in Exotic Snail in Lake Onalaska. Wildlife Health Bulletin 07-01. National Wildlife Health Center, Madison, WI. Van Haitsma, J. P. 1931. Studies on the trematode family Strigeidae (Holostomidae). No. XXII. Cotylurus flabelliformis (Faust) and its life-history. Papers of the Michigan Academy of Science and Arts and Letters 13:447–483. West, A. F. 1961. Studies on the biology of Philopthalmus gralli Mathis and Iyer 1910 (Trematoda: Digenea). American Midland Naturalist 66:363–383.
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Willey, C. H. 1941. The life history and bionomics of the trematode, Zygocotyle lunata (Paramphistomidae). Zoologica: Scientific Contributions of the New York Zoological Society 26:65–88. Willey, C. H., and H. W. Stunkard. 1942. Studies on pathology and resistance in terns and dogs infected with the heterophyid trematode, Cyathocotyle lingua. Transactions of the American Microscopical Society 61:236–253. Wilson, W. D., P. T. J. Johnson, D. R. Sutherland, H. Mon´e, and E. S. Loker. 2005. A molecular phylogenetic study of the genus Ribeiroia (Digenea) trematodes known to cause limb malformations in amphibians. Journal of Parasitology 91:1040–1045. Yamaguti, S. 1971. Synopsis of Digenetic Trematodes of Vertebrates. Interscience Publishers, Inc., New York, 979 pp.
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13 Schistosomes Jane E. Huffman and Bernard Fried further 60 years before the life cycle was elucidated in 1912 by Fujinami in Japan (Mahmoud 2004). An 1855 description by LaValette of an erythemous maculopapular eruption is presumed to have been cercarial dermatitis, although its relationship to the avian schistosomes of waterfowl was not yet known. Karube, a maculopapular rash, was also common among rice farmers who spent a great deal of time in cercariaeinfested water. The first avian schistosome life cycle was described by Oiso (1927) for Bilharziella yokogawai from domestic ducks. In the American scientific literature, Cort (1950) was the first to demonstrate that swimmer’s itch was caused by the cercariae of nonhuman schistosomiasis in 1928. Prior to that, it was believed that all such eruptions were caused by human schistosomiasis.
INTRODUCTION The avian schistosomes are a specialized group of trematodes that develop as adults within the circulatory system or nasal tissue of their avian hosts. They comprise the largest and most diverse clade of the family Schistosomatidae (Brant et al. 2006) and include nine genera: Allobilharzia, Austrobilharzia, Bilharziella, Dendritobilharzia, Gigantobilharzia, Jilinobilharzia, Macrobilharzia, Ornithobilharzia, and Trichobilharzia. Like other trematodes (Chapter 12), avian schistosomes have a two-host life cycle and use freshwater snails as intermediate hosts. Trichobilharzia is the most extensively studied genus (Hor´ak and Kol´aov´a 2005). Schistosomes are important pathogens of birds in areas where intimate contact with infected snails occurs. Pulmonary lesions may be evident for species that live in blood vessels of the visceral organs, while neurological signs may be evident for species of nasal schistosomes that undergo larval development within tissues of the central nervous system (CNS). Avian schistosomes are frequently responsible for human cercarial dermatitis or “swimmer’s itch”—a skin rash caused by penetration of the skin by free living cercarial stages of species of Gigantobilharzia, Ornithobilharzia, Trichobilharzia, and Austrobilharzia.
HOST RANGE AND DISTRIBUTION Migratory water birds, including shorebirds, ducks, and geese, are the most typical hosts for avian schistosomes and movements of infected birds along major migratory flyways may play an important role determining their distribution in North America (Jarcho and van Burkalow 1952), Asia and the Pacific basin (Chu 1958), and Europe and Africa (Moreau 1972). Species of Austrobilharzia, Ornithobilharzia, Bilharziella, Trichobilharzia, Gigantobilharzia, and Dendritobilharzia are cosmopolitan in distribution, while others appear to be more regional. Nasal schistosomes appear to be frequent parasites of birds in central Europe (Rudolfov´a et al. 2002), but it is not clear if this is a sampling artifact. Their small size, threadlike shape, and cryptic life in the nasal mucosa may make them difficult to detect if not specifically searched for. Schistosomes are associated primarily with freshwater habitats in all temperate and tropical regions of the world and typically mirror the distribution of their snail intermediate hosts (Table 13.1). Larval Ornithobilharzia and Austrobilharzia are both parasites of marine caenogastropods and are found as adults primarily
SYNONYMS Trichobilharziasis, cercarial dermatitis, schistosomiasis, swimmer’s itch.
HISTORY Schistosomiasis has a very long history. The first description of the disease in humans occurs some 3,000 years ago in the Egyptian medical papyrus, the Papyrus Ebers, and clearly described the symptoms, including blood in the urine. Bilharz working in Egypt discovered the adult human parasites in 1852, but it was a
246 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Schistosomes Table 13.1. Known intermediate and final hosts of avian schistosomes (Horak ´ et al. 2002). Parasite genus
Geographic location
Intermediate host
Habitat
Allobilharzia Austrobilharzia
Iceland Worldwide
Unknown Prosobrachia
U SW
Bilharziella Dendritobilharzia
Northern Hemisphere Worldwide
FW FW
Gigantobilharzia
Worldwide
Pulmonata Pulmonata Opisthobrachia Pulmonata Opisthobrachia
Jilinobilharzia Macrobilharzia Ornithobilharzia
China Worldwide Northern Hemisphere
Pulmonata Unknown Prosobrachia
FW U SW
Trichobilharzia
Worldwide
Pulmonata
FW
FW, SW
Avian host order Anseriformes Anseriformes Charadriiformes Ciconiiformes Anseriformes Anseriformes Phoenicopteriformes Anseriformes Charadriiformes Ciconiiformes Passeriformes Anseriformes Pelecaniformes Charadriiformes Pelecaniformes Anseriformes Ciconiiformes Columbiformes Coraciiformes Galliformes Passeriformes Pelecaniformes
FW, larval development in fresh water; SW, larval development in sea water; U, Unknown. in gulls. Larval Bilharziella, Trichobilharzia, and Gigantobilharzia are parasites of pulmonate snails and as adults are found in a broad diversity of birds, including ducks, geese, grebes, and passerine birds. Dendritobilharzia larvae are parasites of pulmonate snails, but as adults occur only in ducks and grebes. ETIOLOGY General descriptions of the family Schistosomatidae and taxonomic histories of these parasites can be found in Farley (1971) and Gibson et al. (2002). Members of the Schistosomatidae have common life histories and patterns of transmission. All have brevifurcate furcocercous or fork-tailed cercariae that develop to sexual maturity following direct penetration of the host (Yamaguti 1975) and typically occupy the circulatory system as adults. Schistosomes are venous specialists with the exception of Dendritobilharzia pulverulenta, which inhabits the mesenteric arteries of ducks (Vande Vusse 1979; Platt and Brooks 1997). There are relatively few morphological distinctions between the male and female avian schistosomes when compared to other members of this family. Males are considerably larger than females and possess a gynecophoric canal. This canal is a ventral groove run-
ning the length of the male schistosome into which the threadlike female worm fits. Presence or absence of this canal, relative size of the canal, presence or absence of an oral sucker, and morphological characteristics of the intestinal ceca are important characters for distinguishing the genera of avian schistosomes (Gibson et al. 2002). The Schistosome Group Prague has made significant advances in the systematics and biology of these worms in Europe. The genus Trichobilharzia is the most species-rich genus within the family, with over 40 recognized species. The entire mitochondrial genome of Trichobilharzia regenti has been recently sequenced and annotated (Webster et al. 2007). The gross features of the genome of T. regenti are similar to those of mammalian schistosomes that have been characterized, and the mitochondrial genome is identical to the human parasite, Schistosoma japonicum, in terms of gene order. Intrinsic properties of the mitochondrial genome of T. regenti include potentially useful markers, repeat regions and multiple individual genes that may be useful for development of molecular markers for diagnostic, epidemiological, and population level studies. Other studies of the phylogenetics of this group have used molecular markers, morphological characters, intermediate and definitive host associations, and
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biogeography to investigate relationships among genera (Snyder and Loker 2000; Lockyer et al. 2003). Results of these studies indicate that the avian clade consists of six genera of exclusively avian parasites and two genera of mammalian flukes from North America. This study provides little evidence concerning the identity of ancestral molluscan or vertebrate schistosome hosts but does demonstrate that host switching has been an important feature of schistosome evolution. Evidence also indicates that the reduced sexual dimorphism characteristic of some avian schistosomes is derived evolutionarily. EPIZOOTIOLOGY Adult schistosomes usually reside in veins around the intestine of their avian hosts and release eggs that make their way into the digestive tract and then pass out of the host with the feces. If the eggs are deposited in water, they will hatch within an hour if conditions are right and release a ciliated, free-swimming, nonfeeding aquatic stage, the miracidium. The miracidium has enough energy to keep moving for about a day. Once the miracidium comes in contact with a suitable snail, either it will penetrate the integument or it may be ingested through the mouth. Avian schistosomes exhibit a high specificity toward snail intermediate hosts. The molecules involved in attraction are soluble macromolecular miracidiaattracting glycoproteins, the carbohydrate moieties of which are responsible for signal specificity (Hass 2003). Within the snail, the miracidium will elongate to form a reproductive sac called the sporocyst. This germinating structure will produce a second generation of sporocysts. In approximately 30 days, the sporocysts produce cercaria. The cercariae are furcocercous (have a forked tail). After leaving the snail, the cercariae swim freely in a zigzag pattern. They are negatively geotropic, positively phototropic, and rest occasionally by grasping the water surface or debris with the ventral sucker (Rind 1991). Water temperature and exposure to sunlight are principal determinants of the life span of cercariae. Cercarial die-off increases during hot and sunny days (Mulvihill and Burnett 1990). The life span for the cercariae is variable but averages about 24 h. Infective schistosome cercariae normally gain entry to a mammalian or avian host by attaching to the skin with the ventral sucker and using a number of proteolytic enzymes to digest a route to reach blood capillaries or lymphatic vessels (McKerrow and Salter 2002; Mountford and Trottein 2004). Cercariae can also be ingested and then enter the blood vessels in the walls of the pharynx or esophagus. Cercariae of Trichobilharzia ocellata exhibit low specificity in recognizing their definitive host species and will there-
fore penetrate mammals (Hass and van de Roemer 1998). Cercarial attachment to, and enduring contact with, the vertebrate skin can be stimulated by temperature and chemical signals (ceramides and cholesterol), whereas the penetration itself is triggered by fatty acids (Hass 2003). Upon registering relevant stimuli, the cercariae start to release the contents of their penetration glands. Several proteins with activities probably playing a role in penetration have been detected in cercarial hom*ogenates and/or secretions (Hor´ak and Kol´arˇov´a 2005). Once in the skin, schistosomula, the immature forms of a schistosome after they have entered the blood vessels of their host, need to navigate an appropriate route to the target tissue. They move toward deeper skin layers and search for a blood vessel. They are able to monitor concentration gradients of chemical stimuli (d-glucose and l-arginine) and exhibit chemotactic orientation (Grabe and Haas 2004a). The schistosomes may use negative photo-orientation to move away from the light source into deeper skin layers (Grabe and Haas 2004b). Schistosomes traverse the skin of their primary host within days, and the vast majority enter the circulation and migrate to specific locations in the host to complete their development (Table 13.2). The CNS is the most likely route to the nasal cavity for T. regenti and other species that complete their development in this location. Once sexual maturity is reached, adult worms produce large numbers of eggs that are placed precisely in the venous system because they are released against the blood flow. Eggs are sequestered usually within the portal system of the avian host, thus restricting egg dispersal. Male and female schistosomes are permanently paired while they inhabit the bloodstream of their vertebrate hosts. Female schistosomes produce eggs only when they are in intimate association with a male. The natural elasticity of the vessel serves to hold the eggs in place against the flow of blood (Basch 1991). Endothelial cells actively migrate over the eggs and passively transfer the egg to the perivascular space, where they are subject to the host immune response (File 1995). A significant number of eggs may escape into the external environment before a heavily infected host is incapacitated by or dies from the infection (Platt and Brooks 1997). Avian schistosomes usually complete their life cycle in 2 months; however, the specific time varies with each species. Transmission of avian schistosomes depends on their ability to find, recognize, penetrate, and prosper within appropriate intermediate and final hosts. This requires relevant host signals monitored by the parasite for orientation and migration purposes, but also the ability of parasites to evade host immune reactions and
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Schistosomes Table 13.2. Anatomical location of some common species of adult avian schistosomes. Tissue Veins of nasal fossae
Dorsal aorta Hepatic portal, mesenteric and intestinal veins Renal veins Interlobular bile ducts Pulmonary arteries Central nervous system, spinal cord, and brain
Parasite species Trichobilharzia duboisi Trichobilharzia arcuata Trichobilharzia nasicola Trichobilharzia rodhaini Trichobilharzia spinulata Trichobilharzia regenti Trichobilharzia aureliani Trichobilharzia australis Dendritobilharzia pulverulenta Austrobilharzia spp. Bilharziella spp. Gigantobilharzia spp. Trichobilharzia spp. Ornithobilharzia spp. Bilharziella spp. Trichobilharzia spp. Bilharziella spp. Trichobilharzia spp. Bilharziella spp. Trichobilharzia spp. Trichobilharzia regenti
manipulate other regulatory systems of the host (Hor´ak and Kol´aˇrov´a 2005). A serine protease was characterized from T. ocellata by Bahgat and Ruppel (2002) which may aid in facilitating the migration of avian schistosomes through host skin. Environmental factors such as water temperature, degree of pollution, extent of algal growth/aquatic plant habitats, aquatic bird, and coincident appropriate snail intermediate host populations influence the distribution and incidence of avian schistosomes. Most birds probably become infected in nesting areas; however, birds may also transport mature (within the host) or larval (within snails transported on legs or plumage) schistosomes to and from the wintering locations (Woodruff and Mulvey 1997; Wesselingh et al. 1999). CLINICAL SIGNS Clinical signs of avian schistosomiasis are nonspecific and include weight loss, lameness, and “ill-thrift” (Wojcinski et al. 1987). After experimental exposure to the cercariae of T. ocellata, American Black Ducks (Anas rubripes), Blue-winged Teal (Anas discors), Muscovy Ducks (Cairina moschata), and Rouen ducks
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(Anas platyrhynchos) develop inflammation of the feet marked by engorgement of blood vessels and petechial hemorrhages (Rau et al. 1974). Listlessness, compulsive swallowing, respiratory distress, and occasional mild pulmonary hemorrhage are evident at 2 and 5 days after exposure to cercariae. Severity of these signs vary, at times being detectable only when the ducks were excited or under stress. Most birds produce mucoid, blood-flecked feces when they excrete large numbers of eggs. Neurological signs have been observed in domestic ducklings infected experimentally with T. regenti, including leg paralysis and problems with orientation and balance (Hor´ak et al. 1999). Infections with schistosomes can also affect host hematology. Total leukocytes increase significantly in chickens infected with Austrobilharzia variglandis and peak at day 21 days postinfection (PI) (Ferris and Bacha 1986). Leukocyte counts decline over the next 3 weeks, returning to normal by day 42 PI. Increases in heterophils and monocytes are related to schistosome egg burden. PATHOGENESIS AND PATHOLOGY Schistosome infections in mammals cause chronic proliferative vascular lesions associated with the presence of adult parasites in the lumen of mesenteric and portal veins. In birds, however, these lesions have never been reported. Lesions in avian hosts include obliterative endophlebitis associated with the presence of adult schistosomes in intestinal and portal veins, moderate to severe lymphocytic and granulocytic enteritis associated with release of eggs by adult worms (van Bolhuis et al. 2004), and inflammatory reactions associated with migration of larvae in a variety of tissues, including the CNS. Most of what is known about the pathogenesis of avian schistosome infections is based on experimental studies that have documented migration of larvae, maturation of adults, and their associated host reactions. Neurological signs are associated with the pathogenesis of nasal schistosomes and related to development of migrating larval stages in the CNS. Larval schistosomes are evident in the thoracic spinal cord of domestic ducklings with experimental infections of T. regenti by day 3 PI and are present in the synsacral and cervical portions of the spinal cord by day 6–7 PI (Hor´ak et al. 1999). Worms are present in the cerebellum, cerebral hemispheres, ocular lobes/nerves, and nasal lobes, between day 10 and 13 PI. Adults appear in the nasal region by day 13 PI, and eggs can be detected by day 14 PI. Eggs have never been observed in the CNS. In the CNS, the parasites develop outside the blood vessels, directly in the nerve tissue, and on
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nerve-associated cells. An inflammatory reaction with lymphocytic infiltration and undetermined degenerative changes develops in response to parasites or their tracks during migration in the nerve tissue (Hor´ak et al. 1999). During development of T. regenti in the CNS, immature parasites are located either in meninges or in various parts of the spinal cord and brain (Kol´aˇrov´a et al. 2001). In the spinal cord, the submeningeal location causes a strong inflammatory reaction around migrating schistosomula, resulting in eosinophilic meningitis. In the white and gray matter of the spinal cord and in the white matter of the brain, a cellular infiltration and spongy tissue surrounds the immature parasites. Dystrophic and necrotic changes of neurons, perivascular eosinophilic inflammation in the spinal cord and brain, and cell infiltration around the central canal of the spinal cord have been observed (Kol´aˇrov´a et al. 2001). Both adults and eggs are eventually detected in the nasal mucosa of infected ducklings. As eggs age, various host reactions are evident, ranging from focal accumulation of inflammatory cells to the formation of granulomas. Avian schistosomes that develop in blood vessels surrounding abdominal organs typically migrate through the lungs before reaching their final destinations. Following experimental infection of Muscovy Ducks (Cairina moschata) and American Black Ducks (Anas rubripes) with cercaria of T. ocellata, parasites are found in the lungs and kidneys at 19 h PI and in the liver at 24 h PI (Bourns et al. 1973). In the lung, schistosomula break free into the air spaces. They appear first in air capillaries and parabronchi and later in secondary bronchi where they reinvade the bronchial epithelium and gain entrance into veins. Worms in the liver are found in the sinusoids but mainly in hepatic portal veins. From here, worms move initially to peripheral veins of the small intestine, but later penetrate deep into the mucosa, sometimes approaching the tips of the villi. Adult worms tended to be evenly distributed between Meckel’s diverticulum and the ceca while eggs are most abundant in or near Meckel’s diverticulum and the lymphatic tissue immediately posterior to it (Bourns et al. 1973). Birds release viable eggs of T. ocellata as early as 2 weeks PI. Mallard ducklings (Anas platyrhynchos) experimentally infected with T. ocellata had liver damage, but the lesions were not discussed (McMullen and Beaver 1945). In Mallards, parasites reach the lungs within 24 h PI and remain at this site for at least 5 days. Considerable growth and development takes place in the lungs, and damage may be sufficient to cause death. Among other species of schistosomes that migrate through the lungs, a host inflammatory reaction and development of nodules composed of infiltrated lympho-
cytes, heterophils, eosinophils, and macrophages may occur in association with migrating schistosomula. These structures form around the blood vessels and in the gas-exchange tissues of the parabronchial walls and, consequently, in the walls of secondary bronchi in domestic ducks with experimental infections of Trichobilharzia szidati (Chanov´a et al. 2007). Extensive inflammation of secondary bronchi and parabronchi may be evident. Host responses to eggs released by adult worms are common in schistosome infections. In one detailed study of egg deposition by A. variglandis in the mesenteric veins of domestic chickens, paired adults were observed in the mesenteric veins, or branches, paralleling the mesenteric border of the intestine (Wood and Bacha 1983). Females traveled from vessels in the serosa through the muscularis, squeezed into the small veins of the mucosa, and then withdrew back to the serosa after releasing eggs. As the infection progressed, deposition of eggs occurred in more peripheral layers of the intestine as smaller vessels became constricted from edema and cellular infiltration. This led to edema of the lamina propria and longer villi and expanded crypts were noted in the intestinal mucosa (Wood and Bacha 1983). Granulomatous responses to the presence of eggs were observed from day 12 to 18 weeks PI. Granulomas contained combination of macrophages and lymphocytes, giant cells, epithelioid cells, plasma cells, fibroblasts, eosinophils, and heterophils. The granulomas ranged from dense accumulations of macrophages and lymphocytes to fully developed granulomas. Phagocytosis by giant cells and the Hoeppli phenomenon was reported. Among wild birds with natural schistosome infections, the host response to infection depends both on schistosome and on host species. Most lesions are associated with release of eggs that become lodged in veins associated with adult worms. These include development of granulomas, infiltration of heterophils and leukocytes, proliferation of connective tissue, and calcification of parasite eggs. Depending on species, lesions may develop in the mesenteric and pelvic veins, intestinal mucosa, liver, lungs, pancreas, cerebellum, and gizzard (Table 13.3). In abnormal hosts, adult schistosomes may be found in scattered locations within the arterial system of a wide range of tissues. Here they typically produce few eggs that usually fail to embryonate. Mortality may occur when birds are translocated into new habitats and exposed to schistosome species they would not normally encounter. The death of 36 Brant (Branta bernicla hrota) was attributed to infections with D. pulverulenta and Trichobilharzia spp. when birds were translocated from a marine environment to a freshwater
251 Continental US North America Manitoba, Canada North America North America Japan Africa China New Mexico, USA East Africa North America Japan North America
Visceral tissue Intestinal veins Visceral tissue Visceral tissue Visceral tissue Intestinal veins NA Portal veins Mesenteric veins Mesenteric veins Mesenteric veins Mesenteric veins Visceral tissue, liver
huronensis huttoni lawayi nettapi plectropteri sturniae tantali crecci pulverulenta
Trichobilharzia
Ornithobilharzia
Pelecaniformes Charadriiformes Passeriformes Anseriformes
North America North America Africa North America
acotylea adami ardeola gyrauli
Gigantobilharzia
baeri canaliculata emberizae adamsi
Charadriiformes Charadriiformes Ciconiiformes Anseriformes Passeriformes Passeriformes Anseriformes Charadriiformes NA Charadriiformes Passeriformes Ciconiiformes Anseriformes Pelecaniformes
Chile
Visceral tissue, pancreas, spleen, liver, kidneys Gastrosplenic vein Visceral tissue Visceral tissue Visceral tissue
sp.
Dendritobilharzia
Jilinobilharzia Macrobilharzia
Phoenicopteriformes
North America, Europe North America, New Zealand
variglandis
Charadriiformes Ciconiiformes Anseriformes Charadriiformes Ciconiiformes Anseriformes Anseriformes Pelecaniformes
Canada, continental US, Hawaii, and Australia Worldwide
polonica Mesenteric and portal veins pulverrulenta Visceral tissue, heart
Mesenteric veins
terrigalensis
Austrobilharzia
Anseriformes
Host order
Iceland
Geographic location
Strohm et al. (1981) Leigh (1955) Farley (1963) Farley (1963) Farley (1963) Oshima et al. (1991, 1992) Fain (1955) Liu and Bai (1976) Price (1929), Fain (1955), Baugh (1963), and Kohn (1964) Fain (1955) Kol´aˇrov´a et al. (1997) Uchida et al. (1991) McDonald (1969) (continues)
Ulmer (1968) Farley (1963) Farley (1963) Guth et al. (1979)
Kol´aˇrov´a et al. (1997) Cheatum (1940), Kinsella et al. (2004), and Davis (2006) Pare and Black (1999)
Barber and Caira (1995)
Rohde (1977)
Kol´aˇrov´a et al. (2006)
Reference
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Bilharziella Dendritobilharzia
Intestinal blood vessels and mesenterium Mesenteric veins
Tissue
visceralis
Species
Allobilharzia
Genus
Table 13.3. Genera of avian schistosomes with the location of the parasite in the host, geographic location, vertebrate host order, and selected references.
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Genus North America Central Africa Africa Australia Australia Central Africa North America Japan North America North America Africa Japan Africa North America North America, Europe Europe North America Asia South America, Japan Asia North America Europe
Visceral tissue Visceral tissue, cloacal vein Visceral tissue Visceral tissue Visceral tissue Nasal tissue Visceral tissue, venules of the intestinal wall Visceral tissue Hepatic and enteric veins Visceral tissue Visceral tissue Visceral tissue Visceral tissue Visceral tissue, cloacal vein Visceral tissue, hepatic vessels
brevis burnetti
cameroni
cerylei corvi
duboisi elvae
filiformis
franki
horiconensis
indica jequitibaensis jianensis kegonsensis
kossarewi
Geographic location
Visceral tissue Visceral tissue Nasal tissue Nasal blood vessels Nasal tissue Visceral tissue Serosal and mesenteric veins
Tissue
alaskensis anatina aureliani australis arcuata berghei brantae
Species
Table 13.3. (Continued)
252
Anseriformes
Anseriformes Anseriformes Anseriformes Anseriformes
Anseriformes Dwarf mallards* Anseriformes
Anseriformes Passeriformes Coraciiformes Passeriformes Galliformes Anseriformes Anseriformes Passeriformes Anseriformes
McMullen and Beaver (1945) Baugh (1963) Leite et al. (1978) Liu and Bai (1976) McMullen and Beaver (1945) Rudolfov´a et al. (2005)
Fain (1956)b McMullen and Beaver (1945) McMullen and Beaver (1945); Rudolfov´a (2001) M¨uller and Kimmig (1994)
Fain (1956)b Ito (1960)
Becker (1956) Fain (1955) Fain (1956)a Hor´ak et al. (1998a) Islam (1986) Fain (1955) Farr and Blankemeyer (1956) Uchida et al. (1991) McMullen and Beaver (1945) Wu (1953)
Reference
October 15, 2008
Anseriformes Anseriformes
Anseriformes Anseriformes Podicipediformes Anseriformes Anseriformes Anseriformes Anseriformes
Host order
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China Australia North America, Japan Poland, Ukraine North America Europe, Australia Central Africa
Visceral tissue Visceral tissue Nasal tissue Nasal tissue
polonica querquedulae
regenti rodhaini
253 Visceral tissue Nasal tissue Visceral tissue, intestinal veins Lungs, hepatic-portal system Visceral tissue, intestinal and cloacal veins Europe North America
Central Africa East Africa North America
Anseriformes Anseriformes Charadriiformes Passeriformes Anseriformes Anseriformes
Anseriformes Anseriformes Ciconiiformes Anseriformes
Anseriformes Anseriformes Anseriformes, Columbiformes, Passeriformes Anseriformes Anseriformes
Simon-Martin and Simon-Vicente (1999) Fain (1955) Fain (1955) McMullen and Beaver (1945) Neuhaus (1952) McMullen and Beaver (1945)
˙ Zbikowska (2002) McMullen and Beaver (1945) Hor´ak et al. (1998)b Fain (1955)
Hu et al. (1994) Islam and Copeman (1986) McDonald (1969) and Yokogawa et al. (1976)
McMullen and Beaver (1945) Farley (1971) Kruatrachue et al. (1968) Fain (1955) Loken et al. (1995) and DeGentile et al. (1996) McDonald (1969)
NA, Not available. * Experimental hosts. † The taxonomical validity of Trichobilharzia ocellata is doubtful. Trichobilharzia ocellata should be regarded as species inquirenda (Rudolfov´a et al. 2005). In this chapter, we have used the original terminology used by the cited authors.
szidati waubesensis
schoutedeni spinulata stagnicolae
salmanticensis Visceral tissue
paoi parocellata physellae
Anseriformes
Passeriformes Anseriformes Anseriformes Anseriformes
Anseriformes
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Europe
North America Asia Africa Europe, North America, Asia North America
Visceral tissue Visceral tissue Nasal tissue Intestinal veins, lungs, liver, intestine Visceral tissue, portal, cecal and intestinal veins Visceral tissue Visceral tissue Liver, mesenteric veins
littlebi maegraithi nasicola ocellata†
oregonensis
Europe, Asia
Visceral tissue
kowalewskii
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pond. The birds became emaciated and dehydrated with reduced pectoral muscle mass and prominent keels. Pathological findings were attributed both to eggs and to adult schistosomes and included emaciation, thrombosis of the caudal mesenteric vein and its branches, fibrinohemorrhagic colitis, and in some birds, heptomegaly and pulmonary congestion. Gallbladders were distended with bile and gastrointestinal tracts were devoid of ingesta (Wojcinski et al. 1987). DIAGNOSIS Live birds can be readily checked for nasal schistosomiasis by making a smear of nasal mucous using a cotton swab soaked in 0.85% saline. Eggs can be recovered but usually only when infections are intense (Blair and Ottesen 1979). Adult worms recovered from naturally infected definitive hosts may be identified by morphological characteristics (Farley 1971) and are the most valuable for making identifications to level of species (Blair and Islam 1983). However, adult schistosomes may be knotted together or fragmented and are difficult to remove intact from infected hosts (Basch 1966). Intact specimens of Trichobilharzia can be successfully collected from ducks by exsanguination and retrograde perfusion of the descending aorta (Li et al. 1999). Large numbers of living adult worms were collected by this method. Species of Dendritobilharzia inhabit the arterial system of their avian hosts. All other bird schistosomes live in the venous system (Platt and Brooks 1997). Eggs, miracidia, and cercariae can also provide diagnostic characteristics, but are most useful when the life cycle, developmental stages, and host specificity of the parasites are known. Keys are available for adult males of the genus Trichobilharzia (McDonald 1981). Descriptions of eggs of eight different avian schistosomes from birds in South Africa are available and were used to successfully place parasites in one of four possible genera: Austrobilharzia, Gigantobilharzia, Trichobilharzia, or Ornithobilharzia (Appleton 1986). Schistosome infections can also be detected in birds by allowing miracidia to hatch from eggs in the feces. It is possible to determine the relative intensity of infection by weighing the fecal content and then counting the number of miracidia that hatch from 1 g of feces. Molecular methods are becoming increasingly important for diagnosing and identifying schistosome infections, both in avian and intermediate hosts and in environmental samples. Internal transcribed spacers (ITS1 and ITS2) and the 5.8s ribosomal RNA gene of three European species of Trichobilharzia have been used successfully for identification of species within this genus, and primers developed for these genes may
be particularly useful for diagnosing infection with T. regenti, a potential neuropathogen (Dvo´ak 2001; Dvo´ak et al. 2002). Primers based on a 396 bp tandem repeated DNA sequence (T1323) that was cloned from DNA isolated from T. ocellata have made it possible to identify Trichobilharzia franki, T. ocellata, and T. regenti in both snails and plankton collections. The T1323 sequence represents between 1 and 2% (7,000– 14,000 copies) of the genome of the three Trichobilharzia species. Polymerase chain reaction primers, based on the T1323 sequence, are much more sensitive and also highly specific. They are sensitive enough to identify as few as one cercaria in a 0.5 g plankton sample and two cercariae in a 0.5 g sample of snail (Lymnea stagnalis) tissue (Hertl et al. 2002). Avian schistosomes also induce production of specific antibodies, which may provide a certain level of protection in birds and which can be used to identify infected individuals. Assays have been developed to detect antibodies specific to cercarial antigens (Kol´aˇrov´a et al. 1994) and to gut-associated antigens of immature and adult flukes (Kouˇrilov´a and Kol´aˇrov´a 2002). Isoforms of gut-associated cathepsin B have recently been isolated from schistosomula of T. regenti and may be a useful candidate antigen for diagnostic purposes. Sera collected from ducks experimentally infected with T. regenti have antibodies that bind histological sections of the gut surface of schistosomula (Dvo´ak et al. 2005) and Western blots of recombinant cathepsin B (Hor´ak and Kol´aˇrov´a 2005).
IMMUNITY Schistosomiasis has been referred to as an immunologic disease, and the pathogenesis of acute and chronic schistosomiasis appears to involve immunologic mechanisms, either humoral or cell mediated, that affect both the duration of the infection in birds and the severity of lesions. There are fundamental mechanisms of immune evasion that dictate whether schistosomes succeed in intravascular environments in humans and other vertebrate hosts that are not completely understood (Brant and Loker 2005). Challenge infections lead to a stronger inflammatory response around migrating parasites (T. ocellata in the lungs of birds) (Bourns and Ellis 1975). Avian schistosomes also induce production of specific antibodies that may provide a certain level of protection in birds. This has been shown in ducks, where transfer of large amounts of immune serum from donor birds to recipients was followed by partial or complete reduction in the number of eggs of T. ocellata in the feces and retardation of worm growth (Ellis et al. 1975).
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Schistosomes PUBLIC HEALTH CONCERNS Schistosome dermatitis is known as swimmer’s itch, clam digger’s itch, cercarial dermatitis, Gulf Coast itch, and sea bather’s eruption. It is caused by the penetration of cercariae of nonhuman schistosomes into the skin of humans. On first exposure, it produces a mild erythema and edema, but on repeated exposure a marked reaction occurs with pruritus, vesicle formation, and marked papule formation. Skin lesions may be accompanied by a systemic febrile response that runs for 5–7 days and resolves spontaneously (Hoeffler 1974). The life cycle of avian schistosomes also favors cercarial dermatitis—peak cercarial production occurs in the hottest months when bathing is most common (Cort 1950). A number of species of schistosomes have been implicated as the causative agents of cercarial dermatitis. Foci of infection can be found along migratory routes of waterfowl (Blair and Islam 1983). In Europe and North America, the species most commonly associated with cercarial dermatitis in freshwater habitats are Trichobilharzia stagnicola, Trichobilharzia physellae, and T. ocellata. A. variglandis is a cosmopolitan species and is associated with cercarial dermatitis in saltwater. Treatment may not be necessary when there are only a few itching spots. An antihistaminic or mild corticosteroid cream purchased over the counter in pharmacies can be beneficial. If the initial itching is severe, then scratching can cause abrasions and skin infections may develop. DOMESTIC ANIMAL HEALTH CONCERNS Farming of domestic ducks, geese, or swans on reservoirs with wild waterfowl and intermediate hosts that may contain avian schistosomes may leave these animals vulnerable to infection and disease. The aigamo method of rice farming relies on ducks to eat insects and weeds. This method was developed in 1989 by Takao Furuno, and is used in South Korea, China, Vietnam, the Philippines, Thailand, and Iran (Furuno 1996). In the Philippines, China, and Vietnam, duck pasturing has been implicated in paddy field dermatitis caused by Trichobilharzia paoi (Hu et al. 1994). WILDLIFE POPULATION IMPACTS Large die-offs of birds as the result of avian schistosomes are rare and most reports involve small mortality events or reports of disease in isolated individuals. The two largest documented die-offs involved 45 Ring-billed Gulls (Larus delawarensis) on the shore of the St. Lawrence River, Canada (Dallaire 2006) and 40 wild-caught Brant that were maintained in captivity on a freshwater pond (Wojcinski et al. 1987). An intense, mixed infection with Trichobilharzia and Dendritobil-
255
harzia was considered the primary cause of death in the Brant, while the schistosome infection in the Ringbilled Gulls was not identified to genus. TREATMENT AND CONTROL Control of avian schistosomes is difficult and depends on breaking cycles of transmission. This may require chemical, mechanical, and biological approaches to reduce or eliminate snail intermediate hosts (Hor´ak et al. 2002). Niclosamide has been widely used in mollusk control programs since the 1960s (World Health Organization 1965) and is still the molluscicide of choice (Perrett and Whitfield 1996). It is highly effective at all stages of the life cycle of snails (Webbe 1987) and does not adversely affect economically important crop plants, although certain algae and aquatic plants are damaged and fish mortality may occur at concentrations used to control snails (Andrews 1983). Other effective molluscicides include B-2 (sodium 2,5dichloro-4-bromophenol), copper sulfate, and sodium pentachlorophenate (Perrett and Whitfield 1996). Mechanical elimination of snail habitat has been successfully used to control populations of snails close to roosting habitats for some avian hosts and may be an effective way to control cercarial dermatitis (Leighton et al. 2000). Mechanical disturbance of epilithic habitat with a boat-mounted rototiller or tractor and rake successfully eliminated almost all snails when done in shallow areas of high snail concentration during the breeding and early development of the mollusks. Treatment of infected birds with praziquantel is also effective in interrupting transmission when used in baits for dabbling ducks (Blankespoor and Reimink 1988, 1991) or for treating dwarf Mallards and Mallards infected with Trichobilharzia (M¨uller et al. 1993; Reimink et al. 1995). This method has some drawbacks. When treated baits were used, it was necessary to capture the primary definitive host, Mergus merganser, a diving fish-eater, and inject each bird as part of an overall wildlife management scheme, because these hosts would not consume the bait (Blankespoor and Reimink 1988, 1991). Dosage may also affect efficacy of the methods. When dwarf Mallards and Mallards infected with Trichobilharzia were treated with praziquantel (M¨uller et al. 1993), only a threefold application of 200 mg/duck at 24-h intervals led to permanent reduction of detectable miracidia. Application of praziquantel in low doses (30 or 40 mg/duck) did not reduce the number of released miracidia. Medication with praziquantel led to a strong shift of adult worms located in the enteric veins of the ducks to the liver in as little as 3 h. During prepatency, doses of 22.5 mg praziquantel per duck per day, given for 1 week, were sufficient to completely stop the release of miracidia.
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In spite of these drawbacks, treatment of wild birds can be effective. Treatment of natural populations of Mallards infected with avian schistosomes in Michigan, USA, was an effective therapeutic agent for reducing natural infections of the parasite. One year after treatment, prevalence in Mallards was 1.8% versus 14.6% in an untreated control group (Reimink et al. 1995). MANAGEMENT IMPLICATIONS Management of avian schistosomiasis at an effective scale is difficult because of difficulties in delivering treatments and their high costs. Three basic strategies are possible: (1) prevention of the introduction of disease, (2) control of existing disease, and (3) eradication. Host–parasite interactions may differ by habitat type, host specificity may vary, and identification of species of schistosomes may be difficult, making eradication and control difficult. While snails that harbor the larval stages of avian schistosomes may be destroyed by molluscicides or mechanical treatments, this method is cost-effective only in small areas. Development of better detection methods for identification of schistosome cercariae in reservoirs could help target eradication methods for the parasite (Graczyk and Shiff 2000). LITERATURE CITED Andrews, P. 1983. The biology and toxicology of molluscicides, Bayluscide. Pharmacology and Therapeutics 19:245–295. Appleton, C. C. 1986. Occurrence of avian schistosomatidae (Trematoda) in South African birds as determined by fecal survey. South African Journal of Zoology 21:60–67. Bahgat, M., and A. Ruppel. 2002. Biochemical comparison of the serine protease (elastase) activities in cercarial secretions from Trichobilharzia ocellata and Schistosoma mansoni. Parasitology Research 88:495–500. Barber, K. E., and J. N. Caira. 1995. Investigation of the life cycle and adult morphology of the avian blood fluke Austrobilharzia variglandis (Trematoda: Schistosomatidae) from Connecticut. Journal of Parasitology 81:584–592. Basch, P. F. 1966. The life cycle of Trichobilharzia brevis n. sp. an avian schistosome from Malaya. Zeitschrift fur Parasitenkunde 27:242–251. Basch, P. F. 1991. Schistosomes: Development, Reproduction, and Host Relations. Oxford University Press, New York, 248 pp. Baugh, S. C. 1963. Contributions to our knowledge of digenetic trematodes VI. Zeitschrift fur Parasitenkunde 22:303–315.
Becker, D. A. 1956. The Morphology of Trichobilharzia alaskensis Harkema, McKeever and Becker, 1956 (Trematoda: Schistosomatidae; Bilharziellinae) with Notes on its Life History. M.S. Thesis North Carolina State College. Blair, D., and K. S. Islam. 1983. The life-cycle and morphology of Trichobilharzia austrais n. sp. (Digenea: Schistosomatidae) from the nasal blood vessels of the black duck (Anas superciliosa) in Australia, with a review of the genus Trichobilharzia. Systematic Parasitology 5:89–117. Blair, D., and P. Ottesen. 1979. Nasal schistosomiasis in Australian anatids. Journal of Parasitology 65:982–984. Blankespoor, H. D., and R. L. Reimink. 1988. Control of swimmer’s itch in Michigan: Past, present, future. The Michigan Riparian 10:19. Blankespoor, H. D., and R. L. Reimink. 1991. The control of swimmer’s itch in Michigan: Past, present, future. Michigan Academician 24:7–23. Bourns, T. K., and J. C. Ellis. 1975. Attempts to transfer immunity to Trichobilharzia oscellata (Trematoda: Schistosomatidae) passively via lymphoid cells and/or serum. Transactions of the Royal Society of Tropical Medicine and Hygiene 69:382–387. Bourns, T. K. R., J. C. Ellis, and M. E. Rau. 1973. Migration and development of Trichobilharzia ocellata (Trematoda: Schistosomatidae) in its duck hosts. Canadian Journal of Zoology 51: 1021–1030. Brant, S. V., and E. S. Loker. 2005. Can specialized pathogens colonize distantly related hosts? Schistosome evolution as a case study. PloS Pathogens 1(3):167–169. Brant, S. V., J. A. T. Morgan, G. M. Mkoji, S. D. Synder, R. P. V. J. Rajapakse, and E. S. Loker. 2006. An approach to revealing blood fluke life cycles, taxonomy, and diversity: Provision of key reference data including DNA sequence from single life cycle stages. Journal of Parasitology 92:77–88. Chanov´a, M., S. Vuong, and P. Hor´ak. 2007. Trichobilharzia szidati: The lung phase of migration within avian and mammalian hosts. Parasitology Research 100:1243–1247. Cheatum, E. L. 1940. Dendritobilharzia anatinarum n. sp., a blood fluke from the mallard. Journal of Parasitology 26:165–170. Chu, G. W. 1958. Pacific area distribution of fresh-water and marine cercarial dermatitis. Pacific Science 12:299–312. Cort, W. W. 1950. Studies on schistosome dermatitis. XI. Status of knowledge after more than twenty years. American Journal of Hygiene 52:251–307. Davis, N. E. 2006. Identification of an avian schistosome recovered from Aythya novaeseelandia and infectivity
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Schistosomes of its miracidia to Lymnaea tomentosa snails. Journal of Helminthology 80:225–233. DeGentile, L., H. Picot, P. Bourdeau, R. Bardet, A. Kerjan, M. Piriou, A. LeGuennic, C. Bayssade Dufour, D. Chabasse, and K. E. Mott. 1996. Cercarial dermatitis in Europe: A new public health problem? Bulletin of the World Health Organization 74:159–163. Dallaire, A. D. 2006. Schistosomiasis-related mortality in ring billed ducks. Wildlife Health Center Newsletter 12:9. Dvo´ak, J. 2001. Molecular analysis of European Trichobilharzia species. Helminthologia 4:243–250. Dvo´ak, J., M. Delcroix, and A. Rossi. 2005. Multiple cathepsin B isoforms in schistosomula of Trichobilharzia regenti: Identification, characterization and putative role in migration and nutrition. International Journal of Parasitology 35:895–910. ˇ Vanacova, V. Hampl, J. Flegr, and P. Hor´ak. Dvo´ak, J., S. 2002. Comparison of European Trichobilharzia species based on ITS1 and ITS2 sequences. Parasitology 124:307–313. Ellis, J. C., T. K. R. Bourns, and M. E. Rau. 1975. Migration, development and condition of Trichobilharzia ocellata (Trematoda: Schistosomatidae) in hom*ologous challenge infections. Canadian Journal of Zoology 53:1803–1811. Fain, A. 1955. Recherches sur les schistosomes d’oiseaux au Ruanda-Urundi (Congo belge). Revue de Zoologie et de Botanique Africaines 51:373–387. Fain, A. 1956a. Nasal trichobilharziasis: A new avian schistosomiasis. Nature 177:389. Fain, A., 1956b. Les schistsomes d’oiseaux du genre Trichobilharzia Skrjabin et Zakharov, 1920 au Ruanda-Urundi. Revue de Zoologie et de Botanique Africaines 4:147–178. Farley, J. 1963. A redescription of Gigantobilharzia lawayi Brackett, 1942. Journal of Parasitology 49:465–467. Farley, J. 1971. A review of the family Schistosomatidae: Excluding the genus Schistosoma from mammals. Journal of Helminthology 4:289–320. Farr, M. M., and V. G. Blankemeyer. 1956. Trichobilharzia brante n. sp. (Trematoda): Schistosomatidae from the Canada goose (Branta canadensis). Journal of Parasitology 42:321–325. Ferris, M., and W. J. Bacha, Jr. 1986. Response of leucocytes in chickens infected with the avian schistosome Austrobilharzia variglandis (Trematoda). Avian Diseases 30:683–686. File, S. 1995. Interaction of schistosome eggs with vascular endothelium. Journal of Parasitology 81:234–238.
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14 Cestodes J. Daniel McLaughlin 1983; Holmes 1994). Infected animals usually grow normally and exhibit few, if any, signs of disease (Arme et al. 1983). Similarly, infected birds seldom display clinical signs, and cestodes are not considered a problem unless present in massive numbers (Greve 1986) or in malnourished (Wobeser 1981) or otherwise debilitated hosts. For example, in waterfowl, perhaps the best-studied group of wild birds, only 16 of the 264 cestode species listed by McDonald (1969a) have been associated with disease or mortality. Nevertheless, a number of reports have associated cestodes with disease or mortality in wild or captive birds. Species of Gastrotaenia are known pathogens of waterfowl (Wobeser 1981), and a number of other cestodes cause lesions at the site of attachment or damage the intestinal mucosa. Fatal infections have been reported in Common Eiders (Somateria mollissima) (Grenquist et al. 1972; Kulachkova 1973, Persson et al. 1974; Hario et al. 1992), various species of swans (Cygnus spp.) (Jennings et al. 1961; Czaplinski 1965; Maksimova 1972; Papazahariadou et al. 1994), Arctic Loons (Gavia arctica) (Bayle 1983), Houbara Bustards (Chlamydotis undulata) (Jones et al. 1996a), and Willow Ptarmigan (Lagopus lagopus) (Holmstad et al. 2005). In some of these studies, however, the interpretation may have been confounded by the presence of other helminth species in the affected hosts. Although rare, larval cestodes have also been implicated as causes of avian mortality (Raethel 1977; Toplu et al. 2006). This chapter is not intended to be an exhaustive review of all reports of morbidity and mortality attributed to cestode infections in wild birds. Nor is it meant to document detailed pathological responses in all examples covered. These can be found in the works cited. Rather, the objective here is to summarize some of the general effects of cestode infection on wild or captive birds and to assess their significance in natural populations and their potential effects on domesticated hosts.
INTRODUCTION Cestodes or tapeworms (class Cestoda, phylum Platyhelminthes) are extremely common parasites of birds. Most species infect the intestine, but a few can be found in the ceca or under the gizzard lining. They are readily distinguished from other worm parasites (trematodes, nematodes, and acanthocephalans) by their segmented appearance. Birds have the most diverse cestode fauna of any vertebrate group. Over 1,700 of the approximately 4,000 nominal species listed by Schmidt (1986) infect birds, and that number continues to grow as new species are recognized and described (McLaughlin 2003). Wild birds are often infected with large numbers of cestodes and average prevalence can be quite high. As reported in 16 studies from North America and Eurasia, average prevalence ranged from 18 to 69% in samples of up to 3,089 birds from 232 avian species. In each study, prevalence of cestode infection exceeded that of any other helminth group (Rausch 1983). Depending on host species, apparently healthy birds may be infected with tens, hundreds, or, in some cases, thousands of cestodes (Cornwell and Cowan 1963; Bush and Holmes 1986; Stock and Holmes 1987; Bush 1990). The literature on avian cestodes is replete with studies of the distribution, systematics, and life histories of these parasites, but few address other aspects of host– parasite relationships or disease. Many cestodes are large enough to detect without magnification, and because they are so common, they are often observed in sick or dying birds (e.g., Kinsella and Forrester 1999). It is likely that earlier authors may have erroneously implicated cestodes as causes of disease or mortality when no other agents were evident (Rausch 1983). Further, wild birds may be infected with several species of helminths, making it difficult to ascribe effects to a particular parasite (Rausch 1983; Chapter 1). There is little evidence to suggest that adult cestodes have an adverse effect on animals or birds (Rausch
261 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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SYNONYMS Cestodiasis, hymenolepididiasis, drepanidotaeniasis, fimbriariasis, gastrotaeniasis. HISTORY Early references to mortality and disease in birds due to cestodes can be found in Sprehn (1932). Most of these studies refer to infections in domesticated species. Virtually all the existing information on wild birds is based on incidental observations made during studies that had other goals. Experimental studies on avian cestodes are rare and most data come from studies of domesticated species. Even here, few controlled studies exist (Reid 1983). Few species that cause visible pathology in wild birds have been studied either by experimental methods or by detailed studies of naturally infected birds. Some of the best examples include Gastrotaenia dogieli and Gastrotaenia cygni, two species that live under the gizzard lining of waterfowl (Wolffhugel 1938; Heck 1969; Egizbaeva and Erbolatov 1975; Egizbaeva and Basyvekova 1978; Kulukbaeva 1985), and Schistotaenia tenuicirrus, a species that causes intestinal diverticulae in Pied-billed Grebes (Podilymbus podiceps) and Horned Grebes (Podiceps auritus) (Boertje 1974). ETIOLOGY Cestodes belong to the phylum Platyhelminthes. Most species infect the intestine, a few species infect the ceca, and Gastrotaenia infects the gizzard. Occasionally, cestodes invade abnormal sites including the ureters (Wobeser 1974) and the gizzard muscle (McOrist 1989; Mondal and Baki 1989). Adult cestodes are white and translucent when alive. They range from 1–2 mm to 1 m long (Rausch 1983), but many are less than 10 cm. Cestode bodies consist of
a holdfast (scolex), a short neck, and a body (strobila) made up of repeated units (proglottids) that give it a segmented appearance. A mature strobila consists of three zones: a zone of immature proglottids posterior to the scolex, a zone of sexually mature proglottids with functional reproductive systems, and a postreproductive (gravid) zone consisting of proglottids that contain eggs ready for dispersal from the host. Three of the 14 orders recognized by Jones et al. (1994) are represented in the cestode fauna of birds. About 70 species belong to the orders Tetrabothriidea and Pseudophyllidea and the remainder belong to the Cyclophyllidea (Schmidt 1986). Most cyclophyllidean species that infect birds are found in the families Hymenolepididae, Dilepididae, and Davaineidae. Molecular evidence indicates that the Cyclophyllidea is the most highly derived order. The Tetrabothriidea is its closest relative and the Pseudophyllidea occupies a more basal position. The Pseudophyllidea (sensu Bray et al. 1994) is polyphyletic with only one family that is found in birds, the Diphyllobothriidae. Recent evidence indicates that the Diphyllobothriidae is independent and ancestral to the Pseudophyllidea (Olsen and Tkach 2005), but formal classification of the cestodes has yet to reflect molecular results. The family Diphyllobothriidae will be treated as though it has ordinal status in this chapter. These three groups can be distinguished by morphology of the scolex and mature proglottids (Table 14.1). Cyclophyllidean scolices have four muscular suckers. Most species also have a rostellum (a muscular organ within the scolex) that can be projected from its apex. The rostellum is usually armed with hooks and the number, shape, and size of these are of taxonomic importance. Tetrabothriidean scolices have four large leaflike suckers called bothridia and lack a rostellum. Scolices of the diphyllobothriids have one dorsal and one ventral groove (bothria) instead of suckers or bothridia.
Table 14.1. Morphological comparison of scolices and mature proglottids of adult diphyllobothriid, tetrabothriid, and cyclophyllidean cestodes. Characteristic
Diphyllobothriidae
Tetrabothriidea
Scolex with: Rostellum Genital pore (position)
2 Bothria None Ventral midline
4 Bothridia None Lateral margin
Uterine pore Uterine pore (position) Uterus (structure) Vitelline gland Vitelline gland (position)
Present Ventral midline Tubular, coiled Follicular Follicles visible throughout proglottid
Present Dorsal midline Saccular Compact Preovarian
Cyclophyllidea 4 Acetabula Present/absent Lateral margin (ventral in Mesocestoididae) Absent None Saccular Compact Usually postovarian
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Cestodes Cestodes have complex reproductive systems. Mature proglottids of these three groups can be distinguished by the type and location of the vitelline (yolk) glands, the structure of the uterus, the position of the genital pore, and by the presence or absence of a uterine pore (Table 14.1). Larval infections are less common. Larval cestodes are also white and range from several millimeters to several centimeters long. Larval Mesocestoides (Cyclophyllidea) occur mainly in the body cavity but can be found in all internal organs of heavily infected birds (Kugi 1983). Larval diphyllobothriids occur in the body cavity (Raethel 1977; Kuntz 1979) and in muscle (Kuntz 1979). HOST RANGE Every parasite has limits on the range of species it can infect. An infection is possible only if a series of environmental (contact) and physiological (compatibility) criteria are met (Combes 2001). This is most likely to occur in closely related species or in species that share the same diet and habitat. Most cestodes tend to occur in a single order of birds (Fuhrmann 1932); however, a given cestode may infect multiple host species, genera, or families within an order and may also infect birds of different orders (Table 14.2). Others are restricted to only a few host species. Fimbriaria fasciolaris, for example, infects over 60 species (7 families, 30 genera) of waterfowl (Anseriformes) worldwide (McDonald 1969a). By contrast, other species of cestodes infect less diverse orders and are found in fewer species. For example, species of Schistotaenia are specific to grebes (Rausch 1983; Stock and Holmes 1987), while species of Parorchites infect penguins (Cielecka et al. 1992). In each case, hosts are related phylogenetically, share habitats, and feed on the intermediate hosts to varying degrees. Exchange of cestode species between related hosts is common, particularly in wetland habitats. Here, the presence of multiple host species, limited foraging space, and similar diets ensure contact between pools of infective larval stages established by each host species (Nerassen and Holmes 1975; Stock and Holmes 1987). When this happens, natural selection will likely favor host switching if parasites can successfully reproduce, eventually selecting for adaptations in the life cycle that enhance continued contact with these new hosts. Many cestodes have been reported from multiple host orders (see Rausch 1983 and Table 14.2). However, such records by themselves can be misleading. For example, 26 cyclophyllidean species that normally infect orders of birds other than Anseriformes have been reported 37 times in waterfowl. Of these, 1 species
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has been reported 5 times, 2 species 3 times, 3 species 2 times, and 20 species once in over 100 surveys of wild or domestic waterfowl (McDonald 1969a, b). Fifteen of these species normally infect other aquatic birds: Charadriiformes (10 spp.), Podicipediformes (3 spp.), and Pelecaniformes (2 spp.). Thirteen of these species have been reported once in waterfowl. The normal hosts of these cestodes are aquatic birds that can share habitat, foraging areas, and at least some prey items with ducks. The other 11 species (e.g., species of Raillietina and Amoebotaenia) normally infect Galliformes (10 spp.) or Passeriformes (1 sp.) and were reported in domestic ducks that were apparently raised in proximity with chickens. Other examples of natural infections in phylogenetically different hosts exist. Rausch (1983) reported a species of Schistotaenia that is normally found in grebes in a crow (Corvus sp.). He also suggested that ecological factors rather than physiological factors or phylogenetic relationships were more important determinants of successful infections. GEOGRAPHIC DISTRIBUTION Geographic distribution of cestodes can be considered at different spatial scales that are ultimately dependent on the overlap of both avian and intermediate hosts and successful transmission of the parasites. On a global scale, the cestode fauna of birds has been well documented throughout the Holarctic. However, studies in the southern hemisphere have been less intensive (Rausch 1983), and the cestode fauna is less well known. Many of the species listed in Table 14.2 have cosmopolitan or Holarctic distributions. Others, like Parorchites in penguins, have more restricted distributions that reflect the distribution of suitable hosts (Cielecka et al. 1992). At a more local scale, cestode species may be present in some areas and absent in others. This is common in migratory birds where cestode species may be acquired on the breeding grounds and then transported to wintering areas. Some cestode species may persist while others may disappear if parasite life spans are short and local transmission is not possible. This can lead to seasonal declines in cestode diversity in migratory birds particularly on the wintering grounds (Buscher 1965; Hood and Welch 1980; Wallace and Pence 1986). Alternatively, migrant birds may acquire new species, particularly if they winter in coastal areas. Transmission of cestodes with aquatic life cycles tends to be restricted to either marine or freshwater environments, possibly reflecting osmotic effects on egg or oncosphere survival. Species of Tetrabothrius, Ophryocotyle, and Kowalewskiella are transmitted in marine environments (Stock and Holmes 1987; Bush 1990).
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Table 14.2. A partial list of cestode species that have been reported to cause pathology, disease, or mortality in wild birds. Parasite
Distribution
Host order
Source
Reference
ADULT CESTODES Order Cyclophyllidea Family Hymenolepididae Gastrotaenia cygni
NA, SA
A
Gastrotaenia dogieli
Eu, As
A
Fimbriaria fasciolaris
C
A*, Ch, Po, Gal, Gr, F, Pe, Pi A*, Ca, Ci, Gal, Pa A A*, Ch, Gal, Gr
E C, D?
Heck (1969) Willers and Olsen (1969) Egizbaeva and Erbolatov (1975) Kulukbaeva (1985) Basu et al. (1982)
C, D? C, D? ? C, D?
Gitter et al. (1974) Basu et al. (1982) ˇ Slais (1961) Basu et al. (1982)
Microsomacanthus collaris Microsomacanthus parvula Dicranotaenia coronula Sobolevicanthus gracilis Aploparaksis furcigera Aploparaksis penetrans Cloacotaenia megalops Hispaniolepis falcata Family Davaineidae Otiditaenia conoideis Otiditaenia macqueeni Raillietina sp. Family Dilepididae Choanotaenia infundibulum Parorchites zederi Family Gryporhynchidae† Paradilepis delachauxi Paradilepis scolecina Paradilepis sp. Family Amabiliidae Schistotaenia scolopendra Schistotaenia srivastavi Schistotaenia tenuicirrus Family Paruterinidae Ascometra choriotidis Metroliasthes lucida Cyclophyllidean sp. Order Tetrabothriidea Tetrabothrius skoogi Tetrabothrius sp.
W W E
Eu, As, NA Eu, As, Af, NA Eu, As, Af, NA Eu, As, NA Eu, As, NA C Af, As
A*, Gal, Co
C, D?
Basu et al. (1982)
A*, Gr, Po Ch A*, Gal, Gr Gr
? W W W
ˇ Slais (1961) Spasskaya (1966, Figure 65) Wobeser (1974) Jones et al. (1996b)
Af, Eu, As Af, As NA
Gr Gr Gal
C C W
Jones et al. (1996a) Jones et al. (1996a) Thomas (1985)
C Antarctica
A, Co, F, Gal*, C, D? Gr, Pa, St W Pr W
Basu et al. (1982) McOrist (1989) Fuhrmann (1921)
Af, As C As
Pe Pe Pe
W W W
Baer (1959) Karstad et al. (1982) Matta and Ahluwalia (1977)
SA NA NA
Po Po Po
W W W, E
Baer (1940) Rausch (1970) Boertje (1974)
As NA
Gr Gal
C W
NA via Af
Ph
C
Jones et al. (1996a) ´ Angeles Rebolloso et al. (2006) Poynton et al. (2000)
As Au
Pr Pr
W W
Nishigai et al. (1981) Obendorf and McColl (1980) (continues)
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Cestodes Table 14.2. (Continued ) Parasite Family Diphyllobothriidae Schistocephalus solidus
Ligula intestinalis
Distribution C
C
Host order
Source
A, Ch*, Ci, Co, W F, Gal, Gav, Gr, Pa, Pe, Po, Pr W W A, Ci, Ch*, F, W Pe*, Po*, W Gav*, Pa
Reference Grenquist et al. (1972)
Persson et al. (1974) Hario et al. (1992) Bayle (1983) Betke et al. (2003)
LARVAL CESTODES Family Mesocestoididae Mesocestoides sp.
C
Mammals
W W W
Toplu et al. (2006) Mill´an et al. (2003) Kugi (1983)
Family Diphyllobothriidae Ligula intestinalis Spirometra sp.
C C
See above Mammals
C W
Raethel (1977) Kuntz (1979)
Note: Classification follows Khalil et al. (1994). Distribution refers to the known geographic distribution as presented in McDonald (1969a, b) or Schmidt (1986). Host orders for adult cestodes are based on data in Fuhrmann (1932), McDonald (1969a, b), and Schmidt (1986). Asterisk (*) indicates the major order(s) of host(s) for a specific parasite. Host orders: A, Anseriformes; Ca, Caprimulgiformes; Ch, Charadriiformes; Ci, Ciconiiformes; Co, Columbiformes; F, Falconiformes; Gal, Galliformes; Gav, Gaviiformes; Gr, Gruiformes; Pa, Passeriformes; Pe, Pelecaniformes; Ph, Phoenicopteriformes; Pi, Piciformes; Po, Podicipediformes; Pr, Procellariiformes; Str, Strigiformes;. Asterisks when present indicate the major host order(s) of a particular species. Source of material: W, wild; E, experimental; C, captive; D, domestic; ?, source unknown. Distribution: NA, North America; SA, South America; Eu, Europe; As, Asia; C, Cosmopolitan; Af, Africa. † The family Gryporhynchidae is not included in Khalil et al. (1994).
They infect pelagic and coastal birds but migrant species from freshwater habitats that winter in coastal areas may also become infected. Species of these three genera have been found in birds on breeding areas on the Canadian prairies (Stock and Holmes 1987; Bush 1990), thousands of kilometers from where they were acquired.
EPIZOOTIOLOGY Life Cycle Cestode life cycles are indirect and each stage must be eaten by the next host for transmission to occur. One or two intermediate hosts may be required to complete the life cycle. Life cycles are similar for species within each family of cestodes; however, their intermediate hosts may differ. Except for the Tetrabothriidea, scolices of the infective larval stages are identical to those of the adult worm.
Cyclophyllidea Cyclophyllidean eggs are infective when passed from the host. Each egg consists of a larva (oncosphere) that is armed with three pairs of hooks and surrounded by one or two delicate membranes. In suitable conditions, the eggs of many species can survive for several months at low temperatures and some can survive short periods of freezing (Lee et al. 1992). With the exception of the Gryporhynchidae and Mesocestoididae, life cycles of most cyclophyllidean families require one intermediate host. In most families, the intermediate hosts are invertebrates. Crustaceans, insect larvae, and annelids serve as intermediate hosts of species that infect aquatic birds. Insects, annelids, and mollusks serve as intermediate hosts of species that infect terrestrial birds. Cladotaenia and Paruterina, which infect raptors, use rodents as intermediate hosts (Rausch 1983). Among cyclophyllideans, the oncosphere is released from the egg following ingestion and penetrates the gut
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Figure 14.1. Representative life cycles of cyclophyllidean and diphyllobothriid cestodes. Larval stages of the Gryporhynchidae redrawn after Chervy (2002).
of the intermediate host. It localizes in the hemocoel, coelom, or digestive gland where it develops into a cysticercoid larva that is infective to the avian host (Figure 14.1). Species of the family Gryporhynchidae infect pelicans, cormorants, and other piscivorous birds. Copepods and fresh or brackish water fish are required for transmission (Figure 14.1). The oncosphere develops into a procercoid larva in the hemocoel of the copepod host. When eaten by a fish, the larva migrates to the mesenteries, liver, or gall bladder where it develops into the second larval stage, the merocercoid, which is infective to the avian host (Chervy 2002). The life cycle of the Mesocestoididae is believed to require two intermediate hosts. The first intermediate host is unknown but is thought to be an arthropod. The stage that develops in what is believed to be the second intermediate host is now considered a merocerocoid (Chervy 2002), but the more familiar term tetrathyridium will be used here. This stage is infective to the definitive host. Second intermediate hosts
may be amphibians, reptiles, birds, and mammals. The final hosts are carnivorous mammals or, rarely, birds (Rausch 1994). Under optimal conditions, developmental times of cyclophyllidean cysticercoids range from 6 days to 4 weeks (McDonald 1969a; Reid 1983). Some hymenolepidid species mature to adults in 4 days (Podesta and Holmes 1970), although 10–14 days appears to be the norm (McDonald 1969a). Dilepidid, davaneiid, and gryporhynchid species may require up to 3 weeks (McDonald 1969a; Reid 1983; Scholz et al. 2004). The adult life spans of species infecting wild birds are unknown.
Tetrabothriidea Complete life cycles of this family are unknown; however, larval stages occur in marine crustaceans, teleosts, and cephalopods. Unlike other cestodes, the scolex undergoes further development within the final host (Hoberg 1994).
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Diphyllobothriidae Two intermediate hosts are required to complete Diphyllobothriid life cycles (Figure 14.1). The first host is a copepod and the second is a vertebrate, usually a fish. The exception is Spirometra whose species use all vertebrates except fish as second intermediate hosts (Bray et al. 1994). The eggs are thick shelled, operculate, and require a period of development in water before hatching. The egg eventually releases a coracidium larva, which is essentially an oncosphere surrounded by a ciliated covering. The oncosphere penetrates the gut of the copepod host and develops into a procercoid larva in the hemocoel. When eaten by a fish, the procercoid penetrates the host gut, resumes development in the body cavity, visceral organs or musculature, and transforms into the pleurocercoid stage. This stage is infective to the avian host. Developmental times for diphyllobothriids are 4– 12 days for coracidia, 7–15 days for procercoids, and several months for pleurocercoids. Pleurocercoids of Schistocephalus require 4–6 months while those of Ligula and Diphyllobothrium require 10–14 months. Adults mature rapidly in the avian host and survive for 2–12 days (see McDonald 1969a).
1981) and Little Penguins (Eudyptula minor) (Obendorf and McColl 1980) infected with Tetrabothrius spp., and in an Arctic Loon infected with Ligula intestinalis (Bayle 1983). In the latter case, the loon also displayed generalized weakness and diarrhea. A few species of cestodes in ducks and grebes can cause varying degrees of hemorrhage where they attach to the intestinal mucosa that might be detected as bloody feˇ ces (Slais 1961; Heck 1969; Boertje 1974). Unusual changes in feeding behavior have also been reported in Common Eiders infected with S. solidus. Eiders normally dive for food in deep water, but heavily infected individuals fed in shallow water by tipping up like dabbling ducks until they died (Hario et al. 1992).
CLINICAL SIGNS Cestode infections in wild birds are normally asymptomatic, but when clinical signs are present, they are nonspecific and similar to those reported in poultry (Reid 1983). Emaciation, weakness, and occasionally diarrhea and hemorrhage may be accompanied by changes in posture or locomotory and feeding behavior. Interpretation of clinical signs may be confounded by the common occurrence of mixed helminth and protozoan infections in wild birds. Experimental studies have helped to identify signs that are associated with cestode infection. Weakness and inappetence have been documented in ducklings within 6–8 days after experimental infection with G. dogieli. These signs became worse with time, leading to anorexia and spasmodic head and limb movements by day 21 postinfection (PI) and death by day 30 PI (Kulukbaeva 1985; Egizbaeva and Kulukbaeva 1985). Similar signs have been reported in domestic ducklings infected with Microsomacanthus collaris, including difficulty in walking, an abnormal backward arching of the neck, and unusual huddling behavior (Gitter et al. 1974). Among naturally infected wild birds, emaciation has been observed in common eiders infected with Schistocephalus solidus and Lateriporus sp. (Hario et al. 1992), in a Long-tailed Duck (Clangula hyemalis) infected with G. cygni (Heck 1969), in Short-tailed Shearwaters (Puffinus tenuirostris) (Nishigai et al.
Gizzard Infections: GASTROTAENIA Species of Gastrotaenia normally live under the softer areas of the gizzard lining in waterfowl but are also found under the grinding plates. The lesions appear as roughened, friable, eroded areas on the lining and are usually discolored. Ecchymoses and necrosis are usually present around the edges of the grinding plates (Heck 1969; Egizbaeva and Erbolatov 1975). The gizzard muscles are weakened, portions of the lining may detach from the underlying tissue, and necrosis of the glandular layer occurs (Willers and Olsen 1969; Kulukbaeva 1985). The area occupied by the cestodes is depressed, inflamed, and may show signs of hemorrhage (Heck 1969; Willers and Olsen 1969). Embedded worms cause extensive necrosis of the lining that is accompanied by atrophy of the underlying glandular area and altered mucous secretion (Egizbaeva and Erbolatov 1975). Areas surrounding the cestodes and lesions become inflamed and are infiltrated by polymorphonuclear leukocytes, lymphocytes, and eosinophils that may be accompanied by localized hemorrhage (Heck 1969; Willers and Olsen 1969; Egizbaeva and Erbolatov 1975; Kulukbaeva 1985).
PATHOLOGY OF ADULT CESTODES Adult cestodes may cause damage to the gizzard lining (Gastrotaenia), intestinal blockage, localized damage to the intestinal wall at the site of attachment, or irritation of the intestinal lining. Inflammation is the most common host response to cestode infection and appears to be most intense where prolonged contact occurs between the host and parasite. Cyclophyllidea
Intestinal Infections Potentially fatal intestinal occlusion has been reported in waterfowl infected with various hymenolepidid
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species (Czaplinski 1965; Maksimova 1972; Kulachkova 1973; Gitter et al. 1974; Basu et al. 1982), in Houbara Bustards infected with Ascometra choriotidis (Jones et al. 1996a), and in Wild Turkeys (Meleagris gallopavo) infected with Metroliasthes ´ lucida (Angeles Rebolloso et al. 2006). A number of hymenolepidid species and Choanotaenia infundibulum produce enteritis in waterfowl (Basu et al. 1982; Schmidt et al. 1987) that may be accompanied by a distention of the intestine and an accumulation of hemorrhagic and/or mucous exudates (Basu et al. 1982). Thomas (1985) provided evidence that intestinal hypertrophy in Willow Ptarmigan infected with Raillietina sp. is positively correlated with the number of cestodes present. Various hymenolepidid species from ducks, S. tenuicirrus from grebes, and species of Otiditaenia from bustards cause inflammaˇ tion in the intestinal lining (Slais 1961; Kulachkova 1973; Boertje 1974; Basu et al. 1982; Jones et al. 1996a). Effects of scolices. Scolices of several cyclophyllidean species penetrate deeply into the intestinal wall. Parorchites zederi and S. tenuicirrus produce large diverticulae in the intestine of penguins and grebes, respectively, that are visible on the serosal surface of the organ (Fuhrmann 1921; Boertje 1974; Cielecka et al. 1992). A similar condition caused by an unidentified cyclophyllidean has been reported from a Lesser Flamingo (Phoenicopterus minor) (Poynton et al. 2000). In each case, the diverticulum contained the scolex and a portion of the strobila. Diverticulae produced by S. tenuicirrus are the most complex and consist of four large vesicles averaging 15 mm in diameter (Boertje 1974). Scolices of Schistotaenia scolopendra, Schistotaenia srivastavi, and three species of Paradilepis also penetrate deeply into the intestinal wall of various grebes and cormorants (Baer 1940, 1959; Rausch 1970; Matta and Ahluwalia 1977; Karstad et al. 1982). Species of Paradilepis, including Paradilepis scolecina, produce nodules (1–2 mm) that are visible on the serosal surface of the intestine (Figure 14.2) (Matta and Ahluwalia 1977; Karstad et al. 1982). The large scolices of Schistotaenia and Paradilepis cause extensive damage to the mucosal, submucosal, and muscular layers (Baer 1940; Rausch 1970; Boertje 1974; Matta and Ahluwalia 1977; Karstad et al. 1982) (Figure 14.3). The scolex may be surrounded by a thick fibrous capsule which, in the case of P. scolecina, consists of multinucleate giant cells and fibrocytes (Karstad et al. 1982). Leukocyte infiltration, inflammation, and hemorrhage have been reported at the attachment sites of Schistotaenia spp. (Rausch 1970; Boertje 1974).
Figure 14.2. Nodules on the intestine of a Great Cormorant (Phalacrocorax carbo) infected with Paradilepis scolecina. Reproduced from Karstad et al. (1982), with permission of the Journal of Wildlife Diseases. Scolices of Aploparaksis penetrans have been reported to produce small nodules in various species of Charadriiformes (Spasskaya 1966). In this case, however, it is the tip of the rostellum rather than the scolex that produces the nodule. Smaller scolices produce less damage. Jones et al. (1996b) reported denudation of mucosal epithelium at the attachment site of Hispaniolepis falcata in Houbara Bustards and a hypergenerative response adjacent to it but found little response associated with attachment sites of other species. Scolices of C. infundibulum and three species of hymenolepidids produced necrotic foci in domestic ducks (Basu et al. 1982), whereas those of Aploparaksis furcigera and Microsomacanthus parvula in ducks produced a local inflammation dominated by eosinophils
Figure 14.3. Section through an intestinal nodule infected with Paradilepis scolecina. Reproduced from Karstad et al. (1982), with permission of the Journal of Wildlife Diseases.
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Cestodes ˇ and plasma cells (Slais 1961). Varying degrees of hemorrhage have been reported in ducks infected with ˇ A. furcigera and M. parvula (Slais 1961). Effects of strobilae. The strobila represents virtually all the biomass of a cestode and much of it is in contact with the mucosal surface at any given time. Strobilae do not penetrate the mucosa or cause open lesions, but continuous contact with the mucosa causes irritation that may be exacerbated by the extension and contraction of the worm during normal activity. The fragmentary information available on host responses to strobilae in wild birds is similar to what has been reported from poultry (Reid 1983; Padhi et al. 1986). Physical changes include desquamation, necrosis, and shortening of villi in ducks infected with C. infundibulum and various hymenolepidids (Basu et al. 1982; Kishore and Sinha 1989). Structural changes in Kori Bustards (Ardeotis kori) infected with Otiditaenia conoideis range from mild atrophy to collapse and fibrosis of the intestinal musosa, and unspecified damage to the mucosa has been described in Houbara Bustards infected with O. conoideis and Hymenolepis falsata (Jones et al. 1996a). Inflammation of the intestinal mucosa is common in cestode infections in ducks (Basu et al. 1982; Schmidt et al. 1987) and in bustards (Jones et al. 1996a). In general, intestines of bustards infected with cestodes are inflamed but the degree of inflammation varies among host species. Infiltration of monocytes, lymphocytes, eosinophils, heterophils, and plasma cells occurs to varying degrees. This is accompanied in some cases by the proliferation of connective tissue and enlargement of lymph nodules (Basu et al. 1982; Kishore and Sinha 1989; Jones et al. 1996a). In Red-crested Bustards (Eupodotis ruficrista) infected with Otiditaenia macqueeni, small inflammatory nodules consisting of plasma cells, lymphoid cells, and clumps of hemosiderin-laden macrophages are present on the mucosa (Jones et al. 1996a). Infections in abnormal sites. Cestodes rarely invade abnormal sites but when they do, they can produce irritation or atypical lesions (Wobeser 1974; McOrist 1989). Examples include abnormal development of Cloacotaenia megalops, a cloacal cestode of waterfowl, in the ureters of ducks (Wobeser 1974), abnormal development of C. infundibulum, an intestinal parasite of species of Galliformes and Passeriformes, in the gizzard lining of Barn Owls (Tyto alba) (McOrist 1989), and development of immature F. fasciolaris in the gizzard muscles of a domestic duck (Mondal and Baki 1989). Lesions in ducks infected with C. megalops include enlargement and inflammation of the ureters,
269
flattening and atrophy of the epithelial lining, and a diffuse infiltration of heterophils in the walls of the ureters and in the kidneys (Wobeser 1974). Barn Owls infected with C. infundibulum develop conspicuous lesions in the gizzard lining that have attached cestodes and dark, bloody material (McOrist 1989). Tetrabothriidea Little is known about host responses to infection with adult tetrabothriid cestodes. Nishigai et al. (1981) reported large numbers of Tetrabothrius skoogi in emaciated Short-tailed Shearwaters that died of apparent malnutrition and anemia off the coast of Japan, but gross and microscopic lesions were not reported. Diphyllobothriidae Intestinal distention and occlusion have been reported in very intense infections with diphyllobothriid cestodes. As many as 340 adult S. solidus were recovered from a duck with a fatal infection (Callot and Desportes 1934). Both intestinal distention and occlusion were present in Common Eiders that died with intense infections of this parasite (Grenquist et al. 1972; Persson et al. 1974). Death of a Marabou Stork (Leptoptilos crumeniferus) chick resulted from an intestinal infection of L. intestinalis (Betke et al. 2003). Pathology of Larval Cestodes Cyclophyllidea Larval cestodes develop in the body cavity, internal organs, or musculature. Transitory damage occurs during larval migration from the gut and chronic lesions may develop in tissues where larvae become established (Kugi 1983; Roberts and Janovy 2005; Toplu et al. 2006). Tetrathyridia (Mesocestoides) usually occur in the body cavity (Kugi 1983; Mill´an et al. 2003), but may also infect visceral organs when present in large numbers (Kugi 1983). Generally, there are no visible reactions; however, Toplu et al. (2006) reported nonsuppurative granulomatus pleuritis and peritonitis and a yellow, serous fluid containing tetrathyridia in the abdominal and thoracic cavities of a captive peafowl. Granulomas containing degenerating tetrathyridia were also present on the parietal and pleural peritoneum. These were surrounded by macrophages, lymphocytes, and eosinophils. Additional granulomas containing tetrathyridia were present in the muscles of the proventriculus. The tetrathyridia of Mesocestoides have been reported in livers of Green Pheasants (Phasianus
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versicolor) (Kugi 1983). Livers were congested and a slight infiltration of lymphocytes was present in the lungs. No changes were evident in hepatic cells despite the presence of parasites in the liver parenchyma. Similarly, no apparent lesions were observed in Red-legged Partridges (Alectoris rufa) infected with Mesocestoides (Mill´an et al. 2003).
Diphyllobothriidae Pleurocercoid larvae have been found in the body cavities of a variety of birds (Raethel 1977; Kuntz 1979; Lafuente et al. 1999). Raethel (1977) described fatalities in captive Pink-backed Pelicans (Pelecanus rufescens), a Brown Pelican (Pelecanus occidentalis), a Double-crested Cormorant (Phalacrocorax auritus), a Guanay Cormorant (Phalacrocorax bougainvillii), and a Wood Duck (Aix sponsa) infected with migrating pleurocercoids of L. intestinalis. Partial penetration of the intestine was seen in some individuals and a pleurocercoid had penetrated the abdominal wall of one bird. A putrescent fibrinous serositis was present and what appeared to be intestinal contents were found in the body cavity. Fibrous deposits were present on the serosa and in most birds fibrous lesions had fused several intestinal loops together. Birds are normally the definitive hosts of L. intestinalis, and the presence of Ligula pleurocercoids in the body cavity is unusual. Larvae found by Lafuente et al. (1999) were not identified; those reported by Kuntz (1979) were believed to be species of Spirometra, which are known to use birds as second intermediate hosts. No pathology was associated with infections of Spirometra pleurocercoids in the body cavity of various birds; however, several pleurocercoids were found embedded in subcutaneous connective tissue of the breast muscles (Kuntz 1979), indicating tissue migration by some of the pleurocercoids in these birds. DIAGNOSIS The presence of eggs, gravid proglottids, or cestode fragments in the feces of the host is diagnostic for cestode infection. Although it is not possible to identify cestodes to species in this manner, cyclophyllidean and tetrabothriidean cestodes can be distinguished from diphyllobothriid cestodes on the basis of egg morphology (Figure 14.1) and proglottid structure (Table 14.1). Diphyllobothriid eggs have hard operculate shells (Figure 14.1), but may be difficult to distinguish from eggs produced by trematodes. The gravid proglottids of some families are sufficiently distinct to permit identification to that level. Identification of cestodes to order, family, and generic levels requires microscopic study of adult spec-
imens and evaluation of the morphology of the scolex and reproductive systems (Table 14.1). Identification to species level requires evaluation of the presence or absence of the rostellar hooks on the scolex and their number, shape, and size. The position, size, and shape of components of the male and female reproductive systems are also important, including the number of testes and their spatial relationships within the mature proglottid. Keys to the generic level are available in Schmidt (1986) and Khalil et al. (1994). Schmidt included species lists for each genus. Unfortunately, few species keys are available. Existing keys, taxonomic revisions, and descriptions of new species can be located through Helminthological Abstracts or similar abstracting services. IMMUNITY There is little evidence to suggest that birds develop immunity to cestode infections. Chickens with existing infections of Raillietina laticanalis are not immune to reinfections (Ueta and Avancini 1994). No comparable data exist for wild hosts, although parasitological surveys and studies of cestode life histories indicate that repeated infections occur. Juvenile and adult birds are typically infected by the same species, suggesting that individuals infected as juveniles are reinfected as adults. Many cestode species found in waterfowl are transmitted by copepods and ostracods (McDonald 1969a) that can support only one or two larvae because of their small size. Birds with cestode populations in excess of this would have to ingest multiple intermediate hosts, most likely at different times. It is not uncommon to find mature and immature specimens of the same species in a wild bird, which indicates that cestode recruitment is a continuous process, at least at certain times of the year. Finally, aquatic birds in particular are normally infected by multiple species of cestodes, some of which are transmitted by different species of intermediate hosts (Bush and Holmes 1986; Stock and Holmes 1987; Bush 1990). Acquisition of different components of the cestode community likely occurs over a period of time, indicating that prior infections provide little or no immunity to superinfection with the same or different species. Collectively, these observations argue against the presence of an effective immune response to cestode infection in birds. PUBLIC HEALTH CONCERNS Adult cestodes found in birds cannot be transmitted directly to humans and do not pose a health threat. However, the larval stages of Mesocestoides and Spirometra can infect humans when consumed in raw or undercooked meat (Beaver and Jung 1985; Roberts
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Cestodes and Janovy 2005). The few case reports found in a search of the Helmintological Abstracts database from 1990 to the present, however, suggest that human infections with either parasite are relatively rare. Ingestion of larval Mesocestoides can result in the establishment of adult cestodes in the intestine (Beaver and Jung 1985). Ingestion of Spirometra pleurocercoids results in their transfer, without further development, to a new host individual. Infections with this type of pleurocercoid are known as sparganosis (Roberts and Janovy 2005). In sparganosis, the parasite may localize in the body cavity among the viscera or it may migrate to organs or muscles. It frequently appears as a lump under the skin that is usually treated surgically. A more serious situation may occur if the larva invades an internal organ and proliferates, in which case it can cause extensive damage (Roberts and Janovy 2005). DOMESTIC ANIMAL HEALTH CONCERNS Transmission of cestodes from wild to domestic birds requires a wild reservoir host to provide a source of cestode eggs and intermediate hosts to support development of larval stages of the parasites. The practice of raising ducks and geese on reservoirs and natural wetlands frequented by wild waterfowl can lead to outbreaks of cestode infection, disease, and losses in domestic waterfowl (Gitter et al. 1974). Losses of domestic waterfowl as a result of infection with G. dogieli have been documented in Eastern Europe and the former USSR (Egizbaeva and Erbolatov 1975). Similarly, contamination of local ponds by wild anatids can also lead to outbreaks in captive waterfowl in zoological collections (Kotecki 1970). Cestode diversity in captive birds at the Warsaw Zoo was less than that in wild birds, presumably because a suitable range of intermediate hosts was not present. Cestode larvae develop rapidly and may be infectious to captive or domestic waterfowl in fewer than 2 weeks after a wetland is contaminated by wild birds (McDonald 1969a). Once established, the parasites can be maintained locally by domestic and wild ducks alike. Passerines are often infected with common parasites of poultry such as C. infundibulum and Raillietina echinobothrida (Reid 1983; Ibrahim 2006) and are a potential source of infection for chickens. In contrast, Zetterman et al. (2005) found two species of cestodes in wild Greater Rheas (Rhea americana), but none in captive birds. Other parasites were present in both groups, suggesting that either the intermediate hosts for the cestodes were absent or the area had not been contaminated with eggs. Diphyllobothriid cestodes found in birds use fish as second intermediate hosts. The pleurocercoids of
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Ligula (Cozma and Friciu 1997; Loot et al. 2001; Heckmann 2005) and Diphyllobothrium (Rodger 1991) are pathogenic to fish. Infected birds, attracted to freshwater aquaculture facilities, can easily transmit infections to farmed fish if suitable copepod intermediate hosts are available. Pleurocercoid infections are common in farmed fish (Kitit*yna and Nikitenko 1986; H˚astein and Lindstad 1991; Cozma and Friciu 1997) and are a common cause of disease and mortality in these intermediate hosts (Kurovskaya 1993; Rhakonen et al. 1996). Pleurocercoids of some Diphyllobothrium species may be transmissible to humans (Roberts and Janovy 2005) when consumed in raw or poorly cooked fish and pose a potential health threat. WILDLIFE POPULATION IMPACTS The potential influence of parasites on host population dynamics is difficult to assess (Peterson 2004). Adult cestodes are not generally considered pathogenic or a threat to avian populations under normal conditions (Cornwell and Cowan 1963; Wobeser 1981; Harradine 1982; Thomas 1985; Greve 1986; Sasseville et al. 1988; Purvis et al. 1998; Delahay 1999; Haukos and Neaville 2003). Absence of clinical signs makes it difficult to detect infected birds. Sick birds usually die unnoticed and, if they are found, they are usually infected with a variety of parasites, making it impossible to attribute the condition to a specific agent. Reports of mortality in the field need to be interpreted with caution, particularly if mixed infections are involved. For example, cestodes have been associated with emaciation and starvation of large numbers of birds during sudden cold snaps (James and Llewellyn 1967; Jaramillo and Rising 1995), but their role in these deaths remains unresolved. Parasitism by cestodes may affect reproduction and mate selection, with corresponding impacts on population size. Mortality in breeding Willow Ptarmigan infected with Hymenolepis microps increased with intensity of infection, with subsequent reductions in the annual growth rate of the host population (Holmstad et al. 2005). Similarly, female Common Eiders with heavy cestode infections may forgo breeding (Hario et al. 1992) rather than produce smaller clutches that may have to be abandoned later. Eiders reach sexual maturity later than most ducks, produce comparatively small clutches, and do not renest if the first one is lost. In species such as these, reduction either in survival or in the number of nesting females could have a significant impact on population numbers at least at the local level. Infection with cestodes may also affect plumage quality and sexual ornamentation, with subsequent effects on mate selection. For example, Bar-tailed
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Godwits (Limosa lapponica) in good body condition may undergo a partial molt during spring migration at staging points in the Wadden Sea, while lighter birds do not (Piersma et al. 2001). Heavy, well-ornamented birds replace some of their contour feathers and display more extensive breeding plumage than do nonmolting birds. Birds undergoing the molt had fewer cestodes and, among females, quality of the breeding plumage was inversely associated with intensity of infection. TREATMENT AND CONTROL Wild birds brought in from the field for propagation or relocation programs are usually infected with cestodes. Stress associated with capture or confinement may exacerbate the effects of cestode infections (Jones et al. 1996a). Treatment recommendations vary but in general niclosamide (Yomesin) has been recommended for species of Gruiformes (Carpenter 1986), Anseriformes (except for geese) (Humphries 1986), Falconiformes, and Strigiformes (Ward 1986). Praziquantel is effective in species of Columbiformes (Zwart 1986), starlings and other Sturnidae (Letcher 1986), and in bustards (Jones et al. 1996a). Both niclosamide and flubendazole are effective in controlling infection in captive flocks of bustards (Jones et al. 1996a). Fockema et al. (1985) successfully treated Ostriches infected with Houttuynia struthionis with fenbendazole. There is no practical way to control cestode infections in wild birds. Control measures require a disruption in life cycles by reducing or eliminating contact with potential intermediate hosts. This may be possible on a limited scale for captive flocks but is not feasible on a scale that would affect wild populations. MANAGEMENT IMPLICATIONS There are no effective management options available to control cestode infections in wild birds. The vagility of these hosts ensures that the parasites will be spread widely in local habitats and that those of migratory species will be spread over even broader geographic areas. Birds to be transported into new areas either as captives or for release should be treated for cestodes prior to shipment to reduce the possibility of introducing novel species. Similarly, the cestode fauna of local birds should be studied before restoration projects are undertaken to access potential risks to translocated or introduced species (Kocan et al. 1979). ACKNOWLEDGMENTS I thank Research Librarians Dubrava Kapa and Ruth Noble and Ms Annie Ciarlo, Concordia University for their assistance. Dr David Stallknecht, Editor,
Journal of Wildlife Diseases, kindly granted permission to reproduce Figures 14.2 and 14.3 from the paper by Karstad, Sileo, Okech, and Khalil (1982) published in Journal of Wildlife Diseases, Vol. 18, pp. 507–508. A special thanks to Dr Paul Albert, Department of Biology, Concordia University, who made the copies of Figures 14.1–14.3. This work was supported through the Natural Sciences and Engineering Research Council of Canada Discovery Grant A6979 to J. D. McLaughlin.
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Schmidt, G. D. 1986. Handbook of Tapeworm Identification. CRC Press, Boca Raton, FL. Schmidt, K., D. Duwel, and D. Bartzuki. 1987. Helminthosen des wassergelfl¨ugels und ihre behandlung. Erkrankungen der Zootiere 29:45–55. Scholz, T., R. A. Bray, R. Kuchta, and R. Repov´a. 2004. Larvae of gryporhynchid cestodes (Cyclophyllidea) from fish: A review. Folia Parasitologica 51:131–152. ˇ Slais, J. 1961. Darmschleimhautsch¨adigung durch Bandw¨urmer Aploparaxsis furcigera (Rudolphi) und Hymenolepis parvula (Kowalewski) bei Enten. Helminthologia 3:316–321. Spasskaya, L. P. 1966. Cestodes of Birds of the USSR (In Russian). Izdatelstvo “Nauka,” Moscow. Stock, T. M., and J. C. Holmes. 1987. Host specificity and exchange of intestinal helminths among four species of grebes (Podicipedidae). Canadian Journal of Zoology 65:669–676. Sprehn, C. E. W. 1932. Lehrbuch der Helminthologie. Eine Naturgeschichte der in S¨augtieren deutschenund V¨ogeln schmarotzenden W¨urmer, unterbesonderer Ber¨ucksichtigung der Helminthen des Menschen, der Haustiere, und wichtigsten Nutziere. Borntraeger, Berlin. Thomas, V. G. 1985. Body condition of willow ptarmigan parasitized by cestodes during winter. Canadian Journal of Zoology 64:251–254. Toplu, N., O. Sarimehmetoglu, N. Metin, and H. Eren. 2006. Pleural and peritoneal tetrathyridiosis in a peafowl. Veterinary Record 158:102–103. Ueta, M. T., and R. M. P. Avancini. 1994. Studies on the influence of age in the infection of caged chickens by
Raillietina laticanalis and on the susceptibility to reinfection. Veterinary Parasitology 52:157–162. Ward, H. P. 1986. Raptors (Falconiformes and Strigeiformes) parasites and their treatment in birds of prey. In Zoo and Wild Animal Medicine, 2nd ed., M. E. Fowler (ed.). W. B. Saunders, Philadelphia, PA, pp. 425–430. Wallace, B. M., and D. B. Pence. 1986. Population dynamics of the helminth community from migrating blue-winged teal: Loss of helminths without replacement. Canadian Journal of Zoology 64:1765–1773. Willers, W. B., and O. W. Olsen. 1969. Incidence of infection by Gastrotaenia cygni (Cestoda: Aporidea) of waterfowl in eastern Colorado. Avian Diseases 13:415–416. Wobeser, G. 1974. Renal coccidiosis in mallard and pintail ducks. The Journal of Wildlife Diseases 10:249–255. Wobeser, G. 1981. Diseases of Wild Waterfowl, 2nd ed. Plenum Press, New York. Wolffhugel, K. 1938. Nematoparataeniidae. Zeitschrift f¨ur infektionskrankheiten, parasit¨are krankheiten und hygiene der haustiere 49:9–42. Zetterman, C. D., A. A. Nascimento, J. A. Tebaldi, and M. J. P. Szabo. 2005. Observations on helminth infection of free-living and captive rheas (Rhea americana) in Brazil. Veterinary Parasitology 129:169–172. Zwart, P. 1986. Pigeons and Doves (Columbiformes). In Zoo and Wild Animal Medecine, 2nd ed., M. E. Fowler (ed.). W. B. Saunders, Philadelphia, PA.
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15 Acanthocephala Dennis J. Richardson and Brent B. Nickol more species than do mammals, which in turn host more species than do amphibians. Species of acanthocephalans from avian hosts are mainly represented by a few broadly distributed genera and are harbored by birds of relatively few taxonomic orders. Waterfowl (Anseriformes) are the most heavily parasitized group of birds. Species of the genus Corynosoma and Polymorphus are the most common forms in waterfowl. Acanthocephalans of the genera Arhythmorhynchus and Plagiorhynchus are the principle forms in shorebirds (Charadriiformes). Species of Lueheia, Mediorhynchus, and Plagiorhynchus comprise most of the acanthocephalans in perching birds (Passeriformes). Hawks (Falconiformes) and owls (Strigiformes) most frequently are parasitized by species of Centrorhynchus, Sphaerirostris, and Oligacanthorhynchus.
INTRODUCTION Worms of the phylum Acanthocephala (Greek: akantha, spine or thorn + kephale, head) are known as spiny-headed or thorny-headed worms due to the nature of their holdfast organ, called a proboscis. Acanthocephalans are dioecious pseudocoelomate worms remarkably adapted to a parasitic lifestyle in that there is no mouth or digestive system. Worms absorb nutrients directly through their integument. Adult acanthocephalans vary greatly in size from a few millimeters to over 10 cm long, depending on species, and occur exclusively in the vertebrate small intestine. All acanthocephalans exhibit an indirect life cycle in which the vertebrate definitive host becomes infected by ingesting larvae, known as cystacanths, contained in the hemocoel (body cavity), of an arthropod intermediate host. Although they are capable of causing extreme pathology and death and may be responsible for epizootic outbreaks under certain circ*mstances, by and large, acanthocephalans cause little overt pathology in their avian hosts.
ETIOLOGY In the most recent complete list, Golvan (1994) considered Acanthocephala to comprise slightly more than 1,100 valid species. The most important character from a taxonomic standpoint is the spiny holdfast structure or proboscis. The retractable and invagin*ble proboscis is used by the worm to attach to the intestinal wall of its vertebrate definitive host. Representative proboscides of acanthocephalans parasitizing avian hosts are shown in Figure 15.1. Basic acanthocephalan anatomy is shown in Figure 15.2. The review by Miller and Dunagan (1985) should be consulted for a more comprehensive account of functional morphology. Starling (1985) and Taraschewski (2000) reviewed nutrition and metabolism of Acanthocephala.
SYNONYMS Acanthocephalosis, Acanthocephaliasis. HISTORY Acanthocephalans were first recognized from the intestine of eels by Redi (1684). Since then approximately 1,100 species of acanthocephalans have been described (Golvan 1994), with approximately 400 species being recognized from birds. A concise history of the study of acanthocephalans can be found in Amin (1985).
EPIZOOTIOLOGY Acanthocephalan species for which life cycles have been confirmed require vertebrates for definitive hosts and arthropods as intermediate hosts. Schmidt (1985) provided a summary of known life cycles. Adult
HOST RANGE AND DISTRIBUTION Animals of all vertebrate classes serve as definitive hosts for acanthocephalans. Bony fishes are the most parasitized group and reptiles are the least. Birds harbor
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Figure 15.1. Proboscides of some common acanthocephalans of birds. (a) Centrorhynchus robustus, an acanthocephalan of owls. Bar = 250 μm. Redrawn from Richardson and Nickol (1995). (b) Polymorphus cucullatus from a Hooded Merganser (Lophodytes cucullatus). Bar = 500 μm. Redrawn from Van Cleave and Starrett (1940). (c) Mediorhynchus centurorum from a Red-bellied Woodpecker (Melanerpes carolinus). Bar = 220 μm. Redrawn from Nickol (1969). (d)) Plagiorhynchus cylindraceus from an American Robin (Turdus migratorius). Bar = 1 mm. Redrawn from Schmidt and Olsen (1964).
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Figure 15.2. Male Plagiorhynchus cylindraceus from an American robin (Turdus migratorius). P, proboscis; PR, proboscis receptacle; L, lemnisci; T, testes; CG, cement glands; SP, Saefftigen’s pouch; CB, copulatory bursa. Bar = 1 mm.
female worms release eggs that are passed in the feces of the definitive host. Only eggs exist outside of a host, free in the environment, and transmission from one definitive host to another requires that appropriate invertebrate intermediate hosts ingest eggs. A typical acanthocephalan life cycle is shown in Figure 15.3. Intermediate hosts are known for only about 7% of the species that parasitize birds. Those with terrestrial life cycles usually have insects, frequently species of Coleoptera or Orthoptera, or terrestrial isopods for intermediate hosts. Decapods and microcrustaceans, usually species of Amphipoda or Isopoda, are intermediate hosts for those with aquatic life cycles.
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A larval stage, the acanthor, emerges from the egg upon its ingestion by an arthropod. After penetration of the wall of the alimentary canal, the acanthor undergoes development within the body cavity of its intermediate host, ultimately achieving the final ontogenetic stage, the cystacanth, which is infective to potential definitive hosts. “Cystacanth” has achieved general usage as a name for the stage infective to a final host regardless of whether it is found in the arthropod intermediate host or in a vertebrate paratenic host (Van Cleave 1953). No species of Acanthocephala has been demonstrated to require more than the arthropod intermediate host in order to develop infectivity to vertebrates. However, in the life cycle of some species, another vertebrate host occurs between the arthropod intermediate and vertebrate definitive host. In such hosts, cystacanths penetrate the intestinal wall and localize in mesenteries or visceral organs, but do not attain sexual maturity. Although such intercalated hosts may be required to complete transfer of acanthocephalans from intermediate hosts at the trophic level which potential definitive hosts feed, there is no evidence that they are essential for achievement of infectivity to the final host. The term “paratenic host” has attained wide usage for such animals in which ontogeny does not proceed (Baer 1951; Beaver 1969). An example of a life cycle of an acanthocephalan utilizing a paratenic host is shown in Figure 15.4. Many, if not most, birds acquire acanthocephalans from an intermediate rather than a paratenic host. This is the route of transmission for a large number of species of Corynosoma and Polymorphus found in waterfowl and for species of Plagiorhynchus that occur in charadriiform and passerine birds. Species of Mediorhynchus are also transmitted to passerine birds in this manner. Piscivorous birds frequently acquire acanthocephalans from fishes in their diet although other poikilothermic vertebrates are paratenic hosts for some species. Southwellina hispida has a broad geographical distribution in the Black-crowned Night Heron (Nycticorax nycticorax) and it occurs in mesenteries of fish, frogs, and snakes (Van Cleave 1925; Yamaguti 1935, 1939). Amphibians and reptiles also serve as paratenic hosts for some acanthocephalan species that mature in flesh-eating birds. Species of Centrorhynchus and the related Sphaerirostris are well known as cystacanths in frogs, lizards, and snakes. Adults occur in raptors and other kinds of carnivorous birds. Golvan (1956) and Schmidt and Kuntz (1969) listed many of the definitive and paratenic hosts for species of these genera. Likewise several species of Oligacanthorhynchus occur as
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Figure 15.3. Life cycle of Corynosoma constrictum, a common acanthocephalan of waterfowl, particularly ducks, throughout North America and parts of South America. Corynosoma constrictum uses amphipods (Hyallela azteca) as intermediate hosts.
adults in birds of prey and the literature abounds with worldwide reports of oligacanthorhynchid cystacanths in the viscera of reptiles, usually snakes. Acanthocephalans are found infrequently in extraintestinal sites in birds, but they seem to be important paratenic hosts only for some species of Oncicola. Oncicola canis and Oncicola oncicola, parasites of canine and feline definitive hosts in the Americas, have been found in the outer surface of the esophagus and crop of the Northern Bobwhite (Colinus virginianus), and subcutaneously in the musculature of domestic chickens (Cram 1931; Zeledon and Arroyo 1960). Australian and Asian species of Oncicola also use birds for paratenic hosts (Schmidt 1983).
CLINICAL SIGNS Little is understood about clinical signs in birds infected with acanthocephalans. There are many reports of paralyzed and moribund birds with acanthocephalans (e.g., Jones 1928; Webster 1943; Holloway 1966; McOrist and Scott 1989). Many of these cases have involved American Robins (Turdus migratorius) infected with Plagiorhynchus cylindraceus (Figure 15.5). Birds with high-intensity infections are frequently emaciated and stunted (Hynes and Nicholas 1963).
PATHOGENESIS AND PATHOLOGY It has long perplexed helminthologists that, in some cases, acanthocephalan infections of low intensity seem to have serious adverse effects on an infected animal. In other instances, clinical signs are absent in conspecific animals with infections of high intensity of the same species (Soulsby 1958). On the basis of their extensive study of Polymorphus minutus in domestic ducks, Hynes and Nicholas (1963) suggested that under normal circ*mstances infections build slowly and that density-dependent establishment might limit infections to subclinical levels. However, infections of high intensity with clinical consequences might result if an uninfected bird were suddenly exposed to a large number of acanthocephalans. In contrast to mortality attributed to intense acanthocephalan infections, it is far more common to find equally intense infections in birds that show no sign of disease (Hynes and Nicholas 1963; Schmidt 1972; Moore and Bell 1983). There are very few studies that assist with assessing the importance of infections of low intensity or infections, even if intense, in which clinical effects appear to be lacking. The extent of pathogenesis is likely influenced by the nutritional status of the host (Holmes 1987) and by environmental stress (Grenquist 1970). Connors and
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Figure 15.4. Paratenic transmission of Lueheia inscripta. Reptiles become infected by ingesting cystacanths contained within the body cavity of co*ckroaches. Within the reptilian paratenic host the cystacanths localize in the mesenteries or visceral organs but do not obtain sexual maturity. Passerine birds may become infected when they ingest cystacanths contained within paratenic hosts. Nickol (1991) demonstrated that P. cylindraceus has a significant detrimental effect on the flow of food energy through infected European Starlings (Sturnus vulgaris). Both male and female starlings show reductions in standard metabolic rates as a result of infection, indicating that their basal metabolism and thermal regulatory abilities are altered. Infected male
birds have an increased consumption and excretion of energy, and they average lower daily body weights than do uninfected males when they are temperaturestressed (Connors and Nickol 1991). There seems little question that subclinical infections can become serious in times of nutritional or environmental stress.
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Figure 15.5. Plagiorhynchus cylindraceus in the intestine of an American Robin (Turdus migratorius) found frozen near Owensboro, Kentucky. The robin harbored 69 acanthocephalans. Courtesy of D. F. Oetinger.
The significance of acanthocephalans in dying or dead animals (Figure 15.5) is difficult to interpret because the link is always circ*mstantial and because intensities are often no greater in affected animals than in conspecific individuals showing no adverse effect (Hynes and Nicholas 1963; Schmidt 1972; Moore and Bell 1983); thus, acanthocephalan infection at any intensity should be considered to have pathogenic potential. Attachment of the acanthocephalan proboscis sometimes causes formation of fibrinous nodules on the serosal surface of the intestine. In Red-bellied Woodpeckers (Melanerpes carolinus), nodules that form around the proboscis, neck, and foretrunk of Mediorhynchus centurorum are frequently at least 4 mm long (Nickol 1969). Adult worms of most acanthocephalan species have not been reported to induce nodules in their hosts. Others, such as P. minutus in domestic ducks, may or may not induce nodule formation (Nicholas and Hynes 1958). Still others, such as Profilicollis botulus, seem always to cause nodules to form (Bishop and Threlfall 1974; Bourgeois and Threlfall 1982). Upon necropsy, acanthocephalans occasionally are found protruding from the intestine into the coelom, having perforated the intestinal wall. In Western Bluebirds (Sialia mexicana) infected with P. cylindraceus and in Common Eiders (Somateria mollissima) and Mute Swans (Cyngus olor) infected with P. minutus, peritonitis resulting from perforation has been linked to mortality (Clark et al. 1958; Sanford 1978; ThompsonCowley et al. 1979). Although perforation through acanthocephalan-induced nodules sometimes occurs (Bishop and Threlfall 1974), perforation apparently is
Figure 15.6. Mediorhynchus centurorum in the intestine of a Red-bellied Woodpecker (Melanerpes carolinus). The trunk of the worm has eroded villi, allowing blood vessels to leak. The resulting pus (arrow) contains blood cells, macrophages, polymorphonuclear cells, clotted fibrin, and damaged columnar epithelial cells. C, circular muscle of worm body wall; L, longitudinal muscle of worm body wall; LC, lacunar canal of worm tegument; P, pseudocoelom of worm; V, abraded villus of bird mucosa. Bar = 50 μm. independent of nodule formation (Thompson-Cowley et al. 1979). Histological damage caused by acanthocephalans has been studied more extensively in fish and mammals (Bullock 1963; Chaicharn and Bullock 1967; Szalai and Dick 1987; Richardson and Barnawell 1995) than in birds, with a few notable exceptions (e.g., Nicholas and Hynes 1958; Petrochenko 1958; Schmidt 1963; Sanford 1978; Moore and Bell 1983; Taraschewski and Hofmann 1991). Along the trunks of Mediorhynchus gallinarum and M. centurorum, villi and basal glands of domestic fowl and woodpeckers, respectively, are compressed and eroded (Figure 15.6) in much the same manner as has been described for mammals (Nath and Pande 1963; Nelson and Nickol 1986; Richardson and
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Barnawell 1995) and fish (De Buron and Nickol 1994) infected with acanthocephalans. At sites of attachment of the proboscis, frequent microscopic changes for birds are acute, multifocal, necrotizing transmural inflammation and necrosis and ulcerative enteritis resulting from penetration of the intestinal wall by the proboscis (Bolette 1987). Chronic inflammation at the site where the proboscis attaches may cause fibrinous adhesions that bind viscera, reduce mobility of the gut, and cause emaciation (Bishop and Threlfall 1974).
taxonomic keys available for acanthocephalans found in birds. Most are limited to specific acanthocephalan groups or to individual taxa of birds. The original systematic literature is the most reliable means of specific identification.
DIAGNOSIS Acanthocephalan infections are detected in live birds by observation of eggs in feces of infected animals. Acanthocephalan eggs are recognized by their characteristic membranes that enclose a spined acanthor (Figure 15.7a). Generic identification is possible from the eggs (Figure 15.7). Specific identification may be made by a specialist in many instances. Because acanthocephalan eggs do not float readily in standard floatation mixtures, sedimentation techniques are preferred. The ethyl acetate/formalin sedimentation procedure, developed by Ritchie (1948) and modified by Markell et al. (1999), works well. Diagnosis can also be made by identifying worms obtained at necropsy. Lack of properly prepared specimens is one of the primary reasons for the paucity of information concerning acanthocephalans from wild birds. The bird should be examined as soon after death as possible and carcasses should not be frozen. The intestine should be removed and carefully dissected longitudinally. Proboscides of firmly attached acanthocephalans should be removed from the intestinal wall using insect pins or fine needles. Care must be taken not to cut or puncture the worm. For identification, the proboscis must be evagin*ted and the armature preserved intact. After removal from the intestine, all acanthocephalans should be held in tap water for at least 24 h to promote full evagin*tion of the proboscis. If the proboscis fails to evagin*te in tap water, worms may be placed in distilled water for an additional 24 h. After the period in water, worms may be fixed in a solution of 85% ethanol, formalin, and glacial acetic acid mixed at a ratio of 85:10:5 (Van Cleave 1953). After 24 h in glacial acetic acid, specimens may be stored in 70% ethanol. Richardson (2006) provided detailed information on preparing specimens for microscopic examination, and Richardson (2005) discussed how to properly assess proboscis morphometrics. The most recent comprehensive keys for identification are those by Yamaguti (1963). McDonald (1988) provided a key helpful for identification of forms found in waterfowl. There are, however, few comprehensive
DOMESTIC ANIMAL HEALTH CONCERNS Although acanthocephalans rarely constitute an economic consideration in agricultural operations, three species that occur naturally in wild birds can infect domestic poultry. Plagiorhynchus cylindraceus (=Plagiorhynchus formosus), a cosmopolitan species found in passerine birds, is of occasional concern as a potential parasite of domestic chickens in the US. (Jones 1928; Holloway 1966). This species, common in American Robins and European Starlings, can infect many kinds of birds, including domestic fowl. M. gallinarum occurs in wild and domestic galliform birds throughout Asia and Africa (Schmidt and Kuntz 1977). Significant consequences, however, have not been reported for either of these acanthocephalan species. Polymorphus minutus and the similar Polymorphus magnus have been implicated as important parasites of domestic ducks in Europe (Antipin 1956; Petrochenko 1958; Hynes and Nicholas 1963). These polymorphids are frequent parasites of anatid birds and can be transmitted from amphipod intermediate hosts to domestic fowl that have access to water frequented by wild waterfowl. Mortality in birds at the National Aviary in Pittsburgh, Pennsylvania, was attributed to Mediorhynchus orientalis that apparently was present in an imported bird. The eggs of the parasite-infected species of co*ckroaches and cystacanths developed for transmission to other birds (Bolette 1990, 2000). Acanthocephalans are relatively specific for intermediate hosts. Consequently, importation of exotic species seldom results in continuation of the life cycle through transmission to other animals in the collection. Occurrences such as that in the Pittsburgh aviary apparently are exceptional, but they attest to the fact that domestic and captive animals are at potential risk from acanthocephalans present in wildlife.
PUBLIC HEALTH CONCERNS None of the five acanthocephalan species reported from humans occur in birds (Counselman et al. 1989; Richardson 2003). It is doubtful that wild birds serve as reservoirs for human acanthocephalan infections.
WILDLIFE POPULATION IMPACTS Dead or dying birds are often discovered with large numbers of acanthocephalans (Figure 15.5).
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Figure 15.7. Eggs of some common acanthocephalans of birds. Scale bars = 10 μm. (a) Line drawing of egg of Centrorhynchus microcephalus from the intestine of a Groove-billed Ani (Crotophaga sulcirostris). Note the series of envelopes or membranes that surround the spined acanthor. Terminology follows Awachie (1966). A, acanthor; ENM, embryonic nuclear mass; IE, inner envelope of egg; ME, middle envelope of egg; OE, outer envelope of egg. (b) Egg of Profilicollis botulus that was removed from the body cavity of a gravid worm taken from a Common Eider (Somateria mollissima). Courtesy of D. W. T. Crompton. (c) Egg of Polymorphus minutus removed from the body cavity of a gravid worm taken from a domestic duck. Photograph courtesy of D. W. T. Crompton. (d) Egg of Mediorhynchus grandis removed from the body cavity of a gravid worm taken from a Western Meadowlark (Sturnella neglecta).
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Acanthocephala Thompson-Cowley et al. (1979) considered large numbers of P. cylindraceus to be a contributory cause of death among Western Bluebirds. Perry (1942) found up to 1,842 specimens of Profilicollis altmani in dead or dying Surf Scoters (Melanitta perspicillata) and White-winged Scoters (Melanitta fusca deglandi). Up to 3,500 specimens of P. minutus occurred in Common Eiders found dead during an epizootic in Finland (It¨amies et al. 1980). Acanthocephalan-induced epizootics with extensive mortality occur infrequently. Among birds, populations of Common Eiders seem especially vulnerable to epizootics that are usually caused by P. botulus and occasionally by P. minutus. Liat and Pike (1980) reviewed reports of many of these occurrences throughout the Northern Hemisphere. Populations of swans also experience occasional decimation due to acanthocephalan infections. Large numbers of the acanthocephalan species P. minutus and Filicollis anatis were considered contributors to increased mortality in a flock of Mute Swans in central Scotland (Pennycott 1998). An earlier epizootic caused by P. magnus and Polymorphus mathevossianae led to loss of 40% of the cygnets inhabiting a lake in the Kurgal’dzhin Nature Reserve in Russia (Maksimova 1972). The Scottish and Russian epizootics followed drastic reductions in water levels that probably concentrated intermediate hosts and let to unusually intense infections. Acanthocephalans that normally occur in birds may pose a threat to other kinds of wildlife when feeding activities cause exposure to large numbers of cystacanths. Mortality in California sea otters (Enhydra lutris) due to acanthocepalan-induced peritonitis has been reported in Monterey, California (Thomas and Cole 1996; Mayer et al. 2003). Otters most likely become infected by ingesting the infective cystacanths of P. altmani, normally a parasite of shore birds, in a sand crab (Emerita analoga) intermediate host (Nickol et al. 2002). Presence of gravid females in the body cavity of otters observed by Richardson (unpublished data) suggests that otters may also become infected through postcyclic transmission (ingestion of adult worms harbored in the intestine of a prey animal) while feeding on infected birds. Predation of seabirds by sea otters is rare but appears to have increased in recent years (Riedman and Estes 1988). Significant climatic events, such as El Nino and global warming, might lead to changes in feeding behavior, alter the normal cycles of transmission between parasites and hosts, and lead to greater frequency of aberrant pathogenic infections. In at least one instance, acanthocephalans of birds may have commercial impact on an intermediate host population. Southwellina dimorpha, an acanthocephalan of the White Ibis (Eudocimus albus) and Whooping
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Crane (Grus americana) uses the commercially important cultured red crawfish (Procambrus clarki) as an intermediate host. Although infrequently reported, S. dimorpha may occur in high enough prevalence in crawfish in the southeastern US to have a commercial impact (Lantz 1974; Richardson and Font, 2006). TREATMENT AND CONTROL Although many courses of chemotherapy have been credited with efficacy in individual instances (Petrochenko 1958; Hynes and Nicholas 1963; Goldsmid et al. 1974; Counselman et al. 1989; Richardson 2003), no satisfactory treatment for acanthocephalan infection is known. Prevention depends on keeping birds away from environments that harbor infected intermediate hosts (Nicholas and Hynes 1958, Sanford 1978) although such control for wild birds seems impractical. LITERATURE CITED Amin, O. A. 1985. Classification. In Biology of the Acanthocephala, D. W. T. Crompton and B. B. Nickol (eds). Cambridge University Press, Cambridge, UK, pp. 27–72. Antipin, D. N. 1956. Acanthocephaliases of Livestock. In Parasitology and Parasitic Diseases of Livestock, V. S. Ershov (ed.). 1960 English translation by The Israel Program for Scientific Translations, Office of Technical Services, U.S. Department of Commerce, Washington, DC.. State Publishing House for Agricultural Literature, Moscow, pp. 248–257. Awachie, J. B. E. 1966. The development and life-history of Echinorhynchus truttae Schrank 1788 (Acanthocephala). Journal of Helminthology 40:11–32. Baer, J. C. 1951. Ecology of Animal Parasites. University of Illinois Press, Urbana, IL. Beaver, P. C. 1969. The nature of visceral larval migrans. Journal of Parasitology 55:3–12. Bishop, C. A., and W. Threlfall. 1974. Helminth parasites of the common eider duck, Somateria mollissima (L.), in Newfoundland and Labrador. Proceedings of the Helminthological Society of Washington 41(1):25–35. Bolette, D. P. 1987. Acanthocephaliasis. Veterinary Technician 8(1):19–24. Bolette, D. P. 1990. Intermediate host of Mediorhynchus orientalis (Acanthocephala: Gigantorhynchidae). Journal of Parasitology 76:575–577. Bolette, D. P. 2000. Descriptions of cystacanths of Mediorhynchus orientalis and Mediorhynchus wardi (Acanthocephala: Gigantorhynchidae). Comparative Parasitology 67(1):114–117.
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Bourgeois, C. E., and W. Threlfall. 1982. Metazoan parasites of three species of scoter (Anatidae). Canadian Journal of Zoology 60:2253–2257. Bullock, W. L. 1963. Intestinal histology of some salmonid fishes with particular reference to the histopathology of acanthocephalan infections. Journal of Morphology 112(1):23–44. Chaicharn, A., and W. L. Bullock. 1967. The histopathology of acanthocephalan infections in suckers with observations on the intestinal histology of two species of catastomid fishes. Acta Zoologica 48:19–42. Clark, G. M., D. O’Meara, and J. W. Van Weelden. 1958. An epizootic among eider ducks involving an acanthocephalid worm. Journal of Wildlife Management 22(2):204–205. Connors, V. A., and B. B. Nickol. 1991. Effects of Plagiorhynchus cylindraceus (Acanthocephala) on the energy metabolism of adult starlings, Sturnus vulgaris. Parasitology 103:395–402. Counselman, K., C. Field, G. Lea, B. Nickol, and R. Neafie. 1989. Moniliformis moniliformis from a child in Florida. American Journal of Tropical Medicine and Hygiene 41:88–90. Cram, E. B. 1931. Recent findings in connection with parasites of game birds. Transactions of the American Game Conference 18:243–247. De Buron, I., and B. B. Nickol. 1994. Histopathological effects of the acanthocephalan Leptorhynchoides thecatus in the ceca of the green sunfish, Lepomis cyanellus. Transactions of the American Microscopical Society 113(2):161–168. Goldsmid, J. M., M. E. Smith, and F. Fleming. 1974. Human infections with Moniliformis sp. in Rhodesia. Annals of Tropical Medicine and Parasitology 68:363–364. Golvan, Y. J. 1956. Le genre Centrorhynchus Luhe 1911 (Acanthocephala-Polymorphidae). Bulletin de l’Institut Francais d’Afrique Noire, Series A, 18:732–791. Golvan, Y. J. 1994. Nomenclature of the Acanthocephala. Research and Reviews in Parasitology 54:135–205. Grenquist, P. 1970. Vakakarsamatojen aiheuttamasta haahkojen kuolleisuudesta. Suomen Riista 22:24–34. Holloway, H. L., Jr. 1966. Prosthorhynchus formosum (Van Cleve, 1918) in songbirds, with notes on acanthocephalans as potential parasites of poultry in Virginia. Virginia Journal of Science 17(3):149–154. Holmes, J. H. 1987. Pathophysiology of parasitic infections. Parasitology 94(Suppl):S29–S51. Hynes, H. B. N., and W. L. Nicholas. 1963. The importance of the acanthocephalan Polymorphus minutus as a parasite of domestic ducks in the United Kingdom. Journal of Helminthology 37(3):185–198.
It¨amies, J., E. T. Valtonen, and H.-P. fa*gerholm. 1980. Polymorphus minutus in eiders and its role as a possible cause of death. Annales Zoologici Fennici 17:285–289. Jones, M. 1928. An acanthocephalid, Plagiorhynchus formosus, from the chicken and the robin. Journal of Agricultural Research 36(9):773–775. Lantz, K. E. 1974. Acanthocephalan occurrence in cultured red crawfish. In Proceedings of the 27th Annual Conference of the Southeastern Association of Game and Fish Commissioners 1973:735–738. Liat, L. B., and A. W. Pike. 1980. The incidence and distribution of Profilicollis botulus (Acanthocephala), in the eider duck, Somateria mollissima, and in its intermediate host the shore crab, Carcinus maenas, in north east Scotland. Journal of Zoology 190: 39–51. Maksimova, A. P. 1972. Death of young swans due to helminths in the Kurgal’dzhin reserve. Izdatel’stvo “FAN” Uzbekskoi, SSR 1972:125–129. Markell, E. K., D. T. John, and W. A. Krotski. 1999. Markell and Voge’s MedicalParasitology, 8th ed. W. B. Saunders Philadelphia, PA. McDonald, M. E. 1988. Key to Acanthocephala Reported in Waterfowl. Fish and Wildlife Service Resource Publication 173. United States Department of the Interior, Washington, DC. Mayer, K. A., M. D. Dailey, and M. A. Miller. 2003. Helminth parasites of the southern sea otter Enhydra lutris nereis in central California: Abundance, distribution and pathology. Diseases of Aquatic Organisms 53:77–88 McOrist, S., and P. C. Scott. 1989. Parasitic enteritis in superb lyrebirds (Menura novaehollandiae). Journal of Wildlife Diseases 25:420–421. Miller, D. M., and T. T. Dunagan. 1985. Functional Morphology. In Biology of the Acanthocephala, D. W. T. Crompton and B. B. Nickol (eds). Cambridge University Press, Cambridge, UK, pp. 73–123. Moore, J., and D. H. Bell. 1983. Pathology (?) of Plagiorhynchus cylindraceus in the starling, Sturnus vulgaris. Journal of Parasitology 69(2):387–390. Nath, D., and B. P. Pande. 1963. A note on the acanthocephalan infection of domestic fowl. Indian Journal of Helminthology 15:31–35. Nelson, M. J., and B. B. Nickol. 1986. Survival of Macracanthorhynchus ingens in swine and histopathology of infection in swine and raccoons. Journal of Parasitology 72:306–314. Nicholas, W. L., and H. B. N. Hynes. 1958. Studies on Polymorphus minutus (Goeze, 1782) (Acanthocephala) as a parasite of the domestic duck. Annals of Tropical Medicine and Parasitology 52:36–47.
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Acanthocephala Nickol, B. B. 1969. Acanthocephala of Louisiana picidae with description of a new species of Mediorhynchus. Journal of Parasitology 55(2):324–328. Nickol, B. B., R. W. Heard, and N. F. Smith. 2002. Acanthocephalans from crabs in the Southeastern U.S., with the first intermediate hosts known for Arythmorhynchus frassoni and Hexaglandula corynosoma. Journal of Parasitology 88(1):79–83. Pennycott, T. W. 1998. Lead poisoning and parasitism in a flock of mute swans (Cygnus olor) in Scotland. Veterinary Record 142:13–17. Perry, M. L. 1942. A new species of the acanthocephalan genus Filicollis. Journal of Parasitology 28:385–388. Petrochenko, V. I. 1958. Acanthocephala of Domestic and Wild Animals, Vol. 2. Izdatel’stvo Akademii Nauk SSSR, Moscow. Redi, F. 1684. Osservazioni Intorno Agli Animali Viventi che si Trovano Regli Animali Viventi. Accademia della Crusca, Firenze, Italy. Richardson, D. J. 2003. Other noteworthy zoonotic helminths. In North American Parasitic Zoonoses, D. J. Richardson and P. J. Krause (eds). Kluwer Academic Publishers, Boston, MA, pp. 85–111. Richardson, D. J. 2005. Identification of cystacanths and adults of Oligacanthorhynchus tortuosa, Macracanthorhynchus ingens, and Macracanthorhynchus hirudinaceus based on proboscis and hook morphometrics. Arkansas Academy of Science 59:205–209. Richardson, D. J. 2006. Life cycle of Oligacanthorhynchus tortuosa (Oligacanthorhynchidae), an acanthocephalan of the Virginia opossum (Didelphis virginiana). Comparative Parasitology 73(1):1–6. Richardson, D. J., and E. B. Barnawell. 1995. Histopathology of Oligacanthorhynchus tortuosa (Oligacanthorhynchidae) infection in the Virginia opossum (Didelphis virginiana). Journal of the Helminthological Society of Washington 62:253–256. Richardson, D. J., and W. F. Font. 2006. The Cajun dwarf crawfish (Cambarellus shufeldtii): An intermediate host for Southwellina dimorpha (Acanthocephala). Journal of the Arkansas Academy of Science 60:192–193. Richardson, D. J., and B. B. Nickol. 1995. The genus Centrorhynchus (Acanthocephala) in North America with description of Centrorhynchus robustus n. sp., redescription of Centrorhynchus kuntzi, and a key to species. Journal of Parasitology 81(5):767–772. Riedman, M. L., and J. A. Estes. 1988. Predation on seabirds by sea otters. Canadian Journal of Zoology 66:1396–1402. Ritchie, L. S. 1948. An ether sedimentation technique for routine stool examinations. Bulletin of the U.S. Army Medical Department 8(5):326.
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Sanford, S. E. 1978. Mortality in mute swans in southern Ontario associated with infestation with the thorny-headed worm, Polymorphus boschadis. Canadian Veterinary Journal 19: 234–236. Schmidt, G. D. 1963. Histopathology of a robin infected with the acanthocephalan parasite Prosthorhynchus formosus Van Cleave. Journal of the Colorado-Wyoming Academy of Science 5:54. Schmidt, G. D. 1972. Acanthocephala of captive primates. In Pathology of Simian Primates, Part II, R. N. T. -W.-Fiennes (ed.). S. Karger, Basel, Switzerland, pp. 144–156. Schmidt, G. D. 1983. What is Echinorhynchus pomatostomi Johnston and Cleland, 1912? Journal of Parasitology 69:397–399. Schmidt, G. D. 1985. Development and life cycles. In Biology of the Acanthocephala, D. W. T. Crompton and B. B. Nickol (eds). Cambridge University Press, Cambridge, UK, pp. 273–305. Schmidt, G. D., and R. E. Kuntz. 1969. Centrorhynchus spilornae sp. n. (Acanthocephala), and other Centrorhynchidae from the Far East. Journal of Parasitology 55:329–334. Schmidt, G. D., and R. E. Kuntz. 1977. Revision of Mediorhynchus Van Cleave 1916 (Acanthocephala) with a key to species. Journal of Parasitology 63(3):500–507. Schmidt, G. D., and O. W. Olsen. 1964. Life cycle and development of Prosthorhynchus formosus (Van Cleave, 1918) Travassos, 1926, an acanthocephalan parasite of birds. Journal of Parasitology 50(6):721–730. Soulsby, E. J. L. 1958. Parasitological finding in viscera sent for examination by wildfowlers. Bulletin of the British Ornithology Club 78:21–22. Starling, J. A. 1985. Feeding, nutrition and metabolism. In Biology of the Acanthocephala, D. W. T. Crompton and B. B. Nickol (eds). Cambridge University Press, Cambridge, UK, pp. 125–212. Szalai, A. J., and T. A. Dick. 1987. Intestinal pathology and the site specificity of the acanthocephalan Neoechinorhynchus carpiodi Dechtiar, 1968, in quillback, Carpiodes cyprinus (Lesueur). Journal of Parasitology 73:467–475. Taraschewski, H. 2000. Host–parasite interactions in Acanthocephala: A morphological approach. Advances in Parasitology 46:1–179. Taraschewski, H., and U. Hofmann. 1991. Host–parasite interface of Filicollis anatis (Palaeacanthocephala) in domestic ducks. Diseases of Aquatic Organisms 11:155–162. Thomas, N. J., and R. A. Cole. 1996. The risk of disease and threats to the wild population. Endangered Species Update 13(12):23–27.
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Thompson-Cowley, L. L., D. H. Helfer, G. D. Schmidt, and E. K. Eltzroth. 1979. Acanthocephalan parasitism in the western bluebird (Sialia mexicana). Avian Diseases 23(3):768–771. Van Cleave, H. J. 1925. Acanthocephala from Japan. Parasitology 17:149–156. Van Cleave, H. J. 1953. Acanthocephala of North American mammals. Illinois Biological Monographs 23:1–179. Van Cleave, H. J., and W. C. Starrett. 1940. The Acanthocephala of wild ducks in central Illinois, with descriptions of two new species. Transactions of the American Microscopical Society 59(3):348–353. Webster, J. D. 1943. Helminths from the robin, with the
description of a new nematode, Porrocaecum brevispiculum. Journal of Parasitology 29:161–163. Yamaguti, S. 1935. Studies on the helminth fauna of Japan. Part 8. Acanthocephala, I. Japanese Journal of Zoology 6:247–278. Yamaguti, S. 1939. Studies on the helminth fauna of Japan. Part 29. Acanthocephala, II. Japanese Journal of Zoology 13:317–351. Yamaguti, S. 1963. Systema Helminthum, Vol. V.: Acanthocephala. Interscience Publishers, New York. Zeledon, R., and G. Arroyo. 1960. Presencia de formas larvarias de Oncicola oncicola (Acanthocephala) en una gallina domestica. Revista de Biologia Tropical 8:197–199.
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16 Eustrongylidosis Marilyn G. Spalding and Donald J. Forrester 1977) followed by additional epizootics in Louisiana and Florida (Roffe 1988; Spalding et al. 1993). Individual bird mortalities, however, involving Great Blue Herons (Ardea herodius) and Black-crowned NightHerons (Nycticorax nycticorax) were noted before that in the Washington, DC, area (Chapin 1926; Cram 1933). These parasites were first known in fish and were subsequently shown to be the same as those infecting piscivorous birds (Leuckart 1868). Interest in these parasites increased when outbreaks in Romania in 1927 decreased the value of commercial fish collected from freshwater lakes. Ciurea (1938) was able to demonstrate through experimental infections that larvae from these fish could infect cormorants. This led to early management suggestions that the parasite could be controlled by removing piscivorous birds and draining and refilling lakes. More recent work has been done to resolve the taxonomy (Measures 1988a), the life cycle (Coyner et al. 2003a), and to uncover relationships between nutrient pollution and epizootic outbreaks of disease (Spalding et al. 1993; Coyner et al. 2002).
INTRODUCTION Eustrongylidosis is a disease of piscivorous birds caused by infection with a large dioctophymoid nematode of the genus Eustrongylides. Although a large number of species have been described, only three are considered valid at present: Eustrongylides tubifex, Eustrongylides excisus, and Eustrongylides ignotus (Measures 1988a). Only E. ignotus seems to cause significant disease when infecting herons and egrets. When infected fish containing larval stages of E. ignotus are consumed by herons, egrets, and longlegged wading birds, the parasites perforate the stomach wall and cause severe tubular fibrinous to fibrous peritonitis. Young birds are particularly sensitive and often die from hemorrhage or secondary bacterial infection. Older birds may survive infections of low intensity, but subsequently develop chronic peritonitis. Eustrongylidosis is the most commonly documented cause of epizootic mortality in nestling wading birds in some areas. Up to 80% of nestlings in some locations in Florida have been documented to have died directly or from complications related to infection with this parasite (Spalding et al. 1993). Eutrophication of foraging sites has been associated with increased prevalence of eustrongylidosis in both nestlings and adult birds (Spalding et al. 1993). Morbidity and mortality is less common with infections of E. tubifex, although this species can cause a nodular mass in the proventriculus of mergansers. Morbidity and mortality have not been reported in cormorants infected with E. excisus.
HOST RANGE AND DISTRIBUTION The distribution of Eustrongylides spp. in both fish and birds is worldwide and includes both temperate and tropical climates (Figure 16.1). Remarkably, reports of its occurrence are spotty in spite of the large size and bright red color of the parasites. Reports of larval worms from fish intermediate hosts are much more common than those from avian definitive hosts where adult, sexually reproducing worms are found, undoubtedly because of the economic importance of the fish as food for humans. Species-specific distribution information is limited by confusion over identification of the adult parasites in the definitive avian host, identification of larvae in intermediate or paratenic hosts, and complicated by occurrence of multiple species in the same definitive host (Jagerskiold 1909b, Karmanova
SYNONYMS Verminous peritonitis, eustrongylidiasis. HISTORY Epizootics of eustrongylidosis were first reported in the US in the 1970s. Large numbers of herons and egrets died at a colony in Delaware in 1976 (Wiese et al.
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Figure 16.1. Map illustrating the distribution of Eustrongylides spp. in fish and birds throughout the world. When known, the species is indicated. Square denotes E. ignotus; triangle, E. tubifex; circle, E. excisus; diamond, Eustrongylides sp. 1986). Eustrongylides ignotus and E. tubifex are found most frequently in the Americas and Europe, whereas E. excisus is most common in Asia. The geographic locations of infected birds are listed in Table 16.1 and include such diverse groups as herons, egrets, spoonbills, penguins, cormorants, coots, eagles, ducks, geese, gulls, and even some passeriform birds. The definitive hosts of E. ignotus appear to be limited to species of Ciconiiformes of the family Ardeidae (Spalding and Forrester 1993), even though parasites and lesions are found in other Ciconiiformes and Pelecaniformes. For example, although infections are found in White Ibis (Eudocimus albus) and Roseate Spoonbills (Platalea ajaja) (family Threskiornithidae), only larval stages have been reported. The range of E. ignotus includes the Americas, from Ontario, Canada, to Brazil and from the East Coast of the US to California. There is a single verified record from a Little Pied Cormorant (Phalacrocorax melanoleucos brevirostris) in New Zealand (Measures 1988a). Great Blue Herons in Ontario, Canada, were infected with both E. tubifex and E. ignotus (Measures
1988c); therefore, species of Ardeidae appear to be susceptible to infections with more than one species of Eustrongylides. The failure to find E. tubifex in piscivorous birds in Florida indicates that there may be regional differences in the distribution of the two species (Spalding et al. 1993). It is possible that locations where transmission occurs are more geographically limited than suggested by definitive host records because of migratory behavior of these hosts. Infected fish are commonly reported throughout Central and South America, but remarkably little is known about distribution and prevalence of infections in avian hosts there. Eustrongylides ignotus was identified in all avian cases in Central and South America except for a single report of E. tubifex in a Cocoi Heron (Ardea cocoi) from Brazil (Measures 1988a). Important fish intermediate hosts of E. ignotus in North America are livebearers (Poeciliidae), especially the eastern mosquitofish (Gambusia holbrooki), sunfish (Centrarchidae), and killifish (Fundulus heterocl*tus). By contrast, piranhas (Serrasalmus nattereri) are important intermediate hosts in South America
291
Podicipediformes
E. tubifex
Ukraine
E. tubifex E. sp. E. sp. E. ignotus E. sp. E. tubifex E. sp. E. tubifex E. tubifex E. sp. E. tubifex E. sp. E. sp.
Florida, USA E. tubifex Nova Scotia, Canada E. tubifex Ontario, Canada E. tubifex
E. tubifex E. tubifex E. tubifex
E. tubifex
Ukraine Europe Finland Russia
E. sp. E. tubifex E. sp. E. tubifex
Species
Pennsylvania, USA Europe Europe Russia
Locality
Finland Russia Russia Russia Europe England Europe Yugoslavia Japan Japan Great Crested Grebe (Podiceps cristatus) Yugoslavia Europe Russia
Ruff (Philomachus pugnax) Mew Gull (Larus canus) Black-headed Gull (Larus ridibundus) Common Murre (Uria aalge) Murre (Uria sp.) species not given Little Grebe (Tachybaptus ruficollis)
Common Loon (Gavia immer)
Arctic Loon (Gavia arctica)
Jackass Penguin (Spheniscus demersus) Red-throated Loon (Gavia stellata)
Host species
N N N N N N N N N N N N N
N N N
N
N N N
N
C N N N
NG NG NG 1/1 NG NG NG 3/18 1/1 NG 1/12 NG NG
3/45 1/NG 1/25
NG
NG NG 1/NG
NG
NG NG NG 2/NG
Number infected/ Number of Type examinations
fa*gerholm (1979) Skrjabin (1920)* Shigin (1961)* Morish*ta (1930) Jagerskiold (1909b) Measures (1988a) Jagerskiold (1909b) Brglez (1980) Murata et al. (1997) Yamaguti (1935)* Brglez (1980) Jagerskiold (1909b) Shigin (1957)* (continues)
Winsor (1948) Measures (1988a) Jagerskiold (1909b) Kontrimavichus and Bakhmet’eva (1960)* Smogorzhevskaya (1954, 1959,1962)* Measures (1988a) Karmanova (1986) Kontrimavichus and Bakhmet’eva (1960)* Smogorzhevskaya (1954, 1959, 1962)* Forrester and Spalding (2003) USNPC† # 87045 Measures (1988a)
Reference October 15, 2008
Charadriiformes
Sphenisciformes Gaviiformes
Host order
Table 16.1. Avian hosts of Eustrongylides spp.
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Black-faced Cormorant (Phalacrocorax fuscescens) Cormorant (Phalacrocorax sp.) species not given
Double-crested Cormorant (Phalacrocorax auritus) Little Pied Cormorant (Phalacrocorax melanoleucos) Pygmy Cormorant (Phalacrocorax pygmaeus) Long-tailed Cormorant (Phalacrocorax africanus) Great Cormorant (Phalacrocorax carbo)
Horned Grebe (Podiceps auritus)
Host species
E. excisus E. ignotus
South America
E. excisus
Ukraine Australia
E. excisus E. excisus E. excisus E. sp. E. ignotus E. tubifex
Europe Australia East Arabia Europe New Zealand Ukraine
E. ignotus N E. excisus N E. sp. N E. excisus N E. sp. N&E
New Zealand Australia Europe Romania Uganda
N
N
N
N N N N N N
N N N E
E. excisus E. excisus E. sp. E. tubifex
Romania Russia Russia Ontario, Canada
N N
N
E. sp. E. excisus
E. sp.
Species
Russia Ukraine
Ukraine
Locality
NG
NG
NG
NG NG NG NG — NG
NG NG NG NG 4
NG NG NG 8/10
NG NG
NG
Number infected/ Number of Type examinations
Caballero (1982)
Jagerskiold (1909b) Johnston and Mawson (1941) Measures (1988a) Jagerskiold (1909b) Dickinson (1951)* Smogorzhevskaya (1954, 1959,1962)* Smogorzhevskaya (1954, 1962, 1964)* Johnston and Mawson (1941)
Measures (1988a) Johnston and Mawson (1941) Jagerskiold (1909b) Ciurea (1938)* Paperna (1974)
Smogorzhevskaya (1954, 1962, 1964)* Kosupko (1963)* Smogorzhevskaya (1954, 1962, 1964)* Ciurea (1938)* Karmanova (1986) Nikol’skaya (1939)* Measures (1988a)
Reference
October 15, 2008
Pelecaniformes
Host order
Table 16.1. (Continued)
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Gray Heron (Ardea cinerea) Great Blue Heron (Ardea herodias)
293
(continues)
Forrester and Spalding (2003)
E. sp. N 1/4 Forrester and Spalding (2003) E. sp. N 2/23 Dyer et al. (2002) E. sp. N NG Brglez (1981) E. ignotus N 5/5 Ziegler et al. (2000) E. ignotus N 45/225 Spalding et al. (1993) E. ignotus N 2/25 Winterfield and Kazacos (1977) E. ignotus N 1/NG Locke (1961) E. sp. N NG Measures (1988a) E. sp. N NG Measures (1988a) E. ignotus N 1/NG Bowdish (1948) E. ignotus N NG Measures (1988a) E. sp. N 8/23 Cooper et al. (1978a) E. tubifex N 3/68 Measures (1988c) E 1/1 Measures (1988c) E. tubifex E. ignotus N NG Measures (1988a) E. sp. N NG Evans (1990) E. ignotus N 1/NG Chapin (1926) E. sp. N NG Measures (1988a) E. ignotus E. sp. NG Measures (1988a)
1/6
Florida, USA Puerto Rico Yugoslavia Delaware, USA Florida, USA Indiana, USA Maryland, USA Maryland, USA New Jersey, USA New Jersey, USA New York, USA Ohio, USA Ontario, Canada Ontario, Canada Virginia, USA Virginia, USA Washington, DC, USA Washington, DC, USA Washington, DC, USA
N
Forrester and Spalding (2003) Measures (1988a) Rego and Vicente (1988) Johnston and Mawson (1941) Jagerskiold (1909a) Jagerskiold (1909a)
E. sp.
1/1 NG NG — NG NG
Florida, USA
N N N N N N
E. ignotus E. ignotus E. ignotus E. excisus E. sp. E. sp.
Florida, USA Florida, USA Brazil Australia Sudan Sudan
October 15, 2008
Ciconiiformes
Pink-backed Pelican (Pelecanus rufescens) American White Pelican (Pelecanus erythrorhynchos) Brown Pelican (Pelecanus occidentalis)
Darter (Anhinga melanogaster)
Anhinga (Anhinga anhinga)
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Host order
294
Little Egret (Egretta garzetta) Chinese Pond-heron (Ardeola bacchus) Cattle Egret (Bubulcus ibis) Green Heron (Butorides virescens)
Snowy Egret (Egretta thula)
Little Blue Heron (Egretta caerulea)
Tricolored Heron (Egretta tricolor)
Florida, USA Louisiana, USA Maryland, USA Delaware, USA Texas, USA Ohio, USA Russia Brazil Brazil Florida, USA Florida, USA Delaware, USA California, USA Delaware, USA Florida, USA Florida, USA Delaware, USA Texas, USA California, USA Rhode Island, USA China China Taiwan Florida, USA E. ignotus E. sp. E. ignotus E. ignotus E. sp. E. sp. E. sp. E. ignotus E. ignotus E. ignotus E. ignotus E. ignotus E. ignotus E. ignotus E. ignotus E. ignotus E. ignotus E. sp. E. sp. E. sp. E. sp. E. excisus E. excisus E. ignotus
E. ignotus E. sp.
Brazil Sudan
Goliath Heron (Ardea goliath) N N N N N N N N C N E N C N N N N N N N N N N N
C —
N
N
94/427 5/5 2/NG 2/NG 8/10 4/36 NG NG 30/42 49/472 3/3 2/4 NG 1/7 25/360 82/439 50/53 9/9 3/15 1/10 NG NG 1/NG 1/11
7/9 —
NG
—
Number infected/ Number of Type examinations Windingstad and Swineford (1981) Rego and Vicente (1988); Measures (1988a) Pinto et al. (2004) Jagerskiold (1909a); Measures (1988a) Spalding et al. (1993) Roffe (1988) Locke (1961) Wiese et al. (1977) Franson and Custer (1994) Cooper et al. (1978b) Dubinin and Dubinina (1940) Rego and Vicente (1988) Pinto et al. (2004) Spalding et al. (1993) Spalding et al. (1994) Wiese et al. (1977) Measures (1988a) Wiese et al. (1977) Spalding et al. (1993) Spalding et al. (1993) Wiese et al. (1977) Franson and Custer (1994) Franson and Custer (1994) Franson and Custer (1994) Wu and Liu (1943)* Hoeppli et al. (1929)* Sugimoto (1933)* Spalding et al. (1993)
Reference
October 15, 2008
Great Egret (Ardea alba)
E. ignotus
Brazil
E. ignotus
Species
Cocoi Heron (Ardea cocoi)
Locality Wisconsin, USA
Host species
Blue Heron (species not given)
Table 16.1. (Continued)
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295
Maguari Stork (Ciconia maguari) Marabou Stork (Leptoptilos crumeniferus)
Rufescent Tiger-Heron (Tigrisoma lineatum) Pinnated Bittern (Botaurus pinnatus) American Bittern (Botaurus lentiginosus) Wood Stork (Mycteria americana)
E. ignotus E. ignotus E. sp. E. ignotus (l) E. ignotus E. ignotus E. sp. E. sp.
Brazil Ohio, USA Florida, USA Brazil Brazil Sudan Uganda
E. ignotus E. sp. E. excisus E. ignotus E. ignotus E. ignotus E. ignotus E. sp. E. sp. E. sp. E. excisus E. sp. E. ignotus E. ignotus E. ignotus
Brazil California, USA China Delaware, USA Florida, USA Maryland, USA New Jersey, USA Ohio, USA Rhode Island, USA Russia Taiwan Texas, USA Washington, DC, USA Florida, USA Brazil Brazil
E. ignotus
Brazil
NG
15/35 1/15 NG 2/4 1/4 NG 1/NG 23/36 1/10 NG NG 1/8 1/NG 1/6 NG
NG
Rego and Vicente (1988); Schaffer et al. (1990) Pinto et al. (2004) Franson and Custer (1994) Hoeppli et al. (1929)* Wiese et al. (1977) Spalding et al. (1993) Measures (1988a) Bowdish (1948) Cooper et al. (1978a) Franson and Custer (1994) Dubinina (1937) Sugimoto (1933)* Franson and Custer (1994) Cram (1933) Spalding et al. (1993) Rego and Vicente (1988) and Schaffer et al. (1990) Rego and Vicente (1988)
N C N N N
3/15 3/10 NG NG 1/6
(continues)
Forrester and Spalding (2003) Pinto et al. (2004) Vicente et al. (1995) Jagerskiold (1909a) Moriearty et al. (1972)
N NG Rego and Vicente (1988) N 1/NG Cooper et al. (1978a)
N
C N N N N N N N N N N N N N N
N
October 15, 2008
Yellow-crowned Night-Heron (Nyctanassa violacea)
Black-crowned Night-Heron (Nycticorax nycticorax)
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Eurasian Spoonbill (Platalea leucorodia)
Black-headed Ibis (Threskiornis melanocephalus) Bare-faced Ibis (Phimosus infuscatus) White Ibis (Eudocimus albus) Glossy Ibis (Plegadis falcinellus)
Host species
296 Eustrongylides sp. Eustrongylides ignotus Eustrongylides ignotus Eustrongylides sp. Eustrongylides tubifex Eustrongylides tubifex Eustrongylides sp. Eustrongylides tubifex Eustrongylides sp. Eustrongylides sp. Eustrongylides excisus Eustrongylides sp. Eustrongylides sp. Eustrongylides sp. Eustrongylides sp. Eustrongylides sp. Eustrongylides tubifex
Eustrongylides sp. Eustrongylides sp.
Kazakhstan Russia
Eustrongylides ignotus (l) Eustrongylides sp. Eustrongylides sp. Eustrongylides sp.
Florida, USA Yugoslavia Russia Ukraine Eustrongylides sp.
Eustrongylides ignotus
Brazil
Azerbaidzhan
Eustrongylides excisus
Species
India
Locality
N N N N E E N E N N NG N N N N N E
N N
N
N N N N
N
N
NG 1/136 1/129 NG NG 5/74 NG 4/11 NG NG NG NG NG NG NG NG NG
NG NG
NG
5/171 1/2 NG NG
NG
NG
Number infected/ Number of Type examinations
Skryabin (1916, 1923)* Dubinin and Dubinina (1940)* Graber (1975) Sepulveda et al. (1994) Spalding and Forrester (1993) Madsen (1952) Fastzkie and Crites (1977) Cooper et al. (1978a) Ryzhikov (1950)* Measures (1988c) Madsen (1952) Sugimoto (1932)* Sugimoto (1931)* Ryzhikov (1950)* Ryzhikov (1950)* Lyubimov (1926)* Measures (1988a) Ryzhikov (1963)* Measures (1988a)
Rego and Vicente (1988) and Schaffer et al (1990) Spalding et al. (1993) Brglez (1980) Dubinin and Dubinina (1940) Smogorzhevskaya (1954, 1962, 1964)* Feizullaev (1962, 1963)*
Ali (1971)
Reference
October 15, 2008
African Spoonbill (Platalea alba) Ethiopia Roseate Spoonbill (Platalea ajaja) Florida, USA Florida, USA Anseriformes Greylag Goose (Anser anser) Denmark Mallard (Anas platyrhynchos) Ohio, USA Ohio, USA Georgia (former USSR) Domestic White Pekin (Anas Ontario, Canada platyrhynchos) Denmark Japan Taiwan Northern Pintail (Anas acuta) Georgia (former USSR) Garganey (Anas querquedula) Georgia (former USSR) USSR Northern Shoveler (Anas clypeata) The Netherlands Tufted Duck (Aythya fuligula) Russia White-winged Scoter (Melanitta Alaska, USA fusca)
Host order
Table 16.1. (Continued)
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297 Eustrongylides tubifex Eustrongylides sp. Eustrongylides excisus Eustrongylides sp. Eustrongylides sp.
N N N N N
NG 1/2 1/NG 3/3 NG
NG
Tuggle and Schmeling (1982) Oshmarin (1963)* Ciurea (1938)* Acosta et al. (1988) Rudolphi (1819)* (see Measures (1988a))
Shillinger (1936)
l, identification based upon larval nematode; N, native or wild captured; C, captive; E, experimental infection; NG, no prevalence data given; —, reference not seen or not in translated abstract. * Information taken from Karmanova (1986), original paper not seen. † United States National Parasite Collection in Beltsville, MD, USA.
Gruiformes Passeriformes
Falconiformes
Measures (1988a) Jagerskiold (1909b) Measures (1988c)
N
NG NG 7/10
Jagerskiold (1909b) Smogorzhevskaya (1964)*
Eustrongylides tubifex
N N E
NG NG
N 3/10 Forrester and Spalding (2003) N 36/36 Locke et al. (1964) N 6/6 Cooper et al. (1978a) N 2/50 Measures (1988c) E 2/2 Measures (1988c) E NG Measures (1988c) N NG Jagerskiold (1909b) N 22/106 Measures (1988c) E 27/34 Measures (1988c) N 5/435 Measures (1988c)
Eustrongylides tubifex Eustrongylides sp. Eustrongylides tubifex
Germany Europe Ontario, Canada
N N
Eustrongylides sp. Eustrongylides sp. Eustrongylides sp. Eustrongylides tubifex Eustrongylides tubifex Eustrongylides tubifex Eustrongylides sp. Eustrongylides tubifex Eustrongylides tubifex Eustrongylides tubifex
Eustrongylides sp. Eustrongylides tubifex
Europe Ukraine
October 15, 2008
Florida, USA Virginia, USA Ohio, USA Ontario, Canada Ontario, Canada Common Merganser (Mergus Germany (merganser) Europe Ontario, Canada Ontario, Canada New Brunswick, Canada Merganser (species not given) British Columbia, Canada Bald Eagle (Haliaeetus leucocephalus) Alaska, USA Greater Spotted Eagle (Aquila clanga) Russia Lesser Spotted Eagle (Aquila pomarina) Russia Eurasian Coot (Fulica atra) Spain Eurasian Nutcracker (Nucifraga NG caryocatactes)
Hooded Merganser (Lophodytes cucullatus) Red-breasted Merganser (Mergus serrator)
Long-tailed Duck (Clangula hyemalis) Common Goldeneye (Bucephala clangula) Smew (Mergellus albellus)
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Parasitic Diseases of Wild Birds
(Wiese et al. 1977; Hirshfield et al. 1983; Weisberg et al. 1986; Rego and Vicente 1988; Coyner et al. 2002). E. excisus has the widest geographic distribution and is a common parasite of cormorants and species of Ciconiiformes in Asia, although infections have been recorded in a few other species of Pelecaniformes from Australia and Europe. It has not yet been reported from the Americas. The most important fish intermediate hosts are gobies (Neogobius kessleri and Neogobius melanostomus) and the Caspian roach (Rutilus rutilus caspicus) (Karmanova 1986). Other secondary reservoir hosts include predatory fish, especially perch (Perca fluviatilis), and frogs and snakes. Eustrongylides tubifex is found most commonly in mergansers and cormorants, and occasionally in species of Ciconiiformes in North America, but also causes lesions in loons and grebes in Europe and Asia. There are only two reports of mortality due to E. tubifex, and these involved mergansers in British Columbia and Virginia (Shillinger 1936; Locke et al. 1964). Records for E. tubifex in North America south of Ohio are infrequent. Eustrongylides ignotus was the only species found in a survey of species of Ciconiiformes in Florida (Spalding et al. 1993). However, a few Common Loons (Gavia immer) were found infected with E. tubifex in Florida (Forrester and Spalding 2003), but it is likely that they acquired the infections in northern North America where they breed. Other than a single record of an infected Cocoi Heron in Brazil (Measures 1988a), most other records in South and Central America are for E. ignotus (Rego and Vicente 1988). A similar pattern is evident in Europe and Asia and most records of E. tubifex are from northern latitudes. Pumpkinseed (Lepomis gibbosus) and yellow perch (Perca flavescens) serve as important intermediate hosts in North America (Measures 1988b). There is some evidence that raptors may act as definitive hosts of Eustrongylides, but such reports are not very common. Tuggle and Schmeling (1982) reported Eustrongylides spp. in a Bald Eagle (Haliaeetus leucocephalus) from Alaska, and these were later identified as gravid females of E. tubifex (J. R. Lichtenfels, personal communication). Lesions from one of two Greater Spotted Eagles (Aquila clanga) from Russia are suggestive of a mature eustrongylid infection (Oshmarin 1963). Unfortunately, the species and stage of these nematodes were not given. ETIOLOGY Eustrongylides spp. are large, red, grossly visible nematodes in the order Enoplida, superfamily Dioctophymatoidea, and family Dioctophymidae. They are distinguished from other members of the superfamily by the lack of a posterior sucker. Adults measure
about 4–15 cm or greater in length and 1–4 mm in diameter (Measures 1988a). The taxonomy of the genus Eustrongylides was reviewed by Karmanova (1986) and revised by Rautela and Malhotra (1984), Fastzkie and Crites (1977), and recently by Measures (1988a) who recognized three species: Eustrongylides tubifex (synonym = E. perpapillatus), E. excisus (synonyms = E. formosensis, E. indicus, E. phalacrocoracis, E. plotinus, E. tricolor, and E. excisus amoyensis), and E. ignotus. An additional description of E. ignotus was given by Rego and Vicente (1988). Distinguishing features include characteristics of the labial papillae around the mouth and the perimeter of the caudal sucker of the male and are illustrated by Measures (1988a) (Figures 16.2 and 16.3). A number of other poorly defined species were listed by Measures (1988a) as species inquirendae. These include E. africanus, E. mergorum, E. papillosus, E. rubrum, E. sinicus, E. spinispiculum, and those described from larvae in fish—E. acrochordi, E. gadopsis, and E. wenrichi. EPIZOOTIOLOGY Our understanding of the epizootiology of Eustrongylides, although initiated in the early 1900s, still remains very incomplete despite the significant harm these worms cause in birds and fish. The relatively recent discovery of an association between nutrient pollution and increases in the prevalence of these parasites appears to be fundamental to understanding the epizootiology of these organisms. Factors that must come together for an epizootic to occur include eutrophic wetlands that support high densities of fish, attraction of infected birds to feed on the fish, and nutrientpolluted sites that provide habitat for the appropriate oligochaete intermediate hosts. Thus, the ultimate initiator of this system is nutrient input. These characteristics are important for transmission of E. ignotus in Florida and have been suggested for the other two species of Eustrongylides (Karmanova 1986; Measures 1988b; and Coyner et al. 2002). Life History Transmission to the definitive host is by ingestion of infected fish or, in some cases, by ingestion of a paratenic host. The life history of all three species of Eustrongylides is believed to involve piscivorous avian definitive hosts infected with fourth-stage (L4) larvae and mature, sexually reproducing adults, oligochaete first intermediate hosts infected with first(L1), second- (L2), and third-stage (L3) larvae, and fish second intermediate hosts infected with L3 and L4 larvae (Karmanova 1986; Anderson 2000). Coyner et al. (2003a), however, were able to infect fish directly with larvated eggs of E. ignotus without passage through
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a
b Figure 16.2. Diagnostic features of the cranial end of Eustrongylides spp. (a) Eustrongylides ignotus and (b) Eustrongylides tubifex. Note the relatively larger inner circle papillae (narrow arrow) of E. ignotus. E. tubifex is the only species in which the inner labial papillae are smaller than the outer (wide arrow). Some lateral field papillae (small arrowheads) and ventral papilla (vp) are also visible. Scale bar = 100 μm (insets = 10 μm). From Measures (1988a), with permission of the Canadian Journal of Zoology.
a
b
c Figure 16.3. Diagnostic features of the caudal end of male Eustrongylides spp. (a) Eustrongylides ignotus, (b) Eustrongylides tubifex, and (c) Eustrongylides excisus. Note the projections from the ruffled edge (large arrow) on the rim of E. tubifex, and the wide cuticular hem on the outer perimeter of the sucker (wide arrow) of E. ignotus. Small white arrows and black arrowheads indicate groups of caudal papillae. The dorsal wall of the posterior part of the rectum of E. ignotus bears spines and protrudes from anus (inset). Note the ventral cleft in the drawing of E. excisus. Scale bar = 100 μm (insets = 10 μm). From Measures (1988a), with permission of the Canadian Journal of Zoology.
299
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Parasitic Diseases of Wild Birds
Figure 16.4. Life cycle of Eustrongylides sp. (1) Definitive host, a piscivorous bird, consumes infected fish. The encysted fourth-stage larva (L4) burrows through the stomach wall and matures to an adult. Adults mate and eggs develop in the uterus of the female. (2) Eggs are produced and pass out in feces. The zygote cleaves. (3) The egg matures to the infective stage containing a first-stage larva (L1). (4) A fish intermediate host consumes an egg containing an L1. The L1 encysts and matures to an infective L4, or (5) an egg containing an L1 is consumed by an oligochaete intermediate host and matures to an infective third-stage larva (L3). (6) A fish intermediate host consumes an oligochaete intermediate host containing an L3. The larvae encyst in the fish and develop to an L4. (7) A definitive host consumes a fish containing an encysted L4. (8) A predatory fish or other paratenic or accidental host consumes a fish intermediate host and the L4 encysts. The short cycle can be completed in 115 days, the long cycle takes >330 days. oligochaetes (Figure 16.4). This mode of transmission shortens the life cycle by 78–156 days, but its importance in natural cycles or in other species is unknown. It could be significant in situations where rapid infection of the fish intermediate host is critical or where overwintering of larvae is not necessary. With the exception of E. excisus, the survival time of adult worms in the avian host is not known. Developmental times of the parasite in the environment and in oligochaete, fish, and avian hosts have been determined by experimental studies (Table 16.2). For E. tubifex, the first appearance of L3 larvae in oligochaetes takes longer than for E. ignotus. Multiple infections, with an average intensity of 5.5 larvae in each oligochaete worm (Limnodrilus hoffmeisteri), have been documented experimentally for E. tubifex.
Like E. ignotus, the development time is temperature dependent. The length of time for development in a fish to the L4 stage is not known for E. tubifex and E. excisus. Adult parasites die and nodular lesions resolved within 30 days (Measures 1988c). Under experimental conditions, it takes 5–5.5 months for the larva E. excisus to develop to infective L3 larvae in oligochaetes, but Karmanova (1986) suspected that natural transmission might occur faster based upon evidence of reinfection of avian hosts in the Volga Delta. The natural life cycle from egg to death of adult worms is reported to be 8–9 months in Great Cormorants (Phalacrocorax carbo) (Karmanova 1986). Overwintering of larval E. excisus is thought to occur in fish since cormorants arrive in the Volga Delta in the spring with no evidence of infection. Young nematodes
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301
Eustrongylidosis Table 16.2. The range of development times in days for the life history stages of Eustrongylides ignotus, Eustrongylides tubifex, and Eustrongylides excisus as determined through experimental studies.
Eustrongylides ignotus Eustrongylides ignotus Eustrongylides Eustrongylides excisus‡ without oligochaete* with oligochaete tubifex† Stage Egg in feces to infective L1 in egg L2 to infective L3 in oligochaete L3 (or L2) to L4 in fish L4 to egg in bird feces Days to complete life cycle
Min
Max
Min
Max
Min
Max
Min
Max
17
28
17
28
23
26
22
—
35
77
70
109
150
165
84 20 121
105 23 156
127 20 199
184 23 312
— 10 —
— 17 —
— 12 —
— 14 —
L1, first-stage larvae; L2, second-stage larvae; L3, third-stage larvae; L4, fourth-stage larvae. * From Coyner et al (2003a), the first column omits the oligochaete phase, the second column includes it. Third-stage larvae (L3) can survive up to 284 days in an oligochaete. † From Measures (1988c), infection of fish with oligochaetes containing third-stage larvae (L3) was not successful. Larvae could be maintained in captive fish for 18 months (Crites 1982). ‡ From Karmanova (1986), development of second- to third-stage larvae (L2–L3) is expected to occur faster in a natural system. can be found in cormorants at the end of May to early June, and sexually mature nematodes are found by the end of June. Immature nematodes were recovered from avian hosts throughout the summer, indicating that reinfection was occurring. Cormorants appeared to be free of infection when they migrated for the winter (Karmanova 1986). The range of oligochaete intermediate and/or paratenic hosts is also wide. Identification is complicated both by the difficulty in finding naturally infected oligochaetes and also by the necessity to pass larval nematodes through fish intermediate and avian definitive hosts in order to identify these stages to species. Oligochaetes infected naturally with larval nematodes have not been documented with certainty, but one was suspected to be parasitized by E. tubifex (Lichtenfels and Stroup 1985). Evidence for inclusion of oligochaetes in the life cycle is based almost entirely on experimental work. In Florida, L. hoffmeisteri seems to be the most important, if not the only oligochaete intermediate host for E. ignotus (Spalding et al. 1993; Coyner et al. 2002, 2003a). Both L . hoffmeisteri and Tubifex tubifex can be experimentally infected with E. tubifex (Measures 1988d). Based on experimental studies, the oligochaete intermediate hosts of E. excisus are Limnodrilus sp., T. tubifex, and Lumbriculus variegatus (Karmanova 1986). There is evidence that larval Eustrongylides (L4, and possibly L3) can be transmitted from fish to fish and
to amphibians and reptiles by predation. This has been demonstrated in the laboratory for E. ignotus (Coyner et al. 2003a), E. tubifex (Measures 1988b), and is presumed to occur in the field for E. excisus (Karmanova 1986). This route of transmission allows the parasite to accumulate in a paratenic or reservoir host and may account for high prevalence and intensity of infection in larger predatory fish (Karmanova 1986; Measures 1988b; Coyner et al. 2002). In an experimental setting, infected fish are more likely to be captured and are more quickly captured by predatory fish than uninfected fish (Coyner et al. 2001). Larval Eustrongylides have been collected from various mammals, reptiles, and amphibians (Table 16.3), but these are likely aberrant hosts. In some cases, these may be small enough to be consumed by the definitive host birds and thereby act as paratenic hosts. There is a single report of sturgeon (Acipenser gueldenstaedtii) with encysted mature E. excisus that contained eggs, indicating that it may be possible for fish also to act as definitive hosts (Mikailov et al. 1992). There have been no other reports of adult Eustrongylides spp. in fish, but they could be overlooked since histologic examination of encysted nematodes is not routinely done. Environmental Limitations Eutrophication and nutrient input are significant environmental factors affecting transmission of
302
Fer-de-lance (Bothrops atrox) Copperhead (Agkistrodon contortrix) Javan wart snake (Acrochordus javanicus) Jararaca (Bothrops jararaca) Green anaconda (Eunectes murinus) Coral snake (Micrurus sp.)
Grass snake (Natrix tessellata) Blue Racer (Coluber constrictor) Whip snake (Masticophis flagellum) Pine snake (Pituophis melanoleucus)
Indigo snake (Drymarchon corais) Garter snake (Thamnophis sirtalis)
Reptiles Brown water snake (Nerodia taxispilota) Northern water snake (Nerodia sipedon) Florida, USA Virginia, USA Pennsylvania, USA Florida, USA Maryland, USA Pennsylvania, USA Russia Pennsylvania, USA Pennsylvania, USA Pennsylvania, USA Pennsylvania, USA Pennsylvania, USA Pennsylvania, USA Australia Brazil Brazil Brazil
Maryland, USA Oregon, USA Russia Ontario, Canada Japan Maryland, USA Maryland, USA Brazil Maryland, USA New Jersey, USA New York, USA
Locality
E. sp. E. sp. E. sp. E. sp. E. sp. E. sp. E. excisus E. sp. E. sp. E. sp. E. sp. E. sp. E. sp. E. sp. E. ignotus E. ignotus E. ignotus
E. sp. E. sp. E. excisus (l) E. sp. E. sp. E. sp. E. sp. E. ignotus E. sp. E. ignotus E. sp.
Species*
N N C N E C N C C C C C C N N N N
N N N N N E E E N N N
Type
1/5 18/18 NG 1/12 20/25 NG NG NG NG NG NG NG NG 1/8 NG NG NG
1/NG 12/29 6/15 1/1 NG NG 6/6 7/10 2/NG 1/NG 1/NG
Number infected/ Number of examinations
Ferenc et al. (1986) Bursey (1986) Winsor (1948) Foster et al. (2000) Lichtenfels and Lavies (1976) Winsor (1948) Karmanova (1986) Winsor (1948) Winsor (1948) Winsor (1948) Winsor (1948) Winsor (1948) Winsor (1948) Jones (1978) Rego and Vicente (1988) Rego and Vicente (1988) Rego and Vicente (1988)
Abram and Lichtenfels (1974) Hoberg et al. (1997) Shchupakov (1936)* Gibson and McKiel (1972) Morish*ta (1923) von Brand and Cullinan (1943) Shirazian et al. (1984) Barros et al. (2004) Gunby (1982); Guerin et al. (1982) Eberhard et al. (1989) Arias (1989)
References
October 15, 2008
Human (hom*o sapiens)
Caspian seal (Phoca caspica) Muskrat (Ondatra zibethica) Raccoon Dog (Nyctereutes procyonoides) “Rat” New Zealand white rabbit (Oryctolagus cuniculus)
Mammals River otter (Lutra canadensis)
Host
Table 16.3. Paratenic and accidental hosts of Eustrongylides spp.
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303
E. ignotus (l) E. tubifex E. sp. E. sp. E. sp. E. sp. E. sp. E. excisus E. sp. E. sp. E. ignotus E. sp. E. sp. E. sp. E. sp.
Florida, USA Ohio, USA Maryland, USA Maryland, USA Louisiana, USA Nevada, USA Cuba Russia Turkey Mexico Brazil California, USA Maryland, USA Maryland, USA Louisiana, USA
N E E N E N N N N N N N E E N
N N N E N E E E E N N 3/11 NG 7/7 NG 20/25 – – NG 3/258 10/80 NG 1/230 1/1 1/1 NG
NG NG NG NG NG NG NG 3/3 1/1 3/115 NG Spalding and Forrester (1991) Cooper et al. (1978b) von Brand (1944) von Brand (1944) Modzelewski and Culley (1974) Babero and Golling (1973) Coy-Otero and Martinez (1987) Karmanova (1986) Dusen and Oz (2006) Lezama and Sarabia (2002) Rego and Vicente (1988) Kuperman et al. (2004) von Brand (1944) von Brand (1944) Panesar and Beaver (1979)
Rego and Vicente (1988) Rego and Vicente (1988) Rego and Vicente (1988) Cooper et al. (1978b) Caballero (1982) Cooper et al. (1978b) Cooper et al. (1978b) von Brand (1944) von Brand (1944) Goldberg et al. (1991) Rego and Vicente (1988)
l, identification based upon larval nematode; N, native or wild captured; C, captive; E, experimental infection; NG, no prevalence data given; –, reference not found or not in translated abstract. * Information taken from Karmanova (1986), original paper not seen.
Bigfoot leopard frog (Rana megapoda) Smokey jungle frog (Leptodactylus pentadactylus) African clawed frog (Xenopus laevis) Mudpuppy (Necturus maculosus) Three-toed amphiuma (Amphiuma tridactylum)
Lake frog (Rana ridibunda)
E. ignotus E. ignotus E. ignotus E. tubifex E. sp. E. tubifex E. tubifex E. sp. E. sp. E. sp. E. ignotus
Brazil Brazil Brazil Ohio, USA Texas, USA Ohio, USA Ohio, USA Maryland, USA Maryland, USA Paraguay Brazil
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Bullfrog (Rana catesbeiana)
Blandings turtle (Emys blandingi) Spiny softshell turtle (Apalone spinifer) Painted turtle (Chrysemys picta) American alligator (Alligator mississippiensis) Paraguayan caiman (Caiman yacare) Spectacled caiman (Caiman crocodilus) Amphibians Pig frog (Rana grylio) Northern leopard frog (Rana pipiens)
Cazadora brown-lined snake (Dryadophis bifossatus) Mussurana (Clelia clelia) Graceful brown snake (Rhadinea merremii) Snapping turtle (Chelydra serpentina)
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Eustrongylides. Infected fish are never found in undisturbed, naturally oligotrophic habitats in Florida, even under the naturally eutrophic conditions that seasonally occur under wading bird colonies (Spalding et al. 1993; Coyner et al. 2002). The physical and chemical conditions associated with sites containing infected fish are dramatically different from more natural, usually oligotrophic sites. Sites containing infected fish generally have a long-term history of nutrient input such as sewage outflow or storm water runoff. The changes are characteristic of eutrophic water bodies and include physical alterations of the substrate, higher densities of fish and oligochaetes, decreased dissolved oxygen, increased total nitrogen, total phosphorus and chlorophyll a in surface water, higher soil oxygen demand and total phosphorus in sediment, larger grain size, and higher percentage of emergent vegetation and grasses (Coyner et al. 2003b). Eggs of Eustrongylides spp. are tolerant of a wide range of temperatures (Karmanova 1986), but both development and development time may be affected by temperature, salinity, and desiccation. Development of the eggs of E. tubifex ceases at 0–15◦ C in the laboratory, but resume at higher temperatures (Measures 1988d). The eggs of E. ignotus are not tolerant of hypersaline (>33 ppt) water or desiccation, but develop normally in fresh or estuarine water (20 ppt) (Coyner et al. 2004). The range of fish intermediate hosts of Eustrongylides is very wide (Karmanova 1986; Measures 1988b; Coyner et al. 2002). Freshwater fish are much more commonly infected than marine fish, and there is some evidence that transmission of E. ignotus may not occur in saltwater in Florida (Spalding et al. 1993). Almost no infections are found in wading birds nesting in marine environments (Spalding et al. 1993). The few exceptions are colonies located close to freshwater foraging sites. This may be due to the distribution of the oligochaete intermediate hosts that are not able to reproduce in marine systems. Marine fish may become infected when visiting estuarine or freshwater sites, or when preying on infected fresh or estuarine fish that move into marine systems, leading to the low levels of infections that were found. There is some evidence that E. excisus may be more tolerant of marine environments, based on recovery of E. excisus from sturgeon in the southern portion of the Caspian Sea where higher salinities are more typical of ocean environments (Sattari and Mokhayer 2005). Prevalence and Intensity Prevalence in species of Ciconiiformes can be quite high and range from 11 to 100%, depending on species (Cooper et al. 1978a) (Table 16.1). In a large sample of birds that were collected by gunshot in Ohio, all
Red-breasted Mergansers (Mergus serrator) were infected. The species of eustrongylid was not identified, but Mallards (Anas platyrhynchos) experimentally fed with yellow perch from the area became infected with E. tubifex. Franson and Custer (1994) collected nestling Snowy Egrets (Egretta thula), Black-crowned Night-Herons, and Great Egrets (Ardea alba) in Texas, Rhode Island, and California. Overall, 31% of broods were infected with the highest prevalence in Texas (35%) where 100% of Snowy Egret, 80% of Great Egret, and 12% of Black-crowned Night-Heron broods were infected. A conservative estimate of 400 dead young birds for the year from eustrongylidosis was made for a colony of 900 pairs of Great Egrets in Louisiana (Roffe 1988). For E. ignotus, an overall prevalence of 14% was reported for all species of Ciconiiformes in Florida, with prevalences ranging from 0% in Cattle Egrets (Bubulcus ibis), a species that is not piscivorous, to 33% in Great Blue Herons (Spalding et al. 1993). It was estimated that 80% of nestlings died of eustrongylidosis in one colony. Prevalence was higher in juveniles and adults when compared with nestlings; however, infection was more likely to cause mortality in younger birds. In Great Egret, nestlings prevalences varied among years between 5 and 28% at a colony in central Florida (Spalding et al. 1994). Prevalence increased in fledgling Great White Herons (Ardea herodias occidentalis) when they dispersed from marine to freshwater sites in southern Florida (Spalding et al. 1993). Intensity of E. excisus in Great Cormorants and Pygmy Cormorants (Phalacrocorax pygmaeus) ranged from 3 to 62 worms per individual in a study to discover the life cycle of the parasite that was impacting commercial fisheries in Romania (Ciurea 1938). In Russia, 50% of young cormorants and 40% of adults were infected (Dubinin 1949). Prevalence of infection was similarly high in an earlier study, with 67% of adults and 40% of young infected (Nikol’skaya 1939). Increases in prevalence of Eustrongylides in infected fish have been reported by several researchers, suggesting that this parasite has become more common in recent years. Prevalence of E. tubifex in channel catfish (Ictalurus punctatus) from Lake Erie increased from 0% in 1939 (Bangham and Hunter 1939) to 15% by the 1970s (Cooper et al. 1978a). Similar increases in prevalence from 3 to 42% between 1930 and 1978 have been observed in fish collected at the same sites in Florida (Frederick et al. 1996). CLINICAL SIGNS Clinical signs in species of Ardeidae of all ages include ataxia, lethargy, depression, emaciation, and pale mucous membranes (Spalding and Forrester 1993;
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Ziegler et al. 2000). Little information is available about changes in clinical chemistry or hematology, although anemia and eosinophilia have been reported (Ziegler et al. 2000). In experimental studies, nestling Tricolored Herons (Egretta tricolor) that were infected at a few days of age consumed less food, regurgitated more frequently, and had lower bill and mass growth rates than uninfected control chicks (Spalding et al. 1994). Regurgitation or attempts to regurgitate may be associated with pain from parasite perforation (Measures 1988c; Spalding and Forrester 1993). Reports of clinical signs in other avian families are limited. Red-breasted Mergansers had severe nervous spasms of the head and neck and ocular twitching just before death (Locke et al. 1964). PATHOLOGY Gross Pathology Lesions in species of Ardeidae infected with E. ignotus have been described by several authors (Locke 1961; Roffe 1988; Schaffer et al. 1990; Spalding and Forrester 1993; Ziegler et al. 2000; Pinto et al. 2004). Larvae that perforate the ventricular portion of the stomach generally cause minor local hemorrhage. In very small chicks, there can be significant hemorrhage associated with the perforation. Larvae that have recently perforated the stomach can be seen either free in the coelom or just below the serosa. In subacute infections, parasites are surrounded by a thin, friable tube composed of fibrin and cellular debris of host origin and bacteria. At this stage, the parasites can easily be removed intact. The tubules in more chronic infections (Figure 16.5) are larger, prominent, firm, and are composed of granulomatous fibrosis. The tubules of adult nematodes, especially large females, are frequently twisted and contain 180◦ turns, making removal of intact adult females almost impossible. Fibrous tubules that adhere to the ventral abdominal wall may be visible or palpable on the exterior of the abdomen. Tubules communicate with the lumen of the stomach and rarely the small intestine through a raised portal with either the head or the tail of the worm protruding. Adhesions and penetrating tubules may also involve intestine, liver, airsacs, gall bladder, proventriculus, cloaca, abdominal wall, pancreas, and pericardium in decreasing frequency. The tubules remain when parasites die and gradually resolve to irregularly shaped, firm, often black nodules on the serosal surface of the stomach. In very severe chronic cases, the entire abdominal contents may be woven together with an inseparable mat of fibrous tubules. Bacteria cultured from the abdominal cavity and various organs can include a wide range of species typically found in the intestinal tract of fish-
Figure 16.5. Chronic eustrongylidosis in a Snowy Egret (Egretta thula). Note tubules (arrow) on the surface of the ventriculus (V). The smaller tubules are from a more recent infection. From Spalding and Forrester (1993), with permission of the Journal of Wildlife Diseases.
eating birds (Wiese et al. 1977; Roffe 1988; Spalding and Forrester 1993; Ziegler et al. 2000). Lesion appearance may be more dependent on host species than on parasite species. For example, a much more generalized and severe peritonitis occurs in nestling White Ibises (family Threskiornithidae) than in species of nestling Ardeidae (Spalding and Forrester 1993). Lesions similar to those caused by E. ignotus occur in Great Blue Herons and Common Loons infected with E. tubifex, but appear as nodular masses in the proventriculus of mergansers (Measures 1988c). Nothing is known about whether hosts respond immunologically to the parasite, the bacteria or gastric fluids, or to the tissue damage associated with the perforation. Measures (1988c) described the nodular or cystlike lesions found in the proventriculus of mergansers
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infected with E. tubifex to be 5 mm to 3 cm in diameter and projecting into the peritoneal cavity (Figure 16.6). The nematodes communicated with the lumen of the proventriculus. It is interesting to note that von Brand and Cullinan (1943) were not able to infect ducks and chicks per os and speculated that the parasites were killed in the grinding gizzard (ventriculus). That may explain why lesions in mergansers (with grinding gizzards) develop in the proventriculus and esophagus, whereas E. ignotus perforates the nongrinding, poorly muscled ventricular stomach in species of Ciconiiformes. It does not explain, however, why cystlike lesions occur in species of Pelecaniformes as they do with E. excisus in cormorants. Lesions caused by E. excisus in the proventriculus of cormorants are similar to those produced by E. tubifex (Ciurea 1938). Nematodes are coiled in a capsule that projects from the serosal surface like a pea with the head and tail ends protruding from the mucosal surface into the lumen. During the course of infection, the middle portion of the nematode quickly becomes covered with a connective tissue capsule. Dubinin (1949) also described nematodes in the proventriculus, between the glandular and muscular layers with the anterior and posterior ends in the lumen. More severe and fatal lesions were described by Locke et al. (1964) in Red-breasted Mergansers during a die-off in Virginia. Most had peritoneal hematomas with nematodes burrowing into and causing destruction of the liver, especially the left lobe, the proventriculus, and airsacs. Involvement of the lung, heart, and kidney was less severe. Microscopic Pathology Acute lesions in species of Ardeidae infected with E. ignotus are generally associated with larval stages and located in the tunica muscularis or below the serosa of the stomach (Figure 16.7) (Locke 1961; Roffe 1988; Spalding and Forrester 1993). Occasionally in very early infections, or in severely debilitated birds, no inflammation is present. More commonly and in more subacute lesions, the parasite is surrounded by fibrin with associated erythrocytes, granulocytes, and monocytes. Subserosal hematomas are frequent (Spalding and Forrester 1993). The parasites are easily distinguished from other nematode species and have prominent ventral cords, coelomyarian musculature, and a pseudocoelomic membrane (Figure 16.7). Chronic lesions are observed more frequently and are characterized by a parasite within the lumen of a tubule surrounded by cellular debris, free bacteria, and eustrongylid eggs. Often the cuticle remains, surrounding the nematode. In some cases, degenerate pieces of nematode are present. The tubule itself is composed of
a prominent layer of fibrous connective tissue lined by multinuclear giant cells on the luminal side. Fibrin can often be seen on the peritoneal surface. Chronology of experimental E. tubifex infections in Common Mergansers (Mergus merganser) has been described in detail by Measures (1988c). Fibrinous tubes on the serosal surface of the proventriculus are evident at 2 days postinfection. By 6 days postinfection, the tubules are surrounded by a fibrous connective capsule that contains inflammatory cells, bacteria, and erythrocytes. As these encapsulated lesions expand, they compress the proventricular glands. By 30 days postinfection, only resolving granulomas are present. We found no microscopic descriptions for infections with E. excisus.
DIAGNOSIS Diagnosis in species of live Ciconiiformes is possible. Palpation of the abdomen can be quite reliable in species of Ardeidae if the lesions are subacute to chronic (Spalding 1990). The tortuous tracts can also be seen sometimes on radiographs and by laproscopy or laparotomy. Necropsy remains the most common method of diagnosis. Once observed, the tubular serpininous tracts over the surface of the stomach and intestines and occasionally coursing through organs are pathognomonic. When the parasite dies, the tracts degenerate into black multifocal nodules adherent to the serosa, indicating a chronic resolved infection. The only possible confusion is with occasional perforating ascaroid nematodes or foreign object perforation, which never cause the characteristic serpiginous tracts of eustrongylidosis. These tracts have only been described thus far in birds of the order Ciconiiformes. Patent infections can be demonstrated by direct microscopic examination of feces or by fecal flotation techniques. This method frequently results in false negatives (Spalding 1990). When present, the eggs are highly crenulated, making the diagnosis relatively easy (Figure 16.8). Identification of the nematodes to species can be made by light microscopy by an experienced specialist, but is best made by examination of adult nematodes by scanning electron microscopy. Female specimens of E. ignotus are distinguished from those of E. tubifex by the larger size of the six inner circle labial papillae, but are difficult to differentiate from female specimens of E. excisus (Figure 16.2). Male specimens of E. ignotus have a caudal sucker that lacks a cuticular cleft, while a cuticular cleft is present in the caudal sucker of male specimens of E. excisus. Male specimens of E. tubifex have projections on the inner perimeter of the cuticular hem, while male E. ignotus do not (Figure 16.3).
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(a)
(b)
Figure 16.6. Gross lesions caused by Eustrongylides tubifex in a Common Merganser (Mergus merganser). (a) Note the large tumors on the serosal surface of the proventriculus. (b) The nodules protrude from the serosal surface. From Measures (1988c), with permission of the Canadian Journal of Zoology.
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Figure 16.7. Cross section of a fourth-stage larva in adipose tissue below the serosa of the ventriculus of a Great Blue Heron (Ardea herodias occidentalis). Note the large ventral chord (arrow), the coelomyarian musculature (m), pseudocoelomic membrane (arrowhead), and intestine (i). Bar = 100 μm. From Spalding and Forrester (1993), with permission of the Journal of Wildlife Diseases.
Additional electron micrographs of nematodes can be found in Measures (1988a) and Coyner et al. (2003a). Diagnosis in fish to the level of species is difficult. Larval stages of E. ignotus and E. tubifex are illustrated by Coyner et al. (2003a) and Measures (1988b), respectively. Larvae in fish can be distinguished from Dioctophyma renale by size, cephalic and reproductive morphology, and the tendency for E. ignotus to be associated with the mesentery rather than the musculature (Lichtenfels and Madden 1980; Measures and Anderson 1985; Measures 1988b). Small fish that are infected with larval Eustrongylides have swollen abdomens that affect appearance when swimming. In advanced infections, the coiled L4 larvae can be seen through the abdominal wall (Figure 16.9).
PUBLIC HEALTH AND DOMESTIC ANIMAL CONCERNS Humans can become infected and die from ingesting poorly cooked or raw infected fish, but only a few re-
ports exist (Table 16.3). Generally humans do not eat raw fish, or if they do, the parasites are easy to see if the flesh is carefully examined. Most reported cases are the result of fisherman eating baitfish that are usually collected from locations with high fish density and risk of infection with larval worms is high. In humans, the perforation of the intestinal tract is very painful. One patient underwent surgery for appendicitis, whereupon a larval eustrongylid was discovered in the peritoneal cavity (Gunby 1982; Arias 1989). There are no reports of infections in domestic animals, although consumption of raw fish by cats and dogs may lead to infection.
WILDLIFE POPULATION IMPACTS Eustrongylidosis is probably the most common cause of disease-related mortality in nestling wading birds from North America, but is frequently underreported. Similar mortality likely goes unnoticed throughout the range of E. ignotus and other species. Although
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Figure 16.8. Electron micrograph of the egg of Eustrongylides ignotus. Note the depressions. These depressions can also be seen at the light microscope magnification. From Coyner et al. (2003a), with permission of the Journal of Parasitology.
the prevalence of E. ignotus infections in species of Ardeidae is high, morbidity, mortality, and effects on recruitment of young birds into the breeding population are less well studied. Most mortality occurs within colonies that are visited rarely. Presence of dead nestlings is often not considered alarming because wading birds routinely lay and hatch more nestlings than will fledge. As a result, this group of parasites has gone relatively unnoticed as a cause of significant and increasing wildlife mortality at the population level. Eustrongylides tubifex causes rare sporadic mortality in mergansers. It is interesting that despite the availability of prevalence data and intensive life history work on these parasites, there is no indication that either E. tubifex or E. excisus have significant effects on
the health of their avian hosts. However, like E. ignotus in species of Ardeidae, such information may go unreported until an interest is taken in the health of the avian species involved. TREATMENT AND CONTROL Treatment of infected birds with anthelmintics has not been tried in any systematic fashion and is unlikely to be practical. Mortality of large numbers of nematodes within the host might do more harm than good. MANAGEMENT IMPLICATIONS The presence of nutrient pollution in waters used by wading birds increases risk for eustrongylidosis and
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Figure 16.9. Eastern mosquitofish (Gambusia holbrooki ) with a single eustrongylid larva (top) which has been dissected from the coelomic space and removed from its capsule (bottom). From Spalding et al. (1993), with permission of the Journal of Wildlife Diseases. ultimately nest failure. Nutrients increase fish density and thus indirectly increase the attraction of wading birds to a potentially hazardous site. They also enhance conditions for completion of the parasite life cycle. Reduction or elimination of nutrient release into water bodies that can be used by wading birds may be the most practical large-scale management option. Wading birds are attracted to such sites because increased productivity results in higher fish densities (Frederick and McGehee 1994). Coyner et al. (2003b) documented the decline of infected fishes in an urban watershed that coincided with the decline of nutrient pollution, suggesting that this may be an effective solution. Another approach can be to exclude or discourage birds from using sites with nutrient-rich polluted water by preventing access or by ensuring that wastewater releases occur in locations unlikely to be visited by wading birds.
ACKNOWLEDGMENTS We thank Garry Foster and Jaimie Miller for assistance with finding literature and producing the distribution map. We also thank Lena Measures for reading the manuscript and offering many helpful suggestions for improvement. LITERATURE CITED Abram, J. B., and J. R. Lichtenfels. 1974. Larval Eustrongylides sp. (Nematoda: Dioctophymatoidae) from otter, Lutra canadensis, in Maryland. Proceedings of the Helminthological Society of Washington 41:253. Acosta, I., S. Hernandez-Rodgiguez, S. Martinez-Cruz, and F. Martinez-Gomez. 1988. Parasitoses of aquatic birds in Cordoba. Erkrankungen der Zootiere 1988:327–331.
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Eustrongylidosis Ali, M. M. 1971. A new species of dioctophimid nematode, Eustrongylides indicus n. sp., from an Indian bird. Rivista di Parassitologia 32:47–50. Anderson, R. C. 2000. Nematode Parasites of Vertebrates. Their Development and Transmission, 2nd ed. CABI Publishing, New York. Arias, V. E. 1989. A worm in the peritoneum. Risks of eating raw fish and shellfish. Revista Espanola de las Enfermedades del Aparato Digestivo 76:669. Babero, B. B., and K. Golling. 1973. Some helminth parasites of Nevada bull frogs. Revista de Biologia Tropical 21:207–220. Bangham, R. V., and G. W. I. Hunter. 1939. Studies on fish parasites of Lake Erie. Distribution Studies. Zoologica 24:385–448. Barros, L. A., R. Tortelly, R. M. Pinto, and D. C. Gomes. 2004. Effects of experimental infections with larvae of Eustrongylides ignotus and Contracaecum multipapillatum in rabbits. Arquivo Brasileiro de Medicina Veterinaria e Zootecnia 56:325–332. Bowdish, B. S. 1948. Heron mortality caused by Eustrongylides ignotus. The Auk 65:602–603. Brglez, J. 1980. Some dioctophymid species (Dioctophymidae, Railliet, 1915) in birds from Yugoslavia. Acta Parasitologica Yugoslavica 11:889. Brglez, J. 1981. Incidence and pathogenicity in Yugoslavia in birds of nematodes of the family Dioctophymidae Railliet, 1915. Erkrankungen der Zootiere. Verhandlungsbericht des XXIII Internationalen Symposiums uber die Erkrankungen der Zootiere 24028:233–235. Bursey, C. R. 1986. Histological aspects of natural eustrongyloid infections of the northern water snake, Nerodia sipedon. Journal of Wildlife Diseases 22:527–532. Caballero, R. G. 1982. Nematoda. In Aquatic Biota of Mexico, Central America, and the West Indies, S.H. Hurlbert and A. Villalobos-Figueroa (eds). San Diego State University, San Diego, CA, pp. 101–120. Chapin, E. A. 1926. Eustrongylides ignotus Jagerskiold in the United States. Journal of Parasitology 13:86–87. Ciurea, J. 1938. Sur une infestation massive des poissons rapaces du las Greaca par les larves d’Eustrongylides excisus Jagerskiold. Grigore Antipa Hommage a son oeuvre. Bucaresti, pp. 149–162. (Original not seen, in Karmanova 1986) Cooper, C. L., J. L. Crites, and D. J. Springkle-Fastzkie. 1978a. Population biology and behavior of larval Eustrongylides tubifex (Nematoda: Dioctophymatida) in poikilothermous hosts. Journal of Parasitology 64:102–107. Cooper, C. L., J. L. Crites, and J. Sprinkle Fastzkie. 1978b. Experimental and natural infections of
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Eustrongylides sp. (Nematoda: Dioctophymatidae) in waterfowl and shore birds. Avian Diseases 22:790–792. Coyner, D. F., S. R. Schaack, M. G. Spalding, and D. J. Forrester. 2001. Altered predation susceptibility of mosquitofish infected with Eustrongylides ignotus. Journal Wildlife Diseases 37:556–560. Coyner, D. F., M. G. Spalding, and D. J. Forrester. 2002. Epizootiology of Eustrongylides ignotus in Florida: Distribution, density, and natural infections in intermediate hosts. Journal of Wildlife Diseases 38:483–499. Coyner, D. F., M. G. Spalding, and D. J. Forrester. 2003a. Epizootiology of Eustrongylides ignotus in Florida: Transmission and development of larvae in intermediate hosts. Journal of Parasitology 89:290–298. Coyner, D. F., M. G. Spalding, and D. J. Forrester. 2003b. Influence of treated sewage on infections of Eustrongylides ignotus (Nematoda: Dioctophymatoidea) in eastern mosquitofish (Gambusia holbrooki) in an urban watershed. Comparative Parasitology 70:205–210. Coyner, D. F., M. G. Spalding, and D. J. Forrester. 2004. Influences of salinity and desiccation on development of first stage larvae in the egg of Eustrongylides ignotus and their impact on the epizootiology of Eustrongylides ignotus in Florida, U.S.A. Comparative Parasitology 71:262–263. Coy-Otero, A., and J. J. Martinez. 1987. Nuevo hallazgo en Cuba de larvas del genero Eustrongylides Jaegerskiold, 1909 (Nematoda: Dioctophymidae) en la rana toro (Rana catesbeiana). Miscelanea Zoologica (Havana) 3:32. Cram, E. B. 1933. Eustrongylides ignotus from the black-crowned night heron. Journal of Parasitology 20:71. Crites, J. S. 1982. Impact of the nematode parasite Eustrongylides tubifex on yellow perch in Lake Erie. Commercial Fisheries Research and Development Project, Ohio. U.S. Department of Commerce. National Oceanic and Atmospheric Administration, Washington, DC. Dickinson, P. 1951. Stomach contents of land shags in New Zealand. Australian Journal of Marine and Fresh-Water Resources 2:245–253. Dubinin, V. B., and M. N. Dubinina. 1940. [Parasite fauna of colonial birds of the Astrachan Sanctuary] (in Russian). Trudy Astrakhanskogo Gosudarstvennogo Zapovednika 3:190–298. Dubinin, V. M. 1949. [Experimental studies on the cycle of development of some worm parasites of animals of the Volga delta] (in Russian). Parazitologicheskii Sbornik Zoologicheskii In-ta AN SSSR 11:154–156. (Original not seen, in Karmanova 1986)
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Dubinina, M. N. 1937. [Parasite fauna of the Night Heron, Nycticorax nycticorax L., and changes in it due to migration of the host] (in Russian). Zoologicheskii Zhurnal 16:547–573. Dusen, S., and M. Oz. 2006. Helminths of the marsh frog, Rana ridibunda Pallas, 1771 (Anura: Ranidae), from Antalya Province, Southwestern Turkey. Comparative Parasitology 73:121–129. Dyer, W. G., E. H. Williams, Jr., A. A. Mignucci-Giannoni, N. M. Jimenez-Marrero, L. Bunkley-Williams, D. P. Moore, and D. B. Pence. 2002. Helminth and arthropod parasites of the Brown Pelican, Pelecanus occidentalis, in Puerto Rico, with a compilation of all metazoan parasites reported from this host in the Western Hemisphere. Avian Pathology 31:441–448. Eberhard, M. L., H. Hurwitz, A. M. Sun, and D. Coletta. 1989. Intestinal perforation caused by larval Eustrongylides (Nematoda: Dioctophymatoidae) in New Jersey. American Journal of Tropical Medicine and Hygiene 40:648–650. Evans, R. H. 1990. Ask the D.V.M. Wildlife Rehabilitation Today, Winter 1990:43–44. fa*gerholm, H. P. 1979. Nematode length and preservatives, with a method for determining the length of live specimens. Journal of Parasitology 65:334–335. Fastzkie, J. S., and J. L. Crites. 1977. A redescription of Eustrongylides tubifex (Nitzsch 1819) Jagerskiold 1909 (Nematoda: Dioctophymatidae) from mallards (Anas platyrhynchos). Journal of Parasitology 63:707–712. Feizullaev, N. A. 1962. [Fauna and ecology of helminths of waterfowl (Ciconiiformes) of low-lying regions of Azerbaizhan] (in Russian). Avtoref. Kand. Diss., Baku, pp. 1–17. (Original not seen, in Karmanova 1986) Feizullaev, N. A. 1963. [Helminth fauna (Nematoda, Cestoda, Acanthocephala) of birds of the order Ciconiiformes of low-lying regions in Azerbaidzhan] (in Russian). Izvestiya Akademii Nauk Azerbaidzhanskogo SSR, Seriia Biologicheskaia I Meditsinski Naukite 2:61–67. Ferenc, S. A., T. L. Tallevast, and C. H. Courtney. 1986. Experimental infection of the brown water snake, Nerodia taxispilota, with Sebekia mississippiensis (Pentastomida). Proceedings of the Helminthological Society of Washington 53:296–297. Forrester, D. J., and M. G. Spalding. 2003. Parasites and Diseases of Wild Birds in Florida. University Press of Florida, Gainesville, FL. Foster, G. W., P. E. Moler, J. M. Kinsella, S. P. Terrell, and D. J. Forrester. 2000. Parasites of eastern indigo snakes (Drymarchon corais couperi) from Florida, U.S.A. Comparative Parasitology 67:124–128.
Franson, J. C., and T. W. Custer. 1994. Prevalence of eustrongylidosis in wading birds from colonies in California, Texas, and Rhode Island. Colonial Waterbirds 17:168–172. Frederick, P. C., and S. M. McGehee. 1994. Wading bird use of wastewater treatment wetlands in central Florida, USA. Colonial Waterbirds 17: 50–59. Frederick, P. C., S. M. McGehee, and M. G. Spalding. 1996. Prevalence of Eustrongylides ignotus in mosquitofish (Gambusia holbrooki) in Florida: Historical and regional comparisons. Journal of Wildlife Diseases 32:552–555. Gibson, G. G., and D. A. McKiel. 1972. Dracunculus insignis (Leidy, 1858) and larval Eustrongylides sp. in a muskrat from Ontario, Canada. Canadian Journal of Zoology 50:897–901. Goldberg, S. R., C. R. Bursey, and A. L. Aquino-Shuster. 1991. Gastric nematodes of the Paraguayan caiman, Caiman yacare (Alligatoridae). Journal of Parasitology 77:1009–1011. Graber, M. 1975. Helminths and Helminthiases of Domestic and Wild Animals of Ethiopia, Vol. 2. Institut d’Elevage et de Medecine Veterinaire des Pays Tropicaux, Maisons-Alfort, France. Guerin, P. F., S. Marapendi, L. McGrail, C. L. Moravec, and E. L. Schiller. 1982. Intestinal perforation caused by larval Eustrongylides–Maryland. Center for Disease Control, Morbidity and Mortality Weekly Reports 31:383–389. Gunby, P. 1982. One worm in the minnow equals too many in the gut. Journal of the American Medical Association 248:163. Hirshfield, M. F., R. F. Morin, and D. J. Hepner. 1983. Increased prevalence of larval Eustrongylides (Nematoda) in the mummichog, Fundulus heterocl*tis (L.), from the discharge canal of a power plant in the Chesapeake Bay. Journal of Fish Biology 23: 135–142. Hoberg, E. P., C. J. Henny, O. R. Hedstrom, and R. A. Grove. 1997. Intestinal helminths of river otters (Lutra canadensis) from the Pacific Northwest. Journal of Parasitology 83:105–110. Hoeppli, R., H. F. Hsu, and H. W. Wu. 1929. Helminghologische Beitrage aus f*ckien und Chekiang. Archiv fur Schiffs und Tropen-Hygiene, Pathologie und Therapic Exotischer Krankheiten 33:1–44. Jagerskiold, L. A. 1909a. Nematodes aus Aegypten und dem Sudan. Results of the Swedish Zoological Expedition to Egypt and the White Nile, 1901 3:1–66. Jagerskiold, L. A. 1909b. Zur Kenntnis der Nematoden Gattungen Eustrongylides und Hystrichis. Nova Acta Regiae Societatis Scientiarum Upsaliensis. Series IV 2:1–48.
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Eustrongylidosis Johnston, T. H., and P. M. Mawson. 1941. Additional nematodes from Australian birds. Transactions of the Royal Society of South Australia 65:254–262. Jones, H. I. 1978. Gastrointestinal nematodes from aquatic Australian snakes. Memoirs of the Queensland Museum 18:243–263. Karmanova, E. M. 1986. Dioctophymidea of Animals and Man and Their Causation of Disease, Essentials of Nematology, Vol. 20, K. I. Skrjabin (ed.). Amerind Publishing Company, New Delhi, India, pp. 1–383. Kontrimavichus, V. L., and T. L. Bakhmet’eva. 1960. [Helminth fauna of auks of the lower flow of the Lena River] (in Russian). Trudy Gelan 10:124–133. Kosupko, G. A. 1963. [Study of nematodes of piscivorous birds at the Astrachan’ sanctuary] (in Russian). Materialy Nauchogo Konferentsii Vsesoyuzn Obshchestva Gel’mintologov. Moscow 1:158– 159. Kuperman, B. I., V. E. Matey, R. N. Fisher, E. L. Ervin, M. L. Warburton, L. Bakhireva, and C. A. Lehman. 2004. Parasites of the African clawed frog, Xenopus laevis, in Southern California, U.S.A. Comparative Parasitology 71:229–232. Leuckart, R. 1868. Die mehnschlichen Parasiten. Leipzig 2:381–387. (Original not seen, in Karmanova 1986) Lezama, J. R., and D. O. Sarabia. 2002. Histological lesions in skeletal muscles caused by Eustrongylides sp. (Nematoda: Dioctophymatoidea) larvae in edible frogs from Lake Cuitzeo, in the state of Michoacan, in Mexico. Veterinaria Mexico 33:335–341. Lichtenfels, J. R., and B. Lavies. 1976. Mortality in red-sided garter snakes, Thamnophis sirtalis parietalis, due to larval nematodes, Eustrongylides sp. Laboratory Animal Science 26:465–467. Lichtenfels, J. R., and P. A. Madden. 1980. Cephalic papillae of giant kidney nematode Dioctophyma renale (Goeze, 1782) and comparison with Eustrongylides spp. Proceedings of the Helminthological Society of Washington 47:55–62. Lichtenfels, J. R., and C. F. Stroup. 1985. Eustrongylides sp. (Nematoda: Dioctophymatoidea): First report of an invertebrate host (Oligochaeta: Tubificidae) in North America. Proceedings of the Helminthological Society of Washington 52:320–323. Locke, L. N. 1961. Heron and egret losses due to verminous peritonitis. Avian Diseases 5: 135–138. Locke, L. N., J. B. DeWitt, C. M. Menzie, and J. A. Kerwin. 1964. A merganser die-off associated with larval Eustrongylides. Avian Diseases 8:420–427. Lyubimov, M. P. 1926. [Study of the helminth fauna of domestic and wild ducks of the Soviet Union] (in Russian). Trudy Gosudarstvennogo Instituta Ekspermentalnoi Veterinarii 3:13–26.
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Canavan 1929, occurring encapsulated in the tissues of Amphiuma in Louisiana. Journal of Parasitology 65:96–104. Paperna, I. 1974. Hosts, distribution, and pathology of infections with larvae of Eustrongylides (Dioctophymidae, Nematoda) in fishes from East African lakes. Journal of Fisheries Biology 6:67–76. Pinto, R. M., L. A. Barros, L. Tortelly, R. F. Teixeira, and D. C. Gomes. 2004. Prevalence and pathology of helminths of Ciconiiform birds from the Brazilian swamplands. Journal of Helminthology 78:259–264. Rautela, A. S., and S. K. Malhotra. 1984. Nematode fauna of high altitude avian hosts in Garhwal Himalayan ecosystems. I. Eustrongylides spinispiculum n. sp. and revised key to the species of genus Eustrongylides Jagerskiold (1909). Korean Journal of Parasitology 22:242–247. Rego, A. A., and J. J. Vicente. 1988. Eustrongylides ignotus Jagerskiold, 1909 (Nematoda: Dioctophymatoidea), plarasito de peixes, anfibios, repteis e aves. Distribuicao e taxonomia. Ciencia e Cultura 40:478–483. Roffe, T. J. 1988. Eustrongylides sp. epizootic in young common egrets (Casmerodius albus). Avian Diseases 32:143–147. Rudolphi, C. A. 1819. Entozoorum Synopsis cui Accedunt Mantissa Dublex et Indices Locupletissimi. Berolini. Ryzhikov, K. M. 1950. [Report on the 268th united helminthological expedition to western Georgia during 1948] (in Russian). Trudy Gel’mintologicheskoi Laboratorii 5:252–260. Ryzhikov, K. M. 1963. [Nematodes of geese of Kamchatka] (in Russian). Trudy GELAN 13:131–143. Sattari, M., and B. Mokhayer. 2005. Occurrence and intensity of some parasitic worms in Acipenser gueldenstaedti, A. nudiventris and Huso huso (Chondrostei: Acipenseridae) from the southwest of the Caspian Sea. Turkish Journal of Veterinary and Animal Sciences 29:1279–1284. Schaffer, G. V., A. A. Rego, and G. C. Pavanelli. 1990. [A study of the lesions caused by Eustrongylides ignotus Jagerskiold, 1909 (Nematoda: Dioctophymatoidea) in fish eating birds of Brazil.] (in Portuguese) Revista UNIMAR 12:201–208. Sepulveda, M. S., M. G. Spalding, J. M. Kinsella, R. D. Bjork, and G. S. McLaughlin. 1994. Helminths of the Roseate Spoonbill, Ajaia ajaja, in southern Florida. Journal of the Helminthological Society of Washington 61:179–189. Shchupakov, I. T. 1936. [Parasite fauna of the Caspian seal] (in Russian). Uchenge Zapiski LGU, Number 7, Series Biology 3:135–143. Shigin, A. A. 1957. [Worm parasites of herons and grebes of Rybinsk reservoir] (in Russian). Trudy
Darvinskogo Gosurdatstvennogo Zapovednika 4:245–289. Shigin, A. A. 1961. [Helminth fauna of gulls of Rybinsk reservoir] (in Russian). Trudy Darvinskogo Gosurdatstvennogo Zapovednika 7:309–362. Shillinger, J. E. 1936. Committee on parasitic diseases: Parasites in wildlife. Journal of the American Veterinary Medical Association 41:423–430. Shirazian, D., E. L. Schiller, C. A. Glaser, and S. L. Vonderfecht. 1984. Pathology of larval Eustrongylides in the rabbit. Journal of Parasitology 70:803–806. Skrjabin, K. I. 1916. Ne’matodes des oiseaux du Turkestan Russe. Ottisk iz Ezhegodnika Zoologicheskogo Museia Imperatorskoi Akademii Nauk, Petrograd 20:457–557. Skryabin, K. I. 1920. [Second Azov-Don helminthological expedition] (in Russian). Izviestiia Donskogo Veterinarnago Instituta, Novocherkassk 1:1–18. Skryabin, K. I. 1923. Pyatay Rossiikaya gel’mintologicheskaya ekspeditsiya v Turkestanskii krai [Fifth Russian helminth expedition in Turkestan region]. Trudy GTos. Instituta Experimental’noi Veterinarii, Vol. 1, No. 1, pp. 12–47. Smogorzhevskaya, L. A. 1954. [Helminth fauna of piscivorous birds of the Dnieper valley] (in Russian). Avtoreferat Kandidatskoi Dissertatsii 1–17. Smogorzhevskaya, L. A. 1959. [Ecological characteristics of piscivorous birds of the Dnieper valley] (in Russian). Voprosy Ekologii 3:222– 231. Smogorzhevskaya, L. A. 1962. [Nematodes of piscivorous birds of the Dnieper valley] (in Russian). Materialy Do Vichenya Istorii ta Prirodi R-ku Kanivs’k Zapovidn. Kiev. pp. 118–126. Smogorzhevskaya, L. A. 1964. [Extent of study of helminth fauna of waterfowl in UkrSSR] (in Russian). Trudy Ukrainskogo Respublikanskogo Nauchogo Obshchestva Parazitologov 3:125–188. Spalding, M. G. 1990. Antemortem diagnosis of eustrongylidosis in wading birds (Ciconiiformes). Colonial Waterbirds 13:75–77. Spalding, M. G., and D. J. Forrester. 1991. Effects of parasitism and disease on the reproductive success of colonial wading birds (Ciconiiformes) in southern Florida. Florida Game and Fresh Water Fish Commission Final Report, Tallahassee, FL, 130 pp. Spalding, M. G., and D. J. Forrester. 1993. Pathogenesis of Eustrongylides ignotus (Nematoda: Dioctophymatoidea) in Ciconiiformes. Journal of Wildlife Diseases 29:250–260. Spalding, M. G., G. T. Bancroft, and D. J. Forrester. 1993. The epizootiology of eustrongylidosis in wading birds (Ciconiiformes) in Florida. Journal of Wildlife Diseases 29:237–249.
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Proceedings of the Helminthological Society of Washington 10:29–33. Weisberg, S. B., R. P. Morin, and E. A. Ross. 1986. Eustrongylides (Nematoda) infection in mummichogs and other fishes of the Chesapeake Bay region. Transactions of the American Fisheries Society 115:776–783. Wiese, J. H., W. R. Davidson, and V. F. Nettles. 1977. Large scale mortality of nestling Ardeids cause by nematode infection. Journal of Wildlife Diseases 13:376–382. Windingstad, R. M., and D. M. Swineford. 1981. Eustrongylides and pesticide levels in a Great blue heron shot in Wisconsin. The Prairie Naturalist 13:61–62. Winsor, H. 1948. Hosts of eustrongyloid worms from Fairmont Park Aquarium and Philadelphia Zoo. Proceedings of the Pennsylvania Academy of Science 22:68–72. Winterfield, R. W., and K. R. Kazacos. 1977. Morbidity and mortality of great blue herons in Indiana caused by Eustrongylides ignotus. Avian Diseases 21:448–451. Wu, H. K., and C. K. Liu. 1943. Helminthological notes. III. Sinensia. Contributions of the Metropolitan Museum of Natural History in Nanking 14:99–105. Yamaguti, S. 1935. Eustrongylides elegans (Olfers) in wall of proventriculus of Podiceps ruficollis, Japan. Japanese Journal of Zoology 6:420. Ziegler, A. F., S. C. Welte, E. A. Miller, and T. J. Nolan. 2000. Eustrongylidiasis in eastern great blue herons (Ardea herodias). Avian Diseases 44:443–448.
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17 Trichostrongylus Daniel M. Tompkins INTRODUCTION Members of the genus Trichostrongylus are small, fine, reddish worms that occur primarily in the ceca, but also the small intestine of birds, as well as the small intestine of rodents, lagomorphs, and ruminants. Two species are currently described from birds. Infection in wild birds has been across Europe, Asia, North America, Africa, and Australasia, reported primarily in Anseriformes (waterfowl) and Galliformes (fowl), but also in Gruiformes (cranes) and Otidiformes (bustards). Species of Trichostrongylus are monoxenous, requiring only a single host to complete the life cycle. The free-living, infective third-stage larvae tend to crawl onto vegetation where they are ingested. As with many other gastrointestinal parasites, intense infections of Trichostrongylus worms can cause morbidity, with signs that include loss of appetite, plumage dullness, and emaciation (Wehr 1971). However, intense infections such as these tend to occur naturally only in Red Grouse (Lagopus lagopus scotica, referred to as Willow Ptarmigan in The Clements Checklist of Birds of the World) infected with Trichostrongylus tenuis on mainland UK shooting estates. Here, prevalence of infection is almost 100% (Hudson 1992), geometric average worm burdens have been estimated at 2,471 per bird (Webster 2005), and parasite impacts on both individuals and populations have been characterized (Hudson 1986a; Shaw and Moss 1990; Dobson and Hudson 1992; Hudson et al. 1992a, 1998; Delahay and Moss 1996). Infection of T. tenuis in Red Grouse was also the first system in which regulation of host population size by parasites was experimentally demonstrated in the field (Hudson et al. 1998; Tompkins and Begon 1999).
HISTORY Trichostrongylus tenuis (Mehlis, in Creplin 1846) Raillet and Henry, 1909, was the first of the avian Trichostrongylus species to be described, originally as Strongylus tenuis from the European Ring-necked Pheasant (Phasianus colchicus). Other synonyms for Trichostrongylus tenuis included Strongylus pergracilis Cobbold, 1873; Strongylus serratus Linstow, 1876; Trichostrongylus pergracilis (Cobbold 1873) Raillet and Henry, 1909. Trichostrongylus tenuis was subsequently recognized as the trichostrongylid inhabiting the ceca of many Old-World birds, although it was first described in Red Grouse as T. pergracilis (Cobbold 1873) Raillet and Henry, 1909. Cram (1925) applied the latter name to the cecal worm from the Northern Bobwhite (Colinus virginianus) in the US. She later recognized two species: Trichostrongylus tenuis from 10 bird species (including waterfowl) and T. pergracilis from the Northern Bobwhite and Red Grouse (Cram 1927). After further study of material from many different hosts, including the type host (Ring-necked Pheasant) and from Red Grouse, the type host of T. pergracilis, Cram and Wehr (1934) synonymized both taxa as T. tenuis. Recently, however, the trichostrongylid infecting Northern Bobwhite has been recognized as a distinct North American species, Trichostrongylus cramae, with the name T. tenuis reserved for Old-World hosts (Durette-Desset et al. 1993). This designation has been independently supported by experimental challenges of Northern Bobwhites, in which infective Trichostrongylus larvae of Northern Bobwhite origin readily established while those of Red Grouse origin failed to do so (Freehling and Moore 1993). In light of this, the status of species of Trichostrongylus infecting other wild birds in North America needs reassessment (Anderson 1996), a process that has already started. Trichostrongylus tenuis has been associated with losses in managed populations of Red Grouse in England and Scotland for more than 100 years (Cobbold 1873; Shipley 1909; Lovat 1911). The development and transmission of T. tenuis in Red Grouse was the
SYNONYMS Cecal strongyle, caecal threadworm, grouse disease, hairworms, trichostrongylosis, trichostrongyliasis.
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Trichostrongylus subject of an in-depth investigation by several authors in the early 1900s. In their collated findings (Leslie and Shipley 1912), the authors relate the prevalence, intensity, and pathogenicity of T. tenuis to “grouse disease” and fluctuations in grouse numbers on moors in the UK. Most of the basic findings reported by these authors have been confirmed by more recent investigations, culminating in the classic experimental demonstration of the role that T. tenuis plays in regulating the population size of Red Grouse in the wild (Hudson et al. 1998). ETIOLOGY The genus Trichostrongylus belongs to the superfamily Trichostrongyloidea, the largest superfamily in the group known as the “bursate nematodes” (order Strongylida). Avian species of Trichostrongylus are readily observed under the microscope as small, fine reddish worms, after extraction (usually through sieves) from the cecal and intestinal contents (see Doster and Goater 1997 for extraction methods). Adult male worms are approximately 4–8 mm long and adult females are 6–10 mm, with width gradually increasing from approximately 10 μm at the anterior end to approximately 50 μm in front of the caudal bursa (Figure 17.1). The size and tapered morphology of these worms, in addition to characteristic cuticular striations (see below), makes them unlikely to be confused with other helminths found in the ceca and small intestine of birds. Two characters can be easily and reliably used to distinguish adult T. tenuis from T. cramae (Durette-Desset et al. 1993). First, transverse striations on T. tenuis start from behind the excretory pore of both sexes and cover three- to four-fifths of the body surface. While male specimens of T. cramae also have pronounced transverse striations, they are present in females only on 35–55 μm of the cephalic region in front of the secretory pore (Figure 17.1a). Second, the ratio of the distance between bursal papillae 3 and 4 to the distance between papillae 2 and 3 for T. tenuis ranges from 1:2.2 to 1:3.9. In T. cramae, the ratio ranges from 1:1.3 to 1:1.7 (Figure 17.2b). The two species also differ in the configuration of the caudal bursa dorsal array (Durette-Desset et al. 1993). EPIZOOTIOLOGY Being monoxenous (requiring only a single host), the life cycle of avian species of Trichostrongylus is relatively simple, consisting of egg, four larval stages, and adult (Figure 17.2). Unlike other parasites in the superfamily Trichostrongyloidea, paratenic “transport” hosts are unknown for species of Trichostrongylus. For T. tenuis, eggs measuring approximately 75 × 46 μm are excreted in the fecal droppings of infected hosts at
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the morula (64 cells) stage of development. Eggs are not resistant to desiccation or extreme cold (−15◦ C), with little development occurring at temperatures below freezing point (Shaw et al. 1989; Connan and Wise 1994). In moist feces at suitable temperatures (>5◦ C), eggs hatch in approximately 36–48 h. Emerging firststage larvae measure approximately 360 μm in length. Larvae molt into the second stage after approximately 36–48 h, and then into the third (infective) stage after 8–16 days, depending on temperature. Like the eggs, the larvae are susceptible to desiccation and extreme cold. Continuous temperatures of −15◦ C are lethal to infective larvae within 12 days (Shaw et al. 1989; Connan and Wise 1994). However, they are capable of withstanding winter temperatures on grouse moors in Yorkshire, England, where temperatures fall as low as −10◦ C), and remain infective for up to 3 weeks (Connan and Wise 1994). However, little to no transmission to new hosts occurs during the winter months (Shaw 1988). Similar environmental limits on infection are seen with infections of T. tenuis in wild Red-legged Partridges (Alectoris rufa) in Spain, where parasite abundance is inversely correlated to latitude and directly correlated to yearly mean temperature (Calvete 2003). Host infection by species of Trichostrongylus is via ingestion of third-stage larvae that are still sheathed by the cuticle of the second stage; skin penetration is unknown in the genus. For T. tenuis on UK moors, third-stage larvae have been experimentally demonstrated to crawl to the growing tips of moist heather plants (Calluna vulgaris), the staple diet of adult Red Grouse. Here, they increase their probability of infecting a suitable host by accumulating in drops of water at the most likely point of ingestion by an adult grouse. Movement to heather tips is governed by negative geotaxis, positive phototaxis, and may also be influenced by responses to chemical cues given off by heather (McGladdery 1984; Saunders et al. 2001). The resultant pattern is a highly aggregated distribution of infective larvae among heather plants in the field (Saunders et al. 1999, 2000). Larvae are more available on plants during daylight hours in experimental diurnal periods and on plants collected from the field in the afternoon rather than the morning, indicating that larvae migrate back down plants at night (Saunders et al. 2000). This behavior most likely evolved to avoid desiccation. Indeed, time-series analyses suggest that some of the year-to-year variation in parasite egg numbers that are passed by Red Grouse is explained by rainfall in the previous summer, likely caused by greater transmission during wetter summers (Moss et al. 1993). Furthermore, there is strong evidence that the interactions between T. tenuis and climate act to synchronize the dynamics of Red Grouse
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(b)
(a)
(c)
(d)
Figure 17.1. (a) Trichostrongylus tenuis caudal bursa, ventral view. (b) T. tenuis female, anterior end, lateral view. (c) Trichostrongylus cramae caudal bursa, ventral view. (d) T. cramae female, anterior end, lateral view. Numbers on part (a) denote ray papillae. Modified from Durette-Desset et al. (1993), with permission of Annales de Parasitologie.
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Figure 17.2. Life cycle of Trichostrongylus tenuis infecting Red Grouse (Lagopus lagopus scotica). Parasites are not drawn to scale.
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populations in northern England (Cattadori et al. 2005). Exsheathment, the process by which a third-stage larva escapes from the cuticle of the second stage, occurs within a few days of ingestion. Unsheathed thirdstage larvae invade the small intestine and cecal mucosa (Shaw 1988). At this point, as with other species of Trichostrongylus, the larvae may either develop directly into fourth-stage larvae and then return to the gut lumen where they mature into adults within a few days, or undergo a period of “arrest” at the third stage within the mucosa. In his analysis of the larval parasite population present within individual grouse in different seasons, Shaw (1988) provides strong evidence for this “arrestment” by T. tenuis in the UK. In late summer, most larvae are in the fourth stage but some third-stage larvae are sheathed, indicating that transmission is still taking place. In winter, ensheathed larvae are absent but exsheathed larvae are present, suggesting that few new larvae are being ingested during this period and that the exsheathed third-stage larvae are arrested. In spring, the proportion of third-stage larvae decreases significantly in the parasite population and sheathed larvae are absent, indicating the development of overwintering third-stage larvae to fourth and adult stages. This synchronized development results in a “spring rise” in worm egg production that is evident in grouse feces (Shaw 1988; Shaw et al. 1989). A second peak of recruitment into the adult worm population occurs during late summer that reflects increased numbers of larvae on heather in the preceding weeks. These increases are associated with optimal conditions for transmission where mean temperatures are in the range of 10–15◦ C and mean monthly rainfall is in the range of 20–50 mm. When development is direct for T. tenuis in Red Grouse, the prepatent period between ingestion of infective stages and the appearance of parasite eggs in the feces is approximately 7–8 days. Regardless of whether development is direct or arrested, however, the resulting distribution of parasites among hosts retains the characteristic aggregation of macroparasite infections (Shaw and Dobson 1996). Few host individuals have many parasites, while most host individuals have few parasites (Wilson 1983; Hudson et al. 1992a). Infected host individuals can carry worm intensities of up to 24,000 worms (Hudson 1986b). Adult worms favor the proximal region of the cecal mucosa, threading themselves into the mucosal tissue with anterior and posterior ends protruding into the cecal lumen (Watson et al. 1987, 1988). Eggs are produced at a rate of approximately 5 × 105 eggs per worm per year (Shaw and Moss 1989a; Hudson et al. 1992a). Adult worms can survive for more than 2 years, although parasite egg output decreases with age of the worm population, especially in winter (Shaw and Moss
1989a). There is no evidence of density-dependent suppression of parasite establishment or egg production at high worm intensities (Hudson and Dobson 1997; Seivwright et al. 2004). There are notable differences between the biology of T. tenuis in Red Grouse and the biology of T. cramae in Northern Bobwhites (Freehling and Moore 1993). While T. tenuis produces a chronic infection in Red Grouse that increases throughout the life of the bird (Potts et al. 1984), T. cramae in Northern Bobwhites in northern Florida is a seasonally occurring parasite. Moore et al. (1986) demonstrated a winter peak in acquisition of larvae that was followed by a peak in adult worms, with a decrease in prevalence and intensity occurring during warmer months. Infections of T. tenuis in other host species also appear to be more similar to those of T. cramae, with the high burdens and chronic infections observed in Red Grouse likely being an exception rather than the rule (Holmstad et al. 2004; Millan et al. 2004; Fedynich et al. 2005; Schei et al. 2005; Webster et al. 2007). This may very well be a product of the unnaturally high densities of managed Red Grouse populations in the UK. CLINICAL SIGNS Hosts carrying high intensities of parasites show evidence of appetite loss, malnutrition (including dullness of the plumage), diarrhea, and emaciation (“grouse disease”). Among Red Grouse, signs peak during spring, coinciding with the “spring rise” in worm egg production, and in the autumn in young hatch-year birds. PATHOLOGY Being threaded into the cecal mucosa, adult worms cause trauma, atrophy, and flattening of the epithelial cells. This likely interferes with the normal digestion of heather and other plant material (Watson et al. 1987, 1988). As is common for other cecal nematodes, heavy infections may cause hemorrhagic typhlitis, and in chronic cases, the contents of the ceca may contain a yellowish white material of cheesy consistency. Since the remarkably large ceca of Red Grouse play a key role in food digestion and nutrient recovery in this species (Hudson 1986b), the disruption in normal cecal function by high-intensity infections with T. tenuis leads to a loss of body condition, increased mortality (Hudson 1986b; Hudson et al. 1992a), and decreased fecundity with reductions in both clutch size and brood size (Hudson 1986a; Shaw and Moss 1990; Hudson et al. 1992a; Delahay and Moss 1996). Negative associations between the abundance of T. tenuis and host body mass, body condition, and breeding survival have also been reported for Willow Ptarmigan in Norway
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Trichostrongylus (Holmstad et al. 2005). For T. cramae, no impacts related to disease have been noted on either Northern Bobwhites or Attwater’s Prairie Chicken (Tympanuchus cupido attwateri), a subspecies of the Greater Prairie-Chicken (Tympanuchus cupido pinnatus) (Purvis et al. 1998), although T. cramae has been recorded at a mean intensity of 1,019 worms per bird in the latter species (Peterson et al. 1998). The pathological effects of these infections should be investigated further in this species. Both redness of the comb of male birds (a secondary sexual character of Red Grouse that functions in both intra- and intersexual selection; Redpath et al. 2006a) and the concentration of plasma carotenoids (the molecules responsible for the redness; Mougeot et al. 2007a) negatively correlate with abundance of T. tenuis (Mougeot et al. 2005a, 2007b). This link has been experimentally confirmed, through observations of plasma carotenoid concentration and comb redness both increasing following reduction of infections with T. tenuis (Martinez-Padilla et al. 2007) and decreasing following challenge with T. tenuis (Mougeot et al. 2007b). DIAGNOSIS The recovery of parasite individuals as fragments in feces or, preferably, intact parasites from postmortem examinations is required for reliable diagnosis. Infective stages (eggs or larvae in feces) are also highly suggestive of infection. However, diagnosis based on infective stages should be confirmed by a postmortem examination to verify the presence of adult Trichostrongylus in the cecae and to establish the extent of infection and damage. Fecal egg counts can be used as a reliable estimate of relative parasite intensity among individuals at the same time of year (Shaw and Moss 1989a; Seivwright et al. 2004), but should not be considered reliable enough to assess impacts on populations. Presence of parasites alone is not indicative of disease in infections with Trichostrongylus. The impacts of disease are only likely to be common when intensity of infection is in the thousands of worms per individual, as has been observed in the field for T. tenuis in Red Grouse and T. cramae in Attwater’s Prairie Chicken. IMMUNITY Watson et al. (1988) demonstrated that young domestic chickens acquire resistance to T. tenuis on repeated exposure to the parasite, such that artificially induced infections are rejected and worms are actively expelled. These findings are supported by observations of higher parasite prevalence and intensity in growers as opposed to adult free-range chickens (Magwisha et al. 2002). While the potential immune responses of wild birds to
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infection by T. cramae need further investigation, studies with Red Grouse have found no evidence of such “acquired” immunity to infection. Here, the number of worms present in the cecae increases throughout the life of the bird (Wilson 1983). A similar increase with age has also been observed in infections of T. tenuis in Red-legged Partridges in Spain (Millan et al. 2004). In a study on transmission dynamics and host–parasite interactions, Hudson and Dobson (1997) used ageintensity data to show that the rate of uptake of T. tenuis by Red Grouse increases during the first 6 months of a bird’s life, reflecting an increase in feeding rate with age, with no sign of self-cure. Furthermore, reinfection rates of adults that were treated to reduce parasite intensities were not significantly different from infection rates of na¨ıve immature grouse. This continued rise of infection, observed over 18 months, demonstrates a lack of any strong host-mediated response against the parasite. However, Red Grouse do vary in their innate susceptibility to infection with T. tenuis, with wide and repeatable variation in individual resistance (Shaw and Moss 1989b). The intensities of T. tenuis and other parasite species positively covary in the Willow Ptarmigan, suggesting that repeatable variation in innate susceptibility to gastrointestinal nematode infection also occurs in this species (Holmstad and Skorping 1998). Susceptibility to infection by T. tenuis also varies with season in Red Grouse, with rates of parasite establishment in co*cks being higher in autumn than in summer. Shaw and Moss (1989b) hypothesized that this is likely linked with behavior, since co*cks show higher levels of territoriality in the autumn. Increased corticosteroid levels from stress associated with such behavior could lower resistance to parasites. Recent research has partly supported this hypothesis, indicating that elevated testosterone (not corticosteroid) levels in male Red Grouse increases susceptibility to infection by T. tenuis (Mougeot et al. 2005b, 2006; Seivwright et al. 2005). Furthermore, the opposing effects of testosterone and T. tenuis on carotenoid availability in Red Grouse, whereby infection by T. tenuis decreases availability (as noted above) while testosterone enhances comb redness but tends to deplete plasma carotenoids (Mougeot et al. 2007b), likely makes the red comb an “honest” signal of male health for mate choice by females (Mougeot et al. 2004). DOMESTIC ANIMAL HEALTH CONCERNS Avian species of Trichostrongylus tend to occur only at low prevalence in domestic poultry, geese, and ducks, with infected individuals tending not to exhibit signs of disease (Svoboda 1992; Romaniuk and Lipinski 1999; Poulsen et al. 2000; Magwisha et al. 2002; Irunga et al. 2004). Cross-infection from wild birds can occur,
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although prevalence is generally low without the need for preventative management. WILDLIFE POPULATION IMPACTS The genus Trichostrongylus has been detected in many wild populations of Galliformes and Anseriformes, although, as mentioned above, high intensities of infection have only been observed for T. tenuis infecting Red Grouse and T. cramae infecting Attwater’s Prairie Chicken (Hudson 1986b; Watson and Shaw 1991; Holstadt et al. 1994; Peterson et al. 1998; Purvis et al. 1998; Millan et al. 2004). To date, impacts of such high-intensity infections have only been demonstrated in Red Grouse—disease impacts of T. cramae on Attwater’s Prairie Chicken require further investigation (Peterson 2004). For Red Grouse with infections of T. tenuis, the direct effects of infection on mortality and fecundity interact with other factors to impact individual hosts. Heavily infected birds also experience increased predation, most likely due to increases in emission of scent (Hudson et al. 1992b; Dobson and Hudson 1995). In addition, birds with intense infections are less able to maintain territories due to the energetic consequences of parasitism by T. tenuis (Delahay et al. 1995) and its interaction with testosterone (Mougeot et al. 2006). Indeed, when parasites are removed, aggressive territorial behavior increases (Fox and Hudson 2001; Mougeot et al. 2005c). Finally, parasite infection also influences mating success of male Red Grouse via the interactions with carotenoids discussed above. Impacts of parasites at the scale of host populations have been both extensively modeled and experimentally demonstrated for T. tenuis in Red Grouse on heather moors in northern England. Populations of red grouse have characteristic cyclical dynamics, with cycle periods between 4 and 8 years. Involvement of T. tenuis in these cyclic dynamics has long been suspected, because of the association between population crashes and high parasite intensities recorded from individuals (Leslie and Shipley 1912). Dobson and Hudson (1992) demonstrated in an experimentally parameterized modeling framework that the interaction with T. tenuis was a potential cause of these cycles. They then proved that this was indeed the case by using anthelmintic application to drastically reduce the extent of population crashes at replicated sites (Hudson et al. 1998). This work was the first experimental demonstration of parasite regulation of host numbers in any wildlife population, but did not prove that the population dynamics observed were a function of T. tenuis parasitism alone (Tompkins and Begon 1999). Indeed, the weight of evidence from more recent research supports the hypothesis that it is an interaction between
parasitism, aggressiveness, and territoriality (as observed at the individual scale) that drives the observed dynamics, with differences in the interaction driving observed differences in cyclic behavior of populations between different parts of the UK (Mougeot et al. 2005d; Redpath et al. 2006a, b). TREATMENT AND CONTROL Adult and developing worms can be eliminated via the oral application of standard anthelmintics, such as fenbendazole or levamisole hydrochloride (Samour 2000), although anthelmintic treatment is not highly effective against arrested larvae in the gut mucosa. Anthelmintic resistance has not yet been reported for species of Trichostrongylus of birds (Webster et al. 2007). MANAGEMENT IMPLICATIONS Trichostrongylus parasites are currently only of real concern to managed populations of Red Grouse in northern England and Scotland, where T. tenuis plays a role in cyclical population crashes. Although these populations rapidly recover following crashes, the game industry is interested in maintaining high numbers of birds at all times. Wild populations containing infected individuals may also act as reservoirs, increasing risks of infection on sympatric populations of more susceptible host species. It has been hypothesized that such a relationship may occur in North America between Northern Bobwhite and the endangered Attwater’s Prairie Chicken (Peterson 2004), although further research is required to investigate this idea. Large-scale management of T. tenuis in Red Grouse, through the indirect application of anthelmintics via “medicated grit,” has been experimentally proven at the estate scale. This treatment reduces parasite burdens in grouse hens and increases breeding success by improving chick survival (Newborn and Foster 2002). The increase in grouse numbers arising from the use of medicated grit is a viable method for enhancing hunting for Red Grouse, since the cost of treatment is low relative to the return from an increased harvest. Similar large-scale application of anthelmintics may also be applicable to the management of endangered species that are affected by infections with Trichostrongylus, particularly since anthelmintic resistance is not known for this genus of parasites in birds. LITERATURE CITED Anderson, R. C. 1996. Nematode Parasites of Vertebrates—Their Development and Transmission, 2nd ed. CABI Publishing, Wallingford, UK.
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Trichostrongylus Calvete, C. 2003. Correlates of helminth community in the red-legged partridge (Alectoris rufa L.) in Spain. Journal of Parasitology 89:445–451. Cattadori, I. M., D. T. Haydon, and P. J. Hudson. 2005. Parasites and climate synchronize red grouse populations. Nature 433:737–741. Cobbold, T. S. 1873. The grouse disease. A statement of facts tending to prove the parasite origin of the epidemic. “The field” office, London. Connan, R. M., and D. R. Wise. 1994. Further studies on the development and survival at low temperatures of the free-living stages of Trichostrongylus tenuis. Research in Veterinary Science 57(2):215–219. Cram, E. B. 1925. New records of economically important nematodes in birds. Journal of Parasitology 12:113. Cram, E. B. 1927. Bird parasites of the nematode suborders Strongylata, Ascaridata and Spirurata. U.S. National Museum Bulletin 140:334–361. Cram, E. B., and E. E. Wehr. 1934. The status of the species of Trichostrongylus of birds. Parasitology 26:335. Delahay, R., and R. Moss. 1996. Food intake, weight changes and egg production in captive red grouse before and during laying: Effects of the parasitic nematode Trichostrongylus tenuis. Condor 98(3): 501–511. Delahay, R. J., J. R. Speakman, and R. Moss. 1995. The energetic consequences of parasitism—Effects of a developing infection of Trichostrongylus tenuis (nematode) on red grouse (Lagopus lagopus scoticus) energy-balance, body-weight, and condition. Parasitology 110:473–482. Dobson, A. P., and P. J. Hudson. 1992. Regulation and stability of a free-living host-parasite system—Trichostrongylus tenuis in red grouse. 2. Population models. Journal of Animal Ecology 61(2):487– 498. Dobson, A. P., and P. J. Hudson. 1995. The interaction between the parasites and predators of red grouse Lagopus Lagopus scoticus. Ibis 137:87–96. Doster, G. L., and C. P. Goater. 1997. Collection and quantification of avian helminthes and protozoa. In Host–Parasite Evolution: General Principles and Avian Models, D. H. Clayton and J. Moore (eds). Oxford University Press, Oxford, pp. 396–418. Durette-Desset, M.-C., A. G. Chabaud, and J. Moore. 1993. Trichostrongylus cramae n. sp. (Nematoda), a parasite of bob-white quail (Colinus virginianus). Annales de Parasitologie 68(1):43–48. Fedynich, A. M., R. S. Finger, B. M. Ballard, J. M. Garvon, and M. J. Mayfield. 2005. Helminths of Ross’ and greater white-fronted geese wintering in south Texas, USA. Comparative Parasitology 72:33– 38.
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Fox, A., and P. J. Hudson. 2001. Parasites reduce territorial behaviour in red grouse (Lagopus lagopus scoticus). Ecology Letters 4(2):139–143. Freehling, M., and J. Moore. 1993. Host-specificity of Trichostrongylus tenuis from red grouse and northern bobwhites in experimental infections of northern bobwhites. Journal of Parasitology 79(4):538–541. Holmstad, P. R., and A. Skorping. 1998. Covariation of parasite intensities in willow ptarmigan, Lagopus lagopus L. Canadian Journal of Zoology 76:1581– 1588. Holmstad, P. R., O. Holstad, G. Karbol, J. O. Revhaug, E. Schei, V. Vandvik, and A. Skorping. 2004. Parasite tags in ecological studies of terrestrial hosts: A study on ptarmigan (Lagopus spp.) dispersal. Ornis Fennica 81:128–136. Holmstad, P. R., P. J. Hudson, and A. Skorping. 2005. The influence of a parasite community on the dynamics of a host population: A longitudinal study on willow ptarmigan and their parasites. Oikos 111: 377–391. Holstadt, O., G. Karbol, and A. Skorping. 1994. Trichostrongylus tenuis from willow grouse (Lagopus lagopus) and ptarmigan (Lagopus mutus) in northern Norway. Bulletin of the Scandinavian Society for Parasitology 4:9–13. Hudson, P. J. 1986a. The effect of a parasitic nematode on the breeding production of red grouse. Journal of Animal Ecology 55:85–92. Hudson, P. J. 1986b. Red Grouse: The Biology and Management of a Wild Gamebird. The Game Conservancy Trust, Fordingbridge, UK. Hudson, P. 1992. Grouse in Space and Time. Game Conservancy Ltd, Fordingbridge, UK. Hudson, P. J., and A. P. Dobson. 1997. Transmission dynamics and host–parasite interactions of Trichostrongylus tenuis in red grouse (Lagopus lagopus scoticus). Journal of Parasitology 83(2):194– 202. Hudson, P. J., D. Newborn, and A. P. Dobson. 1992a. Regulation and stability of a free-living host-parasite system—Trichostrongylus tenuis in red grouse. 1. Monitoring and parasite-reduction experiments. Journal of Animal Ecology 61(2):477–486. Hudson, P. J., A. P. Dobson, and D. Newborn. 1992b. Do parasites make prey vulnerable to predation? Red grouse and parasites. Journal of Animal Ecology 61: 681–692. Hudson, P. J., A. P. Dobson, and D. Newborn. 1998. Prevention of population cycles by parasite removal. Science 282(5397):2256–2258. Irunga, L. W., R. N. Kimani, and S. M. Kisia. 2004. Helminth parasites in the intestinal tract of indigenous poultry in parts of Kenya. Journal of the South African Veterinary Association 75:58–59.
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Leslie, A. S., and A. E. Shipley. 1912. The Grouse in Health and Disease. Being the Popular Edition of the Report of the Committee of Inquiry on Grouse Disease. Smith, Elder and Co., London. Lovat, L. 1911. The Grouse in Health and Disease. Report of the Committee of Inquiry on Grouse Disease. Smith, Elder and Co., London. Magwisha, H. B., A. A. Kassuku, N. C. Kyvsgaard, and A. Permin. 2002. A comparison of the prevalence and burdens of helminth infections in growers and adult free-range chickens. Tropical Animal Health and Production 34:205–214. Martinez-Padilla, J., F. Mougeot, L. Perez-Rodriguez, and G. R. Bortolotti. 2007. Nematode parasites reduce carotenoid-based signaling in male red grouse. Biology Letters 3:161–164. McGladdery, S. E. 1984. Behavioural responses of Trichostrongylus tenuis (Cobbold) larvae in relation to their transmission to the red grouse, Lagopus lagopus scoticus. Journal of Helminthology 58:295–299. Millan, J., C. Gortazar, and R. Villafuerte. 2004. Ecology of nematode parasitism in red-legged partridges (Alectoris rufa) in Spain. Helminthologia 41:33–37. Moore, J., M. Freehling, and D. Simberloff. 1986. Gastrointestinal helminths of the northern bobwhite in Florida: 1968 and 1983. Journal of Wildlife Diseases 22:497–501. Moss, R., A. Watson, J. B. Trenholm, and R. Parr. 1993. Caecal thread worms Trichostrongylus tenuis in red grouse Lagopus lagopus scoticus: Effects of weather and host density upon estimated worm burdens. Parasitology 107:199–209. Mougeot, F., J. R. Irvine, L. Seivwright, S. M. Redpath, and S. Piertney. 2004. Testosterone, immunocompetence, and honest sexual signaling in male red grouse. Behavioural Ecology 15:930–937. Mougeot, F., S. M. Redpath, and F. Leckie. 2005a. Ultra-violet reflectance of male and female red grouse, Lagopus lagopus scoticus: Sexual ornaments reflect nematode parasite intensity. Journal of Avian Biology 36:203–209. Mougeot, F., S. M. Redpath, S. B. Piertney, and P. J. Hudson. 2005b. Separating behavioral and physiological mechanisms in testosterone-mediated trade-offs. American Naturalist 166:158–168. Mougeot, F., S. A. Evans, and S. M. Redpath. 2005c. Interactions between population processes in a cyclic species: Parasites reduce autumn territorial behaviour of male red grouse. Oecologia 144:289–298. Mougeot, F., S. B. Piertney, F. Leckie, S. Evans, R. Moss, S. M. Redpath, and P. J. Hudson. 2005d. Experimentally increased aggressiveness reduces population kin structure and subsequent recruitment in red grouse Lagopus lagopus scoticus. Journal of Animal Ecology 74:488–497.
Mougeot, F., S. M. Redpath, and S. B. Piertney. 2006. Elevated spring testosterone increases parasite intensity in male red grouse. Behavioural Ecology 17:117–125. Mougeot, F., J. Martinez-Padilla, L. Perez-Rodriguez, and G. R. Bortolotti. 2007a. Carotenoid-based coloration and ultraviolet reflectance of the sexual ornaments of grouse. Behavioural Ecology and Sociobiology 61:741–751. Mougeot, F., L. Perez-Rodriguez, J. Martinez-Padilla, F. Leckie, and S. M. Redpath. 2007b. Parasites, testosterone and honest carotenoid-based signaling of health. Functional Ecology 2007:886–898. Newborn, D., and R. Foster. 2002. Control of parasite burdens in wild red grouse Lagopus lagopus scoticus through the indirect application of anthelmintics. Journal of Applied Ecology 39:909–914. Peterson, M. J. 2004. Parasites and infectious diseases of prairie grouse: Should managers be concerned? Wildlife Society Bulletin 32:35–55. Peterson, M. J., J. R. Purvis, J. R. Lichtenfels, T. M. Craig, N. O. Dronen, and N. J. Silvy. 1998. Serologic and parasitological survey of the endangered Attwater’s prairie chicken. Journal of Wildlife Diseases 34:137–144. Poulsen, J., A. Permin, O. Hindsbo, L. Yelifari, P. Nansen, and P. Block. 2000. Prevalence and distribution of gastro-intestinal helminthes and haemoparasites in young scavenging chickens in upper eastern region of Ghana, West Africa. Preventive Veterinary Medicine 45:237–245. Potts, G. R., S. C. Tapper, and P. J. Hudson. 1984. Population fluctuations in red grouse: Analysis of bag records and a simulation model. Journal of Animal Ecology 53:21–36. Purvis, J. R., M. J. Peterson, N. O. Dronen, J. W. Lichtenfels, and N. J. Silvy. 1998. Northern bobwhites as disease indicators for the endangered Attwater’s Prairie Chicken. Journal of Wildlife Diseases 34:348–354. Redpath, S. M., F. Mougeot, F. M. Leckie, D. A. Elston, and P. J. Hudson. 2006b. Testing the role of parasites in driving the cyclic population dynamics of a gamebird. Ecology Letters 9:410–418. Redpath, S. M., F. Mougeot, F. M. Leckie, and S. A. Evans. 2006a. The effects of autumn testosterone on survival and productivity in red grouse Lagopus lagopus scoticus. Animal Behaviour 71:1297–1305. Romaniuk, K., and Z. Lipinski. 1999. Prevalence of nematodes in breeding and fattening flocks of geese. Medycyna Weterynaryjna 55:672–673. Samour, J. 2000. Avian Medicine. Mosby, London. Saunders, L. M., D. M. Tompkins, and P. J. Hudson. 1999. The dynamics of nematode transmission in the red grouse (Lagopus lagopus scoticus): Studies on the
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Trichostrongylus recovery of Trichostrongylus tenuis larvae from vegetation. Journal of Helminthology 73(2):171–175. Saunders, L. M., D. M. Tompkins, and P. J. Hudson. 2000. Spatial aggregation and temporal migration of free-living stages of the parasitic nematode Trichostrongylus tenuis. Functional Ecology 14(4): 468–473. Saunders, L. M., D. M. Tompkins, and P. J. Hudson. 2001. Strategies for nematode transmission: Selective migration of Trichostrongylus tenuis infective larvae. Journal of Helminthology 75(4):367–372. Schei, E., P. R. Holmstad, and A. Skorping. 2005. Seasonal infection patterns in willow grouse (Lagopus lagopus L.) do not support the presence of parasiteinduced winter losses. Ornis Fennica 82:137–146. Seivwright, L. J., S. M. Redpath, F. Mougeot, L. Watt, and P. J. Hudson. 2004. Faecal egg counts provide a reliable measure of Trichostrongylus tenuis intensities in free-living red grouse Lagopus lagopus scoticus. Journal of Helminthology 78:69–76. Seivwright, L. J., S. M. Redpath, F. Mougeot, F. Leckie, and P. J. Hudson. 2005. Interactions between intrinsic and extrinsic mechanisms in a cyclic species: Testosterone increases parasite infection in red grouse. Proceedings of the Royal Society B—Biological Sciences 272:2299–2304. Shaw, D. J., and A. P. Dobson. 1996. Patterns of macroparasite abundance and aggregation in wildlife populations: A quantitative review. Parasitology 111: S111–S133. Shaw, J. L. 1988. Arrested development of Trichostrongylus tenuis as third-stage larvae in red grouse. Research in Veterinary Science 48:59–63. Shaw, J. L., and R. Moss. 1989a. The role of parasite fecundity and longevity in the success of Trichostrongylus tenuis in low density red grouse populations. Parasitology 99:253–258. Shaw, J. L., and R. Moss. 1989b. Factors affecting the establishment of the caecal threadworm Trichostrongylus tenuis in red grouse (Lagopus lagopus scoticus). Parasitology 99:259–264. Shaw, J. L., and R. Moss. 1990. Effects of the cecal nematode Trichostrongylus tenuis on egg-laying by
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captive red grouse. Research in Veterinary Science 48(1):59–63. Shaw, J. L., R. Moss, and A. W. Pike. 1989. Development and survival of the free-living stages of Trichostrongylus tenuis, a cecal parasite of red grouse Lagopus lagopus scoticus. Parasitology 99:105–113. Shipley, A. E. 1909. Internal parasites of birds allied to the grouse. Proceedings of the Zoological Society of London 2:363. Svoboda, J. 1992. Seasonal dynamics of the occurrence of gastrointestinal parasites in domestic-fowl. Veterinarni Medicina 37:543–547. Tompkins, D. M., and M. Begon. 1999. Parasites can regulate wildlife populations. Parasitology Today 15: 311–313. Watson, H., and J. L. Shaw. 1991. Parasites and Scottish ptarmigan numbers. Oecologia 88:359–361. Watson, H., D. L. Lee, and P. J. Hudson. 1987. The effect of Trichostrongylus tenuis on the caecal mucosa of young, old and anthelmintic-treated wild red grouse, Lagopus lagopus scoticus. Parasitology 94:405–411. Watson, H., D. L. Lee, and P. J. Hudson. 1988. Primary and secondary infections of the domestic chicken with Trichostrongylus tenuis (Nematoda), a parasite of red grouse, with observations on the effect on the caecal mucosa. Parasitology 97:89–99. Webster, L. M. I. 2005. The Effects of Gene Flow on Local Adaptation in a Natural Host–Parasite System. Division of Environmental and Evolutionary Biology, Institute for Biomedical and Life Sciences, University of Glasgow, Glasgow, UK. Webster, L. M. I., P. C. D. Johnson, A. Adam, B. K. Mable, and L. F. Keller. 2007. Macrogeographic populations structure in a parasitic nematode with avian hosts. Veterinary Parasitology 144:93–103. Wehr, E. E. 1971. Nematodes. In Infectious and Parasitic Diseases of Wild Birds, J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (eds). Iowa State University Press, Ames, IA, pp. 185–233. Wilson, G. R. 1983. The prevalence of caecal threadworms (Trichostrongylus tenuis) in red grouse (Lagopus lagopus scoticus). Oecologica 58:265– 268.
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18 Dispharynx, Echinuria, and Streptocara Ramon A. Carreno INTRODUCTION Nematodes of the genera Dispharynx, Echinuria, and Streptocara are parasitic in the proventriculus and gizzard of many avian taxa. All three are spirurid nematodes belonging to the superfamily Acuarioidea, the acuarioid nematodes. The acuarioids are characterized by their large pseudolabia and modified cuticular structures at the anterior end known as cordons (Figure 18.1). Twenty-eight acuarioid genera were listed by Chabaud (1975) and most are parasites of birds. Of the three genera that are discussed in this chapter, several species exist that are significant pathogens. Dispharynx nasuta is pathogenic mostly in passerine birds and Galliformes, while Streptocara species and Echinuria uncinata are pathogens primarily of waterfowl.
America (Brazil). Streptocara crassicauda has been reported from Asia (Azerbaijan, China, Kazakhstan, Russia, Uzbekistan), Australia, Europe (Austria, Bulgaria, Denmark, France, Italy, the Netherlands, Norway, Spain, UK), and North America (Canada, US). There is evidence that some acuarioid nematodes such as those parasitizing shorebirds have distinct distributions that correspond to migratory flyways (Anderson et al. 1996). However, pathogenic species of Echinuria, Streptocara, and Dispharynx are cosmopolitan in their distribution, probably because of their spread through domestic and migratory birds and because of the abundance of possible intermediate hosts that can support development of the parasites.
HOST RANGE AND DISTRIBUTION Streptocara spp. and Echinuria spp. are parasitic primarily in waterfowl while D. nasuta is most frequently reported from the Galliformes, Columbiformes, and Passeriformes (Table 18.1). Dispharynx nasuta is the most widespread of the three and reports of Echinuria spp. and Streptocara spp. from Africa and South America are rare. Dispharynx nasuta has been reported from Africa (Algeria, Congo, Egypt, Morocco, South Africa, Tunisia, Zimbabwe), Asia (China, Georgia, India, Kazakhstan, Pakistan, Russia, Turkestan, Uzbekistan), Australia, Central America (Costa Rica, Guatemala), Cuba, Europe (Austria, Denmark, France, Italy, Spain), Guam, North America (Canada, US), Puerto Rico, and South America (Argentina, Brazil, Ecuador, Venezuela). Echinuria uncinata has been reported from Africa (Algeria), Asia (Afghanistan, Azerbaijan, India, Japan, Russia), Australia, Europe (Czech Republic, France, Germany, Italy, Poland, Spain, UK), New Zealand, North America (Canada, US), and South
SYNONYMS Gizzard worms, proventricular worms, acuariosis, dispharagosis, dispharyngosis, dispharynxiasis, grouse disease.
DISPHARYNX
HISTORY The first description of D. nasuta was from the proventriculus of a House Sparrow (Passer domesticus) by Rudolphi in Austria (Goble and Kutz 1945). Both this and similar species were placed in various genera and subgenera until the genus Dispharynx was eventually erected by Skrjabin (1916). Piana (1897) first reported the intermediate hosts, terrestrial isopods (sow bugs or pillbugs), and the life cycle of D. nasuta has been further studied throughout the twentieth century (Cram 1931; Birova et al. 1974a, b; Nagy et al. 1977). The pathogenicity of D. nasuta (often recorded as Dispharynx spiralis) has been recognized at least since the early twentieth century. Early descriptions of dispharynxiasis were reported in wild game birds such
326 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Dispharynx, Echinuria, and Streptocara
(b) (a)
(c)
c1
co d
co
p
Figure 18.1. (a) Dispharynx nasuta, male, anterior, from a Rock Pigeon (Columba livia). Note the recurring cordons (co) and absence of cuticular spines. Scale bar = 50 μm. (b) Echinuria uncinata, male, anterior, from a Mute Swan (Cygnus olor). Note the cordons (co) and cuticular spines (sp). Scale bar = 100 μm. (c) Streptocara crassicauda, female, anterior from a King Eider (Somateria spectabilis). Note the collarette (cl), prominent deirids (d), and somewhat inflated cuticle between the collarette and the deirids. Scale bar = 25 μm. as grouse, and in recent years, outbreaks of the disease have also occurred in private collections, outdoor poultry farms, and in zoos. ETIOLOGY Taxonomic Synonyms: Dispharynx nasuta (Rudolphi 1819) Stiles and Hassall, 1920; Spiroptera nasuta Rudolphi, 1819; Dispharagus spiralis Molin, 1858; Acuaria spiralis (Molin 1858) Railliet, Henry, and Sisoff, 1912; Dispharagus nasutus (Rud. 1819) Dujardin, 1844; Filaria nasuta (Rud. 1819) Schneider, 1866; Dispharagus tentaculatus Colucci, 1893; Dispharagus spiralis columbae Bridr´e, 1910; Acuaria nasuta (Rud. 1819) Railliet, Henry, and Sisoff, 1912; Cheilospirura nasuta (Rud. 1819) Ransom, 1916; Dispharynx stonae Harwood, 1933. The species of Dispharynx comprise a genus of acuarioid nematodes that includes D. nasuta, the spiral stomach worm. About 15 species of Dispharynx have been described, and D. nasuta is considered to be the most pathogenic of these.
327
Adults of D. nasuta have prominent cordons that, at their distal ends, curve back toward the anterior of the worm in a recurrent pattern (Figure 18.1a). The posterior ends of the males are often spirally coiled. Males are about 5 mm long and females are 5–9 mm in length. The phylogenetic relationships of the Acuarioidea have not been assessed using both morphological and molecular data. Consequently, the sister groups of Dispharynx species and their precise phylogenetic position within the spirurids are unknown. EPIZOOTIOLOGY The life cycle of D. nasuta requires development of larvae in intermediate hosts, usually in various terrestrial isopods commonly known as “pillbugs.” Porcellio scaber, Armadillidium vulgare, and Oniscus species (Anderson 2000) as well as Venezillo evergladensis and Oscelloscia floridana (Rickard 1985) are suitable intermediate hosts. Eggs of D. nasuta from Ruffed Grouse (Bonasa umbellus) undergo development to third-stage larvae in 26 days when fed to P. scaber and A. vulgare (Cram 1931) and 16 days when fed to Porcellionides pruinosus from Cuba (Birova et al. 1974a). Temperature is a likely factor that influences development of the larvae. Infected isopods behave differently from uninfected isopods and are observed at greater frequencies on light-colored surfaces (Moore and Lasswell 1986). This altered behavior may make infected isopods more susceptible to predation by definitive avian hosts. Following ingestion of an infected isopod, full development to adults occurs in 27 days in Ruffed Grouse, Northern Bobwhite (Colinus virginianus), and Rock Pigeons (Columba livia) according to Cram (1931), and 21 days in domestic chickens (Gallus gallus) (Birova et al. 1974b). Ingested infective larvae spend a short period of time (up to 15 days) in the lumen of the proventriculus before penetrating into the mucosa (Nagy et al. 1977). It is at the latter stage when pathogenic effects are induced. In experimental infections of domestic turkeys with eggs of D. nasuta originating from Boat-tailed Grackles (Quiscalus major), Blue Jays (Cyanocitta cristata), Northern Cardinals (Cardinalis cardinalis), American Crows (Corvus brachyrhynchos), and Wild Turkeys (Meleagris gallopavo), eggs were not detected in the feces of infected turkeys until 42 days postinfection (PI). An experimentally infected grackle became patent at 32 days PI (Rickard 1985). CLINICAL SIGNS Young birds with high intensities of D. nasuta exhibit droopiness, inactivity, and weight loss (Cram 1928; Hwang et al. 1961; Nagy et al. 1977). They undergo
Arctic Loon (Gavia arctica) Red-throated Loon (Gavia stellata) Western Grebe (Aechmophorus occidentalis) Horned Grebe (Podiceps auritus) Great Crested Grebe (Podiceps cristatus) Red-necked Grebe (Podiceps grisegena) Great Cormorant (Phalacrocorax carbo) Great Bittern (Botaurus stellaris) Chilean Flamingo (Phoenicopterus chilensis) Fulvous Whistling-Duck (Dendrocygna bicolor) White-faced Whistling-Duck (Dendrocygna viduata) West Indian Whistling-Duck (Dendrocygna arborea) Mute Swan (Cygnus olor) Tundra Swan (Cygnus columbianus) Whooper Swan (Cygnus cygnus) Black-necked Swan (Cygnus melancoryphus) Black Swan (Cygnus atratus) Trumpeter Swan (Cygnus buccinator) Greater White-fronted Goose (Anser albifrons) Swan Goose (Anser cygnoides) Emperor Goose (Chen canagica) Canada Goose (Branta canadensis) Barnacle Goose (Branta leucopsis) Brant (Branta bernicla) Red-breasted Goose (Branta ruficollis) Hawaiian Goose (Branta sandvicensis)
Gaviiformes
— + + — — + — + — — — — — +
+ — + + + + + + + + + + + + — — — — — — — — — — — — — —
328
— — — — — — — + — — — — — —
—
— — — — — — — — + — —
+ + + + + + + + — — — —
SI
SC
+
— — — — — — — — — + +
EU
—
— — — — — — — — — — —
DN
McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) Wood (1974) McDonald (1969) McDonald (1969) Wood (1974) McDonald (1969) Bailey and Black (1995) and Lapage (1961)
Wood (1974)
McDonald (1969) McDonald (1969) Denny (1969) Denny (1969) McDonald (1969) Denny (1969) McDonald (1969) McDonald (1969) Fox et al. (1974) Wood (1974) McDonald (1969)
Reference
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Pelecaniformes Ciconiiformes Phoenicopteriformes Anseriformes
Podicipediformes
Host species
Host order
Table 18.1. Host distribution of Dispharynx nasuta (DN), Echinuria uncinata (EU), Streptocara crassicauda (SC), and Streptocara incognita (SI).
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— — — — — — — — — — — — — — — — — — — — — — — — — — — —
Orinoco Goose (Neochen jubata) Egyptian Goose (Alopochen aegyptiaca) Ruddy Shelduck (Tadorna ferruginea) Muscovy Duck (Cairina moschata) White-winged Duck (Cairina scutulata) Comb Duck (Sarkidiornis melanotos) Hartlaub’s Duck (Pteronetta hartlaubii) Cotton Pygmy-Goose (Nettapus coromandelianus) African Pygmy-Goose (Nettapus auritus) Ringed Teal (Callonetta leucophrys) Mandarin Duck (Aix galericulata) Wood Duck (Aix sponsa) Blue Duck (Hymenolaimus malacorhynchos) Northern Pintail (Anas acuta) White-cheeked Pintail (Anas bahamensis) American Wigeon (Anas americana)
Eurasian Wigeon (Anas penelope) Chiloe Wigeon (Anas sibilatrix) Cape Teal (Anas capensis) Chestnut Teal (Anas castanea) Cinnamon Teal (Anas cyanoptera) Northern Shoveler (Anas clypeata) Blue-winged Teal (Anas discors)
Baikal Teal (Anas formosa) Falcated Duck (Anas falcata) Yellow-billed Pintail (Anas georgica) Mallard (Anas platyrhynchos) Silver Teal (Anas versicolor)
— — — + —
+ — — + —
McDonald (1969) Wood (1974) McDonald (1969) McDonald (1969) and Mason (1988) Wood (1974) Wood (1974) McDonald (1969) Wood (1974) Wood (1974) Wood (1974) McDonald (1969) McDonald (1969) Wood (1974) McDonald (1969) Lapage (1961) McDonald (1969) and McLaughlin and McGurk (1987) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) Lapage (1961) and McDonald (1969) McDonald (1969) McDonald (1969) and McLaughlin and McGurk (1987) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) Wood (1974) (continues)
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329 — + + + +
— — — — — — —
+ — — — + + +
+ + + + + + +
— — — — — — — — — — — — — — — —
— — + + — — — — — — — — — + — +
+ + + + + + + + + + + + + + + +
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Host order
— — — — — — — — — — — — — —
Ferruginous Pochard (Aythya nyroca) Tufted Duck (Aythya fuligula) Canvas back (Aythya valisineria) Common Eider (Somateria mollissima) King Eider (Somateria spectabilis) Harlequin Duck (Histrionicus histrionicus) Long-tailed Duck (Clangula hyemalis) White-winged Scoter (Melanitta fusca) Black Scoter (Melanitta nigra) Surf Scoter (Melanitta perspicillata) Bufflehead (Bucephala albeola)
Barrow’s Goldeneye (Bucephala islandica) Smew (Mergellus albellus) Hooded Merganser (Lophodytes cucullatus)
DN — — — — — — — — — — — — — —
Host species
Garganey (Anas querquedula) American Black Duck (Anas rubripes) African Black Duck (Anas sparsa) Pacific Black Duck (Anas superciliosa) Philippine Duck (Anas luzonica) Gadwall (Anas strepera) Spectacled Duck (Anas specularis) Yellow-billed Duck (Anas undulata) Red-crested Pochard (Netta rufina) Rosy-Billed Pochard (Netta peposaca) Lesser Scaup (Aythya affinis) White-eyed Duck (Aythya australis) Greater Scaup (Aythya marila) Redhead (Aythya americana)
Table 18.1. (Continued)
— — — — — — — — — — — + — —
+ + + + + + + + + + +
330 + + + — + —
McDonald (1969) McDonald (1969) Lapage (1961) Clark (1977) Wood (1974) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) da Silveira et al. (2006) McDonald (1969) Wood (1974) McDonald (1969) McDonald (1969) and McLaughlin and McGurk (1987) McDonald (1969) McDonald (1969) and Wood (1974) McDonald (1969) McDonald (1969) McDonald (1969); unpublished observations Wood (1974) and McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) and McLaughlin and McGurk (1987) McDonald (1969) McDonald (1969) McDonald (1969)
Reference September 29, 2008
— + + + + + — — — — +
— — — — — — — — — — — — — —
+ — — — — + — — + — + — + +
+ + + + + + + + + + + + + +
SI
SC
EU
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Charadriiformes
— — — — — — — — — — — — — — — — — — — — — — —
+ + + + — + + + + + + + + + + + + — + — — — —
Plain Chachalaca (Ortalis vetula) Wild Turkey (Meleagris gallopavo) Dusky Grouse (Dendragapus obscurus) Ruffed Grouse (Bonasa umbellus) Sharp-tailed Grouse (Tympanuchus phasianellus) California Quail (Callipepla californica) Northern Bobwhite (Colinus virginianus) Barbary Partridge (Alectoris barbara) Gray Partridge (Perdix perdix) Common Quail (Coturnix coturnix) Red Junglefowl (Gallus gallus)
Ring-necked Pheasant (Phasianus colchicus) Golden Pheasant (Chrysolophus pictus) Indian Peafowl (Pavo cristatus) Helmeted guineafowl (Numida meleagris)
Whooping Crane (Grus americana) Sandhill Crane (Grus canadensis) Eurasian Coot (Fulica atra) African Jacana (Actophilornis africanus) Eurasian Oystercatcher (Haematopus ostralegus) Northern Lapwing (Vanellus vanellus) Snowy Plover (Charadrius alexandrinus) Common Greenshank (Tringa nebularia)
— — — — — — — —
+ + +
— — — —
— — — — — —
— — — — —
— — +
— — + — +
— — — +
— — — — — +
— — — — +
+ + +
McDonald (1969) McDonald (1969) McDonald (1969) (continues)
Moore et al. (1988) Goble and Kutz (1945) Goble and Kutz (1945) Goble and Kutz (1945) Baruˇs and Sonin (1983) Goble and Kutz (1945) and McDonald (1969) Goble and Kutz (1945) Goble and Kutz (1945) Goble and Kutz (1945) Goble and Kutz (1945) and McDonald (1969) Spalding et al. (1996) Forrester et al. (1975) McDonald (1969) Schulman et al. (1992) Borgsteede et al. (1988)
Wood (1974) and McDonald (1969) McDonald (1969) McLaughlin and McGurk (1987) and McDonald (1969) Christensen and Pence (1977) Goble and Kutz (1945) Bendell (1955) Goble and Kutz (1945) McDonald (1969)
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Gruiformes
Galliformes
+ — +
— — —
Common Merganser (Mergus merganser) Red-breasted Merganser (Mergus serrator) Ruddy Duck (Oxyura jamaicensis)
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Passeriformes
Psittaciformes Cuculiformes Coraciiformes Piciformes
— — + — — — — — — — — — — — — — — — — — — — — — —
— — — — — — — — — — — — + + + + + + + + + + + + +
Common Redshank (Tringa totanus) Spotted Sandpiper (Actitis macularius) Ruff (Philomachus pugnax) European Herring Gull (Larus argentatus) Mew Gull (Larus canus) Great Black-headed Gull (Larus ichthyaetus) Thayer’s Gull (Larus thayeri) Common Tern (Sterna hirundo) Common Murre (Uria aalge) Razorbill (Alca torda) Pigeon Guillemot (Cepphus columba) Horned Puffin (Fratercula corniculata) Rock Pigeon (Columba livia) Mourning Dove (Zenaida macroura) Luzon Bleeding-heart (Gallicolumba luzonica) Alexandra’s Parrot (Polytelis alexandrae) Smooth-billed Ani (Crotophaga ani) European Roller (Coracias garrulus) Red-headed Woodpecker (Melanerpes erythrocephalus) Yellow-bellied Sapsucker (Sphyrapicus varius) Northern Flicker (Colaptes auratus) Least Flycatcher (Empidonax minimus) Carolina Wren (Thryothorus ludovicianus) Long-billed Gnatwren (Ramphocaenus melanurus) Gray Catbird (Dumetella carolinensis)
EU
DN
Host species
—
—
— — — — —
— — — — — — — — — — — — — — — — — — —
+ + + + + + + + + + + + — — — — — — — — — — — —
SI
SC
Goble and Kutz (1945)
Baruˇs (1971) Bolette (1998)c Bolette (1998)a Goble and Kutz (1945) Zhang et al. (2004)
McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) Gibson (1968) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) Goble and Kutz (1945) Forrester et al. (1983) Lindquist and Strafuss (1980) Bolette (1998)b Macko et al. (1974) Goble and Kutz (1945) Cooper (1974)
Reference
September 29, 2008
Columbiformes
Host order
Table 18.1. (Continued)
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— — — — — — — — — — — — — — — — — — — — —
+ + + + + + + + + + + + + + + + + +
333 + + + — — —
— — — —
— — — — — — — — — — — — — —
— — —
— — — —
— — — — — — — — — — — — — —
Rickard (1985) Baruˇs and Garrido (1968) Goble and Kutz (1945)
Rickard (1985) Zhang et al. (2004) Baruˇs and Garrido (1968) Zhang et al. (2004)
Macko et al. (1974) Goble and Kutz (1945) Goble and Kutz (1945) Zhang et al. (2004) Rickard (1985) Kinsella (1974) Goble and Kutz (1945) Baruˇs and Garrido (1968) Goble and Kutz (1945) Goble and Kutz (1945) Baruˇs and Garrido (1968) Barus and Garrido (1968) Zhang et al. (2004) Zhang et al. (2004)
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Geographic range may include domestic or captive host populations. Host names follow Clements (2000).
Northern Mockingbird (Mimus polyglottos) Eastern Bluebird (Sialia sialis) American Robin (Turdus migratorius) Clay-colored Robin (Turdus grayi) Blue Jay (Cyanocitta cristata) Florida Scrub-Jay (Aphelocoma coerulescens) American Crow (Corvus brachyrhynchos) Cuban Crow (Corvus nasicus) European Starling (Sturnus vulgaris) House Sparrow (Passer domesticus) Prairie Warbler (Dendroica discolor) Blackburnian Warbler (Dendroica fusca) Tennessee Warbler (Vermivora peregrina) Gray-crowned Yellowthroat (Geothlypis poliocephala) Northern Cardinal (Cardinalis cardinalis) Blue-gray Tanager (Thraupis episcopus) Rose-breasted Grosbeak (Pheucticus ludovicianus) Black-faced Grosbeak (Caryothraustes poliogaster) Boat-tailed Grackle (Quiscalus major) Greater Antillean Grackle (Quiscalus niger) Brown-headed Cowbird (Molothrus ater)
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Parasitic Diseases of Wild Birds
retarded development, yet retain a ravenous appetite. When present in large numbers, D. nasuta may inhibit growth and cause anorexia and inability to thrive. Emaciation and death may follow (Cram 1928; Shanthikumar 1987). PATHOGENESIS AND PATHOLOGY Development of both juvenile and adult specimens of D. nasuta in the proventriculus causes dispharynxiasis. Lesions are produced when the worms bury their heads into the lamina propria of the proventriculus, causing an inflammatory response that leads to thickening of the mucosa and functional obstruction of the digestive tract (Schulman et al. 1992). This inflammatory response can result in death by starvation due to decreased digestive capability (Blasdel and Lasswell 1986). At necropsy, characteristic lesions consist of ulceration and inflammation of the proventriculus, which may be as large as the gizzard (Shanthikumar 1987). The mucosa is often completely destroyed, and parasites are found buried in a mass of degenerated and necrotic tissue (Shanthikumar 1987). The presence of D. nasuta can cause swelling, ulceration, cellular infiltration, and destruction of the proventricular glands (Figure 18.2). As much as a threefold increase in the size of the proventriculus has been reported from infected Boattailed Grackles and other birds (Rickard 1985). In the vicinity of the worms, multifocal petechial hemorrhages are found, the mucosa of the proventriculus is thickened, and the worms are covered in an acellular, catarrhal exudate (Hwang et al. 1961; Rickard 1985; Schulman et al. 1992). The pathogenesis of D. nasuta in domestic chickens occurs in three phases that coincide with development of the worms (Nagy et al. 1977). In the first phase, 1–11 days PI, petechial hemorrhages and light hyperplastic processes are evident. In the second, 12–22 days PI, the proventriculus begins to increase in size, and congestion and hemorrhage become more marked. In the third phase at 23 days PI when mature females begin to shed eggs, the serosal surface of the proventriculus is covered with whitish areas from which a yellowish exudate emanates. These whitish regions have been described as necrotic ulcers and contain developing juveniles (Ramaswamy and Sundaram 1985). In addition to the pathogenic effects described above, an extreme papillomatous proliferation of the mucosal surface has been seen in a naturally infected American Crow, a naturally infected Boat-tailed Grackle, and two naturally infected Northern Cardinals. In one case, proliferative tissue almost completely occluded the proventricular lumen (Rickard 1985).
Figure 18.2. Dispharynx nasuta. Proventriculus of a Rock Pigeon (Columba livia) with parasites in situ (arrows). Reproduced with permission from Avian Diseases (Hwang et al. 1961).
In histological sections, extensive desquamation of the superficial mucosa, hypersecretion of mucus, and cellular masses containing bacteria can be observed. At 8 days PI in domestic chickens, a severe nonkeratinizing squamous cell metaplasia of the lining epithelium occurs in the lamina propria, and proventricular glands containing juvenile worms are surrounded by necrotic cell debris and desquamated epithelial cells (Ramaswamy and Sundaram 1985). By 14 days PI, deep ulcers, necrotic foci, lymphoid hyperplasia, and severe cellular infiltration with granulocytes occur in the lamina propria. Many of these histological changes, including the occurrence of a mononuclear transmural infiltrate through the proventriculus, were also evident in experimentally infected domestic turkeys and a Brown-headed Cowbird (Molothrus ater) (Rickard 1985).
DIAGNOSIS Both demonstration of the eggs of D. nasuta in feces and the presence of clinical signs can be used to make
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Dispharynx, Echinuria, and Streptocara an antemortem diagnosis. The eggs are ellipsoidal, 36– 40 μm long and 21 μm wide, and are embryonated when passed in feces. These features distinguish eggs of D. nasuta from those of many other parasitic nematodes such as capillarids and ascarids. However, the possibility of infection with other spirurid nematodes cannot be excluded based on this type of egg morphology. Necropsy is necessary to recover adults from the proventriculus and to verify the presence of recurrent cordons. Pathogenic effects of the worms can occur before the infections become patent and eggs may be passed sporadically. A possible method for antemortem diagnosis involves contrast radiography, whereby distinct radiographic changes of increased proventricular to ventricular size ratio and proventricular filling can be detected (Schulman et al. 1992). Postmortem observation of the characteristic proventriculitis and the presence of adults and juveniles of D. nasuta (Figure 18.2) can confirm diagnosis. IMMUNITY The immune response to D. nasuta has not been studied in detail. Experimental infections in domestic and wild birds have shown that some hosts appear to be more susceptible to dispharynxiasis than do others. This might suggest the presence of different strains of D. nasuta, some of which are more pathogenic (Rickard 1985), or may be reflective of a longer host– parasite coevolution of D. nasuta in some hosts. Although survey data and information from published case reports are limited, infections appear to be more common in younger birds than in older ones. A list of infected galliform and passeriform birds by Goble and Kutz (1945) reported consistently higher prevalences of infection in juvenile birds than adults. Mechanisms of age resistance to infection, if present, are unknown.
ECHINURIA SYNONYMS Acuariasis, echinuriasis. HISTORY The genus Echinuria (family Acuariidae) was erected in 1912 by Soloviev, and E. uncinata (Rudolphi 1819) Soloviev, 1912 was eventually placed here (see list of synonyms below). The intermediate hosts of E. uncinata were discovered by Hamann (1893), who demonstrated development of the larvae in cladoceran crustaceans of the genus Daphnia. Descriptions of echinuriasis were given in the late nineteenth century
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(Hamann, 1893), and the disease has been documented frequently throughout the later half of the twentieth century (Buxton et al. 1952; Cornwell 1963; Gr¨afner and Graubmann 1967; Takla and Thiel 1983; Griffiths et al. 1985). In the latter period, studies of the disease using experimental infections with E. uncinata have provided additional information (Guilhon et al. 1971; Ould and Welch 1980). ETIOLOGY Taxonomic synonyms: Echinuria uncinata (Rudolphi 1819) Soloviev, 1912; Spiroptera uncinata Rudolphi, 1819; Filaria uncinata Schneider, 1866; Dispharagus uncinatus (Rud. 1819) Railliet, 1893; Acuaria (Hamannia) uncinata (Rud. 1819) Stiles and Hassall, 1912; Echinuria jugadornata Soloviev, 1912. Approximately 14–15 species of Echinuria are recognized (Ali, 1968; McDonald, 1974). Members of the genus occur in the proventriculus of avian hosts. Although various Echinuria spp. have been reported from other birds (mostly waterfowl), E. uncinata from anatids is the most significant pathogen of the genus. This species, like other members of the genus, has recurving cordons extending posteriorly from the anterior end and, unlike Dispharynx species, has cuticular spines running along the length of the body (Figure 18.1b). Adult males are 8–10 mm in length and females are 12–18 mm long. EPIZOOTIOLOGY The ingestion of intermediate hosts containing infective larvae of E. uncinata results in the development of the parasites in the proventriculus. Cladoceran crustaceans, usually of the genus Daphnia (e.g., D. pulex and D. magna), are suitable intermediate hosts for E. uncinata (Anderson 2000). In addition to species of Daphnia, Ceriodaphnia spp., Simocephalus vetulus, Gammarus lacustris, the ostracod Heterocypris incongruens, and the conchostracan Lynceus brachyurus have also been shown to support development of third-stage larvae (Misiura 1970; Austin and Welch 1972; Anderson 2000). On the basis of studies of the life cycle in domestic ducks, embryonated eggs are ingested by intermediate hosts and the larvae penetrate through the digestive tract and undergo development in the body cavity of the cladoceran intermediate hosts (Guilhon et al. 1971). The larvae molt once approximately 8–9 days after ingestion and molt again to the third stage after 14–15 days, depending on the temperature. In temperate climates, numbers of larvae in intermediate hosts reach their highest numbers in midsummer (Austin and Welch 1972). Upon ingestion of infected species of Daphnia by the avian
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definitive hosts, the third-stage larvae are liberated and disseminate throughout the proventriculus. Fully formed adults are noticeable after 45 days, and the prepatent period is approximately 30 days. In Mallards (Anas platyrhynchos), parasites are located under the soft mucosa of the proventriculus and gizzard, usually along the region connecting the two organs. Groups of worms occur in fibrotic nodules and large granulomas form after 40–50 days. Sexually mature male worms are found 30 days PI, and mature females shed viable eggs 40 days PI (Austin and Welch 1972). Some species of waterfowl are apparently more susceptible to infection with E. uncinata than others. On the basis of experimental infections of ducklings with infective larvae, Northern Shovelers (Anas clypeata), which feed frequently on Daphnia species, have very low susceptibility to infection (Austin and Welch 1972). Mallards, Gadwalls (Anas strepera), Common Eiders (Somateria mollissima), Northern Pintails (Anas acuta), and domestic geese are the most susceptible to infection with E. uncinata, while Northern Shovelers (Anas clypeata), Blue-winged Teal (Anas discors), Ruddy Ducks (Oxyura jamaicensis), and American Coots (Fulica americana) are generally resistant (Austin and Welch 1972). Experimental infections in Anas spp. generally yield higher average numbers of worms than in other experimentally infected waterfowl (Austin and Welch 1972). CLINICAL SIGNS Birds with infections of high intensity are usually in a weak and emaciated condition without apparent inappetance (Kock et al. 1987). Wild ducks have a
prominent keel as a result of severe emaciation, fading, discoloring, and poor grooming of the feathers, an inability to sustain flight, or to fly at all, and a near absence of wariness (Cornwell 1963). Gr¨afner and Graubmann (1967) described the clinical signs of infection as weakness, awkward gait, and diminished appetite. Throughout the progression of the disease, the animals reeled, crossed their legs, and eventually developed lameness and difficulty in breathing. Decreased intake of food occurred. Attempts to eat resulted in retching or choking, and the birds became completely emaciated and soon died. PATHOGENESIS AND PATHOLOGY The presence of the nematodes in the proventriculus results in a strong immune response that leads to formation of nodules that contain the worms. In heavy infections, these nodules can become so numerous that the proventriculus becomes occluded, preventing successful feeding. Infection with E. uncinata can result in the formation of encapsulated lesions in the proventriculus (Figure 18.3), usually at its junction with the esophagus (Kock et al. 1987). The overall size of the proventriculus increases, and the proventricular mucosae can be covered by thick, white, slimy exudates (Griffiths et al. 1985). The nodules often contain creamy white caseous material, and worms are often not present. The nodules eventually become granulomas. In fatal infections, the nodules become so large that the proventriculus becomes occluded, preventing the passage of food. In histological sections, abscess-like lesions in the wall of the proventriculus are evident. These lesions are lined by cells resembling macrophages and giant cells and surrounded by extensive proliferation
Figure 18.3. Echinuria uncinata. Ulcerative cyst in the proventriculus of a Trumpeter Swan (Cygnus buccinator). There is a caseous tissue reaction and some nematodes (arrow) are visible in situ. Reproduced with permission from Canadian Journal of Zoology (Cornwell 1963).
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Dispharynx, Echinuria, and Streptocara of fibrous tissue and inflammatory cells (Kock et al. 1987). There is some evidence of ulceration in the lining epithelium at the junction of the proventriculus and esophagus (Kock et al. 1987). DIAGNOSIS Methods for diagnosis of echinuriasis in live birds include the recovery of eggs from fecal samples. Eggs shed in the feces of infected birds are approximately 35 μm × 20 μm (Guilhon et al. 1971); however, eggs will not be present if the prepatent period has not been completed (Respaldiza et al. 1979). At postmortem, the characteristic enlarged proventriculus, proventricular lesions (0.5–2.5 cm nodules and granulomas), and presence of whole specimens of E. uncinata inside the nodules can confirm diagnosis. Species of Echinuria differ from most other acuarioid nematodes in having cuticular spines that extend posteriorad from the anterior end (Figure 18.1b). Other nematodes that may occur in the proventriculus include other acuarioid nematodes such as Streptocara spp. as well as Tetrameres spp. and the trichostrongyloid Amidostomum. Adult morphology, including extreme sexual dimorphism, absence of cordons, and different lesions can be used to differentiate Tetrameres infections from those of E. uncinata. Species of Streptocara and Amidostomum differ morphologically from Echinuria and are more often found in the gizzard.
STREPTOCARA SYNONYMS Streptocariasis. HISTORY Species of Streptocara are acuarioid nematodes in the family Seuratiinae that occur in the gizzard and occasionally the proventriculus and esophagus of waterfowl. Records of various species of Streptocara in waterfowl have been known since the nineteenth century. The life cycle remained unknown, however, until Garkavi (1949) established that amphipods (G. lacustris) are suitable intermediate hosts. There have been scattered reports of outbreaks of streptocariasis in waterfowl throughout the twentieth century, but some of the most detailed studies of the disease were carried out in the 1980s and early 1990s (Sterner and Stackhouse 1987; Mason 1988; Laberge and McLaughlin 1991). ETIOLOGY Taxonomic synonyms: Streptocara crassicauda (Creplin 1829) Skrjabin, 1916; Spiroptera crassicauda
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Creplin, 1829; Dispharagus crassicauda (Creplin 1829) Molin, 1860; Streptocara crassicauda crassicauda (Creplin 1829) Gibson, 1968; Spiroptera pectinifera Neumann, 1900; Streptocara pectinifera (Neumann 1900) Skrjabin, 1916; Streptocara crassicauda anseri Skrjabin, 1916; Streptocara crassicauda charadrii Skrjabin, 1916; Streptocara crassicauda skrjabini Liubimov, 1927. Adult Streptocara spp. have cordons that expand only on the cephalic region, forming a collarette (Figure 18.1c) (see also Chabaud, 1975). The deirids (cervical papillae) are located posterior to the cephalic region and have 5–9 teeth. Although up to 12 species have been listed in various monographs, only 4 species and 2 subspecies were included by Gibson (1968) in a review of the genus. Additional species have been described from various avian hosts since 1968. One of the common species that has been shown to be pathogenic is S. crassicauda. Males of these thin, rather delicate parasites are 3.4–5.1 mm long and females are 5.5–12.9 mm in length (Gibson 1968). The cuticle has an annular pattern and is slightly expanded at the cephalic end (Figure 18.1c). Another species of Streptocara, S. incognita Gibson, 1968, has also been shown to be pathogenic in waterfowl. This species lacks the inflated cuticle and annuli between the collarette and the cervical papillae are also different from those of S. crassicauda (Gibson 1968; McDonald 1974). EPIZOOTIOLOGY Definitive hosts become infected by ingesting amphipod intermediate hosts such as species of Gammarus and Hyalella (Anderson 2000). Eggs of S. crassicauda are shed in the feces and develop in various marine and freshwater amphipod crustaceans. Development from eggs to infective third-stage larvae occurs in 19– 25 days in G. lacustris, depending on the temperature (Garkavi 1949). Larvae from six species of fish were also infective to ducks (Kovalenko 1960). The amphipod, Hyalella azteca, also serves as a suitable intermediate host (Denny 1969; Laberge and McLaughlin 1989). First-stage larvae molt as early as 11 days PI at 18–20◦ C in these amphipods, with most molting after 15–17 days. Second-stage larvae molt as early as 15 days PI, with a peak on day 17 PI (Laberge and McLaughlin 1989). The earliest third-stage larvae are found 19 days PI. When infective larvae from H. azteca were fed to domestic ducks, female worms became gravid as early as 9 days PI. The first eggs detected in feces were observed 26 days PI (Laberge and McLaughlin 1989). The life cycle of S. incognita has not been determined, but is likely similar to that of S. crassicauda.
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In comparing infections in Blue-winged Teal, Gadwall, and Lesser Scaup (Aythya affinis), Laberge and McLaughlin (1991) suggested that dabbling ducks such as Blue-winged Teal and Gadwall are suitable hosts for S. crassicauda but play a minor role in maintaining transmission of the parasite. Experimentally infected ducklings have short-lived infections and lesions in Blue-winged Teal and Gadwall are larger than those in diving ducks, such as Lesser Scaup, that feed primarily on amphipods (Laberge and McLaughlin 1991). This suggests that dabbling ducks are more likely to become infected where they overlap in distribution with Lesser Scaup (McLaughlin and McGurk 1987; Laberge and McLaughlin 1991). The smaller lesions in Lesser Scaup may be attributed to a greater degree of tolerance to infection with Streptocara spp. Since fish can serve as paratenic hosts of S. crassicauda (Kovalenko 1960), other waterfowl, including fish-eating ducks, may have evolved a similar tolerance for the parasites. On the basis of the large number of fish-eating definitive hosts for S. crassicauda (Table 18.1), piscine paratenic hosts may play a significant role in transmitting infective larvae to these avian hosts. However, the role of paratenic hosts in the natural transmission of Streptocara to ducks and other birds is poorly understood. CLINICAL SIGNS Birds infected with Streptocara spp. may exhibit weakness, loss of appetite, and sneezing (Mason 1988). Weight loss and an inability to swallow have been observed as additional clinical signs (Dalton 1980). PATHOGENESIS AND PATHOLOGY Infective larvae of Streptocara spp. penetrate the cuticle of the gizzard and burrow into the mucosa. Their presence gives rise to local hemorrhage, ulceration, and necrosis (Boughton 1969). The burrowing of the worms into the mucosa results in local hemorrhage and later ulceration and necrosis of the tissues. Sloughing of the cornified layer of the gizzard can occur, leaving large areas unsuitable for grinding food (Sterner and Stackhouse 1987). Infections with Streptocara can cause a necrotizing esophagitis, an inability to swallow, and death from inanition (Dalton 1980). The presence of S. crassicauda in the gizzard and S. incognita in the gizzard, proventriculus, and esophagus can thus lead to destruction of tissue, inability of the host to feed properly, and death. One- to two-cm-diameter ulcers have been reported at the junction of the gizzard and proventriculus and ulcerative lesions were also found on the proventriculus and gizzard of Chilean Flamingos (Phoenicopterus
chilensis) infected with S. incognita (Fox et al. 1974). The primary lesion associated with infection with Streptocara spp. among a variety of species of ducks in Australia was a necrotic, diphtheritic, plaque-like mass overlying, and adherent to the larynx and pharynx and adjacent mucosae (Mason 1988). In these infections, the laryngeal opening was totally or severely obstructed by the plaque-like mass, and death was considered to be a result of asphyxiation. When parts of the digestive tract are damaged, the lesions can cause difficulty in swallowing and result in starvation. In histological sections from Mallards infected with S. crassicauda and S. incognita, worms are surrounded by varying amounts of necrotic debris and a cellular infiltrate consisting of mononuclear and multinucleated phagocytes, lymphocytes, and heterophiles (Sterner and Stackhouse 1987). In addition, granulomatous inflammation surrounding amorphous debris was observed in the gizzard musculature. In Chilean Flamingos infected with S. incognita, necrotic cellular debris and polymorphonuclear cells formed a pseudomembrane over eroded portions of the tissues at the junction of the gizzard and proventriculus (Fox et al. 1974). DIAGNOSIS A combination of clinical signs and the presence of spirurid-type eggs in feces may be used for antemortem diagnosis of streptocariasis. The eggs of S. crassicauda are 37 μm long and 18 μm wide and those of S. incognita are 36 μm long and 20 μm wide (Gibson 1968). However, lesions may develop before infections become patent, making the detection of eggs in feces unreliable as a diagnostic method. At postmortem, the nematodes can be extracted from lesions and identified by their characteristic morphology (Figure 18.1c). The morphology of adult species of Streptocara is characteristic and distinguishable from most of these other nematodes by the presence of a collarette at the cephalic end. With the exception of several closely related acuarioid nematodes, the collarette is absent in these other groups (Figure 18.1c). There is some confusion regarding which species of Streptocara cause disease. Streptocara crassicauda appears to be limited to the gizzard, while S. incognita is found in the esophagus and proventriculus in addition to the gizzard. On the basis of experimental infections in three species of ducks, Laberge and McLaughlin (1991) believe that S. crassicauda is unlikely to cause the esophageal and proventricular lesions reported in other studies and that S. incognita is more likely to be responsible for lesions occurring outside the gizzard. A recent report of parasitic esophagitis in Muscovy Ducks (Cairina moschata) in Italy caused by S. incognita supports this
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Dispharynx, Echinuria, and Streptocara hypothesis (Bano et al. 2005). Other nematode parasites that may occur in the gizzard include other acuarioid nematodes as well as the trichostrongyloid Amidostomum and the ascaridoid parasites, Contracaecum and Porrocaecum. Other nematodes that may also occur in the proventriculus and esophagus include ascarids and species of Cyrnea, Echinuria, Tetrameres, and Capillaria.
DISPHARYNX, ECHINURIA, AND STREPTOCARA DOMESTIC ANIMAL HEALTH CONCERNS Dispharynx nasuta is potentially pathogenic in domestic poultry. In experimentally infected chickens, Ramaswamy and Sundaram (1985) observed a mortality rate of 29.16% in chicks. Mortality due to dispharynxiasis was also observed in experimentally infected chickens in Cuba (Nagy et al. 1977). Echinuria uncinata is common in waterfowl occurring in the wild, and the cross-transmission to captive waterfowl populations occurring in outdoor facilities is a serious threat. There have been numerous outbreaks in domestic ducks that have usually resulted in the deaths of the animals (Buxton et al. 1952; Gr¨afner and Graubmann 1967; Respaldiza et al. 1979; Griffiths et al. 1985; Kock et al. 1987). This susceptibility also exists for outbreaks of streptocariasis (Boughton 1969; Fox et al. 1974; Mason 1988). Often, a search in the water of aviaries in which enzootics of streptocariasis have occurred will reveal the crustacean intermediate hosts. WILDLIFE POPULATION IMPACTS Dispharynxiasis can cause mortality in wild birds such as Ruffed Grouse (Goble and Kutz 1945). Dispharynx is considered to be one of the most important parasites of game birds in the eastern US (Wehr 1971), and fatal infections have been reported. The widespread range of D. nasuta is of particular importance in the management of declining wild species that may be susceptible to dispharynxiasis. Infections have recently been reported in experimentally introduced Whooping Cranes (Grus americana) in Florida (Spalding et al. 1996). Although infections were of low prevalence and intensity and no pathogenic effects were noted, the potential for Dispharynx to cause pathology in a broad range of avian hosts is a cause for concern. Careful monitoring of these birds and other wild populations is warranted. A species of Dispharynx has also been reported from domestic chickens in the Galapagos Islands, raising concerns about its potential transmission to endemic host species (Gottdenker et al. 2005). Species of Streptocara and Echinuria have similar potential for causing disease in threatened or en-
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dangered populations of wild birds. The endangered Laysan Duck (Anas laysanensis) with a single population in the northwestern Hawaiian Islands is susceptible to infection with E. uncinata. Mortality caused by this parasite was recently reported (Work et al. 2004). TREATMENT AND CONTROL There are few good options for controlling infections with Dispharynx, Echinuria, and Streptocara in freeliving and captive populations, since definitive and intermediate hosts have a cosmopolitan range. Although D. nasuta, E. uncinata, and S. crassicauda are common and widespread in their natural hosts, there is a serious risk of disease and mortality when captive birds in zoos, private collections, and other aviaries are exposed to these parasites. Dispharynx nasuta can be transmitted to many captive, valuable, and endangered birds (Lindquist and Strafuss 1980; Stauber and Schussman 1985; Blasdel and Lasswell 1986; Bolette 1998a, b). Parasite eggs shed from locally infected wild birds can result in the spread of infection to captive animals through the isopod intermediate hosts. Preventing the spread of acuarioid nematodes in domestic birds and in zoo collections is dependent on minimizing exposure to infected wild birds and in reducing or eliminating intermediate hosts. House Sparrows and Rock Pigeons are hosts of D. nasuta, and both are often found in zoos with outdoor displays and in the vicinity of larger aviaries. As E. uncinata is globally widespread, it could occur in any zoos or private collections of waterfowl that are kept on open water where wild birds can deposit eggs of E. uncinata and where suitable intermediate hosts, particularly Daphnia species, can serve as intermediate hosts. Similarly, the overlap of captive and domestic birds with wild waterfowl should be avoided in order to prevent the introduction of Streptocara species into the captive populations. Controlling outbreaks of dispharynxiasis in captive birds is dependent on elimination of the isopod intermediate hosts. This is particularly difficult in outdoor aviaries. However, the use of mild insecticides such as pyrethrin preparations may help to reduce their numbers (Shanthikumar 1987). Various anthelmintics can be used to treat dispharynxiasis. Schulman et al. (1992) treated infected African jacanas (Actophilornis africanus) successfully with ivermectin. Shedding of eggs stopped and preventricular swelling resolved following treatment. Other anthelmintics that are effective include thiabendazole, levamisole, and mebendazole (Stauber and Schussman 1985; Shanthikumar 1987). One of the most reliable means of controlling echinuriasis in captive ducks is to reduce or eliminate the presence of the daphnid intermediate hosts in duck
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ponds and holding facilities. Species of Daphnia survive well in stagnant ponds and lakes with little or no current, and increasing the flow of water should reduce their numbers (Wood 1974). Adult E. uncinata can overwinter in ducks, and it may be possible for larvae in the intermediate hosts to also survive winter (Austin and Welch 1972). Therefore, elimination or reduction of Daphnia or removal of birds from infested areas can control this parasite in captive host populations. Early attempts to treat echinuriasis with anthelmintics were largely unsuccessful. More recently however, the treatment of Red-breasted Geese (Branta ruficollis) with ivermectin has shown promise (Kock et al. 1987). Additional treatment with antibiotics may control secondary bacterial infections, and granulomas may resolve after death of the worms. Eradication of crustaceans with copper sulfate treatment has been suggested for control of streptocariasis (Fox et al. 1974). In zoological collections, birds should be carefully screened for possible infection with Streptocara by examination of feces before they are introduced to a new facility. Various anthelmintics including fenbendazole, levamisole, and mebendazole may be efficacious in treating streptocariasis (Denev et al. 1977; Dalton 1980). Administration of a liquid gruel diet may also assist recovery. LITERATURE CITED Ali, M. M. 1968. Studies on spiruroid parasites of Indian birds. Part II. A new genus and five new species of Acuariidae, together with a key to the genus Echinuria. Journal of Helminthology 42:221– 242. Anderson, R. C. 2000. Nematode Parasites of Vertebrates. Their Development and Transmission, 2nd ed. CABI Publishing, CAB International, Wallingford, UK. Anderson, R. C., P. L. Wong, and C. M. Bartlett. 1996. The acuarioid and habronematoid nematodes (Acuarioidea, Habronematoidea) of the upper digestive tract of waders: A review of observations on their host and geographic distributions and transmission in marine environments. Parasite 4:303–312. Austin, F. G., and H. E. Welch. 1972. The occurrence, life cycle, and pathogenicity of Echinuria uncinata (Rudolphi, 1819) Soloviev, 1912 (Spirurida, Nematoda) in waterfowl at Delta, Manitoba. Canadian Journal of Zoology 50:385–393. Bailey, T., and J. Black. 1995. Parasites of wild and captive Nene Branta sandvicensis in Hawaii. Wildfowl 46:59–65. Bano, L., A. Natale, M. Vascellari, D. Comin, F. Mutinelli, and F. Agnoletti. 2005. First report of parasitic esophagitis by Streptocara incognita in
muscovy ducks (Cairina moschata domesticus) in Italy. Avian Diseases 49:298–300. Baruˇs, V. 1971. A survey of parasitic nematodes of piciform birds in Cuba. Folia Parasitologica 18:315–321. Baruˇs, V., and O. H. Garrido. 1968. Nematodes parasitic in birds of the order Passeriformes in Cuba. Folia Parasitologica 15:147–160. Baruˇs, V., and M. D. Sonin. 1983. Survey of nematodes parasitizing the genus Coturnix (Galliformes) in the Palaearctic region. Helminthologia 20:175–186. Bendell, J. F. 1955. Disease as a control of a population of blue grouse (Dendragapus obscurus fuliginosus (Ridgway) Candian Journal of Zoology 33:195–223. Birova, V., J. K. Macko, and L. Espaine. 1974a. The life cycle of Dispharynx nasuta (Rudolphi, 1819) in experimentally infested intermediate hosts in Cuba. Helminthologia 15:693–713. Birova, V., J. K. Macko, and D. Ovies. 1974b. The life cycle of Dispharynx nasuta (Rudolphi, 1819) in experimentally infested chickens in Cuba. Helminthologia 15:715–740. Blasdel, T., and J. Lasswell. 1986. Dispharynx nasuta in the Houston Zoological Gardens bird collection. In Proceedings of the Annual Meeting of the American Association of Zoo Veterinarians, Chicago, Illinois, M. S. Silberman and S. D. Silberman (eds), pp. 101–102. Bolette, D. P. 1998a. Dispharynxiasis in a least flycatcher, Empidonax minimus (Passeriformes: Tyrannidae), and a golden-breasted starling, Cosmopsarus regius (Passeriformes: Sturnidae). Journal of the Helminthological Society of Washington 65:117–118. Bolette, D. P. 1998b. Dispharynxiasis in a captive princess parrot. Journal of Wildlife Diseases 34:390–391. Bolette, D. P. 1998c. Gastrointestinal helminths of some yellow-shafted flickers, Colaptes auratus luteus (Aves: Picidae), from Allegheny County, Pennsylvania. Journal of the Helminthological Society of Washington 65:114–116. Borgsteede, F. H. M., E. Van Den Broek, and C. Swennen. 1988. Helminth parasites of the digestive tract of the oystercatcher, Haematopus ostralegus, in the Wadden Sea, The Netherlands. Netherlands Journal of Sea Research 22:171–174. Boughton, E. 1969. On the occurrence of oesophageal worms, Streptocara crassicauda, in ornamental ducks in Hampshire. Journal of Helminthology 43:273–280. Buxton, J. C., C. M. Ford, and I. B. Munro. 1952. Infestation of domestic ducks with Acuaria (Echinuria) uncinata. Veterinary Record 64:5–6. Chabaud, A. G. 1975. Keys to genera of the order Spirurida. Part 2. Spiruroidea, Habronematoidea, and
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Dispharynx, Echinuria, and Streptocara Acuarioidea. In CIH Keys to the Nematode Parasites of Vertebrates, No. 3, R. C. Anderson, A. G. Chabaud, and S. Willmott (eds). Commonwealth Agricultural Bureaux, Farnham Royal, Buckinghamshire, pp. 29–58. Christensen, Z. D., and D. B. Pence. 1977. Helminths of the plain chachalaca, Ortalis vetula macalli, from the South Rio Grande Valley. Journal of Parasitology 63:830. Clark, W. C. 1977. Echinuria uncinata in a duck. New Zealand Veterinary Journal 25:39. Clements, J. F. 2000. Birds of the World: A Checklist, 5th ed. Ibis Publishing, Vista, CA. Cooper, C. L. 1974. Occurrence of helminth parasites in avian hosts from South Bass Island, Ohio. Ohio Journal of Science 74:60–62. Cornwell, G. 1963. Observations on waterfowl mortality in southern Manitoba caused by Echinuria uncinata (Nematoda, Acuariidae). Canadian Journal of Zoology 41:699–703. Cram, E. B. 1928. Nematodes of pathological significance found in some economically important birds in North America. United States Department of Agriculture Technical Bulletin no. 49, Washington, DC. Cram, E. B. 1931. Developmental stages of some nematodes of the Spiruroidea parasitic in poultry and game birds. United States Department of Agriculture Technical Bulletin no. 227, Washington, DC. Dalton, P. J. 1980. Streptocara infestation of ducks. Veterinary Record 107:384. da Silveira, E. F., J. F. R. Amato, and S. B. Amato. 2006. Echinuria uncinata (Rudolphi) (Nematoda, Acuariidae) in Netta peposaca (Vieillot) (Aves, Anatidae) in South America. Revista Brasileira de Zoologia 23:520–528. Denev, I., R. Kostov, and I. Vasilev. 1977. Efficacy of fenbendazole against Amidostomum anseris and Streptocara crassicauda infections in geese. Veterinarna Sbirka 75:29–31. Denny, M. 1969. Life cycles of helminth parasites using Gammarus lacustris as an intermediate host in a Canadian lake. Parasitology 59:795–827. Forrester, D. J., A. O. Bush, and L. E. Williams, Jr. 1975. Parasites of Florida sandhill cranes, Grus canadensis pratensis. Journal of Parasitology 61:547–548. Forrester, D. J., J. A. Conti, J. D. Shamis, W. J. Bigler, and G. L. Hoff. 1983. Ecology of helminth parasitism of mourning doves in Florida. Proceedings of the Helminthological Society of Washington 50:143–152. Fox, J. G., S. B. Snyder, G. D. Schmidt, and L. H. Campbell. 1974. Infection with the nematode Streptocara incognita in the Chilean flamingo. Journal of Wildlife Diseases 10:66–69. Garkavi, B. L. 1949. A study of the life-cycle of the nematode Streptocara crassicauda (Creplin, 1829),
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parasitic in domestic and wild ducks (in Russian). Dokladi Akademii Nauk SSSR 65:421–424. Gibson, G. G. 1968. Species composition of the genus Streptocara Railliet et al., 1912 and the occurrence of these avian nematodes (Acuariidae) on the Canadian Pacific coast. Canadian Journal of Zoology 46:629–645. Goble, F. C., and H. L. Kutz. 1945. The genus Dispharynx (Nematoda: Acuariidae) in galliform and passeriform birds. Journal of Parasitology 31:323–331. Gottdenker, N. L., T. Walsh, H. Vargas, J. Merkel, G. U. Jim´enez, R. E. Miller, M. Dailey, and P. G. Parker. 2005. Assessing the risks of introduced chickens and their pathogens to native birds in the Gal´apagos Archipelago. Biological Conservation 126:429–439. Gr¨afner, G., and H. D. Graubmann. 1967. Echinuriose und Tetramerose bei Zoov¨ogeln. Erkrankungen der Zootiere. Verhandlungsbericht des IX. Internationalen Symposiums u¨ ber die Erkrankungen der Zootiere, Prague, May 31–June 4, 1967, pp. 171–173. Griffiths, G. L., D. Hopkins, R. H. Wroth, and W. Gaynor. 1985. Echinuria uncinata infestation in mute swan cygnets (Cygnus olor). Australian Veterinary Journal 62:132. Guilhon, J., A. Saratsiotis, and G. Jolivet. 1971. Echinuriose exp´erimentale du canard. Recueil de M´edecine V´et´erinaire 147:1–21. Hamann, O. 1893. Die Filarienseuche der Enten und der Zwischenwirt von Filaria uncinata R. Centralblatt f¨ur Bakteriologie und Parasitenkunde 14:555–557. Hwang, J. C., N. Tolgay, W. T. Shalkop, and D. S. Jaquette. 1961. Case report—Dispharynx nasuta causing severe proventriculitis in pigeons. Avian Diseases 5:60–65. Kinsella, J. M. 1974. Helminth fauna of the Florida Scrub Jay: Host and ecological relationships. Proceedings of the Helminthological Society of Washington 41:127–130. Kock, R. A., G. M. Henderson, E. C. Appleby, C. M. Hawkey, and R. N. Cinderey. 1987. Acuariasis in water fowl at Whipsnade zoo. Erkrankungen der Zootiere: Verhandlungsbericht des 29. Internationalen Symposiums u¨ ber die Erkrankungen der Zootiere, May 20–24, Cardiff, pp. 65–73. Kovalenko, I. I. 1960. Study of the life-cycles of some helminths of domestic ducks from farms on the Azov coast (in Russian). Dokladi Akademii Nauk SSSR 133:1259–1261. Laberge, R. J. A., and J. D. McLaughlin. 1989. Hyalella azteca (Amphipoda) as an intermediate host of the nematode Streptocara crassicauda. Canadian Journal of Zoology 67:2335–2340. Laberge, R. J. A., and J. D. McLaughlin. 1991. Susceptibility of blue-winged teal, gadwall, and lesser scaup ducklings to experimental infection with
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Streptocara crassicauda. Canadian Journal of Zoology 69:1512–1515. Lapage, G. 1961. A list of the parasitic protozoa, helminths and Arthropoda recorded from species of the family Anatidae (ducks, geese, and swans). Parasitology 51:1–109. Lindquist, W. D., and A. C. Strafuss. 1980. (Dispharynx nasuta) may cycle within avian zoo populations. Journal of Zoo Animal Medicine 11:120–122. Macko, J. K., V. Birova, and J. Hovorka. 1974. The distribution of Dispharynx nasuta and the morphology of D. resticula in free-living birds in Cuba. Helminthologia 15:865–880. Mason, R. W. 1988. Laryngeal streptocariasis causing death from asphyxiation in ducks. Australian Veterinary Journal 65:335–336. McDonald, M. E. 1969. Catalogue of helminths of waterfowl (Anatidae). Bureau of Sport Fisheries and Wildlife Special Scientific Report—Wildlife, no. 126. Washington, DC., 692 pp. McDonald, M. E. 1974. Key to nematodes reported in waterfowl. Department of the Interior, Bureau of Sport Fisheries and Wildlife, Resource Publication 122, Washington, DC., 44 pp. McLaughlin, J. D., and B. P. McGurk. 1987. An analysis of gizzard worm infections in fall migrant ducks at Delta, Manitoba, Canada. Canadian Journal of Zoology 65:1470–1477. Misiura, M. 1970. Development of Echinuria uncinata (Rud., 1819) larvae (Nematoda) in Cladocera and Ostracoda. Acta Parasitologica Polonica 17:247–251. Moore, J., and J. Lasswell. 1986. Altered behavior in isopods (Armadillidium vulgare) infected with the nematode Dispharynx nasuta. Journal of Parasitology 72:186–189. Moore, J., M. Freehling, J. Crawford, and P. Cole. 1988. Dispharynx nasuta (Nematoda) in California quail (Callipepla californica) in western Oregon. Journal of Wildlife Diseases 24:564–567. Nagy, G., V. Birova, and D. Ovies. 1977. Sobre la patomorfolog´ıa de la disfarinxosis de las aves. Revista Cubana de Ciencias Veterinarias 8:75–84. Ould, P., and H. E. Welch. 1980. The effect of stress on the parasitism of mallard ducklings by Echinuria uncinata (Nematoda: Spirurida). Canadian Journal of Zoology 58:228–234. Piana, G. P. 1897. Osservazioni sul Dispharagus nasutus Rud. dei polli e sulle larve nematoelmintiche delle mosche e dei porellioni. Atti Societa Italiana delle Scienze Naturelle, Milano 36:239–262. Ramaswamy, K., and R. K. Sundaram. 1985. Histopathological changes in the proventriculus of fowl given experimental monospecific infection with Acuaria spiralis. Veterinary Parasitology 17:309–317. Respaldiza, E., A. Sanz, A. Juncosa, A. L. Garcia del Campo, and J. L. Valls. 1979. Observaci´on de
acuariosis (Echinuria uncinata) en el Cygnus olor, en Espa˜na. Anales del Instituto Nacional de Investigaciones Agrarias, Serie Higiene y Sanidad Animal 4:69–81. Rickard, L. G. 1985. Proventricular lesions associated with natural and experimental infections of Dispharynx nasuta (Nematoda: Acuariidae). Canadian Journal of Zoology 63:2663–2668. Schulman, F. Y., R. J. Montali, and S. B. Citino. 1992. Pathology, diagnosis, and treatment of Synhimantus nasuta infection in African jacanas (Actophilornis africana). Journal of Zoo and Wildlife Medicine 23:313–317. Shanthikumar, S. R. 1987. Helminthology. In Diseases of Cage Birds, E. W. Burr (ed.). T.F.H. Publications, Neptune City, NJ, pp. 135–146. Skrjabin, K. I. 1916. N´ematodes des oiseaux du Turkestan russe. Annuaire du Mus´ee Zoologique. Academie Imperiale du Sciences, Petrograd 20:457–557. Soloviev, N. F. 1912. Vers parasitaires des oiseaux du Turkestan. Annuaire du Mus´ee Zoologique (Akademiia Nauk), St. Petersbourg, Russie 17:86– 115. Spalding, M. G., J. M. Kinsella, S. A. Nesbitt, M. J. Folk, and G. W. Foster. 1996. Helminth and arthropod parasites of experimentally introduced whooping cranes in Florida. Journal of Wildlife Diseases 32:44–50. Stauber, E., and S. Schussman. 1985. Parasites of pet and aviary birds–part 1. Modern Veterinary Practice 66:457–460. Sterner, M. C., and L. Stackhouse. 1987. Parasitic ulcerative ventriculitis in mallards (Anas platyrhynchos). Journal of Wildlife Diseases 23:680–682. Takla, M., and W. Thiel. 1983. Echinuriose bei 2 schwarzen Schw¨anen. Ein kasuistischer Beitrag zur Diagnose. Tier¨arztliche Praxis 11:331–338. Wehr, E. E. 1971. Nematodes. In Infectious and Parasitic Diseases of Wild Birds, J. W. Davis, R. C. A. Anderson, L. Karstad, and D. O. Trainer (eds). Iowa State University Press, Ames, IA, pp. 185–233. Wood, N. A. 1974. Waterfowl and Acuaria. Avicultural Magazine 80:59–64. Work, T. M., C. U. Meteyer, and R. A. Cole. 2004. Mortality in Laysan ducks (Anas laysanensis) by emaciation complicated by Echinuria uncinata on Laysan Island, Hawaii, 1993. Journal of Wildlife Diseases 40:110–114. Zhang, L., D. R. Brooks, and D. Causey. 2004. Two species of Synhimantus (Dispharynx) Railliet, Henry, and Sisoff, 1912 (Nematoda: Acuarioidea: Acuariidae), in passerine birds from the Area de Conservacion Guanacaste, Costa Rica. Journal of Parasitology 90:1133–1138.
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19 Tracheal Worms M. A. Fernando and John R. Barta recorded in 1779 from poultry by Weisenthal, as a “worm from poultry,” it was named in 1811 by Montagu as Fasciola trachea from a pheasant and partridge. In 1886, Seibold recognized the worm as a nematode that he called S. trachealis and provided the first description. It was called Syngamus trachealis until Chapin (1925) corrected the nomenclature to S. trachea. Since this time, as noted below, S. trachea has been found in many wild and domestic gallinaceous and passeriform birds. Some of these syngamid nematodes that did not reside in permanent copula were placed in the genus Cyathostoma (cf. Syngamus spp. that remain paired even when removed from the animal or placed in fixative, see Borgsteede and Okulewicz, 2001). The genus Cyathostoma was first described in 1849 by Blanchard on the basis of his description of Cyathostoma lari from the orbital cavity of the black-headed gull, Larus ridibundus. Since then, a number of Cyathostoma species have been described from the orbital cavity and throughout the respiratory tract of a wide range of wild and domestic waterfowl and a wide range of wild and captive raptors (Ali 1970).
INTRODUCTION Tracheal worms are cosmopolitan, strongylid nematodes that infect the respiratory tract of wild and domestic birds. Throughout this chapter we discuss gapeworms (Syngamus spp.) and cyathostomes (Cyathostoma spp.) separately. The widely reported gapeworm, Syngamus trachea, is relatively common in wild and range-reared domestic fowl such as turkeys, chickens, pheasants, and perching birds and has been found much less commonly in waterfowl, herons and storks, pelicans, and woodpeckers. Uncommonly, S. trachea has been shown to be a primary pathogen in young game birds, especially pheasants (Lister 1989). Among cyathostomes, the most commonly recorded species, Cyathostoma bronchialis, is likewise found globally, most commonly in waterfowl (Anseriformes). There are a few rare reports of mortality in wild populations (e.g., Karstad and Sileo 1971), but these parasites are usually not regarded as serious pathogens in the wild. However, confined rearing of susceptible hosts and the resulting enhanced transmission can make these serious parasites of young birds. Cyathostoma (Hovorkonema) variegatum has been recovered from wild birds in respiratory distress, suggesting that these parasites may be an unrecognized cause of wild bird mortality (Krone et al. 2007). Only a few worms are capable of causing dramatic clinical signs under some circ*mstances. Many wild birds may act as reservoirs of infection to domestic animals (and vice versa), and transmission of both S. trachea and C. bronchialis between wild and domestic hosts has been demonstrated.
ETIOLOGY Tracheal worms of birds are parasitic nematodes belonging to the order Strongylida, superfamily Strongyloidea, family Syngamidae (Lichtenfels 1980). Species in the subfamily Syngaminae belong to the genera Syngamus or Cyathostoma and are usually considered to occur only in birds (Chapin 1925). Cyathostoma species have been assigned to one of two subgenera (e.g., Cyathostoma (Cyathostoma) spp. or Cyathostoma (Hovorkonema) spp.) on the basis of details of the male copulatory bursa and spicule length (Lichtenfels 1980; Borgsteede and Okulewicz 2001). Some confusion surrounds the naming of cyathostomes infecting birds in various surveys and case reports. For example, Cyathostoma (Hovorkonema) variegatum has been referred to as a Hovorkonema sp. in the literature
SYNONYMS Gapes, gapeworm infection, cyathostomiasis, syngamiasis, tracheal worm infection. HISTORY Madsen (1950) gives a short history of the records of S. trachea from poultry and other birds. Originally
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(Krone et al. 2007 and others). Throughout this chapter, we refer to all cyathostomes as Cyathostoma spp. belonging to the subgenera Hovorkonema or Cyathostoma as appropriate (see Lichtenfels 1980; Borgsteede and Okulewicz 2001). A detailed examination of the taxonomic relationships among the various syngamid worms infecting birds is beyond the scope of this chapter. For the sake of simplicity, we discuss “gapeworms” (Syngamus species) that all reside in permanent copulation in the trachea of birds (Table 19.1) and the closely related “cyathostomes” (Cyathostoma spp.) that reside in the trachea, air sacs, or orbital sinuses of birds and that do not pair up permanently (Table 19.2). Morphology Gapeworms and cyathostomes in birds are typical strongylid nematodes with a moderate to large buccal capsule or mouth and prominent copulatory bursa in the male worms. These parasites are associated with the mucosal surface of the respiratory tract of birds. Determining whether syngamid worms found in the respiratory tract belong to the genera Sygamus or Cyathostoma is relatively straightforward; species of Syngamus are found in permanent copulation as adults and are found attached to the trachea of infected hosts, whereas species of Cyathostoma are not permanently joined in copulation and can be found in the trachea, air sacs, or orbital sinuses of their hosts. Although there are multiple species in each genus, the morphology of the most common species of each is described below. Differentiating species within each of these genera is difficult and relies on detailed morphological and morphometric observations of the adult worms. The general morphology of the remaining species in each genus is similar. SYNGAMUS TRACHEA Both the male and female are bright red, permanently in copulo, and their bodies form a Y shape (Figure 19.1a). The buccal capsule is large and well developed with eight (sometimes nine) teeth at the base, which allows them to live attached to the wall of the trachea (Figure 19.1b). Here, they often form nodules. The males are 2–6 mm long and 200 m in diameter with a bursa and slender spicules 55–82 m long at the posterior end. The females are 5–40 mm in length and 350 m in diameter. They shed ellipsoidal eggs 85–90 m × 50 m that possess distinct opercula at each end. CYATHOSTOMA BRONCHIALIS In contrast to S. trachea, these worms are not permanently in copulo. The buccal capsule is cup shaped
with six to seven teeth at the base (Figure 19.1c). The males are 4–6 mm long, the bursa is well developed and the spicules are about 0.5 mm in length. The females are 16–30 mm in length and their eggs measure 75–83 m × 50–62 m with a single indistinct operculum at the narrow end of the egg. DISTRIBUTION AND HOST RANGES Gapeworms (Syngamus spp.) have been reported mainly from Europe and North America but likely infect birds globally in tropical and temperate to cold temperate climates (Table 19.1). Not surprisingly, many of the reported host species are ground-feeding birds such as chickens, grouse, turkeys, and pheasants or invertebrate-feeding birds such as robins, herons, cranes, gulls, and jays. Surprisingly, cyathostomes have been reported quite commonly from predatory birds such as merlins, hawks, and owls (Table 19.2) that do not generally feed directly on the invertebrates that act as intermediate hosts for these worms. Several authors have suggested that predatory birds acquire infections by eating rodents or smaller birds with infected intermediate hosts containing infective larvae of Cyathostoma spp. in their alimentary tracts (Simpson and Harris 1992; Lavoie et al. 1999). LIFE CYCLE AND EPIZOOTIOLOGY Syngamus trachea Eggs laid by adult female S. trachea within the trachea are coughed up, usually swallowed, and passed in the feces of the host. Under optimal conditions at 29◦ C, the larvae take about a week to develop to the infective third stage within the eggs. The larvae can hatch spontaneously. Development is much longer under less favorable conditions. Eggs do not develop at ambient temperatures at or below 16◦ C (Baruˇs 1966b). Larvae developing within the eggshell reach infectivity in 42 days at 17◦ C, 25–28 days at 19◦ C, 16 days at 21◦ C, 13–14 days at 25◦ C, and only 9 days at 27◦ C. Hatched larvae are susceptible to desiccation (Ortlepp 1923). Both hatched infective larvae and eggs containing infective larvae are infective when ingested by the host. In addition to direct transmission via ingestion of embryonated eggs or larvae from the environment, S. trachea frequently incorporates an optional transport (paratenic) host in its life cycle. Earthworms act as transport hosts and may be the principal means for infective larvae to overwinter in colder climates. The larvae of S. trachea have been shown to remain viable and infective for more than 3 years, encapsulated in earthworm muscle. In experimentally infected earthworms (Eisenia foetida), encysted third-stage larvae
Corvus corone Corvus frugilegus Corvus monedula Pica pica Sturnus vulgaris Passer domesticus Calidris maritima Rhea americana Meleagris gallopavo Trachyphonus erythrocephalus Turdus philomelos Turdus iliacus Turdus merula Turdus migratorius Megaceryle alcyon Gallinago stenura Phalacrocorax carbo Philomachus pugnax Tringa erythropus Nucifraga caryocatactes Acrocephalus schoenobaenus Motacilla alba Cicinnurus magnificus
Carrion Crow Rook Eurasian Jackdaw Eurasian Magpie European Starling House Sparrow Purple Sandpiper Greater Rhea Wild Turkey Red-and-yellow Barbet
Pintail Snipe Great Cormorant Ruff Spotted Redshank Eurasian Nutcracker Sedge Warbler White Wagtail Magnificent Bird-of-paradise
Syngamus gibbocephalus Syngamus microspiculum Syngamus palustris
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Syngamus sp. (eggs only)
Ryzhikov (1949) Skrjabin (1915) Ryzhikov (1949) Ryzhikov (1949)
Russia
Varghese (1987)
Welte and Kirkpatrick (1986) Boyd and Fry (1971)
Delaware, USA Massachusetts, USA and Ontario, Canada Russia Turkestan Russia
Papua New Guinea
Campbell (1935)
Britain
Note: Syngamus trachealis Siebold 1936, Syngamus parvus Chapin 1925, Syngamus gracilis Chapin 1925, and Syngamus merulae Baylis 1926 are considered to be synonyms of Syngamus trachea (Montagu, 1811) as discussed in detail by Madsen (1950). Also called “the gapeworm.”
Syngamus taiga
Syngamus alcyone
De Wit (1995) Zavadil (1966); Baruˇs (1966a) Nevarez et al. (2002)
USA Commercially reared in Eastern Europe In aviary in USA
Britain
Pavlovic et al. (2003) Moynihan and Musfeldt (1950) Campbell (1935) Wissler and Halvorsen (1975) Campbell (1935)
Reference
Commercially reared in Serbia Commercial flock, BC, Canada Britain Norway
Country reported
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Song Thrush Redwing Eurasian Blackbird American Robin Belted Kingfisher
Lagopus lagopus
Syngamus trachea
Willow Ptarmigan
Host species Phasianus colchicus
Host common name Ring-necked Pheasant
Syngamus species
Table 19.1. Examples of Syngamus species reported from wild birds.
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Cyathostoma sp.
C. (Hovorkonema) variegatum
C. bronchialis
Black-headed Gull Rook Eurasian Jackdaw European Starling Gray Heron Eurasian Kestrel Eurasian Buzzard Eurasian Sparrow Hawk European Herring Gull Northern Goshawk Eurasian Buzzard Rough-legged Hawk Broad-winged Hawk Northern Goshawk Canada Goose Swan Goose Geese Mallard Eurasian Buzzard Northern Goshawk Eurasian Sparrow Hawk Eurasian Buzzard Western Marsh-Harrier White-tailed Eagle Common Crane Whooping Crane Barred Owl Snowy Owl Northern Harrier Northern Goshawk Broad-winged Hawk Belted Kingfisher
Host common name Larus ridibundus Corvus frugilegus Corvus monedula Sturnus vulgaris Ardea cinerea Falco tinnunculus Buteo buteo Accipiter nisus Larus argentatus Accipiter gentilis Buteo buteo Buteo lagopus Buteo platypterus Accipiter gentilis Branta canadensis Anser cygnoides Various Anser spp. Anas platyrhynchos and others Buteo buteo Accipiter gentilis Accipiter nisus Buteo buteo Circus aeruginosus Haliaeetus albicilla Grus grus Grus americana Strix varia Bubo scandiacus Circus cyaneus Accipiter gentilis Buteo platypterus Megaceryle alcyon
Host species Pemberton (1959)
Simpson and Harris (1992) Threlfall (1965) Borgsteede and Okulewicz (2001) Lavoie et al. (1999) Fernando et al. (1973) Karstad and Sileo (1971) Chapin (1925) Sanmartin et al. (2004) Krone et al. (2007)
Spalding et al. (1996) Lavoie et al. (1999)
Boyd and Fry (1971)
Britain Britain The Netherlands Quebec, Canada Ontario, Canada Europe and USA Spain Germany
Florida, USA Quebec, Canada
Massachusetts, USA.
Reference
Britain
Country
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C. (Hovorkonema) americanum
C. lari
Species of Cyathostoma
Table 19.2. Examples of Cyathostoma species reported from wild birds.
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Tracheal Worms of S. trachea are found in the body wall of the earthworm, deeply embedded in the muscle in a thin hyaline cyst. Encysted larvae do not affect vitality of the earthworms and hence they can act as cumulative reservoirs (Clapham 1934). Slugs, ants, beetles, and many other invertebrates are capable of acting as paratenic hosts for this parasite and these may provide additional opportunities for dissemination of infections. When eggs of S. trachea are ingested by a suitable avian host, larvae hatch within the intestines and can be recovered from the liver shortly thereafter (Fernando et al. 1971). This suggests that most larvae arrive in the lungs via the bloodstream. The larvae are seen in the pulmonary connective tissue and air sac capillaries as early as 4 h postinfection (PI) and the atria of the lungs by 24 h (Figure 19.2a). Worms are recovered from the lungs up to 7 days PI and from the trachea as early as 7 days PI. Most worms are in the trachea by 11 days PI and the females are fertile by 14 days PI. Worms have been found attached to the cartilage of the tracheal rings at day 14 PI. Male worms are always found permanently attached to the tracheal mucosa. This is normal. In some instances, they penetrate the tracheal rings and the host response forms a nodule around the anterior end of the male worm. The females are attached to the males in permanent copula but are not attached permanently to the tracheal mucosa. Female worms feed at multiple locations around the attachment site of their male partner. The first eggs are found in the feces 17–20 days PI (Wehr 1937; Clapham 1939; Fernando et al. 1971). Gapeworm infection has been documented in commercially raised pheasants (Moynihan and Musfeldt 1950; Pavlovic et al. 2003) and turkeys (Zavadil 1966). In Serbia, S. trachea is the most prevalent nematode species (37.2%) in commercially raised pheasants under the age of 14 weeks (Pavlovic et al. 2003). Commercially raised turkeys may be particularly susceptible throughout life to infection with S. trachea resulting in “mass-perishing” of these birds (Zavadil 1966). In a longitudinal study of seasonally range-reared breeding turkeys in Eastern Europe, newly acquired infections arose each spring upon access to infected earthworms. Transmission peaked in late summer, possibly because larval maturation within eggs occurs most quickly at higher temperatures and because earthworm populations are larger or more available (Baruˇs 1966a). In the same study, almost 7% of the earthworms collected within the turkey runs contained 1–12 infective larvae of S. trachea. Free-living hosts of Syngamus spp. include Rooks (Corvus frugilegus), crows, pheasants, robins, jays, magpies, starlings, and many other species (Campbell 1935). Prevalence of this parasite varies widely by host
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Figure 19.1. (a) and (b) Syngamus trachea from a Ring-necked Pheasant (Phasianus colchicus). (a) Pairs of male and female worms in copulo. (b) Buccal capsule of young adult (L5) recovered 5 days postinfection (PI). (c) Cyathostoma bronchialis from a Canada Goose (Branta canadensis). Buccal capsule of young adult (L5) recovered 5 days PI. Note triangular teeth (arrows) at the base of the buccal cavity. Reproduced from Fernando et al. (1973), with permission from the Journal of Parasitology.
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Parasitic Diseases of Wild Birds old birds 40%, and adults 8.6% (Campbell 1935). This drop in prevalence occurs even over a single breeding season. Prevalence of S. trachea in young Rooks was 98% in May, 85% in June, and 62% in August, suggesting that Rooks are able to expel the worms after a period of infection (Rice 1929). Intensity of infection with S. trachea also decreased as the birds aged. Young Rooks had up to 40 pairs of S. trachea in their trachea but most infections in adult birds had only 1 or a few pairs of worms (Elton and Buckland 1928). In contrast to most wild birds, prevalence of infection in adult and juvenile Ring-necked Pheasants (Phasianus colchicus) is similar (Campbell 1935). Infections with S. trachea in wild birds have been implicated as a cause of outbreaks on game bird farms, particularly pheasant farms, as well as in range poultry (Elton and Buckland 1928; Clapham 1934; Campbell 1935).
Figure 19.2. (a) Syngamus trachea in lung of a Ring-necked Pheasant (Phasianus colchicus) at 24 h postinfection (PI). (b) Larvae of Cyathostoma bronchialis distending parabronchus in lung of a Canada Goose (Branta canadensis) at 3 days PI. Reproduced from Fernando et al. (1973), with permission from the Journal of Parasitology.
species and host age class. For example, in a series of examinations of wild birds over a number of years in the UK, starlings had a prevalence of about 18–35% (Lewis 1925, 1926; Taylor 1928; Campbell 1935) with intensity of infections ranging from 1 to 5 pairs of worms per bird. In a large survey examining gapeworm infections in more than a thousand birds of 53 species, from 1 to 82% of individuals of 13 different species of birds were infected with gapeworms (Campbell 1935). In this same study, the prevalence of gapeworm infections and their intensities in most hosts were shown to decrease with the age of the host. The most striking example of this was observed in the Rook in which the young of the year had a prevalence of 99%, 1-year-
Cyathostoma bronchialis Eggs of C. bronchialis are coughed up, swallowed, and passed in the feces of the host. Under favorable environmental conditions, larvae take about 10 days to develop and then the eggs hatch spontaneously to release larvae. In contrast to the larvae of S. trachea, hatched larvae or eggs containing third-stage larvae are not directly infective to captive Snow Geese (Chen caerulescens) or Canada Geese (Branta canadensis) (Cram 1927; Fernando et al. 1973). Earthworms containing infective C. bronchialis larvae are, however, sources of infection when fed to susceptible geese (Fernando et al. 1973). Earthworms are necessary for transmission and are therefore obligate intermediate hosts. Like larvae of S. trachea, infective larvae of C. bronchialis can survive and overwinter in earthworms for several years but details about this host– parasite association are not known. It is perhaps surprising to find cyathostome infections in birds that do not generally feed on invertebrates such as the Merlin (Falco columbarius), the Northern Goshawk (Accipiter gentilis), and the Snowy Owl (Bubo scandiacus) (Lavoie et al. 1999). In these cases, infections are most likely transmitted through infected invertebrates present in the alimentary tract of normal prey species. Sparrow hawks may acquire Cyathostoma infections in a similar manner (Simpson and Harris 1992). After an infected earthworm is ingested, larvae of C. bronchialis can be recovered from abdominal air sacs and lungs as early as 1–4 h PI (Figure 19.2b). The route of migration from the intestinal tract to the pulmonary spaces is not known. Worms move to the trachea by 6 days PI. By 13 days PI, most males and females attain their maximum size, have mated, and
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Tracheal Worms eggs are found in the tracheal mucus. Eggs are shed in the feces shortly thereafter (Fernando et al. 1973). Wild birds are usually not heavily or commonly infected with cyathostomes. For example, infections with cyathostomes were found in only 7 of 20 species of wild birds during an extended study at three different locations in Germany. Prevalence of infection ranged from <1% to almost 20%, but intensities of infection ranged from only 1 to 5 worms per bird (Krone et al. 2007). In Canada, 394 wild-caught birds of prey belonging to 24 species were examined for cyathostome infections. Seven species of hawks and owls were found to be infected at prevalences that ranged from 4 to 33% (Lavoie et al. 1999). In the latter study, intensity of infection varied widely from 2 to approximately 100 worms per bird. Most birds were infected with only a few worms.
CLINICAL SIGNS Clinical signs of infection with S. trachea vary depending on the size of the bird and the intensity of infection. Although the parasite infects a wide range of avian species, pheasants appear to be particularly susceptible to infection and clinical outbreaks of disease have been recorded (Pavlovic et al. 2003). Young birds are severely affected by migration of larvae through the lungs and develop pneumonia. Approximately 2 weeks after infection adult worms can block the trachea, giving rise to the typical clinical sign of “gaping” or gasping for air with outstretched necks and open mouths (Figure 19.3). Head shaking and bouts of coughing are seen in some birds. Severely affected young birds may stop drinking, deteriorate rapidly, and
Figure 19.3. Syngamus trachea. Ring-necked Pheasant (Phasianus colchicus) with intense infection at 2 weeks postinfection. The outstretched neck and open mouth are typical clinical signs of “gaping” or gasping for air.
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die (Clapham 1934; Levine 1968). Adult birds show few clinical signs except an occasional cough. Even though C. bronchialis can be a significant pathogen in young captive waterfowl (Karstad and Sileo 1971), most wild birds infected with cyathostomes show no signs of disease. In those rare instances where clinical signs are present, they include emaciation and/or dyspnea resulting from impaired respiratory function associated with air sacculitis (Lavoie et al. 1999). Clinically affected birds had higher numbers of worms in their respiratory tracts. PATHOGENESIS AND PATHOLOGY Gapeworms The pathogenesis of the lesions induced by S. trachea in the lungs and trachea in chickens and pheasants has been studied in detail (Wetzel and Fortmeyer 1964; Baruˇs and Blazek 1965). Grossly visible white foci, approximately 0.2 mm in diameter, do not appear in the lungs of pheasants until 7–9 days after experimental infection. However, microscopic lesions are seen as early as 3 days PI (Fernando et al. 1971). Early pulmonary lesions associated with larval migration through the lungs include an increase in the number of lymphocytes within the connective tissue and the parabronchi, cuffing of larger vessels by lymphocytes through migration of lymphocytes and heterophils from the blood stream, disappearance of the normal architecture of the air capillaries, and consolidation of pulmonary lobules. Giant cells and granulocytes fill the atria by 4 days PI and the lumina of the parabronchi by 7 days PI. Infiltration of the lamina propria of the secondary and primary bronchi and collapse of some of the secondary bronchi may be seen from the seventh day onward. Most pathogenic changes appear associated with an inflammatory host response to the presence of larvae and their antigens. Worms in the trachea and bronchi give rise to hemorrhagic tracheitis and bronchitis. Worms are found attached to the cartilage of the tracheal rings by day 14 PI in pheasants (Figure 19.4a). Worms cause direct mechanical damage to the mucosa and produce hemorrhage as they feed on blood. Host-mediated inflammatory responses to these mechanical lesions and perhaps worm antigens and/or secretory products exacerbate the mechanical damage. In Ring-necked Pheasants, pale tracheal nodules form at the point of attachment of the anterior end of male worms and may be visible on the interior or exterior surfaces (Figures 19.4b and 19.4c). These nodules are mainly composed of hyperplastic peritracheal connective tissue (Fernando et al. 1971). Male worms can perforate the mucosa and attach to the tracheal cartilage
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(a)
(b)
(c)
(d)
Figure 19.4. Syngamus trachea from ring-necked pheasant (Phasianus colchicus). (a) Worms blocking trachea at 14 days postinfection (PI). (b) Pea-sized nodules on the mucosal surface of the trachea at the points of attachment of the anterior end of male worms at 14 days after experimental infection. (c) Large nodule on the outer surface of the trachea 6 weeks after experimental infection. The head and anterior half of the body of the male worm is deeply embedded in the connective tissue of the nodule. (d) Buccal capsule attached to tracheal cartilage 14 days PI. (Figure 19.4d). Male worms sometimes penetrate so deeply that they cause rupture and proliferation of the perichondrium, lysis of tracheal cartilage, and, not uncommonly, perforation of the tracheal rings (Clapham 1935; Carrara 1961; Fernando et al. 1971). Cyathostomes Cyathostoma bronchialis causes bronchitis of the primary, secondary, and tertiary bronchi during development in geese (Fernando et al. 1973). Worms move up the trachea by day 6 PI, so they exert the most pathogenic effects in the lower respiratory tract on the day 5 and day 6 PI (Fernando et al. 1973). Lymphoid cuffing of interlobular arteries and lymphoid replacement of air capillaries appears to be a reaction to presence of worms in the parabronchi and superficial secondary bronchi. Severe bronchitis in the sec-
ondary and primary bronchi probably also reflects the period of most rapid growth of the developing worms. By 14 days PI, the bronchial lesions are completely resolved. Varied lesions may be seen in the bronchi at later stages of infection (28 days PI in experimental studies) and presumably are caused by aspiration of adult worms and eggs. Eggs, cuticular material shed by developing worms, and whole worms elicit a pyogranulomatous bronchitis or pneumonia in the lungs. Aspiration pneumonia in goslings with patent infections may also be caused by aspiration of eggs and adult worms. This contrasts with the behavior of S. trachea, which either remains in the trachea after migration through the lungs or is coughed out. Concurrent lesions of aspergillosis have been reported in a “blue goose” (presumably a Snow Goose, Chen caerulescens), a Canada Goose and goslings, and goslings of domestic
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Tracheal Worms geese (Christenson 1932; Fernando et al. 1973). These observations suggest that C. bronchialis may predispose birds to other respiratory infections. The association of fungal hyphae with cuticular remnants in the parabronchi and secondary bronchi of these birds supports this possibility. Cyathostoma americanum and other unidentified Cyathostoma spp. have been implicated as causes of diffuse pyogranulomatous air sacculitis, pneumonia, and bronchitis in wild raptors in North America (Lavoie et al. 1999) and elsewhere. Lesions are generalized and appear to be associated primarily with inflammatory reactions to eggs in the air sacs (Hunter et al. 1993). Prevalence of Cyathostoma (Hovorkonema) variegatum was 3% among raptors and 9% in Goshawks, and this parasite was associated with thickened air sac walls and granulomatous lesions at the site of infection in 7 of 12 cases (Krone et al. 2007). DIAGNOSIS Syngamus trachea A diagnosis is usually made on the basis of the typical clinical signs and confirmed by observation of worms in the trachea during necropsy. Characteristic eggs will be found in the feces. Eggs of S. trachea are approximately 80–100 m × 50–60 m and are bipolar with clearly visible opercula at both ends. Each egg contains a morula (about an eight-cell embryo) when freshly passed. Eggs of S. trachea superficially resemble those of Capillaria spp. However, the eggs of Capillaria spp. are usually smaller (<60 m in length), have a thicker (and frequently brownish) eggshell, possess pronounced polar plugs, and contain a single cell when passed. Cyathostoma bronchialis Clinical signs related to infections with cyathostomes are neither common nor diagnostic. On necropsy, finding nematodes morphologically consistent with cyathostomes within the trachea and bronchi is diagnostic. Antemortem, patent infections can be diagnosed by finding ovoid morulated eggs with an indistinct polar operculum at the narrow end measuring 75–90 m × 45–60 m in feces. IMMUNITY In infections with either S. trachea or C. bronchialis, younger birds are more susceptible to the pathogenic effects of infection. Following an initial infection with S. trachea, pheasants acquire at least partial immunity to further infection (Olivier 1942). Immunity persists after the adult worms are lost. Turkeys are, how-
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ever, susceptible throughout life to gapeworm infection (Varga 1971). Prevalence of infection with Syngamus drops considerably in free-living populations as birds age (see above), suggesting that most natural hosts of these parasites acquire resistance to infection as the birds mature. Approximately 95% of chickens vaccinated with irradiated embryonated eggs and larvae of S. trachea are resistant on challenge (Zeigler 1966, 1968). Development of the worms is inhibited before they enter the trachea, suggesting that the host immune response acts on the migratory phase of the parasite. This immunity may reduce the number of adult worms that become established in the trachea, even in conditions where birds are exposed to significant numbers of larvae. PUBLIC HEALTH CONCERNS Gapeworms of birds (Syngamus or Cyathostoma species) are not known to infect mammals, including humans, and therefore pose no public health concerns. Tracheal worms infecting mammals (species of Mammomanogamus) are relatively common globally and have been reported as accidental infections of humans. Previously, these mammalian tracheal worms were considered to be species of Syngamus and thus there are a number of case reports of “syngamosis” in humans (e.g., Leers et al. 1985). No avian gapeworm has ever been implicated in a human infection. WILDLIFE POPULATION IMPACTS Clinical gapeworm and cyathostome infections in wild populations are uncommon, although subclinical infections with these parasites are frequent. For example, 23 of 107 wild raptors in Quebec, Canada, were infected with species of Cyathostoma, but clinical signs of emaciation and dyspnea attributable to gapeworm infection were evident in only 4 cases (Lavoie et al. 1999). All the clinically affected birds had a severe diffuse pyogranulomatous air sacculitis. Reports of birds clinically affected by species of Syngamus are equally uncommon despite high prevalence and high intensity in some hosts (e.g., young Rooks in Britain, see Campbell 1935). DOMESTIC ANIMAL HEALTH CONCERNS Wild birds including Rooks, pheasants, robins, jays, crows, magpies, and starlings can act as reservoir hosts of S. trachea. These reservoirs have been implicated as the source of outbreaks on game bird farms, particularly pheasant farms, as well as range poultry (LeaMaster 2007). Gapeworms (Syngamus spp.) are not host specific and S. trachea, at least, can be
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transmitted among many orders and families of birds. Likewise, wild waterfowl are expected to act as reservoirs for species of Cyathostoma that infect domestic ducks and geese. Earthworms and perhaps other invertebrates are an important source of infection where poultry and game birds are reared on soil.
TREATMENT AND CONTROL Gapeworm infections can be treated with a number of anthelmintic medications. Benzimidazole anthelmintics such as thiabendazole, mebendazole, flubendazole, and fenbendazole have demonstrated efficacy against gapeworm infections (Wehr and Hwang 1967; Ssenyonga 1982; Draycott et al. 2006). In the US, thiabendazole is registered for the control of S. trachea in pheasants. In Peru, Agrovet Market S.A. produces a broad-spectrum anthelmintic, Gallomec Plus® ; each tablet contains 0.2 mg ivermectin, 30 mg fenbendazole, and 10 mg praziquantel, and is used to control S. trachea in poultry, particularly younger birds. Extralabel use of injectable ivermectin has been used with variable success to treat gapeworm and cyathostome infections in poultry and wild birds, including raptors. Despite treatment, clinical signs of coughing may persist for a period of time because of the presence of dead and dying worms in the respiratory tract. In heavy infections with species of Cyathostoma, treatment may produce fatal pneumonia from the aspiration of dead and dying worms and inflammatory responses to the dead parasites. Opportunistic secondary bacterial or fungal pneumonias may require specific treatment in addition to anthelmintics to remove the worms. Although gapeworm and cyathostome infections appear to be of minor significance in free-living wild birds, they can be a serious problem in commercially reared pheasants, chukars, and waterfowl as well as range-reared turkeys and chickens. The presence of large numbers of susceptible young birds in a relatively confined area can lead to severe and clinically apparent infections and mortality. Restricting access of young birds to heavily contaminated yards, older birds, or wild bird reservoir hosts will reduce the intensity of infections and thereby reduce clinically significant infections. Earthworms are particularly important in maintaining an infective environment because thirdstage larvae of gapeworms can survive in earthworms for extended periods (Baruˇs 1966a) and earthworms are obligate intermediate hosts for cyathostomes. For this reason, restricting access of young birds to open ground may reduce the impact of these parasites in domestic species.
LITERATURE CITED Ali, M. M. 1970. A review and revision of the subfamily Cyathostominae Nicoll, 1927 (Nematoda, Syngamidae). Acta Parasitologica Polonica 17:237–246. Baruˇs, V. 1966a. Seasonal dynamics of the invasion extensity of the nematode Syngamus trachea (Montagu, 1811) in breeding turkeys (Meleagris gallopavo f. domestica). Helminthologia 7:29–37. Baruˇs, V. 1966b. The effect of temperature and air humidity on the development and the resistance of eggs of the nematode Syngamus trachea (Montagu, 1811). Helminthologia 7:103–106. Baruˇs, V., and K. Blazek. 1965. Revision der exogenen und endogenen phase des Entwicklungszyklus und der Pathogenitat von Syngamus (Syngamus) trachea. (Montagu, 1811) Chapin, 1925 im Organismus des Endwirtes. Ceskoslovenska Parasitologie 12:47–70. Borgsteede, F. H. M., and A. Okulewicz. 2001. Justification of the species Cyathostoma (Hovorkonema) americanus (Chapin, 1925) Syngamidae—Nematoda). Helminthologia 38:151–154. Boyd, E. M., and A. E. Fry. 1971. Metazoan parasites of the eastern belted kingfisher, Megaceryle alcyon alcyon. Journal of Parasitology 57:150–156. Carrara, O. 1961. Contributo alla conoscenza della granulomatosi da Syngamus trachea nel fa*giano. La Clinica Veterinaria 84:277–283. Campbell, J. W. 1935. The Gapeworm (Syngamus) in wild birds. Journal of Animal Ecology 4(2):208– 215. Chapin, E. A. 1925. Review of the nematode genera Syngamus Sieb. and Cyathostoma E. Blanch. Journal of Agricultural Research 30:557–570. Christenson, R. O. 1932. An epizootic in wild geese due to nematode and fungus infection. The North American Veterinarian 13:57–59. Clapham, P. A. 1934. Experimental studies on the transmission of gapeworm (Syngamus trachea) by earthworms. Proceedings of the Royal Society of London. Series B, 115(791):18–29. Clapham, P. A. 1935. On nodules occasioned by gapeworms in pheasants. Journal of Helminthology 13:9–12. Clapham, P. A. 1939. On the larval migration of Syngamus trachea and its causal relationship to pneumonia in young birds. Journal of Helminthology 17:159–162. Cram, E. B. 1927. Bird parasites of the nematode suborders Strongylata, Ascaridata, and Spirurata. U.S. National Museum Bulletin 140:41–48. De Wit, J. J. 1995. Mortality of rheas caused by a Syngamus trachea infection. Veterinary Quarterly 17:39–40.
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Tracheal Worms Draycott, R. A. H., M. I. A. Woodburn, D. E. Ling, and R. B. Sage. 2006. The effect of an indirect anthelmintic treatment on parasites and breeding success of free-living pheasants Phasianus colchicus. Journal of Helminthology 80:409–415. Elton, C., and F. Buckland. 1928. The gapeworm (Syngamus trachea Mont.) in Rooks (Corvus frugilegus L.). Parasitology 20:448–450. Fernando, M. A., P. H. G. Stockdale, and O. Remmler. 1971. The route of migration, development, and pathogenesis of Syngamus trachea (Montagu, 1811) Chapin, 1925, in pheasants. Journal of Parasitology 57:107–116. Fernando, M. A., I. J. Hoover, and S. G. Ogungbade. 1973. The migration and development of Cyathostoma bronchialis in geese. Journal of Parasitology 59:759–764. Hunter, D. B., K. McKeever, and C. Bartlett. 1993. Cyathostoma infections in screech owls, saw-whet owls, and burrowing owls in southern Ontario. In Raptor Biomedicine, P. T. Redig, J. E. Cooper, J. D. Remple, and D. B. Hunter (eds). University of Minnesota Press, Minneapolis, MN, pp. 54–56. Karstad, L., and L. Sileo. 1971. Causes of death in captive wild waterfowl in the Kortright Waterfowl Park, 1967–1970. Journal of Wildlife Diseases 7:236–241. Krone, O., D. Friedrich, and M. Honisch. 2007. Specific status and pathogenicity of syngamid nematodes in bird species (Ciconiformes, Falconiformes, Gruiformes) from Germany. Journal of Helminthology 81:67–73. Lavoie, M., I. Mikaelian, M. Sterner, A. Villeneuve, G. Fitzgerald, J. D. McLaughlin, S. Lair, and D. Martineau. 1999. Respiratory nematodiases in raptors in Quebec. Journal of Wildlife Diseases 35:375–380. LeaMaster, B. R. 2007. Parasites Important to Poultry in Hawaii and Their Control. Cooperative Extension Service, College of Tropical Agriculture and Human Resources, University of Hawaii at Manoa. Livestock Management #18, April 2007, 5 pp. Leers, W.-D., M. K. Sarin, and K. Arthurs. 1985. Syngamosis, an unusual cause of asthma: The first reported case in Canada. Canadian Medical Association Journal 132:269–270. Levine, N. D. 1968. Nematode Parasites of Domestic Animals and Man. Burgess Publishing Co., Minneapolis, MN, 600 pp. Lewis, E. L. 1925. Starlings as distributors of “gapes.” Journal of Helminthology 3:81–82. Lewis, E. L. 1926. Starlings as distributors of “gapes.” Journal of Helminthology 4:43–48. Lichtenfels, J. R. 1980. Keys to the genera of the subfamily Strongyloidea. In CIH Keys to the Nematode Parasites of Vertebrates, No. 7, R. C.
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Anderson, A. G. Chabaud, and S. Willmott (eds). Commonwealth Agricultural Bureaux Slough, Buckinghamshire, pp. 1–41. Lister, S. 1989. Diseases of game birds. In Practice 11:170–174. Madsen, H. 1950. On the systematica of Syngamus trachea (Montagu, 1811) Chapin, 1925. Journal of Helminthology 24:33–46. Moynihan, I. W., and Musfeldt, I. W. 1950. Gapeworm infestation of pheasants. Canadian Journal of Comparative Medicine 14:308–310. Nevarez, J. G., K. C. Gamble, and T. N. Tully, Jr. 2002. Syngamus trachea infection in two red-and-yellow Barbets (Trachyphonus erythrocephalus). Journal of Avian Medicine and Surgery 16(1):31–33. Olivier, L. 1942. Acquired resistance to the gapeworm, Syngamus trachea, in the turkey and ringnecked pheasant. Journal of Parasitology 28(Suppl):20. Ortlepp, R. J. 1923. The life-history of Syngamus trachealis (Montagu) v. Siebold, the gape-worm of chickens. Journal of Parasitology 1:119–140. Pavlovic, I., D. Jakic-Dimic, Z. Kulisic, and I. Florestean. 2003. Most frequent nematode parasites of artificially raised pheasants (Phasianus colchicus L.) and measures for their control. Acta Veterinaria (Beograd) 53:393–398. Pemberton, R. T. 1959. Life cycle of Cyathostoma lari, Blanchard 1849 (Nematoda, Strongyloidea). Nature 184:1423. Rice, J. P. 1929. The rook as a source of gapeworm infection. Journal of the Ministry of Agriculture, Northern Ireland 2:84–87. Ryzhikov, K. M. 1949. Syngamidae of Domestic and Wild Animals. In Essentials of Nematology, Vol. 1, K. I. Skrjabin (ed.). Izdatl’stvo Akademii Nauk SSSR, Moscow, USSR, pp. 1–164. Skrjabin, K. 1915. [Syngamus in birds of Turkestan] (In Russian). Vestnik Obshej Vetereinarii Saint Petersburga 27(17):645–658. Sanmartin, M. L., F. Alvarez, G. Barreiro, and J. Leiro. 2004. Helminth fauna of Falconiform and Strigiform birds of prey in Galicia, Northwest Spain. Parasitology Research 92:255–263. Simpson, V. R., and E. A. Harris. 1992. Cyathostoma lari (Nematoda) infection in birds of prey. Journal of Zoology 227:655–659. Spalding, M. G., J. M. Kinsella, S. A. Nesbitt, M. J. Folk, and G. W. Foster. 1996. Helminth and arthropod parasites of experimentally introduced whooping cranes in Florida. Journal of Wildlife Diseases 32:44–50. Ssenyonga, G. S. 1982. Efficacy of fenbendazole against helminth parasites of poultry in Uganda. Tropical Animal Health Production 14:163–166.
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Taylor, E. L. 1928. Syngamus trachea from the starling transferred to the chicken, and some physiological variation observed. Annals of Tropical Medicine and Parasitology 22:307–318. Threlfall, W. 1965. Life-cycle of Cyathostoma lari. Nature 206:1167–1168. Varga, I. 1971. The pathophysiological effects of Syngamus trachea infection in turkeys. Journal of Comparative Pathology 81:63–69. Varghese, T. 1987. Endoparasites of Birds of Paradise in Papua New Guinea. Veterinary Parasitology 26:131–144. Wehr, E. E. 1937. Observations on the development of the poultry gapeworm Syngamus trachea. Transactions of the American Microscopical Society 56:72–78. Wehr, E. E., and J. C. Hwang. 1967. Anthelmintic activity of thiabendazole against the gapeworm (Syngamus trachea) in turkeys. Avian Diseases 11:44–48.
Welte, S. C., and C. E. Kirkpatrick. 1986. Syngamiasis in juvenile American robins (Turdus migratorius) with a note on the prevalence of other fecal parasites. Avian Diseases 30(4):736–739. Wetzel, R., and H. P. Fortmeyer. 1964. Modifications histo-pathologiques chez les poussins infestes par Syngamus trachea. Economie et M´edecine Animales 5:78–86. Wissler, K., and O. Halvorsen. 1975. The occurrence of gapeworm (Syngamus trachea) in willow grouse. Journal of Wildlife Diseases 11:245–247. Zavadil, R. 1966. Problems of syngamosis in large breeds of turkey-hens. Sbornik Vysoke Skoly Zemedelske v Brne (Rada B) 14:427– 439. Zeigler, K. 1966. Vakcinace kurat proi syngamoze. Veterin´arn´ı Medic´ına 11:569–578. Zeigler, K. 1968. Vakcinace proti syngamose kurat, mladych bazantu a krutat. Veterin´arn´ı Medic´ına 13:97–105.
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20 Amidostomum and Epomidiostomum Alan M. Fedynich and Nancy J. Thomas
SYNONYMS Gizzard worm, ventricular nematodiasis, amidostomiasis, epomidiostomiasis.
a report of Amidostomum sp. in wild Canada Geese (Branta canadensis), Snow Geese (Chen caerulescens), Ross’ Geese (Chen rossii), Northern Pintail (Anas acuta), and Green-winged Teal (Anas carolinensis) in California (O’Roke 1928). Shortly thereafter, Amidostomum spatulatum was reported from Canada Geese (wild or captive not reported), which were also infected with Epomidiostomum crami (Wehr 1933a). Although early literature from the Old World clearly established that A. anseris is pathogenic in anatids, evidence of morbidity and mortality in hosts from North America first emerged in 1926 when Canada Goose goslings were found to be particularly susceptible (Wickware 1941). Subsequent observations indicated that A. anseris could be a contributing factor in mortalities observed in wintering Canada Geese (Herman and Wehr 1954). In the early twentieth century, parasitological surveys of birds (particularly game birds) began to provide a better picture of the worldwide distributions of species of Amidostomum and Epomidiostomum in their respective hosts. Later, a few studies focused on understanding the life histories of several species of gizzard worms occurring in North America (Cowan 1955; Leiby and Olsen 1965). However, the life histories of most species in both genera have not been thoroughly examined.
HISTORY Species of Amidostomum and Epomidiostomum were first reported from gizzards of anatids in Europe beginning as early as 1791 when Froelich described Strongylus mucronatus (=Amidostomum anseris). Linstow, Lundahl, Molin, Rudolphi, and Zeder in the 1800s and Baylis, Boulenger, Maplestone, Seurat, Skrjabin, Travassos, Wehr, and Wetzel in the 1900s found and described species in both genera (Yamaguti 1961; McDonald 1969). In North America, A. anseris was first reported from domestic geese in New York (Cram 1925), followed by
DISTRIBUTION AND HOST RANGE Species of Amidostomum and Epomidiostomum have been found in at least 96 and 60 species of wild birds, respectively (Tables 20.1 and 20.2). Both Amidostomum and Epomidiostomum occur principally in the host family Anatidae within the order Anseriformes, which includes the ducks, geese, and swans. The distribution of these parasites reflects the geographic ranges of their hosts. In migratory waterfowl, Amidostomum acutum, A. anseris, A. spatulatum, and Epomidiostomum uncinatum are widely distributed,
INTRODUCTION Gizzard worms of the genera Amidostomum and Epomidiostomum are commonly found in waterfowl within the family Anatidae. They have direct life cycles and are classified in the order Strongylida (bursate nematodes). Adult worms of both genera live under the koilin lining of the gizzard (ventriculus) and feed on blood. Pathology caused by ventricular nematodiasis has been reported in wild geese (Jerstad 1937; Herman and Wehr 1954; Harradine 1982; Tuggle and Crites 1984), ducks (MacNeill 1970; Crichton and Welch 1972; Turner and Threlfall 1975), and swans (MacNeill 1970). Intense infections can result in damage to the koilin lining and associated muscle and lead to gizzard dysfunction, emaciation, weakness, and potentially poor growth rates of juveniles (MacNeill 1970; Tuggle and Friend 1999). Debilitated hosts may be more susceptible to predation or infection by other pathogens (Tuggle and Friend 1999). Additionally, migratory birds in poor condition from ventricular nematodiasis may be unable to cope with the demands of migration and increased competition for limited food resources during winter (Herman and Wehr 1954).
355 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
356 NR The Netherlands Canada Wyoming, USA North America North America Canada Australia NR NR Newfoundland Sweden Massachusetts, USA
Amidostomum anseris Amidostomum cygni Amidostomum acutum Amidostomum anseris Amidostomum anseris Amidostomum acutum Amidostomum acutum Amidostomum anseris Amidostomum spatulatum Amidostomum acutum Amidostomum anseris Amidostomum acutum Amidostomum acutum Amidostomum acutum Amidostomum anseris Amidostomum fulicae
Black Swan (Cygnus atratus) Blue-winged Teal (Anas discors)
Canvasback (Aythya valisineria) Cape Barren Goose (Cereopsis novaehollandiae) Chiloe Wigeon (Anas sibilatrix) Cinnamon Teal (Anas cyanoptera) Common Eider (Somateria mollissima)
Brant (Branta bernicla) Bufflehead (Bucephala albeola) Canada Goose (Branta canadensis)
Canada The Netherlands NR Australia North America
Europe Texas, USA
Amidostomum spatulatum Amidostomum acutum Amidostomum acutum
NR Canada NR Canada Washington, USA NR NR NR The Netherlands Canada Europe
Location
Amidostomum anseris Amidostomum acutum Amidostomum anseris Amidostomum acutum Amidostomum anseris Amidostomum anseris Amidostomum acutum Amidostomum spatulatum Amidostomum anseris Amidostomum anseris Amidostomum anseris
Parasite
Bourgeois and Threlfall (1982) Borgsteede et al. (2006) Gower (1939) Mawson (1980) Turner and Threlfall (1975) and Wallace and Pence (1986) McDonald (1969) Borgsteede et al. (2006) McLaughlin and McGurk (1987) NWHC (voucher no. 48808)* Jerstad (1937) and Wehr and Herman (1954) Wehr (1933a) and Purvis et al. (1997) McLaughlin and McGurk (1987) Mawson (1980) McDonald (1969) McDonald (1969) Bishop and Threlfall (1974) Persson et al. (1974) NWHC (voucher no. 48809)
McDonald (1969) Mahoney and Threlfall (1978) McDonald (1969) McLaughlin and McGurk (1987) Jerstad (1937) Gower (1939) Czapli´nski (1962) McDonald (1969) Borgsteede et al. (2006) MacNeill (1970) Balicka-Ramisz et al. (2000) and Borgsteede et al. (2006) Baylis (1932) and Macko et al. (2002) Fedynich et al. (1996a)
Reference
September 29, 2008
Black-bellied Whistling-Duck (Dendrocygna autumnalis) Black Scoter (Melanitta nigra)
Ashy-headed Goose (Chloephaga poliocephala) Baikal Teal (Anas formosa) Bar-headed Goose (Anser indicus) Barnacle Goose (Branta leucopsis) Barrow’s Goldeneye (Bucephala islandica) Taiga Bean-Goose (Anser fabalis)
American Wigeon (Anas americana)
Anseriformes: Anatidae African Black Duck (Anas sparsa) American Black Duck (Anas rubripes)
Host
Table 20.1. Avian hosts infected with Amidostomum spp., geographic location where infection was reported, and reporting authors.
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357
Green-winged Teal (Anas carolinensis)
Greater White-fronted Goose (Anser albifrons)
Gray Teal (Anas gracilis) Greater Scaup (Aythya marila)
Garganey (Anas querquedula)
Freckled Duck (Stictonetta naevosa) Gadwall (Anas strepera)
Europe Canada Canada Europe NR The Netherlands NR NR
Amidostomum acutum Amidostomum anseris Amidostomum acutum Amidostomum acutum Amidostomum fulicae Amidostomum acutum Amidostomum acutum
Europe NR NR Europe Europe Australia Canada The Netherlands Amidostomum anseris NR Amidostomum fulicae NR Amidostomum acutum Europe Amidostomum anseris NR Amidostomum auriculatum Europe Amidostomum acutum Australia Amidostomum acutum The Netherlands Amidostomum anseris NR Amidostomum acutum Europe Amidostomum anseris Texas, USA Europe Amidostomum spatulatum Europe Texas, USA Amidostomum acutum Europe Taiwan Canada Amidostomum anseris Texas, USA Amidostomum auriculatum Europe
Amidostomum anseris Amidostomum acutum Amidostomum anseris Amidostomum acutum Amidostomum anseris Amidostomum acutum
Amidostomum acutum
Petrova (1987) and Borgsteede et al. (2006) Gower (1939) McDonald (1969) Czapli´nski (1962) Czapli´nski (1962) Mawson (1980) McLaughlin and McGurk (1987) Borgsteede et al. (2006) McDonald (1969) McDonald (1969) Czapli´nski (1962) Gower (1939) Lomakin (1988) Mawson (1980) Borgsteede et al. (2006) Gower (1939) Czapli´nski (1962) Purvis et al. (1997) Balicka-Ramisz et al. (2000) and Borgsteede et al. (2006) Czapli´nski (1962) Purvis et al. (1997) Czapli´nski (1962) Schmidt and Kuntz (1972) Turner and Threlfall (1975) Canaris et al. (1981) Lomakin (1988) (continues)
Czapli´nski (1962) Mahoney and Threlfall (1978) MacNeill (1970) Czapli´nski (1962) McDonald (1969) Borgsteede et al. (2006) McDonald (1969) Czapli´nski (1962)
September 29, 2008
Falcated Duck (Anas falcata) Ferruginous Pochard (Aythya nyroca)
Cotton Pygmy-goose (Nettapus coromandelianus) Eurasian Wigeon (Anas penelope)
Common Shelduck (Tadorna tadorna)
Common Pochard (Aythya ferina)
Common Goldeneye (Bucephala clangula)
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Northern Pintail (Anas acuta)
Musk Duck (Biziura lobata) Mute Swan (Cygnus olor)
Mandarin Duck (Aix galericulata) Maned Duck (Chenonetta jubata) Marbled Teal (Marmaronetta angustirostris) Mottled Duck (Anas fulvigula)
Long-tailed Duck (Clangula hyemalis) Mallard (Anas platyrhynchos)
Lesser White-fronted Goose (Anser erythropus)
McDonald (1969) McDonald (1969) McDonald (1969) Petrova (1987) and Bolte et al. (2000) McDonald (1969) McDonald (1969) McDonald (1969) Bailey and Black (1995) Czapli´nski (1962) McDonald (1969) Czapli´nski (1962) McLaughlin and McGurk (1987) McDonald (1969) McDonald (1969) McDonald (1969) Czapli´nski (1962) Czapli´nski (1962) Mawson (1980) McLaughlin and McGurk (1987) and Gray et al. (1989) Crichton and Welch (1972) Petrova (1987) Nakamura and Asakawa (2001) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) Kinsella and Forrester (1972) Fedynich et al. (1996b) Mawson (1980) Borgsteede et al. (2006) Yamaguti (1961) McLaughlin and McGurk (1987) Petrova (1987) and Borgsteede et al. (2006) Crichton and Welch (1972)
NR NR NR Europe NR NR NR Hawaii, USA NR NR North America Canada NR NR NR NR Europe Australia North America Canada Bulgaria Japan NR NR NR NR Florida, USA Texas, USA Australia The Netherlands Belgium Canada Europe Canada Amidostomum fulicae Amidostomum henryi Amidostomum acutum Amidostomum anseris Amidostomum spatulatum Amidostomum acutum Amidostomum anseris Amidostomum anseris Amidostomum acutum Amidostomum anseris Amidostomum cygni Amidostomum acutum Amidostomum anseris Amidostomum anseris Amidostomum spatulatum Amidostomum acutum Amidostomum acutum
Amidostomum anseris
Amidostomum acutum Amidostomum anseris Amidostomum cygni Amidostomum acutum
Amidostomum fulicae Amidostomum acutum Amidostomum anseris Amidostomum acutum Amidostomum acutum
Amidostomum anseris
Reference
Location
Parasite
September 29, 2008
Lesser Scaup (Aythya affinis)
Hawaiian Goose (Branta sandvicensis) King Eider (Somateria spectabilis)
Harlequin Duck (Histrionicus histrionicus)
Greylag Goose (Anser anser)
Host
Table 20.1. (Continued)
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Amidostomum acutum
North America Europe Amidostomum anseris NR Pacific Black Duck (Anas superciliosa) Amidostomum acutum Australia Pink-footed Goose (Anser brachyrhynchus) Amidostomum anseris NR Amidostomum spatulatum NR Radjah Shelduck (Tadorna radjah) Amidostomum acutum Australia Red-breasted Goose (Branta ruficollis) Amidostomum ryzhikovi Russia (zoo) Red-crested Pochard (Netta rufina) Amidostomum acutum NR Amidostomum fulicae NR Redhead (Aythya americana) Amidostomum acutum Canada Amidostomum anseris Washington, USA Ring-necked Duck (Aythya collaris) Amidostomum acutum Canada Ross’ Goose (Chen rossii) Amidostomum anseris Texas, USA Amidostomum spatulatum NR Ruddy Duck (Oxyura jamaicensis) Amidostomum acutum Canada Ruddy Shelduck (Tadorna ferruginea) Amidostomum acutum NR Amidostomum anseris NR Smew (Mergellus albellus) Amidostomum acutum NR Snow Goose (Chen caerulescens) Amidostomum anseris North America Amidostomum cygni NR Amidostomum spatulatum North America Spectacled Duck (Anas specularis) Amidostomum anseris NR Spectacled Eider (Somateria fischeri) Amidostomum fulicae Alaska, USA Spot-billed Duck (Anas poecilorhyncha) Amidostomum acutum India NR Amidostomum fulicae Surf Scoter (Melanitta perspicillata) Amidostomum acutum Canada Amidostomum fulicae Virginia, USA Trumpeter Swan (Cygnus buccinator) Amidostomum anseris Canada Tufted Duck (Aythya fuligula) Amidostomum acutum The Netherlands Amidostomum anseris NR
Northern Shoveler (Anas clypeata)
Broderson et al. (1977) and McLaughlin and McGurk (1987) Petrova (1987) and Borgsteede et al. (2006) McDonald (1969) Mawson (1980) McDonald (1969) McDonald (1969) Mawson (1980) Kosupko and Lomakin (1985) Czapli´nski (1962) McDonald (1969) McLaughlin and McGurk (1987) Jerstad (1937) McLaughlin and McGurk (1987) Fedynich et al. (2005) McDonald (1969) McLaughlin and McGurk (1987) McDonald (1969) McDonald (1969) McDonald (1969) Tuggle and Crites (1984) and Purvis et al. (1997) McDonald (1969) Tuggle and Crites (1984) and Purvis et al. (1997) McDonald (1969) NWHC (voucher no. 48802, 48803) Dubey and Pande (1964) McDonald (1969) Bourgeois and Threlfall (1982) NWHC (voucher no. 48810) MacNeill (1970) Czapli´nski (1962) and Borgsteede et al. (2006) Gower (1939) (continues)
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359
360 Amidostomum acutum Amidostomum acutum Amidostomum acutum Amidostomum acutum Amidostomum cygni Amidostomum anseris Amidostomum acutum
Gaviiformes: Gaviidae Red-throated Loon (Gavia stellata)
Podicipediformes: Podicipedidae Little Grebe (Tachybaptus ruficollis)
Gruiformes: Aramidae Limpkin (Aramus guarauna)
Amidostomum spatulatum Amidostomum acutum Amidostomum anseris Amidostomum acutum Amidostomum acutum
Galliformes: Phasianidae Black-billed Capercaillie (Tetrao parvirostris) Eurasian Capercaillie (Tetrao urogallus) Hazel Grouse (Bonasa bonasia) Rock Ptarmigan (Lagopus muta)
Yellow-billed Duck (Anas undulata) Yellow-billed Pintail (Anas georgica)
Wood Duck (Aix sponsa)
Amidostomum anseris Amidostomum acutum Amidostomum anseris Amidostomum cygni
Amidostomum spatulatum Amidostomum anseris Amidostomum anseris Amidostomum acutum
Amidostomum cygni
Amidostomum acutum Amidostomum anseris
Parasite
Florida, USA
NR
North America
Europe Europe NR NR
NR Canada Japan Washington DC, USA (zoo) Europe NR Falkland Islands NR Canada The Netherlands NR NR NR Bulgaria Japan NR North America NR Africa NR
Location
Conti et al. (1985)
McDonald (1969)
Czapli´nski (1962)
Baruˇs et al. (1984) Baruˇs et al. (1984) Czapli´nski (1962) Czapli´nski (1962)
McDonald (1969) MacNeill (1970) Nakamura and Asakawa (2001) Wehr (1933b) Czapli´nski (1962) McDonald (1969) Harradine (1982) McDonald (1969) Bourgeois and Threlfall (1982) Borgsteede et al. (2006) Gower (1939) McDonald (1969) McDonald (1969) Petrova (1987) Nakamura and Asakawa (2001) Czapli´nski (1962) Thul et al. (1985) McDonald (1969) Alexander and McLaughlin (1997) McDonald (1969)
Reference
September 29, 2008
Whooper Swan (Cygnus cygnus)
Upland Goose (Chloephaga picta) White-headed Duck (Oxyura leucocephala) White-winged Scoter (Melanitta fusca)
Tundra Swan (Cygnus columbianus)
Host
Table 20.1. (Continued)
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361 Amidostomum acutum Amidostomum acutum Amidostomum acutum Amidostomum acutum Amidostomum acutum
Charadriiformes: Recurvirostridae Black-winged Stilt (Himantopus himantopus) Pied Avocet (Recurvirostra avosetta) Pied Stilt (Himantopus leucocephalus)
Charadriiformes: Scolopacidae Common Snipe (Gallinago gallinago)
Wood Sandpiper (Tringa glareola)
Czech Republic
Canada Taiwan NR
France The Netherlands Australia
Bulgaria
NR NR Australia NR Spain Bulgaria NR NR California Iraq Europe NR
Ryˇsav´y et al. (1955)
Threlfall (1970) Schmidt and Kuntz (1972) McDonald (1969)
Czapli´nski (1962) Borgsteede et al. (2006) Mawson (1980)
Petrova (1987)
Roudabush (1942) McDonald (1969) Mawson (1980) Gower (1939) Acosta et al. (1992) Petrova (1987) Czapli´nski (1962) Gower (1939) Yamaguti (1961) Mahmoud and Mohammad (1989) Borgsteede et al. (2006) Czapli´nski (1962)
Note: Some host–parasite reports found only in the review literature did not include geographic location and often included hosts from wild, domestic, and private and public zoological collections. NR, not reported. * NWHC = USGS National Wildlife Health Center, Diagnostic Parasitology Laboratory, Madison, WI, USA; specimens are deposited at the Harold W. Manter Laboratory of Parasitology at the University of Nebraska State Museum, Lincoln, NE, USA; specimen voucher numbers are provided.
Amidostomum anseris
Amidostomum henryi
Columbiformes: Columbidae Eurasian Collared-Dove (Streptopelia decaocto)
Amidostomum fulicae
Spotted Crake (Porzana porzana)
Amidostomum acutum Amidostomum fulicae Amidostomum tribonyx Amidostomum anseris Amidostomum fulicae Amidostomum quasifulicae Amidostomum acutum Amidostomum anseris Amidostomum fulicae
September 29, 2008
Charadriiformes: Charadriidae Northern Lapwing (Vanellus vanellus)
Eurasian Coot (Fulica atra)
Black-tailed Native-hen (Gallinula ventralis) Common Moorhen (Gallinula chloropus)
Gruiformes: Rallidae American Coot (Fulica americana)
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Common Pochard (Aythya ferina)
Common Eider (Somateria mollissima) Common Goldeneye (Bucephala clangula)
Canvasback (Aythya valisineria) Cape Shoveler (Anas smithii) Cape Teal (Anas capensis) Cinnamon Teal (Anas cyanoptera) Comb Duck (Sarkidiornis melanotos)
Brant (Branta bernicla) Canada Goose (Branta canadensis)
Black-bellied Whistling-Duck (Dendrocygna autumnalis) Black-necked Swan (Cygnus melancoryphus) Blue-winged Teal (Anas discors) Argentina North America New Jersey, USA North America NR Canada Africa Africa NR India India India NR Sweden Canada NR NR Bulgaria
Epomidiostomum crami* Epomidiostomum crami Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum asymmetricum Epomidiostomum sarkidiorni Epomidiostomum sultanai Epomidiostomum uncinatum Epomidiostomum orispinum Epomidiostomum crami Epomidiostomum orispinum Epomidiostomum orispinum Epomidiostomum penelopi
NR India NR NR Europe NR NR NR Texas, USA
Canada North America
Location
Epomidiostomum vogelsangi† Epomidiostomum uncinatum
Epomidiostomum uncinatum Epomidiostomum skrjabini* Epomidiostomum orispinum Epomidiostomum crami Epomidiostomum orispinum Epomidiostomum uncinatum Epomidiostomum orispinum Epomidiostomum uncinatum Epomidiostomum uncinatum
Epomidiostomum uncinatum Epomidiostomum uncinatum
Parasite
Oliveira (1970) Turner and Threlfall (1975) and Shaw and Kocan (1980) NWHC (voucher no. 48801)‡ Wetzel (1931) and Purvis et al. (1997) McDonald (1969) McLaughlin and McGurk (1987) Alexander and McLaughlin (1997) Alexander and McLaughlin (1997) McDonald (1969) Jairajpuri and Siddiqi (1970) Jairajpuri and Siddiqi (1970) Ali (1971a) McDonald (1969) Persson et al. (1974) Mahoney and Threlfall (1978) McDonald (1969) Gower (1939) Petrova (1989)
Mahoney and Threlfall (1978) Shaw and Kocan (1980) and McLaughlin and McGurk (1987) McDonald (1969) Ali (1971b) McDonald (1969) McDonald (1969) Macko et al. (2002) McDonald (1969) Gower (1939) Gower (1939) Fedynich et al. (1996a)
Reference
September 29, 2008
Black Scoter (Melanitta nigra)
Baikal Teal (Anas formosa) Bar-headed Goose (Anser indicus) Barnacle Goose (Branta leucopsis) Taiga Bean-Goose (Anser fabalis)
Anseriformes: Anatidae American Black Duck (Anas rubripes) American Wigeon (Anas americana)
Host
Table 20.2. Avian hosts infected with Epomidiostomum spp., geographic location where infection was reported, and reporting authors.
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363
Greylag Goose (Anser anser)
Green-winged Teal (Anas carolinensis)
Gray Teal (Anas gracilis) Greater Scaup (Aythya marila) Greater White-fronted Goose (Anser albifrons)
Garganey (Anas querquedula)
Fulvous Whistling-Duck (Dendrocygna bicolor) Gadwall (Anas strepera)
Epomidiostomum crami
Epomidiostomum subquadratum Epomidiostomum uncinatum Epomidiostomum subquadratum Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum crami Epomidiostomum orispinum Epomidiostomum skrjabini Epomidiostomum uncinatum Epomidiostomum orispinum Epomidiostomum querquedulae Epomidiostomum subquadratum Epomidiostomum uncinatum
China NR NR NR Alaska, USA NR NR China Bulgaria China NR NR India NR India Texas, USA China North America China Bulgaria NR NR Texas, USA Bulgaria Russia NR NR Egypt China Taiwan Canada Bulgaria NR
Shen (1981) McDonald (1969) McDonald (1969) McDonald (1969) NWHC (voucher no. 48804) McDonald (1969) Gower (1939) Shen and Wu (1973) Petrova (1989) Shen and Wu (1973) McDonald (1969) Ali (1971b) Ali (1971a) McDonald (1969) Ali (1971a) Fedynich et al. (1996a) Shen and Wu (1973) Buscher (1965) Shen and Wu (1973) Petrova (1989) McDonald (1969) McDonald (1969) Purvis et al. (1997) Petrova (1989) Yamaguti (1961) Gower (1939) Gower (1939) Yamaguti (1961) Shen and Wu (1973) Schmidt and Kuntz (1972) Turner and Threlfall (1975) Petrova (1989) McDonald (1969) (continues)
September 29, 2008
Ferruginous Pochard (Aythya nyroca)
Falcated Duck (Anas falcata)
Emperor Goose (Chen canagica) Eurasian Wigeon (Anas penelope)
Common Shelduck (Tadorna tadorna)
Epomidiostomum serratum Epomidiostomum uncinatum Epomidiostomum crami Epomidiostomum uncinatum Epomidiostomum crami Epomidiostomum crami Epomidiostomum orispinum Epomidiostomum subquadratum Epomidiostomum uncinatum Epomidiostomum subquadratum Epomidiostomum uncinatum Epomidiostomum orispinum Epomidiostomum sultanai Epomidiostomum uncinatum Epomidiostomum uncinatum
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Redhead (Aythya americana) Ring-necked Duck (Aythya collaris)
Red-billed Duck (Anas erythrorhyncha) Red-crested Pochard (Netta rufina)
Northern Shoveler (Anas clypeata)
Epomidiostomum serratum Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum crami Epomidiostomum querquedulae Epomidiostomum subquadratum Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum crami
Epomidiostomum cygni Epomidiostomum querquedulae Epomidiostomum uncinatum Epomidiostomum alii Epomidiostomum crami Epomidiostomum querquedulae Epomidiostomum uncinatum
Mute Swan (Cygnus olor)
Northern Pintail (Anas acuta)
Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum uncinatum
Epomidiostomum orispinum Epomidiostomum skrjabini Epomidiostomum uncinatum Epomidiostomum orispinum Epomidiostomum uncinatum Epomidiostomum crami Epomidiostomum orispinum Epomidiostomum petalum§ Epomidiostomum serratum Epomidiostomum uncinatum
Parasite Bulgaria Russia NR NR NR NR Bulgaria China China North America Europe NR NR Florida, USA Texas, USA China NR NR India NR NR North America Bulgaria China North America Africa NR India China NR Canada Canada
Location Petrova (1989) Yamaguti (1961) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) Petrova (1989) Yen and Wu (1959) Shen (1981) Crichton and Welch (1972) Birov´a et al. (1990) McDonald (1969) McDonald (1969) Kinsella and Forrester (1972) Fedynich et al. (1996b) Shen and Wu (1964) McDonald (1969) McDonald (1969) Ali (1971a) McDonald (1969) McDonald (1969) Buscher (1965) Petrova (1989) Shen (1981) Buscher (1965) Alexander and McLaughlin (1997) McDonald (1969) Ali (1971a) Shen and Wu (1973) McDonald (1969) McLaughlin and McGurk (1987) Noseworthy and Threlfall (1978)
Reference
September 29, 2008
Mandarin Duck (Aix galericulata) Marbled Teal (Marmaronetta angustirostris) Mottled Duck (Anas fulvigula)
Lesser White-fronted Goose (Anser erythropus) Long-tailed Duck (Clangula hyemalis) Mallard (Anas platyrhynchos)
Host
Table 20.2. (Continued)
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365
Gower (1939) Petrova (1989)
NR Bulgaria
Epomidiostomum orispinum
Fedynich et al. (2005) McLaughlin and McGurk (1987) Ali (1971a) McDonald (1969) Purvis et al. (1997) and Tuggle and Crites (1984) McDonald (1969) Shen (1981) Shen and Wu (1973) McDonald (1969) Gower (1939) McDonald (1969) McDonald (1969) McDonald (1969) McDonald (1969) Bourgeois and Threlfall (1982) McDonald (1969) Thul et al. (1985) Alexander and McLaughlin (1997)
Epomidiostomum orispinum
Texas, USA Canada India NR North America NR China China NR NR NR NR NR NR Canada NR North America Africa
Note: Some host–parasite reports found only in the review literature did not include geographic location and often included hosts from wild, domestic, and private and public zoological collections. NR, not reported. * Ali (1971b) redescribed E. skrjabini, which has been considered a synonym of E. orispinum (McDonald 1969), and regards E. crami to be a synonym of E. skrjabini. † Some references (Yamaguti 1961; McDonald 1969) report E. vogelsangi as a synonym of E. orispinum; however, Ali (1971b) considers E. vogelsangi a valid species. ‡ NWHC = USGS National Wildlife Health Center, Diagnostic Parasitology Laboratory, Madison, WI, USA; specimens are deposited at the Harold W. Manter Laboratory of Parasitology at the University of Nebraska State Museum, Lincoln, NE, USA; specimen voucher numbers are provided. § Described from domestic Mallard (A. platyrhynchos var. domestica).
Gruiformes: Rallidae Eurasian Coot (Fulica atra) Charadriiformes: Charadriidae Northern Lapwing (Vanellus vanellus)
Whooper Swan (Cygnus cygnus) Wood Duck (Aix sponsa) Yellow-billed Duck (Anas undulata)
Tundra Swan (Cygnus columbianus) White-headed Duck (Oxyura leucocephala) White-winged Scoter (Melanitta fusca)
Epomidiostomum crami Epomidiostomum uncinatum Epomidiostomum querquedulae Epomidiostomum uncinatum Epomidiostomum crami Epomidiostomum petalum Epomidiostomum serratum Epomidiostomum subquadratum Epomidiostomum uncinatum Epomidiostomum orispinum Epomidiostomum uncinatum Epomidiostomum crami Epomidiostomum uncinatum Epomidiostomum orispinum Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum uncinatum Epomidiostomum uncinatum
September 29, 2008
Tufted Duck (Aythya fuligula)
Snow Goose (Chen caerulescens) Spot-billed Duck (Anas poecilorhyncha)
Ross’ Goose (Chen rossii) Ruddy Duck (Oxyura jamaicensis) Ruddy Shelduck (Tadorna ferruginea)
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Parasitic Diseases of Wild Birds
occurring on 4–5 continents (McDonald 1969; Tables 20.1 and 20.2). Other species such as Amidostomum tribonyx, Epomidiostomum alii, Epomidiostomum asymmetricum, Epomidiostomum sarkidiorni, Epomidiostomum serratum, and Epomidiostomum sultanai have been reported in relatively few hosts and seem to be restricted to specific geographic regions (Tables 20.1 and 20.2). Other species of Amidostomum have been reported infrequently in six other taxonomic orders of birds (Table 20.1). Amidostomum fulicae is a parasite of the Rallidae in the order Gruiformes, which is found infrequently in anatids (Table 20.1). Species of Epomidiostomum appear more restricted to the Anatidae, and have been reported in one host species each from the Gruiformes and Charadriiformes (Table 20.2). With the notable exception of certain dove, grouse, and ptarmigan species, hosts infected with species of Amidostomum and Epomidiostomum are typically associated with aquatic habitats (Tables 20.1 and 20.2). There appears to be some association between the two genera of gizzard worms and taxonomic tribe within the Anatidae. For example, A. anseris and E. crami are principally found in Anserini, A. acutum and E. uncinatum occur mainly in Anatini and Aythyini, and Amidostomum cygni is found primarily in Cygnini (Czapli´nski 1962; McDonald 1969). However, some early authors did not distinguish between A. acutum and A. anseris, particularly in species of ducks, so the affinities these two species of nematodes may have for specific tribes is masked in the summarized scientific literature (Czapli´nski 1962; Table 20.1). It remains unclear whether associations are related to host specificity or other factors such as differences in host diet, feeding strategies, or specific life cycle requirements of the parasites. For example, successful experimental infections in hatchling chickens (Phasianidae) with Amidostomum raillieti (=Amidostomum fulicae, normally found in coots) and in hatchling chickens and pigeons (Columbidae) with E. uncinatum (primarily found in ducks) suggest a lack of host specificity (Leiby and Olsen 1965). However, laboratory experiments using A. anseris were unsuccessful in infecting swan, coot, avocet, godwit, gull, blackbird, crow, dove, chicken, and turkey (Ruff 1984 citing Enigk and Dey-Hazra 1968a). In another experiment, chickens were infected with larval stages of A. anseris, but the worms did not mature, suggesting that they are abnormal hosts (Phuc and Varga 1975). ETIOLOGY Species of Amidostomum and Epomidiostomum are classified in the phylum Nemata (Nematoda), class Secernentea, order Strongylida (bursate nematodes), and
family Amidostomatidae (Anderson 2000). The subfamily Amidostomatinae contains the genus Amidostomum, and the subfamily Epomidiostomatinae contains the genus Epomidiostomum (Anderson 2000). In 1962, the genus Amidostomum was revised by Czapli´nski (1962) and a number of species entered into synonymy, resulting in six recognized species: Amidostomum acutum, A. anseris, A. cygni, A. fulicae, Amidostomum henryi, and A. spatulatum. Since then, three additional species—Amidostomum auriculatum, Amidostomum ryzhikovi, and Amidostomum tribonix—have been described (Mawson 1980; Kosupko and Lomakin 1985; Lomakin 1988) and one (Amidostomum biziurae) redescribed and taken out of synonymy with A. acutum (Mawson 1980). Recently, there has been an effort to revise the classification of species within the subfamily Amidostomatinae. On the basis of morphological characteristics, Lomakin (1993) proposed a revision of the genera in which Amidostomum (after Czapli´nski 1962) is expanded into four genera—Amidostomum with three species: Amidostomum anseris, A. cygni, and A. spatulatum; Mesamidostomum with one species: Mesamidostomum skrjabini; Amidostomoides with six species: Amidostomum acutum, A. auriculatum, A. henryi, A. monodon, A. petrovi, and A. tribonix; and Quasiamidostomum with one species: Q. fulicae. However, this revised classification has not been readily adopted, indicating a need for genetic analyses to help delineate species within the Amidostomatinae (Borgsteede et al. 2006). Depending on the taxonomic authority, there are at least 14 species in the genus Epomidiostomum (Table 20.2) including E. alii, E. asymmetricum, E. crami, Epomidiostomum cygni, Epomidiostomum orispinum, Epomidiostomum penelopi, Epomidiostomum petalum, Epomidiostomum querquedulae, E. sarkidiorni, E. serratum, Epomidiostomum subquadratum, E. sultanai, E. uncinatum, and Epomidiostomum vogelsangi. As with Amidostomum, some species have been synonymized, then redescribed and separated, thereby making it difficult to determine the true number of species within this genus based solely on morphological differences. Adult Amidostomum spp. are typically found in the koilin lining of the gizzard, whereas Epomidiostomum spp. can also invade the gizzard muscle (Tuggle and Crites 1984). They often appear white to red and are relatively long and narrow in shape, having a threadlike appearance. In both genera, adult males are shorter and thinner than females. Depending on the species, the males of Amidostomum range in length from 5.9 to 18 mm, and females range from 6.5 to 27.7 mm (Czapli´nski 1962). Males of Epomidiostomum range in length from 5.3 to 11 mm, and females range from 7.4 to 19 mm (Cram 1927; Wetzel 1931; Jairajpuri and
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Amidostomum and Epomidiostomum Siddiqi 1970; Ali 1971a, b; Macko et al. 2002). Descriptions and figures of morphological features of each species can be found in taxonomic reviews (Czapli´nski 1962; Ali 1971b; Petrova 1987; Lomakin 1993) and original species descriptions or redescriptions (see Literature Cited section). EPIZOOTIOLOGY Life Cycle Species of Amidostomum and Epomidiostomum have a direct life cycle. Partially developed eggs are released from the adult female worm into the gastric lumen of the infected host and excreted into the environment with the feces. Survivability of eggs and larvae in the environment varies. Eggs and larvae require moist conditions for survival (Anderson 2000). The inability of eggs to withstand desiccation helps to explain the close association with the Anatidae. Winter temperatures will inhibit larval development within the egg (Stradowski 1971) and prevent hatching (Cowan 1955). Eggs of A. anseris are able to withstand periods of freezing, but most larvae are killed (Stradowski 1975). Warmer temperatures of 18–23◦ C increase the rate of embryo development to third-stage larvae, with eggs hatching in 23–72 h (Cowan 1955; Leiby and Olsen 1965; McDonald 1969). In A. anseris, the rate of embryo development and hatching is asynchronous, which may be a survival strategy for geographic regions that have unpredictable late winter and early spring conditions (Stradowski 1975). After a short free-living stage of about 4–9 days, third-stage larvae become infective to susceptible hosts and can remain viable up to 20 days under suitable environmental conditions (Leiby and Olsen 1965; McDonald 1969). One study suggests that larvae of A. anseris are infective to domestic geese (Anser anser dom.) within as little as an hour after hatching (Stradowski 1971). Susceptible hosts become infected when they ingest the infective third-stage larvae during feeding or drinking. Additionally, larvae of A. anseris can penetrate the skin and migrate to the lungs and trachea prior to invading the gizzard lining (Anderson 2000 citing Enigk and Dey-Hazra 1968b, 1969, 1970). Larvae penetrate the thinner portions of the gizzard epithelium at the junction of the gizzard and proventriculus after ingestion (Leiby and Olsen 1965; Anderson 2000). Development to the fourth stage requires 2–4 days (Leiby and Olsen 1965), and approximately 12–25 days are required for worms to mature to adults (McDonald 1969). Host age can influence the rate at which worms mature. For example, the prepatent phase of A. anseris in geese is shorter in goslings than in adults (Cowan 1955; Stradowski 1977).
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Prevalence, Intensity, and Abundance1 Gizzard worms are some of the most commonly found species of helminths infecting waterfowl. Parasitological surveys examining more than 50 host individuals of the same species have found prevalances of infection with Amidostomum spp. greater than 50% (Herman and Wehr 1954; Persson et al. 1974; Bourgeois and Threlfall 1982; Thul et al. 1985; McLaughlin and McGurk 1987; Fedynich and Pence 1994; Nowicki et al. 1995). Additionally, some studies have reported prevalences of greater than 90%. For example, 96% of 94 Blue-winged Teal (Anas discors) were infected with A. acutum (Wallace and Pence 1986), 98% of 329 Canada Geese were infected with A. anseris (Herman and Wehr 1954), and 100% of 117 Common Eiders (Somateria mollissima) were infected with A. acutum (Borgsteede et al. 2006). A few studies have found prevalence of infection with species of Epomidiostomum that exceeds 50% (Kinsella and Forrester 1972; Wallace and Pence 1986; McLaughlin and McGurk 1987), but it appears that Amidostomum spp. often occur at a higher prevalence. Additional research is needed to determine why Amidostomum spp. may be more prevalent than Epomidiostomum spp. Numerous host intrinsic and environmental extrinsic factors are likely to play a role in determining prevalence, intensity, and abundance of gizzard nematodes. Differences in one or more of these factors have been found among waterfowl tribes. For example, in a study conducted at Delta, Manitoba, Canada, during autumn 1979, E. uncinatum was found in each of seven species of dabbling ducks (Anatini), whereas only three of six species of diving ducks (Aythyini) were infected (McLaughlin and McGurk 1987). Rapid development of infective stages in shallow wetlands, coupled with generalistic and intermediate foraging strategies of Anas spp., may account for these observed differences in prevalence (McLaughlin and McGurk 1987). Within host species, the most commonly examined variables (and most easily measured) include host age (usually juvenile and adult), host sex, geographic location, and season of collection. Several studies have reported age-related effects. For example, prevalence, mean intensity, and abundance of A. acutum and E. uncinatum were higher in juvenile Mallards (Anas platyrhynchos) than in adults during autumn 1979 at Delta, Manitoba (McLaughlin and McGurk 1987). Rank abundance of A. acutum in Mallards from the 1
Studies reporting prevalence, intensity, and abundance represent observations made at specific points in time and, thus, do not necessarily indicate present dynamics of hosts and parasites.
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Southern High Plains was higher in juveniles than in adults during two consecutive winter and summer periods (Fedynich and Pence 1994). Rank abundance of A. acutum in Blue-winged Teal collected from the Texas panhandle did not vary by age, but E. uncinatum was higher in juveniles than in adults (Wallace and Pence 1986). Juvenile geese and swans often are more severely infected than adults (Herman and Wehr 1954; Pennycott 1998). Although age-dependent immunity is thought to influence infections of certain helminths (Wehr and Herman 1954), research is needed to elucidate the relationship, if any, between host age and the ability to suppress infections of gizzard nematodes. Host sex, coupled with age, has been found to influence prevalence, mean intensity, and abundance of some gizzard nematodes. For example, abundance of A. acutum in adult male Lesser Scaup (Aythya affinis) was higher than that found in adult females (McLaughlin and McGurk 1987). Mean intensity of E. uncinatum was higher in adult female and juvenile female American Wigeon (Anas americana) than in adult males and juvenile males (McLaughlin and McGurk 1987). It is uncertain why differences in prevalence or intensity of gizzard worms is associated by host sex in some species and not others. There is some speculation that acquisition of helminth species may be influenced by variation in habitat use, foraging time, and/or diet (Fedynich et al. 2005), which permits different exposure probabilities to infective parasitic stages. Seasonal variation in prevalence, mean intensity, and abundance has been reported for Amidostomum spp. and Epomidiostomum spp., and could be attributed to a wide range of factors such as seasonal changes in diet or differences in exposure rates among breeding, migrating, and wintering birds. For example, prevalence of A. acutum in wild Mallards was lowest in January (5%) and highest in June (43%), whereas prevalence of E. uncinatum was lowest in September (19%) and highest in May (58%) (Birov´a et al. 1990). Abundance of E. uncinatum was lower in Blue-winged Teal collected in the fall than in those collected during spring (Wallace and Pence 1986). Prevalence and intensity of A. anseris and E. crami were similar between the incubation and brood-rearing periods in adult female Snow Geese at La P´erouse Bay, Manitoba, whereas prevalence and intensity of A. spatulatum were higher during the incubation period and declined during the brood-rearing period (Clinchy and Barker 1994). Findings in Snow Geese suggest a geographic and seasonal component to infections where transmission of E. crami was occurring throughout summer on the breeding grounds and A. spatulatum was being acquired primarily on the wintering grounds (Clinchy and Barker 1994).
CLINICAL SIGNS Mild infections are usually accompanied by no clinical signs. Signs of severe infections in wild birds are nonspecific, and include unthriftiness, loss of appetite, anemia, emaciation, and general weakness (Herman and Wehr 1954; MacNeill 1970; Tuggle and Friend 1999). Trumpeter Swans (Cygnus buccinator) infected with A. anseris exhibit disorientation, staggered gait, and impaction of the esophagus (MacNeill 1970). In Canada Geese, the intensity of infection with A. anseris was negatively correlated with body condition (Herman and Wehr 1954).
PATHOGENESIS Tissue injury by Amidostomum spp. is caused primarily by mechanical disruption of the gizzard and to a lesser extent by the host inflammatory response. The pathogenesis of infections with species of Epomidiostomum is probably similar but has not been reported. Thirdstage larvae of Amidostomum spp. penetrate the soft area of the mucosa near the junction of the proventriculus or intestine with the gizzard. They develop to fourth-stage larvae with little migration within or beneath the mucosa (Leiby and Olsen 1965). Mature worms migrate and feed on blood in the gizzard mucosa causing hemorrhage, leakage of plasma proteins, and potential ischemia resulting in erosions and ulcers (Bunyea and Creech 1926; Enigk et al. 1969). Parasite activity and hemorrhage in the mucosa beneath the koilin lining and within the koilin disrupt the attachment between the glandular mucosa and koilin, leading to cracks and loss of portions of the gizzard pads. In severe infections, the result may be impaired gizzard function and nutritional deprivation (Bunyea and Creech 1926). In times of nutritional stress, severe infections with Amidostomum spp. may substantially contribute to fatal debilitation (Herman and Wehr 1954; Borgsteede 2005). It has been proposed that the parasite releases toxins that contribute to localized formation of ulcers; to generalized signs of depression, emaciation, dyspnea, and dysphagia; and to inflammatory cell reactions in organs remote from the parasite (Bunyea and Creech 1926; Vet´esi et al. 1976), but this has not been verified by experimental studies.
PATHOLOGY Birds with severe ventricular nematodiasis may be thin to emaciated. Gross lesions associated with ventricular nematodiasis are present in the gizzard, and, even in mild infections, red to brown discoloration may be found at the margins of the koilin lining, between the koilin pads, and in the soft mucosa close to the
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Amidostomum and Epomidiostomum
Figure 20.1. Infection with Amidostomum sp. in the gizzard of a Canada Goose (Branta canadensis). Hemorrhage is present at the margins of the koilin pads, which are cracking and separating from the mucosa. Thin nematodes lie atop focal mucosal hemorrhage between the pads (arrow). (Photograph by J. Runningen, US Geological Survey, National Wildlife Health Center files.) duodenal orifice (Tuggle and Crites 1984). The small threadlike nematodes are buried in the discolored mucosa and/or beneath the koilin. Multiple genera and species of gizzard worms can occur within a single individual bird, and several species may be found occupying different locations in the gizzard (Tuggle and Crites 1984). In Snow Geese, A. anseris was found close to the duodenal orifice and at the periphery of the koilin; A. spatulatum was found in similar locations but also beneath the koilin; and E. crami was found beneath the koilin, partially within the mucosa, and in the gizzard muscle (Tuggle and Crites 1984). In severe gizzard worm infections, the koilin lining is undermined by parasite activity and hemorrhage so that it develops cracked margins, becomes friable, erodes, and separates from the glandular mucosa (Figure 20.1). In severe infections with Epomidiostomum, tracts containing the parasite and its eggs can also be found in the underlying muscle (Figure 20.2) (Tuggle and Crites 1984). Severe lesions were found in Canada Geese with more than 150 A. anseris, in Snow Geese with more than 30 A. anseris and/or A. spatulatum, and in Snow Geese with more than 25 E. crami (Herman and Wehr 1954; Tuggle and Crites 1984). Gross lesions are similar in various species of waterfowl, but in Tundra Swans (Cygnus columbianus) and Trumpeter Swans, severe infections with Amidostomum spp. may be associated with esophageal impaction (McKelvey and MacNeill 1981). Lesions in other avian species have not been described. Reports of Amidostomum and Epomidios-
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Figure 20.2. Species of Epomidiostomum may invade the gizzard muscle beneath the koilin pads and form granulomatous tracts (arrows), as in this Snow Goose (Chen caerulescens). (Photograph by J. Runningen, US Geological Survey, National Wildlife Health Center files.) tomum in avian families other than Anatidae generally have focused on morphological descriptions of the parasites rather than the effects on the hosts. In microscopic sections of gizzard, Amidostomum spp. are commonly visible between the mucosa and the koilin lining, within the deep layers of the koilin, and in the lumina of mucosal glands (Bunyea and Creech 1926; Vet´esi et al. 1976). With time, parasites may spread throughout the koilin layer and become surrounded by eggs (including embryonated eggs). Sections of the parasites are found less frequently in the mucosal propria (although the parasites must feed there), suggesting that the majority of the parasite body resides in more superficial layers. Mucosal glands containing parasites may be dilated and the epithelial cells flattened. The koilin often is separated from the mucosa and fragmented, and mucosal erosions may be present. Hemorrhage fills clefts between the mucosa and koilin lining, within the koilin, and the mucosal propria. Early inflammation consists of diffuse eosinophils in the propria, and later these are mixed with lymphocytes and macrophages (Vet´esi et al. 1976). Lymphoid hyperplasia progresses with time and consists of perivascular infiltrates of lymphocytes and histiocytes as well as lymphoid follicles in the deep mucosal propria; plasma cells are not a feature (Vet´esi et al. 1976). Fibroblasts and fibrosis also increase with time. Similar inflammatory lesions may occur in the proventriculus where larvae may have initially invaded. In infections with Epomidiostomum spp., granulomatous inflammation and granulomas form around the parasites in the ventricular muscularis in addition to presence of parasites and inflammation in the mucosa (Tuggle and Crites
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1984). The parasites in muscle are surrounded by reactive macrophages, hemorrhage, multinucleated giant cells, fibroblasts, and fibrosis; fibrosis may be severe in gizzard muscle (Tuggle and Crites 1984). DIAGNOSIS The diagnosis of ventricular nematodiasis is made by observation of worms in the koilin lining and gizzard muscle or eggs in the feces. Eggs of both genera have similar features and may require trained personnel for identification to genus (Tuggle and Friend 1999). Identification to species requires examination of adult male and female worms and comparison of morphological characteristics found in established keys and species descriptions. A diagnosis of morbidity or mortality should be based on the severity of lesions and complicating factors, as well as elimination of other causes. For example, thickening and irregularity of gizzard pads as well as upper gastrointestinal impaction can be present in lead-poisoned waterfowl. IMMUNITY In studies of A. anseris in domestic geese, precipitating antibodies were not detected in infected goslings (Vet´esi et al. 1976 citing Bausov 1969), and goslings as well as older geese were susceptible to reinfection after initial infection with intact or irradiated larvae (Phuc and Varga 1973; Stradowski 1977 citing Georgiev 1963). In the absence of reinfection, there is evidence that A. anseris has a limited life span in the host, although the mechanism responsible for loss of worms and involvement of host immunity in the process is unknown (Stradowski 1977). Age-related susceptibility has been documented experimentally in domestic geese (Phuc and Varga 1973; Stradowski 1977) and observed in wild geese (Herman and Wehr 1954; Nowicki et al. 1995). The mechanism is uncertain but may be linked to innate or acquired immunity. After experimental inoculations in domestic geese, the development of A. anseris was correlated with the age of the bird, in which the parasites matured more rapidly and had a longer life span in younger geese (Phuc and Varga 1973; Stradowski 1977). The greatest intensity of infection with A. anseris in wild Canada Geese in North Carolina, USA, occurred in younger geese, and young geese were predominantly involved in winter mortality to which Amidostomum was considered to be a contributor (Herman and Wehr 1954). In a later study in Illinois, USA, immature Canada Geese were found to have greater intensity of gizzard worm infections, greater prevalence of A. anseris infection, and a greater proportion of mature A. anseris than adult geese (Nowicki et al. 1995).
There is evidence that A. anseris infection may interfere with development of an immune response to other pathogens. The mechanism is unknown but could be related to stress, nutritional compromise, or protein loss. Antibody production in response to vaccination was compared in domestic geese with and without subclinical gastrointestinal parasite infections (specifically a combination of A. anseris, Capillaria anatis, and Heterakis gallinarum) in which A. anseris was most prevalent (Ziomko et al. 1998). Antibody production was lower and significantly delayed in parasitized geese (Ziomko et al. 1998).
PUBLIC HEALTH CONCERNS There is no evidence that species of Amidostomum and Epomidiostomum present human health concerns. For human consumption, however, it is recommended that gizzards are thoroughly cooked or discarded if nematode-damaged tissues are apparent because of possible secondary bacterial infections (Tuggle and Friend 1999).
DOMESTIC ANIMAL HEALTH CONCERNS Ventricular nematodiasis in domestic anatids has long been recognized throughout the world (Cram 1925; Gower 1939; Levine 1968; McDonald 1969). Instances of heavy losses and severe pathology have been reported among geese infected with A. anseris (Ruff 1984). This species is a concern in commercially raised ducks, geese, and pigeons (Ruff 1984; Permin and Hansen 1998). Chickens experimentally inoculated with A. anseris experienced some hemorrhage and destruction of the koilin, demonstrated acute or subacute inflammation of the mucosa, and had lymphoid hyperplasia, but the worms were subsequently eliminated by this host within 15 days postinoculation (Vet´esi et al. 1976). Additionally, Amidostomum skrjabini (=A. acutum) has produced pathology in young ducks (Ruff 1984). Other species of Amidostomum and species of Epomidiostomum are not reported or emphasized as species of concern for poultry (Ruff 1984; Permin and Hansen 1998; Merck Veterinary Manual 2006). There is potential for transmission of gizzard worms between domestic and wild birds within the Anatidae. Species reported in domestic ducks and geese include A. acutum, A. anseris, and E. uncinatum (McDonald 1969). Consequently, unenclosed zoological gardens, open-air commercial farms, and backyard flocks may be at risk from wild anatids infected with these species of gizzard nematodes.
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Amidostomum and Epomidiostomum WILDLIFE POPULATION IMPACTS AND MANAGEMENT IMPLICATIONS Although numerous studies have demonstrated pathology in individual birds resulting from ventricular nematodiasis (Jerstad 1937; Herman and Wehr 1954; MacNeill 1970; Crichton and Welch 1972; Turner and Threlfall 1975; Harradine 1982; Tuggle and Crites 1984), there presently is no evidence that robust wild populations of anatids are negatively impacted. However, mortality in Canada Goose goslings has been associated with ventricular nematodiasis (Wickware 1941; Herman and Wehr 1954). This could affect recruitment and possibly have a negative population effect if direct or indirect mortalities resulting from infections are additive. Additionally, this may be a problem if Canada Geese are being managed at the subspecies level, particularly if they are differentially impacted by gizzard worms because of host geographic-isolating mechanisms that permit differences in exposure probabilities among host subspecies. Concerns about introducing diseases and parasites have been raised in regard to releasing captive-raised infected birds to supplement or reestablish wild populations of threatened or endangered species (Bailey and Black 1995). However, few specific examples are available regarding negative effects on small host populations caused by species of Amidostomum and Epomidiostomum, and further study seems warranted. Infections are relatively uncommon in host families not associated with wetland habitats. However, the report of Eurasian Collared-Doves (Streptopelia decaocto) becoming infected with A. anseris at a goose farm (Ryˇsav´y et al. 1955) suggests potential routes of entry into wild bird populations in situations where they are exposed to high concentrations of infective larvae originating from domestic anatids. Additional research is needed to determine whether local population impacts occur in these alternate hosts.
TREATMENT AND CONTROL No methods have been developed for treatment and control of species of Amidostomum and Epomidiostomum in wild host populations, presumably because there is little evidence that ventricular nematodiasis is an important cause of epizootic die-offs. Treatment and control of ventricular nematodiasis are appropriate for captive flocks of wild species, threatened or endangered species, and where wild and domesticated waterfowl intermingle in commercial farming operations. Where applicable, providing larger feeding areas for captive herbivorous grazing species such as the Hawaiian Goose (Branta sandvicensis) may decrease infections of density-dependent
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disease agents (Bailey and Black 1995), which would be applicable to gizzard worms. Other strategies used for domestic fowl may be applicable to captive wild anatids, including raising young and old birds separately to prevent exposure of uninfected young individuals to infected older individuals and maintaining good sanitation practices (Levine 1968). Where possible, efforts should be made to prevent access of wild anatids to captive or domesticated flocks, particularly if anthelmintics are being used, because reinfection can rapidly occur following treatment. Anthelmintics have been used effectively on infected captive wild anatids and commercial flocks, but in these instances, treatments were focused on eliminating A. anseris because of its economic importance to commercial enterprises. Specific anthelmintics used to kill A. anseris include flubendazole (Vanparijs 1984), mebendazole (Ruff 1984 citing Enigk et al. 1973; Bailey et al. 1990), cintrin (Merck Veterinary Manual 2006), cambendazole (Ruff 1984 citing Enigk and Dey-Hazra 1971), pyrantel (Ruff 1984), and a combination of neguvon, atropine sulfate, and piperazine sulfate (Georgiev 1968).
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Slo´nsk Nature Reserve. Biological Bulletin of Pozna´n 37:267–271. Baruˇs, V., M. D. Sonin, F. Tenora, and R. Wiger. 1984. Survey of nematodes parasitizing the genus Tetrao (Galliformes) in the Palaearctic region. Helminthologia 21:3–15. Bausov, I. A. 1969. Immunity in amidostomiasis (in Russian). Uchenye Zapiski Kurskogo Gosudarstvennogo Pedagogicheskogo Instituta 59:133–142. Baylis, H. A. 1932. A comparison of certain species of the nematode genus Amidostomum, with a description of a new species. Annals and Magazine of Natural History 10:281–286. ˇ Birov´a, V., M. Spakulov´ a, and J. K. Macko. 1990. Seasonal dynamics of the invasive cycle of nematodes and acanthocephalans in the wild (Anas platyrhynchos L.) and domestic duck (Anas platyrhynchos f. dom.). Helminthologia 27:291–301. Bishop, C. A., and W. Threlfall. 1974. Helminth parasites of the common eider duck, Somateria mollissima (L.), in Newfoundland and Labrador. Proceedings of the Helminthological Society of Washington 41:25–35. Bolte, A. L., W. Lutz, and E. F. Kaleta. 2000. Untersuchungen zum Vorkommen von Infektionserregern bei freilebenden Graug¨ansen (Anser anser Linn´e, 1758) (English summary). Zeitschrift f¨ur Jagdwissenschaft 46:176–179. Borgsteede, F. H. M. 2005. The gizzard worm, Amidostomum acutum (Lundahl, 1848) Seurat, 1918 in common eiders (Somateria mollissima L.) in the Netherlands. Helminthologia 42:215–218. Borgsteede, F. H. M., K. M. Kavetska, and P. E. F. Zoun. 2006. Species of the nematode genus Amidostomum Railliet and Henry, 1909 in aquatic birds in the Netherlands. Helminthologia 43:98–102. Bourgeois, C. E., and W. Threlfall. 1982. Metazoan parasites of three species of scoter (Anatidae). Canadian Journal of Zoology 60:2253–2257. Broderson, D., A. G. Canaris, and J. R. Bristol. 1977. Parasites of waterfowl from southwest Texas: II. The shoveler, Anas clypeata. Journal of Wildlife Diseases 13:435–439. Bunyea, H., and G. T. Creech. 1926. The pathological significance of gizzard-worm disease of geese. North American Veterinarian 7:47–48. Buscher, H. N. 1965. Dynamics of the intestinal helminth fauna in three species of ducks. Journal of Wildlife Management 29:772–781. Canaris, A. G., A. C. Mena, and J. R. Bristol. 1981. Parasites of waterfowl from southwest Texas: III. The green-winged teal, Anas crecca. Journal of Wildlife Diseases 17:57–64.
Clinchy, M., and I. K. Barker. 1994. Dynamics of parasitic infections at four sites within lesser snow geese (Chen caerulescens caerulescens) from the breeding colony at La P´erouse Bay, Manitoba, Canada. Journal of Parasitology 80:663–666. Conti, J. A., D. J. Forrester, and S. A. Nesbitt. 1985. Parasites of limpkins, Aramus guarauna, in Florida. Proceedings of the Helminthological Society of Washington 52:140–142. Cowan, A. B. 1955. Some preliminary observations on the life history of Amidostomum anseris Zeder, 1800. Journal of Parasitology 41(Suppl):43–44. Cram, E. B. 1925. New records of economically important nematodes in birds. Journal of Parasitology 12:113–114. Cram, E. B. 1927. Bird Parasites of the Nematode Suborders Strongylata, Ascaridata, and Spirurata. Smithsonian Institution, U.S. National Museum Bulletin 140. U.S. Government Printing Office, Washington, DC. Crichton, V. F. J., and H. E. Welch. 1972. Helminths from the digestive tracts of mallards and pintails in the Delta Marsh, Manitoba. Canadian Journal of Zoology 50:633–637. Czapli´nski, B. 1962. Nematodes and acanthocephalans of domestic and wild Anseriformes in Poland. I. Revision of the genus Amidostomum Railliet et Henry, 1909. Acta Parasitologica Polonica 10:125–164. Dubey, J. P., and B. P. Pande. 1964. A note on some helminths of the wild duck (Anas poecilorhyncha). Indian Journal of Helminthology 16:27–32. Enigk, K., and A. Dey-Hazra. 1968a. On the host specificity of Amidostomum anseris Strongyloidea Nematoda. Zeitschrift f¨ur Parasitenkunde 31:266–275. Enigk, K., and A. Dey-Hazra. 1968b. Die perkutane infektion bei Amidostomum anseris (Strongyloidea, Nematoda). Zeitschrift f¨ur Parasitenkunde 31:155–165. Enigk, K., and A. Dey-Hazra. 1969. Zum verhalten der exogen entwicklungformen von Amidostomum anseris (Strongyloidea, Nematoda). Archiv f¨ur Gefl¨ugelkunde 33:259–273. Enigk, K., and A. Dey-Hazra. 1970. The behaviour of exogenous stages of Amidostomum anseris (Strongyloidea, Nematoda). In H. D. Srivastava Commemoration Volume, K. S. Singh and B. K. Tandan (eds). Indian Veterinary Research Institute, Izatnagar, India, pp. 603–619. Enigk, K., and A. Dey-Hazra. 1971. Zur behandlung der h¨aufigsten nematodeninfektionen des hausgefl¨ugels. Deutsche Tier¨arztliche Wochenschrift 78:178–181.
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Purvis, J. R., D. E. Gawlik, N. O. Dronen, and N. J. Silvy. 1997. Helminths of wintering geese in Texas. Journal of Wildlife Diseases 33:660–663. Roudabush, R. L. 1942. Parasites of the American coot (Fulica americana) in central Iowa. Iowa State College Journal of Science 16:437–441. Ruff, M. D. 1984. Nematodes and acanthocephalans. In Diseases of Poultry, 8th ed., M. S. Hofstad, H. J. Barnes, B. W. Calnek, W. M. Reid, and H. W. Yoder, Jr. (eds). Iowa State University Press, Ames, IA, pp. 614–648. Ryˇsav´y, B., J. Mich´alek, and V. Fidler. 1955. Zur Frage der M¨oglichkeit einer adaption des in G¨ansen parasitierenden wurmes Amidostomum anseris (Zeder, 1800) Railliet und Henry, 1909 an Wirtsv¨ogel anderer Ordnungen (in Russian, German summary). Folia Biologica 1:276–281. Schmidt, G. D., and R. E. Kuntz. 1972. Nematode parasites of Oceanica XVII. Schistorophidae, Spiruridae, Physalopteridae and Trichostrongylidae of birds. Parasitology 64:269–278. Shaw, M. G., and A. A. Kocan. 1980. Helminth fauna of waterfowl in central Oklahoma. Journal of Wildlife Diseases 16:59–64. Shen, S. 1981. On new species of nematodes parasitic in birds from Taihu Region, Jiangsu Province, China (in Chinese, English summary). Acta Zootaxonomica Sinica 6:359–364. Shen, S., and S. Wu. 1964. A preliminary survey of trematodes and nematodes parasitic in aquatic birds from Ulasu Hai, Inner Mongolia, China (in Chinese, English summary). Acta Zoologica Sinica 16:398–415. Shen, S., and S. Wu. 1973. A survey on the helminths of birds from Bai-Yang-Dien Lake, Hopeh Province, China. I. Nematodes (in Chinese). Acta Zoologica Sinica 19:26–34. Stradowski, M. 1971. The age of Amidostomum anseris (Zeder, 1800) Railliet et Henry, 1909 larvae and their invading activity. Acta Parasitologica Polonica 19:63–68. Stradowski, M. 1975. Wintering of Amidostomum anseris (Zeder, 1800) eggs under climatic conditions in central Poland. Acta Parasitologica Polonica 23:373–379. Stradowski, M. 1977. Duration of prepatent, patent and postpatent periods of the Amidostomum anseris (Zeder, 1800) infection in domestic geese. Acta Parasitologica Polonica 24:249–258. Threlfall, W. 1970. A preliminary check list of the helminth parasites of the common snipe, Capella gallinago (Linnaeus). The American Midland Naturalist 84:13–19. Thul, J. E., D. J. Forrester, and C. L. Abercrombie. 1985. Ecology of parasitic helminths of wood ducks, Aix
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21 Tetrameridosis John M. Kinsella and Donald J. Forrester The first species of Microtetrameres was described by Travassos (1914) from a Yellow-fronted Woodpecker (Melanerpes flavifrons) in Brazil, and the first species of Geopetitia from a Coal Tit (Periparus ater) in France (Chabaud 1951). The earliest description of lesions caused by a species of Tetrameres appears to be that of Rust (1908) in Germany. Ellis (1970) published the first account of the pathogenicity of a species of Microtetrameres. It was not until comparatively recently that the first reports on disease due to Geopetitia have appeared (Bartlett et al. 1984; Tscherner et al. 1997).
INTRODUCTION The term tetrameridosis includes diseases caused by species of nematodes belonging to three genera of the family Tetrameridae: Tetrameres, Microtetrameres, and Geopetitia. Females of Tetrameres and Microtetrameres are typically found embedded in the gastric glands of the proventriculus. One or more of the much smaller males may be either associated with the females in the glands or free in the lumen of the proventriculus. A few species of Tetrameres (e.g., Tetrameres strigiphila) are found encysted in a fibrous capsule in the tunica muscularis of the proventriculus, with the female tail extending through an opening into the lumen of the proventriculus for the deposition of eggs. Species of Geopetitia occupy similar cysts in the wall of the esophagus, proventriculus, or the gizzard, but often have multiple females per cyst rather than one (Anderson 2000). Infections of high intensity with species of the genus Tetrameres have led to emaciation, anemia, and death in domestic ducks and chickens. There is some evidence (Ellis 1970) that the damage done to the avian proventriculus by Microtetrameres spp. is similar to that of Tetrameres. One species of Geopetitia has caused widespread morbidity and mortality in zoo birds, but no similar reports are available for wild populations.
HOST RANGE AND DISTRIBUTION The distribution of species of Tetrameres is cosmopolitan. The majority of the species parasitize aquatic birds, especially Anseriformes, Ardeiformes, Gruiformes, and Charadriiformes, although some species are found in land birds such as Passeriformes and occasionally Galliformes. In contrast to species of Tetrameres, species of Microtetrameres usually parasitize terrestrial birds, with the vast majority of species found in Passeriformes, Accipitriformes, and Strigiformes. The exceptions prove the rule in this case, since two of the species found in Ardeiformes, Microtetrameres spiralis and Microtetrameres egretes, parasitize the Cattle Egret (Bubulcus ibis), which feeds principally on land. The distribution of species of Microtetrameres is also cosmopolitan. Eight species of Geopetitia are known; seven occur in wild Falconiformes, Cuculiformes, Coraciiformes, Piciformes, or Passeriformes in France, the former Soviet Union, Taiwan, Australia, India, Madagascar, Ghana, Congo, and Cuba. The natural host of the eighth species, Geopetitia aspiculata, remains unknown. It was originally described from the “purple sugarbird” (Coerulea coerulea) at the National Zoological Park, Washington, DC, USA. (Webster 1971). Unfortunately, purple sugarbird is not a recognized common
HISTORY The first species of Tetrameres was described by Diesing (1835) from raptors in Brazil. The extreme sexual dimorphism led to much early confusion and, in one case, the globular female was thought to be a trematode, not a nematode. It was some time before the males of these species were recognized and associated with the females. Even today it is not always possible to pair males and females with assurance, since mixed infections of different species are not uncommon. This confusion has led to some problems with taxonomy, many of which are still not resolved.
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name, and Coerulea is not listed among world bird species either as a genus or as a species (Clements 2000), so the type host of G. aspiculata remains a mystery. A good possibility is the Purple Honeycreeper (Cyanerpes caeruleus), since honeycreepers have sometimes been called sugarbirds. It has since been recorded from a variety of Passeriformes, Coraciiformes, and Charadriiformes at the Assiniboine Park Zoo, Winnipeg, Canada (Bartlett et al. 1984), the Lincoln Park Zoological Gardens, Chicago, Illinois, USA (French et al. 1994), and four different zoos in Austria and Germany (Tscherner et al. 1997; Juncker-Voss et al. 2001). ETIOLOGY Tetramerid nematodes are characterized by marked sexual dimorphism. Males have a typical nematode filiform appearance, but females are swollen by the greatly distended uterus and may be globular (Tetrameres spp.) (Figure 21.1a), spirally coiled (Microtetrameres spp.) (Figure 21.1b), or have the posterior extremity more slender than the remainder of the body, with the distal end inflated (Geopetitia spp.) (Figure 21.1c) (Yamaguti 1961). The taxonomy of the genus Tetrameres remains in a state of confusion, and it is difficult to estimate the number of valid species described since Yamaguti (1961) listed a total of 46. In an unpublished doctoral dissertation, Mollhagen (1976) provided an extensive host list and proposed 5 new species and a number of synonymies. However, much of his dissertation remains unpublished in peer-reviewed journals and, as a result, his conclusions are not recognized as valid under the rules of the International Code of Zoological Nomenclature. Nevertheless, his dissertation is an invaluable resource for identification of species of Tetrameres, and the extensive bibliography covering the years 1835–1985 has been published separately (Mollhagen 1991). Although some authors (e.g., Chabaud 1975) have considered Microtetrameres to be a subgenus of Tetrameres, Anderson (2000) believed that differences in larval and adult morphology justified its retention as a separate genus. Ellis (1971) listed host and geographical distribution for 35 species of Microtetrameres and gave measurements for those species described from the Western Hemisphere. Mawson (1977) described 15 new species of Microtetrameres from Australian birds and presented a key to the males of 52 species. Seven additional species have been described since 1977, bringing the total known species to 59. There is currently no review of the taxonomy of the genus Geopetitia available. Webster (1971) listed
Figure 21.1. Line drawings of representatives of the three genera of Tetrameridae. (a) Adult male (left) and female (right) of Tetrameres strigiphila from a Barred Owl (Strix varia). Courtesy of D. B. Pence and the Journal of Parasitology. (b) Adult female of Microtetrameres aquila from a golden eagle (Aquila chrysaetos). Courtesy of S. C. Schell and Transactions of the American Microscopical Society. (c) Posterior extremity of an adult female Geopetitia aspiculata from a Purple Sugarbird (Coerulea coerulea). Courtesy of W. A. Webster and the Proceedings of the Helminthological Society of Washington.
eight species, but considered three of these doubtful because they had been described from females only. In light of the notable lack of host specificity of G. aspiculata, the validity of existing species needs to be reexamined.
EPIZOOTIOLOGY Life History Information on life cycles of seven species of Tetrameres was summarized by Anderson (2000). The majority of species of this genus parasitize aquatic
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birds such as anseriforms and ciconiiforms and intermediate hosts are usually crustaceans. A smaller number of species parasitize terrestrial birds, especially passeriforms and galliforms, and intermediate hosts are normally insects, most commonly orthopterans and coleopterans. Kovalenko (1960) implicated fish as paratenic hosts of Tetrameres fissispina, but the role of carrier hosts in the genus remains largely undefined. The following are examples of typical aquatic and terrestrial life cycles. Larvae of T. fissispina, a common parasite in anseriforms, reach the third or infective stage in the intermediate host, Gammarus lacustris, in 8–18 days (Garkavi 1949). In ducklings, males and females occur together in glands at 10 days postinfection, but by 18 days only females are found, indicating that insemination of the females occurs early in infections, followed by death of the males (Cvetaeva 1960). The intermediate host of Tetrameres cardinalis, a parasite of the Northern Cardinal (Cardinalis cardinalis), was determined to be an orthopteran, Locusta migratoria. Infective third-stage larvae are encapsulated in the fat body at 11 days (Quentin and Barre 1976), and when fed to a cardinal developed to the fourth stage by 11 days postinfection. The life history of Microtetrameres corax, a parasite of the Black-billed Magpie (Pica hudsonia) in Colorado, was described by Bethel (1973). Eggs were fed to grasshoppers (Melanopus spp.), and third-stage larvae were found in the thoracic region of the hemocoel and among the fat bodies of the abdominal cavity as early as 27 days postinfection. Grasshoppers were fed to laboratory-born magpies and adult males and females of M. corax were recovered 48 days later. Quentin et al. (1986) infected Tylotropidus patagiatus and Locusta migratoria (Orthoptera: Acrididae) with eggs of Microtetrameres inermis from Orange Weavers (Ploceus aurantius). Third-stage larvae were found free in the hemocoel 18 days later. The life cycle of G. aspiculata has been studied only in zoos and the natural definitive host remains unknown. Bartlett et al. (1984) placed gravid female worms on apple peels and fed them to crickets (Acheta domestica). Infective third-stage larvae were found by 48 days, and when administered orally to a Cut-throat (Amadina fasciata), two adults were found in a cyst on the proventricular serosa 35 days later. French et al. (1994) infected crickets (Acheta domestica) and co*ckroaches (Blattella germanica and Supella supellectilium) with eggs from G. aspiculata, and obtained infective larvae at 35 days. Larvae were found at the junction of the proventriculus and the ventriculus 24–48 h postinfection in Zebra Finches (Taeniopygia guttata), and raised nipplelike nodules on the serosal surface of the proventriculus were
observed at 2 weeks. A mass of mature worms was found in a proventricular nodule at 14 weeks. Prevalence and Intensity As with many nematode infections, prevalences and intensities of Tetrameres spp. tend to be higher in confined domestic birds than in wild populations. For example, Czaplinski (1962) reported a 46% prevalence of T. fissispina in domestic Mallards (Anas platyrhynchos) in Poland, but only a 27% prevalence in wild Mallards. In East Slovakia, T. fissispina occurred at higher prevalences in wild mallards, but higher intensities in domestic Mallards (Birova et al. 1990). Kinsella (1973) found 66% of American Coots (Fulica americana) infected with Tetrameres globosa in Florida, while only 25% of Purple Gallinules (Porphyrio martinica) and 6% of Common Moorhens (Gallinula chloropus) from the same area were infected with the same species (Kinsella et al. 1973), presumably reflecting differences in feeding preferences for intermediate hosts. Mollhagen (1976) states that intensities of infections of Tetrameres spp. in wild birds usually average fewer than ten worms per bird, although intensities in the hundreds have been reported occasionally. Only a few studies of seasonal and long-term trends in Tetrameres have been done in wild birds. In a study of Northern Bobwhites (Colinus virginianus) in 1968– 1969 in northern Florida, Davidson et al. (1980) found more than 70% of juveniles and adult birds infected with Tetrameres pattersoni, with intensities as high as 65 per bird. A summer peak of relative abundance was found, coinciding with an abundance of arthropod intermediate hosts. But in a 1983–1984 survey of the same population, Moore et al. (1986) found only 5% of adults and no juveniles infected with T. pattersoni, possibly due to the decline of these intermediate hosts. In a 5-year study of Northern Bobwhites in southern Florida, Forrester et al. (1984) reported T. pattersoni to be totally absent in 1 year. From 1976 to 1983, 25 birds of 14 species (Passeriformes: 12, Coraciiformes: 1, Charadriiformes: 1) that died at the Assiniboine Park Zoo in Winnipeg were found infected with G. aspiculata. Intensities could not be exactly determined but were estimated to range from 6 to 50 (Bartlett et al. 1984). At the Lincoln Park Zoological Gardens in Chicago, 96 birds of 25 species (Passeriformes: 19, Piciformes: 5, Ciconiiformes: 1), and at three zoos in Germany, 77 birds of 41 species (Passeriformes: 32, Piciformes: 5, Columbiformes: 2, Coraciiformes: 1, Charadriiformes: 1) were infected with G. aspiculata (French et al. 1994; Tscherner et al. 1997) but prevalences and intensities were not recorded. Between 1993 and 1998, G. aspiculata was reported as the cause of death of
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Tetrameridosis 44 birds at the Viennese Zoo aviary (Juncker-Voss et al. 2001). CLINICAL SIGNS Clinical signs of infections with Tetrameres spp. in pigeons, ducks, and chickens include weakness, loss of appetite, diarrhea, and emaciation with atrophy of the thoracic musculature (Wehr 1971; Endo and Inoue 1989). Fink et al. (2004) infected three groups of chickens with doses of 25, 100, and 400 larvae of Tetrameres americana, respectively, and found that infected chickens exhibited no difference in weight gain compared to controls. Higher doses influenced the establishment rate and intensity of infection, but not worm size, or host mortality. There is virtually no information on clinical signs of disease in wild birds due to species of the genus Microtetrameres. Bethel (1973) infected two laboratoryborn Black-billed Magpies with 110 and 74 females of M. corax and found experimental birds less aggressive and less active than controls at 35 days postinfection. Many zoo birds infected with G. aspiculata showed no specific signs other than sudden death. Others had a history of weight loss and abdominal distension (Bartlett et al. 1984; French et al. 1994). PATHOLOGY Clinical Pathology Hematological changes were studied in 12 chicks experimentally infected with Tetrameres mohtedai by Ramaswamy and Sundaram (1983). A marked eosinophilia was found during the early stages of infection when juveniles migrated through the wall of the proventriculus, followed by heterophilia and lymphocytopenia when the nematodes developed in the proventricular glands. A normocytic and normochromic anemia in early stages of development progressed to a macrocytic, hypochromic anemia as the worms matured. We know of no information on clinical pathology of birds infected with species of Microtetrameres or Geopetitia. Gross Pathology Infections of high intensity with Tetrameres spp. are characterized by enlargement of the proventriculus, with thickening of the wall. The females within the glands can be seen through the wall as dark red, slightly raised foci 2–3 mm in diameter, resembling hematomas. The infected proventricular glands may be covered with a foamy, necrotic material. The small
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intestinal mucosa may be swollen and covered with mucus and the contents greenish due to the presence of bile, or granular and golden-brown due to the presence of red blood cells (Popova 1954). No information is available on gross pathology due to Microtetrameres spp. Cysts of G. aspiculata range in size from small plaques on the serosal surface of the esophagus, proventriculus, or gizzard to large cysts containing up to 50 worms and filling the whole abdominal cavity. Adhesions may form between the cysts and the serosa of the liver or spleen causing inflammation and fibrosis (Bartlett et al. 1984; French et al. 1994). Younger cysts are delicate and membranous and the worms can be seen through the wall while older cysts become hard, friable masses. Histopathology Bergan et al. (1994) described lesions caused by Tetrameres striata in Mallards. In most cases, the gravid females caused pressure atrophy and necrosis of the proventricular gland mucosa, with complete loss of acini, but little or no inflammatory response around the parasites or in the compacted mucosa or submucosa (Figure 21.2). Occasional lesions were noted in the submucosa surrounded by a thin layer of fibrous material forming a cyst, with or without adjacent inflammatory cells. A similar pattern of pressure atrophy and loss of acini was reported by Mascarenhas and Ghosh (1992) in proventricular lesions caused by T. mohtedai in fowl, but gravid females were also found deep within the muscular layer of the gizzard. These females were found to cause atrophy of the adjacent muscle fibers and severe cellular reaction characterized by infiltration of lymphocytes, plasma cells, eosinophils, and macrophages, and proliferation of fibrovascular granulation tissue. Ellis (1970) conducted an in situ study of sections of adult female Microtetrameres centuri in Eastern and Western Meadowlarks (Sturnella magna and Sturnella neglecta) and demonstrated nucleated red blood cells within the lumen of the intestine, indicating this species to be hematophagous. Mechanical growth of the females within the proventricular glands caused dilatation of the gland lobule with mild pressure atrophy of the epithelium. Involved gland lobules were nonfunctional and mild hyperemia of the lamina propria was noted. There was no marked inflammatory response, but increased mucus secretion and a mild proventriculitis were observed. Bethel (1973) noted a similar pattern in natural infections of M. corax in Black-billed Magpies, with pressure atrophy of the glandular tissue around the worms but no connective tissue or cellular response. Loss of secretory function in infected glands
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Figure 21.2. Section of a proventriculus of a Mallard (Anas platyrhynchos) infected with an adult male (M) and female (F) Tetrameres striata. Note the lack of an inflammatory response. Bar = 500 μm. Courtesy of D. B. Pence and the Journal of Wildlife Diseases.
was also reported in Microtetrameres nestoris infections of the New Zealand Kaka (Nestor meridionalis) (Clark et al. 1979). Within the cyst, adults of G. aspiculata are surrounded by fibrinous exudate or granulation tissue containing scattered fibroblasts, giant cells, heterophils, and lymphocytes. Both live and dead worms may be surrounded by erythrocytes, degranulated heterophils, and giant cells, and thick-shelled, larvated nematode eggs may be seen both free in the cysts and within female worms (Bartlett et al. 1984). In areas penetrated by the posterior end of the worms (Figure 21.3), the wall of the proventriculus may lose its structural integrity, and fibrinous exudate, scattered heterophils, and sloughed proventricular gland cells may be present in the proventricular lumen. Mucosal damage may render the host susceptible to secondary bacterial and fungal invasion (French et al. 1994).
DIAGNOSIS Diagnosis of tetramerid infections in a living bird by examination of fecal samples is difficult and usually
Figure 21.3. Section of the proventriculus of a White-lined Tanager (Tachyphonus rufus) showing a specimen of Geopetitia aspiculata protruding from a cyst through the proventricular wall. Bar = 500 μm. Courtesy of C. M. Bartlett and the Journal of Wildlife Diseases.
not recommended. Eggs are oval, thick-shelled, larvated, and, in some species, polar tufts of filaments are present. It would be difficult to distinguish them from other common spirurid eggs found in bird feces such as Dispharynx nasuta. Tscherner et al. (1997) used endoscopic examination of some birds to confirm the characteristic cysts of Geopetitia on the wall of the proventriculus. At necropsy, lesions of Tetrameres and Microtetrameres can often be seen as dark red, slightly raised foci (which contain the female worms, swollen with ingested blood) on the serosal wall of the unopened proventriculus. Compression of the fundic glands without dissection will often express the females intact. Care should be taken to search for the smaller, colorless male worms since identification of species is primarily based on the males. Females of Tetrameres are globular and divided into four sectors by longitudinal grooves while females of Microtetrameres are twisted into two or three tight coils. Males of Tetrameres often have two to four rows of spines on the body, although
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Tetrameridosis one subgenus, Gynaecophila, is spineless. Males of Microtetrameres are always spineless. Both sexes of the adults of Geopetitia may be coiled as in Microtetrameres spp., but the tail of female Geopetitia spp. are drawn out into a posterior inflation separated by a constriction. IMMUNITY No experimental studies have been conducted on immunity to infections with this family of nematodes. Neither Davidson et al. (1980) nor Forrester et al. (1984) found any significant differences in prevalences or intensities of infection of T. pattersoni between juvenile and adult Northern Bobwhites. However, in a study of free-ranging chickens in Tanzania, Fink et al. (2005) found that prevalences of T. americana were significantly higher in chicks than in growers or adults, although the mean intensity did not differ significantly. Whether this difference was related to age-specific immunity remains to be proven. PUBLIC HEALTH CONCERNS There is no evidence that these nematodes infect humans. DOMESTIC ANIMAL HEALTH CONCERNS Many species of Tetrameres are shared between wild and domestic birds, including T. fissispina and Tetrameres crami in wild and domestic Mallards (Czaplinski 1962; McDonald 1969; Birova et al. 1990), and Tetrameres zakharowi in wild and domestic geese (McDonald 1969; Mollhagen 1976). Both wild and domestic birds may then act as sources for the infection of intermediate hosts. For example, Hudina and Pavlovic (1989) reported “a great number of deaths” in flocks of carrier pigeons (Rock Pigeons, Columba livia) attributed to infections of T. fissispina acquired en route during a competition, presumably from feeding on intermediate hosts in the wild. WILDLIFE POPULATION IMPACTS Data are lacking on the impact of most members of the Tetrameridae on wild populations of birds. An exception might be T. fissispina, which has been listed as a cause of morbidity and mortality in waterfowl (McDonald 1969), although this needs further study. As with other parasitic infections, factors such as crowding and change in availability of food supplies can increase stress on the host and potentiate the effect of infections. Crowding can also increase transmission and lead to infections of higher intensities.
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TREATMENT AND CONTROL Several treatment regimes have been used to treat tetramerid infections in captive birds. These include a single oral dose of Levamisole to control T. americana in Rock Pigeons (Panigrahy et al. 1982). A subcutaneous dose of ivermectin or a combination of Panacur and ivermectin given intraperitoneally has been used to control infections of G. aspiculata in zoo birds (Tscherner et al. 1997). To prevent transmission of G. aspiculata at the Assiniboine Park Zoo, feeding of farm-raised crickets was stopped and efforts were made to reduce the numbers of feral insects, resulting in a decrease from nine birds infected per year in 1981 to three in 1983 (Bartlett et al. 1984). Similarly, control of co*ckroaches at the Viennese Zoo aviary significantly reduced mortalities due to G. aspiculata (Juncker-Voss et al. 2001). We are not aware of any studies on treatment of infections of species of Microtetrameres, but the abovementioned drugs might be effective. MANAGEMENT IMPLICATIONS Although there are no reports of deaths in birds in the wild attributed to species of Geopetitia, the demonstrated epizootic and pathogenic potential of G. aspiculata in zoo environments warrants further study. The natural hosts of G. aspiculata are assumed to be tropical birds of the order Passeriformes, since they were the host group most often infected in zoos. Because of the well-documented lack of host specificity of G. aspiculata, there is legitimate concern that it could spread to the local avifauna in both Europe and North America if infected birds are kept in outdoor flight cages (Bartlett et al. 1984). LITERATURE CITED Anderson, R. C. 2000. Nematode Parasites of Vertebrates: Their Development and Transmission, 2nd ed. Commonwealth Agricultural Bureaux, Farnham Royal, Buckinghamshire, UK. Bartlett, C. M., G. J. Crawshaw, and R. G. Appy. 1984. Epizootiology, development, and pathology of Geopetitia aspiculata Webster, 1971 (Nematoda: Habronematoidea) in tropical birds at the Assiniboine Park Zoo, Winnipeg, Canada. Journal of Wildlife Diseases 20:289–299. Bergan, J. F., A. A. Radomski, D. B. Pence, and O. E. Rhodes, Jr. 1994. Tetrameres (Petrowimeres) striata in ducks. Journal of Wildlife Diseases 30:351–358. Bethel, W. M. 1973. The life cycle and notes on the developmental stages of M. corax Schell, 1953 (Nematoda: Tetrameridae). Proceedings of the Helminthological Society of Washington 40:22–26.
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Birova, B., M. Spakulova, and J. K. Macko. 1990. Seasonal dynamics of the invasive cycle of nematodes and acanthocephalans in the wild (Anas platyrhynchos L.) and domestic duck (Anas platyrhynchos f. dom.). Helminthologia 27:291–301. Chabaud, A. G. 1951. Description d’un nematode parasite d’m´esange, Geopetitia pari n. g., n. sp., interm´ediaire entre Tetrameridae et Crassicaudidae et hypoth`eses sur l’interpr´etation phylog´en´etique des helminthes de ce groupe. Annales de Parasitologie Humaine et Compar´ee 26:190–200. Chabaud, A. G. 1975. Keys to genera of the order Spirurida. Part 2. Spiruroidea, Habronematoidea and Acuarioidea. In CIH Keys to the Nematode Parasites of Vertebrates No. 3, R. C. Anderson, A. G. Chabaud, and S. Willmott (eds). Commonwealth Agricultural Bureaux, Farnham Royal, Buckinghamshire, UK, pp. 29–58. Clark, W. C., H. Black, and D. M. Rutherford. 1979. M. nestoris new species (Nematoda, Spirurida), a parasite of the North Island New Zealand kaka, Nestor meridionalis septentrionalis (Aves, Psittaciformes). New Zealand Journal of Zoology 6:1–6. Clements, J. F. 2000. Birds of the World: A Checklist, 5th ed. Ibis Publishing, Vista, CA. Cvetaeva, N. P. 1960. [Pathom*orphological changes in the proventriculus of ducks by tetrameriasis] (in Russian). Helminthologia 2:143–150. Czaplinski, B. 1962. Nematodes and acanthocephalans of domestic and wild Anseriformes in Poland. III. General comment. Acta Parasitologica Polonica 10:277–319. Davidson, W. R., F. E. Kellogg, and G. L. Doster. 1980. Seasonal trends of helminth parasites of bobwhite quail. Journal of Wildlife Diseases 16:367–375. Diesing, K. M. 1835. Tropisurus und Thysanosoma, zwey neue Gattungen von Bunnenw¨urmen (Entozoen) aus Brasilien. Medicinische Jahrb¨ucher ¨ Oesterreichischen Staates 7:83–116. Ellis, C. J. 1970. Pathogenicity of Microtetrameres centuri Barus, 1966 (Nematoda: Tetrameridae) in meadowlark. Journal of Nematology 2:33–35. Ellis, C. J. 1971. Comparative measurements and host and geographical distribution of species of Microtetrameres (Nematoda: Tetrameridae). Iowa State Journal of Science 46:29–47. Endo, M., and I. Inoue. 1989. Observations on tetrameriasis in race pigeons, Columba livia var. domestica. Bulletin of the College of Agriculture and Veterinary Medicine of Nihon University 46: 83–88. Fink, M., A. Permin, H. B. Magwisha, and K. M. V. Jensen. 2004. Tetrameres americana Cram (1927) populations in chickens infected with different dose levels. Veterinary Parasitology 124:239–247.
Fink, M., A. Permin, H. B. Magwisha, and K. M. V. Jensen. 2005. Prevalence of the proventricular nematode Tetrameres americana Cram (1927) in different age groups of chickens in the Morogoro Region, Tanzania. Tropical Animal Health and Production 37:133–137. Forrester, D. J., J. A. Conti, A. O. Bush, L. D. Campbell, and R. K. Frohlich. 1984. Ecology of helminth parasitism of bobwhites in Florida. Proceedings of the Helminthological Society of Washington 51:255–260. French, R. A., K. S. Todd, T. P. Meehan, and J. F. Zachary. 1994. Parasitology and pathogenesis of Geopetitia aspiculata (Nematoda: Spirurida) in zebra finches (Taeniopygia guttata): Experimental infection and new host records. Journal of Zoo and Wildlife Medicine 25:403–422. Garkavi, B. L. 1949. [Elucidation of cycle of development of the nematode Tetrameres fissispina, parasite of domestic and wild ducks] (in Russian). Doklady Academii Nauk SSSR 66:1215–1218. Hudina, V., and I. Pavlovic. 1989. Finding of the nematode Tetrameres fissispina in carrier pigeons and its control. Veterinarski Glasnik 43:1063–1066. Juncker-Voss, M. A. Kubber-Heiss, E. Kutzer, P. Kanzler, D. Schratter, and H. Prosl. 2001. Geopetitia aspiculata: autochtones Vorkommen bei Zoovogeln in Osterreich. Wiener Tierarztliche Monatsschrift 88:106–113. Kinsella, J. M. 1973. Helminth parasites of the American coot, Fulica americana americana, on its winter range in Florida. Proceedings of the Helminthological Society of Washington 40:240–242. Kinsella, J. M., L. T. Hon, and P. B. Reed, Jr. 1973. A comparison of the helminth parasites of the common gallinule (Gallinula chloropus cachinnans) and the purple gallinule (Porphyrula martinica) in Florida. American Midland Naturalist 89:467–473. Kovalenko, I. I. 1960. [Study of the life cycles of some helminths of domestic ducks from farms on the Azov coast] (in Russian). Doklady Academii Nauk SSSR 133:1259–1261. Mascarenhas, A. R., and R. C. Ghosh. 1992. Occurrence of Tetrameres mohtedai (Bhalarao and Rao, 1944) infection in the proventriculus and gizzard of fowl: Gross and histopathological changes. Indian Veterinary Journal 69:498–500. Mawson, P. M. 1977. The genus Microtetrameres Travassos (Nematoda, Spirurida) in Australian birds. Records of the South Australian Museum 17:239– 259. McDonald, M. E. 1969. Catalogue of Helminths of Waterfowl (Anatidae). Bureau of Sport Fisheries and Wildlife, Special Scientific Report No. 126, 692 pp. Mollhagen, T. R. 1976. A Study of the Systematics and Hosts of the Parasitic Nematode Genus Tetrameres
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Tetrameridosis (Habronematoidea: Tetrameridae). Ph.D. Dissertation. Texas Tech University, Lubbock, TX. Mollhagen, T. R. 1991. A study of the parasitic nematode genus Tetrameres. I. The literature, 1835–1985. Special Publications, The Museum, Texas Tech University 35:1–76. Moore, J., M. Freehling, and D. Simberloff. 1986. Gastrointestinal helminths of the northern bobwhite in Florida: 1968 and 1983. Journal of Wildlife Diseases 22:497–501. Panigrahy, B., J. E. Grimes, S. E. Glass, S. A. Naqi, and C. F. Hall. 1982. Diseases of pigeons and doves in Texas: Clinical findings and recommendations for control. Journal of the American Veterinary Medicine Association 181:384–386. Popova, Z. G. 1954. [Pathological–morphological changes in the glandular stomach of the duck caused by Tetrameres fissispina (Diesing, 1861)] (in Russian). Raboty po Gel’mintologii k 75-Letiu Akademika K. I. Skrjabin, Akademia Nauk SSSR, Moskva 547–551. Quentin, J. C., and N. Barre. 1976. Description et cycle biologique de Tetrameres (Tetrameres) cardinalis n. sp. Annales de Parasitologie Humaine et Compare´e 51:65–81. Quentin, J. C., C. Seureau, and S. D. Kulo. 1986. Life cycle of Tetrameres inermis, a tetramerid nematode parasite of the weaver Ploceus aurantius in Togo. Annales de Parasitologie Humaine et Compare´e 61:321–332.
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Ramaswamy, K., and R. K. Sundaram. 1983. Hematological changes in fowls experimentally infected with a monospecific Tetrameres mohtedai infection. Kerala Journal of Veterinary Science 14:35–44. Rust, W. 1908. Entenerkrankung durch Tropidocerca fissispina. Ver¨offentlichungen Jahres-Veterin¨arBerichten der Beamteten Tier¨arzte Preussens 6:30. Travassos, L. 1914. Contribui¸co˜ es para o conhecimento da fauna helmintolojica brazileira. 3. Sobre as especies brazileiras do genero Tetrameres Creplin, 1846. Memorias do Instituto Oswaldo Cruz 6:150–162. Tscherner, W., U. Wittstatt, and R. G¨oltenboth. 1997. Geopetitia aspiculata Webster, 1971—ein pathogener Nematode tropischer V¨ogel in Zoologischen G¨arten. Zoologische Garten 67:108–120. Webster, W. A. 1971. Geopetitia aspiculata sp. n. (Spirurida) from Coerulea coerulea and other imported birds in the National Zoological Park, Washington, DC. Proceedings of the Helminthological Society of Washington 38: 64–68. Wehr, E. E. 1971. Nematodes. In Infectious and Parasitic Diseases of Wild Birds, J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (eds). Iowa State University Press, Ames, IA, pp. 185–233. Yamaguti, S. 1961. Systema Helminthum, Vol. 3. The Nematodes of Vertebrates. Interscience Publishers, New York.
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22 Avioserpensosis John M. Kinsella galliardi Chabaud and Campana, 1949, and Avioserpens mosgovoyi Supryaga, 1965. A fourth species, Avioserpens sichuanensis, was described by Li (1983) from domestic ducks in China. Adults are ovoviviparous and are characterized by extreme sexual dimorphism. The females reach lengths of up to 64 cm while the males only range from 6 to 14 mm.
INTRODUCTION The term avioserpensosis is applied to infections and diseases caused by nematodes of the genus Avioserpens from the family Dracunculidae. This family contains only two genera—Dracunculus, parasitic in the body cavities and subcutaneous tissues of reptiles and mammals; and Avioserpens, parasitic in the subcutaneous tissues of waterbirds (Chabaud 1975). Avioserpens spp. produce tumorlike swellings in the submandibular region and legs, and sometimes on the shoulders and shanks of ducks and other waterbirds. These swellings can lead to interference with swallowing, asphyxiation, and death. At least three species of Avioserpens have caused epizootics in domestic birds in Eurasia, but similar outbreaks have not been reported thus far in wild birds.
HOST RANGE AND DISTRIBUTION Gibson (1973) summarized host and distribution records for the three species of Avioserpens recognized by Supryaga (1971a). Avioserpens taiwana is a parasite of species of Anseriformes in southeast Asia and the US, and has also been reported from the Anhinga and a “white heron” (either Ardea alba or Egretta thula) in the US. A. galliardi appears to be primarily a parasite of species of Ciconiiformes in France, Spain, Russia, Canada, and the US, but has also been reported from species of Anseriformes and Gaviiformes. Avioserpens mosgovoyi is a parasite of species of Gruiformes, Podicipediformes, Anseriformes, and Gaviiformes in Spain and Russia, and has recently been found for the first time in a Least Sandpiper (Calidris minutilla) in the US (J. Kinsella, R. Brannian, and C. Roderick, unpublished data). Avioserpens sichuanensis has so far been recorded only from domestic ducks.
HISTORY Sugimoto (1919) described Filaria taiwana from the Mallard (Anas platyrhynchos) in Formosa. Wehr and Chitwood (1934) first proposed the genus Avioserpens for a new species, Avioserpens denticulophasma, from a “white heron” and the Anhinga (Anhinga anhinga) from North America. These two species of Avioserpens are now considered to be synonyms (Supryaga 1971a). The first detailed report of disease due to Avioserpens taiwana was by Truong-Tan-Ngog (1937) in domestic ducks in Indochina. More recently, there have been a series of reports on outbreaks of disease due to Avioserpens spp. in domestic ducks, geese, and chickens in Russia (Garkavi et al. 1972; Garkavi and Golub 1974) and China (Li et al. 1981, 1983, 1990; Wang et al. 1983). Similar outbreaks have not been reported in Europe or North America.
EPIZOOTIOLOGY Transmission and Life History The females of A. taiwana are found in tumorlike swellings in subcutaneous tissues, mainly in the submandibular areas. The anterior ends of gravid females pierce the end of the swelling, producing an opening through which the first-stage larvae escape into the water. Here, they are ingested by copepods, including Cyclops sternuus, Eucyclops serratulus, Mesocyclops leuckarti, and Thermocyclops hyalinus (Wang et al. 1983). The larvae then penetrate through the gut wall of
ETIOLOGY The genus Avioserpens was reviewed by Supryaga (1971a), who recognized only three valid species: Avioserpens taiwana (Sugimoto 1919), Avioserpens
384 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Avioserpensosis the copepods into the hemocoel, molting to the second stage after 3–4 days at 28–30◦ C, and the third stage at 7 days. In experimental studies with ducklings, males were found in the mesenteries 18 days postinfection and females in the subcutaneous tissue at 20 days. The life cycle of A. mosgovoyi is quite similar, with the molt to the second stage in copepods taking place at 6–8 days postinfection and the third stage at 10–11 days (Supryaga 1969, 1971b). Paratenic or transfer hosts include fish (roach, gobies, and sticklebacks), frogs, and dragonfly larvae. Paratenic hosts can ingest infective copepods and infective, encysted larvae will persist in their tissues for up to 2.5 months. Thus, definitive hosts can become infected by ingestion of either infected copepods or infected paratenic hosts. After ingestion by Eurasian Coots (Fulica atra), the third molt takes place in the body cavity, and the fourth-stage larvae migrate via the air sacs to the subcutaneous tissues, where they molt to adults on days 12–14 postinfection. Adult females start to discharge larvae at 4 weeks postinfection. Males persist in tissues much longer than do females, up to 260 days. The life history of A. sichuanensis from Sichuan province, China, is similar (Li et al. 1988). Intermediate hosts include the copepods, Mesocyclops leuckarti, Thermocyclops hyalinus, Thermocyclops taihokuensis, Eucyclops serratulus, Macrocyclops fuscus, and Tropocyclops prasinus. In experimentally infected domestic ducklings, females begin producing larvae at 29 days postinfection, and then gradually shrivel and die over a period of 3 days. The entire life cycle, including development in the copepods, takes about 36– 40 days.
Prevalence and Intensity Although there are considerable data on prevalence and intensity of Avioserpens spp. in domestic birds, many of the records in wild birds are based on reports from a single infected host. In a study of 6 species of herons and egrets in Spain, Nogueserola et al. (2002) reported 4 of 65 hosts infected with A. galliardi, each with a single nematode. Similar infections of low prevalence and intensity have been reported for an Avioserpens spp. in Common Mergansers (Mergus merganser) from Utah (1/8 individuals, 13%) (McDonald 1974), and for A. galliardi in Great Egrets (Ardea alba) from Florida (4/63 individuals, 6%) (Sepulveda et al. 1999).
CLINICAL SIGNS Swellings in the throat may lead to difficulty in swallowing, anorexia, retarded growth, sluggishness, and eventually even asphyxiation. Swellings of the shank
385
may interfere with the ability to swim (Truong-TanNgog 1937).
PATHOLOGY In some cases, swellings containing worms are small. After adult worms die, they are slowly resorbed and replaced by fibrous tissues. In other cases, swellings are very large and occasionally as large as the head of the bird. In early stages of the infection, the swelling is painless and soft, but as the infection progresses, it becomes hard, voluminous, and painful. An open lesion is usually present through which the larvae are released. This opening may eventually close without complication by formation of scar tissue or may lead to secondary bacterial infection, abscesses, and fatal infections (Wehr 1971).
DIAGNOSIS Since adult Avioserpens spp. are ovoviviparous and the larvae are released directly into the external environment, diagnosis must be based on characteristic swellings in the submandibular areas that may also occur occasionally on the shoulders or shanks. The extremely long females are often entangled and may be difficult to remove intact. Identification of species depends on morphological features of the much smaller males, which have darkly chitinized and permanently extruded spicules (Figure 22.1a). Males are not usually present within the swelling, but may be found separately in the mesenteries or subcutaneous tissues. Both males and females have a characteristic inflation of the esophagus, which is diagnostic for members of the Dracunculidae (Figure 22.1b).
IMMUNITY Although no experimental studies on immunity have been done, a number of authors have remarked that this is primarily a disease of young birds. Garkavi and Golub (1974) found that adult domestic ducks were resistant to infection by A. mosgovoyi, and Garkavi et al. (1972) found only goslings infected with an Avioserpens species. The most severe losses in duck flocks in China due to A. sichuanensis infections were reported in ducklings 1–2 months of age (Chen et al. 1984).
PUBLIC HEATH CONCERNS There are no reports of infections of Avioserpens spp. in humans and infected birds pose no risk to public health.
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Parasitic Diseases of Wild Birds or vice versa. It is probable that crowded conditions among domestic fowl lead to very intense infections in copepod intermediate hosts which in turn lead to high prevalence and intensity of infection in susceptible avian hosts. In 14 flocks consisting of 8,173 ducklings in Sichuan province, mortality due to A. sichuanensis ranged from 50 to 62% (Li et al. 1983).
WILDLIFE POPULATION IMPACTS Reports of Avioserpens spp. infections in wild birds are sporadic and isolated. Epizootics in domestic birds demonstrate their potential for causing significant morbidity and mortality, especially in young birds, but no epizootics have been reported in wild populations, possibly because characteristic lesions have not been recognized. Biologists should pay particular attention to occurrence of swellings on the head, neck, sternum, and legs of wild birds.
TREATMENT AND CONTROL Treatment of Avioserpens infections by systemic anthelmintics has not been reported. Some cures in China have been achieved by direct injection of anthelmintics into the parasitic swellings, although this is not common practice in western veterinary medicine (Li et al. 1981; Chen et al. 1984). Prevention of infections in ducklings can be achieved by providing uncontaminated food and water and preventing contact with paratenic hosts. For example, an outbreak of A. mosgovoyi in domestic ducklings in the former USSR was attributed to ingestion of fish that had been collected from a river inhabited by infected “bald coots” (most likely Eurasian Coot, Fulica atra). Withdrawal of fish from the diet eradicated the infection (Garkavi and Golub 1974). Figure 22.1. Fixed and cleared specimens of Avioserpens mosgovoyi. (a) Posterior end of male showing darkly chitinized, permanently extruded spicules. (b) Anterior end of female showing characteristic inflation of the esophagus. Courtesy of C. Roderick and R. Brannian, U.S. Geological Survey, National Wildlife Health Center, Madison, WI.
DOMESTIC ANIMAL HEALTH CONCERNS Although significant outbreaks of disease due to Avioserpens spp. have been reported in domestic ducks and chickens in Russia and China (Garkavi and Golub 1974; Li et al. 1981, 1990; Chen et al. 1984), there is no evidence of these infections spreading to wild birds
LITERATURE CITED Chabaud, A. G. 1975. Keys to genera of the order Spirurida. Part 1. Camallanoidea, Dracunculoidea, Gnathostomoidea, Physalopteroidea, Rictularoidea, and Thelazoidea. In CIH Keys to the Nematode Parasites of Vertebrates, R. C. Anderson, A. G. Chabaud, and S. Willmott (eds). Commonwealth Agricultural Bureaux, Farnham Royal, Buckinghamshire, UK, pp. 1–27. Chen, F., C. Y. Xue, X. L. Zhou, K. R. Zhang, and G. Z. Ying. 1984. The finding of Avioserpens sichuanensis in ducks in Huangyan County, Zhejiang Province, China (in Chinese). Chinese Journal of Veterinary Medicine, Zhongguo Shouyi Zazhi 10:23– 25.
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Avioserpensosis Garkavi, H. L., and V. E. Golub. 1974. Avioserpens mosgovoyi in ducks (in Russian). Veterinariya, Moscow 9:73–74. Garkavi, B. L., G. I. Monko, and N. S. Marchenko. 1972. An outbreak of Avioserpens infection in geese (in Russian). Nauchnye Trudy Krasnodarskogo Nauchno Issledovatel’skoi Veterinarnoi Stantsii Bolezni sel’skokhozyaistvennykh zhivotnykh 5:311. Gibson, G. G. 1973. Cardiofilaria pavlovskyi Strom, 1937 and Avioserpens sp. (Nematoda) from Canadian ciconiiform birds. Canadian Journal of Zoology 51: 847–851. Li, M. Z. 1983. Description and supplementary information on the morphology of Avioserpens sichuanensis sp. nov (in Chinese). Chinese Journal of Veterinary Medicine, Zhongguo Shouyi Zazhi 9:6–25. Li, M. Z., G. R. Sa, and T. F. Zhang. 1981. A survey of Avioserpens szechuanensis in ducks. II. Prevalence and treatment with tetramisole (in Chinese). Chinese Journal of Veterinary Medicine, Zhongguo Shouyi Zazhi 7:2–6. Li, M. Z., M. X. Huang, S. Z. Zhao, B. Y. Ni, W. Y. Hu, H. P. Li, H. Y. Gong, G. T. Zhang, C. Z. Liang, D. H. Zhang, X. W. Jiang-XW, S. K. Li, Y. B. Yan, and M. Zhang. 1983. A survey of Avioserpens sichuanensis infection (filariasis of the duck) in Sichuan Province (in Chinese). Chinese Journal of Veterinary Medicine, Zhongguo Shouyi Zazhi 9:2–4. Li, M. Z., M. W. Li, and T. J. Cheng. 1988. Studies on the life history and morphology of Avioserpens sichuanensis. In Waterfowl Production. Proceedings of the International Symposium, September 11–18, 1988 Beijing, China. International Academic Publishers, Beijing, China, pp. 299–305. Li, M. Z., Z. J. Cheng, and X. Q. Gou. 1990. A preliminary report on Avioserpens infection in domestic chickens in China (in Chinese). Acta Veterinaria et Zootechnica Sinica 21:64. McDonald, M. E. 1974. Nematode parasites of waterfowl (Anseriformes) from western United States. Wildlife Disease 64:17.
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Nogueserola, M. L., P. Navarro, and J. Lluch. 2002. Helmintos par´asitos de Ardeidae en Valencia (Espana). Anales de Biologia 24:139–144. Sepulveda, M. S., M. G. Spalding, J. M. Kinsella, and D. J. Forrester. 1999. Parasites of the great egret (Ardea albus) in Florida and a review of the helminths reported for the species. Journal of the Helminthological Society of Washington 66: 7–13. Sugimoto, M. 1919. List of Zooparasites of the Domesticated Animals in Formosa. Taihoku, Formosa, Japan. Supryaga, A. M. 1969. The life-span of Avioserpens mosgovoyi in final hosts (in Russian). Problemy Parazitologii 1:245–246. Supryaga, A. M. 1971a. Revision of Avioserpens Wehr et Chitwood, 1934 (Camallanata, Dracunculidae) (in Russian). Trudy Gel’mintologicheskoi Laboratorii Teoreticheskie Voprosy Obshchei Gel’mintologii 22:196–200. Supryaga, A. M. 1971b. Life cycle of Avioserpens mosgovoyi (Camallanata: Dracunculidae), nematodes of domestic birds (in Russian). In Sbornik Rabot po Gel’mintologii pos vyashchen 90-letiyu so dnya rozhdeniya Akademika K. I. Skrjabina. Izdatel-stvo Kolos, Moscow, pp. 374–383. Truong-Tan-Ngog. 1937. Filariose du canard domestique en Cochinchine due a` Oshimaia taiwana (Suginoto, 1919). Bulletin Societ´e Pathologique Exotique 30:775–778. Wang, P., Y. Sun, and Y. Zhao. 1983. Studies on the life history and epidemiology of Avioserpens taiwana of the domestic duck in Fujian (China) (in Chinese). Acta Zoologica Sinica 29:350–357. Wehr, E. E. 1971. Nematodes. In Infectious and Parasitic Diseases of Wild Birds, J. W. Davis, R. C. Anderson, L. Karstad, and D. O. Trainer (eds). Iowa State University Press, Ames, IA, pp. 185–233. Wehr, E. E., and B. G. Chitwood. 1934. A new nematode from birds. Proceedings of the Helminthological Society of Washington 1:10–11.
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23 Heterakis and Ascaridia Alan M. Fedynich 1926; Canavan 1929, 1931). Information from these early studies provided important insights into variation in pathogenicity of these worms among different host species. For example, reports of nodular typhlitis associated with severe infections of Heterakis isolonche in several species of captive pheasants helped to establish that some hosts are highly susceptible to these parasites (Schwartz 1924; Griner et al. 1977). Subsequent studies established that Heterakis gallinarum and Ascaridia galli are particularly pathogenic to Helmeted Guineafowl (Numida meleagris) (Ayeni et al. 1983), but not to Northern Bobwhite (Colinus virginianus) (Cram et al. 1931; Davidson et al. 1982). Growing interest in the early twentieth century both in the potential exchange of parasites between wild and domestic populations and in the importance of parasites as causes of mortality in game birds led to a series of parasitological surveys of game species in North America and Europe, including Northern Bobwhite (Cram et al. 1931), Ruffed Grouse (Bonasa umbellus) (Morgan and Hamerstrom 1941; Erickson et al. 1949), Wild Turkey (Meleagris gallopavo) (Maxfield et al. 1963), Ring-necked Pheasant (Phasianus colchicus) (Gilbertson and Hugghins 1964), and European species of grouse, partridges, and pheasants (Madsen 1941, 1952). Consequently, early studies focusing on wild game birds contributed significantly to our understanding about the occurrence and geographic distribution of species of Heterakis and Ascaridia.
INTRODUCTION The genera Heterakis and Ascaridia are members of the nematode order Ascaridida, a relatively large group of parasitic nematodes that have direct life cycles and occur primarily in the gastrointestinal tracts of their hosts. Both genera occur worldwide, and one or more species have been reported from most of the major taxonomic orders of birds. Pathogenicity varies widely within both genera and is dependent on host group and history of host–parasite association. More species of Ascaridia appear to negatively affect birds than species of Heterakis. However, this may reflect disproportionate attention on those parasites that have economic impacts. Species of Heterakis and Ascaridia have caused disease and/or reduced fitness in economically important domesticated birds such as chickens, pheasants, and turkeys (Schmidt and Kuntz 1970; Norton et al. 1999; Menezes et al. 2003), wild-caught captive birds in zoological gardens (Griner et al. 1977; Callinan 1987; Balaguer et al. 1992), pen-raised game birds (Woodburn 1994; Mill´an et al. 2004a), and wild birds (Wehr and Shalkop 1963; Ayeni et al. 1983; Rizzoli et al. 1999; Draycott et al. 2000; Tompkins et al. 2002; Daehlen 2003). SYNONYMS Heterakiasis, heterakosis, ascariasis, ascarosis. HISTORY The heterakid and ascarid nematodes of birds have been recognized since the late 1700s. Many of the first descriptions of these parasitic nematodes were provided by early investigators, including Bloch, Dujardin, Fr¨ohlich, Gmelin, Linstow, Railliet, Rudolphi, Schneider, and Schrank (Skrjabin et al. 1951; Yamaguti 1961). By the early 1900s, new species were described and diseases noted from necropsies performed primarily on captive wild hosts from zoological gardens (Baylis and Daubney 1922; Chandler
DISTRIBUTION AND HOST RANGE Species of Heterakis and Ascaridia are widely distributed geographically, and at least one species from each genus has been found on all continents except Antarctica (Tables 23.1 and 23.2). Heterakid nematodes have been found in at least 107 wild and captive bird species (Table 23.1), and ascarid nematodes have been reported from at least 139 host species (Table 23.2). Several species are cosmopolitan (e.g.,
388 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Table 23.1. Nonexhaustive list of avian hosts infected with Heterakis spp., geographic location where infection was reported, and reporting authors. Host STRUTHIONIFORMES Rheidae Greater Rhea (Rhea americana) TINAMIFORMES Tinamidae Gray Tinamou (Tinamus tao) Red-winged Tinamou (Rhynchotus rufescens) Small-billed Tinamou (Crypturellus parvirostris) Solitary Tinamou (Tinamus solitarius) Spotted Nothura (Nothura maculosa) Tataupa Tinamou (Crypturellus tataupa) Undulated Tinamou (Crypturellus undulates) Variegated Tinamou (Crypturellus variegates) Yellow-legged Tinamou (Crypturellus noctivagus)
Heterakis sp.
Location
Heterakis gallinarum France (zoo)
Cram (1927)
Heterakis nattereri* Brazil Heterakis gallinarum Brazil
Vicente et al. (1993) Vicente et al. (1993)
Heterakis inglisi
Brazil
Vicente et al. (1993)
Heterakis nattereri* Heterakis gallinarum Heterakis spiculata Heterakis spiculata Heterakis spiculata
Brazil Brazil South America Brazil Brazil
Vicente et al. (1993) Vicente et al. (1993) Kaseta (1973) Pinto et al. (2006) Vicente et al. (1993)
Heterakis inglisi
Brazil
Vicente et al. (1993)
Heterakis spiculata Heterakis spiculata
Brazil Brazil
Pinto et al. (2006) Vicente et al. (1993)
CICONIIFORMES Ardeidae Black-crowned Night-Heron (Nycticorax Heterakis pavonis Japan nycticorax) Ciconiidae Maguari Stork (Ciconia maguari) Heterakis Brazil valdemucronata* Marabou Stork (Leptoptilos Heterakis gallinarum Uganda crumeniferus) ANSERIFORMES Anatidae Ashy-headed Goose (Chloephaga poliocephala) Barnacle Goose (Branta leucopsis) Black Swan (Cygnus atratus) Canada Goose (Branta canadensis) Cape Barren Goose (Cereopsis novaehollandiae) Common Shelduck (Tadorna tadorna) Eurasian Wigeon (Anas penelope) Gadwall (Anas strepera) Greater White-fronted Goose (Anser albifrons) Greylag Goose (Anser anser) Hawaiian Goose (Branta sandvicensis) Lesser White-fronted Goose (Anser erythropus) Mallard (Anas platyrhynchos)
Reference
Madsen (1950)
Cram (1927) Bwangamoi et al. (2003)
Heterakis dispar
England (zoo)
Madsen (1950)
Heterakis dispar Heterakis dispar Heterakis gallinarum Heterakis dispar Heterakis dispar
NR Germany (zoo) Germany (zoo) USA India
Cram (1927) Cram (1927) Johnston (1912) Cram (1927) Maplestone (1932)†
Heterakis dispar Heterakis gallinarum Heterakis dispar Heterakis dispar Heterakis dispar
NR NR NR NR Texas, USA
Cram (1927) Cram (1927) McDonald (1969) McDonald (1969) Purvis et al. (1997)
Heterakis dispar Heterakis dispar Heterakis dispar
NR NR NR
McDonald (1969) Cram (1927) McDonald (1969)
Heterakis dispar
NR
McDonald (1969) (continues)
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Table 23.1. (Continued ) Host
Heterakis sp.
Location
Reference
Maned Duck (Chenonetta jubata) Muscovy Duck (Cairina moschata) Ross’ Goose (Chen rossii) Snow Goose (Chen caerulescens)
Heterakis dispar Heterakis dispar Heterakis dispar Heterakis dispar
Australia (zoo) NR Texas, USA North America
Speckled Teal (Anas flavirostris) Swan Goose (Anser cygnoides) Taiga Bean-Goose (Anser fabalis) Upland Goose (Chloephaga picta)
Heterakis dispar Heterakis dispar Heterakis dispar Heterakis dispar
West Indian Whistling-Duck (Dendrocygna arborea) Whooper Swan (Cygnus cygnus) Wood Duck (Aix sponsa)
Heterakis dispar
NR NR NR Chile Falkland Islands NR
Johnston (1912) Cram (1927) Fedynich et al. (2005) Swinyard (1931) and Forbes et al. (1999) McDonald (1969) McDonald (1969) Cram (1927) Gonz´alez et al. (2005) Harradine (1982) McDonald (1969)
Heterakis dispar Heterakis dispar
NR Germany (zoo)
McDonald (1969) Cram (1927)
FALCONIFORMES Cathartidae King Vulture (Sarcoramphus papa) GALLIFORMES Cracidae Red-billed Curassow (Crax blumenbachii) Megapodiidae Australian Brush-turkey (Alectura lathami) Maleo (Macrocephalon maleo) Odontophoridae California Quail (Callipepla californica) Northern Bobwhite (Colinus virginianus)
Heterakis gallinarum NR
Madsen (1950)
Heterakis nattereri*
Brazil
Cram (1927)
Heterakis oscari
Brazil
Yamaguti (1961)
Heterakis gallinarum NR
Madsen (1950)
Heterakis gallinarum NR
Cram (1927)
Heterakis isolonche Oregon, USA Heterakis gallinarum North America
Moore et al. (1989) Venard (1933) and Davidson et al. (1982) Cram et al. (1931) and Venard (1933)
Heterakis isolonche Phasianidae Barbary Partridge (Alectoris barbara)
North America
Black Grouse (Tetrao tetrix)
Heterakis gallinarum Canary Islands Heterakis tenuicauda Algeria Heterakis gallinarum Europe
Blood Pheasant (Ithaginis cruentus)
Heterakis isolonche
Dusky Grouse (Dendragapus obscurus) Blue Eared-Pheasant (Crossoptilon auritum)
Heterakis gallinarum Heterakis isolonche
Brown Eared-Pheasant (Crossoptilon mantchuricum) Caspian Snowco*ck (Tetraogallus caspius) Ceylon Junglefowl (Gallus lafayetii) Cheer Pheasant (Catreus wallichi) Chinese Francolin (Francolinus pintadeanus)
Heterakis gallinarum Heterakis isolonche Heterakis macroura Heterakis pusilla* Heterakis gallinarum Heterakis gallinarum
390
Foronda et al. (2005) Madsen (1950) Madsen (1952) and Bezubik (1960) India (zoo) Baylis and Daubney (1922) North America Buss et al. (1958) Canada Webster (1982) USA (zoo, origin Canavan (1929) China) NR Madsen (1950) USA (zoo) Griner et al. (1977) Turkestan Yamaguti (1961) Africa Cram (1927) NR Madsen (1950) NR Madsen (1950)
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Table 23.1. (Continued ) Host Chukar (Alectoris chukar) Common Quail (Coturnix coturnix) Copper Pheasant (Syrmaticus soemmerringii) Crested Fireback (Lophura ignita) Crested Partridge (Rollulus rouloul) Double-spurred Francolin (Francolinus bicalcaratus) Eurasian Capercaillie (Tetrao urogallus) Golden Pheasant (Chrysolophus pictus)
Gray Junglefowl (Gallus sonneratii)
Gray Partridge (Perdix perdix) Gray Peaco*ck-Pheasant (Polyplectron bicalcaratum) Greater Prairie-Chicken (Tympanuchus cupido) Greater Sage-Grouse (Centrocercus urophasianus) Green Peafowl (Pavo muticus) Green Pheasant (Phasianus versicolor) Hazel Grouse (Bonasa bonasia) Helmeted Guineafowl (Numida meleagris) Hill Partridge (Arborophila torqueola)
Himalayan Monal (Lophophorus impejanus)
Heterakis sp.
Location
Heterakis gallinarum India Heterakis gallinarum Russia Turkey Heterakis gallinarum NR Heterakis pavonis Japan Heterakis gallinarum India Heterakis isolonche India (zoo) Heterakis gallinarum Canada Heterakis Africa brevispiculum Heterakis gallinarum Europe
Reference Maplestone (1932)† Wetherbee (1961) Kurtpinar (1957) Madsen (1950) Madsen (1950) Maplestone (1932)† Chandler (1926) Webster (1982) Cram (1927)
Bezubik (1960) and Baruˇs et al. (1984) Heterakis gallinarum USA (zoo, origin Canavan (1929, 1931) China) Heterakis isolonche USA (zoo) Cram (1927) and Griner et al. (1977) Heterakis Asia Levine (1968) beramporia India (zoo) Madsen (1950) Heterakis isolonche India Maplestone (1932)† Heterakis gallinarum Europe Clapham (1935), Madsen (1941, 1952) North America Yocom (1943) Heterakis Africa, Brazil, Levine (1968) brevispiculum Puerto Rico India (zoo) Chandler (1926) Heterakis isolonche India (zoo) Maplestone (1932)† Heterakis gallinarum North America Morgan and Hamerstrom (1941) and Harper et al. (1967) Heterakis gallinarum North America Simon (1940) and Wehr (1940a) Heterakis hamulus* Germany (zoo) Cram (1927) Heterakis gallinarum NR Cram (1927) Heterakis pavonis Japan Madsen (1950) Heterakis gallinarum NR Skrjabin et al. (1951) Heterakis silindae China Madsen (1950) Heterakis Africa Madsen (1950) brevispiculum Heterakis gallinarum Africa Ayeni et al. (1983) Heterakis gallinarum India (zoo) Baylis and Daubney (1922) Heterakis India (zoo) Chandler (1926) vulvolabiata Heterakis gallinarum India (zoo) Baylis and Daubney (1922) Heterakis isolonche
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India (zoo)
Baylis and Daubney (1922) (continues)
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Table 23.1. (Continued ) Host
Heterakis sp.
Himalayan Snowco*ck (Tetraogallus himalayensis) Indian Peafowl (Pavo cristatus)
Heterakis macroura
Lesser Prairie-Chicken (Tympanuchus pallidicinctus) Long-billed Partridge (Rhizothera longirostris)
Heterakis isolonche
Heterakis gallinarum Heterakis hamulus* Kalij Pheasant (Lophura leucomelanos) Heterakis gallinarum Heterakis isolonche Lady Amherst’s Pheasant Heterakis gallinarum Heterakis isolonche (Chrysolophus amherstiae)
Ocellated Turkey (Meleagris ocellata) Red Junglefowl (Gallus gallus)
Red Spurfowl (Galloperdix spadicea) Red-legged Partridge (Alectoris rufa)
Heterakis interlabiata
Location Turkestan
Madsen (1950)
NR Germany (zoo) India India NR Germany USA (zoo, origin China) Texas, USA
Cram (1927) Cram (1927) Maplestone (1932)† Maplestone (1932)† Madsen (1950) Yamaguti (1961) Canavan (1931) Pence and Sell (1979)
Africa
Yamaguti (1961)
Asia England (zoo) USA (zoo) Asia
Cram (1927) Madsen (1950) Griner et al. (1977) Cram (1927)
Heterakis isolonche Heterakis beramporia Heterakis South America brevispiculum Heterakis gallinarum Asia Heterakis indica Heterakis kumaoni Heterakis nainitalensis Heterakis parva Heterakis pusilla* Heterakis gallinarum Heterakis gallinarum
Asia India India Asia Africa India (zoo) Europe
Red-necked Francolin (Francolinus Heterakis silindae Africa afer) Reeves’ Pheasant (Syrmaticus reevesii) Heterakis gallinarum NR Ring-necked Pheasant (Phasianus Heterakis gallinarum Europe colchicus) North America North America, Russia Heterakis pavonis Asia Heterakis dispar Turkey Heterakis gallinarum Italy Turkey Heterakis tenuicauda Italy Turkestan Heterakis isolonche
Rock Partridge (Alectoris graeca)
392
Reference
Yamaguti (1961) Schmidt and Kuntz (1970) and Arya (1990) Schmidt and Kuntz (1970) Arya (1990) Arya (1990) Schmidt and Kuntz (1970) Cram (1927) Baylis and Daubney (1922) Clapham (1935) and Mill´an et al. (2004b) Yamaguti (1961) Madsen (1950) Clapham (1935) and Madsen (1952) Gilbertson and Hugghins (1964) Yamaguti (1961) Levine (1968) K¨oroˇglu and Tasan (1996) Rizzoli et al. (1999) K¨oroˇglu and Tasan (1996) Rizzoli et al. (1999) Madsen (1950)
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Table 23.1. (Continued ) Host Rock Ptarmigan (Lagopus muta)
Heterakis sp.
Location
Heterakis gallinarum Europe North America
Ruffed Grouse (Bonasa umbellus)
Satyr Tragopan (Tragopan satyra)
Heterakis gallinarum North America Heterakis isolonche
North America
Heterakis bosia
India (zoo)
Heterakis gallinarum India (zoo) Heterakis isolonche See-see Partridge (Ammoperdix griseogularis) Sharp-tailed Grouse (Tympanuchus phasianellus) Silver Pheasant (Lophura nycthemera)
India (zoo)
Heterakis tragopanis India (zoo) Heterakis tenuicauda Turkestan Heterakis gallinarum North America
Willow Ptarmigan (Lagopus lagopus)
Madsen (1952) (citing Galli-Valerio 1931 and Redi 1708) Braun and Willers (1967) (see references therein) Wehr (1940a) and Erickson et al. (1949) Wehr (1940a) and Kalla et al. (1997) Baylis and Daubney (1922) Baylis and Daubney (1922) Baylis and Daubney (1922) Lal (1942) Madsen (1950)
Heterakis pedioecetes Heterakis beramporia Heterakis dispar Heterakis gallinarum Heterakis isolonche Heterakis parva Heterakis pavonis
Canada
Morgan and Hamerstrom (1941) Mawson (1956)
Asia
Levine (1968)
Heterakis gallinarum Heterakis tenuicauda Heterakis gallinarum Heterakis meleagris Heterakis gallinarum
India Africa North America China Europe North America
India India India India (zoo) Asia India (zoo) Sooty Grouse (Dendragapus fuliginosus) Heterakis gallinarum North America Swamp Francolin (Francolinus gularis) Heterakis gallinarum India (zoo) Vulturine Guineafowl (Acryllium vulturinum) Wild Turkey (Meleagris gallopavo)
Reference
GRUIFORMES Otididae Great Bustard (Otis tarda)
Heterakis gallinarum Asia, Germany (zoo) Houbara Bustard (Chlamydotis undulate) Heterakis gallinarum India Little Bustard (Tetrax tetrax) Heterakis gallinarum Asia, Germany (zoo) Psophiidae Dark-winged Trumpeter (Psophia viridis) Heterakis psophiae Brazil Heterakis skrjabini* Brazil
Maplestone (1932)† Maplestone (1932)† Maplestone (1932)† Maplestone (1932)† Levine (1968) Maplestone (1932)† Beer (1944) Baylis and Daubney (1922) Maplestone (1932)† Yamaguti (1961) Maxfield et al. (1963) Yamaguti (1961) Madsen (1950) Cram (1927)
Cram (1927) Madsen (1950) Cram (1927) Cram (1927) Cram (1927) (continues)
393
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394
Parasitic Diseases of Wild Birds
Table 23.1. (Continued ) Host CHARADRIIFORMES Alcidae Crested Auklet (Aethia cristatella)
Whiskered Auklet (Aethia pygmaea) COLUMBIFORMES Columbidae Eared Dove (Zenaida auriculata) Eurasian Collared-Dove (Streptopelia decaocto) PSITTACIFORMES Cacatuidae Pink co*ckatoo (Cacatua leadbeateri) STRIGIFORMES Strigidae Eurasian Pygmy-Owl (Glaucidium passerinum)
Heterakis sp.
Location
Heterakis kurilensis
Russia
Heterakis kurilensis
Russia
Reference
Yamaguti (1961) and Muzaffar and Jones (2004) (citing Baruˇs et al. 1978) Yamaguti (1961)
Heterakis gallinarum Chile Heterakis gallinarum Czechoslovakia
Gonz´alez et al. (2004) Baruˇs (1966a)
Heterakis gallinarum NR
Madsen (1950)
Heterakis dispar
NR
Cram (1927)
Note: Some host–parasite reports found only in the review literature did not include geographic location and often included hosts from wild, domestic, and private and public zoological collections; where possible, hosts from zoos and host origin of capture are noted. NR, not reported. * Reported as species inquirenda by Madsen (1950). † Maplestone (1932) included hosts pooled together from zoo and/or domestic birds. Heterakis dispar, H. gallinarum, Ascaridia columbae, A. galli), which reflects the worldwide distribution of their hosts. Other species tend to occur in specific geographic regions (e.g., Heterakis pavonis in Japan, Ascaridia geei in China) or tend to occur in particular host families (e.g., Heterakis kurilensis in Alcidae, Ascaridia aegyptiaca in Caprimulgidae, and Ascaridia hermaphroditia and Ascaridia platyceri in Psittacidae). ETIOLOGY Heterakis and Ascaridia are classified in the phylum Nemata (Nematoda), class Secernentea, order Ascaridida, and superfamily Heterakoidea. Members of this superfamily have monoxenous (direct) life cycles, develop within the gastrointestinal tract of the definitive host, produce eggs that embryonate outside the host, and have a prominent preanal sucker encompassed by a cuticularized ring (Anderson 2000). The superfamily Heterakoidea includes the families Heterakidae and Ascaridiidae. The subfamily Heterakinae is placed within the Heterakidae and contains the genera Africana, Ganguleterakis, Heraldakis, Heter-
akis, Odontoterakis, Paraspidodera, Spinicauda, and Strongyluris (Skrjabin et al. 1951; Inglis 1991). Depending on taxonomic authority, there are about 29 species of Heterakis (Table 23.1). Some of these are incompletely described and will likely be synonymized after additional study (Madsen 1950). The family Ascaridiidae includes the subfamily Ascaridiinae with the single genus Ascaridia (Skrjabin et al. 1951). At least 41 species of Ascaridia have been reported from birds (Table 23.2). Species of Heterakis are small (5.5–31 mm; males slightly smaller than females), white-to-tan colored nematodes (Figure 23.1) that typically occur in the ceca of the definitive host (Cram 1927; Maplestone 1932; Madsen 1950). Like other members of their subfamily, heterakid nematodes have a transversely striated cuticle; three lips that encircle the mouth, with each lip having two papillae; lateral flanges or alae (may be absent); a three-part esophagus composed of a pharynx that is narrow anteriorly, broadens posteriorly, and ends with a well-developed posterior bulb; and a valvular apparatus within the bulb of the esophagus (Figure 23.2) (Skrjabin et al. 1951; Yamaguti 1961; Yorke and Maplestone 1962).
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Table 23.2. Nonexhaustive list of avian hosts infected with Ascaridia spp., geographic location where infection was reported, and reporting authors. Host
Ascaridia sp.
Location
Reference
STRUTHIONIFORMES Rheidae Greater Rhea (Rhea americana)
Ascaridia orthocerca
Cram (1927)
Struthionidae Ostrich (Struthio camelus)
Brazil, Italy (zoo)
Ascaridia struthionis
Italy (zoo)
Skrjabin et al. (1951)
Ascaridia pintoi
Brazil
Skrjabin et al. (1951)
Ascaridia brasiliana
Brazil
Yamaguti (1961)
Ascaridia aegyptiaca
Egypt
Cram (1927)
Ascaridia galli
Africa
Bwangamoi et al. (2003)
Ascaridia galli Ascaridia galli Ascaridia galli Ascaridia galli Ascaridia styphlocerca Ascaridia galli Ascaridia galli
NR NR NR NR Africa NR NR
McDonald (1969) McDonald (1969) Skrjabin et al. (1951) Gower (1939) Gower (1939) Gower (1939) Cram (1927)
TINAMIFORMES Tinamidae Red-winged Tinamou (Rhynchotus rufescens) Spotted Nothura (Nothura maculosa) CICONIIFORMES Ardeidae Little Egret (Egretta garzetta) Ciconiidae Marabou Stork (Leptoptilos crumeniferus) ANSERIFORMES Anatidae Black Scoter (Melanitta nigra) Common Pochard (Aythya ferina) Greylag Goose (Anser anser) Mallard (Anas platyrhynchos) Muscovy Duck (Cairina moschata) Northern Pintail (Anas acuta)
FALCONIFORMES Accipitridae Crested Goshawk (Accipiter trivirgatus) Ascaridia galli Taiwan Eastern Marsh-Harrier (Circus Ascaridia dolichocerca New Guinea spilonotus) Sharp-shinned Hawk (Accipiter striatus) Ascaridia galli Massachusetts, USA GALLIFORMES Cracidae Ascaridia sergiomeirai Brazil Black-fronted Piping-Guan (Pipile jacutinga) Ascaridia serrata Brazil
Su and Fei (2004) Cram (1927) Rankin (1946)
Yamaguti (1961) Skrjabin et al. (1951)
Megapodiidae Australian Brush-turkey (Alectura lathami)
Ascaridia catheturina
Australia
Skrjabin et al. (1951)
Odontophoridae Northern Bobwhite (Colinus virginianus)
Ascaridia compar
NR
Ascaridia galli Ascaridia cordata
North America Mexico
Baylis and Daubney (1922) Cram et al. (1931) Cram (1927)
Ascaridia francolina
Africa
Skrjabin et al. (1951)
Scaled Quail (Callipepla squamata) Phasianidae Ahanta Francolin (Francolinus ahantensis)
(continues)
395
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Table 23.2. (Continued ) Host
Ascaridia sp.
Location
Barbary Partridge (Alectoris barbara) Black Grouse (Tetrao tetrix)
Ascaridia galli Ascaridia compar Ascaridia galli Ascaridia magnipapilla
Canary Islands Europe Denmark Europe
Blood Pheasant (Ithaginis cruentus)
Ascaridia galli
India (zoo)
Dusky Grouse (Dendragapus obscurus) Ascaridia bonasae Chukar (Alectoris chukar) Ascaridia compar
Ascaridia galli Ascaridia numidae Common Quail (Coturnix coturnix) Crested Guineafowl (Guttera pucherani) Double-spurred Francolin (Francolinus bicalcaratus) Eurasian Capercaillie (Tetrao urogallus)
Gray Partridge (Perdix perdix) Greater Prairie-Chicken (Tympanuchus cupido) Hazel Grouse (Bonasa bonasia) Helmeted Guineafowl (Numida meleagris) Himalayan Snowco*ck (Tetraogallus himalayensis) Indian Peafowl (Pavo cristatus) Ocellated Turkey (Meleagris ocellata) Red Junglefowl (Gallus gallus)
Ring-necked Pheasant (Phasianus colchicus) Rock Partridge (Alectoris graeca)
Ascaridia compar Ascaridia numidae Ascaridia francolina
Canada India (zoo) USA (zoo, origin Asia) Nevada, USA USA (zoo, origin Asia) Russia Africa Africa
Reference Foronda et al. (2005) Bezubik (1960) Madsen (1952) Cram (1927) and Baruˇs (1966b) Baylis and Daubney (1922) Bendell (1955) Baylis and Daubney (1922) Canavan (1929) Tibbitts and Babero (1969) Canavan (1929) Wetherbee (1961) Skrjabin et al. (1951) Cram (1927)
Lund (1946) and Baruˇs et al. (1984) Ascaridia galli Europe Lund (1946) and Baruˇs (1966b) Ascaridia magnipapilla Europe Baruˇs (1966b) Ascaridia compar Czechoslovakia Baruˇs (1966b) Denmark Madsen (1941) Ascaridia galli Denmark Madsen (1952) Ascaridia galli North America Morgan and Hamerstrom (1941) Ascaridia compar Europe Skrjabin et al. (1951) Ascaridia galli NR Skrjabin et al. (1951) NR Baylis and Daubney Ascaridia compar (1922) Ascaridia galli Nigeria Ayeni et al. (1983) Ascaridia numidae Africa Cram (1927) Ascaridia skrjabini Russia Yamaguti (1961) Ascaridia compar
Europe
Skrjabin et al. (1951) Yamaguti (1961) Skrjabin et al. (1951) Baylis and Daubney (1922) Ascaridia galli Asia Schmidt and Kuntz (1970) Brazil Cram (1927) Ascaridia styphlocerca Africa Yamaguti (1961) Ascaridia compar Czechoslovakia Baruˇs (1966b) Ascaridia galli Europe Madsen (1952) North America Gilbertson and Hugghins (1964) Ascaridia compar Italy Rizzoli et al. (1999) Ascaridia numidae Africa Skrjabin et al. (1951) Ascaridia columbae Ascaridia galli Ascaridia galli Ascaridia compar
396
NR NR NR NR
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Table 23.2. (Continued ) Host
Ascaridia sp.
Location
Ascaridia borealis Ascaridia compar Ascaridia galli
Europe Alaska, USA Norway Alaska, USA
Ascaridia bonasae
North America
Ascaridia galli
North America
Sharp-tailed Grouse (Tympanuchus phasianellus)
Ascaridia galli
North America
Vulturine Guineafowl (Acryllium vulturinum) White-tailed Ptarmigan (Lagopus leucura)
Ascaridia numidae Ascaridia compar
USA (zoo, origin Africa) Alaska, USA Babero (1953)
Ascaridia galli
Alaska, USA
Wild Turkey (Meleagris gallopavo)
Ascaridia dissimilis
Willow Ptarmigan (Lagopus lagopus)
Ascaridia galli Ascaridia borealis Ascaridia compar
Europe North America North America Europe Europe North America Alaska, USA
Rock Ptarmigan (Lagopus muta)
Ruffed Grouse (Bonasa umbellus)
Ascaridia galli
GRUIFORMES Cariamidae Red-legged Seriema (Cariama cristata) Ascaridia pterophora Gruidae Black Crowned-Crane (Balearica pavonina) Blue Crane (Anthropoides paradiseus)
Reference Skrjabin et al. (1951) Babero (1953) Madsen (1952) Braun and Willers (1967) (citing DeLeonardis 1952) Rankin (1946) and Mawson (1956) Connell and Doremus (1937) and Wehr (1940a, b) Morgan and Hamerstrom (1941) and Wehr (1940a) Canavan (1929)
Braun and Willers (1967) (citing DeLeonardis 1952) Levine (1968) Maxfield et al. (1963) Maxfield et al. (1963) Skrjabin et al. (1951) Madsen (1952) Babero (1953) Braun and Willers (1967) (citing DeLeonardis 1952)
Brazil
Cram (1927)
Ascaridia cristata
India (zoo)
Ascaridia stroma
Baylis and Daubney (1922) Canavan (1929)
Common Crane (Grus grus)
Ascaridia stroma
Demoiselle Crane (Anthropoides virgo)
Ascaridia stroma
Gray Crowned-Crane (Balearica regulorum) Sandhill Crane (Grus canadensis) Sarus Crane (Grus antigone)
Ascaridia cristata Ascaridia pterophora Ascaridia cristata
Whooping Crane (Grus americana)
Ascaridia stroma Ascaridia pterophora
USA (zoo, origin Africa) India (zoo) Baylis and Daubney (1922) USA (zoo, Canavan (1931) origin Asia) NR Baylis and Daubney (1922) Florida, USA Spalding et al. (1996) India (zoo) Baylis and Daubney (1922) NR Cram (1927) Florida, USA Spalding et al. (1996)
Ascaridia fasciata
Africa
COLUMBIFORMES Columbidae African Green-Pigeon (Treron calvus)
Skrjabin et al. (1951) (continues)
397
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Table 23.2. (Continued ) Host
Ascaridia sp.
Band-tailed Pigeon (Patagioenas Ascaridia columbae fasciata) Black-billed Cuckoo-Dove (Macropygia Ascaridia australis nigrirostris) Common Wood-Pigeon (Columba Ascaridia columbae palumbus) Eurasian Collared-Dove (Streptopelia decaocto) Eurasian Turtle-Dove (Streptopelia turtur) Luzon Bleeding-heart (Gallicolumba luzonica) Madagascar Green-Pigeon (Treron australis) Mourning Dove (Zenaida macroura) Oriental Turtle-Dove (Streptopelia orientalis) Picui Ground-Dove (Columbina picui) Rameron Pigeon (Columba arquatrix) Rock Pigeon (Columba livia)
Ruddy Ground-Dove (Columbina talpacoti) Ruddy Quail-Dove (Geotrygon montana) Scaled Pigeon (Patagioenas speciosa) Spotted Dove (Streptopelia chinensis) White-winged Dove (Zenaida asiatica) Yellow-footed Pigeon (Treron phoenicopterus) PSITTACIFORMES Cacatuidae co*ckatiel (Nymphicus hollandicus)
Ascaridia columbae Ascaridia columbae Ascaridia columbae
Reference
Colorado, USA
Olsen and Braun (1980)
Australia
Cram (1927)
Czechoslovakia Baruˇs (1966b) (citing Vojtechovsk´aMayerov´a 1952) Czechoslovakia Baruˇs (1966a) Florida, USA Bean et al. (2005) Czechoslovakia Baruˇs (1966a) Canada India (zoo)
Ascaridia longecirrata Belgian Congo
Webster (1982) Baylis and Daubney (1922) Yamaguti (1961)
Ascaridia maculosa Ascaridia razia Ascaridia columbae Ascaridia magalh˜aesi Ascaridia magalh˜aesi
North America NR China NR NR Africa Chile Czechoslovakia India North America Germany India NR Brazil Brazil
Lee et al. (2004) Skrjabin et al. (1951) Skrjabin et al. (1951) Cram (1927) Cram (1927) Skrjabin et al. (1951) Toro et al. (1999) Baruˇs (1966a) Wajihullali et al. (1982) Wehr and Hwang (1964) Yamaguti (1961) Skrjabin et al. (1951) Cram (1927) Yamaguti (1961) Cram (1927)
Ascaridia columbae Ascaridia columbae Ascaridia columbae
NR NR North America
Ascaridia columbae
India (zoo)
Cram (1927) Skrjabin et al. (1951) Conti and Forrester (1981) Baylis and Daubney (1922)
Ascaridia platyceri
Canada Germany
Ascaridia columbae Ascaridia columbae Ascaridia geei Ascaridia columbae Ascaridia columbae Ascaridia maculosa Ascaridia columbae
Ducorps’ co*ckatoo (Cacatua ducorpsii) Ascaridia platyceri Yellow-crested co*ckatoo (Cacatua Ascaridia sulphurea) hermaphrodita Ascaridia platyceri Psittacidae Alexandra’s Parrot (Polytelis alexandrae)
Location
Ascaridia columbae
Ascaridia platyceri
398
Webster (1982) Hartwich and Tscherner (1979) Czech Republic Kajerova et al. (2004b) South America Cram (1927) Czech Republic Kajerova et al. (2004b) Australia, Brazil Kajerova et al. (2004a) (citing Johnston and Mawson 1941 and Ferrola et al. 1976) Australia Mines (1979)
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Table 23.2. (Continued ) Host Austral Parakeet (Enicognathus ferrugineus) Australian King-Parrot (Alisterus scapularis) Black-billed Parrot (Amazona agilis)
Ascaridia sp. Ascaridia platyceri Ascaridia columbae Ascaridia platyceri Ascaridia hermaphrodita Ascaridia platyceri
Black-winged Lovebird (Agap*rnis taranta) Blue-and-yellow Macaw (Ara ararauna) Ascaridia hermaphrodita Bluebonnet (Northiella haematogaster) Ascaridia platyceri Blue-crowned Parakeet (Aratinga Ascaridia acuticaudata) hermaphrodita Blue-fronted Parrot (Amazona aestiva) Ascaridia hermaphrodita Blue-headed Parrot (Pionus menstruus) Ascaridia hermaphrodita Bourke’s Parrot (Neophema bourkii) Ascaridia columbae Ascaridia platyceri Brown-throated Parakeet (Aratinga pertinax) Budgerigar (Melopsittacus undulatus)
Ascaridia hermaphrodita Ascaridia columbae Ascaridia platyceri
Caatinga Parakeet (Aratinga cactorum)
Ascaridia hermaphrodita Ascaridia platyceri
Crimson Rosella (Platycercus elegans) Cuban Parrot (Amazona leucocephala) Dusky Parrot (Pionus fuscus) Eastern Rosella (Platycercus eximius)
Elegant Parrot (Neophema elegans) Festive Parrot (Amazona festiva) Fischer’s Lovebird (Agap*rnis fischeri) Golden-winged Parakeet (Brotogeris chrysoptera) Green-cheeked Parakeet (Pyrrhura molinae) Green-rumped Parrotlet (Forpus passerinus) Hispaniolan Parakeet (Aratinga chloroptera)
Ascaridia hermaphrodita Ascaridia platyceri Ascaridia hermaphrodita Ascaridia galli Ascaridia platyceri Ascaridia galli Ascaridia platyceri Ascaridia hermaphrodita Ascaridia platyceri
Location
Reference
Germany
Hartwich and Tscherner (1979) Australia Kajerova et al. (2004a) Czech Republic Kajerova et al. (2004b) South America Cram (1927) Germany Brazil
Hartwich and Tscherner (1979) Skrjabin et al. (1951)
Australia South America
Mines (1979) Kajerova et al. (2004a)
South America
Cram (1927)
South America
Kajerova et al. (2004a)
Australia Australia Germany South America
Kajerova et al. (2004a) Mines (1979) Hartwich and Tscherner (1979) Kajerova et al. (2004a)
Australia, Brazil Canada Czech Republic Brazil
Kajerova et al. (2004a) Webster (1982) Kajerova et al. (2004b) Skrjabin et al. (1951)
Czech Republic Kajerova et al. (2004b) Germany Hartwich and Tscherner (1979) South America Cram (1927) Czech Republic Kajerova et al. (2004b) Brazil Skrjabin et al. (1951) United Kingdom Peirce and Bevan (1973) Czech Republic Kajerova et al. (2004b) Germany Hartwich and Tscherner (1979) England Peirce and Bevan (1973) Australia Mines (1979) South America Cram (1927)
Czech Republic Kajerova et al. (2004a, b) (origin Africa) Ascaridia sergiomeirai Brazil Kajerova et al. (2004a) Ascaridia Brazil hermaphrodita Ascaridia sergiomeirai Brazil
Skrjabin et al. (1951)
Brazil
Skrjabin et al. (1951)
Ascaridia hermaphrodita
399
Yamaguti (1961)
(continues)
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Table 23.2. (Continued ) Host Hyacinth Macaw (Anodorhynchus hyacinthinus) Jandaya Parakeet (Aratinga jandaya) Long-tailed Parakeet (Psittacula longicauda) Mallee Ringneck (Barnardius barnardi) Mealy Parrot (Amazona farinose) Orange-winged Parrot (Amazona amazonica) Port Lincoln Parrot (Barnardius zonarius) Puerto Rican Parrot (Amazona vittata)
Ascaridia sp.
Ascaridia Germany (zoo) hermaphrodita Ascaridia platyceri Czech Republic Ascaridia nicobarensis Great Nicobar Island (India) Ascaridia platyceri Australia Ascaridia Brazil hermaphrodita Ascaridia USA (zoo) hermaphrodita Ascaridia ornata Brazil Ascaridia platyceri Czech Republic
Ascaridia hermaphrodita Red-and-green Macaw (Ara Ascaridia chloropterus) hermaphrodita Red-lored Parrot (Amazona autumnalis) Ascaridia hermaphrodita Red-necked Parrot (Amazona arausiaca) Ascaridia hermaphrodita Red-rumped Parrot (Psephotus Ascaridia platyceri haematonotus) Red-winged Parrot (Aprosmictus erythropterus) Regent Parrot (Polytelis anthopeplus)
Location
Ascaridia platyceri
Reference Hartwich and Tscherner (1979) Kajerova et al. (2004b) Soota et al. (1971) Mines (1979) Skrjabin et al. (1951) Canavan (1931) Kajerova et al. (2004a) Kajerova et al. (2004b)
South America
Kajerova et al. (2004a)
Argentina
Mart´ınez et al. (2003)
Nicaragua
Schmidt and Neiland (1973) Kajerova et al. (2004a)
South America Australia Germany
Mines (1979) Hartwich and Tscherner (1979) Czech Republic Kajerova et al. (2004b) Australia Mines (1979)
Ascaridia platyceri
Australia Germany
Rose-ringed Parakeet (Psittacula krameri)
Ascaridia hermaphrodita Ascaridia platyceri
Rosy-faced Lovebird (Agap*rnis roseicollis) Sapphire-rumped Parrotlet (Touit purpuratus) Scaly-headed Parrot (Pionus maximiliani)
Ascaridia platyceri
USA (zoo, origin Brazil) Czech Republic Kajerova et al. (2004a, b) (origin Africa) Czech Republic Kajerova et al. (2004a, b) (origin Africa) Russia Kajerova et al. (2004a)
Scarlet Macaw (Ara macao) Scarlet-chested Parrot (Neophema splendida) Sun Parakeet (Aratinga solstitialis)
Ascaridia hermaphrodita Ascaridia hermaphrodita Ascaridia sergiomeirai Ascaridia hermaphrodita Ascaridia platyceri
Ascaridia hermaphrodita Tui Parakeet (Brotogeris sanctithomae) Ascaridia sergiomeirai Turquoise Parrot (Neophema pulchella) Ascaridia platyceri Vinaceous Parrot (Amazona vinacea) Ascaridia hermaphrodita White-eared Parakeet (Pyrrhura Ascaridia leucotis) hermaphrodita
400
Mines (1979) Hartwich and Tscherner (1979) Canavan (1931)
Brazil
Skrjabin et al. (1951)
Brazil Brazil
Pinto et al. (1993) Skrjabin et al. (1951)
Australia Mines (1979) Czech Republic Kajerova et al. (2004b) South America Cram (1927) Brazil Australia South America
Skrjabin et al. (1951) Mines (1979) Cram (1927)
South America
Kajerova et al. (2004a) (continues)
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401
Heterakis and Ascaridia Table 23.2. (Continued ) Host
Ascaridia sp.
Location
White-eyed Parakeet (Aratinga leucophthalma)
Ascaridia Brazil hermaphrodita Ascaridia sergiomeirai Brazil Yellow-collared Lovebird (Agap*rnis Ascaridia platyceri Czech Republic personatus) New Zealand Yellow-crowned Parrot (Amazona Ascaridia South America ochrocephala) hermaphrodita Yellow-fronted Parakeet (Cyanoramphus Ascaridia platyceri Germany auriceps) CUCULIFORMES Cuculidae Common Cuckoo (Cuculus canorus) Greater Coucal (Centropus sinensis)
Reference Skrjabin et al. (1951) Pinto et al. (1993) Kajerova et al. (2004b) Weeks (1981) Cram (1927) Hartwich and Tscherner (1979)
Russia Thailand Asia
Skrjabin et al. (1951) Skrjabin et al. (1951) Cram (1927)
STRIGIFORMES Strigidae Eurasian Eagle-Owl (Bubo bubo) Ascaridia galli New Britain Hawk-Owl (Ninox odiosa) Ascaridia australis
NR Australia
McDonald (1969) Skrjabin et al. (1951)
CAPRIMULGIFORMES Caprimulgidae Nacunda Nighthawk (Podager nacunda) Ascaridia amblimoria
Brazil
Skrjabin et al. (1951)
Ascaridia galli
NR
McDonald (1969)
Ascaridia galli
NR
McDonald (1969)
Ascaridia galli
NR
Yamaguti (1961)
PASSERIFORMES Emberizidae Yellowhammer (Emberiza citronella) Passeridae House Sparrow (Passer domesticus) Turdidae Mistle Thrush (Turdus viscivorus)
Ascaridia cuculina Ascaridia circularis Ascaridia trilabium
Note: Some host–parasite reports found only in the review literature did not include geographic location and often included hosts from wild, domestic, and private and public zoological collections; where possible, hosts from zoos and host origin of capture are noted. NR, not reported. Ascaridia are large (16–120 mm; males typically smaller than females), opaque white-colored worms (Figure 23.1) found in the intestinal tract (Cram 1927; Ruff 1984). Like other members of their subfamily, ascarid nematodes have lips that lack an interlabia, a long club-shaped esophagus without a posterior bulb, cuticular lateral flanges (may be absent), and lack a ventriculus and ceca (Skrjabin et al. 1951; Yamaguti 1961; Yorke and Maplestone 1962). EPIZOOTIOLOGY Life History Most life history information about heterakid and ascarid nematodes is based on studies of species that occur in economically important host species such as pigeons and poultry. These include A. columbae, As-
caridia dissimilis, A. galli, Ascaridia numidae, H. gallinarum, and H. isolonche. In general, gravid females release unembryonated eggs into the intestinal tract of the host and the eggs are subsequently voided with the feces (Ruff 1984; Anderson 2000). Embryonation occurs after eggs exit the host, and transmission is complete when a susceptible host ingests the embryonated egg (Anderson 2000). Embryonation is dependent on environmental conditions. Optimal development of the eggs of H. gallinarum occurs at 18–33◦ C and eggs take from 7 to 17 days to become infective (Anderson 2000). By contrast, eggs of A. galli embryonate in about 5–14 days at 22–34◦ C (Anderson 2000). At least two species, H. gallinarum and A. galli, use earthworms as transport hosts. Larvae of both nematode species have been found in Lumbricus terrestris, Allolobophora caliginosa, and Eisenia foetida,
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Parasitic Diseases of Wild Birds
(a)
(b)
Figure 23.1. (a) Heterakis and Ascaridia (b). Courtesy of R. Davidson, Southeastern Cooperative Wildlife Disease Study; originally published in Field Manual of Wildlife Diseases in the Southeastern United States, 2nd ed. Southeastern Cooperative Wildlife Disease Study, Athens, GA.
but only larvae of H. gallinarum appear to accumulate (Lund et al. 1966; Augustine and Lund 1974). Eggs hatch after they are ingested by the earthworm and transmission occurs when the earthworm is itself eaten by the definitive host (Lund et al. 1966; Augustine and Lund 1974). Eggs and larvae of A. galli are retained briefly by earthworms before being expelled into the soil (Augustine and Lund 1974). Other transport hosts have been documented for H. gallinarum, including sow bugs (Porcellio scaber) (Spindler 1967) and grasshoppers (Bruneria brunnea, Camnula pellucida, and Melanoplus spp.) (Frank 1953). Flies (Musca domestica and Lucilia sp.) can also serve as mechanical carriers of the eggs of H. gallinarum (Frank 1953). In chickens, eggs of H. gallinarum hatch in the upper intestinal tract and larvae migrate to the ceca within 24 h (Ruff 1984). Here the larvae occur in close associ-
ation with the cecal mucosa and may invade the crypts of the mucosa (Vatne and Hansen 1965). Peak association with cecal tissue is about 3 days and by day 12, most larvae have migrated to the distal third of the cecal lumen (Vatne and Hansen 1965) where they develop into adults. Females mature in about 24–36 days (Levine 1968). For H. isolonche and possibly other species, larvae penetrate the cecal mucosa where they develop into adults (Schwartz 1924). They are often found encapsulated on the cecal wall (Schwartz 1924; Callinan 1987; Balaguer et al. 1992). The eggs of A. galli hatch in the proventriculus and upper intestinal tract and larvae live in the lumen of the duodenum and jejunum for about 9 days (Ackert 1931; Ruff 1984). They subsequently invade the spaces between the intestinal villi and penetrate the mucosa (Ackert 1931). Larvae live in the intestinal mucosa for
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Heterakis and Ascaridia about 17 days before they move into the intestinal lumen to mature (Ackert 1931). Maturation of A. columbae takes about 35–40 days in Rock Pigeons (Columba livia) (Wehr and Hwang 1964). By contrast, maturation times for A. galli and A. dissimilis range from 28 to 30 days (Wehr 1942; Ruff 1984). Ectopic migration has been reported for several species of ascarid nematodes including A. columbae in Rock Pigeons (Wehr and Shalkop 1963; Wajihullali et al. 1982), A. dissimilis in domestic turkeys (Norton et al. 1999), and Ascaridia perspicillum (=Ascaridia galli) in Indian Peafowl (Pavo cristatus) (Rao et al. 1981). Larvae of A. columbae have been found in the liver, intestinal mesenteries, lungs, and gizzard lining (Wehr and Hwang 1964; Wajihullali et al. 1982), whereas larvae of A. dissimilis have been found in the liver, bile duct, and portal veins (Norton et al. 1999). Larval migration may be the result of overcrowding in the intestines (Wajihullali et al. 1982). Larvae of A. columbae and A. dissimilis fail to develop in these tissues, suggesting this is not a normal pathway for larval development (Wehr and Hwang 1964; Wajihullali et al. 1982; Norton et al. 1999). However, several adult A. perspicillum (=Ascaridia galli) have been found in the bile duct of an Indian Peafowl chick (Rao et al. 1981), which suggests they either migrated as adults from the intestinal tract or reached the duct as larvae and subsequently matured. Eggs subjected to normal environmental conditions can remain viable for extended periods. Farr (1956, 1961) reported that the eggs of H. gallinarum remained infective in soil at a Maryland, USA, outdoor test facility for up to 229 weeks (4.4 years) while eggs of Ascaridia spp. (likely A. galli and/or A. dissimilis) remained viable for up to 136 weeks (2.6 years). A temperature of 57◦ C was lethal to eggs of H. gallinarum, although eggs of A. galli survived slightly higher temperatures (Christenson et al. 1942). Eggs of Ascaridia lineata (=Ascaridia galli) were killed when exposed to a temperature of 43◦ C for 12 h or a temperature of about −12◦ C for 22 h (Ackert 1931). Prevalence and Intensity The prevalence and intensity of infections with Heterakis and Ascaridia vary widely, ranging from 0 to 100% prevalence and intensities of up to thousands of worms within a single host. Differences in intrinsic factors such as host resistance, immunity, co-occurring susceptible host species, behavior, diet, age, and sex and extrinsic environmental factors such as season, precipitation, and geographic location may explain this variability. For example, prevalence and intensity of infection of H. gallinarum in Ring-necked
(a)
(b)
Figure 23.2. Heterakis gallinarum head (a) and tail (b). Modified from Ruff (1984) Diseases of Poultry, 8th ed., after Skrjabin and Shikhobalova (1949) and Lane (1917), respectively, and reproduced with permission from Blackwell Publishing.
Pheasants have ranged from 78 to 100% and from 1 to 2,116 worms, respectively, at a hunting estate in Great Britain (Draycott et al. 2000). By contrast, prevalence and intensity in Ring-necked Pheasants in the upper Midwest US have been much lower (Gilbertson and Hugghins 1964; Greiner 1972). Some studies have shown a host age effect. For example, higher prevalence of H. gallinarum was found in juvenile Greater Prairie-Chickens (Tympanuchus cupido) than in adults (Harper et al. 1967). Prevalence of infection in other host species may be higher in adult birds. Examples include H. gallinarum in Ringnecked Pheasants (Gilbertson and Hugghins 1964), Heterakis bonasae (=Heterakis isolonche) in Northern Bobwhites (Blakeney and Dimmick 1971), A. columbae in White-winged Doves (Zenaida asiatica) (Glass et al. 2002), and Ascaridia compar in Willow Ptarmigan (Lagopus lagopus) (Wissler and Halvorsen 1977).
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Prevalence and intensity of infections may vary between host sexes. For example, prevalence of H. gallinarum was higher in male Greater Prairie-Chickens (Harper et al. 1967), whereas prevalence of H. bonasae (=Heterakis isolonche) was similar between male and female Ruffed Grouse, but mean intensity was higher in females (Kalla et al. 1997). Others have reported no relationship between host sex and infections of H. isolonche in Northern Bobwhites (Moore et al. 1987) or of A. columbae in White-winged Doves (Glass et al. 2002). The effect of season has been examined in some detail. Prevalence and intensity of H. gallinarum was higher in spring than fall in Wild Turkeys (McJunkin et al. 2003). Prevalence was similar among seasons for H. bonsae (=Heterakis isolonche) in Ruffed Grouse, but mean intensity was highest in February and lowest in October–December (Kalla et al. 1997). Prevalence of A. compar in Willow Ptarmigan and Eurasian Capercaillie (Tetrao urogallus) was highest in summer and fall and lowest in winter (Lund 1946, reporting data from H¨ulphers 1930; Wissler and Halvorsen 1977). CLINICAL SIGNS Signs of heterakiasis and ascariasis in wild birds have not been adequately described, probably because moribund individuals are more susceptible to predation and thus are rarely found. Nonspecific signs of heterakiasis in captive pheasants include emaciation, weakness, diarrhea, dyspnea, and terminal gasping (Helmboldt and Wyand 1972; Balaguer et al. 1992). Nonspecific clinical signs described as unthriftiness have also been observed in Rock Pigeons that had heavy infections of A. columbae (Wehr and Shalkop 1963). PATHOLOGY Little information is available regarding pathological responses of wild birds to heterakid and ascarid nematodes. Much of our understanding about pathological responses comes mainly from poultry and captive wild birds, particularly pheasants, which seem highly susceptible to several species of Heterakis. The ceca of Ring-necked Pheasants infected with H. gallinarum may exhibit congestion, thickening and petechiation of the mucosa, intussusception, and cecal abscesses (Menezes et al. 2003). Nodular typhlitis has been reported in pheasants infected with H. gallinarum (Menezes et al. 2003) and H. isolonche (Schwartz 1924; Griner et al. 1977; Callinan 1987; Balaguer et al. 1992). Nodules appear pale white or pink to dark brown and are approximately 1–8 mm in size (Griner et al. 1977; Menezes et al. 2003). A host inflammatory response is evident during early nodule
development when heterakid nematodes become surrounded by lymphocytes, macrophages, and fibroblasts (Callinan 1987; Balaguer et al. 1992). Older nodules containing degenerated heterakids have a preponderance of epithelioid cells, plasma cells, and giant cells (Balaguer et al. 1992). A pathological response to ascarid nematodes has been reported for chickens (Ackert 1923) and Rock Pigeons (Wajihullali et al. 1982) and is likely similar in susceptible wild hosts. Ascarid nematodes irritate the mucosal lining of the intestine and can produce edema, hyperemia, hemorrhage and destruction of villi, dilation of the crypts of Lieberk¨uhn, and infiltration by eosinophils and lymphoid cells (Ackert 1923; see reviews in Mozgovoi 1953). High intensities of adult A. numidae in the lumen are associated with localized excess mucus formation indicative of mild catarrhal enteritis (Matta and Ahluwalia 1979). Acute catarrhal enteritis caused by infections with A. perspicillum (=Aseradia galli) has been reported in Indian Peafowl (Rao et al. 1981). Intestinal obstruction was reported in a Chukar (Alectoris chukar), caused by over 260 worms of A. galli (Tibbitts and Babero 1969). Intestinal dilation and obstruction have been observed in heavy infections of A. columbae in Rock Pigeons (Wajihullali et al. 1982). Several species of ascarids have been known to cause nodulation in susceptible hosts. Small pinhead sized nodules have been found on the intestinal wall surrounding A. columbae in Rock Pigeons (Wajihullali et al. 1982). Larvae of A. numidae in Helmeted Guineafowl are surrounded by macrophages, some eosinophils and giant cells, and are contained within collagen fiber capsules (Matta and Ahluwalia 1979). The host response at sites of ectopic migration varies widely. No pathological effects are associated with larvae of A. columbae within the gizzard lining of Rock Pigeons, while intensive cellular infiltration of leukocytes may occur around ascarid larvae in tissues of the intestine, mesenteries, liver, and lung (Wajihullali et al. 1982). Development of granulomatous lesions in the liver of Rock Pigeons infected with A. columbae initially involves infiltration of host lymphoid cells and some eosinophils around degenerating larvae, followed by an increase in eosinophils, and finally engulfment of the larval debris by giant cells (Wehr and Shalkop 1963). Ascarid-induced hepatic lesions and purulent cholangitis and thickening of bile duct walls resulting from infiltration of lymphocytes, plasma cells, and heterophils have been reported in Indian Peafowl (Rao et al. 1981). Severe catarrhal inflammation accompanied with hyperplasia of mucosal epithelial cells can occur at affected sites in the proventriculus (Rao et al. 1981).
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Heterakis and Ascaridia DIAGNOSIS Definitive diagnosis is by direct observation and identification of adult worms. Identification of species of Heterakis usually requires examination of male features because females in this genus are often indistinguishable (Maplestone 1932; Madsen 1950). Identification of species of Ascaridia usually requires examination of male spicules, determination of number of caudal papilla, and other distinct morphological characteristics that are described in taxonomic keys or original species descriptions. Eggs of H. gallinarum and A. galli can be distinguished from one another on the basis of size and shape (Christenson et al. 1942).
IMMUNITY Little is known about immunity to Heterakis and Ascaridia in wild birds. While some species can cause morbidity and mortality in some hosts, few or no pathological responses are seen in others. For example, H. isolonche is pathogenic in pheasants, but is relatively benign in Northern Bobwhites. Similarly, while infections with H. gallinarum can be acquired by sympatric Ring-necked Pheasants and Gray Partridges (Perdix perdix), substantially higher worm intensities occur in pheasants (Madsen 1941). Whether these findings in pheasants and partridges are the result of innate or acquired immunity or simply host species differences in exposure probabilities to infective parasite stages has yet to be determined. There is speculation that higher prevalence and intensity of infection by certain species of helminths in juvenile birds may be the result, in part, of the bird’s immunological naivety (Buscher 1965; Fedynich et al. 2005). However, this may not be the case for heterakids and ascarids because numerous studies have shown higher prevalence and/or intensity of infection in adults (see Prevalence and Intensity section). Unfortunately, data from wild birds regarding immunological responses caused by heterakids and ascarids are lacking. Study of this topic is particularly problematic because immunological responses can vary between wild and domestic individuals of the same species and birds can only be reliably grouped into two (juvenile and adult) or possibly three age classes (juvenile, subadult, and adult).
PUBLIC AND DOMESTIC ANIMAL HEALTH CONCERNS Human exposure to avian heterakid or ascarid nematodes does not appear to pose a health risk. However,
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there is potential for transmission between domestic and wild birds. Wild birds may serve as sources of infection for poultry flocks, and domestic birds may aide in maintaining cycles of transmission in wild birds or possibly introduce new species to wild populations (Wehr 1940a, 1942; Maxfield et al. 1963). Ascarid nematodes that negatively affect domesticated birds raised for commercial purposes include A. dissimilis in turkeys (Wehr 1942; Norton et al. 1992), A. galli in chickens (Ruff 1984) and Helmeted Guineafowl (Ayeni et al. 1983), and A. numidae in Helmeted Guineafowl (Matta and Ahluwalia 1979). Of these, A. dissimilis causes significant concern in North America because hepatic lesions caused by this ascarid worm can lead to condemnation of affected turkeys (Norton et al. 1999) and domestic flocks can experience high mortality (Norton et al. 1992). Heterakis gallinarum is also a potential threat to domestic and wild Galliformes because it can transmit Histomonas meleagridis, which is the protozoan that causes histomoniasis in domestic and Wild Turkeys (Cole and Friend 1999; Forrester and Spalding 2003; Chapter 7). WILDLIFE POPULATION IMPACTS Our early understanding of Heterakis and Ascaridia infections was derived primarily from observations and experimental studies of individual hosts. Populationlevel impacts have been more difficult to assess, particularly regarding the sublethal effects of these nematodes on predation rates, host competition, and host fitness. Our knowledge of these factors is improving. Recent studies have found negative relationships between A. compar and body weight in juvenile Willow Ptarmigan (Daehlen 2003) and indications that parasite assemblages (including A. compar, Heterakis altacia, H. gallinarum, and Heterakis tencicauda) may influence population cycles of the Rock Partridge (Alectoris graeca) (Rizzoli et al. 1999). Significant negative impacts on wild bird populations can occur when parasitized pen-raised birds are released into the wild. The introduction of new parasite species into native populations is a concern. This may have occurred when 27 Whooping Cranes (Grus americana) and a Sandhill Crane (Grus canadensis) apparently infected with Ascaridia pterophora were released in Florida (Spalding et al. 1996). The only previous record of this ascarid worm was from the Red-legged Seriema (Cariama cristata) in Brazil (Cram 1927). Introduced birds can also help maintain and perpetuate heterakid or ascarid nematodes already present in wild populations. Negative effects of infection with H. gallinarum at the population level have been observed
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in released stocks of pen-reared Ring-necked Pheasants in Britain, including lower host body condition (Robertson and Hillgarth 1993) and lower reproduction and survival than wild pheasants (Woodburn 1994). In Spain, there is a concern that populations of the Redlegged Partridge (Alectoris rufa) may be negatively affected by infected pen-reared stock released into the wild to support recreational hunting activities (Mill´an et al. 2004a; Villan´ua et al. 2007). Parasites may play important roles in facilitating or inhibiting host species invasions and may have significant negative effects on native host species when unintentional introductions of exotic parasites occur (Prenter et al. 2004). Several studies conducted in the UK suggest that maintenance of H. gallinarum in populations of the introduced Ring-necked Pheasant has negatively impacted the native Gray Partridge (Tompkins et al. 2000, 2001, 2002). Heterakid and ascarid nematodes can also be spread to new geographic regions via the pet trade (Webster 1982), via acquisition of exotic birds for zoological gardens (Tables 23.1 and 23.2), via human introductions of exotic birds into native ecosystems, and from natural range expansions of infected hosts. The magnitude of this problem is evident in the current worldwide distribution of psittacine birds and their ascarid parasites as well as the number of infected bird species that occur outside their natural geographic ranges (Tables 23.1 and 23.2).
TREATMENT AND CONTROL Wildlife management agencies have not actively pursued preventative treatment and control activities for heterakid and ascarid nematodes in wild bird populations, likely because of practicality and cost. It is unclear whether broad-scale treatment and control activities have any long-term benefits to wild bird populations. However, for captive breeding programs involving game birds and threatened or endangered species, treatment and control programs are essential, and they typically follow procedures used for poultry. Treatment and control measures are designed to disrupt the transmission cycle of the heterakids and ascarids. Recommendations for captive birds include routinely disinfecting feeders and watering troughs, and raising birds on hardware cloth to prevent birds from contacting contaminated soil and feces (Mozgovoi 1953; Levine 1968). Prevention of cross infection between domestic and wild Galliformes is an important consideration in captive situations. For example, Greater Prairie-Chickens and Sharp-tailed Grouse (Tympanuchus phasianellus) should be kept from areas used by domestic poultry
(Morgan and Hamerstrom 1941). Such recommendations serve to prevent situations such as that noted by Tibbitts and Babero (1969) where Chukars likely became infected by A. galli when they were housed in pens frequented by chickens. Additionally, plans for restocking of wild birds should consider proximity to commercial enterprises that have the same or closely related species. For example, plans for restocking Wild Turkeys should take into consideration the proximity of turkey farms to minimize the potential of cross infection (Maxfield et al. 1963). Anthelmintics have been used extensively on captive flocks of chickens and turkeys and they should be effective for wild or captive wild species. The U.S. Food and Drug Administration approves specific anthelmintics that are used in birds intended for human consumption (i.e., eggs or meat) and currently authorizes fenbendazole, phenothiazine, and hygromycin B for poultry (Kahn 2006). Other drugs include tetramisole and levamisole for controlling H. gallinarum and A. galli (Kahn 2006) and piperazine dihydrochloride for controlling A. galli (Nilsson and Alderin 1988). Additionally, levamisole has been used to control A. columbae in captive flocks of Rock Pigeons (Panigrahy et al. 1982). Other possible anthelmintics for birds may include albendazole, oxfendazole, and ivermectin (Dawe and Hofacre 2002). Ivermectin is effective in eliminating H. gallinarum and A. galli from flocks of Helmeted Guineafowl (Okaeme 1988). However, albendazole and fenbendazole can cause toxicosis when administered to captive columbids (Howard et al. 2002). One study found that albendazole (at the dose administered) was not satisfactory in substantially reducing H. gallinarum in farm-reared Red-legged Partridges that were intended for release into the wild (Villan´ua et al. 2007). There has been success in targeted field applications of anthelmintics. Several studies conducted on hunting estates in Britain have used supplemental feed treated with levamisole hydrochloride (Robertson and Hillgarth 1993; Draycott et al. 2000) and flubendazole (Woodburn et al. 2002) to reduce infections of H. gallinarum, thereby increasing breeding success in free-ranging Ring-necked Pheasants.
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24 Ascaridoid Nematodes: Contracaecum, Porrocaecum, and Baylisascaris Hans-Peter fa*gerholm and Robin M. Overstreet ity of an ascaridoid infection. As noted below, the diseases caused by ascaridoid parasites have been referred to by a number of names depending on the species discussed. Recent molecular studies on Contracaecum have regularly reported the presence of sibling species within individual morpho-species. For example, there are at least five sibling species in the Contracaecum osculatum complex from pennipeds (Bullini et al. 1997). This similarity among species makes identification of strains and species difficult without using molecular techniques (Nadler and Hudspeth 1998; D’Amelio et al. 2007; Mattiucci et al. 2008). Molecular studies should be used to determine the correct geographic distribution and to investigate the ecology of these and other sibling forms. Many may represent new species or strains and their potential role as pathogens in avian hosts remains unknown.
INTRODUCTION The superfamily Ascaridoidea is a medically and economically important entity made up of some 50 genera covering six families (fa*gerholm 1991; Nadler and Hudspeth 1998, 2000). Adult members of few ascaridoid genera infect birds (fa*gerholm 1996), and we treat in this chapter only the anisakid Contracaecum Railliet and Henry, 1912 (Contracaecinae) and the ascaridid Porrocaecum Railliet and Henry, 1912 (Toxocarinae). There are a few records of species of other anisakids in birds such as Duplicaecum Majumdar and Chakravarty, 1963, Heterotyphlum Spaul, 1927, and the ascaridid Amplicaecum Baylis, 1920. Some may represent accidental infections or incorrect identifications (Deardorff and Overstreet 1981b). With the exception of Anisakis sp. (Riley 1972), none of these species are known to harm their avian hosts or constitute a public health risk (e.g., Majumdar and Chakravarty 1963). We also treat juvenile species of the ascaridid Baylisascaris Sprent, 1968 (Ascaridinae) because of their detrimental influence on both birds and people. Members of these ascaridoid genera should not be confused with those of the related superfamily Heterakoidea, such as Heterakis gallinarum (Schrank 1788) (see Kellogg and Reid 1970; Norton et al. 1992; Menezes et al. 2003; Chapter 23). Members of the genera Contracaecum and Porrocaecum also resemble fish nematodes of the genera Hysterothylacium Ward and Magath, 1916 and Raphidascaris Railliet and Henry, 1915 (Raphidascarididae sensu fa*gerholm, 1991) (Deardorff and Overstreet 1981a, b; Nadler et al. 2007). Although the pathogenicity of ascaridoids of birds has not been studied extensively, numerous case studies show or suggest that the host nutritional and immune status may directly or indirectly affect the sever-
CONTRACAECUM AND PORROCAECUM SYNONYMS Infections with the adult anisakids of the genus Contracaecum have been referred to as nematodiasis, anisakiasis, and contracaeciasis. Infections with species of Porrocaecum are referred to as porrocaeciasis. ETIOLOGY Contracaecum contains over 60 nominal species (e.g., Yamaguti 1935; Hartwich 1964, 1974; Baruˇs et al. 1978) that are medium sized (some 1–8 cm long), threadlike parasites of the proventriculus, gizzard, and intestines of birds, seals, and dolphins. Whereas few synonyms of Contracaecum have been used recently,
413 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Figure 24.1. Structure of Contracaecum variegatum (SEM photographs). (a) Anterior end, subdorsal view, with narrow interlabia, with bifid apex (see enlarged insert). (b) Detail of labium with knoblike lateral auricle. One of the six inner labial sense organs, situated laterally on lips, is located in pit in the center of image. (c) Female tail. A subdorsal left phasmid is located near the tip of the tail. (d) Male tail with a row of subventral proximal papillae. Spicules are extruded from the cloacal orifice. (e) Enlargement of the male tail. Two separate paracloacal papillae and four distal papillae and a phasmid can be seen on the tail. The distribution of the caudel sense organs is close to that of C. magnipapillatum (Fig 24.5). (f) Distal end of a spicule. The extended sharp point is demarcated by the sperm channel, two winglike structures that form a tube. Modified in part from fa*gesholm et al. (1996) with permission of Helminthologia. some fish raphidascaridids now accepted in the genus Hysterothylacium (Deardorff and Overstreet 1981a) are still identified in some nontaxonomic literature as species of Contracaecum (fa*gerholm 1978, 1982, 1988b). Concepts of the phylogeny of ascaridoid nematodes and of the systematic position of the superfamily Ascaridoidea within the order Rhabditida have
been reviewed recently by De Ley and Blaxter (2002) and Nadler et al. (2007). Studies based on genetic features corroborate clades obtained on the basis of structural data (fa*gerholm 1991; Nadler et al. 2000, 2005). Species of Contracaecum have been defined on the basis of distribution of caudal sense organs (papillae) in the male (fa*gerholm 1988a) as well as numerous
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Ascaridoid Nematodes: Contracaecum, Porrocaecum, and Baylisascaris other structural features of the adult worms (Figure 24.1). No fewer than 30 fully grown adult worms of each sex should be studied by light microscopy to make identifications because of intraspecific variability in morphological features. Material for genetic analysis and scanning electron microscopy (SEM) should also be collected. While caudal features generally differentiate species from seals from those that occur in birds, morphological data suggest that hostswitching might have occurred from birds to seals for some species. For example, C. ogmorhini Johnston and Mawson, 1941 sensu lato of ottarid seals has morphological features that are very close to C. rudolphii Hartwich, 1964 sensu lato from birds (fa*gerholm and Gibson 1987), suggesting that this seal parasite
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originated from birds. By contrast, there is little evidence to suggest that species of Contracaecum in birds may have evolved from species that occur in seals. The genus Porrocaecum contains over 30 described species that infect the intestine of birds. Good descriptions of common species are given by Baruˇs et al. (1978) and Anderson (2000). Many of these require revision. By definition, species of Porrocaecum differ from Contracaecum by the absence of a ventricular appendix. They have only small interlabia and they also have a delicate denticulated ridge on the lips that differs from the denticles of Phocascaris, a genus systematically close to Contracaecum (see Berland 1963; Paggi and Bullini 1994) (Figure 24.2). The spicules of male
Figure 24.2. Porrocaecum depressum (SEM photographs) from the stomach of a Northern Goshawk (Accipiter gentilis). (a) Anterior lateral view. (b) Male tail with a subventral row of proximal caudal papillae. These serve as sense organs. (c) Male tail with extruded spicule. The tip of the male tail is modified into a knob. (d) Enlargement of the distal end of the male tail. Note left four distal papillae and phasmid (lowermost dimple, with minute opening).
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specimens of Porrocaecum are usually short (1 mm), while those in male specimens of Contracaecum range from 2 to 12 mm in length. Species of Porrocaecum have been differentiated on the basis of the presence or absence of cervical alae and on the projections of the lip pulp (Osche 1955; Baruˇs et al. 1978; Digiani and Sutton 2001). Host Range and Distribution Species of Contracaecum are generally very common in fish-eating birds and have a wide host range and geographic distribution (Figure 24.3). In some aquatic birds, at least four different species of the genus may occur concurrently in the same individual. Common avian species are Contracaecum magnipapillatum Chapin, 1925 (synonym C. magnicollare Johnston and Mawson, 1941) from the proventriculus of migratory terns such as the Black Noddy (Anous minutus) that range throughout the Pacific Ocean (fa*gerholm et al. 1996). Contracaecum variegatum (Rudolphi, 1809) from the Pacific Loon (Gavia pacifica) and Contracaecum himeu Yamaguti, 1941 from cormorants in Japan were recently redescribed (Nagasawa et al. 1999a, b). Contracaecum microcephalum (Rudolphi, 1809) has been described from all main continents from cormorants (Phalacrocorax spp.), pelicans (Pelecanus spp.), ducks, swans, and geese (Anatidae), and herons, egrets, and bitterns (Ardeidae) (Yamaguti 1935; Hartwich 1964). Many of these records may represent a complex of sister cryptic species similar to those described for Contracaecum multipapillatum (Contracaecum multipapillatum (Von Drasche, 1882)) sensu lato from the proventriculus and esophagus of pelicans, herons, cormorants, storks, and Anhinga (Anhinga anhinga) in North America, including Mexico and Cuba (Deardorff and Overstreet 1980b). Other common avian species include Contracaecum pelagicum Johnston and Mawson, 1942 from the Magellanic Penguin (Spheniscus magellanicus) and numerous other fish-eating birds (fa*gerholm et al. 1996; Garbin et al. 2007). Contracaecum rudolphii is considered to be the most common species of Contracaeacum in birds and has been reported from birds in the Neotropics, Nearctic, Palearctic, Ethiopian, and Australian regions (Johnston and Mawson 1941; Whitfield and Heeg 1977; Baruˇs et al. 1978; Torres et al. 1983; Amato et al. 2006). In the Palearctic Region, C. rudolphii sensu lato has been reported from 58 species of birds in 24 genera (Baruˇs et al. 1978). Also considered a senior synonym of C. spiculigerum by Huizinga (1971) and others (Deardorff and Overstreet 1980b), C. rudolphii occurs in the proventriculus of pelicans, herons, mergansers, and cormorants. Like many other species, numerous indi-
viduals are frequently found embedded in the proventriculus (Thomas 1937). The taxonomic status of C. rudolphii needs to be reconsidered since it comprises a complex of at least four genetically isolated sibling forms that are morphologically similar (Bullini et al. 1986; Cianchi et al. 1992; Mattiucci et al. 1994; Li et al. 2005; D’Amelio et al. 2007; Szostakowska and fa*gerholm 2007). One, Contracaecum septentrionale (Kreis, 1955) has been shown to be an accepted species that occurs mainly at higher latitudes (Li et al. 2005). Similarly, Contracaecum multipapillatum from Florida may comprise at least two genetically isolated sibling types (D’Amelio et al. 2007; H. Ma and R. M. Overstreet, unpublished observations). In Australia and Europe, additional sibling species are present (Cianchi et al. 1992; Mattiucci 2006; S. Shamsi, personal communication). Both Contracaecum magnipaillatum and C. variegatum (Rudolphi, 1809) are structurally similar to C. pelagicum Johnston and Mawson, 1942 (see fa*gerholm et al. 1996). Genetic studies are needed to define further these and other morphologically similar species and their host range and distribution. Species of Porrocaecum are cosmopolitan in distribution, but there is a clear need to review their taxonomic status using both structural and modern genetic methods. Common species include Porrocaecum ensicaudatum (Zeder, 1800), a cosmopolitan parasite of passerine birds (Figure 24.4) and Porrocaecum angusticolle (Molin, 1860) from the intestines of raptors from throughout the world. Porrocaecum depressum (Zeder, 1800) has a similar cosmopolitan distribution and has been reported from approximately 47 species of birds of prey, although there is considerable morphological variation in material collected from owls (Morgan and Schiller 1950). Porrocaecum crassum (Deslongchamps, 1824) occurs in European waterfowl, especially ducks (Figure 24.4). Epizootiology Among species of ascaridoids that have been studied, third-stage (L3) larvae hatch from fully embyronated eggs that are passed in the feces of their avian definitive hosts. These are ingested by a crustacean intermediate host and penetrate into the body cavity of these invertebrates. Fish can act as paratenic hosts when they ingest infected crustaceans. The L3 larvae encyst within the intestinal wall, mesentery, liver, or other internal organs of fish, but are not known to occur within muscle tissue. When fish (or crustaceans) are ingested by a suitable avian host, the L3 larvae are released and undergo final molts to fourth-stage (L4) larvae and adults within the proventriculus of the avian host. Under some conditions, fish may serve as a
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C. multipapillatum (Drasche, 1882) s.l. 37
C. magnipapillatum Chapin, 1925 7
Ardea (1,2) 14%
Morus (3) 14%
Phalacro-corax (1) 8%
Anous (1,3,4) 72%
C. microcephalum (Rud., 1819) 67 Anhinga (1,6) 6%
Other (1,2,3,,4) 25% Podiceps (4,?) 6%
Phalacro-corax (4,5,6) 12%
Tachybaptus (?) 7%
Ardea (1,4,5) 17%
Egretta (1,2,4) 14% Pelecanus (1,4) 8%
Other (1,5) 29%
Nycticorax (1,2) 8% Pandion (1) 11%
Pelecanus (1,5) 17%
C. ovale (Linstow, 1907) 29 Aechmo-phorus (1) 11%
Ardea (4) 3% Aythya (4) 3%
Podilymbus (?) 3% Podiceps (1,4.5) 70%
Nycticorax (1,4,5) 12%
C. micropapillatum (Stossich, 1890) 24
Mergus (4) 3%
C. rudolphii s.l. 125 Other (1,2,4,5,6) 44%
Pelecanus (1,4,5,6) 50%
Phalacro-corax (2,4,5) 21%
Larus (1,2,3,4) 7% Phalacrocorax (1,2,3,4,5,6) 32% Podiceps (2,5,4) 7%
C. septentrionale (Kreis, 1956) 6
Pelecanus (1,2,4) 10%
C. variegatum s.l. 16
Other (1,2,5,6) 37%
Phalacrocorax (1,5) 100%
Uria (1,5) 19%
Gavia (?,4,5) 25%
Larus (5) 19%
C. yamaguti Mawson, 1956 6
C. tricuspe (Geodelst, 1916) 8
Nycticorax (5) 17% Mergus (1) 17 %
Phalacrocorax (6) 25% Ardeola (4) 13%
Egretta (1,2) 8%
Other (1,2) 34%
Diomedea (1) 14%
Anhinga (1,4,6) 62%
Bubulcus (5) 17 %
Uria (4) 17%
Ardea (5) 32%
Figure 24.3. Host range of species of Contracaecum. Diagrams are based on published data on percentage of recorded cases of Contracaecum from different genera of birds. Information was retrieved from Gibson et al. (2005). The category “Other” represents records of different birds only occasionally reported to be infected. Number of cases is underlined in the heading. The origin (geographical area) of records is shown in parenthesis next to the genus name: 1, North America; 2, South America; 3, Australia and New Zealand; 4, Asia (including India and Japan); 5, Europe; 6, Africa. Species recently (re)described include Contracaecum bioccai Mattiucci et al., 2008 (in Pelicanus); Contracaecum eudyptulae Johnston et Mawson, 1942 (in Eudyptula); and Contracaecum pelagicum Johnston et Mawson, 1942 (in Diomedea). Recently defined genetically isolated strains of Contracaecum rudolphii sensu lato are not included.
417
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Parasitic Diseases of Wild Birds
P. crassum (Deslonchamps, 1824) 11
P. reticulatum (Zeder, 1800) 17
Vanellus (5) 9%
Podiceps (?) 9%
Aythya (4) 9%
Otus (4) 9% Numida (5) 9%
Other (2) 12%
Anas (5,6) 28%
Tadorna (3) 9%
Dendrocygna (1) 9%
Nycticorax (2,4) 22% Egretta (2) 17%
Milvus (5) 3%
Aegypius (4) 3%
Gypus (5) 3%
Aquila (5) 3%
Falco (5) 8% Circus (5) 10%
Other (5) 10%
Other (1,3,4,5) 25% Turdus (1,3,4,5) 29%
Lanius (1) 5%
Larus (1,5) 8%
Numenius (5) 5%
Tringa (5) 13% Scolopax (5) 5%
P. ensicaudatum (Zeder, 1800) 64 Charadrius (5) 3%
Charadrius (5) 10%
Limosa (5) 5%
Turdus (5) 13%
Bubo (1,5) 3%
Corvus (4,5) 11%
Calidris (5) 5%
Vanellus (5,6) 13%
Asio (5) 3% Buteo (1,5) 23%
Casmerodius (2) 6%
P. semiteres (Zeder, 1800) 39
Accipiter (1,5) 25%
Tyto (1) 5%
Ardeola (?) 11%
Botaurus (5) 9%
P. depressum (Zeder, 1800) 38 Strix (1) 11%
Ardea (1,2,4) 27%
Pluvialis (5) 8%
P. spirale (Rud., 1795) 8
Garrulus (5) 3% Larus (5) 5% Limosa (5) 3%
Other (5) 50%
Strix (4,5) 50%
Quiscalus (5) Sturnus (1,3,4,5) 5% 16%
Figure 24.4. Host range of species of Porrocaecum. Diagrams are based on published data on percentage of recorded cases of Porrocaecum from different genera of birds. Information was retrieved from Gibson et al. (2005). The category “Other” represents records of different birds only occasionally reported to be infected. Number of cases is underlined in the heading. The origin (geographical area) of records is shown in parenthesis next to the genus name: 1, North America; 2, South America; 3, Australia and New Zealand; 4, Asia (including India and Japan); 5, Europe; 6, Africa. ? indicates absence of locality data. true intermediate host and directly transmit the infection (Huizinga 1966, 1967; Køie and fa*gerholm 1995; Bartlett 1996; Dzieko´nska-Rynko and Rokicki 2007; Szostakowska and fa*gerholm 2007; R. M. Overstreet, unpublished observations). Direct “accidental” transmission of adult worms along with regurgitated food from the parent birds to the chicks is an additional important route for infection (Huizinga 1971; fa*gerholm et al. 1996; Kuiken et al. 1999). Many species of fish-eating birds serve as definitive hosts, with some species of Contracaecum being highly specific to their avian host and others infecting a wide range of hosts (Figure 24.3). Since some worms survive for only about 90 days (Huizinga 1971), the total number depends on continual ingestion of infected
hosts. Because of the high percentage of C. multipapillatum in mixed infections, a primary host prey in Mississippi and Louisiana is presumably mullet (Deardorff and Overstreet 1980b). In northern Europe, very intense infections of C. rudolphii have been observed in the Great Cormorant (Phalacrocorax carbo) (Zuchowska 2000; Kijewska et al. 2002; Szostakowska et al. 2002). Anderson (2000) and Moravec (1994) reviewed the biology of juvenile stages of C. rudolphii sensu lato in fishes. As many as 500 L3 larvae of C. rudolphii have been recorded in a single fish. When intense infections occur in fish, their market value may be reduced for esthetic reasons (Szostakowska and fa*gerholm 2007).
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In contrast to Contracaecum, the life cycle of species of Porrocaecum is terrestrial and includes an earthworm intermediate host. After earthworms ingest larvae from fully developed eggs, they reach the main blood vessels of the oligochaete where they remain. Some species, such as P. depressum and P. angusticolle (Molin, 1860) from birds of prey (Osche 1959), use small mammals as paratenic hosts and the larvae are found encysted in the mesentery and on the intestine and reach their avian definitive hosts when these tissues are consumed (Osche 1955; Erkinaro and Heikura 1977). Among other species, no paratenic host is involved as avian hosts may consume earthworms for food. This is the case for both P. crassum and P. ensicaudatum. The biology of the latter species has been studied by several authors. J¨ogis (1970) and also McNeill and Anderson (1990a, b) found that P. ensicaudatum is encountered only in passerine birds and that mature worms are found naturally only in the avian families Sturnidae and Turdidae (Figure 24.4). Juveniles can occur under the lining of the gizzard and in the wall of the duodenum of the final avian host, but apparently most adults occur both free and embedded in the intestine (Mawson 1956). Diagnosis Species of different ascaridoid genera can usually be identified to genus without difficulty. However, differentiation of species within genera may be challenging because of the often-small structural differences among species (Figure 24.5, Table 24.1). This is true for several species of the genus Contracaecum and is certainly the case for sibling species that cannot be distinguished by structural features alone. In these situations morphological features can be used to show that worms belong to a certain species group, but biochemical methods are needed for a correct identification. One must take into account the process of allometric growth where key morphological features such as the relative location of the vulva and spicule length continue to change until the worm has attained full length (fa*gerholm 1989). The centrid, an asymmetrically oriented pair of lateral papillae in the mid-body region, may also change position as a result of allometric growth (fa*gerholm et al. 1996, 1998). During growth, the length of the spicules increases from the anterior (basal) part of these structures (fa*gerholm 1989), and one can argue that structural features of the point of the tip are the most conserved (Figure 24.1d). One should be careful not to rely too strongly on total spicule length if measurements are not consistent among a considerable number of fully grown worms. With recent advances in molecular methods, we have the opportunity to investigate whether
Figure 24.5. Scanning election micrograph (SEM) of the tail of a male specimen of Contracaecum magnipapillatum from the proventriculus of the Black Noddy (Anous minutus), Heron Island, the Great Barrier Reef, Australia. The distribution of the caudal papillae is an important taxonomic character. The proximal papillae (pr) are located in this species anterior to cloacal orifice as two parallel rows. The paracloacal papillae (p) are located posterior to the cloacal orifice as two large pairs that are not joined. Four pairs of distal papillae (d1, d2, d3, and d4) are located on each side of the tail. One pair of phasmids (ph) is located between the outer lateral pair of distal papillae. The left phasmid (on right in figure) is very close to the posterior most distal papilla. A precloacal median sensory area (m) is also present. The “pts zone” (precloacal tranverse striae zone) represents 25 transverse striae that start at the cloacal orifice and extend anteriorly, covering three papillae in this figure. The “aggregation zone” (agr) covers a straight row of five proximal papillae on one (or both) subventral sides of the worm. In this species, the entire region comprising five proximal papillae is evident on the left subventral side of the worm.
Birds
− + − + + − − + − −
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+ −
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Note: Selected somatic features of taxonomic interest include structures of the anterior end, e.g., labia, and the position of phasmids and centrids. Cervical, labial, and interlabial features are useful in defining some species, e.g., the elaborate structure of C. tricuspe (Geodelst, 1916). The position of phasmids and centrids may also be of importance (fa*gerholm et al. 1998, 2004). Important generic features that are not illustrated include the position of the excretory pore just posterior to the ventral interlabium, a short ventricle and the presence of “contracaeca” (an anteriorly directed intestinal caecum and posteriorly directed ventricular appendix). * In addition to the pattern of distribution, the size (diameter or (length + width)/2) of caudal papillae can be used as a taxonomic criterion. † In addition to the form of the distal end of the spicules, the length of the tip can be used as a taxonomic criterion, provided that differences are statistically significant. Spicule length is often used to separate species. In this case, different growth-related factors as well as intraspecific variability need to be considered.
Distribution of proximal caudal papillae (numerous) Pts zone (region of 25 precloacal striae) 6–17 + − 2–3 − + Aggregation (region of row of 5 precloacal papillae) 6–15 + − 5 − + Number present posterior to cloaca 0 − + 1–21 + − Distribution of paracloacal papillae (2 pairs) Pairs joined + − Pairs separate − + Distribution of distal papillae (4 pairs) Anterior 2 pairs joined − − on both sides Posterior 2 pairs joined − − on both sides Size of defined caudal papillae* Papillae, size Papillae, relative size Length of transverse striae in cloacal region diminutive − − Spicules† Distal end of spicules Points pointed − − Points rounded + + Spicule length Spicule relative length
(a) (b) (c) (d) (e) (f) (g) (h) (i) (j) (k) (l) (m) ContraContraContra- Contra- ContraContraContraContraContraContra- Contra- ContraContracaecum spp. caecum caecum caecum caecum caecum caecum caecum caecum caecum caecum caecum caecum from seals, ogmorhini radiatum bioccai magnipamicromicromultirudolphii pelagicum septent- tricuspe variegatum but not: sensu lato pillatum cephphalum papillatum papillatum sensu lato rionale sensu lato
Seals
Table 24.1. Selected caudal features of diagnostic interest in adult male Contracaecum Railliet and Henry, 1912 from seals (a–c) and birds (d–m) as amended from fa*gerholm (1988a), with new data (c.f. Figure 24.1).
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Ascaridoid Nematodes: Contracaecum, Porrocaecum, and Baylisascaris genetically isolated populations are also morphologically distinct. Other structural features that have been used to distinguish species or groups of species within this genus include features of the cuticle and patterns of caudal papillae (Table 24.1) (fa*gerholm 1988a, 1990). CLINICAL SIGNS Infections with adult ascaridoids that belong to the genera Contracaecum and Porrocaecum usually produce no severe disease or clinical signs in birds. Starvation, however, with its associated signs, may be a contributory cause of death in infections of high intensity. This is frequently observed during necropsy of aquatic birds with Contracaecum sp. (Deardorff and Overstreet 1980b) and wild raptors (kestrels) infected with P. depressum (Keymer et al. 1981). Ruffled feathers and an inability to maintain balance in passerine birds have been attributed to infections with P. ensicaudatum (Arnall and Keymer 1975). PATHOGENESIS AND PATHOLOGY Infections of ascaridoids in the alimentary tract can produce a severe inflammatory response, especially when juvenile worms of some species embed and migrate within the walls of the proventriculus, esophagus, or intestine (Figure 24.6). While documented for Contracaecum, detailed studies of host responses to species of Porrocaecum in birds have not been done. Infections can cause anemia and may lead to actual disease when the bird becomes stressed. Highly pathogenic cases usually involve high-intensity infections in nestlings or young juveniles, starved individuals, birds that are stressed by environmental contamination or other causes, and birds with peritonitis or secondary microbial infections that involve other vital organs. Whereas peritonitis can result in death, the direct pathogenic effect by the worms in most cases is probably low (Deardorff and Overstreet 1980b; Greve et al. 1986). Gross lesions caused by several species of Contracaecum are similar and consist of erosive, often yellow ulcers in the mucosal wall of the proventriculus and adjacent portion of the esophagus. These are associated with both petechial and larger hemorrhages. Both adult and juvenile worms attach or embed in clusters within these small to large ulcers as well as individually in isolated ulcers (Figure 24.7). They also remain free in the lumen. Worms are not always present in ulcers and experimental studies indicate that these are scars from previous worm attachments (fa*gerholm 1988b). Embedded juveniles seem to be the most pathogenic stage of infection to the host. In an 18-year-old cormorant
421
maintained for fishing in Sichuan Province, China, many additional nodules were also in the muscular layer and projected from the serosal surface (Sarashina et al. 1987). Focal mucosal ulcerations containing attached worms occur in some fish, reptiles, and marine mammals infected with other ascaridoid genera (Deardorff and Overstreet 1980a; R. M. Overstreet, personal observations). In some avian cases, especially those with only a few worms, all the individuals are free in the proventriculus or intestine and ulceration is not evident (Huizinga 1971; Deardorff and Overstreet 1980b). Microscopic histopathological alterations caused by Contracaecum include compression of the glandular mucosa and ulceration with inflammatory infiltration. An eosinophilic hyaline cap often surrounds the anterior end of worms at the host–parasite interface, especially around juveniles and young adults. The tissue interface is usually a nodular area that typically includes necrosis, granulomatous inflammation, and an abundance of bacteria. Fibrosis is present and usually more abundant in chicks. The lesion can be extensive in the submucosa and muscularis mucosa. In chicks, the nodule may extend through the muscularis externa and adventitia with associated inflammatory exudates, but in older birds the nodule seldom extends farther than the muscularis interna. Recent infections may exhibit lymphocyte infiltration, and all infections include a variety of inflammatory cells, including lymphocytes, plasma cells, heterophils, macrophages, fibroblasts, and even multinucleated giant cells. The latter often occur associated with degenerating hyaline caps (Huizinga 1971; Liu and Edward 1971; Greve et al. 1986; fa*gerholm et al. 1996; Kuiken et al. 1999). IMMUNITY Relatively little is known about immunity to infections with Contracaecum and Porrocaecum. There is evidence that prevalence and intensity of infection with Contracaecum declines with age, with fewer worms in fledgling pelicans than nestling birds. This may be related to change in diet as birds are exposed to fewer worms when they are old enough to select their own prey. Age immunity may also have contributed to the decline in infections (Humphrey et al. 1978). However, Kuiken et al. (1999) recorded an increase in prevalence of Contracaecum rudolphii sensu lato in Doublecrested Cormorants (Phalacrocorax auritus) to 100% of postnestling chicks and adults. An experimental in vitro study by Raybourne et al. (1983) demonstrated that excretory–secretory (E–S) products of juvenile Anisakis “simplex” and Terranova sp. had a potent inhibitory effect on rodent lymphocyte blast transformation when compared with whole
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extracts of the worms and with controls. This suggests that the worms may have immunosuppressive effects in their avian hosts. PUBLIC HEALTH CONCERNS Avian ascaridoids are generally not thought to be a significant risk to public health. However, ascaridoids from humans are often not identified or are degenerated and cannot be identified. It has been suggested that juvenile worms may be able to acclimate to high temperature or develop to fourth-stage juveniles under some conditions and possibly infect humans. This has been reported for C. multipapillatum in Mayan cichlid fish from Mexico and these worms infected the intestine of an experimental kitten, developed into adults, and produced hemorrhaging ulcers (Vidal-Martinez et al. 1994; R. M. Overstreet, unpublished observations). An accidental infection by a nonfertilized species of Contracaecum was found embedded in the damaged brain of a striped dolphin (Martin et al. 1970). In contrast, specimens reported as Contracaecum sp. in a granulomatous lesion of a dog in Japan (Kitayama et al. 1967) appear to represent a species of Hysterothylacium, which matures in fish (Vidal-Martinez et al. 1994). Other than attention-seeking children, most humans do not commonly eat earthworms, but infections with Porrocaecum spp. from earthworms, other unknown intermediate hosts, or paratenic hosts are possible. Because of the difficulty in identifying species of Porrocaecum in human tissue sections, similar appearing species have been misidentified as Porrocaecum. Examples include parasites from the genus Terranova Leiper and Atkinson, 1914 that mature in elasmobranchs and species of Pseudoterranova that mature in marine pinnipeds (Deardorff et al. 1983). Species of Pseudoterranova commonly infect humans, and juveniles of a Terranova sp. were shown to penetrate into the stomach of a rat, ultimately forming a granuloma (Deardorff et al. 1983).
Figure 24.7. Adult Contracaecum multipapillatum sensu lato from a pelican. Close-up image of worms embedded in the proventricular mucosa (compare with Figure 24.6a). DOMESTIC ANIMAL HEALTH CONCERNS Porrocaecum spp. are a potential risk for domestic birds (Hair and Forrester 1970; Coles 1985), especially when earthworms are present or paratenic hosts have access to infected hosts. However, the apparent host specificity of adult worms may reduce the relative risk of infection to domestic hosts. Both juveniles and adults can harm the host when present in high enough numbers, and these in turn can ultimately contaminate the substratum with eggs that are capable of continuing high rates of infection. Relatively little, however, is known about the risk that wild birds pose to domesticated hosts and appropriate experimental studies are needed to help determine this. Species of Contracaecum can have a major influence on aquatic birds being reared commercially, on display in zoos and other parks, or used in research or recovery. Infections can rapidly accumulate in birds that are confined in small amounts of water with fish that will serve as intermediate hosts.
← Figure 24.6. Proventriculus of a Black Noddy (Anous minutus) infected with Contracaecum magnipapillatum (compare with Figure 24.1). (a) Numerous specimens attached to the proventricular mucosa (sp = spleen) (scale bar = 3.3 mm). (b) Normal histological appearance of proventriculus. (c) Invasion by parasite (right) with an associated inflammatory reaction (arrows indicate border of muscularis). Scale bar = 100 μm. (d) Invasion by parasite (right) with less involvement of the muscularis (cf. previous image), however, with muscularis interna obliterated (arrows). Scale bar = 100 μm. (e) Worm with hyaline cap with inflammatory exudate. Scale bar = 40 μm. (f) Hyaline cap (see star), enlarged (cf. previous image). (g) Partly resorbed hyaline cap. Scale bar = 25 μm. (h) Concentration of inflammatory cells in affected regions. Scale bar = 25 μm. Modified from fa*gerholm et al. (1996) with permission of Helminthologia.
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WILDLIFE POPULATION IMPACTS One should be aware that most infections of ascaridoids of average intensity are not harmful and possibly even beneficial. Because adult ascaridoids often entwine among dietary items in the alimentary tracts of birds as well as of other animals, they may well aid in the digestive process and be advantageous, at least when food is abundant (e.g., Owre 1962; Deardorff and Overstreet 1980a, b; fa*gerholm et al. 1996). Even though individual birds can be heavily infected with ascaridoids, these infections probably have little influence on wildlife populations unless they are compromised by environmental conditions or stress. Intense alimentary tract ascaridoid infections can weaken avian hosts and make them more susceptible to predation, trauma from automobiles, or cause abnormal behavior. Large numbers of Contracaecum spp. in pelicans and other waterbirds have been reported. For example, McOrist (1989) reported peritonitis associated with C. spiculigerum (=Contracaecum rudolphii sensu lato) in 2 Rufous Night-Herons (Nycticorax caledonicus) that may have contributed to their starvation and death. Lesions in the birds included fibrinous hemorrhagic exudates over the intestinal serosa. There are reports of mortality in American White Pelicans (Pelecanus erythrorhynchos) and Brown Pelicans (Pelecanus occidentalis) and Double-crested Cormorants from the southeastern US that are associated with intense infections of hundreds of individuals of C. multipapillatum and other ascaridoids (Contracaecum microcephalum and C. rudolphii sensu lato) (Deardorff and Overstreet 1980b). Oglesby (1960) reported more than 1,100 specimens in a mixed infection from an American White Pelican, and Courtney and Forrester (1974) counted 1,192 in a Brown Pelican. Intensity of infections usually number less than 100 worms, but even birds with intense infections may appear healthy. Fatal ascaridoid infections have also been reported from other avian hosts. Two of 92 Eurasian Kestrels (Falco tinnunculus) that died in the British Isles of apparent starvation had 50–70 large specimens of P. depressum that impacted the duodenum and upper small intestine (Keymer et al. 1981). The birds also had a concurrent infection with the coccidian Isospora buteonis, which may have contributed to the deaths. TREATMENT AND CONTROL Unmanageable infections in wild birds result from biological and environmental conditions that concentrate birds and intermediate hosts together. Little can be done about the infections, but measures can be taken to reduce compounding stress from lead shot, pesticides, and other contaminants. In domestic situations, intermediate and paratenic hosts can be eliminated or reduced. Birds can be medicated, if necessary, for treat-
ment of breeding stock or valuable individuals. Greve et al. (1986) and Grimes et al. (1989) determined that albendazole and fenbendazole were effective against C. multipapillatum in the Brown Pelican and piperazine dihydrochloride, clorsulon, and Curatrem were not. Good results have been reported with a single dose of 1-tetramisole but not with arecoline hydrobromide, thiabendazole, or niclosamide (Courtney et al. 1977). Cooking-infected products destined for dispersal or consumption by humans or other animals will kill all stages of Contracaecum and Porrocaecum, even though only the juveniles in the intermediate or paratenic hosts would be infective to humans. The US Department of Agriculture recommends freezing and storing at −20◦ C or below for 7 days or freezing at −35◦ C or below until solid and then storing at −35◦ C or below for 15 h or at −20◦ C or below for 24 h. Heating at 63◦ C for 17 min will also kill all nematodes as well as bacteria. Control practices for some anisakid infections under normal conditions in the wild can be cost-prohibitive, not practical, or totally unnecessary. Mortality of nestling or young birds may occur in aquatic areas resulting from infections with Contracaecum spp. and the mortalities should be investigated for complicating factors. Porrocaecum spp. can be serious for both wild and domestic birds that have access to earthworms or other intermediate and paratenic hosts. Both juveniles and adults can harm the host when present in high enough numbers, and these in turn ultimately contaminate the substratum with eggs capable of causing infections. For Porrocaecum or Contracaecum, control strategies should first consider eliminating or reducing access to the intermediate hosts. For Contracaecum, hosts reared in an aquaculture setting must be prevented from feeding on infected fish and from defecating in the culture system.
BAYLISACARIS SYNONYMS Depending on their location in the host, infections with the juvenile L3 stage of Baylisascaris are referred to as baylisascariasis, larva migrans, visceral larva migrans (VLM), verminous encephalitis, cerebrospinal (or cerebellar or cerebral) nematodiasis, and nonsuppurative meningoencephalitis. Human infections are usually referred to as eosinophilic meningoencephalitis, VLM, baylisascariasis, or raccoon roundworm encephalitis.
ETIOLOGY The genus Baylisascaris contains as many as nine or more species (Sprent 1968), including Baylisascaris
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Ascaridoid Nematodes: Contracaecum, Porrocaecum, and Baylisascaris procyonis (Stefanski and Zarnowski, 1951) from the raccoon (Procyon lotor) and kinkajou (Potos flavus) in North and South America (Overstreet 1970), Baylisascaris columnaris (Leidy, 1856) from skunks (Mephitidae) in North America (Overstreet 1970), Baylisascaris melis (Gedoelst, 1920) from badgers (Mustelidae), Baylisascaris transfuga (Rudolphi, 1819) from bears and pandas (Ursidae and Procyonidae) throughout the world, and Baylisascaris laevis (Leydy, 1856) from large rodents (Rodentia). The larval stages of virtually all these species pose some risk to birds if eggs or larvae are ingested. HOST RANGE AND DISTRIBUTION The larval stages of species of Baylisascaris have been reported from a wide variety of avian hosts from North America (Figure 24.8). The absence of reports from other parts of the world likely reflects inadequate sampling; however, the infective parasites are primarily North American (Overstreet 1970). Larval stages of Baylisascaris are more commonly reported from ground-feeding species such as doves (Zeinaida spp.), where exposure to eggs is more likely. EPIZOOTIOLOGY Adult species of Baylisascaris occur in the intestines of a specific mammalian definitive host and eggs are passed in the feces. Developed eggs of B. procyonis and B. laevis may infect the final, or definitive host directly. In this case, the L3 (often reported as L2) larvae hatch in the intestine and “migrate to the intestinal mucosa” (Kazacos 2001) where they eventually develop into adults. Another route is taken when developed eggs or hatched larvae are ingested by paratenic hosts, primarily small rodents and birds. In these animals, L3 larvae hatch from the eggs and migrate via the circulatory system to the liver and the lungs and then reach different organs via the heart. The severity of the host tissue reactions depends specifically on the species of paratenic host, the site in the host, and the host immune status. Depending on the location in the body and where they cause adverse signs, larvae cause a disease referred to as visceral larva migrans (VLM), ocular larva migrans (OLM), and neural larva migrans (NLM) (Kazacos 1986; Gavin et al. 2005). When an infected paratenic host is consumed by a mammalian definitive host, L3 larvae develop directly into L4 and later into adult worms in the intestine. The life cycle of different species of Baylisascaris should be studied in more detail and compared with other parasites in the same subfamily, for example, Ascaris spp., to establish similarities in the development of the larvae and migration routes in the final host (Murrell et al. 1997; Geenen et al. 1999; fa*gerholm et al. 2000).
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Birds become involved in the life cycle of Baylisascaris as paratenic hosts when they ingest eggs. Reports of infections with Baylisascaris in wild birds are not abundant. However, infections in birds may be more common than generally recognized, given how long infective eggs can persist in the environment. Some studies have shown that each stool from an average infected raccoon contains from 2 to 10 million eggs, that worms can produce eggs for long periods, and that the eggs of B. procyonis can remain infective under refrigeration for at least 12 years and probably 3–4 years in the wild where they are exposed to summer heat and winter freezing (Kazacos et al. 1982). Large outbreaks of VLM have been reported in domestic birds such as chickens, illustrating the potential of the eggs to infect birds (Richardson et al. 1980). DIAGNOSIS When found in histological sections of avian host tissues, larvae of Baylisascaris may be confused with L3 larvae of several other genera of nematodes that cause larva migrans. They are often not identified to species because they have few differentiating features. Diagnostic features are found on both posterior and anterior ends of larvae and in details of somatic structures that are evident in transverse sections. L3 larvae of B. procyonis are longer (1.27–1.56 mm) (Figure 24.9) than larvae of Toxascaris leonina (Linstov, 1902) Leiper, 1907 (0.65–0.75 mm). Larvae of two other genera that may be confused with Baylisascaris—Lagochilascaris sp. and Hexametra sp.—are longer than larvae of Baylisascaris and measure up to 5 mm in length for Lagochilascaris and up to 10 mm in length for Hexametra sp. (Bowman 1987). Additional distinguishing morphological features have been discussed by Bowman (1987) for Toxocara spp., Porrocaecum, Ophidascaris, Travassosascaris, and Polydelphis. A transverse section of the anterior region of an L3 larva of B. columnaris is illustrated in Figure 24.10. High-quality, well-fixed material and considerable experience are needed to make identifications. Circ*mstantial evidence including geographic data and host history may also aid in the diagnosis. While it is also possible to analyze the ontogeny of the cuticula by SEM to differentiate taxonomic groups (fa*gerholm et al. 2000), this is not routinely done. With recent advances in DNA-based technologies (Gasser 2006; Nadler et al. 2007), identifications based on polymerase chain reaction amplification of ribosomal DNA sequences and internal spacers, ITS-1 and ITS-2, can be done with primers known to function in the Ascaridoidea (Nadler et al. 2000). Mitochondrial genes (mtDNA-cox2) have also been useful (Mattiucci et al. 2008) as well as the use of allozymes (Nascetti et al. 1986).
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Baylisascaris procyonis (Stefanski and Zarnowski, 1951) larva, wild birds, 2 Nyctiocorax (1) 2%
Pipilo (1) 1%
Mimus (1) 9%
Corvus (1) Geococcyx (1) 1% Lanius (1) 3% 1% Carpodacus (1) 9% Callipepla (1) 8%
Passer (1) 3%
Psaltriparus (1) Sturnus (1) 3% 3% Toxostoma (1) 3%
Calidris (1) 1% Anas (1) 2% Turdus (1) 3%
Tyto (1) 1%
Aphelocoma (1) 8%
Zenaida (1) 34%
Baylisascaris procyonis (Stefanski and Zarnowski, 1951) larva, captive birds, 4 Struthio 25%
Ara 50%
Dromaius 25%
Baylisascaris transfuga (Rudolphi, 1819) larva (experimental inf.) 1
Gallus (5) 100%
Figure 24.8. Host range of larvae (larva migrans) of species of Baylisascaris. Diagrams are based on published data on percentage of recorded cases of Baylisascaris in different genera of avian hosts. Information was retrieved from Gibson et al. (2005). The number of records is underlined in heading. The origin (geographical area) of records is shown in parenthesis after the genus name: 1, North America; 2, South America; 3, Australia and New Zealand; 4, Asia (including India and Japan); 5, Europe; 6, Africa.
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Figure 24.9. Live specimen (L3 stage) of Baylisacaris procyonis from the brain of paratenic mammalian vertebrate host.
CLINICAL SIGNS Birds with central nervous system (CNS) infections of larval B. procyonis, B. columnaris, B. melis, and perhaps other species may first exhibit head-tilt or slight stumbling. As the infection progresses, the bird may circle, roll, or fall and eventually become recumbent and comatose before dying (Kazacos 1983). In addition to loss of coordination, torticollis, and paralysis of wings, the feathers are ruffled (Reed et al. 1981). PATHOLOGY There are few studies investigating tissue reactions to NLM in birds. Early work by Tiner (1953a, b) demonstrated that pathogenesis of Baylisascaris in the CNS depends on host species, host size, and migratory behavior of the larval worms. Experimental work with chickens has been done with B. transfuga (Papini et al. 1993) and B. procyonis (Kazacos and Wirtz 1983), but
Figure 24.10. Cross section of Baylisacaris columnaris larva (L3) in brain of a paratenic mammalian host. Lateral alae (with expansion of hypodermis) discernable.
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we still know few details about migratory behavior and pathogenesis of infections in avian hosts. In mammals, host species has an effect on where juvenile L3 migrate. For example, most juvenile B. procyonis and B. columnaris are restricted to encapsulations in the thoracic musculature in rodents, but a few migrate to the brain. In mice, about 5–9% of juvenile B. procyonis enter the brain (R. M. Overstreet, unpublished data). Mice fed 25 eggs begin to have problems with orientation about day 14 postinfection (PI), proceed to drag their rear legs or run continuously in circles, and ultimately die at about day 21–23 PI. A single juvenile in the medulla can kill a mouse. In mice given 175 eggs, loss of coordination begins at 6–11 days PI and death occurs by 8–14 days PI. Mice infected with many more eggs show signs at day 6 PI and die at day 8–9 PI. No mouse recovered after it lost coordination. Juvenile Baylisascaris appear to initially migrate within the cerebral hemispheres for a few days and then migrate to the cerebellum, medulla, and upper spinal cord, at which time disorientation becomes evident. From day 8 to day 10 PI, average length of individual worms increases from about 0.30 to 1.00 mm, causing considerable damage to the brain. The same development presumably takes place in the avian brain. Kazacos (1983) investigated infections in a variety of paratenic hosts and found that 5–15% of the juvenile worms reach the brain of most hosts. A single juvenile worm can be fatal in some avian hosts, including Emu (Dromaius novaehollandiae) (Kazacos et al. 1982) and macaws (Ara arauna, Ara macao, and a hybrid cross) (Armstrong et al. 1989). Gross lesions are not evident in Northern Bobwhite (Colinus virginianus) and other birds killed by B. procyonis (Reed et al. 1981; Kazacos and Wirtz 1983), but microscopic lesions are associated with the migratory tract of the parasites and may occur throughout most areas of the brain. Microscopic lesions include sections through juvenile worms and widely disseminated nonsupportive meningoencephalitis with multifocal areas of malacia, necrosis, and intense inflammatory reaction. Longer-surviving birds exhibit focal granulomas. No alteration was observed in sites other than the brain. Chickens infected experimentally with only 200 eggs exhibited few lesions in the brain (Kazacos and Wirtz 1983). IMMUNITY Little is known about immunity to larval stages of Baylisascaris in avian hosts. On the basis of studies of larval anisakid infections in mammals, hosts are likely to have a strong antibody response, strong blood and tissue eosinophil levels, and a strong T-helper type 2 cell response. The intensity of the response is
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species specific, and those in which the juveniles migrate within the brain seem to have a stronger immune response (Sheppard and Kazacos 1997). PUBLIC HEALTH CONCERNS The primary public health risk from B. procyonis is associated with contact with raccoon latrines and pica/geophagia and not interaction with infected birds or other nondefinitive hosts. DOMESTIC ANIMAL HEALTH CONCERNS Baylisascaris can cause fatal or severe CNS disease in pheasants, chickens, domestic quail, partridges, pigeons, exotic turkeys, Emus, and a wide variety of other species (Kazacos 2001). As is the case with human infections, these risks are associated with contact with raccoon latrines and pica/geophagia and not interaction with infected wild birds or other nondefinitive hosts (Kazacos 1983). WILDLIFE POPULATION IMPACTS Most infections of Baylisascaris in wild birds probably consist of no more than a few juveniles and do not develop into disease. However, juvenile Baylisascaris have been documented in the brains of numerous species of passerines as well as in a few shorebirds, quail, and ducks in studies conducted in California, USA (Evans and Tangredi 1985; Evans 2002). A Barn Owl (Tyto alba) was suspected to have become infected by feeding on an infected California ground squirrel (Spermophilus beecheyi). Abundance of passeriform birds was coincidentally low in the study during a period of time when raccoons were fed and heavily protected, presumably from NLM. Evans (2002) studied 18 species of birds and found a total of 87 birds with B. procyonis-associated NLM or/and VLM. In this study, four areas close to raccoon latrines were inspected for birds with abnormal behavior. Few cases are known where wild birds have been observed with signs of larval migrans, but the actual number of clinical cases in wild birds may be underestimated because sick or moribund birds are more susceptible to predation. In east-central Kansas, a decline in Northern Bobwhite populations occurred during a period with increased raccoon and skunk populations. It was suggested that many birds had died from baylisascariasis, although this conclusion was based on only a single case (Williams et al. 1997). Page et al. (1999) found that 15 species of birds regularly visited sites of raccoon latrines in Indiana, USA, providing further evidence that birds may be exposed to infective larvae of B. procyonis in areas populated by raccoons.
TREATMENT AND CONTROL In the case of Baylisascaris spp., especially B. procyonis, ingestion of only a few eggs can result in death. Management strategies for these agents primarily involve keeping raccoons and other hosts (e.g., mustelids, badgers, and ursids) from contaminating the environment with feces. Numerous compounds have been tested for treatment of adult Baylisascaris in the raccoon, and such treatment can help manage infections in domestic birds (e.g., Bauer and Gey 1995).
ACKNOWLEDGMENTS This material is based on work supported by the National Science Foundation under grant numbers 0529684 and 0608603, NOAA, Oceans and Human Health Initiative Award NAOBNO54730, 322, and financial support from Jubileumsfond 1968, Åbo Akademi and Stiftelsen f¨or Åbo Akademi. We acknowledge the technical assistance of Gunilla Henriksson and Esa Nummelin at the Department of Biology, Abo Akademi University. We acknowledge Prof. G¨oran Malmberg, Stockholm University, for the use of SEM equipment (=“Signhild”). LITERATURE CITED Amato, J. F. R., C. M. Monteiro, and S. B. Amato. 2006. Contracaecum rudolphii Hartwich (Nematoda, Anisakidae) from the Neotropical Cormorant, Phalacrocorax brasilianus (Gmelin) (Aves, Phalacrocoracidae) in southern Brazil. Revista Brasileira de Zoologia 23(4):1284–1289. Anderson, R. C. 2000. Nematode Parasites of Vertebrates: Their Development and Transmission, 2nd ed. CABI Publishing, Wallingford, UK. Armstrong, D. L., R. J. Montali, A. R. Doster, and K. R. Kazacos. 1989. Cerebrospinal nematodiasis in macaws due to Baylisascaris procyonis. Journal of Zoo and Wildlife Medicine 20(3):354–359. Arnall, L., and I. F. Keymer. 1975. Bird Diseases: An Introduction to the Study of Birds in Health and Disease. T. F. H. Publications, Neptune City, NJ. Bartlett, C. M. 1996. Morphogenesis of Contracaecum rudolphii (Nematoda: Ascaridoidea), a parasite of fish-eating birds, in its copepode precursor and fish intermediate host. Parasite 4:367–376. Baruˇs, V., T. P. Sergeeva, M. D. Sonin, and K. M. Ryzhikov. 1978. Helminths of Fish-Eating Birds of the Palearctic Region I. Academia, Publishing House of the Czechoslovak Academy of Science, Prague, Czech Republic. Bauer, C., and A. Gey 1995. Efficacy of six anthelmintics against luminal stages of Baylisascaris
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fishes and birds in the northern Gulf of Mexico. Journal of Parasitology 66(5):853–856. Deardorff, T. L., and R. M. Overstreet. 1981a. Review of Hysterothylacium and Iheringascaris (both previously = Thynnascaris) (Nematoda: Anisakidae) from the northern Gulf of Mexico. Proceedings of the Biological Society of Washington 93(4):1035–1079. Deardorff, T. L., and R. M. Overstreet. 1981b. Raphidascaris camura sp. n., Hysterothylacium eurycheilum (Olsen) comb. N., and comments on Heterotyphlum Spaul (Nematoda: Ascaridoidea) in marine fishes. Journal of Parasitology 67(3):426–432. Deardorff, T. L., M. M. Kliks, and R. S. Desowitz. 1983. Histopathology induced by larval Terranova (Type HA) (Nematoda: Anisakinae) in experimentally infected rats. Journal of Parasitology 69(1):191–195. De Ley, P., and M. L. Blaxter. 2002. Systematic position and phylogeny. In The Biology of Nematodes, D. L. Lee (ed.). Taylor and Francis, London, pp. 1–30. Digiani, M. C., and C. A. Sutton. 2001. New reports and a redescription of Porrocaecum heteropterum (Diesing, 1851) (Ascarididae), a rare nematode parasitic in South American threskiornithid birds. Systematic Parasitology 49(1):1–6. Dzieko´nska-Rynko, J., and J. Rokicki. 2007. Life cycle of the nematode Contracaecum rudolphii Hartwig, 1964 (sensu lato) from northern Poland under laboratory conditions. Helminthologia 44(3):95– 102. Erkinaro, E., and K. Heikura. 1977. Dependence of Porrocaecum sp. (Nematoda) occurences on the sex and age of the host (Soricidae) in Northern Finland. Aquilo, Serie Zoologica 17:37–41. Evans, R. H. 2002. Baylisascaris procyonis (Nematoda: Ascarididae) larva migrans in free-ranging wildlife in Orange County, California. Journal of Parasitology 88(2):299–301. Evans, R. H., and B. Tangredi. 1985. Cerebrospinal nematodiasis in free-ranging birds. Journal of the American Veterinary Medical Association 187(11):1213–1214. fa*gerholm, H.-P. 1978. New Implications of the Nematode Infection of Baltic Cod Liver. Section C. Fourth International Congress of Parasitology. Warsaw, Poland. pp. 192. fa*gerholm, H.-P. 1982. Parasites of fish in Finland. VI. Nematodes. Acta Academiae Aboensis, series B 40(6):1–128. fa*gerholm, H.-P. 1988a. Patterns of caudal papillae in Contracaecum osculatum (Rudolphi) (Nematoda) and some related species from different regions of the world. International Journal for Parasitology 18(8):1039–1051. fa*gerholm, H.-P. 1988b. Incubation in rats of a nematode larva from cod to establish its specific
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identity: Contracecum osculatum (Rudolphi). Parasitogy Research 75(1):57–63. fa*gerholm, H.-P. 1989. Intra-specific variability of the morphology in a single population of the seal parasite Contracaecum osculatum (Rudolphi) (Nematoda: Ascaridoidea), with a redescription of the species. Zoologica Scripta 18(1):33–41. fa*gerholm, H.-P. 1990. Systematic position and delimitation of ascaridoid nematode parasites of the genus Contracaecum with a note on the superfamily Ascaridoidea. Acta Aboensis, Series B 48(4):1–28. fa*gerholm, H.-P. 1991. Systematic implications of male caudal morphology in ascaridoid nematode parasites. Systematic Parasitology 19(3):215–229. fa*gerholm, H.-P. 1996. Nematode parasites of marineand shore birds and their role as pathogens. Bulletin of the Scandinavian Society for Parasitology 6(2):16–30. fa*gerholm, H. -P., and D. I. Gibson. 1987. A redescription of the pinniped parasite Contracaecum ogmorhini (Nematoda, Ascaridoidea), with an assessment of its antiboreal circumpolar distribution. Zoologica Scripta 16(1):19–24. fa*gerholm, H.-P., R. M. Overstreet, and I. Humphrey-Smith. 1996. Contracaecum magnipapillatum (Nematoda: Ascaridoidea): Resurrection and pathogenic effect of a common parasite from the proventriculus of Anous minutes from the Great Barrier Reef, with a note on C. variegatum. Helminthologia 33(4):195–207. fa*gerholm, H.-P., P. Nansen, A. Roepstorff, F. Frandsen, and L. Eriksen. 1998. Growth and structural features of the adult stage of Ascaris suum (Nematoda, Ascaridoidea) from experimentally infected domestic pigs. Journal of Parasitology 84:269–277. fa*gerholm, H.-P., P. Nansen, A. Roepstorff, F. Frandsen, and L. Eriksen. 2000. Differentiation of cuticular structures during the growth of the third-stage larva of Ascaris suum (Nematoda, Ascaridoidea) after emerging from the egg. Journal of Parasitology 86(23):421–427. fa*gerholm, H.-P., M. Brunanska, A. Roepstroff, and L. Eriksen. 2004. Phasmid ultrastructure in an ascaridoid nematode. Hysterothylacium auctum. Journal of Parasitology 90(3):499–506. Garbin, L. E., G. T. Navone, J. I. Diaz, and F. Cremonte. 2007. Further study of Contracaecum pelagicum (Nematoda: Anisakidae) in Spheniscus magellanicus (Aves: Spheniscidae) from Argentinean coasts. Journal of Parasitology 93(1):143–150. Gasser, R. B. 2006. Molecular tools—Advances, opportunities and prospects. Veterinary Parasitology 136(2):69–89. Gavin, P. J., K. R. Kazacos, and S. T. Shulman. 2005. Baylisascariasis. Clinical Microbiology Reviews 18(4):703–718.
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Moravec, F. 1994. Parasitic Nematodes of Freshwater Fishes in Europe. Kluwer Academic Publishers, Dordrecht, The Netherlands. Morgan, B. B., and E. Schiller. 1950. A Note on Porrocaecum depressum (Zeder, 1800), (Nematoda: Anisakidae). Transactions of the American Microscopical Society 69(2):210–213. Murrell, K. D., L. Eriksen, P. Nansen, H.-C. Slotved, and T. Rasmussen. 1997. Ascaris suum: A revision of its early migration path and implications for human ascariasis. Journal of Parasitology 83(2):255–260. Nadler, S. A., and D. S. Hudspeth. 1998. Ribosomal DNA and phylogeny of the Ascaridoidea (Nematoda, Secernentea): implications for morphological evolution and classification. Molecular Phylogenetics and Evolution 10(2):221–236. Nadler, S. A., and D. S. Hudspeth. 2000. Phylogeny of the Ascaridoidea (Nematoda: Ascaridida) based on three genes and morphology: Hypothesis of structural and sequence evolution. Journal of Parasitology 86(2):380–393. Nadler, S. A., S. D’Amelio, H.-P. fa*gerholm, B. Berland, and L. Paggi. 2000. Phylogenetic relationships among species of Contracaecum Railliet & Henry, 1912 and Phocascaris Hø´ st, 1932 (Nematoda: Ascaridoidea) based on nuclear rDNA sequence data. Parasitology 121(4):455–463. Nadler, S. A., S. D’Amelio, M. D. Dailey, L. Paggi, S. Siu, and J. A. Sakanari. 2005. Molecular phylogenetics and diagnosis of Anisakis, Pseudoterranova, and Contracaecum from northern Pacific marine mammals. Journal of Parasitology 91(6):1413–1429. Nadler, S. A., R. A. Carreno, H. Mej´ıa-Madrid, J. Ullberg, C. Pagan, R. Houston, and J.-P. Hugot. 2007. Molecular phylogeny of clade III nematodes reveals multiple origins of tissue parasitism. Parasitology 134(10):1421–1442. Nagasawa, K., V. Barus, F. Tenora, and N. Oka. 1999a. Contracaecum variegatum (Nematoda: Anisakidae) from the Pacific Diver (Gavia pacifica) in Japan. International Journal of Biogeography, Phylogeny, Taxonomy, Ecology, Biodiversity, Evolution, and Conservation Biology 1:107–110. Nagasawa, K., V. Barus, F. Tenora, M. Prokes, and N. Oka. 1999b. Validity and Redescription of Contracaecum himeu (Nematoda, Anisakidae), a Parasite of Cormorants in Japan. Bulletin of the National Science Museum. Series A, Zoology 25(3):149–161. Nascetti, G., B. Berland, S. Mattiucci, and L. Paggi. 1986. Twin species in Contracaecum osculatum (Ascaridida: Anisakidae): reproductive isolation and
diagnostic features at the electrophoresis level. Annali Dell’Instituto Superiore di Sanita 22(1):349–351 (In Italian). Norton, R. A., B. A. Hopkins, J. K. Skeeles, J. N. Beasley, and J. M. Kreeger. 1992. High mortality of domestic turkeys associated with Ascaridia dissimilis. Avian Diseases 36(2):469–473. Oglesby, L. C. 1960. Heavy nematode infestation of white pelican. The Auk 77(3):354. Osche, G. 1955. The development, intermediate hosts and morphology of Porrocaecum talpae, Porrocaecum ensicaudatum and Habronema mansioni (Nematoda) Zeitschrift f¨ur Parasitenkunde 17(2):144–164. (in German). Osche, G. 1959. On intermediate and accidental hosts, and on the ontogeny of the labial region in species of Porrocaecum and Contracaecum (in German). Zeitschrift f¨ur Parasitenkunde 19(5):458–484. Overstreet, R. M. 1970. Baylisascaris procyonis (Stefanski and Zarnowski, 1951) from the kinkajou, Potos flavus, in Colombia. Proceedings of the Helminthological Society of Washington 37(2):192–195. Owre, O. T. 1962. Nematodes in birds of the Order Pelecaniformes. The Auk 79(1):114. Page, L. K., R. K. Swihart, and K. R. Kazacos. 1999. Implications of raccoon latrines in the epizootiology of baylisascariasis. Journal of Wildlife Diseases 35(3):474–480. Paggi, L., and L. Bullini. 1994. Molecular taxonomy in anisakids. Bulletin of the Scandinavian Society for Parasitology 4(2):25–39. Papini, R., P. Cavicchio, and L. Casarosa. 1993. Experimental infection in chickens with larvae of Baylisascaris transfuga (Nematoda: Ascaridoidea). Folia Parasitol (Praha) 40(2):141–143. Raybourne, R., R. S. Desowitz, M. M. Kliks, and T. L. Deardorff. 1983. Anisakis simplex and Terranova sp.: Inhibition by larval excretory–secretory products of mitogen-induced rodent lymphoblast proliferation. Experimental Parasitology 55(3):289–298. Reed, W. M., K. R. Kazacos, A. S. Dhillon, R. W. Winterfield, and H. L. Thacker. 1981. Cerebrospinal nematodiasis in bobwhite quail. Avian Diseases 25(4):1039–1046. Richardson, J. A., K. R. Kazacos, H. L. Thacker, A. S. Dhillon, and R. W. Winterfield. 1980. Verminous encephalitis in commercial chickens. Avian Diseases 24(2):498–503. Riley, J. 1972. The pathology of Anisakis nematode infections of the fulmar Fulmarus glacialis. Ibis 114(1):102–104. Sarashina, T., H. Tanyiyjama, and J. Yamada. 1987. A case of Contracaecum spiculigerum (Ascaridoidea:
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25 Diplotriaena, Serratospiculum, and Serratospiculoides Mauritz C. Sterner, III and Rebecca A. Cole Serratospiculum spp. and Serratospiculoides spp. are so similar that the differences can only be appreciated by microscopy. Serratospiculum spp. were first identified by Skrijabin (1916) from the Eurasian Kestrel (Falco tinnunculus) in Russia. The first species reported in the US was Serratospiculum amaculata (later to be renamed Serratospiculoides amaculata) by Wehr (1935) in a Prairie Falcon (Falco mexicanus) from Montana. All three genera belong to the superfamily Filarioidea. Unlike others in Filariidae, where microfilaria are found in the blood and transmitted by biting insects (see Chapter 26), these parasites lay eggs which pass through the respiratory system and leave the body of the host via fecal material (Chabaud 1955). The eggs are subsequently eaten by foraging insects and larval development of the worm takes place within the hemocoel (Anderson 1957). Infected insects are eaten by birds to complete the life cycle (Anderson 1957, 1962, 1992; Sonin 1963). Studies conducted by Anderson (1962) confirmed the life cycle and led to reclassification of Diplotriaena, Serratospiculum, and Serratospiculoides in the order Spirurida. The superfamily Filarioidea was divided into three superfamilies. Two new superfamilies were added, the Diplotriaenoidea and Aproctoidea, representing nematodes with oviparous development (Sonin 1963). The new classification placed the genus Diplotriaena in the family Diplotriaenidae, subfamily Diplotriaeninae and placed the two genera Serratospiculum and Serratospiculoides in the subfamily Dicheilonematinae (Anderson and Bain 1976; Anderson 1992).
INTRODUCTION Diplotriaena and Serratospiculum are the two most common genera of air sac parasites of birds. Nematodes that belong to the genus Serratospiculoides are less frequently encountered but are closely related. All three genera belong to the order Spirurida, suborder Spirurina, and like all members of this suborder, use arthropods as intermediate hosts. These three genera have historically been confused with filariid nematodes because of their long filiform bodies, sexually dimorphic features, and presence in air sacs. Males are much smaller than females and usually have a spiral tail. Adult worms within these three genera can cause degenerative changes within the collagen and muscle layers between the epithelial and mesothelial components of the air sacs, resulting in the thickening of the air sac tissue. These changes in the lung may predispose birds to pneumonia in areas where eggs and bloody edema have blocked secondary bronchi and associated parabronchi, and lead to secondary infections of Pseudomonas spp., Klebsiella spp., and Aspergillus spp. These secondary bacterial infections, in turn, may result in pneumonia, air sacculitis, aspergillosis, and death of the host. Adult worms may also puncture the air sacs and gain access to the body cavity, depositing eggs within the liver and various other organs. Migration of larvae through organs and the presence of embryonated eggs can cause physical damage to tissues as well as occlusions and congestion of various structures such as bile ducts and hepatic veins.
HISTORY The first species of Diplotriaena was described in Europe and later in India, Russia, Burma, Australia, China, and Africa. The first report of this genus in North America was by Walton (1927) from the Florida Scrub-Jay (Aphelocoma coerulescens).
HOST RANGE AND DISTRIBUTION The distribution of species of Diplotriaena, Serratospiculum, and Serratospiculoides is cosmopolitan. Species of Diplotriaena parasitize the air sacs and
434 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Diplotriaena, Serratospiculum, and Serratospiculoides subcutaneous tissue of birds belonging to the orders Anseriformes, Apodiformes Galliformes, Charadriiformes, Columbiformes, Piciformes, and Passeriformes. Families include Columbidae, Apodidae, Picidae, Furnariidae, Dendrocolaptidae, Tyrannidae, Vireonidae, Corvidae, Hirundinidae, Sittidae, Sylviidae, Muscicapidae, Turdidae, Mimidae, Sturnidae, Bombycillidae, Ptilogonatidae, Parulidae, Thraupidae, and Icteridae. Serratospiculum spp. have been reported from North and South America, Australia, Europe, the UK, Asia, Africa, and the Middle East (Bain and Mawson 1981; Quentin et al. 1983; Gomez et al. 1993; Lierz and Remple 1997; Samour and Naldo 2001; Lloyd 2003). Species of Serratospiculum and Serratospiculoides infect the air sacs of carnivorous birds, primarily of the order Falconiformes, but have also been recorded from accipiters in North America (Sterner and Espinosa 1988; Taft et al. 1993). ETIOLOGY All three genera have typical characteristics of filariid nematodes in that they are long filamentous worms found usually in the subcutaneous tissues but also in the heart, lungs, air sacs, and various other organs of the host. Males are much smaller than females and posses a hooked tail (Olsen 1962). With these similarities they were originally placed in the group classified as filarids. However, further study indicated that they produced thick-shelled eggs that developed in insect hosts rather than microfilariae, which led to a reclassification of these three genera to the order Spiurida. The type species of the genus Diplotriaena, Diplotriaena ozouzi, was described by Railliet and Henry in 1909 in Europe from a reptile, Fuadias madagascariensis. The number of species within the genus has been debated for some time. Skrjabin (1949) listed 65 species in this genus. Yamaguti (1961) listed a total of 76 species. Reclassification of this genus by Anderson and Bain (1976) listed over 47 species as being valid. The most recent consensus is that the genus Diplotriaena has 27 valid species that occur throughout the world (Anderson 1959; Levine 1980). Skrijabin (1916) described a new genus Serratospiculum. Because morphological characteristics were similar to filarids, this genus was placed in the superfamily Filarioidea, family Filariidae, subfamily Setarinae (York and Maplestone 1926; Olsen 1962). Later, studies by Chabaud (1964), Anderson (1992), and Anderson and Bain (1976) showed that the species within the genus Serratospiculum did not develop microfilariae. Instead, eggs were eaten by insects and partial development occurred within the insect host. This life cycle resembled those of spirurid nematodes and
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led to the reclassification of the genus Serratospiculum into the order Spirurida, superfamily Diplotriaenoidea, family Diplotriaenidae, subfamily Dicheilonematinae. The number of species within the genera Serratospiculum and Serratospiculoides has remained constant over the last several years. On the basis of length of the spicules, nine species of Serratospiculum are recognized from avian hosts in the order Falconiformes (Samour and Naldo 2001). The type species for the genus Serratospiculoides is Serratospiculoides alii. Serratospiculoides alii was first described in India by Rasheed in 1960, but identified as Hamatospiculum alii. Chabaud et al. (1964) renamed the specimen Serratospiculum ali, based on differences in spicule morphology from other specimens of Hamatospiculum and their similarity to spicules from Serratospiculum amaculatum. Sonin (1968) examined these two species and determined that the morphology of spicules of S. alii and S. amaculatum were the same and intermediate in form between species of Hamatospiculum and Serratospiculum. These differences in spicule characteristics led to reclassification of this species as the new genus Serratospiculoides (Sonin 1968; Samour and Naldo 2001). There are only two species in this genus, S. alii and S. amaculatum. EPIZOOTIOLOGY The life cycles of species of Diplotriaena, Serratospiculum, and Serratospiculoides are indirect and use an intermediate host. Adults of the species in all three genera are found in the abdominal and thoracic air sacs of the host. The final or definitive hosts of species of Diplotriaena include a variety of passerine species and their intermediate hosts are grasshoppers (order Orthoptera). Thick-shelled eggs containing fully developed firststage (L1) larvae are passed out in the feces of the infected bird. The eggs are eaten by the nymphal stage or young adults of coprophagic (dung eating) grasshoppers (Cawthorn and Anderson 1980a; Ansari 1982). Within the grasshopper, L1 larvae are released and actively burrow out of the gut and into the hemocoel. The larvae lodge in fat bodies throughout the hemocoel and begin to develop (Olsen 1962). Within 4 days the worms have molted to become second-stage (L2) larvae. Within 8 days after invading the fat bodies, they molt again to become third-stage (L3) larvae. The L3 larvae are infective to the definitive host and remain dormant within a clear thin encapsulating membrane until the grasshopper is eaten by a suitable avian host. Encapsulated larvae can survive as long as their intermediate hosts, with most transmission during the spring and summer months (Anderson 1956, 1992; Cawthorn and Anderson 1980a; Ansari 1982).
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After infected grasshoppers are eaten by a suitable avian host, larvae are digested out of the capsules and released into the intestine. The L3 larvae penetrate the intestinal lining and enter the hepatic portal system, where they develop into fourth-stage (L4) larvae, or subadults in approximately 20 days (Cawthorn and Anderson 1980a). They migrate as subadults from the hepatic portal system to the lungs via the right side of the heart and the pulmonary arteries. The subadults subsequently break out of the arterial system and into the lungs where they move to the air sac and undergo a final molt to become adult worms, approximately 20 days after entering the lungs. Adults can remain active and alive within the definitive host for several years (Cawthorn and Anderson 1980b). Female worms mature in 4 months and begin laying eggs within the air sacs that move to the lungs through natural movement of air and mucus. Once in the lungs, the eggs are coughed up, swallowed, and pass out in the feces, completing the life cycle (Anderson 1956). The life cycles of species of Serratospiculum and Serratospiculoides are similar. Eggs are eaten by a variety of coprophagic beetles and hatch in the gut of the intermediate hosts. The L1 larvae move to adipose tissue of the host, where they become encapsulated in a thin and transparent capsule. The L1 larvae develop and mature to the infective L3 stage within these capsules. Samour and Naldo (2001) determined that the L3 remain in the capsule until the beetle is eaten by the definitive host. After ingestion, the L3 are digested out of the capsule and penetrate the wall of the proventriculus or ventriculus. There is some histopathological evidence that the L3 go directly to the air sacs and not through the portal system (Gomez et al. 1993). Once in the air sacs, the L3 larvae undergo two molts and become immature adults (Samour and Naldo 2001). Mature adults mate and females produce a large number of thin-shelled, embryonated eggs. The eggs are coughed up into the mouth, swallowed, and passed out in the feces. CLINICAL SIGNS Birds infected with air sac parasites display chronic illness. Signs include lethargy (“poor doers”), labored breathing, below average body weight or size, and poor plumage or unthriftiness. Infected birds may be less likely to breed. Thin- or thick-shelled eggs can sometimes be seen by microscopy in fecal samples. PATHOLOGY Species of Diplotriaena affect many organs both as a result of the presence of the adult worms and the secondary tissue damage arising from larval migration
and the presence of eggs. The lungs, liver, air sacs, and vascular system are frequently affected. Pathological changes associated with subadult, adult worms, and eggs include inflammatory lesions in the liver and lungs, bronchitis caused by lymphoid hyperplasia, edema in airways, fibroplasias within the air sacs and hepatic parenchyma, adhesions between the liver and air sacs, hemorrhage in the liver, periarteritis, endothelial swelling and vacuolation, congestion in the lungs, thrombi present in arteries, pulmonary inflammation, granulomatous pneumonia in the lungs, and mucoid hyperplasia (Cawthorn et al. 1980). As they penetrate the intestine and enter the hepatic portal system, larvae cause petechial hemorrhages along the intestine and the liver. Traumatic liver lesions can result from the encapsulation of dead nematodes and eggs by giant cell granulomas, leading to chronic liver inflammation, including lymphoid hyperplasia (Ansari 1985; Young et al. 1998). During migration to the lungs via arteries, the subadults cause swelling of the endothelial cells and invasion of the tunica media by macrophages, resulting in periarteritis. Once larvae reach the lungs, they cause edema, fibroplasia, hemorrhaging within the secondary bronchi, lymphoid hyperplasia, and thrombosis. Large giant cell granulomas can be found close to the secondary bronchi. As the number of larvae increase within the lungs, tissue swelling and presence of the worms can cause blockage and congestion of the air passages. Once the subadults reach the air sacs, an acute inflammatory reaction occurs around the parasites, resulting in fibrotic thickening of the air sac as well as adhesions among air sacs, the body cavity, and the internal organs (Cawthorn et al. 1980). The majority of the pathological changes associated with species of Serratospiculum and Serratospiculoides occur in association with the adult worms in the air sacs. Air sac membranes become thickened, compromising air exchange (Kocan and Gordon 1976). Congestion, focal hemorrhages, focal necrosis, and moderate infiltration of macrophages occur within the lungs of infected birds. Serratospiculiasis is characterized by necrosis of the crop and esophagus, edema in the media of the arterioles and bronchial passages, congested hepatic veins, squamous metaplasia, hyperplasia of the mesothelium, heterophile infiltration, focal hemorrhages in the lungs, lesions within the lungs and spinal cord, air sacculitis, and pneumonia (Kocan and Gordon 1976). Other lesions include hepatitis, chronic cholangitis, pericholangitis, pericarditis, atelectasis of the air sacs, and degeneration of the collagen-muscle layer of the air sacs. Damage caused to the air sacs by species of Serratospiculum predispose the bird to secondary infections with species of Aspergillus, Pseudomonas, and Klebsiella (Samour and Naldo 2001).
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Diplotriaena, Serratospiculum, and Serratospiculoides DIAGNOSIS The presence of embryonated eggs within the feces as a diagnostic tool is unreliable because other parasites besides air sac worms lay embryonated eggs. Both the location of the adult worms interwoven within the abdominal air sacs and morphological features characteristic of these genera are the only reliable means for diagnosing an infection. PUBLIC AND DOMESTIC ANIMAL HEALTH CONCERNS There is no evidence that these nematodes infect humans. While there have been no major die-offs or reports concerning deaths associated with this parasite in domestic fowl, free-ranging poultry, such as chickens and turkeys, can become infected by feeding on grasshoppers, although reports are rare. WILDLIFE POPULATION IMPACTS Although deaths of individuals have been attributed to infection with species of these genera (Bigland et al. 1964; Ward and Fairchild 1972; Sterner and Espinosa 1988; Ackerman 1992; Young et al. 1998; Hawkins et al. 2001), no widespread mortality events have been reported. TREATMENT AND CONTROL Since the only reliable means for determining whether these parasites are present is to physically find and identify the worms, treatment specifically for air sac nematodes is usually not undertaken. However, infections with species of Serratospiculum or Serratospiculoides have been treated with mebendazole, Panacur, and fenbendazole with some degree of success in raptors used for falconry. In the Middle East, Samour and Naldo (2001) reported that these parasites are routinely removed endoscopically 3–5 days after treatment with ivermectin. The surgery is followed with a second dose of ivermectin 1–2 weeks later (Lierz 2001; Lloyd 2003). Control of these parasites in wildlife populations is generally not undertaken. DISCLAIMER Any use of trade, product, or firm names in this publication is for descriptive purposes only and does not imply endorsem*nt by the U.S. government.
ACKNOWLEDGMENTS We thank the U.S. Geological Survey Wildlife and Terrestrial Resources Program for financial support.
437
LITERATURE CITED Ackerman, N. 1992. Pneumocoelom associated with Serratospiculum amaculata in a bald eagle. Veterinary Radiology and Ultrasound 1(4):351– 355. Anderson, R. C. 1956. Two new filarid nematodes from Ontario birds. Canadian Journal of Zoology 34:213–218. Anderson, R. C. 1957. Observations on the life cycles of Diplotriaenoides translucidus Anderson and members of the Genus Diplotriaena. Canadian Journal of Zoology 35:15–24. Anderson, R. C. 1959. Further observations on diplotriaena in birds. Parasitologia 1:195–307. Anderson, R. C. 1962. On the development, morphology, and experimental transmission of Diplotriaena bargusinica (Filarioidea: Diplotriaenidae). Canadian Journal of Zoology 40:1175–1186. Anderson, R. C. 1992. Nematode Parasites of Vertebrates: Their Development and Transmission. Commonwealth Agricultural Bureaux, Farnham Royal, Buckinghamshire, England. Anderson, R. C., and O. Bain. 1976. Keys to the Genera of the Order Spirurida. Part 3. Diplotriaenoidea Aproctoidea and Filarioidea. Commonwealth Agricultural Bureaux, Farnham Royal, Buckinghamshire, England. Ansari, W. J. A. 1982. Larval development of Diplotriaena tricuspis (Fedtschenko, 1874) (Filaroidea, Diplotriaenidae) in grasshoppers. Helminthologia 19(2):135–140. Ansari, W. J. A. 1985. Histopathological study of Diplotriaena tricuspis (Fedtschenko, 1874) (Filariidae, Nematoda) infection in pigeons. Acta Parasitologica 30(4):53–56. Bain, O., and P. M. Mawson. 1981. Oviparous filarial nematodes mainly from Australian birds. Records of the South Australian Museum 18:265–284. Bigland, C. H., S. Liu, and M. L. Perry. 1964. Five cases of Serratospiculum amaculatum (Nematoda: Filarioidea) infection in Prairie Falcons (Falco mexicanus). Avian Diseases 8:412–419. Cawthorn, R. J., and R. C. Anderson. 1980a. Development of Diplotriaena tricuspis (Nematoda: Diplotriaenoidea), a parasite of Corvidae, in intermediate and definitive hosts. Canadian Journal of Zoology 58:94–108. Cawthorn, R. J., and R. C. Anderson. 1980b. Diplotriaena tricuspis (Fedtschenko, 1874) (Nematoda: Diplotriaenoidea) in crows (Corvus brachyrhynchos Brehm) wintering in Ontario. Journal of Wildlife Diseases 16:363–365. Cawthorn, R. J., R. C. Anderson, and I. K. Barker. 1980. Lesions caused by Diplotriaena tricuspis (Nematoda:
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Diplotriaenoidea) in the American crow, Corvus brachyrhynchos Brehm. Canadian Journal of Zoology 58:1892–1898. Chabaud, A. G. 1955. Remarques sur le cycle evolutif des filaires du genre Diplotriaena et redescription de D. monticellana (Stossich, 1890). Vie et Milieu 6:342–347. Chabaud, A. G. 1964. Class nematoda. Keys to subclasses, orders, and super-families. In CIH Keys to Nematode Parasites of Vertebrates, R. C. Anderson, A. G. Chabaud, and S. Wilmott (eds). Commonwealth Agricultural Bureaux, Farnham Royal, Buckinghamshire, England. Chabaud, A. G., E. R. Brygoo, and J. Richard. 1964. Filaires d’oiseaux malgaches (deuxieme note). Annales de Parasitologie Humaine et Compare´e 39:69–94. Gomez, M., P. Illescas, M. R. Osorio, and F. A. Maza. 1993. Parasitation of Falconiform, Strigiform and Passeriform (Corvidae) birds by helminths in Spain. Review in Parasitology 53:129–135. Hawkins, M. G, S. Couto, L. A. Tell, V. Joseph, and L. J. Lowenstine. 2001. Atypical parasite migration and necrotizing sacral myelitis due to Serratospiculoides amaculata in a prairie falcon (Falco mexicanus). Avian Diseases 45:276–283. Kocan, A. A., and L. R. Gordon. 1976. Fatal air sac infection with Serratospiculum amaculatum in a Prairie Falcon. Journal of the American Veterinary Medical Association 169:908. Levine, N. D. 1980. Nematode Parasites of Domestic Animals and Man, 2nd ed. Burgess Publishing Company, Minneapolis, MN. Lierz, M. 2001. Evaluation of the dosage of Ivermectin in falcons. Veterinary Record 148(19):596–600. Lierz, M., and J. D. Remple. 1997. Endoparasites in falcons in Dubai. Kleintierpraxis 42:757–768. Lloyd, C. 2003. Control of nematodes in captive birds. In Practice 25:198, 201–206. Olsen, W. O. 1962. Animal Parasites: Their Biology and Life Cycles. Burgess Publishing Company, Minneapolis, MN. Quentin, J. C., C. Seureau, and S. D. Kulo. 1983. Life cycle of Diplotriaena sokolowi an oviparous filarial nematode parasite of the kingfisher Halcyon senegalensis in Togo (In French). Annales de Parasitologie Humaine et Compare´e 58:57–70.
Rasheed, S. 1960. The nematode parasites of the birds of Hyderabad (India). Biologia (Lahore) 6:1–116. Samour, J. H., and J. N. Naldo. 2001. Serratospiculiasis in Captive Falcons in the Middle East: A review. Journal of Avian Medicine and Surgery 15:2–9. Skrijabin, K. J. 1916. N´ematodes des oiseaux du Turkestan russe. Annuaire du Mus´ee Zoologique de l’Acad´emie Imp´eriale des Sciences de Petrograd 20:457. Skrjabin, K. I. 1949. Key to Parasitic Nematodes: Volume 1. Spirurata and Filariata. Izdatel’stvo Akademii Nauk SSSR, Moskva, USSR. Sonin, M. D. 1963. Revision of the suborder Filariata Skrjabin, 1915. Helminthologia 4:485–494. Sonin, M. D. 1968. Filariata of animals and man and diseases caused by them. In Essentials of Nematodology, Vol 21, K. I. Skrjabin (ed.). Akademia Nauk SSSR, Moscow, USSR, pp. 186–190. Sterner, M. C., III, and R. H. Espinosa. 1988. Serratospiculoides amaculata in a Cooper’s hawk Accipiter cooperii. Journal of Wildlife Disease 24:378–379. Taft, S., K. Schuchow, and M. V. Horn. 1993. Helminths from some Minnesota and Wisconsin raptors. Journal of the Helminthological Society of Washington 60:260–263. Walton, A. C. 1927. A revision of the nematodes of the Leidy collection. Proceedings of the Academy of Natural Sciences of Philadelphia 79:49–163. Ward, F. P., and D. G. Fairchild. 1972. Air sac parasites of the genus Serratospiculum in falcons. Journal of Wildlife Diseases 8:165–168. Wehr, E. E. 1935. New genera and species of the nematode superfamily Filarioidea. I. Serratospiculum amaculatum. Proceedings of the Helminthological Society of Washington 5:59–60. Yamaguti, S. 1961. Systema Helminthum: Volume 3. The Nematodes of Vertebrates. Interscience Publishers, New York. York, W., and P. A. Maplestone. 1926. The Nematode Parasites of Vertebrates. J. & A. Churchill Publishers, London, England. Young, E. A., T. E. Cornish, and S. E. Little. 1998. Concomitant mycotic and verminous pneumonia in a blue jay from Georgia. Journal of Wildlife Diseases 34(3):625–628.
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26 Filarioid Nematodes Cheryl M. Bartlett ies of wildlife parasites and diseases require accuracy with respect to species and pathogen identifications.
INTRODUCTION Filarioids are highly specialized nematode parasites of the tissues and tissue spaces of birds, mammals, amphibians, and reptiles. About 160 species are known from birds. Infections are probably much more common than this number and that the literature suggest, primarily because adult worms can be notoriously difficult to find and thus are often not sought or are overlooked at necropsy. Also, with notable exceptions, avian filarioids are not pathogenic, and even among those that are, few draw attention by provoking clinical signs. In addition, adult worms of some species are short lived and disappear after microfilariae (MF), the specialized, first-stage larvae produced by adult female worms, are released. This ephemerality of adults (an unusual life history trait) is balanced by longevity of MF. The life cycles of all filarioids involve a hematophagous invertebrate intermediate host (vector) that ingests MF. After release by the adult female, MF generally enter the bloodstream and become available to vectors. Blood-borne MF are relatively easy to detect; indeed, much of our awareness of avian filarioids as common parasites in birds is based on surveys of avian hosts for hematozoa by examination of stained blood smears. However, surveys of birds for helminths do not always include hematozoa. Most have never included microscopic examination of skin, yet skin rather than blood is occupied by MF of some avian filarioids and such MF are probably more widespread than currently realized. Finally, individual birds can harbor more than one filarioid species (same genus or different genera), and this possibility is not always taken into account at necropsy. This chapter provides the first published synopsis of the taxonomy, biology, and pathogenesis of the filarioid parasites of the world’s birds (see www.integrativescience.ca for exhaustive references). It also emphasizes information that can facilitate finding adult filarioids in birds at necropsy since species identifications are based on adults, and modern stud-
HISTORY Modern classification (Anderson et al. 1974) places filarioids in the superfamily Filarioidea, order Spirurida, suborder Spirurina. Some avian nematodes are still initially identified incorrectly at the level of superfamily, reflecting the overly quick (and outdated) tendency to assign the labels of filarid, filariid, filarioid, or filarial worm to any nematode found outside the lumen of the gastrointestinal tract. However, taxonomic clarity was achieved with the CIH Key No. 1 (Chabaud, in Anderson et al. 1974) to nematode superfamilies in conjunction with the CIH Key No. 3 (Anderson and Bain 1976) to genera in superfamilies Diplotriaenoidea, Aproctoidea, and Filarioidea. The important taxonomic understanding is that nematodes producing microfilariae (family Onchocercidae in superfamily Filarioidea) or those that seem to be closely related (family Filariidae in superfamily Filarioidea) are now classified separately from nematodes inhabiting the air sacs of their hosts (superfamilies Diplotriaenoidea and Aproctoidea) (Chapter 25). Diplotriaenoids (which parasitize both birds and reptiles) produce thick-shelled eggs with fully differentiated first-stage larvae that pass to the outside via the respiratory system and gut of the host. They are believed to be a hom*ogeneous group representing a line of evolution fundamentally distinct from filarioids. Similarly, species in Aproctoidea (which parasitize only birds) generally produce thick-shelled eggs containing fully developed first-stage larvae. They inhabit the air sacs and, unlike the diplotriaenoids, also the nasal cavities, orbits, and subcutaneous tissues of the neck and head. Reports of diplotriaenoids and aproctoids in organs (e.g., lungs, kidneys, liver, intestine) are likely incorrect and might result when delicate air sacs are disrupted at necropsy (Anderson 2000). Aproctoids may be more closely related to filarioids than are the
439 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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diplotriaenoids; more information is required about their life cycles and development. It is regrettable that confusion still occurs with respect to the superfamilies Filarioidea, Diplotriaenoidea, and Aproctoidea. Some confusion is understandable given the taxonomic challenge posed by the CIH keys for the nonexpert. Confusion is also fostered by the continued use of “filariid” and “filarioid” for collective reference to the three superfamilies (e.g., Bain et al. 1992; Chabaud and Bain 1994). ETIOLOGY Adult Filarioid Worms All genera of avian filarioids are classified in the family Onchocercidae, and 16 genera are recognized. Only Pelecitus has species that parasitize both avian and nonavian hosts (Bartlett and Greiner 1986; Jim´enez-Ruiz et al. 2004). Approximately 160 species are known, based mainly on descriptions of adult worms. Descriptions based only on MF are discouraged. There is one report of a biologically aberrant dog heartworm (Dirofilaria immitis) infection in a captive Humboldt Penguin (Spheniscus humboldti) (Sano et al. 2005), but this will not be discussed further. Adult avian filarioids are slender, whitish nematodes with a thin cuticle. They are generally 1–5 cm in length, although shorter or longer specimens are not uncommon. Females are generally larger than males. In comparison to other nematode parasites of vertebrates, filarioids have a simple external morphology and thus they pose a taxonomic challenge for the nonexpert. Cephalic papillae are reduced and other cephalic structures (e.g., pseudolabia) are lacking. The caudal end similarly tends to be nonornate although males have caudal papillae and, in some species, caudal alae (wing-like structures) are present. Delicate or marked cuticular cross striations may occur over the length of the body. A few species have nonprominent lateral alae. Other cuticular ornamentation is lacking. Microfilariae Avian filarioids produce MF that are either blood-borne or skin-inhabiting. Microfilarial morphology can provide useful clues about generic identity, but is insufficient as the sole basis for identification of species which must be based on adult worms. Key features of MF are easily and best observed in wet mounts and can be obscured in dry blood smears. The latter is unfortunate as hematozoan surveys generally use dried smears; however, such surveys have contributed greatly to our general awareness of avian filarioids as common parasites of birds. Such surveys are not able to provide reliable
information about the genus or species of filarioid, although reports do exist where identifications are based solely on MF. These should be viewed with caution and used in conjunction with additional information to make definitive identifications.
HOST RANGE AND DISTRIBUTION A summary of the host distributions of avian filarioids is presented in Table 26.1 and can help guide a search for adult worms at necropsy. Here, the 150 plus families of birds (Howard and Moore 1984) are crossreferenced with the 16 genera of avian filarioids. It is clear from this information that Pelecitus is the most broadly distributed genus with reports from 17 avian orders, followed by Chandlerella in 13, and Paronchocerca, Splendidofilaria, and Cardiofilaria in 11. Also evident is how much work remains to be done with respect to the occurrence of these parasites in most avian families. For example, the family Phasianidae has received considerable study because of its economic importance and appears to contain the most genera (8). Similarly, avian families that are well represented in zoological collections and the pet trade have large numbers of reported genera. For example, 7 genera have been reported from the Psittacidae and 7 genera have been reported in both the Muscicapidae and Corvidae. Due to lack of data it is also premature to make conclusions about possible geographic trends in the distribution of avian filarioids, for example, whether some are more common in temperate zones versus tropics, or Nearctic versus Palearctic. However, cross-referenced information in Table 26.1 may help reveal biologically meaningful relationships, since parasite genera are ordered and grouped by subfamily in a scheme that seeks to reflect evolutionary relationships (Anderson and Bain 1976). At the finer level of parasite genera and species, Paronchocerca appears unique among the genera of avian filarioids that contain numerous species. Each of the 17 species seems restricted to a single family of birds and some degree of host–parasite coevolution should be considered in attempting to understand why (Bartlett and Anderson 1986). As a genus, Paronchocerca occurs in a broad range of hosts but it is known predominantly from those orders generally considered “primitive” or those that occupy the “primitive” adaptive zone, that is, the aquatic and shoreline habitats. Paronchocerca may have become established in early Ornithurae and subsequently persisted in some of the “primitive” birds, as well as having transferred to “modern” groups, which now occupy the aquatic adaptive zone originally occupied by the earliest Ornithurae.
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Mf
Eufilaria
+
+
+
+
+
+
+
+ +
+
+
+
+
+
+
+
?
Eulimdana
Sarconema
Aproctiana
Lemdana
Dessetfilaria
Splendidofilaria
Chandlerella
Andersonfilaria
Cardiofilaria
Aproctella
+
Striatofilaria
+
Pseudlemdana
Paronchocerca
Struthioniformes 1 Struthionidae Rheiformes 2 Rheidae Casuariiformes 3 Casuariidae 4 Dromaiidae Apterygiformes 5 Apterygidae Tinamiformes 6 Tinamidae Sphenisciformes 7 Spheniscidae Gaviiformes 8 Gaviidae Podicipediformes 9 Podicipedidae Procellariiformes 10 Diomedeidae 11 Procellariidae 12 Hydrobatidae 13 Pelecanoididae Pelecaniformes 14 Phaethontidae 15 Pelecanidae 16 Sulidae 17 Phalacrocoracidae 18 Anhingidae 19 Fregatidae Ciconiiformes 20 Ardeidae 21 Balaenicipitidae 22 Scopidae 23 Ciconiidae 24 Threskiornithidae 25 Phaenicopteridae Anseriformes 26 Anhimidae 27 Anatidae Falconiformes 28 Cathartidae 29 Pandionidae 30 Accipitridae 31 Sagittariidae 32 Falconidae Galliformes 33 Megapodiidae
Struthiofilaria
Bird orders and families
Pelecitus
Table 26.1. Reports of avian filarioid genera in families of birds.
+
+
+
+ +
+
+
+
+ + +
+
+
+
+ +
+
?
+
+
+ +
+
+ + + + + (continues)
441
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+
+
+
+ +
+
+
+
Mf
Eufilaria
Eulimdana
Sarconema
Aproctiana
+
Lemdana
+
Dessetfilaria
Splendidofilaria
+
Chandlerella
+
Andersonfilaria
Striatofilaria
Pseudlemdana
Paronchocerca +
Cardiofilaria
+ +
Aproctella
34 Cracidae 35 Phasianidae 36 Opisthocomidae Gruiformes 37 Mesitornithidae 38 Turnicidae 39 Pedionomidae 40 Gruidae 41 Aramidae 42 Psophiidae 43 Rallidae 44 Heliornithidae 45 Rhynochetidae 46 Eurypygidae 47 Cariamidae 48 Otididae Charadriiformes 49 Jacanidae 50 Rostratulidae 51 Dromadidae 52 Haematopodidae 53 Ibidorhynchidae 54 Recurvirostridae 55 Burhinidae 56 Glareolidae 57 Charadriidae 58 Scolopacidae 59 Thinocoridae 60 Chionididae 61 Stercorariidae 62 Laridae 63 Rynchopidae 64 Alcidae Columbiformes 65 Pteroclididae 66 Columbidae Psittaciformes 67 Loriidae 68 Cacatuidae 69 Psittacidae Cuculiformes 70 Musophagidae 71 Cuculidae
Struthiofilaria
Bird orders and families
Pelecitus
Table 26.1. (Continued)
+ + + + + +
+
+ +
+
+ +
+ +
+
+ +
?
+
+ +
+
+ +
+
+
+
+ +
+ +
+
+
442
+
+
+
+ +
+ +
+
+
+
+
+
+ +
+
+ + + +
+ +
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+
+
+
Mf +
+ +
+ +
Eufilaria
Eulimdana
Sarconema
Aproctiana
Lemdana
Dessetfilaria
Splendidofilaria
+
Chandlerella
+
Andersonfilaria
Cardiofilaria
Striatofilaria
Pseudlemdana
Paronchocerca
+
Aproctella
Strigiformes 72 Tytonidae 73 Strigidae Caprimulgiformes 74 Steatornithidae 75 Podargidae 76 Nyctibiidae 77 Aegothelidae 78 Caprimulgidae Apodiformes 79 Apodidae 80 Hemiprocnidae 81 Trochilidae Coliiformes 82 Coliidae Trogoniformes 83 Trogonidae Coraciiformes 84 Alcedinidae 85 Todidae 86 Momotidae 87 Meropidae 88 Coraciidae 89 Brachypteraciidae 90 Leptosomatidae 91 Upupidae 92 Phoeniculidae 93 Bucerotidae Piciformes 94 Galbulidae 95 Bucconidae 96 Capitonidae 97 Indicatoridae 98 Ramphastidae 99 Picidae Passeriformes 100 Eurylaimidae 101 Dendrocolaptidae 102 Furnariidae 103 Formicariidae 104 Conopophagidae 105 Rhinocryptidae 106 Cotingidae
Struthiofilaria
Bird orders and families
Pelecitus
Table 26.1. (Continued)
+
+
+
+ +
+
+
+
+ +
+
+
+
+
+ + +
+
+ +
+ + +
+ +
+
+
+ + +
+
+
+
+
+
+
+
+ + + + + + +
(continues)
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107 Pipridae 108 Tyrannidae 109 Oxyruncidae 110 Phytotomidae 111 Pittidae 112 Xenicidae 113 Philepittidae 114 Menuridae 115 Atrichornithidae 116 Alaudidae 117 Hirundinidae 118 Motacillidae 119 Campephagidae 120 Pycnonotidae 121 Irenidae 122 Laniidae 123 Vangidae 124 Bombycillidae 125 Dulidae 126 Cinclidae 127 Troglodytidae 128 Mimidae 129 Prunellidae 130–142 Muscicapidae 143 Aegithalidae 144 Remizidae 145 Paridae 146 Sittidae 147 Certhiidae 148 Rhabdornithidae 149 Climacteridae 150 Dicaeidae 151 Nectariniidae 152 Zosteropidae 153 Meliphagidae 154–158 Emberizidae 159 Parulidae 160 Drepanididae 161 Vireonidae 162 Icteridae 163 Fringillidae 164 Estrildidae 165 Ploceidae 166 Sturnidae 167 Oriolidae
+
+
+
+
+
+
+ +
+ +
+ +
Mf
Eufilaria
Eulimdana
Sarconema
Aproctiana
Lemdana
Dessetfilaria
Splendidofilaria
Chandlerella
Andersonfilaria
Cardiofilaria
Aproctella
Striatofilaria
Pseudlemdana
Paronchocerca
Struthiofilaria
Bird orders and families
Pelecitus
Table 26.1. (Continued)
+ +
+ +
+
+
+
+
+ + + + + + + + +
+ +
+
+
+
+
+
+
+
+
+
+
+
+
+
+ + +
+
+ +
+ +
+
+ + +
+
+
+
+
+
+
+ +
+ + +
+ + + + +
+ +
+ + +
+
+
+ +
444
+
+ + + +
+ + + + + + +
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445
Filarioid Nematodes
168 Dicruridae 169 Callaeidae 170 Grallinidae 171 Artamidae 172 Cracticidae 173 Ptilonorhynchidae 174 Paradisaeidae 175 Corvidae
+
Mf
Eufilaria
Eulimdana
Sarconema
Aproctiana
Lemdana
Dessetfilaria
Splendidofilaria
Chandlerella
Andersonfilaria
Cardiofilaria
Aproctella
Striatofilaria
Pseudlemdana
Paronchocerca
Struthiofilaria
Bird orders and families
Pelecitus
Table 26.1. (Continued)
+ + + +
+
+
+
+
+
+
+
+ +
Note: Parasite genera are arranged to group closely related taxa. Bird families follow Howard and Moore (1984); all families are included to help highlight where information is lacking. The “?” denotes an irresolvable question as to the species of avian filarioid in the genus noted (i.e., pertains to a species inquirenda). References available at www.integrativescience.ca. Finally, at the level of individual avian hosts, concurrent infections with more than one species of Chandlerella and Splendidofilaria have been noted (e.g., Hibler 1963; Bartlett and Anderson 1980a). Concurrent infections with more than one species in the same filarioid genus might be restricted to parasite genera where adults of the different species tend to occupy very different locations. This diversity of anatomical sites is most commonly observed in the genera Chandlerella and Splendidofilaria. The host distribution of avian filarioids is apt to be narrowest when vectors are host specific (e.g., most lice) and broader when vectors feed on a wide range of avian host species (e.g., blood-sucking dipterans). EPIZOOTIOLOGY Life Cycles and Life History Traits The life cycles of avian filarioids follow the standard filarioid pattern that invariably involves a vertebrate definitive host and an invertebrate intermediate host, the latter referred to as the vector. Adult male and female worms in the vertebrate mate, and the females produce microfilariae that enter the host’s blood or skin. Upon ingestion by a hematophagous arthropod, development proceeds to a so-called sausage stage and then to the second and finally to the infective third stage. Infective third-stage larvae migrate to the head and mouthparts of the arthropod and, while the arthropod feeds, break out of these locations and onto the vertebrate’s skin. Given conditions of suitable moisture,
they quickly enter the puncture wound made by the arthropod and gain entrance to the vertebrate’s body. In the vertebrate, development continues to the fourth larval stage and finally the fifth or adult stage of development. An enduring mystery is how filarioids manage to find their highly specific, final locations in the body of the vertebrate host (Anderson 2000). The first details about development of an avian filarioid in its intermediate host were provided by Anderson (1956) working with Splendidofilaria fallisensis, a parasite of American Black Ducks (Anas rubripes). Hibler (1963), working with Splendidofilaria picacardina, Eufilaria longicaudata, and Chandlerella striatospicula in Black-billed Magpies (Pica hudsonia), provided detailed information about development in the intermediate host as well as the first, and to this day only, details about morphological development of fourthstage larvae in the avian host. In na¨ıve magpies exposed to naturally infected vectors, third-stage larvae of S. picacardina migrate to the definitive site where adult worms are normally found, behind the semilunar valves in the myocardium. By 2 weeks postinfection (PI) in magpies, female fourth-stage larvae (N = 4) were 2.55–2.95 mm long (vs. 33–38 mm for mature adult females) and a male fourth-stage larva was 2 mm (vs. 11–20 mm for mature adult males). In laboratoryreared American Coots (Fulica americana) experimentally infected with Pelecitus fulicaeatrae, subadult fourth-stage worms were present in the definitive site of the ankles when birds were first examined 20 days PI (Bartlett and Anderson 1989). Morphologic changes continued during the fifth stage and details were noted
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at 20, 30, and 55 days PI. Morphologic changes during this stage (other than those normally associated with differentiation of the reproductive tract) have been reported for filarioids in other vertebrate hosts, but are unusual among parasitic nematodes. Among species that have blood-borne MF, the prepatent periods of S. picacardina, C. striatospicula, and E. longicaudata are 42–73, 37–53, 34–76 days, respectively (Hibler 1963), 21–36 days for Cardiofilaria nilesi (Niles et al. 1965; Niles and Kulasiri 1970), 30–36 days for S. fallisensis (Anderson 1956), and approximately 3 months for Splendidofilaria californiensis (Weinmann et al. 1979). Little is known about the prepatent period of avian filarioids that have skin-inhabiting MF. MF of P. fulicaeatrae appear in fluid adjacent to adult worms 210–265 days PI (Bartlett and Anderson 1989). Studies of other species of avian filarioids with skin-inhabiting MF are required to determine if they also have such apparently long prepatent periods. Our understandings of the remarkably diverse locations of adult filarioids in birds are congruent with generalizations in Anderson (2000) who emphasized two points: (1) that filarioids have freed themselves from dependency on the food chain of their vertebrate hosts for transmission (cf. most other nematode parasites in vertebrate hosts) and thus have been able to radiate extensively throughout their hosts’ bodies, and (2) that species have been reported from all organ systems and most tissues. Although adult avian filarioids are notoriously difficult to find, various insights and resources (Table 26.2) can help overcome the challenge. In addition, a general understanding of the locations occupied by adult filarioid worms in birds is extremely useful and these can be summarized by parasite subfamily and genus (Table 26.3). A more detailed summary is provided for Paronchocerca, Splendidofilaria, and Chandlerella in Table 26.4. Adult female filarioids produce MF that are either blood-borne or skin-inhabiting, although in an “occult infection” adult worms will be present but not MF (Table 26.2). For many decades, only blood-borne MF were known for avian filarioids. Skin-inhabiting MF in birds were first discovered in Africa in Little Swifts (Apus affinis = Cypselus affinis) infected with Eulimdana cypseli (=Filaria cypseli). Unfortunately, the observation went unnoticed for years, as it was but a brief statement within a larger manuscript discussing a dog filarioid (Nelson 1962). New awareness of skin-inhabiting MF in birds came with studies of P. fulicaeatrae in American Coots (Bartlett and Anderson 1987b) and various species of Eulimdana in birds in the order Charadriiformes (Bartlett et al. 1989; Bartlett and Anderson 1990; Bartlett 1992, 1993). However, we do not yet have a
detailed understanding of the host range of filarioids that produce skin-inhabiting MF in birds. The genera Eulimdana and Pelecitus are now known to have some species that produce skin-inhabiting MF and other species that produce blood-borne MF. Other genera where only blood-borne MF are currently known require additional study as this same diversity may occur elsewhere. Some species of avian filarioids exhibit life history traits that are not known among the filarioids of other vertebrate hosts. Reproductive senescence and ephemerality were first recognized in studies of P. fulicaeatrae in American Coots and species of Eulimdana in charadriiforms (Bartlett et al. 1989; Bartlett and Anderson 1990; Bartlett 1992, 1993). These avian filarioids have skin-inhabiting MF and are transmitted by lice (order Phthiraptera). Reproductive senescence (curtailment of MF production while adult worms continue to live) and emphemerality (death of adult worms soon after females produce MF) both lead to a short period of MF production. This appears to be an adaptation to limit within-host transmission where continuous ingestion of MF might kill the louse vectors. Thus, reproductive senescence may have evolved in species that occupy sites where, if they were to die, they might initiate life-threatening inflammation (e.g., Pelecitus fulicaeatrae near joints in legs) (Anderson and Bartlett 1994). Ephemerality may have evolved in species in which adults occupy sites where, when they die, they are harmlessly resorbed (e.g., species of Eulimdana in the neck) (Anderson and Bartlett 1994). Vectors Since adult filarioids occur in tissue sites that do not connect with the passageways of the gastrointestinal, renal, or respiratory tracts of the bird host, they have evolved specialized means of transmission: all use hematophagous arthropod vectors (Anderson 2000). Vectors are known for 18 species of avian filarioids and include, in order of discovery, lice (order Phthiraptera) and flies (order Diptera, families Simuliidae, Culicidae, and Ceratopogonidae) (Table 26.5). Details of parasite development first emerged in studies using Simuliidae (Anderson 1956) and Ceratopogonidae (Hibler 1963). At the generic level, it is apparent that avian filarioids may use more than one family or order of vector (Table 26.5). At the species level among those that use dipteran vectors, there appears to be restriction to one family of vectors, but more than one species in a particular vector genus may be used. Splendidofilaria fallisensis develops in two species of Simulium (Anderson 1956, 1968), and E. longicaudata, Chandlerella quiscali, and Chandlerella chitwoodae develop in at
447 Know that adult worms will be present but MF will not be found Know that adult worms will be present but MF will not be found
Prepatent infection
In some filarioid species, adults die and are resorbed soon after they produce MF but MF are long lived In some host species, adult female filarioids produce MF but MF become trapped in inflammatory tissues around adult worms MF are not produced when adults are still immature MF are not produced when only female or only males worms are present
Tables 26.3 and 26.4
Table 26.7; Figures 26.2–26.15
Table 26.1
Resources to consult or information to consider
Morphology of MF present Possible filarioid genus or genera present Possible filarioid genus or genera present
+ + +
−
−
−
Host species identification
Other
+ or −
MF in bird*
∗
Note: The order of presentation is not prescriptive; circ*mstances at necropsy should determine use, sequence, and/or combination. This information must be determined by looking for microfilariae (MF) in the blood and skin of the bird. Microfilariae might be found (i.e., are present: +) or they might not be found (i.e., are absent: −).
Sterile infection
Know that adult worms will be present but MF will not be found in blood
Know that adult worms will not be present
Occult infection
Sites occupied by adult worms, by genus Ephemerality
Determine possible filarioid genus or genera present in the species of bird being examined Determine possible filarioid genus or genera present in the individual bird being examined Determine priority location(s) to examine in bird
Use
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Microfilarial morphology
Host distribution
Subject
Additional information required (to be determined by examiner)
Table 26.2. Knowledge, resources, and information to guide searches for adult filarioid worms in birds at necropsy.
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Parasitic Diseases of Wild Birds
least two species of Culicoides (Hibler 1963; Robinson 1971; Bartlett and Anderson 1980b). There may be less vector specificity among filarioids that use louse vectors; Eulimdana bainae and Eulimdana wongae develop in both ischnoceran and amblyceran lice (Bartlett 1993), which are generally viewed as different suborders of Phthiraptera (Chapter 29). Bird–filarioid relationships, including the roles vectors play, have been discussed most recently by Bartlett and Anderson (1980a, b, 1986, 1987b), Bartlett and Greiner (1986), Hoberg (1986), and Bartlett (1992).
Broad host ranges have been suggested for many avian filarioids (Bartlett et al. 1985; Bartlett and Greiner 1986). Filarioids that are not host specific can be sporadic, occult, or common in different bird species in the same bird community (Bartlett and Anderson 1980a). In contrast, Bartlett (1992) suggested narrower host distributions for other avian filarioids, especially those transmitted by lice, which are host specific. A different perspective, namely that most species of avian filarioids have very narrow host ranges, regardless of vector, leads to a proliferation of descriptions
Table 26.3. Sites in the avian host occupied by adult avian filarioids, by subfamily and genus. Subfamily
Genus
Dirofilariinae
Pelecitus
Onchocercinae Splendidofilariinae
Struthiofilaria Paronchocerca Pseudlemdana Striatofilaria Aproctella
Cardiofilaria Andersonfilaria Chandlerella Splendidofilaria Dessetfilaria Lemdaninae
Lemdana Aproctiana Sarconema Eulimdana Eufilaria
Sites
r Near joints of legs, feet, or toes r Other* : subcutaneous in neck and around esophagus Body cavity Diverse, see Table 26.4 r Generally: connective tissue around trachea and esophagus r Occasionally: subcutaneous connective tissue of head, neck, thorax, or thighs r Subcutaneous connective tissue of neck and connective tissue around trachea r Other† : thoracic cavity r Generally: body cavity or in association with heart r Occasionally: subcutaneous tissue of neck r Rarely† : kidney r Generally: body cavity or in association with heart r Rarely† : subcutaneous connective tissue, lungs, hepatic portal vein In fossa in dorsal wall of pelvic girdle under middle region of right kidney Diverse, see Table 26.4 Diverse, see Table 26.4 r Outer wall of aorta r Other‡ : cervical air sacs r Subcutaneous connective tissue of head and neck, or connective tissue around trachea, esophagus, or crop r Other† : body cavity Abdominal cavity r Sarconema eurycerca: heart r Sarconema pseudolabiata: subcutaneous connective tissue of neck r Generally: subcutaneous connective tissue of head and neck, or connective tissue around trachea, esophagus, or crop r Rarely: body cavity r Generally: subcutaneous connective tissue of head and neck, or connective tissue around trachea, esophagus, or crop r Occasionally: subcutaneous connective tissue of groin or legs
* Reports are likely misidentifications of species of Lemdana or Eulimdana (subfamily Lemdaninae). Report(s) requires confirmation and/or clarification. ‡ Report is undoubtedly an error resulting when delicate air sacs are disrupted at necropsy. †
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Table 26.4. Sites in the avian host occupied by adult filarioids in the genera Paronchocerca, Splendidofilaria, and Chandlerella. Sites in host Circulatory system Heart
Paronchocerca
Splendidofilaria
Chandlerella
Paronchocerca ciconiarum Paronchocerca limboonkengi Paronchocerca tonkinensis Paronchocerca rousseloti Paronchocerca mirzai Paronchocerca straeleni Paronchocerca mansoni Paronchocerca bumpae Paronchocerca francolina Paronchocerca sonini
Splendidofilaria pavlovskyi Splendidofilaria travassosi Splendidofilaria brevispiculum Splendidofilaria verrucosa Splendidofilaria californiensis Splendidofilaria kasmirensis Splendidofilaria wehri Splendidofilaria alii Splendidofilaria pachacuteci S. longicaudata Splendidofilaria osmaniae
Chandlerella bosei Chandlerella skrjabini Chandlerella lerouxi Chandlerella pelecani Chandlerella singhi Chandlerella buckleyi Chandlerella himalayansis Chandlerella alii Chandlerella apusi Chandlerella longicaudata Chandlerella sultana
Behind heart valves
Splendidofilaria picacardina Splendidofilaria travassosi Splendidofilaria periarterialis Splendidofilaria algonquinensis Splendidofilaria caperata
Aorta Pulmonary arteries Paronchocerca (walls and/or lumen) ciconiarum Paronchocerca rousseloti Other blood vessels Lumen of blood vessels Abdominal cavity
Paronchocerca bambusicolae Paronchocerca ibanezi Paronchocerca schelupovi
Liver
Spleen Kidney Lungs
Paronchocerca papillatus Paronchocerca thapari Paronchocerca francolina Paronchocerca sonini Paronchocerca struthiones
Connective tissues Trachea/esophagus
Chandlerella sinensis Chandlerella apusi Chandlerella bosei Splendidofilaria smithi Chandlerella bosei Splendidofilaria verrucosa Chandlerella stantchinski∗ Splendidofilaria grettillati Chandlerella sinensis Chandlerella Splendidofilaria columbigallinae kasmirensis Chandlerella lienalis Chandlerella thapari Chandlerella columbae Chandlerella inversa Chandlerella bosei Chandlerella sinensis Chandlerella lienalis Chandlerella hepatica Chandlerella lienalis Chandlerella shaldybini Splendidofilaria falconis Chandlerella bosei Chandlerella sinensis
Splendidofilaria skrjabini
Blood vessels (general) Splenic artery Intestine Lymphatics
Chandlerella sinensis Chandlerella chitwoodae Chandlerella striatospicula Chandlerella chitwoodae Chandlerella striatospicula Chandlerella striatospicula Chandlerella bushi (continues)
449
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Parasitic Diseases of Wild Birds
Table 26.4. (Continued ) Sites in host
Paronchocerca
Subcutaneous tissues Nonspecific locations
Head Neck
Bursa of knee Feet
Chandlerella
Splendidofilaria smithi Chandlerella robinsoni Splendidofilaria fallisensis Splendidofilaria singhi Splendidofilaria columbensis Splendidofilaria hibleri Splendidofilaria singhi
Paronchocerca rousseloti Paronchocerca rousseloti
Splendidofilaria singhi
Chest Thighs Brain Eyes
Splendidofilaria
Splendidofilaria pectoralis Splendidofilaria gedoelsti Paronchocerca helicina
Calcaneal region Subcutaneous tissues
Splendidofilaria smithi Splendidofilaria rotundicephala Splendidofilaria mavis Splendidofilaria bohmi Splendidofilaria tuvensis
Chandlerella quiscali Chandlerella petrowi
Note: References available at www.integrativescience.ca. * Correction of original report as “air sacs.”
of species; this perspective is not held by this author. CLINICAL SIGNS Little has been reported about the clinical signs of infections with avian filarioids because very few species are pathogenic and, even with those, infections are generally subclinical. Moreover, clinical disease, when noted, may involve only a few birds. Reported signs include reduced body weight (Seegar 1979a) and acute depression (www.wildlifeinformation.org1 ) in association with Sarconema eurycerca, swollen joints and/or lameness with Pelecitus spp. (Greve et al. 1982; Paster 1983; Allen et al. 1985; Kummerfeld and Daugschies 1989), feather loss with Eulimdana clava (Eslami 1987; Gharagozlou 1988; Pizarro et al. 1994), and torticollis and progressive ataxia with C. quiscali (Law et al. 1993).
in the walls of the pulmonary arteries of American Crows (Corvus brachyrhynchos) infected with Splendidofilaria caperata. MF are trapped in situ and the infection is “occult” (see Point #4 below). Of note with respect to skin-inhabiting MF is the possibility that they are the cause of the disease associated with E. clava in domestic Rock Pigeons (Columba livia) (see Point #3 below). Few adult avian filarioids provoke overt disease in a live bird or gross tissue damage that can be observed at necropsy. Adults of a species of avian filarioid that are pathogenic in one or more species of birds may not be pathogenic in other species. Similarly, a species that is pathogenic in an individual bird may not be pathogenic in other birds of the same species. Known pathogens include the following:
1. Splendidofilaria eurycerca. This filarioid has been reported from various species of geese PATHOLOGY and swans (Anatidae). Adults live under the MF are generally considered nonpathogenic although epicardium or within the myocardium. Patholthere are exceptions. Of note with respect to bloodogy includes weakness and enlargement of the borne MF is the chronic inflammation caused by MF heart, myocardial hemorrhage, and myocardial inflammation, necrosis, and eventually fibrosis (www.wildlifeinformation.org; Quortrup and Holt 1 S. Boardman, and Bourne, D. C. (eds). Waterfowl: Health 1940; Cowan 1946; Kluge 1967; Irwin 1975; Cole and management waterfowl diseases, Available at http://wildlife1. 1999) (Figure 26.1). Severity of lesions increases wildlifeinformation.org/List Vols/Waterfowl Mod/Wildpro with increasing numbers of worms. Waterfowl Cont.htm; and Sarconema eurycerca, Available at 2. Pelecitus spp. Swellings and nodules in the legs http://wildlife1.wildlifeinformation.org/S/00dis/Parasitic/ and feet, especially near joints, and tenosynovitis Heartworm Sarconema Infection.htm.
451
Eufilari longicaudata Eufilaria bartlettae Eufilaria delicata Eufilaria kalifai
Splendidofilaria picacardina Splendidofilaria californiensis
Aproctella alessandroi Chandlerella striatospicula Chandlerella quiscali Chandlerella chitwoodae
Pelecitus ceylonensis
f*ckuda et al. (2005) Niles (1962); Niles et al. (1965); Dissanaike and Fernando (1965); Dissanaike and Niles (1967) Niles et al. (1965); Dissanaike (1967); Mosquito§ (Mansonia crassipes) Dissanaike and Niles (1967) Blue-gray Tanager (Thraupis episcopus) Bain et al. (1981) Black-billed Magpie (Pica hudsonia) Hibler (1963) Common Grackle (Quiscalus quiscula versicolor) Robinson (1971) Common American Crow (Corvus Bartlett and Anderson (1980b) brachyrhynchos) Black-billed Magpie (Pica hudsonia) Hibler (1963) California Quail (Callipepla californica) Weinmann et al. (1979); Atchley and Wirth (1975) Black-billed Magpie (Pica hudsonia) Hibler (1963) Eurasian Blackbird (Turdus merula) Bain (1980) Eurasian Blackbird (Turdus merula) Bain (1980) Eurasian Magpie (Pica pica pica) Millet and Bain (1984)
Anderson (1956, 1968)
Dutton (1905); Nelson (1962) Bartlett (1992) Bartlett (1992) Seegar et al. (1976); Cohen et al. (1991) Bartlett and Anderson (1987a, b)
Reference
Note: Filarioid species are grouped by vector, chronological order of discovery, and then parasite genus. * The larvae found were presumed to be a species of Splendidofilaria. † The wild bird host was not known. ‡ The wild bird host was not known by Niles (1962) who found infective filarioid larvae in mosquitoes. These were experimentally inoculated into chickens by Niles et al. (1965) and the adult worms recovered were described as the new species Cardiofilaria nilesi by Dissanaike and Fernando (1965). The infective larvae were described by Dissanaike and Niles (1967). § The wild bird host was not known by Niles et al. (1965) who found two types of infective filarioid larvae in mosquitoes. Larvae of one of these types were similar to those of Niles (1962). Larvae of the second type were experimentally inoculated into “ash-doves” by Dissanaike (1967) who described the adult worms recovered as the new species Pelecitus nilesi, and also reported the species in wild “crows.” The infective larvae were described by Dissanaike and Niles (1967).
Family Ceratopogonidae
Little Swift (Apus affinis =Cypselus affinis) Whimbrel (Numenius phaeopus) Marbled Godwit (Limosa fedoa) Tundra Swan (Cygnus columbianus) American Coot (Fulica americana)
Avian or arthropod host
American Black Duck (Anas rubripes); domestic duck (Anas platyrhynchos) Splendidofilara sp. (presumed* ) Blackfly† (Simulium sp.) Cardiofilaria nilesi Mosquito‡ (Mansonia crassipes)
Splendidofilaria fallisensis
Eulimdana cypseli Eulimdana bainae Eulimdana wongae Sarconema eurycerca Pelecitus fulicaeatrae
Filarioid species
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Family Culicidae
Order Diptera Family Simuliidae
Order Phthiraptera
Vector
Table 26.5. Known vectors of avian filarioids.
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Figure 26.1. Heart of a Tundra Swan (Cygnus columbianus) with myocarditis due to Sarconema eurycerca. At this magnification, separation of myofibers is evident with interstitial hemorrhage, and infiltration with mixed inflammatory cells including heterophils, eosinophils, macrophages, and perivascular lymphocytes. Developing embryos are visible in the ovary of the parasite (top section). In more advanced cases, necrosis and fibrosis may be present in the myocardium. Hematoxylin and eosin; 400×. Photomicrograph Courtesy of US Geological Survey, National Wildlife Health Center. have been reported in psittaciformes that harbor Pelecitus spp. (e.g., Greve et al. 1982; Paster 1983; Allen et al. 1985; Kummerfeld and Daugschies 1989). Reports of disease in psittaciformes due to “Pelecitus” in the neck region are likely misidentifications of worms in the genus Eulimdana. The adults of Pelecitus fulicaeatrea occur near the ankles and generally provoke swellings in Red-necked Grebes (Podiceps grisegena) but rarely in American Coots (Bartlett and Anderson 1989). In such rare cases in coots, worms were generally within soft, thin-walled capsules but a note was also made of worms within a fibrous, thick-walled capsule. Histopathologic changes in the tendon sheaths adjacent to adult worms included synovial cell hypertrophy, hypervascularization, inflammatory cell infiltration, and mild fibrosis. 3. Eulimdara clava. Disease has been associated with infections of E. clava in domestic Rock Pigeons in Iran (Eslami 1987; Gharagozlou 1988). Clinical signs included loss of feathers in the head, neck,
back, and wings. One report indicated that MF were not found in blood where they were expected although MF were present in female worms (Eslami 1987). It is possible these clinical signs were related to skin-inhabiting MF, which are now known for other species of Eulimdana and may also occur with E. clava. In 1994, Pizarro et al. reported that a pigeon that harbored adult E. clava (identified as “Pelecitus clavus”) had MF in subcutaneous tissues immediately adjacent to adult worms in the neck but the broader possibility that MF were skininhabiting was not considered. “Peritracheal filariosis” described in pigeons (Guidal and Settnes 1968; Rutherford and Black 1974) undoubtedly refers to adult E. clava and the MF reported in the lungs (i.e., in blood) were probably of a different species and not those of E. clava. 4. Splendidofilaria caperata. In American Crows, infection with S. caperata is accompanied by chronic inflammation that is caused by MF near adults living in tortuous channels within the walls of the pulmonary arteries; MF become trapped in the inflamed tissues and are not found in the blood (Bartlett and Anderson 1981). Such amicrofilaremic infections are called “occult.” Some inflammation was reported around adult worms in Black-billed Magpies infected with this same parasite but MF were present in blood (Hibler 1963). Pathology has not been noted with S. caperata in various other wild birds, including five more passeriform species and a species in each of Strigiformes, Coraciiformes, and Gruiformes (Bartlett and Anderson 1985, 1987a, c). 5. Species with adults in “sensitive” locations. Adults of species of Paronchocerca, Chandlerella, and Splendidofilaria live in various locations in birds (Table 26.4), and lesions have been reported in some cases where adults live in the muscles or lumen of the heart, walls or lumen of major blood vessels, or brain. The occurrence of S. californiensis in nodules on the luminal surface of the aorta elicits a relatively mild inflammation, but can occlude 20–60% of the aortic orifice and interfere with movement of aortic valves, thus compromising cardiac output (Weinmann et al. 1979). In addition to S. caperata, adults of Splendidofilaria algonquinensis live in the pulmonary arteries and cause lesions in House Sparrows (Passer domesticus) (Huizinga et al. 1971), although infections are not occult. Adults of C. quiscali live in the brain. Clinical signs and pathology were associated with this parasite in farmed Emus (Dromaius novaehollandiae) (Law et al. 1993), but lesions have
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453
not been reported in infected wild passeriforms in North America including American Crows, Blue Jays (Cyanocitta cristata), Brown-headed Cowbirds (Molothrus ater), Common Grackles (Quiscalus quiscula versicolor), and (introduced) European Starlings (Sturnus vulgaris). While adult Paronchocerca ciconarium were associated with cardiovascular lesions in a Marabou Stork (Leptoptilos crumeniferus) that died in a zoo (Ensley 1978), the role of Paronchocerca spp. in the deaths of other zoo birds is not clear (Bartlett and Anderson 1986; Nicholls et al. 1995). Splendidofilaria eurycerca in anatids is another example of a filarioid whose adults live in a “sensitive location,” namely, the muscles of the heart. It can be pathogenic although this is not inevitable. Intensity of infection and overall condition of the avian host are important.
DIAGNOSIS Finding adult worms at necropsy is a critical step in identifying species and can be aided by diverse information (Table 26.2) including the genera of filarioids previously reported in the relevant avian host family (Table 26.1) and the locations that adult parasites in these genera tend to occupy in the bird (Tables 26.3 and 26.4). A key to genera has been provided by Bartlett and Anderson (1987a), along with the 31 synonyms for the 16 recognized genera. References for recent taxonomic reviews (or for the original generic proposals) are provided in Table 26.6. An exhaustive list of species, synonyms, taxonomic authorities, and other references can be found at www.integrativescience.ca. The finding of adult filarioid worms at necropsy can be greatly facilitated by determining what morphological type(s) of MF are present (Table 26.7, Figures 26.2– 26.15). It is important, therefore, to use appropriate techniques to look for MF. When a carcass (fresh, refrigerated, or thawed) is examined, a small piece of lung and skin can be torn into tiny pieces in a few drops of physiologic saline on a microscope slide. After removing tissue bits, an equal volume of 2–5% formalin is added, and the preparation is covered with a vaseline-ringed microscope cover glass. The resulting wet mount is then examined by microscopy for MF using a compound microscope with a 10× objective lens or higher magnification. A slight modification involves the addition of a vital stain (brilliant cresyl blue or Giemsa) to a saline-only preparation before the cover glass is applied. Skin snips (2–3 mm2 ) should be taken from sites at or near the locations where adult worms are expected (e.g., legs for Pelecitus spp. and neck for Eulimdana spp.), regardless of whether birds are dead or alive.
Figure 26.2. Morphology of microfilariae of Andersonfilaria africanus, Giemsa-stained thin blood smear. S, sheath (which has detached from body). Note: Microfilaria was obtained from the blood or skin of the avian host unless otherwise indicated. Caution is advised when morphological information based on microfilariae from uteri of female worms is used (see text). Adapted from Figure 73 in Bartlett and Bain (1987) and reproduced with permission of the Comparative Parasitology, formerly Proceedings of the Helminthological Society of Washington. When a live bird is available, blood samples can be taken, preferably using techniques that do not involve drying the blood. The hematocrit centrifuge technique enables study of MF in a wet mount preparation (Woo 1971). After centrifugation, the hematocrit capillary tube is broken at the interface between serum and cells where MF concentrate and MF are then expressed onto
Figure 26.3. Morphology of microfilariae of Aproctella stoddardi, Giemsa’s stain. Note as given in Figure 26.2. Adapted from Figure 7 in Anderson (1957) and reproduced with permission of the Canadian Journal of Zoology.
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Table 26.6. The 16 genera of avian filarioid nematodes from the family Onchocercidae. Subfamily
Reference for recent taxonomic review (or proposal for new genus)
Number of valid species
Dirofilariinae 1. Pelecitus Onchocercinae 2. Struthiofilaria Splendidofilariinae 3. Paronchocerca 4. Pseudlemdana 5. Striatofilaria 6. Aproctella 7. Cardiofilaria 8. Andersonfilaria 9. Chandlerella 10. Splendidofilaria 11. Dessetfilaria Lemdaninae 12. Lemdana 13. Aproctiana 14. Sarconema 15. Eulimdana 16. Eufilaria
16
Bartlett and Greiner (1986)
1
Noda and Nagata (1976)
17 1 1 7 14 2 26 31 2
Bartlett and Anderson (1986) Sonin (1968) Sonin (1968) Anderson (1957) Bartlett and Anderson (1980a) Bartlett and Bain (1987) Bartlett and Anderson (1980a) Bartlett and Anderson (1980a) Bartlett and Bain (1987)
9 1 2 16 14
Bartlett and Anderson (1987a) Sonin (1968) Sonin (1966) Bartlett et al. (1985) and Bartlett (1992) Bartlett and Anderson (1980a)
Note: Genera are grouped by subfamily, with information provided as to the numbers of valid species and references for recent taxonomic reviews or original generic proposals (see www.integrativescience.ca for the taxonomic authorities, 31 generic synonyms, and exhaustive references).
+ + +
+
+
+
+
+
+
+
+ + +
+
+
+ +
+
+
+ +
Eufilaria
+
Eulimdana
+
+
Sarconema
+
Aproctiana
+
Lemdana
+
Dessetfilaria
+
+
Splendidofilaria
+
Chandlerella
+
+ +
Andersonfilaria
+ +
Cardiofilaria
+ +
Aproctella
+
Striatofilaria
+
Pseudlemdana
Paronchocerca
Sheath Present Absent Tail Broadly rounded Sharply pointed Tapering rounded Length ≤200 m ∼ 200 m ≥200 m
Struthiofilaria
Morphological traits of microfilariae
Pelecitus
Table 26.7. Key morphological traits of microfilariae of avian filarioids in different genera.
+
+
+
+
+
+
Note: Parasite genera are arranged to group closely related taxa, and all 16 genera are included to highlight where information is lacking. References available at www.integrativescience.ca. Information in this table should be viewed as “suggestive” rather than “diagnostic” since exceptions may occur in the pattern portrayed for each genus.
454
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455
Figure 26.5. Morphology of microfilariae of Paronchocerca ciconiarum, in glycerin from uterus of female worm. Note as given in Figure 26.2. Adapted from Figure 55 in Bartlett and Anderson (1986) and reproduced with permission of the Canadian Journal of Zoology. Figure 26.4. Morphology of microfilariae of Cardiofilaria pavlovskyi, cresyl blue vital stain in wet mount. Note as given in Figure 26.2. Adapted from Figure 3 in Bartlett and Anderson (1980a) and reproduced with permission of the Systematic Parasitology.
a microscope slide for preparation of a wet mount. Techniques described for mammalian blood do not necessarily work well for the nucleated erythrocytes of avian blood and Seegar (1979b) and Holmstad et al. (2003) have explored more appropriate techniques. Dried, stained thin blood smears are useful within the limitations previously mentioned. Some information about MF morphology in the scientific literature (e.g., Figures 26.5 and 26.6) is based on specimens from the vagin* or uteri of female worms (especially females preserved in fixatives required for taxonomic study). Caution is advised as these MF tend
to be shorter than MF in blood or skin and the presence or absence of a sheath can be difficult to definitively ascertain. IMMUNITY Very little is known about avian immunity in the context of filarioid infections in wild birds. The unusual life history traits of ephemerality and reproductive senescence may be genetically controlled, but it is also possible that they are mediated by host immune processes (Anderson and Bartlett 1994). MF of C. nilesi disappeared from the blood of experimentally infected chickens (Gallus gallus), leading Gooneratne (1969) to suggest an immune response that affects MF but not adults. Niles and Kulasiri (1970) indicated such immunity might not be relevant and further study was required. It should be noted that chickens (order Galliformes) are not the normal host of C. nilesi and that the parasite has been reported
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Figure 26.8. Morphology of microfilariae of Eufilaria longicaudata, cresyl blue vital stain in wet mount. Note as given in Figure 26.2. Adapted from Figure 46 in Bartlett and Anderson (1980a) and reproduced with permission of the Systematic Parasitology.
Figure 26.6. Morphology of microfilariae of Paronchocerca struthionus, in glycerin from uterus of female worm. Note as given in Figure 26.2. Adapted from Figure 21 in Bartlett and Anderson (1986) and reproduced with permission of the Canadian Journal of Zoology.
Figure 26.9. Morphology of microfilariae of Dessetfilaria guianensis, hematein-stained thin smear. Note as given in Figure 26.2. Adapted from Figure 72 in Bartlett and Bain (1987) and reproduced with permission of the Comparative Parasitology, formerly Proceedings of the Helminthological Society of Washington.
Figure 26.7. Morphology of microfilariae of Eulimdana metcalforum, in 2–5% formalin-saline wet mount. Note as given in Figure 26.2. Adapted from Figure 22 in Bartlett (1992) and reproduced with permission of the Systematic Parasitology.
456
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Figure 26.12. Morphology of microfilariae of Splendidofilaria caperata, cresyl blue vital stain in wet mount. Note as given in Figure 26.2. Adapted from Figure 28 in Bartlett and Anderson (1980a) and reproduced with permission of the Systematic Parasitology.
Figure 26.10. Morphology of microfilariae of Chandlerella bushi, Giemsa-stained thin smear. Note as given in Figure 26.2. Adapted from Figure 11 in Bartlett and Anderson (1987c) and reproduced with permission of the Canadian Journal of Zoology.
Figure 26.13. Morphology of microfilariae of Lemdana wernaarti, cresyl blue vital stain in wet mount. Note as given in Figure 26.2. Adapted from Figure 18 in Bartlett and Anderson (1987a) and reproduced with permission of the Canadian Journal of Zoology.
Figure 26.11. Morphology of microfilariae of Struthiofilaria megalocephala, unspecified preparation technique. Note as given in Figure 26.2. Adapted from Figure 8 in Noda and Nagata (1976) and reproduced with permission of the Bulletin of the University of Osaka Prefecture.
457
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Parasitic Diseases of Wild Birds
Figure 26.14. Morphology of microfilariae of Pelecitus fulicaeatrae, cresyl blue vital stain in wet mount. Note as given in Figure 26.2. Adapted from Figure 6 in Bartlett and Anderson (1987b) and reproduced with permission of the Canadian Journal of Zoology.
in wild Spotted Doves (Streptopelia chinensis) (order Columbiformes) (Lee and Amin-Babjee 1986).
PUBLIC AND DOMESTIC ANIMAL HEALTH CONCERNS There are few known public or domestic animal health concerns associated with avian filarioids in wild birds and the possibility has received little study. Clinical signs and pathology were associated with C. quiscali in farmed Emus (Law et al. 1993).
WILDLIFE POPULATION IMPACTS Little is known about the impact of filarioids on bird populations and this aspect of their biology has been rarely studied. Even with S. eurycerca, which is the best known pathogen among all avian filarioids, the overall conclusion is that “this parasite has not received
Figure 26.15. Morphology of microfilariae of Pelecitus tubercauda, Giemsa-stained thin smear. Note as given in Figure 26.2. Adapted from Figure 3 in Vanderburgh et al. (1984) and reproduced with permission of the Canadian Journal of Zoology.
sufficient study for its full host range, its relative frequency of occurrence in different species, or its significance as a mortality factor for wild birds to be determined” (Cole 1999). Lead poisoning, gunshot wounds and other trauma, an abundance of lice, and the presence of other nematode parasites and pathogens confound efforts to determine the impact of S. eurycerca on the health of individual wild birds as well as populations (Quortrup and Holt 1940; Cowan 1946; Holden and Sladen 1968; Irwin 1975; McKelvey and MacNeill 1981; Cohen et al. 1991). Two studies attempted to document population impacts from filarioids in galliforms. Data were inconclusive about the impact of S. californiensis, a heartworm, in populations of California Quail (Callipepla californica) (Weinmann et al. 1979).
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Filarioid Nematodes Splendidofilaria smithi (identified as “Splendidofilaria papillocerca”) in Willow Ptarmigan (Lagopus lagopus) was negatively correlated with population growth rates of the host (Holmstad et al. 2005). TREATMENT AND CONTROL Infections with avian filarioids are rarely of concern and usually no treatment is indicated. Levamisole hydrochloride was ineffective in eliminating MF from the blood of an otherwise clinically normal Marabou Stork in a zoo (Ensley 1978). Use of levamisole, fenbendazole, and mebendazole has been suggested for psittaciformes (Paster 1983). Treatment of Emus with ivermectin appeared to prevent clinical signs (Law et al. 1993) and this compound has been used to treat an Alexandrine Parakeet (Psittacula eupatria) that had adult worms removed from one eye (Kummerfeld and Daugschies 1989). Drugs that kill adult worms in situ may lead to more severe consequences than those posed by the original infection since dead worms may initiate inflammatory responses and lesions that adversely affect normal physiological function. Nonintervention is likely best and is consistent with the fact that neither disease nor pathology has been reported for the vast majority of the 160 known species of avian filarioids. ACKNOWLEDGMENTS This chapter is dedicated to the late Roy C. Anderson, who made substantial contributions to the taxonomy, classification, and biology of the avian filarioids. The author acknowledges, with gratitude, the invaluable assistance of Prune Harris and Kristy Read in the preparation of this chapter and of Cathy Chisholm in the facilitation of online access to the CABI database. LITERATURE CITED Allen, J. L., G. V. Kollias, E. C. Greiner, and W. Boyce. 1985. Subcutaneous filariasis (Pelecitus sp.) in a yellow-collared macaw (Ara auricallis). Avian Diseases 29:891–894. Anderson, R. C. 1956. The life cycle and seasonal transmission of Ornithofilaria fallisensis Anderson, a parasite of domestic and wild ducks. Canadian Journal of Zoology 34:485–525. Anderson, R. C. 1957. Taxonomic studies on the genera Aproctella Cram, 1931 and Carinema Pereira and Vaz, 1933 with a proposal for a new genus Pseudoproctella n. gen. Canadian Journal of Zoology 35:25–33. Anderson, R. C. 1968. The simuliid vectors of Splendidofilaria fallisensis of ducks. Canadian Journal of Zoology 46:610–611.
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Anderson, R. C. 2000. Nematode Parasites of Vertebrates, Their Development and Transmission, 2nd ed. CABI Publishing, Oxon, UK. Anderson R. C., and O. Bain. 1976. Keys to genera of the order Spirurida. Part 3: Diplotriaenoidea, Aproctoidea and Filarioidea. In CIH Keys to the Nematode Parasites of Vertebrates, R. C. Anderson, A. G. Chabaud, and S. Willmott (eds). 3:59–116. Commonwealth Agricultural Bureaux, Farnham Royal, WK. Anderson, R. C., and C. M. Bartlett. 1994. Ephemerality and reproductive senescence in avian filarioids. Parasitology Today 10:33–35. Anderson, R. C., A. G. Chabaud, and S. Wilmott. 1974. Key 1, General Introduction, Glossary of Terms, and Keys to Subclasses, Orders and Superfamilies. CIH Keys to the Nematode Parasites of Vertebrates 1:1–17. Atchley, W. R., and W. W. Wirth. 1975. Two western Culicoides (Diptera: Ceratopogonidae) which are vectors of filaria in the California Valley quail. Canadian Journal of Zoology 53:1421–1423. Bain, O. 1980. Deux filaires du genre Eufilaria chez le merle: d´eveloppement chez Culicoides nubeculosus. Annales de Parasitologie Humaine et Compar´ee 56:527–530. Bain, O., G. Petit, W. J. Kosek, and A. G. Chabaud. 1981. Sur les filaires Splendidofilariinae du genre Aproctella. Annales de Parasitologie Humaine et Comparee 56:95–105. Bain, O., A. G. Chabaud, and W. P. Burger. 1992. Versternema struthionis n. gen., n. sp., filaire archaique a` morphologie peu sp´ecialis´ee. Annales de Parasitologie Humaine et Compar´ee 76:141– 143. Bartlett, C. M. 1992. New, known, and unidentified species of Eulimdana (Nematoda): Additional information on biologically unusual filarioids of charadriiform birds. Systematic Parasitology 23:209–230. Bartlett, C. M. 1993. Lice (Amblycera and Ischnocera) as vectors of Eulimdana spp. (Nematoda: Filarioidea) in charadriiform birds and the necessity of short reproductive periods in adult worms. Journal of Parasitology 79:85–91. Bartlett, C. M., and R. C. Anderson. 1980a. Filarioid nematodes (Filarioidea: Onchocercidae) of Corvus brachyrhynchos in southern Ontario, Canada and a consideration of the epizootiology of avian filariasis. Systematic Parasitology 2:77–102. Bartlett, C. M., and R. C. Anderson. 1980b. Development of Chandlerella chitwoodae Anderson, 1961 (Filarioidea: Onchocercidae) in Culicoides stilobezzioides Foote and Pratt and C. travisi Vargas (Diptera: Ceratopogonidae). Canadian Journal of Zoology 58:1002–1006.
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Bartlett, C. M., and R. C. Anderson. 1981. Occult filariasis in crows (Corvus brachyrhynchos brachyrhynchos Brehm) infected with Splendidofilaria caperata Hibler, 1964 (Nematoda: Filarioidea). Journal of Wildlife Diseases 17: 69–77. Bartlett, C.M., and , R.C. Anderson. 1985. On the filarioid nematodes (Splendidofilaria spp.) from the pulmonary arteries of birds. Canadian Journal of Zoology 63:2373–2377. Bartlett, C. M., and R. C. Anderson. 1986. Paronchocerca struthionus n.sp. (Nematoda: Filarioidea) from ostriches (Struthio camelus), with a redescription of Paronchocerca ciconiarum Peters, 1936 and a review of the genus. Canadian Journal of Zoology 64:2480–2491. Bartlett, C. M., and R. C. Anderson. 1987a. Lemdana wernaarti n.sp. and other filarioid nematodes from Bubo virginianus and Asio otus (Strigiformes) in Ontario, Canada, with a revision of Lemdana and a key to avian filarioid genera. Canadian Journal of Zoology 65:1100–1109. Bartlett, C. M., and R. C. Anderson. 1987b. Pelecitus fulicaeatrae (Nematoda: Filarioidea) of coots (Gruiformes) and grebes (Podicipediformes): Skin-inhabiting microfilariae and development in Mallophaga. Canadian Journal of Zoology 65:2803–2812. Bartlett, C. M., and R. C. Anderson. 1987c. Chandlerella bushi n.sp. and Splendidofilaria caperata Hibler, 1964 (Nematoda: Filarioidea) from Fulica americana (Gruiformes: Rallidae) in Manitoba, Canada. Canadian Journal of Zoology 65:2799–2802. Bartlett, C. M., and R. C. Anderson. 1989. Mallophagan vectors and the avian filarioids: New subspecies of Pelecitus fulicaeatrae (Nematoda: Filarioidea) in sympatric North American hosts, with development, epizootiology, and pathogenesis of the parasite in Fulica americana (Aves). Canadian Journal of Zoology 67:2821–2833. Bartlett, C. M., and R. C. Anderson. 1990. Eulimdana florencae n.sp. (Nematoda: Filarioidea) from Micropalama himantopus (Aves: Charadriiformes): Evidence for neo-natal transmission, ephemerality of adults, and longevity of microfilariae among filarioids of shorebirds. Canadian Journal of Zoology 68:986–992. Bartlett, C. M., and O. Bain. 1987. New avian filarioids (Nematoda: Splendidofilariinae): Dessetfilaria guianensis gen.n., sp.n., Andersonfilaria africanus gen.n., sp.n., and Splendidofilaria chandenieri sp.n. Proceedings of the Helminthological Society of Washington 54:1–14. Bartlett, C. M., and E. C. Greiner. 1986. A revision of Pelecitus Railliet and Henry, 1910 (Filarioidea:
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techniques for detecting microfilariae in avian blood. Ibis 121(1):104–106. Seegar, W. S., E. L. Schiller, W. J. L. Sladen, and M. Trpis. 1976. A mallaghaga, Trinoton anserinum, as a cyclodevelopmental vector for a heartworm parasite of waterfowl. Science 194:739–741. Sonin, M. D. 1966. Filariata of animals and man and diseases caused by them. Part I. Aproctoidea. In Essentials of nematodology, Vol. XVII, K. I. Skrjabin (ed.). Nauka Publishers, Moscow. (Translated by the Israel Program of Scientific Translations, Jerusalem, 1974.) 365 pp. Sonin, M. D. 1968. Filariata of animals and man and diseases caused by them. Part 2. Diplotriaenoidea. In Essentials of Nematology, Vol. XXI, K. I. Skrjabin (ed.). Nauka Publishers, Moscow. (Translated by the Israel Program for Scientific Translations, Jerusalem, 1975.) 411 pp. Vanderburgh, D. J., R. C. Anderson, and T. M. Stock. 1984. Pelicitus tubercauda n. sp. (Nematoda: Filarioidea) from Geothlypis trichas L. and a redescription of P. fulicaeatrae (Diesing, 1861) Lopez-Nevra, 1956. Canadian Journal of Zoology 62:362–367. Weinmann, C. J., K. Murphy, J. R. Anderson, J. C. DeMartini, W. M. Longhurst, and G. Connolly. 1979. Seasonal prevalence, pathology, and transmission of the quail heartworm, Splendidofilaria californiensis (Wehr and Herman, 1956), in northern California. (Nematoda: Filarioidae). Canadian Journal of Zoology 57:1871–1877. Woo, P. T. K. 1971. Evaluation of the haematocrit centrifuge and other techniques for the field diagnosis of human trypanosomiasis and filariasis. Acta Tropica 28:298–303.
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27 Capillarid Nematodes Michael J. Yabsley INTRODUCTION Avian parasites that belong to the genera Baruscapillaria, Capillaria, Echinocoleus, Eucoleus, Ornithocapillaria, Pterothominx, and Tridentocapillaria are collectively referred to as the capillarids. The capillarids are small thin nematodes in the superfamily Trichinelloidea, family Trichuridae, and subfamily Capillarinae, and are related to Trichinella and Trichuris. The taxonomy of this group has changed extensively but currently there are 24 genera of capillarids that infect all classes of vertebrates. These parasitize various tissues including the gastrointestinal tract, respiratory system, urinary bladder, subcutaneous tissues, and liver. Representatives from seven of these genera infect birds and all of them parasitize regions of the gastrointestinal tract, either singly or in combination, which may be specific for particular capillarid species. For example, Eucoleus annulatus and Eucoleus contortus are found in the crop and esophagus of chickens, Pterothominx caudinflata, Pterothominx bursata, and Baruscapillaria obsignata are found in the small intestine, and Capillaria anatis is found in the ceca. Although prevalence of infection in some avian species can be high, clinical disease among freeranging birds is uncommon and effects are more at the level of an individual than at the level of entire populations. This is likely because the intensity of infection is often low. The life cycle of many capillarids is direct, so there is a high risk of transmission and disease among captive birds in poorly maintained facilities where worm burdens can be high. The natural history of several species that infect domesticated birds is well known, but the ecology of capillarids that infect wild birds is poorly understood. Detailed morphologic examination is necessary to identify capillarids to the proper genus and species. Their lack of distinguishing characteristics has led to numerous revisions in classification and large numbers of synonyms for recognized species.
SYNONYMS Capillariasis, hairworm, threadworm, cropworm. HISTORY The capillarids have historically been placed in numerous genera. Many of the early species descriptions were inadequate and based on only a few specimens or, in some cases, fragments of worms. This led to extensive revisions in taxonomy and changes in our understanding of capillarid–host relationships since C. anatis was first described from a clinically normal domestic duck over 200 years ago (see Gower 1939). Skrjabin et al. (1957) conducted an extensive review of capillarid taxonomy and proposed five genera: Capillaria, Eucoleus, Hepaticola, Skrjabinocapillaria, and Thominx. During the next 20 years, 15 additional genera were described. Anderson and Bain (1982) synonymized all 20 genera of capillarids into the genus Capillaria because of confusion stemming from morphologic criteria that were used to define genera. Although none of the proposed schemes are universally accepted, the classification proposed by Moravec (1982) and used for subsequently described genera is the most widely accepted. This taxonomy will be retained in this chapter. Twenty-four genera have been proposed to date (Moravec 1982; Moravec et al. 1987; Barus and Sergejeva 1990a, b; De and Maity 1995; Moravec et al 1999): Amphibiocapillaria, Aonchotheca, Baruscapillaria, Calodium, Capillaria, Capillostrongyloides, Crocodylocapillaria, Echinocoleus, Eucoleus, Freitascapillaria, Huffmanela, Indocapillaria, Ornithocapillaria, Paracapillaria, Paracapillaroides, Paratrichosoma, Pearsonema, Piscicapillaria, Pseudocapillaria, Pseudocapillarioides, Pterothominx, Schulmanela, Tenoranema, and Tridentocapillaria. The capillarid species that infect birds are found in seven of these genera: Baruscapillaria, Capillaria, Echinocoleus, Eucoleus, Ornithocapillaria, Pterothominx, and Tridentocapillaria. Where older species names are still commonly
463 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Parasitic Diseases of Wild Birds
used, I have listed the currently accepted name first and followed it by the old name. Because the taxonomy is confusing, some proposed synonyms are not included because most scientists consider them to be valid species. For example, Madsen (1951) and Cram (1936) consider Eucoleus perforans and E. annulatus to be synonyms of E. contortus, but most researchers accept all three as distinct species. DISTRIBUTION AND HOST RANGE The majority of capillarid species have been reported from two or more related avian species and a few are known to infect a wide range of avian host species (e.g., E. contortus infects birds in nine orders). These highly adaptive species have a cosmopolitan distribution because of the wide geographic range of the numerous hosts they infect. Some capillarids have a narrower host range and are only found in areas where suitable hosts are present. For example, Pterothominx moraveci infects only Port Lincoln Parrots (Barnardius zonarius) while Eucoleus frugilegi infects a range of species in the family Corvidae. Reports of avian genera infected with six cosmopolitan species are shown in Table 27.1. The remaining capillarid species have a narrower host range and a detailed list of these capillarid species and hosts is given in Table 27.2. Table 27.2 only includes reports based on identification of adult worms to species. Reports of adults identified to level of genus, for example, “Capillaria sp.,” or based on detection of capillarid eggs in feces are not included. ETIOLOGY Capillarids are small hairlike nematodes that parasitize the gastrointestinal tract of all classes of vertebrates. Capillarids have an esophagus that is characteristic of nematodes in the superfamily Trichinelloidea. The esophagus has a short muscular anterior part and a long glandular posterior portion (stichosome) that is composed of cuboidal gland cells called stichocytes (Figure 27.1). Eggs of trichurids and capillarids are also distinct in that they have bipolar plugs and may be nearly clear to a deep golden color; the surface ranges from smooth to variably textured (Figure 27.2). EPIZOOTIOLOGY Detailed life history traits are primarily known for those species that infect domestic fowl. With few exceptions, for example, Pterothominx philippinensis, unembryonated eggs are passed in the feces and develop to the first larval stage (L1) in 9–14 days. For
those species with a direct life cycle, eggs containing L1 larvae are infective to the next avian host. For capillarids with indirect life cycles, various species of earthworms serve as required intermediate hosts. One exception, however, is P. philippinensis, which uses fish as a required intermediate host. After eggs are ingested by an earthworm, they hatch in the gastrointestinal tract and larvae migrate through tissues of the earthworm until it is ingested by an appropriate avian host. Among some capillarid species, larvae are immediately infective after hatching in the earthworm, while in other species further development of the larvae must occur in the earthworm prior to ingestion by an avian host. If an earthworm is required in the life cycle, eggs must pass through the worm before they hatch (Moravec et al. 1987; McDougald 2003). Interestingly, an extreme female sex bias has been observed in capillarid infections of birds and other vertebrates. Since most birds infected with capillarids have few adult worms, a bias that favors adult females will increase parasite reproductive potential. The mechanism by which this sex bias develops is unknown (Lewis 1968; Joy and Scott 1997; Poulin 1997; Martin et al. 2006). The study of capillarids in wild birds is complicated by the lack of knowledge of life cycles and taxonomy of many species. Many studies are restricted to reports of capillarid eggs in fecal samples. Fecal surveys likely underestimate the true prevalence of capillarid infection because most free-ranging birds have very low-intensity infections and low fecal egg counts (Prestwood 1968). Ecological studies based on fecal egg detection are limited further because the capillarid species cannot be identified by egg morphology, coinfections with multiple capillarid species may be missed, and coinfecting capillarid species may use different life cycle strategies. The effects of age, sex, and season on capillarid infection seem to vary greatly depending on the geographic region and host and parasite species. For example, greater numbers of adult Wild Turkeys (Meleagris gallopavo) and Rock Pigeons (Columba livia) are infected with Capillaria spp. ( Prestwood 1968) while more young American Woodco*cks (Scolopax minor) and Clapper Rails (Rallus longirostris) are infected with Capillaria spp. (Heard 1967; Pursglove 1969). Senlik et al. (2005) reported that adult pigeons and those sampled in the autumn were more likely to be infected with B. obsignata than fledglings or nestlings (Senlik et al. 2005). Seasonal trends were not associated with capillarid infections in chickens in Tanzania or infections of P. caudinflata in Willow Ptarmigan (Lagopus lagopus) from Norway (Permin et al. 1997; Schei et al. 2005), but were associated with infections of
Cecum, sometimes in the small intestine
Esophagus and crop
Pharynx, esophagus, and crop
Capillaria anatis
Eucoleus annulatus
Eucoleus contortus†
465
Small intestine Small intestine
* References: Walton (1923), Gower (1939), Read (1949), Madsen (1951), Yamaguti (1961), Hodasi (1963), Wakelin (1967), Kellogg and Prestwood (1968b), Hair and Forrester (1970), Wehr (1971), Gibson (1972), Moravec (1982), Moravec et al. (1987), Barus and Sergejeva (1989), Barus and Sergejeva (1990c), Barus and Sergejeva (1990d), Ching (1990), Uchida et al. (1991), Borgsteede et al. (2003), Forrester and Spalding (2003), McDougald (2003). † Some researchers (see Barus and Sergejeva 1989) consider Eucoleus contortus to be a complex of at least two species—Eucoleus contortus from the oral cavity and esophagus of aquatic birds in the orders Anseriformes, Charadriiformes, Ciconiiformes, Pelecaniformes, and Podicipediformes and Eucoleus dispar from the oral cavity and esophagus of terrestrial birds. Other researchers consider these two species to occur in both aquatic and terrestrial birds.
Pterothominx bursata Pterothominx caudinflata
Avian genera reported as host
Aix, Anas, Anser, Aythya, Branta, Bucephala, Chen, and Lophodytes Alectoris, Meleagris, Perdix, and Phasianus Attila, Corvus, Garrulus, and Pica Anas and Cairina Alectoris, Bonasa, Colinus, Gallus, Lyrurus, Meleagris, Numida, Perdix, Phasianus, and Tetrao Anseriformes Aix, Anas, Aythya, Bucephala, Clangula, Netta, Somateria, and Tadorna Charadriiformes Actitis, Alle, Calidris, Capella, Charadrius, Chlidonias, Gallinago, Gelochelidon, Himantopus, Larus, Pavoncella, Philomachus, Pluvialis, Rissa, Tringa, Recurvirostra, Sterna, Thalasseus, Uria, and Vanellus Ciconiiformes Botaurus, Cochlearius, Eudocimus, Plegadis, and Platalea Falconiformes Accipiter, Buteo, Falco, and Haliaeetus Galliformes Alectoris, Bonasa, Colinus, Crossoptilon, Lofortyx, Meleagris, Oreortyx, Perdix, and Phasianus Gruiformes Crex, Fulica, and Gallinula Passeriformes Corvus, Cyanocitta, Erithacus, Passerella, Phoenicurus, and Sturnus Pelecaniformes Pelecanus and Phalacrocorax Podicipediformes Podiceps and Tachybaptus Anseriformes Aix, Branta, Chen, and Anser Ciconiiformes Ardea Columbiformes Columba and Zenaida Galliformes Colinus, Coturnix, Gallus, Meleagris, Pavo, Perdix, and Zenaida Piciformes Ramphastos Psittaciformes Agap*rnis, Barnardius, and Melopsittacus Galliformes Gallus, Meleagris, Numida, Perdix, Phasianus, and Tetrao Anseriformes Anas, Anser, Branta, Cairina, and Chen Columbiformes Columba and Streptopelia Galliformes Alectoris, Bonasa, Callipepla, Coturnix, Chrysolophus, Gallus, Lagopus, Meleagris, Numida, Perdix, Phasianus, and Tetrao Gruiformes Otis Passeriformes Erithacus, Ixoreus, Sturnus, Passer, and Turdus Tinamiformes Nothoprocta Anseriformes Galliformes Passeriformes Anseriformes Galliformes
Avian order
September 29, 2008
Baruscapillaria obsignata Small intestine and cecum
Location in host
Parasite species
Table 27.1. Genera of birds infected with six cosmopolitan capillarids species.*
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466
Crop Esophagus
Small intestine Large intestine
Small intestine, cecum Small intestine, cecum Cloaca
Eucoleus penidoi Eucoleus crypturi Baruscapillaria mergi Baruscapillaria mergi Baruscapillaria ryjikovi
Cloaca
Unknown Small intestine, ceca, and large intestine Large intestine
Baruscapillaria ryjikovi (=Baruscapillaria podicipitis) baruscapillaria ryjikovi Carcharhinus perezi Ornithocapillaria carbonis Baruscapillaria spilculata
Baruscapillaria ryjikovi
Not given
Location in host
Capillaria parvumspinosa
Capillarid species
None reported
None reported
Unknown
None reported
None reported
None reported
None reported
None reported
None reported
None reported
None reported
None reported
Disease signs/lesions
USA
USA
Cuba
Czechoslovakia, Japan, USSR
USA
Czechoslovakia
USSR
Unknown
USA
Brazil
Argentina, Brazil
Europe
Location
Threlfall (1982) and Forrester and Spalding (2003) Fedynich et al. (1997)
Yamaguti (1961)
Baru˘s and Sergejeva (1990c) and Forrester and Spalding (2003) Baru˘s and Sergejeva (1990c) and Uchida et al. (1991)
Baru˘s and Sergejeva (1990c)
Baru˘s and Sergejeva (1990c)
Kinsella and Forrester (1999) Baru˘s and Sergejeva (1990c)
Freitas and Almeida (1935) and Pinto et al. (2006) Yamaguti (1961)
Yamaguti (1961)
References
September 29, 2008
Little grebe (Tachybaptus ruficollis) Pelecaniformes Anhinga (Anhinga anhinga) Double-crested Cormorant (Phalacrocorax auritus)
Struthioniformes Greater Rhea (Rhea americana) Tinamiformes Spotted Nothura (Nothura maculosa) Gray Tinamou (Tinamus tao) Gaviiformes Common Loon (Gavia immer) Red-throated Loon (Gavia stellata) Podicipediformes Red-necked Grebe (Podiceps grisegena) Eared Grebe (Podiceps nigricollis) Pied-billed Grebe (Podilymbus podiceps)
Avian host
Table 27.2. Reports of capillarids in avian hosts excluding Capillaria anatis, Eucoleus annulatus, Eucoleus contortus, Baruscapillaria obsignata, Pterothominx bursata, and Pterothominx caudinflata (Table 27.1). Only reports that included identification of adult worms to species are included. Reports of capillarid eggs in the feces or unspeciated capillarid adults are not included. Unless otherwise specified, all reports are from free-ranging birds.
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Not given Not given
Not given
Bursa of Fabricius and cloaca Not given
Not given
Small and large intestine, cecum Intestine
Baruscapillaria jaenschi Baruscapillaria jaenschi Baruscapillaria jaenschi Ornithocapillaria phalacrocoraxi
467 Baruscapillaria jaenschi Baruscapillaria jaenschi Baruscapillaria mergi Baruscapillaria mergi
None reported USA
None reported USA
None reported Australia
None reported Australia
None reported Tajikistan
None reported Australia
None reported Australia
None reported Australia
None reported Czech Republic None reported Russia, Poland, Czech Republic
Intestine Intestine
Baruscapillaria rudolphii Ornithocapillaria carbonis
None reported Brazil, Mexico, USA None reported Brazil, USA
Large intestine
Baruscapillaria spilculata Large intestine
Ornithocapillaria appendiculata
Courtney and Forrester (1974) (continues)
Kinsella et al. (2004)
Johnston and Mawson (1945)
Johnston and Mawson (1945)
Baru˘s and Sergejeva (1990b)
Johnston and Mawson (1945)
Freitas (1933b), Vicente et al. 1995, and Moravec et al. 2000 Freitas (1933b) and Fedynich et al. (1997) Moravec et al. (1994) Dubinin and Dubinina (1940), Frantova (2001), and Frantova (2002) Johnston and Mawson (1945) Johnston and Mawson (1945)
September 29, 2008
Black-faced Cormorant (Phalacrocorax fuscescens) Little Pied Cormorant (Phalacrocorax melanoleucos) Pygmy Cormorant (Phalacrocorax pygmaeus) Little Black Cormorant (Phalacrocorax sulcirostris) Australian Pelican (Pelecanus conspicillatus) American White Pelican (Pelecanus erythrorhynchos) Brown Pelican (Pelecanus occidentalis)
Great Cormorant (Phalacrocorax carbo)
Neotropic Cormorant (Phalacrocorax brasilianus)
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468
Eurasian Teal (Anas crecca) Mottled Duck (Anas fulvigula) Mallard (Anas platyrhynchos) Ring-necked Duck (Aythya collaris)
Anseriformes Wood Duck (Aix sponsa)
Unknown
Baruscapillaria mergi Capillaria brasiliana
Cecum
Intestine Cecum Cecum Intestine Esophagus Cecum
Capillaria spinulosa Pterothominx exilis Baruscapillaria mergi Capillaria spinulosa Capillaria nyrocinarum Eucoleus perforans Capillaria spinulosa
Gizzard, small intestine, and large intestine
Small intestine
Capillaria avellari
Baruscapillaria mergi
Esophagus
Baruscapillaria mergi
Location in host Small intestine Small and large intestine Small intestine
Capillaria herodiae Capillaria herodiae
Capillarid species
None reported None reported None reported
None reported
None reported
None reported
None reported
None reported
Unknown
None reported
None reported
None reported
None reported None reported
Disease signs/lesions
Japan Asia, Europe USA
USA
USA, England (captive) USSR
USA
USA
Brazil
USA
USA, Brazil
USA
USA USA
Location
Forrester and Spalding (2003) Motohiro et al. (2000) Baru˘s and Sergejeva (1989) Forrester and Spalding (2003)
Ogburn-Cahoon (1979) and Forrester and Spalding (2003) McDonald (1969) and Baru˘s and Sergejeva (1990d) Baru˘s and Sergejeva (1990c)
Sep´ulveda et al. (1994)
Yamaguti (1961) and Forrester and Spalding (2003) Forrester and Spalding (2003) Freitas (1933a) and Yamaguti (1961)
Sep´ulveda et al. (1999) Boyd (1966) and Forrester and Spalding (2003) Sep´ulveda et al. (1996)
References
September 29, 2008
Black-crowned Night-Heron (Nycticorax nycticorax) Roseate Spoonbill (Platalea ajaja)
Ciconiiformes Great Egret (Ardea alba) Great Blue Heron (Ardea herodias) Little Blue Heron (Egretta caerulea) Wood Stork (Mycteria americana)
Avian host
Table 27.2. (Continued)
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Intestine Intestine Cecum Intestine
Intestine Small and large intestine Intestine Intestine
Small and large intestine Intestine Intestine and cecum
Cecum
Small intestine
Capillaria nyrocinarum Capillaria nyrocinarum Capillaria spinulosa Baruscapillaria mergi
Capillaria nyrocinarum Baruscapillaria mergi Capillaria nyrocinarum Capillaria nyrocinarum
469 Baruscapillaria mergi Capillaria nyrocinarum Capillaria nyrocinarum
Capillaria nyrocinarum Baruscapillaria mergi
Surf Scoter (Melanitta perspicillata)
Common Merganser (Mergus merganser)
Black Scoter (Melanitta nigra)
White-winged Scoter (Melanitta fusca)
White-winged Scoter (Melanitta fusca)
None reported Denmark, USSR
None reported Canada
None reported Denmark None reported Canada, Denmark
None reported Denmark
None reported Denmark None reported Canada
None reported Denmark None reported Denmark
None reported Denmark None reported Europe None reported Canada, Denmark
None reported Denmark
Yamaguti (1961) Gower (1939) Yamaguti (1961), Mahoney and Threlfall (1978), and Baru˘s and Sergejeva (1990c) Yamaguti (1961) Gower (1939), Yamaguti (1961), and Baru˘s and Sergejeva (1990c) Yamaguti (1961) U.S. National Parasite Collection, Accession No. 076891 Yamaguti (1961) and Baru˘s and Sergejeva (1990c) Yamaguti (1961) Yamaguti (1961) and U.S. National Parasite Collection Accession No. 076911 U.S. National Parasite Collection, Accession No. 076865 Yamaguti (1961) and Baru˘s and Sergejeva (1990c) (continues)
Yamaguti (1961)
September 29, 2008
Long-tailed Duck (Clangula hyemalis)
Greater Scaup (Aythya marila) Ferruginous Pochard (Aythya nyroca) Common Goldeneye (Bucephala clangula)
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470
Eurasian Sparrow Hawk (Accipiter nisus) Capillaria tenuissima
Baruscapillaria falconis Eucoleus dispar
Small intestine, sometimes gizzard
None reported
Small intestine, sometimes gizzard Small intestine Esophagus
None reported
None reported None reported
None reported
Esophagus
None reported
None reported
Small intestine
Capillaria gigantoteca (some authors consider this a synonum of Baruscapillaria obsignata) Capillaria droummondi
None reported
Intestine
Intestine
Capillaria ellisi
None reported None reported
Unknown
Small intestine Intestine
Baruscapillaria mergi Capillaria nyrocinarum
None reported
None reported
Disease signs/lesions
Unknown
Intestine
Capillaria nyrocinarum
Black-necked Swan (Cygnus melancoryphus) Mute Swan (Cygnus olor) Capillaria pudendotecta Falconiformes Northern Goshawk Eucoleus dispar (Accipiter gentilis) (sometimes as Trichosomum contorta) Capillaria tenuissima
Small intestine
Location in host
Baruscapillaria mergi
Capillarid species
Spain, Germany, Czech Republic
Czech Republic Spain
Germany
Germany, Poland, Spain
Russia
Brazil
Russia
Australia
Denmark Denmark
Netherlands, Denmark, Canada
Denmark
Location
Frantova (2002) Ferrer et al. (2004) and Sanmartin et al. (2004) Krone (2000), Frantova (2002), and Sanmartin et al. (2004)
Okulewicz (1988), Krone (2000), and Ferrer et al. (2004) Krone (2000)
Yamaguti (1961)
Yamaguti (1961)
Johnston and Mawson (1945) Yamaguti (1961)
Yamaguti (1961), Bishop and Threlfall (1974), and Borgsteede et al. (2005) Yamaguti (1961) Yamaguti (1961)
Yamaguti (1961)
References September 29, 2008
King Eider (Somateria spectabilis) Black Swan (Cygnus atratus) Tundra Swan (Cygnus columbianus)
Red-breasted Merganser (Mergus serrator) Common Eider (Somateria mollissima)
Avian host
Table 27.2. (Continued)
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471 None reported Spain
Esophagus Esophagus
Esophagus None reported Spain Small intestine, None reported Spain sometimes gizzard
Eucoleus dispar Eucoleus dispar Eucoleus dispar Capillaria tenuissima
Short-toed Eagle (Circaetus gallicus) Western Marsh-Harrier (Circus aeruginosus) Montagu’s Harrier (Circus pygargus)
None reported Germany, Spain
None reported USA
None reported USA
Eucoleus dispar (reported Esophagus as Capillaria contorta) Baruscapillaria falconis Small intestine
None reported USA
Red-shouldered Hawk (Buteo lineatus)
Baruscapillaria falconis Small intestine Eucoleus dispar (reported Esophagus as Capillaria contorta) Baruscapillaria falconis Small intestine
Esophagus
Eucoleus dispar None reported Netherlands, Germany, Czech Republic, Spain None reported Czech Republic None reported USA
Small intestine, None reported Netherlands, Spain, sometimes gizzard Poland, Germany
None reported Taiwan
None reported Europe
Capillaria tenuissima
Small intestine
Small intestine
(continues)
Lierz et al. (2002) and Ferrer et al. (2004) Sanmartin et al. (2004) Sanmartin et al. (2004)
Okulewicz (1988), Illescas et al. (1993), Krone (2000), Borgsteede et al. (2003), and Sanmartin et al. (2004) Lierz et al. (2002), Borgsteede et al. (2003), and Ferrer et al. (2004) Frantova (2002) Forrester and Spalding (2003) Read (1949) and Forrester and Spalding (2003) Forrester and Spalding (2003) Forrester and Spalding (2003) Ferrer et al. (2004)
Su and Fei (2004)
Read (1949) and Yamaguti (1961)
September 29, 2008
Red-tailed Hawk (Buteo jamaicensis)
Crested Goshawk (Accipiter trivirgatus) Eurasian Buzzard (Buteo buteo)
Baruscapillaria falconis (as Trichosomum contortum) Baruscapillaria falconis
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472
Red Kite (Milvus milvus) Osprey (Pandion haliaetus) Chimango Caracara (Milvago chimango)
Bald Eagle (Haliaeetus leucocephalus)
White-tailed Eagle (Haliaeetus albicilla)
Small intestine Intestine
Baruscapillaria falconis Ornithocapillaria cylindrica Eucoleus dispar
Esophagus Small intestine Small intestine
Eucoleus dispar Baruscapillaria falconis Capillaria tenuissima
Capillaria tenuissima Small intestine Eucoleus dispar (reported Esophagus as Capillaria contorta) Baruscapillaria falconis Small intestine
Eucoleus dispar Esophagus (sometimes as Eucoleus contortus or Eucoleus suppereri) Capillaria tenuissima Small intestine, sometimes gizzard Eucoleus dispar Esophagus
Esophagus
Esophagus
Location in host
Eucoleus dispar
Capillarid species
None reported
Chile
Spain USA
USA
None reported None reported None reported
Germany USA
Germany, Finland
Spain
Spain, Germany
Spain
Cuba
USA
Germany, Spain
Location
None reported None reported
None reported
None reported
None reported
None reported
None reported
None reported
None reported
Disease signs/lesions
Martin et al. (2006)
Forrester and Spalding (2003) Sanmartin et al. (2004) Kinsella et al. (1996)
Krone et al. (2003) and Krone et al. (2006) Krone et al. (2003) Kinsella et al. (1998)
Sanmartin et al. (2004)
Lierz et al. (2002) and Sanmartin et al. (2004)
Ferrer et al. (2004)
Lierz et al. (2002) and Ferrer et al. (2004) Forrester and Spalding (2003) Baru˘s and Sergejeva (1990c)
References
September 29, 2008
Eurasian Hobby (Falco subbuteo) Eurasian Kestrel (Falco tinnunculus)
Peregrine Falcon (Falco peregrinus) American Kestrel (Falco sparverius)
Avian host
Table 27.2. (Continued)
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473 None reported
Cecum Small intestine Esophagus
Small intestine (submucosa) Esophagus Cecum Esophagus Small intestine
Capillaria phasianina Eucoleus perforans Pterothominx blomei (some consider a synonym of Pterothominx caudinflata)
Worldwide
Worldwide
USA USA Worldwide
Uruguay
Denmark (captive)
USA (captive)
Mild to severe Denmark, England necrosis, mortality None reported Worldwide None reported England, Switzerland
Severe esophagitis
None reported None reported None reported
Cecum
Baruscapillaria montevidensis Capillaria phasianina Thomix tridens Eucoleus perforans (=Capillaria or Eucoleus combologiodes) Pterothominx meleagridis Eucoleus perforans
None reported
Cecum
Capillaria phasianina
Weakness, anorexia, vomiting, thickening of mucosa, extensive damage to epithelium None reported
Esophagus
Eucoleus perforans
(continues)
Baru˘s and Sergejeva (1989) and Menezes et al. (2001) Clapham (1949) and Madsen (1951) Baru˘s and Sergejeva (1989) Baru˘s and Sergejeva (1990d)
Baru˘s and Sergejeva (1990d)
Schorr (1988) Davidson et al. (1975) Baru˘s and Sergejeva (1989)
Calzada (1937)
Madsen (1951)
De Rosa and Shivaprasad (1999)
September 29, 2008
Helmeted Guinea Fowl (Numida meleagris) Gray Partridge (Perdix perdix)
Golden Pheasant (Chrysolophus pictus) domestic chicken (Gallus gallus domestica) Wild Turkey (Meleagris gallopavo) and domesticated turkey
Galliformes Vulturine Guinea Fowl (Acryllium vulturinum)
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Ring-necked Pheasant (Phasianus colchicus)
Avian host
Table 27.2. (Continued)
474 Crop and esophagus Small intestine (submucosa) Small intestine
Capillaria uropapillata Pterothominx meleagridis Pterothominx blomei (some consider a synonym of Pterothominx caudinflata)
Small intestine and cecum
Cecum
Echinocoleus cyanopicae (has only been reported in Cyanopica cyaneus, this may be a misidentification)
None reported
None reported
Lethargy, emaciation, diarrhea, thickened, small nodules, congestion, and petechial hemorrhage of the mucosa Necrosis of epithelium and glandular crypts, some inflammation in the mucosa None reported (low burdens) to necrosis, fibrosis, sloughing of the epithelium (heavy burdens) None reported
Disease signs/lesions
Worldwide Poland, Brazil, Romania, Denmark, Hungary (captive, domesticated) Brazil (captive, domesticated) Worldwide (captive, domesticated) England, Switzerland (captive, domesticated)
Romania (captive, domesticated)
Brazil (captive, domesticated)
Location
Baru˘s and Sergejeva (1990d)
Baru˘s and Sergejeva (1990d)
Madsen (1951), Kellogg and Prestwood (1968a), Iulia and Pavlovic (2003), Pinto et al. (2004), Tampieri et al. (2005), and Gassal and Schm¨aschke (2006) Pinto et al. (2004)
Iulia and Pavlovic (2003)
Pinto et al. (2004) and Gassal and Schm¨aschke (2006)
References September 29, 2008
Capillaria phasianiana
Crop and esophagus
Location in host
Eucoleus perforans
Capillarid species
BLBS014-Atkinson 16:25
Gizzard Cecum Cecum Gizzard Small intestine, cecum, and large intestine Gizzard Gizzard Cecum
Intestine Intestine Large intestine Intestine
Eucoleus obtusiuscula Capillaria fulicae Capillaria fulicae Eucoleus obtusiuscula Capillaria fulicae
Eucoleus obtusiuscula Capillaria fulicae Pterothominx totani Pterothominx totani Capillaria cecumitis Pterothominx totani
Eucoleus obtusiuscula
Pterothominx alpina
Duodenum near pylorus Duodenum near pylorus
Pterothominx alpina
475 None reported USSR
None reported USA
None reported Canada
None reported Europe, USSR
None reported USA
None reported Europe
None reported USA
None reported USA (captive) None reported USA (captive)
None reported USA
None reported USA
None reported Africa
None reported Austria, Germany
None reported Austria, Germany
(continues)
Forrester and Spalding (2003) Baru˘s and Sergejeva (1990d)
Yamaguti (1961) and Baru˘s and Sergejeva (1990d) Gibson (1972)
Forrester and Spalding (2003)
Yamaguti (1961)
Spalding et al. (1996)
Forrester and Spalding (2003) Spalding et al. (1996) Forrester and Spalding (2003)
Kinsella (1973)
Baru˘s and Sergejeva (1989)
Baru˘s and Sergejeva (1990d) and Fischbacher (2007) Baru˘s and Sergejeva (1990d) and Fischbacher (2007)
September 29, 2008
Sandhill Crane (Grus canadensis) Common Crane (Grus grus) Purple Gallinule (Porphyrio martinica) Charadriiformes Common Sandpiper (Actitis hypoleucos) Spotted Sandpiper (Actitis macularius) Red Knot (Calidris canutus) Common Ringed Plover (Charadrius hiaticula)
Black Grouse (Tetrao tetrix) Eurasian Capercaillie (Tetrao urogallus) Gruiformes Black-crowned-Crane (Balearica pavonina) American Coot (Fulica americana) Common Moorhen (Gallinula chloropus) Whooping Crane (Grus americana)
BLBS014-Atkinson 16:25
476
Kea (Nestor notabilis)
Small intestine
Pterothominx moraveci Not given
Not given
Capillaria plagiaticia
Capillaria plagiaticia
Intestine
Capillaria vasi
Small intestine
Capillaria recurvirostrae
Intestine
Not given
Pterothominx totani
None reported
Small intestine
Baruscapillaria belopolskaiae Baruscapillaria jaenschi
Gizzard
None reported
Large intestine Large intestine
Capillaria cecumitis Capillaria cecumitis
Eucoleus obtusiuscula
None reported
Not given
None reported
None reported
None reported
None reported
None reported
None reported
None reported
None reported None reported
None reported
Disease signs/lesions
Gizzard
Location in host
Eucoleus obtusiuscula (=Capillaria vanelli) Baruscapillaria jaenschi
Capillarid species
Captive (Germany/Czech Republic) New Zealand (captive)
Brazil (captive)
Brazil
USSR
Europe, Tunisia
USA
Australia
Europe
USA USA
Australia
USA
Location
Wakelin (1967)
Baru˘s et al. (2005)
Freitas et al. (1959)
Madsen (1951)
Johnston and Mawson (1945) Ahern and Schmidt (1976) and Garcia and Canaris (1987) Yamaguti (1961), Bernard (1989), and Baru˘s and Sergejeva (1989) Baru˘s and Sergejeva (1990d)
Baru˘s and Sergejeva (1990c)
Forrester and Spalding (2003) Johnston and Mawson (1945) Ching (1990) Ching (1990)
References
September 29, 2008
Terek Sandpiper (Xenus cinereus) Columbiformes Spot-winged Wood-Quail (Odontophorus capueira) Psittaciformes Caatinga Parakeet (Aratinga cactorum) Port Lincoln Parrot (Barnardius zonarius)
Killdeer (Charadrius vociferus) Whiskered Tern (Chlidonias hybrida) Dunlin (Calidris alpina) Willet (Tringa semipalmata inornata) Common Snipe (Gallinago gallinago) Silver Gull (Larus novaehollandiae) American Avocet (Recurvirostra americana) Northern Lapwing (Vanellus vanellus)
Avian host
Table 27.2. (Continued)
BLBS014-Atkinson 16:25
477
Capillaria tenuissima Eucoleus dispar Capillaria tenuissima
Australian Masked-Owl (Tyto novaehollandiae)
Described as Capillaria strigis (hom*onym so renamed Capillaria newzealandica)
Small intestine Small intestine Esophagus Small intestine Small intestine Small intestine, sometimes gizzard Not given
Small intestine Esophagus Small intestine, sometimes gizzard Small intestine
Baruscapillaria falconis
Spotted Owl (Strix Baruscapillaria falconis occidentalis) Ural Owl (Strix uralensis) Capillaria tenuissima Capillaria tenuissima Barred Owl (Strix varia) Eucoleus dispar Baruscapillaria falconis Capillaria tenuissima Barn Owl (Tyto alba) Capillaria tenuissima
Small intestine
Baruscapillaria falconis
Japan Austria USA USA USA France, Germany None reported New Zealand
None reported None reported None reported None reported None reported None reported
None reported USA
None reported USA None reported Poland None reported Austria, Germany
None reported Canada, USA
Small intestine, None reported Austria, Germany sometimes gizzard Small intestine None reported Spain
Capillaria tenuissima
None reported USA
Small intestine, None reported Austria, Germany sometimes gizzard Small intestine None reported Spain, USA
Small intestine
(continues)
Uchida et al. (1991) Kutzer et al. (1982) Kinsella et al. (2001) Kinsella et al. (2001) Kinsella et al. (2001), Thebault (1988), and Krone (2000) Johnston and Mawson (1944) and Yamaguti (1961)
Read (1949), Ramalingam and Samuel (1978), and Kinsella et al. (2001) Kinsella et al. (2001) Okulewicz (1988) Kutzer et al. (1982) and Krone (2000) Hoberg et al. (1989)
Kutzer et al. (1982) and Krone (2000) Read (1949) and Illescas et al. (1993) Kutzer et al. (1982) and Krone (2000) Illescas et al. (1993)
Read (1949)
September 29, 2008
Tawny Owl (Strix aluco)
Little Owl (Athene noctua) Eurasian Eagle-Owl (Bubo bubo) Great Horned Owl (Bubo virginianus)
Strigiformes Northern Saw-whet Owl Baruscapillaria falconis (Aegolius acadicus) Northern Long-eared Owl Capillaria tenuissima (Asio otus) Baruscapillaria falconis
BLBS014-Atkinson 16:25
478 None reported
Small intestine Small intestine Small intestine Small intestine
Small intestine Small intestine Small intestine Small intestine
Capillaria longistriata Thomix tridens Capillaria venusta Ornithocapillaria picorum Capillaria longistriata Thomix tridens Ornithocapillaria picorum Ornithocapillaria picorum
Ornithocapillaria picorum Tridentocapillaria eurycerca Green Woodpecker (Picus Ornithocapillaria viridis) picorum
None reported
Small intestine
Thomix tridens
None reported None reported None reported
Small intestine Small intestine Small intestine
None reported
None reported
None reported None reported None reported None reported
None reported
None reported
Intestine
Tridentocapillaria hirundinis
None reported
Disease signs/lesions
Small intestine
Location in host
Thomix tridens
Capillarid species
Europe
USSR
Europe
Brazil
Mexico
USA
USA
USA USA Cuba USA
Borneo
Europe
Philippines
Location
Walton (1923) and Yamaguti (1961)
Walton (1923) and Yamaguti (1961) Baru˘s and Sergejeva (1990a)
Walton (1923) and Yamaguti (1961)
Leidy (1856)
Forrester and Spalding (2003) Foster et al. (2002)
Walton (1923) Bolette (1998) Baru˘s (1971) Walton (1923) and Yamaguti (1961)
Baru˘s and Sergejeva (1990a)
Yamaguti (1961)
U.S. National Parasite Collection
References
September 29, 2008
Great Spotted Woodpecker (Dendrocopos major) Pileated Woodpecker (Dryocopus pileatus) Red-bellied Woodpecker (Melanerpes carolinus) Northern Flicker (Colaptes auratus) Golden-olive Woodpecker (Piculus rubiginosus) (=Colaptes maximus) Gray-faced Woodpecker (Picus canus)
Caprimulgiformes Large-tailed Nightjar (Caprimulgus macrurus) Apodiformes Common Swift (Apus apus) Piciformes Maroon Woodpecker (Blythipicus rubiginosus) Northern Flicker (Colaptes auratus)
Avian host
Table 27.2. (Continued)
BLBS014-Atkinson 16:25
479
Tree Pipit (Anthus trivialis) European Greenfinch (Carduelis chloris) Gray-cheeked Thrush (Catharus minimus) Brown Dipper (Cinclus pallasii)
None reported Cuba None reported Japan
Not given
Esophagus
Small intestine Intestine
Intestine Intestine Small intestine Small intestine
Capillaria venusta Eucoleus dispar Thomix tridens Ornithocapillaria ovopunctata Pterothominx exilis Pterothominx longifilla Baruscapillaria emberizae Ornithocapillaria ovopunctata (reported as Capillaria ornate or Baruscapillaria inflexa) Pterothominx longifilla Pterothominx exilis Ornithocapillaria ovopunctata Baruscapillaria cincli
None reported Europe
Intestine
None reported Asia, Europe
None reported France
None reported Asia, Europe None reported Czechoslovakia None reported Japan
Intestine Small intestine Small intestine
None reported USA None reported USA
None reported Europe
None reported Brazil
None reported Brazil
Not given
Capillaria venusta
None reported Brazil
Not given
Capillaria venusta
(continues)
Uchida et al. (1991)
Baru˘s and Sergejeva (1990c)
Baru˘s and Sergejeva (1990d)
Yamaguti (1961)
Yamaguti (1961)
Baru˘s and Sergejeva (1990d) Baru˘s and Sergejeva (1990d) Uchida et al. (1991)
Read (1949) Cooper and Crites (1974b)
Baru˘s and Sergejeva (1989)
Pinto et al. (1996)
Pinto et al. (1996)
Yamaguti (1961)
September 29, 2008
Eurasian Skylark (Alauda arvensis) Olive-backed Pipit (Anthus hodgsoni) Meadow Pipit (Anthus pratensis)
Black-necked Aracari (Pteroglossus aracari) Toco Toucan (Ramphastos toco) Channel-billed Toucan (Ramphastos vitellinus) Passeriformes Sedge Warbler (Acrocephalus schoenobaenus) Red-winged Blackbird (Agelaius phoeniceus)
BLBS014-Atkinson 16:25
480
Eurasian Jackdaw (Corvus monedula)
Rook (Corvus frugilegus)
Carrion Crow (Corvus corone)
Not given
Small intestine
Capillaria graucalina
Baruscapillaria resecta (=Baruscapillaria corvorum) Baruscapillaria resecta (=Baruscapillaria corvorum) Baruscapillaria resecta (=Capillaria corvorum) Eucoleus dispar Baruscapillaria resecta (=Baruscapillaria corvorum) Eucoleus dispar Baruscapillaria resecta (=Baruscapillaria corvorum) Pterothominx exilis Baruscapillaria resecta (=Baruscapillaria corvorum) None reported None reported
None reported None reported
Small intestine Intestine
None reported None reported
None reported
None reported
None reported
None reported
None reported
Disease signs/lesions
Esophagus Intestine
Esophagus Intestine
Intestine
Intestine
Small intestine
Location in host
Ornithocapillaria ovopunctata
Capillarid species
Asia, Europe Europe
Not given Europe
Europe Europe
Tunisia
Europe
Canada
Australia
Nepal
Location
Baru˘s and Sergejeva (1990d) Yamaguti (1961), Fronska-Popiel et al. (2001), and Frantova (2002)
Baru˘s and Sergejeva (1989) Baru˘s and Sergejeva (1990c) and Frantova (2002)
Baru˘s and Sergejeva (1989) Baru˘s and Sergejeva (1990c)
Bernard (1989)
Yamaguti (1961) and Baru˘s and Sergejeva (1990c)
Andrews and Threlfall (1975)
Johnston and Mawson (1941)
Baru˘s and Daniel (1976)
References September 29, 2008
Eurasian Nutcracker (Nucifraga caryocatactes) Common Raven (Corvus corax)
White-capped Redstart (Chaimarrornis leucocephalus) Black-faced Cuckoo-shrike (Coracina novaehollandiae) American Crow (Corvus brachyrhynchos)
Avian host
Table 27.2. (Continued)
BLBS014-Atkinson 16:25
None reported Asia, Europe
Small intestine Small intestine
Baruscapillaria emberizae Pterothominx exilis
Baru˘s and Sergejeva (1990d) Johnston and Mawson (1945) Johnston and Mawson (1947) Yamaguti (1961) Ching (1993)
None reported Asia, Europe None reported Australia
None reported Canada
Small intestine Not given Not given
Capillaria gymnorhinae
481 Tridentocapillaria Intestine hirundinis Ornithocapillaria quiscali Intestine
None reported Europe
None reported Australia
(continues)
Yamaguti (1961) Uchida et al. (1991)
Uchida et al. (1991)
Unknown Russia None reported Japan
None reported Japan
Small intestine Unknown Intestine
None reported Japan
Small intestine
Uchida et al. (1991)
Johnston and Mawson (1944) Baru˘s and Sergejeva (1990d)
Baru˘s and Sergejeva (1990a)
Baruscapillaria emberizae Baruscapillaria emberizae Capillaria avicola Baruscapillaria resecta (=Baruscapillaria corvorum, sometimes reported as Capillaria anatis) Pterothominx exilis Baruscapillaria grallinae
None reported Cuba
Lopez-Neyra (1947) and Baru˘s and Sergejeva (1989) Baru˘s and Sergejeva (1990a)
September 29, 2008
Magpie-lark (Grallina cyanoleuca) Australasian Magpie (Gymnorhina tibicen) Barn Swallow (Hirundo rustica) Varied Thrush (Ixoreus naevius)
None reported New Zealand
Small intestine
Thomix tridens
None reported Cuba
Small intestine
Thomix tridens
Yellow-throated Warbler (Dendroica dominica) Blackburnian Warbler (Dendroica fusca) Yellowhammer (Emberiza citrinella) Reed Bunting (Emberiza schoeniclus) Yellow Bunting (Emberiza sulphurata) Gray Bunting (Emberiza variabilis) Eurasian Jay (Garrulus glandarius)
None reported Spain
Cecum
Echinocoleus cyanopicae
Azure-winged Magpie (Cyanopica cyanus)
BLBS014-Atkinson 16:25
482
USA, Europe
Not given USA
None reported
None reported None reported None reported None reported None reported
None reported None reported
Small intestine
Ornithocapillaria Intestine ovopunctata (reported Baruscapillaria inflexa) Thomix tridens Small intestine Tridentocapillaria parusi Small intestine Intestine Small intestine
Thomix tridens
Capillaria rigidula Capillaria freitasi Baruscapillaria corvorum Intestine (=Capillaria resectum, sometimes reported as Capillaria anatis) Eucoleus dispar Esophagus Baruscapillaria corvorum Small intestine (reported as Capillaria anatis)
Black-billed Magpie (Pica hudsonia)
Great Tit (Parus major) Green-backed Tit (Parus monticolus) Spanish Sparrow (Passer hispaniolensis) Fox Sparrow (Passerella iliaca) Eurasian Magpie (Pica pica pica)
None reported
Small intestine
Thomix tridens
USA
Tunisia
Poland Taiwan
Europe
USA
Cuba
France, Poland
Baru˘s and Sergejeva (1989) Todd et al. (1967)
Yamaguti (1961) and Todd and Worley (1967)
Read (1949)
Bernard (1989)
Okulewicz (1991) Baru˘s and Sergejeva (1990a)
Yamaguti (1961)
Cooper et al. (1973)
Baru˘s and Sergejeva (1990a)
Yamaguti (1961) and Okulewicz (1982)
Hromada et al. (2000) and U.S. National Parasite Collection Uchida et al. (1991)
References September 29, 2008
None reported
None reported
Small intestine
Thomix tridens
Japan
None reported
Baruscapillaria calliopsis Small intestine
Siberian Rubythroat (Luscinia calliope) Common Nightingale (Luscinia megarhynchos) Black-and-white Warbler (Mniotilta varia) Brown-headed Cowbird (Molothrus ater) Blue Rock-Thrush (Monticola solitarius)
Russia, Philippines
Location
None reported
Disease signs/lesions
Small intestine
Location in host
Thomix tridens
Capillarid species
Long-tailed Shrike (Lanius schach)
Avian host
Table 27.2. (Continued)
BLBS014-Atkinson 16:25
483
Eastern Bluebird (Sialia sialis) Eurasian Nuthatch (Sitta europaea)
Yellow-billed Chough (Pyrrhocorax graculus) Common Grackle (Quiscalus quiscula)
None reported Cuba
None reported Nepal None reported Nepal None reported France None reported Czechoslovakia
Small intestine Not given
Small intestine Large Intestine Intestine Small intestine Small intestine Small intestine
Capillaria pomatostomi Ornithocapillaria ovopunctata Capillaria rigidula Capillaria rigidula Pterothominx longifilla Pterothominx exilis Ornithocapillaria ovopunctata
None reported Taiwan None reported Asia, Europe
Tridentocapillaria parusi Pterothominx longifilla Small intestine Small intestine
None reported USA None reported USA None reported England (captive)
Ornithocapillaria quiscali Small intestine Pterothominx exilis Intestine Pterothominx exilis Intestine
None reported USA, Canada
None reported Asia, Europe
None reported Australia
None reported USA
Small intestine
Thomix tridens (reported as Capillaria pirangae) Thomix tridens
None reported Canada
Small intestine
Thomix tridens
(continues)
Baru˘s and Sergejeva (1990a) Baru˘s and Sergejeva (1990d)
Hodasi (1963), Cooper and Crites (1974b), and Badley and Dronen (1979) Read (1949) Cooper and Crites (1974b) Baru˘s and Sergejeva (1990d)
Baru˘s and Daniel (1976) Yamaguti (1961) Baru˘s and Sergejeva (1990d) and Frantova (2002) Baru˘s and Sergejeva (1990d)
Baru˘s and Daniel (1976)
Johnston and Mawson (1945)
Baru˘s and Sergejeva (1990a)
Durbin (1952)
Hodasi (1963)
September 29, 2008
Dunnock (Prunella modularis)
Eastern Towhee (Pipilo erythrophthalmus) Scarlet Tanager (Piranga olivacea) Summer Tanager (Piranga rubra) White-browed Babbler (Pomatostomus superciliosus) Alpine Accentor (Prunella collaris)
BLBS014-Atkinson 16:25
Capillarid species
484
American Robin (Turdus migratorius)
Spotless Starling (Sturnus unicolor) Redwing (Turdus iliacus) Eurasian Blackbird (Turdus merula) Intestine
Ornithocapillaria ovopunctata Pterothominx exilis Ornithocapillaria ovopunctata
Intestine Intestine
Pterothominx exilis Pterothominx exilis
Intestine
Intestine
Esophagus Small intestine
Eucoleus dispar Thomix tridens
None reported
None reported
None reported
None reported None reported
None reported None reported
Unknown
Unknown
None reported
Intestine
None reported
None reported
Small intestine
Intestine
None reported
Disease signs/lesions
Small intestine
Location in host
USA
Asia, Europe, and North Africa Canada
Asia, Europe Europe
United Kingdom, Russia Europe Tunisia
Worldwide
United Kingdom, Poland, Europe
Tadzhikistan
Japan
Location
Baru˘s and Sergejeva (1990d) Yamaguti (1961), Baru˘s and Sergejeva (1990d), and Frantova (2002) Yamaguti (1961) and Frantova (2002) Rayner (1932) and Cooper and Crites (1974a) Cooper and Crites (1974a)
Baru˘s and Sergejeva (1989) Bernard (1989)
Yamaguti (1961), Hair and Forrester (1970), and Baru˘s and Sergejeva (1990d) Yamaguti (1961) and Baru˘s and Sergejeva (1990b) Hair and Forrester (1970)
Baru˘s and Sergejeva (1990b)
Uchida et al. (1991)
References
September 29, 2008
Ornithocapillaria ovopunctata Capillaria similis
Chestnut-cheeked Starling Baruscapillaria (Sturnia philippensis) emberizae Rosy Starling (Pastor Ornithocapillaria roseus) ovopunctata European Starling Pterothominx exilis (Sturnus vulgaris)
Avian host
Table 27.2. (Continued)
BLBS014-Atkinson 16:25
485
White-eyed Vireo (Vireo griseus) Hooded Warbler (Wilsonia citrina)
Europe
None reported Asia, Europe None reported Cuba
Intestine Small intestine Small intestine
Thomix tridens
None reported Cuba
None reported Europe
Intestine
None reported Poland, Czech Republic None reported Canada Unknown Europe None reported Asia, Europe
None reported
Baruscapillaria ovopunctata Pterothominx exilis Thomix tridens
Ornithocapillaria quiscali Intestine Capillaria similis Unknown Pterothominx exilis Intestine
Baruscapillaria Intestine ovopunctata (also reported as Baruscapillaria inflexa) Pterothominx exilis Intestine
Baru˘s and Sergejeva (1990a)
Baru˘s and Sergejeva (1990d) Baru˘s and Sergejeva (1990a)
Yamaguti (1961)
Frantova (2002) and Kopociska ´ et al. (2004) Ching (1993) Yamaguti (1961) Baru˘s and Sergejeva (1990d)
Yamaguti (1961) and Kopociska ´ et al. (2004)
September 29, 2008
Fieldfare (Turdus pilaris) Dark-throated Thrush (Turdus ruficollis) Mistle Thrush (Turdus viscivorus)
Song Thrush (Turdus philomelos)
BLBS014-Atkinson 16:25
BLBS014-Atkinson
486
September 29, 2008
16:25
Parasitic Diseases of Wild Birds
Figure 27.1. Eucoleus contortus stichosome (glandular esophagus). The middle of one of the three individual stichocytes depicted in the photomicrograph is marked with arrow. Scale bar = 100 μm. Eucoleus contortus in Wood Ducks (Aix sponsa) from Louisiana (Ogburn-Cahoon 1979). Details about the natural history of a limited number of the more thoroughly studied capillarids are provided in the following sections. These species primarily infect species of Anseriformes, Columbiformes, or Galliformes or are zoonotic and have been extensively studied because of their importance to avian or human health. Pterothominx caudinflata (=Capillaria caudinflata, =Aonchotheca caudinflata) This species is a cosmopolitan parasite of birds in the orders Anseriformes, Columbiformes, Galliformes, Gruiformes, Passeriformes, and Tinamiformes (Table 27.1). Eggs passed in the feces of hosts must develop in the environment for 11–12 days prior to ingestion and further development in earthworms. Required intermediate hosts include Allolobophora caliginosa, Allolobophora parva, Aporrectodea rosea, Dendrobaena octaedra, Dendrobaena adaiensis (synonym Dendrobaena smidti), Dendrodrilus rubidus, Eisenia foetida, Helodrilus caliginosus, and Lumbricus terrestris (Moravec et al. 1987; McDougald 2003).
Pterothominx bursata This species infects many species of Galliformes (Table 27.1) and adults are found in the mucosa of the small intestine. Development of larvae within the egg takes 8–15 days after eggs are passed in feces. Eggs hatch once they are ingested by a required earthworm intermediate host. Larvae penetrate into the earthworm body cavity to molt and develop further. Larvae are infective to the definitive host approximately 22–25 days after infecting the earthworms. Ingested larvae develop to adults in 20–26 days ( Moravec et al. 1987; McDougald 2003).
Eucoleus contortus (=Thominx spirale, =Eucoleus spiralis, =Capillaria venteli) This is a cosmopolitan species that infects the oral cavity, esophagus, and crop of at least eight orders of birds (Table 27.1). Adults are found in both the mucosa and submucosa (Anderson 2000). The life cycle is direct and eggs passed in feces develop for 24–40 days before they are infective. Under ideal conditions of suitable moisture and no sunlight, embryonated eggs can remain viable for up to 15 months. The prepatent period
BLBS014-Atkinson
September 29, 2008
16:25
Capillarid Nematodes
487
Figure 27.2. Eggs of Eucoleus contortus from Northern Bobwhite (Colinus virginianus). Scale bar = 20 μm. Note the distinctive bipolar plugs that characterize capillarid and trichurid eggs.
is 29–54 days. This species will occasionally cause disease in free-ranging birds including Northern Bobwhite (Colinus virginianus), raptors, and, rarely, other species (Cram 1930; Colglazier et al. 1967; Kellogg and Prestwood 1968b; Helmboldt et al. 1971; Clausen and Gudmundsson 1981; Bosch et al. 2000.
Baruscapillaria obsignata (=Capillaria columbae) This species is a cosmopolitan parasite of Galliformes and Columbiformes, but is also found in birds in four other orders (Table 27.1). Chickens, turkeys, partridges, pigeons, guinea fowl, and quail are the most frequent hosts. The life cycle is direct; eggs passed in the feces develop to the infective stage within 6–7 days (Moravec et al. 1987). Embryonated eggs exposed to temperatures below −3.5◦ C, temperatures higher than 50◦ C, or desiccation for 14 days are killed (Levine 1937; McDougald 2003). When protected from direct sunlight, embryonated eggs of B. obsignata survive exposure to natural environmental conditions for at least 384 days. When exposed to the sun, they can survive at least 129 days (Levine 1937). This species is typically only associated with disease in captive birds.
Eucoleus annulatus (=Capillaria annulata) This cosmopolitan species infects the esophagus and crop of Galliformes and, rarely, Anseriformes. It is considered to be a synonym of E. contortus by some researchers. Earthworms in the genera Eisenia, Allolobophora, Lubricus, and Dendrobaena are required intermediate hosts (Wehr 1936). Larvae develop to the infective stage 14–21 days after eggs are ingested by a suitable earthworm. Although common in domestic turkeys and capable of causing significant disease in captive Wild Turkeys, prevalence in free-ranging Wild Turkeys is <5% nad no disease has been documented (Maxfield et al. 1963; Hurst et al. 1979; Castle and Christensen 1984; Sasseville et al. 1988).
Eucoleus contortus, Eucoleus dispar, Baruscapillaria falconis, and Capillaria tenuissima These four species are common in raptors (Table 27.2). Although most reports of E. contortus and E. dispar (possible synonyms, see Table 27.1 note) are from clinically normal birds (Tables 27.1 and 27.2), capillarids present in oral lesions are rarely identified to species and could represent one or both of these species. Oral
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lesions can occur in stressed birds with extremely intense infections, although presumably healthy wild birds seem to harbor infections of high intensity with no signs of disease. Mortality associated with unidentified oral capillarids has been reported in captive and free-ranging Gyrfalcons (Falco rusticolus), Rednecked Falcon (Falco chicquera), and Peregrine Falcons (Falco peregrinus) (Br¨ull 1932; Trainer et al. 1968; Cooper 1969; Clausen and Gudmundsson 1981). Eucoleus dispar has an indirect life cycle with earthworms as intermediate hosts, but a direct life cycle is suspected (Barus and Sergejeva 1989). The life cycles of B. falconis and C. tenuissima are unknown. If earthworm intermediate hosts are required, rodents are believed to serve as paratenic hosts (Olsen 1974). Pterothominx philippinensis (=Calodium philippinensis, =Capillaria philippinensis) Numerous species of birds are competent definitive hosts for P. philippinensis including White-breasted Waterhens (Amaurornis phoenicurus), Chinese PondHerons (Ardeola bacchus), Black-crowned NightHeron (Nycticorax nycticorax), Cattle Egret (Bubulcus ibis), Yellow Bittern (Ixobrychus sinensis), Common Moorhen (Gallinula chloropus), Greater Paintedsnipe (Rostratula benghalensis), pigeons, ducks (Anas spp.), and chickens (Bhaibulaya and Indra-Ngarm 1979; Cross and Basaca-Sevilla 1983). Many different species of fish can serve as a required intermediate host for P. philippinensis (Bhaibulaya and IndraNgarm 1979; Cross 1992) and eggs of P. philippinensis are not directly infective to avian or mammalian hosts (Cross 1992). The life cycle of P. philippinensis is unusual in that two biologically different populations of adult worms are produced. When larvae are ingested by consumption of an infected fish, they develop into adults in the intestine of the host within 10–11 days. Larviparous females begin to release L1 larvae in 13–14 days (Cross 1992). These L1 reinfect (autoinfect) the same host and develop into adults in 22–24 days. Male and female worms in this second population of adults mate, and females produce typical capillarid eggs that are passed with the feces. Experimentally infected birds with intense infections became listless, anorexic, have mucoid diarrhea, and may die (Bhaibulaya and Indra-Ngarm 1979). Since free-ranging birds are unlikely to develop infections of high intensity because of lower rates of exposure to infected fish, clinical signs and mortality are not expected. Birds are currently considered to be the natural host for this zoonotic parasite on the basis of their susceptibility in experimental studies and a single report from a naturally infected bittern (Ixobrychus sp.) (Cross 1992).
CLINICAL SIGNS Clinical signs are dependent both on locations within a host where particular species of capillarids develop and on intensity of infection. Birds with low numbers of parasites may not exhibit any clinical signs (Pinto et al. 2004). While most free-ranging wild birds have low intensity infections, those that die from capillarid infections are usually found dead and no clinical signs are observed. Birds harboring large numbers of B. obsignata, E. perforans, E. annulatus, or P. caudinflata may have nonspecific signs, which include emaciation, diarrhea, ruffled feathers, anorexia, and reduced water intake. Birds may also exhibit ataxia and weakness and will frequently die if such symptoms are evident (Cram 1936; Reis and Nobrega 1938; Kellogg and Prestwood 1968a; Wehr 1971; Hurst et al. 1979; Mathey and Gutter 1979; De Rosa and Shivaprasad 1999; McDougald 2003; Pinto et al. 2004). Clinical signs may not always be a good indicator of disease. Ring-necked Pheasants (Phasianus colchicus) infected with E. perforans did not exhibit signs of infection; however, gross and histopathologic lesions were apparent (Pinto et al. 2004). Few studies have investigated the clinical pathology of capillarid infections and findings vary depending on intensity of infection, extent of tissue damage, and severity of clinical signs. There are reports of normal white blood cell counts and hematocrit values in chickens infected with B. obsignata (Wakelin 1965) while others describe heterophilia and eosinophilia (Olson and Levine 1939). Some infected chickens may have slightly elevated globulins and total protein (Berghen 1966). In contrast, pigeons with intense infections of B. obsignata have decreased total protein and albumen that is likely due to severe diarrhea (Chubb et al. 1964).
PATHOLOGY As with clinical signs, the severity of gross and microscopic lesions is dependent on intensity of infection, host species, and parasite species. Low numbers of worms in a normal host rarely cause any appreciable lesions, remain in the superficial mucosa, and are incidental findings (Figure 27.3). Significant lesions and secondary bacterial infections can develop in aberrant hosts however when parasites migrate in the submucosa. Capillarid species that infect the upper gastrointestinal tract (oral cavity, esophagus, crop) can cause inflammation (Figure 27.4), dilatation of the crop or esophagus, thickening of mucosa, ulceration, bacterial colonization (Figure 27.4), exudation, and fibrinonecrotic plaques (Figure 27.4) and are generally
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Figure 27.3. Esophagus, Purple Martin (Progne subis). Cross sections of adult capillarids and eggs in the mucosa. Little inflammation or disruption may be caused by low-intensity infections in a normal avian host. Hematoxylin and eosin stain. Scale bar = 100 μm. Courtesy of R. W. Gerhold, University of Georgia.
considered to be more pathogenic than intestinal species (Helmboldt et al. 1971; Hurst et al. 1979). A Blue Jay (Cyanocitta cristata) infected with E. contortus was emaciated and dehydrated and had a diphtheritic membrane extending from the oral cavity to the proventriculus (Helmboldt et al. 1971). Gyrfalcons heavily infected with E. contortus or E. dispar had multifocal aggregates of thick, granular, yellow exudate in the oral cavity, pharynx, and esophagus (Clausen and Gudmundsson 1981). Epithelial necrosis and sloughing with occasional ulceration are evident in birds with lesions. Large numbers of adult parasites and eggs can be present in the lamina propria and are frequently surrounded by prominent mononuclear infiltrates. Similar infiltrates are also present in the epithelium and around mucous glands in the esophagus (Cram 1936; Clausen and Gudmundsson 1981). Marked necrosis and sloughing of the epithelium of the esophagus and crop also occurs in Northern Bobwhite infected with E. contortus (Cram 1936). Pheasants heavily infested with E. perforans have a chronic esophagitis characterized by mononuclear and granulocytic infiltrates in the lamina
propria (Pinto et al. 2004). Although the vast majority of lesions are confined to the mucosa, in extreme cases lesions may extend into the muscularis (Hung 1926). Additionally, hyperplasia or squamous metaplasia of the esophageal mucous glands, hyperkeratosis, severe inflammation, bacterial overgrowth, and intercellular edema may be observed. Intestinal and cecal dwelling capillarid species can cause thickening of the mucosa, erosion or ulceration, intraluminal fluid accumulation, petechiae, exudates, and diarrhea (with or without blood). Microscopic lesions may include epithelial necrosis with erosion or ulceration. Although gross lesions were absent in pheasants infected with Capillaria phasianiana, a chronic typhlitis with mononuclear cell infiltrate was noted by microscopy (Pinto et al. 2004). DIAGNOSIS Diagnosis of capillarid infection can be made by detection of eggs in fecal samples or mucosal scrapings or by detection of eggs or adult worms in histologic sections of tissues (Figures 27.3 and 27.4). Eggs of capillar-
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Figure 27.4. Crop, Red-tailed Hawk (Buteo jamaicensis). Numerous cross sections of adult capillarids and eggs are present in the mucosa. Note heterophilic inflammation in the submucosa (arrowheads) and bacterial colonies (arrows). Hematoxylin and eosin stain. Scale bar = 100 μm. Courtesy of A. E. Ellis, University of Georgia.
ids are easily recognized by the characteristic bipolar plugs (Figure 27.2); however, egg morphology alone cannot be used to reliably identify the parasite to genus or species. Eggs can be demonstrated by direct examination of fecal material or concentration and flotation of feces with either saturated sugar solution (specific gravity = 1.275), zinc sulfate solution (specific gravity = 1.18), or sodium nitrate (specific gravity = 1.2). Because worms are extremely thin, coiled through the mucosa, and easily broken, acquiring intact worms from tissue can be challenging. Gentle teasing apart of fresh tissue is the easiest way to acquire intact worms. Identification of capillarids to species depends on knowledge of the site of infection, examination of the vulvar region of the female worms, and detailed study of the posterior and anterior ends of adult worms of both sexes. However, small thin worms embedded in the mucosa that have a stichosome esophagus can be reliably identified as a capillarid. Infection with capillarids is common in many host species; therefore, diagnosis of disease requires demonstration of appropriate clinical signs and/or lesions (gross or microscopic) and a lack of other pathogens.
Differential diagnoses in sick birds include other agents that can cause weakness, emaciation, diarrhea, ruffled feathers, anorexia, and reduced water intake. Gross lesions associated with oral capillariasis in raptors can mimic lesions caused by trichom*oniasis or “frounce” (Trainer et al. 1968; Clausen and Gudmundsson 1981), yeast infections, or any agent causing caustic damage to the epithelium. IMMUNITY Little is known about the development of immunity to capillarids. Experimental studies with P. philippinensis and Baruscapillaria resecta suggest that some immunity develops following infection. Attempts to reinfect birds that had cleared an infection with P. philippinensis were not successful, suggesting development of immunity (Bhaibulaya and Indra-Ngarm 1979; Cross and Basaca-Sevilla 1983). Antibodies to antigens of B. resecta were detected in sera from infected Eurasian Jackdaw (Corvus monedula) (Fronska-Popiel et al. 2001); however, cross-reactive antigens appear to be present within the superfamily Trichinelloidea
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Capillarid Nematodes because sera samples from humans infected with P. philippinensis reacted with antigens from B. obsignata, Trichinella sprialis, and Trichuris vulpis (Banzon et al. 1975). The exact mechanism of immunity, if any develops, is currently unknown.
PUBLIC HEALTH CONCERNS The only known avian capillarid that poses a zoonotic threat is P. philippinensis. This species is a zoonosis in parts of Southeast Asia, Egypt, India, and Iran (Cross 1992). This parasite can cause severe enteritis in humans and is acquired by ingestion of raw or undercooked freshwater and brackish-water fish (Cross 1998). Unlike all other capillarids, eggs can embryonate within the human host, hatch, and autoinfect (Cross 1998).
DOMESTIC ANIMAL HEALTH CONCERNS Capillariasis is an important disease of many species of domesticated and captive wild birds and is associated with how birds are managed in captivity. Several species of capillarids (Table 27.1) infect both wild and domestic birds; however, wild birds do not pose any unique threat to domestic birds because these parasites are easily maintained in captive flocks without exposure to wild birds.
WILDLIFE POPULATION IMPACTS Capillarid infections rarely cause clinical disease in wild free-ranging birds in spite of being common and large outbreaks have not been reported. Wild birds that are kept in zoological parks or other captive facilities are more likely to develop capillariasis than their free-ranging counterparts. Captive birds are often exposed to large numbers of eggs or infected earthworms in facilities where birds are crowded (Maxfield et al. 1963; Hurst et al. 1979; Castle and Christensen 1984; Sasseville et al. 1988). This problem can be compounded in facilities where large numbers of closely related host species are housed together because several capillarid species have low host-specificity.
TREATMENT, MANAGEMENT IMPLICATIONS, AND CONTROL Management of capillarid infection in wild birds is not feasible and generally not warranted due to the limited pathogenicity associated with most infections. Captive wild birds should be tested and treated as appropriate to prevent the development of clinical disease associated with stress or acquisition of intense infections while in
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captivity. Sanitation of living quarters and raising birds on wire bottom cages will greatly decrease intensity of infection and risk of disease in captive birds because transmission of capillarids is dependent on fecal contamination and exposure to either eggs or infected earthworms. Mixing of wild birds of different species is discouraged and wild birds should be excluded from enclosures. Fenbendazole, febantel, and levamisole are highly efficacious for treatment of capillariasis in numerous avian species including chickens, turkeys, pheasants, partridges. They have been used successfully to treat pigeons infected with B. obsignata and raptors infected with an oral capillarid (Lawrence 1983; Kirsch 1984; Santiagno et al. 1985; Norton et al. 1991; Baert et al. 1993; Taylor et al. 1993; El-Kholy and Kemppainen 2005). Subcutaneous injection of ivermectin is effective against infections with P. caudinflata in captive guinea fowl (Okaeme and Agbontale 1989). Interestingly, albendazole improved body condition of birds coinfected with P. caudinflata and Heterakis gallinarum but, because worm burdens were unaffected, it was not considered an effective treatment (Villan´ua et al. 2007). Piperazine was not effective in treating P. caudinflata in guinea fowl (Ayeni et al. 1983). LITERATURE CITED Ahern, W. B., and G. D. Schmidt. 1976. Parasitic helminths of the American avocet Recurvirostra americana; four new species of the families Hymenolepididae and Acoleidae (Cestoda: Cyclophyllidae). Parasitology 73:381–398. Anderson, R. C. 2000. Nematode Parasites of Vertebrates. Their Development and Transmission. CABI Publishing, Wallingford, Oxon, UK, 650. Anderson, R. C., and O. Bain. 1982. No. 9: Keys to genera of the Superfamilies Rhabditoidea, Dioctophymatoridea, Trichinelloidea and Muspiceoidea. In CIH Keys to the Nematode Parasites of Vertebrates, R. C. Anderson, A. G. Chabaud, and S. Willmott (eds). Commonwealth Agricultural Bureaux, Farnham Royal, UK, pp. 1–26. Andrews, S. E., and W. Threlfall. 1975. Parasites of the common crow (Corvus brachyrhynchos Brehm, 1822) in insular Newfoundland. Proceedings of the Helminthological Society of Washington 42:24–27. Ayeni, J. S., O. O. Dipeolu, and A. N. Okaeme. 1983. Parasitic infections of the grey-breasted helmet guinea-fowl (Numida meleagris galeata) in Nigeria. Veterinary Parasitology 12:59–63. Badley, J. E., and N. O. Dronen. 1979. Some helminth parasites of the common grackle of southern Texas. Proceedings of the Helminthological Society of Washington 46:149–150.
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Baert, L., S. Van Poucke, H. Vermeersch, J. P. Remon, J. Vercruysse, P. Bastiaensen, and P. De Backer. 1993. Pharmaco*kinetics and anthelmintic efficacy of febantel in the racing pigeon (Columba livia). Journal of Veterinary Pharmacology and Therapy 16:223–231. Banzon, T. C., R. M. Lewert, and M. G. Yogore. 1975. Serology of Capillaria philippinensis infection: Reactivity of human sera to antigens prepared from Capillaria obsignata and other helminths. American Journal of Tropical Medicine and Hygiene 24:256–263. Baru˘s, V. 1971. A survey of parasitic nematodes of piciform birds in Cuba. Folia Parasitologica 18:315–321. Baru˘s, V., and M. Daniel. 1976. Capillarids (Nematoda:Capillariidae) from passeriform birds of Nepal. Folia Parasitologica 23:105–110. Baru˘s, V., and T. P. Sergejeva. 1989. Capillarids parasitic in birds in the Palaearctic region (2). Genera Eucoleus and Echinocoleus. Acta Scientiarum Naturalium Academiae Scientiarum Bohemoslovacae 23:1–47. Baru˘s, V., and T. P. Sergejeva. 1990a. A new genus of capillarids from birds, Tridentocapillaria gen. n. (Nematoda: Capillariidae). Folia Parasitologica 37:67–75. Baru˘s, V., and T. P. Sergejeva. 1990b. A new genus of capillarids from birds, Ornithocapillaria gen. n. (Nematoda: Capillariidae). Folia Parasitologica 37:237–248. Baru˘s, V., and T. P. Sergejeva. 1990c. Capillarids parasitic in birds in the Palaearctic Region (3). Genus Baru˘scapillaria. Acta Scientiarum Naturalium Academiae Scientiarum Bohemoslovacae 24:1–53. Baru˘s, V., and T. P. Sergejeva. 1990d. Capillarids parasitic in birds in the Palaearctic Region (4). Genera Pterothominx and Aonchotheca. Acta Scientiarum Naturalium Academiae Scientiarum Bohemoslovacae 24:1–48. Baru˘s, V., V. Kajerova, and B. Koubkova. 2005. A new species of Pterothominx Freitas, 1959 (Nematoda: Capillariidae) parasitising psittacine birds (Psittaciformes). Systematic Parasitology 62:59– 64. Berghen, P. 1966. Serum protein changes in Capillaria obsignata infection. Experimental Parasitology 19:34–41. Bernard, J. 1989. Nematode parasites of birds of the fauna in Tunisia. Archives de I’Institut Pasteur de Tunis 66:22–51. Bhaibulaya, M., and S. Indra-Ngarm. 1979. Amaurornis phoenicurus and Ardeola bacchus as experimental definitive hosts for Capillaria philippinensis in Thailand. International Journal for Parasitology 9:321–322.
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Capillarid Nematodes Clausen, B., and F. Gudmundsson. 1981. Causes of mortality among free-ranging Gyrfalcons in Iceland. Journal of Wildlife Diseases 17:105–109. Colglazier, M. L., E. E. Wehr, R. H. Burtner, and L. M. Wiest. 1967. Haloxon as an anthelmintic against the cropworm Capillaria contorta in quail. Avian Diseases 11:257–260. Cooper, C. L., and J. L. Crites. 1974a. Helminth parasites of the robin from South Bass Island, Ohio. Journal of Wildlife Diseases 10:397–398. Cooper, C. L., and J. L. Crites. 1974b. The helminth parasites of the red-winged blackbirds from South Bass Island, Ohio, including a check list of the helminthes reported from this host. Journal of Wildlife Diseases 10:399–403. Cooper, C. L., E. L. Troutman, and J. L. Crites. 1973. Helminth parasites of the brown-headed cowbird, Molothrus ater ater, from Ohio. Ohio Journal of Science 73:376–380. Cooper, J. E. 1969. Oesophageal capillariasis in captive falcons. Veterinary Record 84:634–636. Courtney, C. H., and D. J. Forrester. 1974. Helminth parasites of the brown pelican in Florida and Louisiana. Proceedings of the Helminthological Society of Washington 41:89–93. Cram, E. B. 1930. Parasitism in game birds. Transactions of the American Game Bird Conference 17:203–206. Cram, E. B. 1936. Species of Capillaria parasitic in the upper digestive tract of birds. U.S. Department of Agriculture Technical Bulletin 516:1–27. Cross, J. H. 1992. Intestinal capillariasis. Clinical Microbiology Reviews 5:120–129. Cross, J. H. 1998. Capillariasis. In Zoonoses: Biology, Clinical Practice, and Public Health Control, S. R. Palmer, L. Soulsby, and D. I. H. Simpson (eds). Oxford University Press, New York, pp. 759–772. Cross, J. H., and V. Basaca-Sevilla. 1983. Experimental transmission of Capillaria philippinensis to birds. Transactions of the Royal Society of Tropical Medicine and Hygiene 77:511–514. Davidson, W. R., F. E. Kellogg, and G. L. Doster. 1975. Capillaria tridens (Dujardin 1845) Travassos 1915, from wild turkeys (Meleagris gallopavo) in the southeastern United States. Journal of Parasitology 61:1115. De, N. C., and R. N. Maity. 1995. Indocapillaria gibsoni n. g., n. sp. (Nematoda: Trichuroidea) from the estuarine fish Sillaginopsis panijus (Hamilton) of West Bengal, India. Systematic Parasitology 30:153–157. De Rosa, M., and H. L. Shivaprasad. 1999. Capillariasis in a vulture guinea fowl. Avian Diseases 43:131–135. Dubinin, V. B., and M. N. Dubinina. 1940. Parasite fauna of bird colonies of the Astrakhan Reserve.
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Trudy Astrakhanskogo Gosudarstvennogo Zapovednika 2:190–298. Durbin, C. G. 1952. A new roundworm, Capillaria pirangae (Nematoda: Trichinellidae) from the scarlet tanager, Piranga erythromelas. Journal of the Washington Academy of Sciences 42:238–239. El-Kholy, H., and B. W. Kemppainen. 2005. Levamisole residues in chicken tissues and eggs. Poultry Science 84:9–13. Fedynich, A. M., D. B. Pence, and J. F. Bergan. 1997. Helminth community structure and pattern in sympatric populations of double-crested and neotropic cormorants. Journal of the Helminthological Society of Washington 64:176–182. Ferrer, D., R. Molina, C. Adelantado, and J. M. Kinsella. 2004. Helminths isolated from the digestive tract of diurnal raptors in Catalonia, Spain. The Veterinary Record 154:17–20. Fischbacher, W. K. 2007. Untersuchungen zum Gesundheitszustand von freilebendem Auer- und ¨ Birkwild in Osterreich. Ph.D. Dissertation, Veterin¨armedizinische Universit¨at Wien, 77. Forrester, D. J., and M. G. Spalding. 2003. Parasites and Diseases of Wild Birds in Florida. University Press of Florida, Gainesville, FL, 1152. Foster, G. W., J. M. Kinsella, E. L. Walters, M. S. Schrader, and D. J. Forrester. 2002. Parasitic helminths of red-bellied woodpeckers (Melanerpes carolinus) from the Apalachicola National Forest in Florida. Journal of Parasitology 88:1140–1142. Frantova, D. 2001. Capillarid nematodes (Nematoda: Capillariidae) parasitic in the common cormorant (Phalacrocorax carbo), with description of Baruscapillaria carbonis (Dubinin et Dubinina, 1940). Folia Parasitologica 48:225–230. Frantova, D. 2002. Some parasitic nematodes (Nematoda) of birds (Aves) in the Czech Republic. Acta Societatis Zoologicae Bohemicae 66:13–28. Freitas, J. F. T. 1933a. Sur deux nouvelles esp`eces du genre Capillaria Zeder, 1800. Comptes Rendus de la Soci´et´e de Biologie, Paris. 114:962–964. Freitas, J. F. T. 1933b. Nouvelles especes du genre Capillaria Zeder, 1800. CRSB Rio de Janeiro 114:1195–1196. Freitas, J. F. T., and J. L. Almeida. 1935. Sobre os nematoda Capillariinae parasitas de esophago e papo de aves. Mem´orias do Instituto Oswaldo Cruz 30:123–56. Freitas, J. F. T., J. M. Mendon¸ca, and J. P. Guimares. 1959. Sobre algumas especies do genero Capillaria Zeder, 1800 parasitas de aves (Nematoda: Trichuroidea). Mem´orias do Instituto Oswaldo Cruz 57:17–32. Fronska-Popiel, A., A. Okulewicz, and A. Perec. 2001. Immune response of jackdaw (Corvus monedula) to
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antigens of Capillaria resecta (Nematoda)—Western blotting analysis. Wiadomoúci parazytologiczne 47:693–698. Garcia, C. A., and A. G. Canaris. 1987. Metazoan parasites of Recurvirostra americana Gmelin (Aves), from Southwestern Texas and Monte Vista National Wildlife Refuge, Colorado, with a checklist of helminth parasites hosted by this species in North America. The Southwestern Naturalist 32:85-91. Gassal, S., and R. Schm¨aschke. 2006. The helminth and coccidial fauna of pheasants (Phasianus colchicus) in view of the specific environmental conditions in pheasantries and in the wild. Berliner und Munchener Tierarztliche Wochenschrift 119:295–302. Gibson, G. G. 1972. Sciadiocara denticulata n. sp. (Acuariidae) from Actitis macularia (L.) and other nematodes from spotted sandpiper and black-bellied plover. Canadian Journal of Zoology. 50:131–136. Gower, W. C. 1939. Host–parasite catalogue of the helminths of ducks. American Midland Naturalist 22:580–628. Hair, J. D., and D. J. Forrester. 1970. The helminth parasites of the Starling (Sturnus vulgaris L.): A checklist and analysis. American Midland Naturalist 83:555–564. Heard, R. W. 1967. Some Helminth Parasites of the Clapper Rail, Rallus longirostris Boddaert, from the Atlantic and Gulf Coasts of the United States. M.S. Thesis, University of Georgia, 32 pp. Helmboldt, C. F., R. P. Eckerlin, L. R. Penner, and D. S. Wyand. 1971. The pathology of capillariasis in the blue jay. Journal of Wildlife Diseases 7:157–161. Hoberg, E. P., G. S. Miller, E. Wallner-Pendleton, and O. R. Hedstrom. 1989. Helminth parasites of northern spotted owls (Strix occidentalis caurina) from Oregon. Journal of Wildlife Diseases 25:246–251. Hodasi, J. K. M. 1963. Helminths from Manitoba birds. Canadian Journal of Zoology 41:1227–1231. Hromada, M., V. Dudi`oa´ k, and R. Yosef. 2000. An inside-out perspective of the true shrikes—A review of the helminthofauna. Ring 22:185–204. Hung, S. L. 1926. Pathological lesions caused by Capillaria annulata. North American Veterinarian 7:49–50. Hurst, G. A., L. W. Turner, and F. S. Tucker. 1979. Capillariasis in penned wild turkeys. Journal of Wildlife Diseases 15:395–397. Illescas, M. P., M. Rodriguez, and F. Aranda. 1993. Parasitation of falconiform, strigiform and passeriform (Corvidae) birds by helminths in Spain. Research and Reviews in Parasitology 53:129–135. Iulia, F., and I. Pavlovic. 2003. The first occurrence of Thominx cyanopicae (Lopez-Neyra, 1947) in pheasants. Acta Veterinaria 53:183–190.
Johnston, T. H., and P. M. Mawson. 1941. Some nematode parasites of Australian birds. Proceedings of the Royal Society of New South Wales 66:250–256. Johnston, T. H., and P. M. Mawson. 1944. Remarks on some parasitic nematodes from Australia and New Zealand. Transactions of the Royal Society of South Australia 68:60–66. Johnston, T. H., and P. M. Mawson. 1945. Capillarid nematodes from South Australian fish and birds. Transactions of the Royal Society of South Australia 69:243–248. Johnston, T. H., and P. M. Mawson. 1947. Some avian and fish nematodes, chiefly from Tailem Bend, South Australia. Records of the South Australian Museum 8:547–553. Joy, J. E., and J. B. Scott. 1997. Amphibiocapillaria tritonispunctati (Nematoda: Trichuridae) infections in the red-spotted newt Notophthalmus v. viridescens from western West Virginia. American Midland Naturalist 138:408–411. Kellogg, F. E., and A. K. Prestwood. 1968a. Case report and differentiating characteristics of Capillaria phasianina from pen-raised pheasants of Maryland. Avian Diseases 12:518–522. Kellogg, F. E., and A. K. Prestwood. 1968b. Gastrointestinal helminthes from wild and pen-raised bobwhites. Journal of Wildlife Management 32:468–475. Kinsella, J. M. 1973. Helminth parasites of the American Coot, Fulica americana americana, on its winter range in Florida. Proceedings of the Helminthological Society of Washington 40:240–242. Kinsella, J. M., and D. J. Forrester. 1999. Parasitic helminths of the common loon, Gavia immer, on its wintering grounds in Florida. Journal of the Helminthological Society of Washington 66:1–6. Kinsella, J. M., R. A. Cole, D. J. Forrester, and C. L. Roderick. 1996. Helminth parasites of the Osprey, Pandion haliaetus, in North America. Journal of the Helminthological Society of Washington 63:262–265. Kinsella, J. M., G. W. Foster, R. A. Cole, and D. J. Forrester. 1998. Helminth parasites of the bald eagle, Haliaeetus leucocephalus, in Florida. Journal of the Helminthological Society of Washington 65:65–68. Kinsella, J. M., G. W. Foster, and D. J. Forrester. 2001. Parasitic helminthes of five species of owls from Florida, USA. Comparative Parasitology 68:130–134. Kinsella, J. M., M. G. Spalding, and D. J. Forrester. 2004. Parasitic helminths of the American white pelican, Pelecanus erythrorhynchos, from Florida, USA. Comparative Parasitology 71:29–36. Kirsch, R. 1984. Treatment of nematodiasis in poultry and game birds with fenbendazole. Avian Diseases 28:311–318.
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Pursglove, S. R., Jr., 1969. A Survey of the Internal Parasite Fauna of American Woodco*ck in the Canaan Valley of West Virginia. M.S. Thesis, University of Georgia, 86 pp. Ramalingam, S., and W. M. Samuel. 1978. Helminths in the great horned owl, Bubo virginianus, and snowy owl, Nyctea scandiaca, of Alberta. Canadian Journal of Zoology 56:2454–2456. Rayner, J. A. 1932. Parasites of wild birds in Quebec, Canada. Journal of Agricultural Science 12:307–309. Read, C. P. 1949. Studies on North American Helminths of the Genus Capillaria Zeder, 1800 (Nematoda): III. Capillarids from the lower digestive tract of North American Birds. Journal of Parasitology 35:240–249. Reis, J., and P. Nobrega. 1938. Sobre as les˜oes produzidas pela Capillaria perforans Kotl´an e Orosz nas aves dom´esticas. Arquivos do Instituo Biologico 9:21–23. Sanmartin, M. L., F. Alvarez, G. Barreiro, and J. Leiro. 2004. Helminth fauna of Falconiform and Strigiform birds of prey in Galicia, Northwest Spain. Parasitology Research 92:255–263. Santiagno C., P. A. Mills, and C. E. Kirkpatrick. 1985. Oral capillariasis in a red-tailed hawk: Treatment with fenbendazole. Journal of the American Veterinary Medical Association 187:1205–1206. Sasseville, V. G., B. Miller, and S. W. Nielsen. 1988. A pathologic study of wild turkeys in Connecticut. Cornell Veterinarian 78:353–364. Schei, E., P. R. Holmstad, and A. Skorping. 2005. Seasonal infection patterns in Willow Grouse (Lagopus lagopus L.) do not support the presence of parasite-induced winter losses. Ornis Fennica 82:137–146. Schorr, L. F. 1988. A Survey of Parasites and Diseases of Pen-Raised Wild Turkeys. M.S. Thesis, The University of Georgia, p. 132. Senlik, B., E. Gulegen, and V. Akyol. 2005. Effect of age, sex and season on the prevalence and intensity of helminth infections in domestic pigeons (Columba livia) from Bursa Province, Turkey. Acta Veterinaria Hungarica 53:449–456. Sep´ulveda M. S., M. G. Spalding, J. M. Kinsella, R. D. Bjork, and G. S. McLaughlin. 1994. Helminths of the Roseate Spoonbill, Ajaia ajaja, in Southern Florida. Journal of the Helminthological Society of Washington 61:179–189. Sep´ulveda, M. S., M. G. Spalding, J. M. Kinsella, and D. J. Forrester. 1996. Parasitic helminths of the little blue heron, Egretta caerulea, in southern Florida. Journal of the Helminthological Society of Washington 63:136–140. Sep´ulveda, M. S., M. G. Spalding, J. M. Kinsella, and D. J. Forrester. 1999. Parasites of the great egret (Ardea albus) in Florida and a review of the helminths
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Section IV: Leeches
Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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28 Leech Parasites of Birds Ronald W. Davies, Fredric R. Govedich, and William E. Moser INTRODUCTION Leeches (Euhirudinea) occur worldwide and form an important component of the benthic invertebrate fauna of most lakes and ponds, with a few species occurring in the quieter flowing sections of streams and rivers. Marine leeches typically feed on marine vertebrates including fish and turtles. Land (terrestrial) leeches can be found in moist, humid tropical and subtropical regions where they inhabit wet rainforests and feed on the blood of vertebrates (Sawyer 1986; Davies and Govedich 2001; Govedich 2001). Most freshwater leeches are predators and feed on chironomids, oligochaetes, amphipods, and mollusks. Others are temporary, sanguivorous (i.e., bloodsucking) ectoparasites of fish, turtles, amphibians, mammals, or birds. One exception is Placobdelloides jaegerskioeldi, which is host-specific for hippopotamus and is an endoparasite found only in the rectum of this aquatic mammal (Oosthuizen and Davies 1994). The majority of terrestrial leeches are sanguivorous, feeding on reptiles, mammals, and birds (Sawyer 1986; Davies and Govedich 2001; Govedich 2001). There are currently 12 families of leeches, 4 of which contain species that parasitize birds: the Glossiphoniidae, Hirudinidae, Ornithobdellidae, and Haemadipsidae. Glossiphoniids feed using a proboscis (Figure 28.1) that protrudes through a mouth located in the anterior sucker. Hirudinids, ornithobdellids, and haemadipsids all feed by using cutaneous jaws with rows of teeth to cut the skin of the host (Sawyer 1986; Davies and Govedich 2001; Govedich 2001). Leeches in the genus Theromyzon are of particular interest because they tend to specialize on waterbirds. They prefer to feed in the nasal cavities and thus have the potential to cause stress. Intense leech infestations have been correlated with increased water bird mortality (Sawyer 1986; Davies and Govedich 2001).
HISTORY Leeches are important bloodsucking parasites and predators in terrestrial, marine, and freshwater ecosystems and have been used in medicine for centuries. Many species are still used for medicinal purposes and have been transported throughout the world, particularly Hirudo medicinalis (Sawyer 1986; Govedich 2004). The history of the discovery of avian leeches was summarized by Bartonek and Trauger (1975) and Trauger and Bartonek (1977). Most research on leech parasitism of birds has been done in North America on a limited number of species in the glossiphoniid genus Theromyzon (Figure 28.2). These studies have led to substantial taxonomic revision of the North American species of the genus Theromyzon (Oosthuizen and Davies 1992, 1993; Davies and Oosthuizen 1993). DISTRIBUTION AND HOST RANGE Leeches have a variety of hosts (Table 28.1) and generally are not host-specific, tending to feed on any available hosts that live in the same habitat. Avian hosts tend to live in or near water and include waterbirds (ducks, coots, loons, herons, etc.), seabirds (shearwaters, penguins, etc.), and even large land birds such as the cassowary (Benham 1907; Mann 1961; Richardson 1969, 1981; Wilkialis 1970; Mason 1976; Sawyer 1986). Although species of Theromyzon are often referred to as duck leeches (Figure 28.3), they parasitize a wide variety of waterbirds in addition to ducks (Trauger and Bartonek 1977) (Table 28.1). They commonly attack grebes (Storer 2000), coots, loons, gulls, geese, swans, herons, and any other bird that comes within their reach. Theromyzon tessulatum has been found in eagles (Collin 1892) and also in the eye of a Hooded Crow (Corvus cornix) (Christiansen 1939). Theromyzon cooperi will feed in the eye socket of Rock Pigeons (Columba livia) in the laboratory (Oosthuizen and Fourie 1985).
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(a)
(b)
Figure 28.1. Anterior sucker of a leech that feeds with (a) a proboscis (Glossiphoniidae) and (b) a jawed leech (Hirudinidae, Ornithobdellidae, or Haemadipsidae).
Species of Placobdella, Placobdelloides, Haementeria, and other glossiphoniids besides Theromyzon have also been known to feed on birds (Table 28.1). For example, Placobdella ornata has been found feeding on North American birds (Moore 1964, 1966; Scudder and Mann 1968), while Placobdella papillifera feeds readily on fresh heparinized Mallard (Anas platyrhynchos domesticus) blood in the laboratory, indicating that this species might feed on waterfowl in the wild (Davies and Wilkialis 1982). Placobdelloides maorica, another glossiphoniid, feeds on Pacific Black Duck (Anas superciliosa) in New Zealand and it is likely that other members of this genus of leech also take blood meals from waterfowl as well as from reptiles and amphibians. The high frequency of leech attacks on waterfowl likely reflects the availability and abundance of birds rather than host specificity of the leeches. Among hirudinid leeches, H. medicinalis has been recorded from Europe and western Asia and was imported into North America in large numbers for medical uses. This species is still farmed and sold worldwide to medical institutions. Whether or not it has established itself in the wild in North America is still questionable, but Davies (1973) collected a specimen of this species from a field site in Alberta, Canada.
Theromyzon tessulatum and Theromyzon maculosum are considered to be European species, but their presence in North America has been well documented. Both species feed on a variety of waterfowl including ducks and geese. Another species, Theromyzon trizonare, has only been recorded from the migratory routes of ducks from the Rocky Mountain flyway in the Canadian provinces and states west of the Great Lakes (Elliott and Mann 1979; Davies and Wilkialis 1980). The African duck leech, T. cooperi, is the only representative of the genus reported from Africa (Oosthuizen 1993). Theromyzon garjaewi is recorded from Lake Baikal, Theromyzon propinquum from South America, and Theromyzon matthaii from India (Singhal and Davies 1986). ETIOLOGY Species of leeches that parasitize birds are found in four families: the Glossiphoniidae, Hirudinidae, Ornithobdellidae, and Haemadipsidae. Glossiphoniids have a “leaflike” body shape and have eyes located near the centerline of the head (Figures 28.4 and 28.5). They feed using a proboscis that is everted through the mouth and located in the anterior sucker (Figure 28.1). Jawed leeches (hirudinids, ornithobdellids, and haemadipsids) are long and slender in shape, typically have eyes located near the margin of the head, and cutaneous jaws to cut the skin of the host (Figures 28.1, 28.4, and 28.5). The body of leeches is composed of 32 postoral segments, an anterior sucker, and a ventrally directed posterior sucker that is generally wider than the body at the point of attachment (Figures 28.1 and 28.5). The mouth is found within the anterior sucker (Figure 28.1) and the anus opens on the dorsal surface just anterior to the posterior sucker. Detailed reviews of the taxonomy, anatomy, ecology, and physiology together with taxonomic keys of the freshwater leeches of North America can be found in Davies and Govedich (2001) and Klemm (1985, 1995). Members of the family Glossiphoniidae have dorsoventrally flattened triannulate bodies that are not differentiated into regions. The ventral anterior sucker is narrower than the body but fused to it. The mouth is a small pore on the ventral surface of the anterior sucker through which a muscular pharyngeal proboscis can be protruded (Figure 28.1). The genus Theromyzon has four pairs of eyes in comparison to Placobdella, Placobdelloides, and Haementaria, which have zero, one, or two pairs of eyes, respectively (Figure 28.4). The Hirudinidae, Ornithobdellidae, and Haemadipsidae have a large mouth occupying the entire cavity of the anterior sucker (Figure 28.1). Hirudinids have three jaws (Figure 28.6), which carry 35–100 acute sharp
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Figure 28.2. Distribution map of species of North American Theromyzon (Glossiphoniidae) (shaded area).
teeth in one (monostichodont) or two (distichodont) rows. The ornithobdellids also have three jaws, but the teeth are blunt, reduced, and vestigial. The haemadipsids, like the hirudinids, have sharp teeth in one row on two or three jaws. All three families have five pairs of eyes (Figure 28.4) arranged in an arch, and the body is elongated in freshwater species and only slightly flattened during swimming. EPIZOOTIOLOGY All leeches are hermaphrodites showing protandry or cosexuality (Davies and Singhal 1988), with reciprocal cross-fertilization being the norm. Fertilization is internal and the eggs are deposited into a cocoon secreted by the cl*tellum. The life cycle of all leeches consists of
egg, juvenile, and mature hermaphrodite adult. Most leeches attach the cocoon to a substrate and abandon the developing eggs and young (Figure 28.7). However, members of the family Glossiphoniidae brood their eggs and young (Kutschera and Wirtz 2001; Govedich 2004) (Figure 28.8). The majority of sanguivorous leeches are iteroparous (i.e., capable of breeding several times) and exhibit saltatory growth after reaching a mature size. Most families of leeches produce thick leathery cocoons that contain both eggs and nutrients. These are attached to a substrate and abandoned by the parent. However, the cocoons of Glossiphoniidae are very thin walled (sometimes called egg cases) and are either deposited on the substrate and immediately covered by the body of the parent or are
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Figure 28.3. Dissected nasal passages of a Mallard (Anas platyrhynchos) with an infestation by Theromyzon sp. Reproduced from Tuggle (1999), with permission of the author.
directly attached to the ventral surface of the parent. In both cases the hatchlings are attached to the ventral body wall and carried by the parent for up to 5 months in the case of T. tessulatum and for about 1 month by T. trizonare (Wilkialis and Davies 1980). Theromyzon trizonare differs behaviorally from all other species of Theromyzon in North America by attaching its cocoon to its ventral body wall so that if disturbed, the parent can move away, carrying the cocoon with it (Davies and Wilkialis 1980). Once on an avian host, leeches generally move to the head where most frequently they attach themselves inside the nasal passages (Figure 28.3) or, more rarely, around the eyes, particularly under the nictitating membrane and eyelids. Leeches in these positions are safe from removal by host scratching behavior. Leeches entering the nares often penetrate further into the trachea, lungs, and into the air spaces of the skull. Less common attachment sites include the mouth, cloaca, and the skin of the legs, feet, and breast. Leeches are also intermediate hosts for some digenetic trematodes that are commonly found in ducks and grebes (Sawyer 1986; Storer 2000). Leeches can also serve as intermediate hosts for cestodes of ducks (Sawyer 1986) and can potentially serve as intermediate or paratenic hosts of cestodes, acanthocephalans, and nematodes. Leeches are sometimes consumed as prey by other invertebrates, fish, snakes, and waterfowl. Leeches respond to the presence of a potential predator by attaching to a substrate and flattening their body, making it difficult for the predator to pull it off of the substrate. Other defensive responses include forming a ball to protect attached eggs and young, and burrowing into soft mud or leaf material. Sanguivorous leeches spend a relatively short time (1–8 h) taking a blood meal on the host (bird) and are
more frequently found free living in the benthos of the lake or river or under vegetation in moist terrestrial habitats. Theromyzon trizonare and T. tessulatum and presumably other Theromyzon species that feed on waterfowl require a minimum of three complete blood meals in order to reach reproductive maturity (Wilkialis and Davies 1980; Davies 1984). This is significantly fewer than H. medicinalis, which requires 10 or more meals to reach maturity (Putter 1907, 1908; Blair 1927). Over 80% of T. trizonare take 3 blood meals from birds in the first 6 months after hatching. The remainder of the population overwinters after 2 blood meals and takes the third meal in the spring, so that all of the population reproduces approximately 12 months after hatching. Individuals that overwinter after 3 meals may decline in weight before the spring and require a fourth meal before reproduction commences. Weight of T. trizonare increases proportionately with each successive blood meal, ranging from 7.3 g after the first meal to 20.4 g after the third (Davies 1984). Also shortly after taking a blood meal, Theromyzon shows a further increase in body weight, presumably from water uptake. Sanguivorous leeches in proximity to a potential host respond to changes in illumination (shadows), vibrations in the water, chemical stimuli, and possibly differences in temperature between the host and the surrounding water. For example, species of Theromyzon attach to and attack a host with a body temperature in the range of 37–40◦ C (Dickinson and Lent 1984). Disturbances in the water can be detected by the sensilla of a hungry leech. These are frequently the first indication of the presence of a potential host and a hungry leech will move in their direction. Although movements toward a potential host by sanguivorous leeches are not specific, the next step, determining whether or not the potential host is an appropriate source of blood, involves selectivity. When a shadow passes over a hungry sanguivorous leech, it reacts by making searching movements or even swimming toward the potential host. After attachment to the potential host with the posterior sucker, the leech makes a number of searching movements with the anterior portion of the body. If the host is not appropriate, the leech detaches. If T. trizonare, which primarily feeds in the nares, attaches to the feathers rather than the beak, it will move about on the bird for up to 30 minutes in search of a suitable location to feed. Suitable host recognition is presumably by chemoreception and/or mechanoreception. When birds are present and available, hungry Theromyzon are found primarily in shallow waters on the upper exposed surfaces of rocks, stones, and plants where the probability of meeting a host is high.
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Leech Parasites of Birds Table 28.1. Host and distribution records for families and species of leeches that parasitize birds (Sawyer 1986). Leech family
Leech species
Host families
Distribution
Glossiphoniidae
Haementeria depressa Oosthuizobdella garoui Placobdella costata Placobdella ornata Placobdella papillifera Placobdelloides maorica Theromyzon sp. Theromyzon affinis Theromyzon bifarium Theromyzon cooperi Theromyzon garjaewi Theromyzon maculosum Theromyzon matthaii Theromyzon mollissimum Theromyzon pallens Theromyzon propinquum Theromyzon rude Theromyzon sexoculatum Theromyzon tessellatoides Theromyzon tessulatum Theromyzon trizonare Hirudobdella antipodum Ornithobdella edentula Aetheobdella hirudoides Myxobdella annandalei Richardsonianus howensis Hirudo medicinalis Hirudo nipponia Dinobdella ferox Haemadipsa zeylanica Chtonobdella bilineata Chtonobdella limbata Placobdella novabritanniae
Anatidae Podicipedidae Rallidae
Cosmopolitan
Ornithobdellinae
Hirudinidae
Haemadipsidae
Anatidae Ardeidae Podicipedidae Gaviidae Rallidae Scolopacidae Recurvirostridae Charadriidae Laridae* Corvidae* Accipitridae*
Procellariidae Spheniscidae Bush birds Ardeidae Birds hypothesized Phasianidae Phasianidae Phasianidae Phasianidae Cracticidae Muscicapinae Casuariidae
New Zealand New Zealand, Snare’s Islands Southeastern Australia Indo-Pacific Australia, Lord Howe Island Europe, Asia Europe, Asia Indo-Pacific Indo-Pacific Indo-Pacific Indo-Pacific Indo-Pacific
*Atypical hosts. Although leeches in general are strongly photonegative, hungry sanguivorous leeches are frequently photopositive. After a meal or when carrying eggs or young, leeches remain in sheltered areas away from light and potential predators. In the winter, when feeding does not occur, leeches move into deeper water under ice cover. Behavior of terrestrial sanguivorous leeches is similar to freshwater species with individuals moving toward shadows and vibrations while searching for potential hosts. Unlike freshwater species, Amicobdella nigra and other terrestrial haemadipsid species react to the presence of carbon dioxide by increasing searching movements and moving toward higher carbon dioxide
concentrations. As a result, leeches move upward to the mouth, eyes, and nostrils where the majority of feeding occurs. Direct passive transfer of leeches can occur during contact between two ducks. Davies et al. (1982) examined passive transfer of two sanguivorous glossiphoniids, T. trizonare and P. papillifera, between ducks. Theromyzon adults were transferred from duck to duck both when the leeches were feeding in the nares of the ducks and while they were on the body under the feathers, where feeding was never observed. Placobdella papillifera were never found in the nares of the ducks, but were carried on the body under the feathers where a low proportion were observed to feed.
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(a)
(b)
(c)
(d)
Figure 28.4. Relative eye positions of the (a–c) Glossiphoniidae and (d) Hirudinidae, Ornithobdellidae, and Haemadipsidae.
Because of the diversity of physiochemical conditions encountered in different aquatic ecosystems and while taking a blood meal, sangivorous leeches have considerable physiological plasticity. They are capable of living in waters with very low salt concentrations as well as in waters exceeding the salinity of sea water. Similarly, some species are capable of surviving in the absence or near absence of oxygen, as might occur when taking a blood meal from the nares of a bird, or in supersaturated water, as might occur when a leech leaves its host and lands in the weedy shallow waters of a warm placid lake or pond. CLINICAL SIGNS Infected birds can be identified by the presence of leeches in or near the nares, around the eyes, or near the cloaca where there are many capillaries and a good blood supply. Leech bite wounds may also be found following feeding, after the leech has left the host. Infested birds exhibit various degrees of discomfort, including shaking their heads, scratching their
(a)
(b)
Figure 28.5. Body shape of (a) jawed leeches and (b) glossiphoniids.
bill with their feet, and/or forcing air through their nasal passages (sneezing), sometimes even when flying (Bartonek and Trauger 1975; Trauger and Bartonek 1977; Sawyer 1986). Engorged leeches protruding from the nares, eyes, or attached elsewhere on the host are readily visible, even from a distance using binoculars (Bartonek and Trauger 1975; Trauger and Bartonek 1977). Periorbital feathers soiled with ocular discharge, conspicuous eye irritations, eyelids matted together, and impairment of vision, including blindness, are also signs of leech infestation (Bartonek and Trauger 1975; Oosthuizen and Fourie 1985). Symptoms of heavy parasitism by Theromyzon spp. are similar to botulism (Oosthuizen and Fourie 1985; Fourie et al. 1986). Heavily infested birds are lethargic and sometimes exhibit various degrees of paralysis. In Africa, after heavy infestation by T. cooperi, some birds are unable to walk or fly because their leg and/or wing muscles have become paralyzed. Later, the neck muscles may also become paralyzed, leading to a drooping head (Fourie et al. 1986). Heavily infested birds often also exhibit short, labored breathing (Bartonek and Trauger 1975). PATHOGENESIS AND PATHOLOGY Parasitism by leeches can result in temporary blindness, clouding of the cornea, and in some cases, collapse of the globe of the eye (Bartonek and Trauger 1975; Tuggle 1999). Theromyzon trizonare has been implicated in waterfowl deaths due to either suffocation from numerous engorged leeches in the nasal passages or stress and general weakness (Davies and Wilkialis 1981). Ducklings that were exposed in the laboratory to numbers of T. trizonare comparable to those recorded from wild birds developed signs of stress and debilitation. These signs included significant increases in biomass of the adrenals, spleen, and thymus, a significant decrease in liver biomass, and a significant decrease in the keel/sternum ratio, a relative measurement of emaciation. Ulceration or inflammation of the nasal mucosa of the parasitized ducks was not observed by microscopy (Davies and Wilkialis 1981). DIAGNOSIS The identification of Theromyzon (family Glossiphoniidae) is based on the number of annuli separating the male and female gonopores. In North America, the taxonomy of the genus has been revised by Oosthuizen and Davies (1992, 1993) and Davies and Oosthuizen (1993), so care must be taken to bring earlier identifications into line with this work. Fourteen species of Theromyzon leeches have been described from ducks
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Leech Parasites of Birds
Jaws Teeth
(a)
(b)
(c)
(d)
Figure 28.6. Jaw placement in (a) hirudinids and ornithobdellinids and (b) haemadipsids. Number of teeth rows on jaws: one row or monostichodont (c) and two rows or distichodont (d).
in different parts of the world. These can be divided into four groups on the basis of the number of annuli (2, 3, 4, or 5) separating the male and female gonopores. In North America, T. trizonare (with three annuli), Theromyzon rude, Theromyzon bifarium, and
Cocoon eggs
+
T. maculosum (each with two) and T. tessulatum (with four) differ in the number of annuli between the gonopores and in several other reproductive characteristics (Oosthuizen and Davies 1993; Davies and Govedich 2001).
Cocoon hatchlings
+
Cocoon juveniles
+
Juveniles leaving cocoon Mature adult
Free-living juvenile
Figure 28.7. Typical life cycle of nonbrooding leeches (including hirudinids, ornithobdellinids, and haemadipsids). The cocoon is attached to a substrate and abandoned.
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Development from egg to juvenile
Adult with eggs
Juvenile
Adult with young
Figure 28.8. Life cycle of glossiphoniid leeches. Eggs (cocoons) may be attached to a substrate (nest) or to the parent (brooding, shown here). Young are brooded on the parent until they are mature.
The African duck leech, T. cooperi, is the only representative of the genus reported from Africa (Oosthuizen 1993) and, like T. garjaewi from Lake Baikal, has two annuli between its gonopores. Both T. propinquum from South America and T. matthaii from India (Singhal and Davies 1986) have three annuli between the gonopores. Freshwater and terrestrial leeches in the families Hirudinidae, Ornithobdellidae, and Haemadipsidae are identified based on external features such as the presence of auricles, ocular areoli (Haemadipsidae only), and the nature of the jaws and teeth (numerous sharp teeth for Hirudinidae; reduced, blunt vestigial teeth in the Ornithobdellidae). Species within these families are determined using combinations of characters such as the color pattern, the structure of the female and male reproductive systems, and the structure of the digestive system. Identification of leeches can be difficult, especially if they are not properly relaxed and preserved. Many
species of leeches require dissection or histological sectioning for species-level identification. Notes on dorsal and ventral coloration and pattern should be made prior to preservation as leech pigmentation typically fades during the preservation process. Leeches should be narcotized before fixation. Otherwise, they contract sharply and obscure structures used in identification. A recommended method is to add 70% ethanol dropwise into a vessel of water containing the leech until the leech no longer reacts to a blunt probe. Other methods of narcotizing leeches and specific preservation protocols are available (Klemm 1985, 1995; Davies and Govedich 2001). A voucher series of specimens narcotized and then fixed in 10% formalin followed by 95% ethanol is recommended. IMMUNITY There is no acquired immunity to leech infestation, but leeches may be mechanically removed or die when
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Leech Parasites of Birds feeding on a host. Mortality among leeches during feeding on ducks varies with each blood meal. Mortality is highest during the first meal and lowest during the third. Most leech mortality occurs when they are ingested by the host as they attempt to feed in the nares. Leeches ingested in this manner or that were fed directly to ducks in the laboratory were completely digested and never recovered from duck feces. Following successful feeding, Theromyzon leeches leave the host by one of two methods. Engorged leeches either drop out of the nares by themselves or are forcibly ejected by sneezing. The latter occurs more frequently during later meals. Species of Theromyzon are susceptible to desiccation and will die if the bill and nares dry out.
PUBLIC HEALTH AND DOMESTIC ANIMAL HEALTH CONCERNS Leeches are typically not host-specific and members of the Hirudinidae and Haemadipsidae have been known to feed on humans and other large mammals as well as birds. Leeches are primarily a nuisance and do not generally cause major problems for humans. However, leeches can cause problems for people allergic to their feeding secretions or who have a suppressed immune system. The site of incision during feeding can also become secondarily infected, and in a few rare cases leeches have caused problems in humans when they have entered the mouth, anus, or vagin*. Members of the genus Theromyzon (Glossiphoniidae) are host-specific to birds although there are two cases from Europe where T. tessulatum was reportedly attached to a human eye and also found inside a human larynx. Both cases required medical attention (AuwHaedrich et al. 1998; Kuehnemund and Bootz 2006). While Kuehnemund and Bootz (2006) described this leech as T. tessulatum, examination of the photographs in the paper suggest the leech was more likely a species of Hirudo. Leeches certainly have the potential to cause stress and mortality in domestic birds, especially cygnets and ducklings. Leeches can be removed by a trained veterinarian and birds usually recover within a few hours after leeches have been removed.
TREATMENT AND CONTROL Several chemotherapeutic remedies have been recommended for alleviation of leech infestation. Each regimen includes immersion or rinsing of the nasal chambers with an aqueous irritant for a few seconds until the leeches detach. Recommended solutions include lactic acid (4–10%), weak gastric juice, sodium chloride (10%), acetic acid, weak ammonia, and opthaine or
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sulfathiazole (Kuznetsova 1955; Ponomarenko 1960; Lang 1969). Removal of conspicuous leeches by hand or with forceps has also been suggested (Bartonek and Trauger 1975; Trauger and Bartonek 1977) but this is not recommended. Measures to reduce leech populations in the field are not presently feasible or advisable as no agent (chemical or physical) selectively kills leeches. Use of nonspecific agents may damage other components of the aquatic ecosystem and have more serious effects on wildfowl populations than the leeches themselves. Keeping birds away from habitats with leeches is theoretically possible but usually impractical.
LITERATURE CITED Auw-Haedrich, C., A. Keim, and M. Kist. 1998. Conjunctival infestation of a child with Theromyzon tessulatum. British Journal of Ophthalmology 82:1093–1094. Bartonek, J. C., and D. L. Trauger. 1975. Leech (Hirudinea) infestations among waterfowl near Yellowknife, Northwest Territories. Canadian Field Naturalist 89:234–243. Benham, W. B. 1907. Two new species of leech in New Zealand. Transactions New Zealand Institute 39:181–193. Blair, W. N. 1927. Notes on Hirudo medicinalis, the medicinal leech, as a British species. Proceedings of the Zoological Society of London 11:999–1002. Christiansen, M. 1939. Protoclepsis tesselata (O. F. Muller), der Entengel, als Ursache von Krankheit, u.a. Konjunktivitis, bei Gansen and Enten. Z. InfektKrankheit parasit. Zeitschrift f¨ur Infektionskrankheiten, parasit¨are Krankheiten und Hygiene der Haustiere 55:75–89. Collin, A. 1892. Kleine mittheilungen u¨ ber W¨urmer (Bipalium und Clepsine). Sitzungsberichte der Gesellschaft Naturforschender Freunde zu Berlin 1892:166–170. Davies, R. W. 1973. The geographic distribution of freshwater Hirudinoidea in Canada. Canadian Journal of Zoology 51:531–545. Davies, R. W. 1984. Sanguivory in leeches and its effects on growth, survivorship, and reproduction of Theromyzon rude. Canadian Journal of Zoology 62:589–593. Davies, R. W., and F. R. Govedich. 2001. Annelida: Euhirudinea and Acanthobdellidae. In Ecology and Systematics of North American Freshwater Invertebrates, 2nd ed., J. H. Thorp and A. P. Covich (eds). Academic Press, San Diego, CA, pp. 465–504. Davies, R. W., and J. H. Oosthuizen. 1993. A new species of duck leech from North America formerly
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confused with Theromyzon rude (Rhynchobdellida: Glossiphoniidae). Canadian Journal of Zoology 71:770–775. Davies, R. W., and R. N. Singhal. 1988. Co-sexuality in the leech, Nephelopsis obscura (Erpobdellidae). International Journal of Invertebrate Reproduction and Development 13:55–56. Davies, R. W., and J. Wilkialis. 1980. The population ecology of the leech (Hirudinoidea: Glossiphoniidae) Theromyzon rude. Canadian Journal of Zoology 58:913–916. Davies, R. W., and J. Wilkialis. 1981. A preliminary investigation on the effects of parasitism of domestic ducklings by Theromyzon rude (Hirudinoidea: Glossiphoniidae). Canadian Journal of Zoology 59:1196–1199. Davies, R. W., and J. Wilkialis. 1982. The ecology and morphology of Placobdella papillifera (Verrill) (Hirudinoidea: Glossiphoniidae) in Alberta, Canada. American Midland Naturalist 107:316–324. Davies, R. W., L. R. Linton, and F. J. Wrona. 1982. The passive dispersal of four species of freshwater leeches (Hirudinoidea) by ducks. Freshwater Invertebrate Biology 5:40–44. Dickinson, M. H., and C. M. Lent. 1984. Feeding behavior of the medicinal leech, Hirudo medicinalis L. Journal of Comparative Physiology 154:449– 455. Elliott, J. M., and K. H. Mann. 1979. A key to the British Freshwater Leeches with Notes on Their Life Cycles and Ecology. Freshwater Biological Association Scientific Publication No. 40. Fourie, F. le R., J. H. Oosthuizen, and M. Cross. 1986. A preliminary report on the effects of parasitism by the leeches Theromyzon cooperi and Placobdella goroui on the physiology of the Red billed Teal Anas erythrothyncha. Comparative Biochemistry and Physiology 84A:573–579. Govedich, F. R. 2001. A Reference Guide to the Ecology and Taxonomy of Freshwater and Terrestrial Leeches (Euhirudinea) of Australasia and Oceania. Identification Guide No. 35. Cooperative Research Centre for Freshwater Ecology, Albury, NSW, Australia, 67 pp. Govedich, F. R. 2004. Tender Loving Leeches. Australasian Science 25:16–22. Klemm, D. J. 1985. A Guide to the Freshwater Annelida (Polychaeta, Naidid and Tubificid Oligochaeta, and Hirudinea) of North America. Kendall/Hunt Publishing, Dubuque, IA, 173 pp. Klemm, D. J. 1995. Identification Guide to the Freshwater Leeches (Annelida: Hirudinea) of Florida and Other Southern States. Florida Department of Environmental Protection, Tallahassee, FL, 82 pp.
Kuehnemund, M., and F. Bootz. 2006. Rare living hypopharyngeal foreign body. Head and Neck 28(11):1046–1048. Kutschera, U., and P. Wirtz. 2001. The evolution of parental care in freshwater leeches. Theory in Biosciences 120:115–137. Kuznetsova, O. N. 1955. Leech parasitism of aquatic birds. Ptit*evodstvo 5:32–34. Lang, D. C. 1969. Infestation of ducklings with leeches. Veterinary Record 85:566. Mann, K. H. 1961. Leeches (Hirudinea) Their Structure, Physiology, Ecology and Embryology. Pergamon Press, Oxford, 201 pp. Mason, J. 1976. Studies on the freshwater and terrestrial leeches of New Zealand. Part 2. Orders Gnathobdelliformes and Phararyngobdelliformes. Journal of the Royal Society New Zealand 6:255–276. Moore, J. E. 1964. Notes on the leeches (Hirudinea) of Alberta. National Museum of Canada Natural History Papers 27:1–15. Moore, J. E. 1966. Further notes on Alberta leeches (Hirudinea). National Museum of Canada Natural History Papers 32:1–11. Oosthuizen, J. H. 1993. Redescription of the African duck leech Theromyzon cooperi (Harding, 1932) (Hirudinea: Glossiphoniidae). South African Journal of Zoology 28:67–70. Oosthuizen, J. H., and R. W. Davies. 1992. Redescription of the duck leech, Theromyzon rude (Baird 1869) (Rhynchobdellida: Glossiphoniidae). Canadian Journal of Zoology 70:2028–2033. Oosthuizen, J. H., and R. W. Davies. 1993. A new species of Theromyzon (Rhynchobdellida: Glossiphoniidae) with a review of the genus in North America. Canadian Journal of Zoology 71:1311–1318. Oosthuizen, J. H., and R. W. Davies. 1994. The biology and adaptations of the hippopotamus leech Placobdelliodes jaergerskioeldi (Glossiphonidae) to its host. Canadian Journal of Zoology 72: 418–422. Oosthuizen, J. H., and F. le R. Fourie. 1985. Mortality amongst waterbirds caused by the African duck leech, Theromyzon cooperi. South African Journal of Wildlife Research 15:98–106. Ponomarenko, V. A. 1960. Opyt bor’by s pijavkami vodoplavajuscej pticy. Ptit*evodstvo 10:31. Putter, A. 1907. Der Stoffwechsel der Blutegels (Hirudo medicinalis L.). I. Zeitschrift fur Allgemeine Physiologie 6:217–286. Putter, A. 1908. Der Stoffwechsel der Blutegels (Hirudo medicinalis). II. Zeitschrift fur Allgemaine Physiologie 7:16–61. Richardson, L. R. 1969. On a distinctive new subequatorial Australian quadrannulate land-leech,
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Leech Parasites of Birds and related matters. The Australian Zoologist 15:201–213. Richardson, L. R. 1981. On the Papuan Elocobdella novabritanniae, the Oceanian Abessebdella palmyrae (Haemadipsoidea: Domanibdellidae), and an Oceanian Barbronid (Hirudinea). Records of the Australian Museum 33:673–694. Sawyer, R. T. 1986. Leech Biology and Behaviour, Vols I–III. Clarendon Press, Oxford, 1065 pp. Scudder, G. G. E., and K. H. Mann. 1968. The leeches of some lakes in the southern interior plateau region of British Columbia. Syesis 1:203–209. Singhal, R. N., and R. W. Davies. 1986. Freshwater leeches (Annelida: Hirudinoidea) of India. Trends in Life Sciences 1:95–102. Storer, R. W. 2000. The Metazoan Parasite Fauna of Grebes (Aves: Podicipediformes) and Its Relationship to the Birds’ Biology. Miscellaneous Publications, No.
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188. Museum of Zoology, University of Michigan, Michigan, 90 pp. Trauger, D. L., and J. C. Bartonek. 1977. Leech parasitism of waterfowl in North America. Wildfowl 28:143–152. Tuggle, B. N. 1999. Nasal leeches. In Field Manual of Wildlife Diseases: General Field Procedures and Diseases of Birds, M. Friend and J. C. Franson (eds). United States Geological Survey, Biological Resources Division, Information and Technology Report 1999–001, Washington, DC, pp. 245–248. Wilkialis, J. 1970. Investigations on the biology of leeches of the Glossiphoniidae family. Zoologica Polska 20:29–57. Wilkialis, J., and R. W. Davies. 1980. The population ecology of the leech (Hirudinoidea: Glossiphoniidae) Theromyzon tessulatum. Canadian Journal of Zoology 58:906–912.
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Section V: Arthropods
Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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29 Phthiraptera, the Chewing Lice Dale H. Clayton, Richard J. Adams, and Sarah E. Bush Price et al. (2003) provided a brief history of subsequent taxonomic work on chewing lice. Blood feeding by amblyceran chewing lice was documented as early as 1778, thus establishing the possibility that chewing lice might elicit allergic reactions, transmit pathogens, and leave cutaneous wounds that can fester (Price and Graham 1997; Prelezov et al. 2006). Although blood feeding by some species has been documented in detail (Derylo and Gogacz 1974), the extent of blood feeding by most species of Amblycera remains poorly understood. Ischnoceran chewing lice are recognized vectors of some filarial worms (Bartlett 1993) and cause thermoregulatory stress (Booth et al. 1993), reduced mating success (Clayton 1990), and reduced survival in wild birds (Clayton et al. 1999).
INTRODUCTION Chewing lice are small, wingless, dorsoventrally compressed insects that parasitize birds and some mammals. They range from ≤1 mm to about 10 mm in length. Avian chewing lice belong to one of two suborders: Amblycera, which occur on feathers and skin, or Ischnocera, which are more restricted to feathers. As a group, chewing lice are among the most host specific of all parasites with many species being found on only one genus or species of host. Some species of chewing lice are less specific, however, occurring on multiple host genera, families, or even orders. Most bird lice feed on feathers, dead skin, and skin products, which they appear to metabolize with the aid of endosymbiotic bacteria. Some species also feed on blood, and a few species of Amblycera feed exclusively on blood. Chewing lice are normally found in small, subclinical infestations that are kept in check by regular host grooming, including preening with the bill and scratching with the feet. Because small numbers of lice have no apparent effect on the host, the conventional wisdom has been that chewing lice are relatively benign parasites. When present in large numbers, however, they can cause severe irritation and reduced host survival and reproductive success. They can also affect the host indirectly by serving as vectors of other parasites, including some species of filarial worms. The time and energy that birds must devote to preening to keep lice in check may also be costly. In this chapter we focus on the impact of lice on wild birds.
HOST RANGE AND DISTRIBUTION Many chewing lice are extremely host specific, but host specificity should never be assumed. The systematics of chewing lice has suffered greatly at the hands of taxonomists who describe new species on the basis of host associations, rather than on the basis of the lice themselves. This practice has made it necessary to synonymize hundreds of species over the years (Price et al. 2003). Clayton et al. (2004) reviewed factors governing the host specificity of chewing lice, and the related phenomenon of host-parasite cospeciation. Cospeciation occurs when speciation in a host group is “mirrored” by parallel speciation in its parasites. Repeated bouts of cospeciation yield congruent host-parasite phylogenies. Page (2003) provided an overview of cospeciation, including many examples involving lice. Nearly all species of birds that have been checked have lice. Since many avian hosts have yet to be examined, new species of lice and new host records undoubtedly await discovery, particularly in the tropics. A thoroughly revised comprehensive world checklist and biological overview of chewing lice was published
SYNONYMS Infestation with lice is technically known as pediculosis, although this term is not often used in reference to wild birds. HISTORY Linnaeus included 23 species of chewing lice in his 10th edition of the Systema Naturae ( Linnaeus 1758).
515 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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by Price et al. (2003). It includes a key to the 253 known genera of bird lice. The first confirmed fossil louse was recently described from Eckfeld maar, Germany (Wappler et al. 2004). This fossil is approximately 44 million years old, and feathers preserved in its foregut confirm that it was a bird louse. Louse diversity varies considerably among avian taxa, ranging from just one species per host, as in the case of the Ostrich (Struthio camelus), to more than 20 species per host, as in the case of some tinamous (Ward 1957). It is not clear what factors govern the diversity of lice among avian taxa. Host ecology and behavior may play a role. For example, birds which dive underwater have fewer genera of lice than their nondiving relatives (Fels˜o and R´ozsa 2006). Host morphology may also play a role. Among species of birds, the abundance component of diversity is correlated with host body size and bill morphology (Clayton and Walther 2001). Abiotic factors such as humidity can also influence louse distributions. Species of lice that acquire water from the air are susceptible to desiccation and death in arid environments (Rudolph 1983). Consequently, birds living in arid environments tend to have fewer lice than similar birds in humid environments (Moyer et al. 2002b). Other factors, such as latitude, do not appear to affect parasite richness or abundance (Clayton et al. 1992). The geographic distribution of lice often corresponds to that of the host. However, some lice show a more restricted distribution, being abundant on certain host populations, but rare or absent on others. For example, the louse Quadraceps ridgwayi occurs on Australasian populations of the Eurasian Oystercatcher (Haematopus ostralegus), but is absent from this species in Africa and Eurasia (Clay 1976). In short, lice can exhibit geographic “specificity” that is nested within their host specificity. Geographic specificity of lice can be helpful in elucidating the ecological history of the host. For example, there is little genetic structure among island populations of the endangered Galapagos Hawk (Buteo galapagoensis). However, the lice Deegeriella regalis and Colpocephalum turbinatum on hawks from different islands are genetically distinct, suggesting that the hawk populations on different islands are, in fact, isolated (Whiteman and Parker 2005). In some cases geographic specificity may explain louse distributions better than the relatedness of their hosts. For example, lice from toucans (Ramphastos spp.) are more likely to be found on distantly related toucans in the same geographic region than on more closely related toucans in different regions (Weckstein 2004).
Table 29.1. Higher level classification of lice (Insecta: Phthiraptera). Suborder Amblycera
Family
Menoponidae Boopidae* Laemobothriidae Ricinidae Gyropidae Trimenoponidae Ischnocera Philopteridae† Trichodectidae Rhynchophthirina Haematomyzidae Anoplura (16 families)
Genera Species 68 8 1 3 9 6 138 19 1 49
1,039 55 20 109 93 18 2,698 362 3 532
Families in boldface are found on birds; the others are found on mammals. Data were compiled from Price et al. (2003). * One genus (Therodoxus) occurs on birds (cassowaries). † One genus (Trichophilopterus) occurs on mammals (lemurs).
ETIOLOGY Chewing lice are members of the insect order Phthiraptera, which also contains the sucking lice (Table 29.1). Modern classifications divide Phthiraptera into four suborders, three of which comprise the chewing lice: Amblycera, Ischnocera, and Rhynchophthirina. Members of these three suborders all have mandibulate, chewing mouthparts. Most species of Amblycera and Ischnocera are parasites of birds, although about 12% of the species, along with the three species of Rhynchophthirina, are parasites of mammals. Members of the fourth suborder, Anoplura, all of which parasitize mammals, are known as sucking lice because they have piercing-sucking mouthparts. Durden (2001) provides an excellent review of the sucking lice, as well as taxa of chewing lice found on mammals. Despite sharing mandibulate mouthparts, the three suborders of chewing lice are not the closest relatives to one another. Ischnocera and Anoplura share a more recent common ancestor than either suborder shares with Amblycera (Lyal 1985; Barker et al. 2002; Johnson and Whiting 2002). For this reason, modern workers classify all lice in the single order Phthiraptera, rather than following the traditional classification in which the chewing lice are placed in one order (Mallophaga), with sucking lice in another order (Anoplura). Recent molecular and morphological evidence (Johnson et al. 2004; Yoshizawa and Johnson 2006) also confirms the hypothesis that Phthiraptera is polyphyletic (Lyal 1985). The sister group of Amblycera is actually Liposcelididae, a family of nonparasitic bark/book lice in the order Pscoptera.
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Phthiraptera, the Chewing Lice The two suborders of lice on birds, Amblycera and Ischnocera, are relatively easy to distinguish. Roughly 30% of all species of bird lice belong to the suborder Amblycera. Members of this suborder feed on blood and feathers, have maxillary palps, and concealed antennae with four segments (Figures 29.1a–29.1c). The majority of species of bird lice (70%) belong to the suborder Ischnocera and feed primarily on feathers and dead skin that are metabolized with the aid of endosymbiotic bacteria (f*ckatsu et al. 2007). Members of
(a)
(d)
the Ischnocera lack maxillary palps and have protruding 3–5 segmented antennae (Figures 29.1d–29.1g). The Amblycera are more mobile than the Ischnocera and will make short forays away from the host and abandon a dead host to search for a new one. In contrast, the Ischnocera, which are often called “feather lice,” are so specialized for life on feathers that many species are not even capable of moving from feathers onto the skin of the host (e.g., Clayton et al. 1999).
(b)
(e)
(c)
(f)
(g)
Figure 29.1. Representatives of the six families of avian chewing lice (see Table 29.1). Dorsal morphology to left of midlines, ventral morphology to right. M, male, F, female. Scale bars = 0.5 mm. (a) Colpocephalum holzenthali (Amblycera: Menoponidae), F, host: Barred Forest-Falcon (Micrastur ruficollis); (b) Ricinus sp. (Amblycera: Ricinidae), F, host: passeriformes spp; (c) Laemobothrion maximum (Amblycera: Laemobothriidae), F, host: Buteo sp.; (d) Philopterus sp. (Ischnocera: Philopteridae), M, host: passeriformes spp.; (e) Columbicola columbae (Ischnocera: Philopteridae) M, host: Rock Pigeon (Columba livia); (f) Goniodes australis (Ischnocera: Goniodidae), F, host: Malleefowl (Leipoa ocellata); (g) Heptapsogaster sp. (Ischnocera: Heptapsogasteridae), F, host: Tinamiformes spp. (a) Redrawn from Clayton and Price (1989); (b) redrawn from Ledger (1980); (c) redrawn from Nelson and Price (1965); (d) redrawn from Price and Hellenthal (1998); (e) by the second author; (f) redrawn from Emerson and Price (1986); (g) by the second author.
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Ecologically speaking, bird lice can be divided into five categories based on overall morphology and how they escape host preening: (1) agile Amblycera that run quickly across the skin or feathers (Figure 29.1a); (2) very large Amblycera that slip sideways between the feathers (Figures 29.1b, c); (3) sluggish, triangularheaded Ischnocera that avoid preening by dwelling mainly on the head and neck (Figure 29.1d); (4) elongate Ischnocera that hide between the barbs of wing and tail feathers (Figure 29.1e); and (5) sluggish Ischnocera that burrow into the lush, downy regions of neck and abdominal feathers (Figures 29.1f, g). Although some species cannot be placed into one of these categories, this scheme illustrates the principal adaptive zones occupied by bird lice. EPIZOOTIOLOGY Chewing lice are obligate, permanent parasites that complete their entire life cycle on the body of the host. This cycle consists of the egg, three nymphal instars, and the adult stage. The eggs incubate for 4 to 10 days, depending on the species, and each nymphal instar requires 3–12 days for completion (Marshall 1981). Most adult lice are thought to live about a month, with females producing about one egg per day, for a total of 12–20 eggs. Transmission of chewing lice among hosts often requires physical contact between birds, such as between mates and parents and their offspring in the nest (Hillgarth 1996; Tompkins et al. 1996). However, ischnoceran lice are also capable of moving between hosts by phoresis or “hitchhiking” on hipposboscid flies (Keirans 1975). Phoresis can be common. For example, lice were attached to 44% of 156 hippoboscid flies that were removed from European Starlings (Sturnus vulgaris) (Corbet 1956). Because hippoboscid flies are not generally as host specific as lice, phoresis may help explain why some lice have a wide range of taxonomically diverse avian hosts (Clayton et al. 2004; Harbison et al. in press). Although phoresis is common among the Ischnocera, it is rare among the Amblycera because members of this suborder appear to be morphologically incapable of attaching to flies (Keirans 1975). CLINICAL SIGNS, PATHOLOGY, AND PATHOGENESIS When present in large numbers, amblyceran lice can cause extensive feather and skin damage, leading to dermititis, puritis (itching), insomnia, and excessive preening and scratching. Although ectoparasites that feed solely on blood can cause anemia in their hosts, this has seldom been reported in the case of avian
lice, perhaps because so few species feed exclusively on blood (Marshall 1981; Price and Graham 1997). Poultry lice cause reductions in food consumption, body mass, and egg production as a result of irritation (Nelson et al. 1977; Arends 1997; Prelezov et al. 2006). For example, infestations of the chicken head louse, Cuclotogaster heterographus, cause severe restlessness and debility (Kim et al. 1973) and sometimes kill chicks outright (Loomis 1978). Grooming rates of chickens infested with the louse Menacanthus stramineus also increase significantly (Brown 1974). Despite their potential effects, poultry lice are considered a relatively minor problem in modern operations because they are relatively easy to control. Lice also have negative effects on wild birds. Severe hemorrhagic ulcerative stomatitis and death has been documented in juvenile American White Pelicans (Pelecanus erythrorhynchos) infested with the menoponid louse, Piagetiella peralis, a species that lives within the pouch of these hosts (Samuel et al. 1982; Dik 2006). Although it is not clear whether lice were the principal cause of death, they clearly contributed to poor condition in heavily infested young pelicans. One of the most thorough case studies of the impact of lice on wild birds involves free-ranging Rock Pigeons (Columba livia). Populations of the ischnoceran lice Columbicola columbae and Campanulotes compar increase dramatically on pigeons with naturally or experimentally impaired preening ability (Clayton et al. 2005). These two species feed on abdominal contour feathers (Figure 29.2) and reduce the density of the plumage. This leads to an increase in thermal conductance and a corresponding increase in the metabolic rates of their avian hosts to maintain normal core body temperatures (Figure 29.3). Metabolic rates increase by an average of 8.5% and heavily infested birds need to draw on fat reserves to keep up with these energetic costs, leading to a chronic decline in body mass over several months (Booth et al. 1993). The end result, not surprisingly, is a significant drop in survival during the winter months (Clayton et al. 1999). The impact of feather lice on energetics may also be responsible for a significant drop in the rate of male courtship display, and thus the ability of heavily infested males to attract mates (Figure 29.3) (Clayton 1990). Studies of several other bird species have also demonstrated reductions in the potential attractiveness of lousy males to females (Clayton 1991b). For example, Barn Swallows (Hirundo rustica) with high louse loads have songs of shorter duration than swallows with few lice (Garamszegi et al. 2005). Adult male Satin Bowerbirds (Ptilonorhynchus violaceus) with the most attractive bowers had low infestations of the louse Myrsidea ptilinorhynchi when they were juveniles (Borgia et al. 2004).
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(a)
(b)
Figure 29.2. Damage to Rock Pigeon (Columba livia) feathers by feeding lice (Columbicola columbae and Campanulotes compar). (a) Abdominal contour feathers with (left to right) no damage, average damage and severe damage. The basal downy region and barbules of the basal and medial regions have been consumed (reprinted with permission from the American Zoologist (Clayton 1990). (b) Magnified view of severe damage showing where barbules have been removed by lice. Neither the barbs, nor the rachis itself (=shaft) are damaged, probably because they are too large for the lice to sever with their mandibles. Reprinted with permission from Oxford University Press (Clayton 1991a). Feather damage from chewing lice can have other consequences. The menoponid louse, Hirundoecus (=Machaerilaemus) malleus, chews holes in the tail feathers of Barn Swallows (Kose et al. 1999). These holes may increase feather breakage as well as per-
meability of the feathers to air, thus altering aerodynamic efficiency (Bonser 2001). Barn Swallows with many holes flap more frequently than other swallows (Barbosa et al. 2002), presumably incurring an energetic cost. The holes also increase the potential
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Figure 29.3. Consequences of experimentally increasing the abundance of chewing lice on Rock Pigeons (Columba livia) (combined results of Clayton 1990; Booth et al. 1993; and Clayton et al. 1999). See the text for discussion. costliness of long tails, which appear to function as sexually selected “handicaps” signaling freedom from parasites (Kose et al. 1999). In another study, Cliff Swallows (Petrochelidon pyrrhonota) infested with lice, fleas, and bugs had significantly lower long-term survivorship relative to fumigated, parasite-free controls (Brown et al. 1995), although it was not possible to assess what fraction, if any, of the survival effect could be attributed specifically to lice, fleas, or bugs. The time and energy required for efficient grooming to control lice may also be costly. Both numbers of lice and their species richness can influence grooming rates and the amount of time devoted to these activities (Brown 1974; Cotgreave and Clayton 1994). More time devoted to grooming may mean less time available for other activities, such as foraging and defense of territories. Increased grooming can also reduce vigilance and increase the risk of being killed by a predator (Redpath 1988). Chewing lice can also have indirect effects on the host by acting as vectors or intermediate hosts of other parasites (Table 29.2). For example, the amblyceran louse Trinoton anserium transmits the common heartworm, Sarconema eurycera, to swans when the louse takes a bloodmeal (Seegar et al. 1976; Cohen et al. 1991; Chapter 26), while ischnoceran lice that serve as intermediate hosts for other filarid nematodes transmit these worms when they are ingested during preening (Bartlett 1993). Viruses and bacteria have also been isolated from chewing lice (Table 29.2), but it is not clear whether lice play a role in their transmission. Saxena et al. (1985) provided a detailed review of chewing lice as intermediate hosts and vectors.
DIAGNOSIS In principle, lice are easy to detect because their life cycle is restricted to the body of the host. In practice,
however, some lice are small and can be difficult to see. Some species are also restricted to microhabitats that are difficult to examine, such as the interior of quill feathers. Even lice that are normally found on the surface of feathers can be hidden in the shafts of developing pin-feathers during molt (Moyer et al. 2002a). Methods for collecting and quantifying lice and other avian ectoparasites were reviewed by Clayton and Walther (1997). The five most commonly used methods for quantifying lice include body washing and post-mortem ruffling for dead birds, and dust ruffling, visual examination, and the use of fumigation chambers for live birds (Clayton and Drown 2001). The authors provided a decision tree for choosing among these five methods. Louse nits (eggs) are usually white and are sometimes easier to detect than lice themselves. Although they are small (usually ≤1 mm in length), unhatched eggs can be obvious because they glisten in reflected light and are often laid in clusters. Hatched eggs remain attached to the feathers and are grayish and flattened in appearance. Many species of lice deposit their eggs in regions that are relatively protected from host grooming, such as the gular region (throat), vent region, or between the barbs of feathers (Nelson and Murray 1971). Most species of lice attach their eggs to the base of feathers. Representative photographs and diagnostic drawings of eggs can be found in Balter (1968), Foster (1969), Nelson and Murray (1971), Marshall (1981), and Cohen et al. (1991).
HOST DEFENSE AND IMMUNITY Birds combat lice using a variety of defenses. The simplest defense is to avoid infection. This may be the principal advantage birds gain from choosing louse-free individuals as mates (Clayton 1991b). The most important defense of infested birds against lice is preening.
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Phthiraptera, the Chewing Lice Table 29.2. Parasites and pathogens isolated from bird lice. Bird host Charadriiformes Marbled Godwit (Limosa fedoa) Charadriiformes Whimbrel (Numenius phaeopus) Apodiformes African Swift (Apus barbatus) Gruiformes American Coot (Fulica americana) Podicipediformes Red-necked Grebe (Podiceps grisegena) Anseriformes Mute Swan (Cygnus olor) Tundra swan (Cygnus columbianus) Galliformes Red Junglefowl (Gallus gallus)
Louse Actornithophilus limosae* Carduiceps clayae†
Parasite/pathogen ‡ ‡
Eulimdana wongae Eulimdana wongae‡
Source Bartlett (1993) Bartlett (1993)
Austromenopon phaeopodis* Eulimdana bainae‡ Eulimdana bainae‡ Lunaceps numenii†
Bartlett (1993) Bartlett (1993)
Dennyus hirundinis*
Filaria cypseli‡
Dutton (1905)
Pseudomenopon pilosum*
Pelecitus fulicaeatrae‡
Bartlett and Anderson (1987)
Pseudomenopon sp.*
Pelecitus fulicaeatrae‡
Bartlett and Anderson (1987)
Trinoton anserinum*
Sarconema eurycera‡
Seegar et al. (1976); Cohen et al. (1991)
Eomenacanthus stramineus Escherichia coli§ Eastern equine¶ (= Menacanthus)* Encephalitis Pasteurella multicoida§ Salmonella gallinarum§ Streptococcus equinus§ Menopon gallinae*
Escherichia coli§ Ornithosis bedsoniae§ (=Chlamydophila) Pasteurella multocida§ Streptococcus equinus§
Derylo and Jarosz (1972) Howitt et al. (1948) Derylo (1970) Derylo (1975) Derylo and Jarosz (1972) Derylo and Jarosz (1972) Eddie et al. (1962) Derylo (1970) Derylo and Jarosz (1972)
* Amblycera. † Ischnocera. ‡ Helminth. § Bacteria. ¶ Virus. Wild birds with bill deformities can have enormous louse populations because they are not able to preen efficiently (reviewed by Pomeroy 1962 and Clayton 1991a). Experimental manipulations of the bill confirm that efficient self-preening is critical for controlling louse populations (Brown 1972; Clayton et al. 2005). Similarly, natural “experiments” confirm that scratching with the feet is critical for controlling louse populations on regions that cannot be preened. Birds that cannot scratch because of leg injuries sometimes have
large numbers of lice and nits on the head and neck, but not on regions that the bird can still preen (Clayton 1991a). Allopreening, in which one bird preens another, may also play a role in louse control, although this possibility has not been tested carefully. Other behaviors that may help control lice include dusting, sunning, anting, and “fumigation” of nests with aromatic green vegetation (Hart 1997). Additional research is needed to determine the importance of these behaviors in louse control.
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Feather chemistry is also important in defense against lice. The feathers and skin of several species of birds in the genus Pitohui contain the neurotoxin found in the skin of poison dart frogs (Dumbacher et al. 1992). When given a choice between these feathers and nontoxic control feathers, lice have higher mortality and avoid feeding or resting on the toxic feathers (Dumbacher 1999). A more common feather compound, the pigment melanin, makes feathers more resistant to mechanical abrasion (Bonser 1995). There is some evidence that lice on Barn Swallows avoid feeding on heavily melanized feathers (Kose et al. 1999), although a diet rich in melanin had no effect on lice of Rock Pigeons (Bush et al. 2006). The avian immune system also provides another probable defense against lice that feed on blood or living tissue (Wikel 1996). Some early studies reported inverse correlations between avian “immunocompetence” and size of louse populations (Saino et al. 1995; Eens et al. 2000; Blanco et al. 2001). A recent comparative study by Møller and R´ozsa (2005) reported a positive correlation between the number of genera of amblyceran lice on different species of altricial birds, and the T-cell-mediated immune responsiveness of nestlings of those species. The authors argued that the positive correlation reflects greater specialization of lice on species with strong immune responses. In contrast, there was no correlation between ischnoceran lice and immune responsiveness by the same species. This result makes sense because feather-feeding lice should be invisible to the immune system. More recently, Whiteman et al. (2005) found an inverse correlation between natural antibody (NAb) levels and the abundance of amblyceran lice (Colpocephalum turbinatum) on Galapagos Hawks. In contrast (but in parallel to Møller and R´ozsa’s (2005) results), the hawks showed no correlation between immunity and ischnoceran lice (Deegeriella regalis). Although the results of these studies are intriguing, they should be interpreted with caution. Assessing immunocompetence on the basis of simple assays of immune function is risky. A decline in one component of the immune system can be offset by up-regulation of other components (innate, cell-mediated, or humoral), or by up-regulation of other aspects of the same component (Salvante 2006; Owen and Clayton 2007). Future studies need to explore the relationship between integrated immune responsiveness and lice, preferably in an experimental context. TREATMENT AND CONTROL Jackson (1985) reviewed the use of pesticides for controlling ectoparasites on wild birds in nestboxes. The safest choice is probably pyrethrum dust or spray,
a “fast knock-down, slow killing” insecticide with no side effects on birds or mammals. Pyrethrum is a biodegradable derivative of chrysanthemums that breaks down within hours or days in the environment. Its kill rate is not 100%, so most commercial products use a combination of pyrethrin, a derivative of pyrethrum, and the synergist piperonyl butoxide. A 1.0% concentration of this mixture kills lice effectively, with no side effects on host nestlings or adults (Clayton and Tompkins 1995). PUBLIC HEALTH CONCERNS Bird lice are of little concern to humans because they cannot survive or reproduce off the body of the avian host. Although some bird lice can bite when infested birds are handled, they do not transmit human pathogens. Arthropods from ledge nesting birds occasionally enter dwellings through ventilation ducts or windows and take blood meals from people; however, such reports usually involve nest mites, not lice. DOMESTIC ANIMAL HEALTH CONCERNS Lice are considered a relatively minor problem in modern poultry operations because they are relatively easy to control (Williams 1992). However, they can still be a major problem for poultry kept under traditional conditions, particularly when birds are crowded or in poor health. Arends (1997) and Price and Graham (1997) review the impact of lice on poultry and other domestic birds, and provide details concerning the control of lice on domestic birds. The host specificity of lice on wild birds means that they pose little threat to domesticated birds. WILDLIFE POPULATION IMPACTS AND MANAGEMENT IMPLICATIONS Wildlife managers should be aware that lice can have negative effects on wild birds under certain conditions (e.g., Samuel et al. 1982; Cohen et al. 1991). Although healthy hosts normally keep their louse populations in check, lice can quickly increase on debilitated hosts. This can lead to blood loss, feather damage, irritation, and possible transmission of endoparasites and pathogens (Table 29.2), with effects on individual hosts or entire breeding populations (Samuel et al. 1982). Overcrowding of birds should be avoided because it facilitates transmission of lice, with a subsequent increase in average louse load (Clayton 1991a). For this reason, highly social birds are probably more at risk than solitary birds. Increases in lice can be either a cause or consequence of poor host health, depending on the situation. Although direct effects of lice are usually correlated with the number of lice present, their
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LITERATURE CITED Arends, J. J. 1997. External parasites and poultry pests. In Diseases of Poultry, B. W. Calnek (ed.). Iowa State University Press, Ames, 1080 pp. Balter, R. S. 1968. The microtopography of avian lice eggs. Medical and Biological Illustration 18:166–179. Barbosa, A., S. Merino, F. de Lope, and A. P. Møller. 2002. Effects of feather lice on flight behavior of male Barn Swallows (Hirundo rustica). The Auk 119:213–216. Barker, S. C., M. Whiting, K. P. Johnson, and A. Murrell. 2002. Phylogeny of the lice (Insecta, Phthiraptera) inferred from small subunit rRNA. Zoologica Scripta 32:407–414. Bartlett, C. M. 1993. Lice (Amblycera and Ischnocera) as vectors of Eulimdana spp. (Nematoda: Filarioidea) in charadriiform birds and the necessity of short reproductive periods in adult worms. Journal of Parasitology 79:85–91. Bartlett, C. M., and R. C. Anderson. 1987. Pelecitus fulicaeatrae (Nematoda, Filarioidea) of coots (Gruiformes) and grebes (Podicipediformes)—skin inhabiting microfilariae and development in Mallophaga. Canadian Journal of Zoology 65:2803–2812. Blanco, G., J. De La Puente, M. Corroto, A. Baz, and J. Colas. 2001. Condition-dependent immune defence in the Magpie: How important is ectoparasitism? Biological Journal of the Linnean Society 72:279–286. Bonser, R. H. C. 1995. Melanin and the abrasion resistance of feathers. Condor 97:590–591. Bonser, R. H. C. 2001. Mites on birds. Trends in Ecology and Evolution 16:18–19. Booth, D. T., D. H. Clayton, and B. A. Block. 1993. Experimental demonstration of the energetic cost of parasitism in free-range hosts. Proceedings of the Royal Society of London, B 253:125–129. Borgia, G., M. Egeth, J. A. Uy, and G. L. Patricelli. 2004. Juvenile infection and male display: testing the bright male hypothesis across individual life histories. Behavioral Ecology 15:722–728. Brown, N. S. 1972. The effect of host beak condition on the size of Menacanthus stramineus populations of domestic chickens. Poultry Science 51:162–164. Brown, N. S. 1974. The effect of louse infestation, wet feathers, and relative humidity on the grooming behavior of the domestic chicken. Poultry Science 53:1717–1719.
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Brown, C. R., M. B. Brown, and B. Rannala. 1995. Ectoparasites reduce long-term survivorship of their avian host. Proceedings of the Royal Society of London, B 262:313–319. Bush, S. E., B. R. Moyer, D. Kim, J. Lever, and D. H. Clayton. 2006. Is melanin a defense against feather-feeding lice? The Auk 123:153–161. Clay, T. 1976. Geographical distribution of the avian lice (Phthiraptera): a review. Journal of the Bombay Natural History Society 71:536–547 Clayton, D. H. 1990. Mate choice in experimentally parasitized Rock Doves: Lousy males lose. American Zoologist 30:251–262. Clayton, D. H. 1991a. Coevolution of avian grooming and ectoparasite avoidance. In Bird-Parasite Interactions: Ecology, Evolution, and Behaviour, J. E. Loye and M. Zuk (eds.). Oxford University Press, Oxford, pp. 258–289. Clayton, D. H. 1991b. The influence of parasites on host sexual selection. Parasitology Today 7:329–334. Clayton, D. H., and D. M. Drown. 2001. Critical evaluation of five methods for quantifying chewing lice (Insecta: Phthiraptera). Journal of Parasitology 87:1291–1300. Clayton, D. H., and R. D. Price. 1989. Colpocephalum holxenthali n. sp. (Malloghaga: Menoponidae) from the Barred Forest-falcon Micrastur ruficollis (Falconidae) in Peru. Journal of Parasitology 75:505–507. Clayton, D. H., and D. M. Tompkins. 1995. Comparative effects of mites and lice on the reproductive success of Rock Doves (Columba livia). Parasitology 110:195–206. Clayton, D. H., and B. A. Walther. 1997. Collection and quantification of arthropod parasites of birds. In Host-Parasite Evolution: General Principles and Avian Models, D. H. Clayton and J. Moore (eds.). Oxford University Press, Oxford, pp. 419–440. Clayton, D. H., and B. A. Walther. 2001. Influence of host ecology and morphology on the diversity of Neotropical bird lice. Oikos 94:455– 467. Clayton, D. H., R. D. Gregory, and R. D. Price 1992. Comparative ecology of neotropical bird lice. Journal of Animal Ecology 61:781–795. Clayton, D. H., P. L. M. Lee, D. M. Tompkins, and E. D. Brodie III. 1999. Reciprocal natural selection on host-parasite phenotypes. American Naturalist 154:261–270. Clayton, D. H., S. E. Bush, and K. P. Johnson. 2004. Ecology of congruence: Past meets present. Systematic Biology 53:165–173. Clayton, D. H., B. R. Moyer, S. E. Bush, D. W. Gardiner, B. B. Rhodes, T. G. Jones, and F. Goller. 2005. Adaptive significance of avian beak morphology for
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ectoparasite control. Proceedings of the Royal Society, B 272:811–817. Cohen, S., M. T. Greenwood, and J. A. Fowler. 1991. The louse Trinoton anserium (Amblycera: Phthiraptera); an intermediate host of Sarconema eurycerca (Filarioidea: Nematoda), a heatworm of swans. Medical and Veterinary Entomology 5:101–110. Corbet, G. B. 1956. The life-history and host relations of a hippoboscid fly Ornithomyia fringillina Curtis. Journal of Animal Ecology 25:402–420. Cotgreave, P., and D. H. Clayton. 1994. Comparative analysis of time spent grooming by birds in relation to parasite load. Behaviour 131:171–187. Derylo, A. 1970. Mallophage as a reservoir of Pasteurella multocida. Acta Parasitologica Polonica 17:301–313. Derylo, A. 1975. Badania nad Szkodliwoscia gospodarcza wszolo (Mallophaga) V. proba ustalenia roli Wszolow Eomenacanthus stramineus (Nitzch) W przenoszeniu Tyfusu U kur. Wiadomosci Parazytologiczne 21:61–68. Derylo, A., and E. Gogacz. 1974. Attempts to determine blood amount taken from hens by bird lice (Eomenacanthus stramineus Nitzsch) by using radioactive chromium as a labelling factor. Bulletin of the Veterinary Institute Pulawy 18:50–51. Derylo, A., and J. Jarosz. 1972. Mikroflora jelitowa niektorych wszolow hematofogicznych. Wiadomosci Parazytologiczne 18:113–119. Dik, B. 2006. Erosive stomatitis in a White Pelican (Pelecanus onocrotalus) caused by Piagetiella titan (Mallophaga: Menoponidae). Journal of Veterinary Medicine, B 53:153–154. Dumbacher, J. P. 1999. Evolution of toxicity in Pitohuis: I. Effects of hom*obatrachotoxin on chewing lice (order Phthiraptera). The Auk 116:957–963. Dumbacher, J. P., B. M. Beehler, T. F. Spande, H. M. Garraffo, and J. W. Daly. 1992. hom*obatrachotoxin in the genus Pitohui: chemical defense in birds? Science 258:799–801. Durden, L. A. 2001. Lice (Phthiraptera). In Parasitic Diseases of Wild Mammals, W. M. Samuel, M. J. Pybus, and A. A. Kocan (eds.). Iowa State University Press, Ames, IA, pp. 3–17. Dutton, J. E. 1905. The intermediary host of Filaria cypseli (Annett, Dutton, Elliot). The filaria of the African Swift, Cypselus affinis. Journal of Tropical Medicine 8:108. Eddie, B., K. F. Meyer, F. L. Lambrecht, and D. P. Furman. 1962. Isolation of Ornithosis bedsoniae from mites collected in turkey quarters and from chicken lice. Journal of Infectious Diseases 110:231–237. Eens, M., E. V. Duyse, and L. Berghman. 2000. Shield characteristics are testosterone dependent in both
male and female Moorhens. Hormones and Behavior 37:126–134. Emerson, K. C., and R. D. Price. 1986. Two new species of Mallophaga (Philopteridae) from the Mallee Fowl (Galliformes: Megapodiidae) in Australia. Journal of Medical Entomology 23:353–355. Fels˜o, B., and L. R´ozsa. 2006. Reduced taxonomic richness of lice (Insecta: Phthiraptera) in diving birds. Journal of Parasitology 92:867–869. Foster, M. S. 1969. The eggs of three species of Mallophaga and their significance in ecological studies. Journal of Parasitology 55:453–456. f*ckatsu, T., R. Koga, W. A. Smith, K. Tanaka, N. Nikoh, K. Sasaki-f*ckatsu, K. Yoshizawa, C. Dale, and D. H. Clayton. 2007. Bacterial endosymbiont of the slender pigeon louse Columbicola columbae allied to endosymbionts of grain weevils and tsetse flies. Applied and Environmental Microbiology 73:6660–6668. Garamszegi, L. Z., D. Heylen, A. P. Møller, M. Eens, and F. de Lope. 2005. Age-dependent health status and song characteristics in the Barn Swallow. Behavioral Ecology 16:580–591. Harbison, C. W., S. E. Bush, J. R. Molenke, and D. H. Clayton. In press. Comparative transmission dynamics of competing parasite species. Ecology Hart, B. L. 1997. Behavioural defence. In Host-Parasite Evolution: General Principles and Avian Models, D. H. Clayton and J. Moore (eds.). Oxford University Press, Oxford, pp. 59–77. Hillgarth, N. 1996. Ectoparasite transfer during mating in Ring-necked Pheasants Phasianus colchicus. Journal of Avian Biology 27:260–262. Howitt, B. F., H. R. Dodge, L. K. Bishop, and R. H. Gorrie. 1948. Virus of Eastern Equine Encephalomyelitis isolated from chicken mites (Dermanyssus gallinae) and chicken lice (Eomenacanthus stramineus). Proceedings of the Society of Experimental Biology and Medicine 68:622–625. Jackson, J. A. 1985. On the control of parasites in nest boxes and the use of pesticides near birds. Sialia 7:17–25. Johnson, K. P., and M. F. Whiting. 2002. Multiple genes and the monophyly of Ischnocera (Insecta: Phthiraptera). Molecular Phylogenetics and Evolution 22:101–110. Johnson, K. P., K. Yoshizawa, and V. S. Smith. 2004. Multiple origins of parasitism in lice. Proceedings of the Royal Society of London, B 271:1771– 1776. Keirans, J. E. 1975. A review of the phoretic relationship between Mallophaga (Phthiraptera: Insecta) and Hippoboscidae (Diptera: Insecta). Journal of Medical Entomology 12:71–76.
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Phthiraptera, the Chewing Lice Kim, K. C., K. C. Emerson, and R. D. Price. 1973. Lice. In Parasites of Laboratory Animals, R. J. Flynn (ed.). Iowa State University Press, Ames, pp. 376–397. Kose, M., R. Mand, and A. P. Møller. 1999. Sexual selection for white tail spots in the Barn Swallow in relation to habitat choice by feather lice. Animal Behaviour 58:1201–1205. Ledger, J. A. 1980. The arthropod parasites of vertebrates in Africa south of the Sahara: Phthiraptera. Publications of the South Africa Institute for Medical Research 56:1–327. Linnaeus, C. 1758. Systema Naturae. Editio Decima, Reformata. Impensis Direct, L. Salvii, Holmiae. I:[chewing lice:610–614]. Loomis, E. C. 1978. External parasites. In Diseases of Poultry, 7th ed. Iowa State University Press, Ames, pp. 667–704. Lyal, C. H. C. 1985. Phylogeny and classification of the Psocodea, with particular reference to the lice (Psocodea: Phthiraptera). Systematic Entomology 10:145–165. Marshall, A. G. 1981. The Ecology of Ectoparasitic Insects. Academic Press, London, 459 pp. Møller, A. P., and L. R´ozsa. 2005. Parasite biodiversity and host defenses: chewing lice and immune response of their avian hosts. Oecologia 142:169– 176. Moyer, B. R., D. W. Gardiner, and D. H. Clayton. 2002a. Impact of feather molt on ectoparasites: looks can be deceiving. Oecologia 131:203–210. Moyer, B. R., D. M. Drown, and D. H. Clayton. 2002b. Low humidity reduces ectoparasite pressure: implications for host life history evolution. Oikos 97:223–228. Nelson, B. C., and M. D. Murray. 1971. The distribution of Mallophaga on the domestic pigeon (Columba livia). International Journal for Parasitology 1:21– 29. Nelson, R. C., and R. D. Price. 1965. The Laemobothrion (Mallophaga: Laemobothriidae) of the Falconiformes. Journal of Medical Entomology 2:249–257. Nelson, W. A., J. F. Bell, C. M. Clifford, and J. E. Keirans. 1977. Review Article: Interaction of ectoparasites and their hosts. Journal of Medical Entomology 13:389–428. Owen, J. P., and D. H. Clayton. 2007. Where are the parasites in the PHA response? Trends in Ecology and Evolution 22:228–229. Page, R. D. M. (ed). 2003. Tangled Trees: Phylogeny, Cospeciation, and Coevolution. University of Chicago Press, Chicago, 378 pp. Pomeroy, D. E. 1962. Birds with abnormal bills. British Birds 55:49–72.
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Prelezov, P. N., N. I. Groseva, and D. I. Goundaheva. 2006. Pathom*orphological changes in the tissues of chickens experimentally infected with biting lice (Insecta: Phthiraptera). Veterinarski Archiv 76:207–215. Price, M. A., and O. H. Graham. 1997. Chewing and Sucking Lice as Parasites of Mammals and Birds. U.S. Department of Agriculture, Technical Bulletin No. 1849, 309 pp. Price, R. D., and R. A. Hellenthal. 1998. Taxonomy of Philopterus (Phthiraptera: Philopteridae) from the Corvidae (Passeriformes), with descriptions of nine new species. Entomological Society of America 91:782–799. Price, R. D., R. A. Hellenthal, R. L. Palma, K. P. Johnson, and D. H. Clayton. 2003. The Chewing Lice: World Checklist and Biological Overview. Illinois Natural History Survey, Spec. Publ. 24. Redpath, S. 1988. Vigilance levels in preening Dunlin Caladris alpina. Ibis 130:555–557. Rudolph, D. 1983. The water-vapour uptake system of the Phthiroptera. Journal of Insect Physiology 29:15-25. Saino, N., A. P. Møller, and A. M. Bolzern. 1995. Testosterone effects on the immune system and parasite infestations in the Barn Swallo (Hirundo rustica): an experimental test of the immunocompetence hypothesis. Behavioral Ecology 6:397–404. Salvante, K. G. 2006. Techniques for studying integrated immune function in birds. The Auk 123:575–586. Samuel, W. M., E. S. Williams, and A. B. Rippin. 1982. Infestations of Piagetiella peralis (Mallophaga: Menoponidae) on juvenile White Pelicans. Canadian Journal of Zoology 60:951–953. Saxena, A. K., G. P. Agarwal, S. Chandra, and O. P. Singh. 1985. Pathogenic involvement of Mallophaga. Zeitschrift fuer Angewandfe Entomologie 99:294–301 (In English). Seegar, W. S., E. L. Schiller, W. J. L. Sladen, and M. Trpis. 1976. A mallophaga, Trinton anserium, as a cyclodevelopmental vector for a heartworm parasite of waterfowl. Science 194:739–741. Tompkins, D. M., T. Jones, and D. H. Clayton. 1996. Effect of vertically transmitted ectoparasites on the reproductive success of swifts (Apus apus). Functional Ecology 10:733–740. Wappler, T., V. S. Smith, and R. C. Dalgleish. 2004. Scratching an ancient itch: an Eocene bird louse fossil. Proceedings of the Royal Society of London, B.271(Suppl.):S255–S258. Ward, R. A. 1957. A study of the host distribution and some relationhips of biting lice (Mallophaga) parasitic on birds of the order Tinamiformes. Part II.
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30 Acariasis Danny B. Pence asitic species may cause several conditions commonly known as cystic and nodular mite disease; air sacculitis; respiratory, tracheal, and nasal acariasis; and subcutaneous acariasis.
INTRODUCTION Although thousands of species are recognized, and hundreds of these are parasitic on or in birds, only a few endo- and ectoparasitic mites are recognized as pathogens (Krantz 1978). Mite infections/infestations in wild birds range in clinical severity from nonpathogenic to life-threatening, causing diseases of the skin, feathers, subcutaneous tissues, or respiratory tract. Clinical conditions resulting from acariasis in birds include anemia from exsanguination; dermatitis; mange on the body, feet, and face; feather damage or loss; and granulomatous inflammation in the subcutaneous tissues and in the respiratory tract (Arends 1997). While they have been known for centuries as common parasites affecting individual birds, only recently have certain mites been recognized as the cause of epizootic disease in wild bird populations (Pence et al. 1999). It is neither the intent nor purpose of this chapter to provide a listing of the many species, genera or even families of mites that occur on avian hosts. For such, the reader is referred to the many publications on taxonomy by W. Ateyo, J. Gaud, A. Fain, and many well-known acarologists specializing on specific taxa of mites (see Krantz 1978 for a partial listing). Herein, those few genera and species that cause epizootics in wild bird populations as well as those that evoke a significant pathogenic response resulting in clinical disease will be discussed. They may include some to all of the life history stages (eggs, larvae, protonymphs, deutonymphs including the heteromorphic hypopi, and adults) of representatives from the superorders Parasitiformes (order Mesostigmata) and Acariformes (orders Astigmata and Prostigmata).
HOST RANGE AND DISTRIBUTION Mites are parasitic on/in almost all bird species worldwide. The few pathogenic species are best known as parasites of domestic birds such as fowl, turkeys, and pigeons or in captive caged birds such as finches and psittiforms. Since they are usually discovered sporadically as isolated cases in wild birds (Arends 1997), there are only a few reports of epizootic acariasis in wild bird populations. An epizootic of epidermoptid mange was reported in the Laysan Albatross (Phoebastria immutabilis) from Midway Atoll (Gilardi et al. 2001). Epizootics of knemidocoptic podoacariasis have occurred in the American Robin (Turdus migratorius) in the central and eastern United States; Red-winged Blackbird (Agelaius phoeniceus), Common Grackle (Quiscalus quiscula), and Brownheaded Cowbird (Molothrus ater) in eastern Canada; Evening Grosbeak (Coccothraustes vespertinus) in the southwestern United States; Chaffinch (Fringilla coelebs) in England and Baltic Russia; Sedge Warbler (Acrocephalus schoenobaenus) and European Beeeater (Merops apiaster) in Nigeria; and Eurasian Tree Sparrow (Passer montanus) in Hong Kong (Pence et al. 1999).
ETIOLOGY The Epidermoptidae are ectoparasitic macroscopic species including Epidermoptes bilobatus and Epidermoptes phasianus that cause head mange in domestic fowl and in Ring-necked Pheasants (Phasianus colchicus) (Fain 1965). A related species, Myialges nudus, is responsible for epizootic epidermoptid mange in Laysan Albatross fledglings (Gilardi et al. 2001). Chigger (Thrombiculidae) dermatitis caused by
SYNONYMS Skin diseases are commonly called bird mange; scaley leg; leg mange; podoacariasis; tassel foot; bumble foot; and scaley face. Feather problems include miteinduced depluming; permeable plumage; and feather lengthening, blackening, soiling, and droop. Endopar-
527 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Figure 30.1. Dorsal (left) and ventral (right) views of adult female Knemidokoptes jamaicensis (original). Wormersia midwayensis also has been reported in this species (Sileo et al. 1990). The Knemidokoptidae are submacroscopic endoparasitic mites in the skin of birds. Knemidokoptes jamaicensis (Figure 30.1) is responsible for isolated cases of scaley leg in many species of mostly passerine birds and for epizootic podoacariasis in several species worldwide (Pence et al. 1999). The lethal head and body mange produced by Knemidokoptes pilae is a problem in captive budgeriegers, parrots, and macaws, and, while it occurs in wild psitticines, epizootics have not been reported (Fain and Elsen 1967). Knemidokoptes derooi causes mouth mange in the African Palm-Swift (Cypsiurus parvus). The depluming mite, Neoknemidokoptes laevis, is sometimes a problem in domestic poultry and captive pheasants but has not been reported as a problem in free-ranging pheasants or the Wild Turkey (Meleagris gallopavo) (Arends 1997). Species in the family Harpyrhynchidae are reported as deplumbing and itch mites of captive pigeons, canaries, and finches, but the consequences of the many species infesting wild birds is unknown. The same is true for the many species of quill mites (Syringophill-
idae) in wild birds that cause feather loss in domestic fowl, pigeons, and canaries. Feather mites (Cheyletiellidae, Analgesidae, and Dermoglyphidae) cause pruritis and feather damage in caged birds (Krantz 1978), but their effect on wild birds is unknown. Of the endoparasitic mites, the cyst mite, Laminosioptes cysticola (Laminosioptidae) (Figure 30.2), is well known as the causative agent for fatal fowl nodular disease of the subcutaneous tissues in domestic fowl, turkeys, pheasants, geese, and pigeons (Arends 1997). Although known to occur in wild turkeys, its impact on these hosts is unknown. Most species of birds are infected with host family-, genera-, or sometimes species-specific nasal mites of the families Rhinonyssidae, Speleognathidae, and/or Turbinoptidae (Fain 1956, 1957, 1963; Pence 1975). Rhinonyssus rhinolethrum is found commonly in ducks and geese. Like most other species of nasal mites, it appears not to evoke serious pathogenic lesions in its hosts. An important exception is the sometimes fatal nasal and respiratory tract rhinonyssid, Sternostoma trachaecolum (Figure 30.3), of caged birds such as canaries and finches (Fain and Hyland 1962). This mite also occurs in many wild birds
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Figure 30.2. Dorsal (left) and ventral (right) views of adult female Laminosioptes cysticola (redrawn after Fain 1981).
worldwide and is very common in certain species such as the Great Crested Flycatcher (Myiarchus crinitus) in the southeastern United States. However, its effect on infected wild birds remains unknown. Another, sometimes pathogenic, air sac mite in domestic birds is the tiny Kitodites (=Cytodites) nudus (Figure 30.4) found in chickens, turkeys, pheasants, pigeons, canaries, and Ruffed Grouse (Bonasa umbellus) (Arends 1997). Most nonpasserine orders of birds are infected with subcutaneous heteromorphic deutonymphs (hypopi) of hypoderatid mites (Hypoderatidae) of several genera, including Phalacrodectes, Neottialges, and Hypodectes (Figure 30.5) (Fain 1967). They are especially
common in the subcutaneous adipose tissue beneath the skin in the axillary and groin area of most birds. EPIZOOTIOLOGY All birds, domestic or wild, are infested with any number of species of mites from several families occupying almost every available niche of the skin and feathers. Some of these may be fairly host specific to the bird genus or family level while other species of mites may infest many different unrelated bird hosts. Thus, wild birds may sometimes be important in the transmission of mites to domestic species. For example, the wild
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Figure 30.3. Dorsal (left) and ventral (right) views of adult female Sternostoma trachaecolum (original). House Sparrow (Passer domesticus) appears to be the reservoir for several dermanyssid poultry mites worldwide, because of the practice of lining their nests with chicken feathers (Arends 1997). Certainly, the role of wild passerines is well known in their “seeding” of the environment with larval chigger mites (Thrombiculidae) that subsequently infest humans as well as other domestic animals and birds (Krantz 1978). All the life history stages including eggs, larvae, nymphs, and adults of most avian acari can be found on/in the same host individual. Transmission between hosts may be by direct contact or at nest sites by actively motile larvae, nymphs, or adults (Krantz 1978). The important exceptions are the Thrombiculidae, Speleognathidae, and Hypoderidae that have parasitic larval and nymphal stages, but free-living adults (Fain 1967). The quiescent hypoderid heteromorphic deutonymphs localized in the subcutaneous fat of many birds become activated at the time of nesting, metamorphose to adults and exit through the skin of adult birds. The submacroscopic adult mites live and mate in the nesting material, the females produce eggs that
hatch into larvae, and the microscopic larvae penetrate the intact skin of the nestlings. The larvae localize in the adipose tissues as heteromorphic deutonymphs that slowly mature over the next year to sixth-stage hypopi. PATHOGENESIS, PATHOLOGY, AND CLINICAL CORRELATIONS Similar to psoroptid mites in mammals, epidermoptid mites burrow and tunnel through the cornified layers of the skin of birds. By mechanical irritation and probably through release of excretory and secretory products, including keratinases, the mites can evoke severe mange in infected birds. Epidermoptids such as M. nudus in Laysan Albatross fledglings can cause granulomatous inflammation, with epithelioid histiocytes as the dominant cell type, accompanied by diffuse hyperkeratosis, multifocal dermal edema, and cellular infiltrates of heterophils with scattered eosinophils and lymphocytes in the dermis of infected birds (Gilardi et al. 2001). In occasionally fatal cases, these lesions may progress to more severe granulomatous inflammation
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Figure 30.4. Dorsal (left) and ventral (right) views of adult female Kytodites nudus (original).
with Langhans’ giant cells replacing large areas of the superficial dermis and obscuring the dermoepithelial junction. Mites can sometimes be observed within dilated feather follicles. Despite the severe granulomatous dermatitis, superficially the skin and feathers of infected birds appear normal (Gilardi et al. 2001). The release of proteolytic enzymes, formation of a feeding stylostome, and subsequent ingestion of liquified tissue are a well-known mechanism of pathogenicity for the dermatitis caused by chiggers (Krantz 1978). There is a density-dependent relationship between the degree of infestation and the severity of the disease in the host. Fatal chigger dermatitis in Laysan Albatross fledglings presents as a severe necrotizing dermatitis. The effects of trombidiosis in this host ranged from mild dermatitis and edema to massive edema, focal hemorrhaging of the subcutaneous tissues of the crura and abdomen, and death with carcasses demonstrating ketosis, generalized pallor, and severe anemia (Sileo et al. 1990). The pathogenesis of the knemidokoptids in birds resembles that of sarcoptid mites in mammals. Depending on the species, they have a predilection for the skin of the legs and feet and/or face and beak area where they burrow to the level of the stratum germativium or on the wings and thigh where they burrow into the
feather follicles. Mechanical trauma from burrowing as well as excretory and secretory products released from the mites in situ contributes to the skin changes. The lesions of epizootic podoknemidokoptiasis or scaley leg caused by K. jamaicensis range from a white powdery scaling to proliferative epidermal overgrowth with massive crusts and scab formation resulting from massive hyperkeratosis and intense dermal inflammation on the unfeathered part of the legs and feet. Grossly, the skin may become markedly thickened and very rough, gray-white in color, desiccated, and fractured (Figure 30.6); under a stereo microscope there are numerous 0.1 mm black orifices on the surface of the skin (Pence 1975). There may be severe diffuse hyperkeratosis sometimes with sloughing of the nails, loss of digits, or traumatic amputation of the entire foot in severe cases (Pence et al. 1999). The abundant submacroscopic mites are often arrayed in a striking honeycombed pattern within the cornified epithelium (Figure 30.7), with scattered aggregates of eosinophils, lymphocytes, and histiocytes in the epidermal scale and in the underlying dermis (Pence 1970). Podoacariasis caused by K. pilae can occur in conjunction with scaley-face mange at the base of the beak and around the eyes and nares in captive, and probably in wild, psitticines (Fain and Elsen 1967). Lesions are similar
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Figure 30.5. Dorsal (left) and ventral (right) views of two hypopi of Hypodectes nudus embedded in the axillary fat of a Mariana Fruit-Dove (Ptilinopus roseicapilla). Bar = 250 μm.
to those caused by K. jamaicensis in passerines, especially the small black orifices on the skin surface and the honeycombed pattern apparent on removing the superficial stratum corneum. Lesions may also be found on the body around the cloacal vent, on the wings, and on the thighs. As in infections with the depluming mite, Neoknemidokoptes laevi, there may be loss of feathers in and around the head and body lesions. Proliferation of keratinaceous overgrowths sometimes results in beak deformities including elongation, flattening, thinning, crossing, clefting, and/or splitting. In cases of depluming caused by knemidokoptids, the mites burrow into the basal shafts of the feathers on the epidermis leaving a well-defined orifice in the feather shaft. They produce the same white powdery material seen on the legs and face which may partly fill the feather follicle, evoke an intense irritation of the shaft and follicle, and cause the bird to eventually pull out the infected feather (Fain and Elsen 1967; Arends 1997). Slight thickening of the surface epithelium is noted in the lateral aspects of the internal surface of the mouth near the commissures of the bill in the African Palm-Swift infected with K. derooi (Fain 1970). Beak lesions and sometimes leg lesions caused by K. fossor are described by
Figure 30.6. Excised leg of an American Robin (Turdus migratorius) showing the extensive hyperkeratosis of scaley leg from an infection with Knemidokoptes jamaicensis. Fain and Elsen (1967) from several captive passerines in the Antwerp Zoo in Belgium. The fowl cyst mite, L. cysticola, is found inside subcutaneous yellowish nodular granulomas of several centimeters diameter (Arends 1997). These transform into caseocalcareous deposits surrounding dead mites and are responsible for the condemnation of carcasses of heavily infected domestic poultry. The consensus of most (Kettle 1990; Arends 1997) is that the inflammation and destruction of muscle fibers cause loss of condition, emaciation, and even death of some heavily infected birds. Others contend that even massive infections of these mites are of little consequence to infected birds (Urquhart et al. 1996). Certainly, dead mites are known to evoke nodular subcutaneous granulomas, with larger numbers of nodules found in older birds. Similar lesions in the lungs are reported to cause
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Figure 30.7. Low power magnification of a histologic section of the dermoepithelial junction of the foreleg of an Americal Robin (Turdus migratorius) with scaley leg from infection with Knemidokoptes jamaicensis. Note the honeycombed pattern in the epidermis containing numerous mites (arrows). H&E. Bar = 200 μm.
death in pigeons (Arends 1997). The effects of infection with any of the several species of laminosioptids that occur in wild birds are unknown (Fain 1981). Tarsal claws are used to perforate the nasal mucosa by rhinonyssid mites such as R. rhinolethrum in the nasal passages of waterfowl; this allows them to feed on tissue and blood (Feider and Mironescu 1972). Mechanical irritation evokes hyperplasia and chronic inflammation of the nasal and respiratory mucosa pro-
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duces an abundance of mucous and causes dilatation of the blood vessels which become engorged with erythrocytes (Feider and Mironescu 1972). So, heavy infestations of even those species of rhinonyssids strictly confined to the nasal passages may not always be inconsequential. Here, an edematous and bloody mucosa throughout the nasal sinuses is occasionally observed (D. B. Pence, unpublished observations). In captive canaries and goldfinches heavy infections of the nasal, tracheal, and air sac rhinonyssid, S. trachaecolum, may cause severe inflammatory tracheitis and airsacculitis leading to a fatal pneumonia (Fain and Hyland 1962). Although many species are reported as hosts of S. trachaecolum (Fain and Hyland 1962; Pence 1975), clinical disease has not been seen in wild free-living birds. It has been suggested that wild birds are more often parasitized in the nasal passages than in the lower respiratory tract and that they are refractory to the more serious stress-induced clinical conditions seen in their captive counterparts (Fain and Hyland 1962). However, it should be remembered that any heavily infected wild bird that is seriously ill with pneumonia undoubtedly would be very quickly removed from the population rendering such conditions in wild birds extremely difficult to determine. The pathogenesis of the cytoditid air sac mites in domestic poultry, pheasants, pigeons, and Ruffed Grouse is density-dependent. Only heavy infections of K. nudus can cause weakness, emaciation, pneumonitis, and/or peritonitis (Arends 1997). Fatal granulomatous pneumonia results from obstruction of air passages from chronic inflammation and accumulation of mucous in the air sacs (Lindt and Kutzer 1965). Clinical signs resemble those of tuberculosis with wheezing, coughing, and weight loss in infected birds. The ramifications of this infection in Ruffed Grouse and other wild birds are unknown. Additionally, there are several other species of Kitodites and the related genus Cytonyssus that infect wild birds (Fain and Bafort 1964); clinical disease has not been reported. Hypopi of the Hypoderatid mites appear as submacroscopic whitish to yellowish cylindrical bodies in the subcutaneous adipose tissue (Figure 30.5). They evoke a very mild to sometimes granulomatous inflammatory response in the subcutaneous tissues, especially in the dermal adipose tissue of many birds (Gr¨unberg and Kutzer 1962; Hendrix et al. 1987; D. B. Pence, unpublished data). Having no mouth or digestive tract, it has been proposed that nutrient transport is through the exoskeleton of the hypopi or perhaps through the openings at the genital plate (Fain 1967). The somatic cells of these hypopi contain large quantities of the same neutral lipids that occur in the adjacent host adipose tissues (D. B. Pence, unpublished data). The inflammatory response to hypoderatids in
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the subcutaneous adipose tissue ranges from a few foamy macrophages and plasma cells in the vicinity of the living hypopi in some hosts (Hendrix et al. 1987) to a diffuse granulomatous inflammation in many heavily infected birds (D. B. Pence, unpublished data). The latter is characterized as having only a few lymphocytes, eosinophils, and fusiform fibroblasts and with mostly epithelioid histiocytes as the predominant cell circumferential to the living hypopus, but without an attendant organized fibroplasia surrounding the lesion (D. B. Pence, unpublished data; Schwan and Sileo 1978). Severe granulomatous inflammation with a peripheral ring of Langhans’ giant cells surrounding masses of epithelioid histiocytes, lymphocytes, plasma cells, and eosinophils is sometimes seen adjacent to degenerating hypopi in the subcutaneous tissues (D. B. Pence, unpublished data). There are usually no clinical signs of disease associated with infections of the subcutaneous hypopi of hyperderatid mites. They are usually discovered, sometimes in large numbers, at necropsy of birds having died of other causes. Hypopi are found most commonly in the axillary and groin fat, but sometimes they are seen in the deeper subcutaneous tissues of the esophagus and trachea, pericardial sac, abdominal cavity, and between the fascia of the large thigh muscles. DIAGNOSIS Because of their submacroscopic to microscopic size and predilection for unusual sites in the host, many kinds of acariases are difficult to diagnose, especially the species of endoparasitic mites. Deep skin scrapings examined in 10% KOH are very helpful in cases of epidermoptid and knemidokoptid mange. The knemidokoptids appear as tiny white immobile spheres. Examination of feather shafts under a stereo microscope is useful in diagnoses of syringophilid quill mites and depluming knemidokoptids. Ectoparasitic mites can be dislodged from the skin and feathers by brushing, especially around the head and neck with a stiff bristled brush such as an old toothbrush. The residue from these brushings should be examined with a stereo microscope. Laminosioptids are most often diagnosed by finding dead mites or their remains in the excised yellowish subcutaneous nodules crushed under a cover glass in a drop of acidulated water and examined microscopically. All rhinonyssid nasal and respiratory mites are macroscopic and usually darkly pigmented such that they are difficult to see in the anterior nasal epithelium, but they are sharply contrasted against the posterior turbinates and the nasal and respiratory mucosa. They are usually discovered at necropsy. The smaller cytoditids are more difficult to diagnose, but close inspection at necropsy may reveal white dots moving
slowly over the surface of the air sacs. Hypopi usually appear at necropsy as scattered to very numerous immobile submacroscopic cylindrical whitish cystlike structures in the subcutaneous tissues, especially in the adipose tissue of the medial and lateral aspects of the thighs and in the inguineal region. Examination under a stereo microscope will reveal the characteristic deeply pigmented chitinized substructure (apodemes) of the anterior and posterior pairs of legs. Mites collected from their hosts can be fixed and stored indefinitely in 70% ethyl alcohol. Species can be identified only after light microscopic study of cleared specimens mounted in Hoyer’s medium. At this point, species can usually be identified by examination of their many varied morphological features.
IMMUNITY There have been no studies on the immune response of wild or domestic birds to acariases.
PUBLIC HEALTH CONCERNS Dermanyssid mites that occur on domestic birds in poultry facilities can sometimes be carried by wild birds, especially House Sparrows. They can cause transient dermatitis in humans exposed to them. Also, many passerines worldwide are important in spreading chiggers (larval trombiculids) that are human pests that cause chigger dermatitis and transmit the rickettsiosis, tsutsugamushi disease (scrub typhus) caused by Orientia tsutsugamushi in Southeast Asia and the Indo Pacific. Otherwise, there are no public health problems associated with acariasis in wild birds.
DOMESTIC ANIMAL HEALTH CONCERNS Host specificity varies dramatically across different species, genera, and families of mites. Thus, while cross transmission of many mite species is possible between their wild and domestic bird hosts, just how frequently this occurs is unknown. Certainly, the House Sparrow and possibly the European Starling (Sturnus vulgaris) and Rock Pigeon (Columba livia) have been responsible for the introduction of any of several dermanyssids such as the chicken mite (Dermanyssus gallinae), northern fowl mite (Ornithonyssus sylviarum), and tropical fowl mite (Ornithonyssue bursa) into domestic poultry flocks (Arends 1997). Recently captured wild passerines should be isolated for some time prior to mixing with long-term captive residents in a facility in order to avoid establishing knemidokoptid and other highly contagious mite infections or infestations.
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Acariasis WILDLIFE POPULATION IMPACTS Epizootic knemidokoptic mange has been reported in numerous passerine populations in North America, Europe, Asia, and Africa (Pence et al. 1999). Many individuals may be lost from the population during such epizootics, but it is unknown whether or not the impact of this disease on these avian populations is compensatory with other mortality factors. The same is true for the impact of epizootic epidermoptid and trombiculid dermatitis in Laysan Albatross populations. And, while heavy infections of many other species of endoparasitic mites (tracheal and respiratory mites), subcutaneous hypopi, and massive infestations of ectoparasitic species (skin and feather mites) may extract their toll from decreased fitness and perhaps reproductive potential for individual birds, their importance to the population dynamics of their host species is unknown. So, while mites are sometimes annoying to the individual bird, and they are occasionally responsible for mortality events, it is difficult to envision any significant population impact that mites could induce on any managed game bird species. Thus, the need for any kind of management intervention in wild bird populations is questionable.
TREATMENT AND CONTROL Permethrin spray may be used to treat for dermanyssids on transport cages or in resident facilities holding captive wild birds. Ivermectin has been used to treat wild birds with knemidokoptic podoacariasis, but it was not always successful as a prophylactic (Pence et al. 1999).
LITERATURE CITED Arends, J. J. 1997. External parasites and poultry pests. In Diseases of Poultry, 10th ed. Iowa State University Press, Ames, IA. Fain, A. 1956. Les acariens de la famille Rhinonyssidae Vitzhum 1935, parasites des fosses nasale des oiseaux au Ruanda-Urundi (note preliminaire). Revue de Zoologie et de Botanique Africaines 53:131–157. Fain, A. 1957. Les acariens des familles Epodermoptidae et Rhinonyssidae parasites des fosses nasales d’oiseaux au Ruanda-Urundi et au Congo belge. Annales du Musee Royal du Congo belge (Tervuren). Serie in Octavo, Sciences Zoologiques 60:1–176. Fain, A. 1963. Chaetotaxy et classification des Speleognathinae (Acari: Thrombidiformes). Bulletin de l’Institute Royale des Sciences Naturelles de Belgique 39:1–80. Fain, A. 1965. A review of the family Epidermoptidae Touessart parasitic on the skin of birds (Acarina: Sarcoptiformes). Verhandelingen van de Koninklijke
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Vlaamse Academie voor Wetenschappen, Letteren en Schone Kunsten Van Belgie, Klasse der Wetenchappen 27:Pt. I, 1–176; Pt. II, 1–144. Fain, A. 1967. Les hypopes parasites des tissues cellulaires des oiseaux (Hypodectidae: Sarcoptiformes). Bulletin de l’Institute Royale des Sciences Naturelles de Belgique 43:1–139. Fain, A. 1970. A new species of Knemidokoptes producing mange in the palm-swift (Acarina: Sarcoptiformes). Revue de Zoologie et de Botanique Africaines 81:220–224. Fain, A. 1981. Notes on the genus Laminosioptes Megnin, 1880 (Acari, Astigmata) with descriptions of three new species. Systematic Parasitology 2:123–132. Fain, A., and J. Bafort. 1964. Les acariens de la famille Cytoditidae (Sarcoptiformes). Description de sept especies nouvelles. Acarologia 6:504–528. Fain, A., and K. E. Hyland. 1962. The mites parasitic in the lung of birds. The variability of Sternostoma trachaecolum Lawrence, 1948, in domestic and wild birds. Parasitology 52:401–424. Fain, A., and P. Elsen. 1967. Les acariens de la famille Knemidokoptidae producteurs de gale chez oiseaux (Sarcoptiformes). Acta Zoologica et Pathologica Antverpiensia 45:1–142. Feider, Z., and I. Mironescu. 1972. Une modalit´e particuli`ere de perforer la muqueuse nasale, utilis´ee par les acariens de la famille Rhinonyssidae Troussart, 1895. Acarologia 14:21–31. Gilardi, K. V. K., J. D. Gilardi, A. Frank, M. L. Goff, and W. M. Boyce. 2001. Epidermoptid mange in Laysan albatross fledglings in Hawaii. Journal of Wildlife Diseases 37:490–498. Gr¨unberg, W., and E. Kutzer. 1962. Deutonymphen von federmilben in der subkutis von Tantalus leucocephalus (indischer nimmersatt). Zeitschrift f¨ur Parasitenkunde 21:542–599. Hendrix, C. M., R. P. Kwapien, and J. R. Porch. 1987. Visceral and subcutaneous acariasis caused by hypopi of Hypodectes propus bubulci in the cattle egret. Journal of Wildlife Diseases 23:693–697. Kettle, D. S. 1990. Medical and Veterinary Entomology. C.A.B International, Wallingford, UK. Krantz, G. W. 1978. A Manual of Acarology, 2nd ed. Oregon State University Bookstores, Inc., Corvallis. Lindt, S., and E. Kutzer. 1965. Luftsackmilben (Cytodites nudus) als Ursache einer granulomatosen Pneumonic biem Huhn. Pathologia Veterinaria 2:264–276. Pence, D. B. 1970. Knemidokoptes jamaicensis Turk from the rufous-sided towhee in Louisiana. Journal of Medical Entomology 7:686. Pence, D. B. 1975. Keys, species and host list, and bibliography for nasal mites of North American birds
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(Acarina: Rhinonyssinae, Turbinoptinae, Speleognathinae, and Cytoditidae). Special Publications of the Museum of Texas Tech University 8:1–148. Pence, D. B., R. A. Cole, K. E. Brugger, and J. R. Fischer. 1999. Epizootic podoknemidokoptiasis in American robins. Journal of Wildlife Diseases 35:1–7. Schwan, T. G., and L. Sileo. 1978. Neottialges (Pelecanectes) evansi (Sarsoptiformes: Hypoderidae)
parasitizing a white-necked cormorant in Kenya. Journal of Medical Entomology 14:22. Sileo, L., P. R. Sievert, and M. D. Samuel. 1990. Causes of mortality of albatross chicks at Midway atoll. Journal of Wildlife Diseases 26:329– 338. Urquhart, G. M., J. Armour, J. L. Duncan, A. M. Dunn, and F. W. Jennings. 1996. Veterinary Parasitology, 2nd ed. Blackwell Science, Oxford.
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31 Black Flies (Diptera: Simuliidae) Douglas C. Currie and D. Bruce Hunter INTRODUCTION Black flies are a relatively small family of nematocerous Diptera with approximately 2,000 described species. These are distributed worldwide, particularly in areas where there are fast-flowing streams that serve as habitat for immature stages. The adults are small (1.2–6.0 mm long) hunch-backed flies with cigarshaped antennae and broad wings. Simuliids are best known for the biting habits of adult females, which typically require a blood meal to develop their eggs. They rank among the world’s most notorious pests for people and livestock, particularly in northern temperate regions. Their painful bites may incite local and/or systemic allergic reactions and can cause significant economic losses when outdoor business and recreational activities are affected. Black flies have caused serious problems in cattle raising areas in Canada and Central/Eastern Europe by harassing cattle, altering their behavior, reducing productivity, and even causing death. Black flies also act as vectors for a number of important pathogens of people, livestock, and birds, including arboviruses such as vesicular stomatitis in cattle, Onchocerca spp. in people and cattle, and Leucocytozoon spp. and Trypanosoma spp. in birds. The role of black flies as the main vector of Leucocytozoon is well documented in wild, captive, and domestic birds (Chapter 4). There is also growing evidence that swarming, direct harassment, and blood feeding may contribute to increased nestling mortality, early fledging, and altered roosting behavior in several species of raptors and passerines.
HISTORY The earliest incontrovertible simuliid fossils date to the late Jurassic (Kalugina 1991; Currie and Grimaldi 2000); however, the fossil record of related families suggests that black flies originated during even earlier times, perhaps suggesting a Pangean or effectively Pangean origin for the family. This early origin of simuliids raises questions about what hosts might have been available to females. Records of bloodsucking on invertebrates and cold-blooded vertebrates have not been confirmed, and it must be presumed that the earliest simuliids blood fed exclusively on homeothermic animals, as they do today. Given that the earliest birds appeared only during mid-Jurassic times and that mammals (although present since the Jurassic) did not begin their radiation until the Tertiary period, it seems likely that simuliids began their sanguinary habits on another, more abundant, group of organisms, namely, homeothermic dinosaurs. Of course, the distinction between birds and dinosaurs during Jurassic times is moot, given that Aves are phylogenetically extant dinosaurs. The earliest example of an ornithophilic black fly is Archicnephia ornithoraptor—an Upper Cretaceous-aged fossil preserved in New Jersey amber (Currie and Grimaldi 2000). The role of black flies as vectors of Leucocytozoon spp. has been known since the early 1930s with the demonstration by O’Roke (1930) and Skidmore (1931) that black flies transmit Leucocytozoon simondi to ducks and Leucocytozoon smithi to turkeys. In spite of extensive work on Leucocytozoon in birds since then (Chapter 4), the pathogenicity of ornithophilic black flies themselves remains poorly documented. It is only relatively recently that attention has been drawn to direct effects of black fly feeding on nestling survival and on behavior of juvenile birds (Hill 1994; Hunter et al. 1997; Smith et al. 1998; Gaard 2001).
SYNONYMS Black fly (also black-fly and blackfly), buffalo gnat (USA), mouche noire (French-speaking Canada), sand fly (New Zealand and Australia), simuliid, reed smut, Kriebelm¨uke (German), pium (Brazil), borrachudo (southern Brazil), jejenes (Venezuela), bocones (Costa Rica), moshka (Russia), potu (India), mawi (Africa).
DISTRIBUTION Simuliids can be found virtually everywhere there is freely flowing, unpolluted water—the habitat of the
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immature stages. Only Antarctica, certain archipelagos (e.g., Hawaii, Falkland Islands) and isolated desert islands are unpopulated by members of the family. Some geographical areas (e.g., South Asia, New Guinea) remain inadequately surveyed for black flies, and it is estimated that the actual number of simuliids will easily exceed 3,000 species (Currie and Adler 2008). HOST RANGE Black flies are known to take blood meals from a wide variety of avian hosts and it seems unlikely that any major group of birds is immune to attack. So far, hematophagy by ornithophilic simuliids has been confirmed for at least 13 orders of birds (Crosskey 1990). The degree of host specificity varies among simuliids, with some species exhibiting a narrow host preference while others are more eclectic. However, the host preferences of most species remain poorly studied. The most host-specific black fly known is Simulium annulus (Lundstr¨om), which has so far been confirmed to feed only on Common Loons (Gavia immer). Extracts from the uropygial gland of loons are believed to play an important role in attracting females of S. annulus to their host (Fallis and Smith 1964; Lowther and Wood 1964). More typically, simuliids have a range of hosts that fit a particular profile (e.g., “small mammals,” “ungulates,” “passeriform birds,” and “galliform birds”). Most female black flies can be assigned to one of two broad categories of blood-feeding habit based on their host preference for mammals (mammalophily) or birds (ornithophily). No species of black flies confines its bloodsucking to humans, but mammalophilic species that include humans as hosts are called anthropophilic. Bloodsucking records on invertebrates and cold-blood vertebrates have not been confirmed. A small number of arctic- or mountain-adapted species have reduced mouthparts that are not capable of piercing the skin (autogenous). Such obligatorily autogenous species are able to develop their eggs exclusively from nutrients gathered during the larval stage. Bloodsucking (anautogenous) species of black flies are classified as ornithophilic or mammalophilic based on the form of the female tarsal claw (Shewell 1955). Ornithophilic species have an accessory tooth of variable size near the base of the claw, which presumably aids in grasping the feathers (Figure 31.1). In contrast, the claw of mammalophilic species is typically in the form of a simple talon, without a markedly developed subbasal tooth. Unfortunately, the distinction between the two feeding groups is not clear in all instances, with certain species taking blood from both birds and mammals regardless of claw structure. For example, the simple-clawed female of Simulium venustum is known
Figure 31.1. Scanning electron micrograph of the bifid claws typical of ornithophilic black flies. These claws are adaptations for grasping feathers. Courtesy of D. A. Craig, University of Alberta. to feed on various species of birds, although their numbers are low relative to well-established ornithophilic species with bifid claws (Bennett 1960). In contrast, S. venustum is recognized as among the most notorious pests of mammals in North America (Hunter et al. 1993; Adler et al. 2004). Despite such exceptions, a blood-meal study of black flies confirms the fidelity of claw type and host: species with bifid claws fed overwhelmingly on avian hosts, whereas species with simple claws fed predominantly on mammalian hosts (Malmqvist et al. 2004). ETIOLOGY The Simuliidae are most closely related to the Ceratopogonidae (biting midges), Chironomidae (nonbiting midges), and Thaumaleidae (solitary or trickle midges), which together comprise the nematoceran superfamily Chironomoidea. There are no comprehensive phylogenies of the Simuliidae, and different schemes of classification are favored by specialists from different geographical regions. Adler et al. (2004) provided a cladistic analysis of the Holarctic Simuliidae, which agreed largely with Moulton’s (2000) suprageneric analysis of world black flies based on his analysis of molecular sequence data. Both studies recognize a two-subfamily arrangement of black flies, including the Parasimuliinae and the Simuliinae. The latter subfamily is further subdivided into two tribes: the Prosimuliini and the Simuliini. This two-subfamily and two-tribe arrangement has gained increased acceptance in recent years, although a more finely divided suprageneric classification continues to be favored by certain specialists. There is no comprehensive
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anterolateral corners of the thorax and project anteriorly or anterodorally into the current (Figure 31.3). The gill, which aids in respiration, typically consists of a number of slender filaments, but in certain species the gill has a tubular or clublike form. Upon completion of a cocoon, the larva retreats inside and transforms into a pupa. Once fully developed, the adult emerges through a T-shaped slit in the pupal thorax and rises to the water surface in a bubble of gas. More detailed information about the life history of black flies can be found in Crosskey (1990). The life span of ornithophilic black flies is not well established and likely varies among species. A study in Algonquin Park, Ontario, based on recapture of female flies (Simulium rugglesi) fed on ducks injected with a radioactive isotope showed that flies can live as long as 28 days, that is, sufficient time for the flies to accomplish two ovipositions (Bennet 1963). Black fly eggs can undergo periods of dormancy (diapause) in order to survive periods of temperature extremes or drought conditions. Diapause is usually ended when environmental conditions become once again favorable for fly survival. Some northern species overwinter in the larval stage, but most overwinter as eggs and hatch when the water temperatures rise in the spring (Crosskey 1990). Water temperature and food supply regulate the rate of larval development. In northern climates usually not more than two generations of flies develop over a season, whereas further south there may be as many as four or five generations of flies.
Growth and Development Like other true flies, black flies pass through four stages to complete their life history: egg, larva, pupa, adult. The first three stages are confined to running waters, which, depending on species, can range in size from tiny headwater trickles to large rivers. Eggs are deposited onto submerged or emergent substrata, or are simply dropped into the water, where they eventually settle into the bottom sediments. Larvae are roughly sausage-shaped creatures that bear a pair of rakelike appendages (labral fans) on the apex of their heads (Figure 31.2). When positioned into a current of water the fans intercept suspended particles as they pass by, providing the larva with food. Algae, bacteria, and even smaller particles, such as dissolved organic matter, are captured by the fans, which are alternatively folded and inserted into the mouth. Larvae typically pass through a series of seven molts. Mature larvae spin a silken cocoon that varies in form from a shapeless sac to an elaborate shoe- or bootshaped structure. The pupa more or less reflects the shape of the future adult, with the body regions and most of the appendages clearly visible through the pupal cuticle. A pair of spiracular gills arise from the
Feeding Behavior Many groups of black flies feed predominantly on birds and phylogenetic evidence suggests that ornithophily arose early in simuliid evolutionary history. It seems likely that simuliids first began blood feeding on the featherless progenitors of modern birds and that the specially modified tarsal claws of females arose shortly after the evolution of feathers. The subfamily Parasimuliinae is a relict group that currently consists only of autogenous (i.e., non-blood-feeding) species. In contrast, the subfamily Simuliinae has birdfeeding lineages in both the Prosimuliini and Simuliini. Phylogenetic evidence is equivocal about whether ornithophily is in the ground plan of the subfamily Simuliinae or whether the habit evolved twice independently in the two tribes (Prosimuliini and Simullini). In either case, it is clear that ornithophily had an early origin in black flies, and is the predominant feeding habit among currently recognized genera. The proportion of ornithophilic species among anautogenous black flies is difficult to assess because many regional faunas are inadequately studied. But in North America—where the simuliid fauna is most
Figure 31.2. Larva of Simulium vittatum. Note the sausage-shaped body and labral feeding fans on head. From Currie (1986).
taxonomic treatment of the world Simuliidae, other than the world inventory of Crosskey and Howard (2004). We follow the generic classification of these authors for the purposes of this chapter.
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Figure 31.3. Pupa of Simulium vittatum. Note the silk cocoon and spiracular gills arising from the anterolateral corners of the thorax. These latter structures are important for respiration in the developing fly. From Peterson (1981).
completely known—approximately 37% of anautogenous black flies are ornithophilic compared to 63% that are mammalophilic (Adler et al. 2004). The incidence of ornithophily increases with latitude, with a proportion of 30% between 30 and 40◦ north and 46% between 50 and 60◦ north. The relative proportion of ornithophilic species at a world scale is impossible to evaluate because there are so few reliable host records outside of North America and Europe. Even in North America, about 60% of blood feeding species lack even a single confirmed host record (Adler et al. 2004). The apparent higher incidence of ornithophily at northern latitudes perhaps reflects a higher incidence of exclusively ornithophilic lineages in the Northern Hemisphere. In contrast, there are relatively few exclusively ornithophilic lineages in South America, and there are proportionally far fewer records of hematophagy on avian hosts (cf. Coscar´on and Coscar´on-Arias 2007). Whether this truly reflects a lower incidence of ornithophily in South America remains to be seen. Such a trend seems counterintuitive, given the relatively higher diversity of birds in tropical regions. Along similar lines, there are relatively few exclusively ornithophilic lineages of black flies in the Afrotropical and Australasian regions, and there are correspondingly few accounts about hematophagy on wild birds. This neglected aspect of simuliid biology requires much greater attention. Bloodsucking (anautogenous) females locate their hosts using a combination of habitat features and host cues. Included among these latter are size, shape, color, odor (especially CO2 ), body temperature, and phagostimulants, such as adenosines (Simmons 1985; Sutcliffe 1986, 1987; Allan et al. 1987). Visual stimuli and carbon dioxide are most important at long range, whereas odor and heat perception are thought to play a role at close range. Host products such as sweat are known to be attractive to female black flies (Schofield 1994).
At least two blood meals are needed for transmission of parasitic diseases—one to acquire the causative agent and another to transmit it. Anautogenous species that produce multiple egg batches, therefore, include the most important vectors of disease agents. Protozoa (Leucocytozoon, Trypanosoma), filarial nematodes (e.g., Splendidofilaria fallisensis), arboviruses (e.g., vesicular stomatitis), and possibly bacteria are transmitted to a wide variety of avian and mammalian hosts (Crosskey 1990; Adler et al. 2004; Adler 2005). The only known simuliid-borne pathogen of humans—the filarial worm Onchocerca volvulus, the cause of river blindness—is restricted to Africa and South and Central America. In northern latitudes black fly activity is seasonal. Black flies emerge a few weeks after the ice leaves the streams and rivers in the spring and continues until the first frost in the fall. Peak black fly activity coincides with the nesting period for most northern bird species. Black flies are active during daylight hours in sheltered areas and particularly during cloudy periods. In open areas, they are most active in the early morning and evening shortly after sunset. Black fly activity increases at the approach of storms; however, rain and cold inhibit activity. Black flies cease flight activity when temperatures drop to 8–10◦ C and resume at 13– 15◦ C (Crosskey 1990). The intensity of feeding (i.e., the prevalence, of hematophagous female black flies) is dependent on season, nest site location (proximity to open running water), when egg laying is initiated, and local weather conditions, including wind. PATHOGENESIS Black flies have bladelike mandibles that are used to pierce the skin and introduce salivary secretions into the wound. These salivary secretions cause vasodilation and promote blood flow to the wound, have anticoagulant activity to facilitate feeding, and modulate
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Black Flies (Diptera: Simuliidae) components of the host immune system (Cupp and Cupp 1997). It takes between 3 and 6 min for females to fully gorge. Salivary secretions have been studied in a number of mammalophilic black fly species. Saliva from Simulium lineatum and Simulium equinum contains histamine and several bioamines including putrescine, spermine, N1-monoacetyl-spermine, and spermidine (Wirtz 1990). Salivary apyrase and antithrombin salivary protein inhibit platelet aggregation and target components of the common pathway of the mammalian coagulation cascade (Cupp and Cupp 1997). Salivary gland extracts from Simulium vittatum modulate host immune responses by altering patterns of cytokine response (Cross et al. 1994). Black fly bites result in an itchy raised bump or wheal, which may persist for up to 2 weeks, depending on the host. Repeated bites or large numbers of bites may lead to severe local or systemic allergic reactions in humans and other mammals.
Impact of Biting and Swarming on Birds Black flies can have a negative impact on birds even without the complication of disease transmission. Blood feeding can disrupt feeding and nesting behaviors, resulting in death in extreme cases (Adler et al. 2004). Whether mortality is the result of anemia or toxemia (or a combination of these two factors) is poorly understood. Nonetheless, the deleterious effects of blood feeding by black flies are most pronounced in young birds. The effects of bloodsucking on birds are best documented in the commercial poultry industry, where persistent attacks are known to result in inflammation of the skin, loss or reduction in appetite, egg abandonment, and death (Swenk and Mussehl 1928; Edgar 1953; Anderson and Voskuil 1963). Attacks on poultry are typically concentrated in the vicinity of the neck and head, especially around the eyes. Attacks have also been documented in the exotic bird industry in North America, with black flies implicated in the deaths of co*ckatoos and parrots (Mock and Adler 2002).
CLINICAL SIGNS AND PATHOLOGY Chickens and turkeys become restless and distracted when harassed by large numbers of black flies crawling on their skin, under the wings, and biting around the eyes or on combs and wattles. Hens abandon nests, food intake decreases, and egg productivity drops. Young chicks and turkey poults may die rapidly from blood loss if infestations are intense (Crosskey 1990). Clinical pathology has not been documented, but it
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is likely that survivors develop a regenerative anemia with marked erythrocyte polychromasia. Wild birds tormented by black flies also show behavioral changes. Nestling Red-tailed Hawks (Buteo jamaicensis) continuously exhibited annoyance behavior during infestations by continuously flapping their wings, vigorously shaking their heads, moving around in the nest, and pecking at exposed areas of their bodies (Smith et al. 1998). Occasional chicks found dead under nest sites had fractured bones but no other signs of disease, possibly the result of nestlings falling or jumping from the nests prematurely in an attempt to escape black fly harassment. Red-tailed Hawk nestlings in the Yukon, Canada, were restless and called incessantly (Doyle 2000). Similar behaviors were not observed in nestlings unaffected by black flies. Rohner et al. (2000) documented that roost site selection in Great Horned Owls (Bubo virginianus) in the Kluane region of the Yukon, Canada, shifted from the winter and late spring to the summer months, coinciding with the emergence of black flies and black fly harassment. During the summer the owls roosted near ground level or on the ground in open areas compared to the traditional concealed mid-canopy locations used during the rest of year. Black fly activity was near zero in open areas at ground level and highest at mid-canopy level. The authors suggest that changes in roost site selection may be influenced by black fly avoidance behavior. Black flies feed mostly on unfeathered areas such as the eyelids, cere, corners of the beak, ventral surface of the mandible, auricular openings, jugular groove, and ventral surfaces of the patagium (Figure 31.4) (Hunter et al. 1997; Smith et al. 1998). Eyelids may be edematous and swollen shut. Raised, inflamed erythematous areas surround small puncture wounds at black fly feeding sites and these areas are often scabbed over with dried blood (Figure 31.5). In heavy infestations adjacent feathers may be matted with blood. Microscopically, the skin lesions are characterized by increased vascularization, local edema, subcutaneous hemorrhage, local tissue necrosis, mixed inflammatory cell infiltration, and thrombosis of adjacent capillaries (Hunter et al. 1997; Smith et al. 1998). Allergic reactions, generalized toxemia, or simuliotoxicosis similar to that described in cattle and so-called black fly fever in people (headache, feverish shivering, nausea, swelling and tenderness of lymph glands, aching joints, and mental depression) have not been reported in birds. DIAGNOSIS Diagnosis is based on the presence of clinical signs or lesions typical of biting black flies and supported by finding the flies on or around the host. Black flies are
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Figure 31.4. Common Loon (Gavia immer) harassed by black flies, likely Simulium annulus (Lundstrom), ¨ the most host-specific of ornithophilic black flies. Note the flies feeding along the unfeathered margins of the bill and cere. Courtesy of N. K. Dawe, Canadian Wildlife Service.
notorious for their structural hom*ogeneity, and specieslevel identification often requires that specimens be examined by a specialist. It is not unusual for a single host to be attacked by more than one species of black fly.
Figure 31.5. Eyelid of a Great Horned Owl (Bubo virginianus) fledgling that has suffered extensive black fly bites to the eyelids and elsewhere. The eyelid is swollen and each dark crusted area is a black fly feeding site. Feathers near the margin of the eyelid are matted with dried blood. Courtesy of D. B. Hunter.
IMMUNITY There is no literature documenting immunity or resistance to black fly bites in birds. The increased susceptibility of nestlings compared to adults is due to their small size and physical and physiological immaturity as well as their inability to escape harassment. Mortality in both Eastern Bluebirds (Sialia sialis) and Redtailed Hawk nestlings was age related and decreased once protective feathering developed. In people, some individuals appear to have natural immunity and are bitten less frequently by black flies than are others. Biting rates have been shown to be partially dependent on interindividual variation in skin secretions and skin temperature (Schofield and Sutcliffe 1997) rather than host immunity. This phenomenon has not been studied in birds. PUBLIC HEALTH CONCERNS Mammalophilic and anthropophilic black flies cause many health concerns in people. These include being a major nuisance, causing localized dermatitis through bite wounds, inciting localized or systemic allergic reactions, and being the main vector for Onchocerca volvulus, the cause of human onchocerciasis or river blindness. Ornithophilic black flies pose no health hazards to humans. DOMESTICATED ANIMAL HEALTH CONCERNS Ornithophilic black flies have long been recognized as important pests for domestic poultry or other bird species that are raised outdoors in areas close to black fly habitats. The importance of black flies in commercial poultry production has decreased as poultry are less frequently raised in open range conditions, reducing exposure to black flies. Black flies still pose a threat to backyard poultry, organic and free range growing operations, pet birds housed in outdoor aviaries, and birds in zoos and private collections. WILDLIFE POPULATION IMPACTS In general, accounts about the effects of hematophagy and harassment are sparsely represented in the wild bird literature, perhaps because there is less impetus to investigate cause of death in wild (as opposed to domesticated) birds. The impact of black flies on wild birds is not well known; however, evidence suggests that it may be just as severe as that observed in the poultry and exotic bird industries. For example, attacks by members of the S. annulus species group are known to induce mortality in Red-tailed Hawk nestlings, either by causing anemia and dehydration or by driving
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Black Flies (Diptera: Simuliidae) individuals from nests (Fitch et al. 1946; Smith et al. 1998; Doyle 2000). In a 4-year study (1992–1995) of Red-tailed Hawks nesting in the Kluane region of the Yukon, Canada, Doyle (2000) monitored black fly numbers at nests every 2–3 days. In these 4 years, whenever >70 flies were recorded on or around the young during a visit, and chicks were less than 20 days of age, all chicks had died by the next nest visit. Over all 4 years, the annual proportion of nest failures was significantly correlated with the mean number of flies seen per nest visit. The time of nest initiation was a factor and late breeders had a higher rate of nest failure. Smith et al. (1998) documented black fly infestations (Simulium canonicolum) at 42 Red-tailed Hawk nests in Wyoming. Black flies caused mortality at 6 of 42 (14%) nests where young hatched (13 of 87 nestlings) and were the only known cause of nestling mortality. The onset of infestations occurred when nestlings were 3–20 days old and usually lasted until nestlings died or fledged. Brown and Amadon (1968) reported that during wet years black fly feeding (Prosimulium sp.) was a significant cause of Red-tailed Hawk nestling mortality in California. Females of Simulium meridionale are known to cause nestling mortality in Purple Martins (Progne subis), Mangrove Swallows (Tachycineta albilinea), and Eastern Bluebirds (Hill 1994; Gaard 2001, 2002, 2003). Affected birds are generally 6–10 days of age and feathering is incomplete. Finally, attacks by females of Cnephia ornithophilia may disrupt the feeding and reproductive behaviors of the critically endangered Attwater’s Prairie-Chicken (Tympanuchus cupido attwateri), a subspecies of the Greater PrairieChicken (Adler et al. 2007). The vulnerability of birds, whether through the effects of blood feeding or leucocytozoonosis, is probably heightened during episodes of meager food supply (Hunter et al. 1997) and severe weather (Smith et al. 1998). The impact of black flies on nestlings and reproductive success could be easily overlooked or missed if nests were not visited at critical times. Baby birds are impacted at an early natal down stage. Standard nesting surveys with two to three nest visits per nesting period could easily miss the impacts of black flies on nesting success. Harassment by black flies can be an important mortality factor in man-made nesting cavities for Eastern Bluebirds and Mangrove Swallows. These problems are generally local and related to the location of the nest boxes near running streams and influenced by weather conditions. Mortality is noticed because of the frequency of observation, but there is little information on black fly related mortality in natural nesting areas that are not monitored as rigorously. Ornithophilic species of black flies can reduce the “fitness” of an avian host in a variety of ways: through
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direct blood loss and harassment during the nestling period; by the increased investment of energy in inflammation and repair and immunological defense; and by transmitting potential pathogens such as Leucocytozoon or Trypanosoma. Each of these factors has an energetic cost for the bird. The combination of black fly feeding and transmission of blood parasites may tip the scale toward mortality in times of stress, decreased food availability as demonstrated in Great Horned Owls in the Yukon (Rohner and Hunter 1996; Hunter et al. 1997), or perhaps during periods of inclement weather. The importance of black flies in reducing “fitness” and influencing other physiological processes deserves further study. It is also interesting that harassment by black flies and the subsequent avoidance behavior by the host may result in other subtle effects and trade-offs. Great Horned Owls may put themselves at greater risk of predation by choosing roosting locations on the open ground, presumably in an effort to avoid black fly harassment. Perhaps nest site selection is influenced by a similar process. The role of black flies in the natural history of birds remains poorly understood.
TREATMENT, CONTROL, AND MANAGEMENT IMPLICATIONS Many strategies have been developed for black fly control because of their importance as vectors of Onchocerca volvulus in West Africa. The World Health Organization Onchocerciasis Control Program (OCP), established in 1974, was based on the spray of insecticides by helicopters and aircraft over larval habitats (aerial larviciding). With the donation of Mectizan® (ivermectin) by Merck & Co., Inc., in 1987, control operations changed from exclusive use of vector control to a combination of vector control and treatment with ivermectin. In some areas, ivermectin alone was used to control infections (see World Health Organization Web site www.who.int/blindness/ partnerships/onchocerciasis OCP/en/index.html). The OCP was officially closed in December 2002 and considered to have been a highly successful program. Currently, most control programs against black flies are directed toward nuisance pests of humans. It is unlikely that large-scale black fly control programs would be used or warranted in protecting wild birds. However, there may be circ*mstances where it should be considered, particularly when endangered species are at risk. Adler et al. (2007) suggested that if management of black flies became necessary to protect the endangered Attwater’s Prairie-Chicken on the Attwater Prairie Chicken National Wildlife Refuge, Texas, USA, then control would include a thorough
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evaluation of local streams and irrigation systems to first identify black fly breeding sites that could then be treated with an environmentally safe, simuliid-specific biopesticide such as Bacillus thuringiensis israelensis (Gray et al. 1996). Measures to control access of black flies to manmade nest boxes for Eastern Bluebirds and Mangrove Swallows include blocking ventilation openings with cotton and duct tape and creating a darker interior to the nesting cavity to reduce black fly activity (Gaard 2002). Pesticides (pyrethrins) have been used within nesting cavities to reduce black fly numbers. These efforts are targeted toward nestlings that are around 6– 10 days of age when they are extremely vulnerable to feeding by black flies. Vector proofing aviaries in captive or breeding collections using fine mesh netting is another possible prevention method.
LITERATURE CITED Adler, P. H. 2005. Black flies, the Simuliidae. In Biology of Disease Vectors, 2nd ed, W. C. Marquardt (ed.). Elsevier Academic Press, San Diego, CA, pp. 127–140. Adler, P. H., D. C. Currie, and D. M. Wood. 2004. The Black Flies (Simuliidae) of North America. Cornell University Press, Ithaca, NY, 941 pp. Adler, P. H., D. Roach, W. K. Reeves, J. P. Flanagan, M. E. Morrow, and J. E. Toepfer. 2007. Attacks on the endangered Attwater’s Prairie-Chicken (Tympanuchus cupido attwateri) by black flies (Diptera: Simuliidae) infected with an avian blood parasite. Journal of Vector Ecology 32:309–312. Allan, S. A., J. F. Day, and J. D. Edman. 1987. Visual ecology of biting flies. Annual Review of Entomology 32:297–316. Anderson, J. R., and G. H. Voskuil. 1963. A reduction in milk production caused by the feeding of blackflies (Diptera: Simuliidae) on dairy cattle in California, with notes on the feeding activity on other animals. Mosquito News 23:126–131. Bennett, G. 1960. On some ornithophilic blood-sucking Diptera in Algonquin Park, Ontario, Canada. Canadian Journal of Zoology 38:377–389. Bennett, G. 1963. Use of P32 in the study of a population of Simulium rugglesi (Diptera: Simuliidae) in Algonquin Park, Ontario. Canadian Journal of Zoology 41:831–840. Brown, L. H., and D. Amadon. 1968. Eagles, Hawks and Falcons of the World, Country Life Books, London, England. Coscar´on, S., and C. L. Coscar´on-Arias. 2007. Neotropical Simuliidae (Diptera: Insecta). In Aquatic
Biodiversity of Latin America (ABLA Series), Vol. 3, J. Adis, J. R. Arias, G. Rueda-Delgado, and K. M. Wantzen (eds). Pensoft Publishers, Sofia-Moscow, Bulgaria, 685 pp. Cross, M. L., E. W. Cupp, and F. J. Enriquez. 1994. Modulation of murine cellular immune responses and cytokines by salivary gland extract of the black fly Simulium vittatum. Tropical Medical Parasitology 45:119–124. Crosskey, R. W. 1990. The Natural History of Blackflies. John Wiley and Sons, Chichester, UK, 711 pp. Crosskey, R. W., and T. M. Howard. 2004. A Revised Taxonomic and Geographical Inventory of World Blackflies (Diptera: Simuliidae). The Natural History Museum, London. Available http://www.nhm.ac.uk/ research-curation/projects/blackflies/. Accessed September 28, 2006. Cupp, E. W., and M. S. Cupp. 1997. Black fly (Diptera: Simuliidae) salivary secretion: importance in vector competence and disease. Journal of Medical Entomology 34:87–94. Currie, D. C. 1986. An annotated list of and keys to the immature black flies of Alberta (Diptera: Simuliidae). Memoirs of the Entomological Society of Canada 134:1–90. Currie, D. C., and P. H. Adler. 2008. Global diversity of black flies (Diptera: Simuliidae) in freshwater. Hydrobiologia 595:469–475. Currie, D. C., and D. Grimaldi. 2000. A new black fly (Diptera: Simuliidae) genus from mid Cretaceous (Turonian) amber of New Jersey. In Studies on Fossils in Amber, with Particular Reference to the Cretaceous of New Jersey, D. Grimaldi (ed.). Backhuys Publishers, Leiden, The Netherlands, pp. 473–485. Doyle, F. I. 2000. Timing of Reproduction by Red-tailed Hawks, Northern Goshawks and Great Horned Owls in the Kluane Boreal Forest of Southwestern Yukon. M.Sc. thesis, University of British Columbia, Vancouver, Canada, 141 pp. Edgar, S. A. 1953. A field study of the effect of black fly bites on egg production of laying hens. Poultry Science 32:779–780. Fallis, A. M., and S. M. Smith. 1964. Ether extracts from birds and CO2 as attractants for some ornithophilic simuliids. Canadian Journal of Zoology 42:723–730. Fitch, H. S., F. Swenson, and D. F. Tillotson. 1946. Behavior and food habits of the red-tailed hawk. The Condor 48:205–237. Gaard, G. 2001. June nest box mortality. Wisconsin Bluebird 16(2):1, 4–5. Gaard, G. 2002. Black fly induced nest mortality—Prevention possibilities. Wisconsin Bluebird 17(1):13–16. Gaard, G. 2003. Eliminating black fly feeding on nestling bluebirds. Wisconsin Bluebird 18(1):1–4.
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Black Flies (Diptera: Simuliidae) Gray, E. W., P. H. Adler, and R. Nolet. 1996. Economic impact of black flies (Diptera: Simuliidae) in South Carolina and development of a localized suppression program. Journal of the Mosquito Control Association 12:676–678. Hill, J. R., III. 1994. What’s bugging your birds? An introduction to the ectoparasites of purple martins. Purple Martin Update 5(1):1–7. Hunter, D. B., C. Rohner, and D. C. Currie. 1997. Mortality in fledgling Great Horned Owls from black fly hematophagy and leucocytozoonosis. Journal of Wildlife Diseases 33:614–618. Hunter, F. F., J. F. Sutcliffe, and A. E. R. Downe. 1993. Blood-feeding host preferences of the isomorphic species Simulium venustum and S. truncatum. Medical and Veterinary Entomology 7:105–110. Kalugina, N. S. 1991. New Mesozoic Simuliidae and Leptoconopidae and blood-sucking origin in the lower dipterans (in Russian with English summary). Paleontologischesky Zhurnal 1991:69–80. Lowther, J. K., and D. M. Wood. 1964. Specificity of a black fly, Simulium euryadminiculum Davies, toward its host, the common loon. Canadian Entomologist 96:911–913. Malmqvist, B., D. Strasevicius, O. Hellgren, P. H. Adler, and S. Bensch. 2004. Vertebrate host specificity of wild-caught blackflies revealed by mitochondrial DNA in blood. Proceedings of the Royal Society of London. Series B. Biology Letters 271:S152–S155. Mock, D. E., and P. H. Adler. 2002. Black flies (Diptera: Simuliidae) of Kansas: Review, new records, and pest status. Journal of the Kansas Entomological Society 75:203–213. Moulton, J. K. 2000. Molecular sequence data resolves basal divergences within Simuliidae (Diptera). Systematic Entomology 25:95–113. O’Roke, E. C. 1930. The incidence, pathogenicity and transmission of Leucocytozoon anatis of ducks. Journal of Parasitology 17:112. Peterson, B. V. 1981. Simuliidae. In Manual of Nearctic Diptera, Vol. 1, Monograph No. 27, J. F. McAlpine, B. V. Peterson, G. E. Shewell, H. J. Teskey, J. R. Vockeroth, and D. M. Wood (eds). Research Branch, Agriculture Canada, Ottawa, Canada, 674 pp.
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Rohner, C., and D. B. Hunter. 1996. First-year survival of Great Horned Owls during a peak and decline of the snowshoe hare cycle. Canadian Journal of Zoology 74:1092–1097. Rohner, C., C. J. Krebbs, D. B. Hunter, and D. C. Currie. 2000. Roost site selection of Great Hormned Owls in relation to black fly activity: An anti-parasite behavior? The Condor 102:950–955. Schofield, S., and J. F. Sutcliffe. 1997. Humans vary in their ability to elicit biting responses from Simulium venustum (Diptera: Simuliidae). Journal of Medical Entomology 34:64–67. Schofield, S. W. 1994. The Bases of Human Attractiveness and Bitability for Black Flies (Diptera: Simuliidae) in Algonquin Park, Ontario. M.S. thesis, Trent University, Peterborough, Ontario, Canada, 133 pp. Shewell, G. E. 1955. Identity of the black fly that attacks ducklings and goslings in Canada (Diptera: Simuliidae). Canadian Entomologist 87:347–349. Simmons, K. R. 1985. Reproductive Ecology and Host-Seeking Behavior of the Black Fly, Simulium venustum Say (Diptera: Simuliidae). Ph.D. thesis, University of Massachusetts, Amherst, MA, 204 pp. Skidmore, L. V. 1931. Leucocytozoon smithi infection in turkeys and its transmission by Simulium occidentale Townsend. Journal of Parasitology 18:130. Smith, R. N., S. L. Cain, S. H. Anderson, J. R. Dunk, and E. S. Williams. 1998. Blackfly-induced mortality of nestling red-tailed hawks. The Auk 115: 368–375. Sutcliffe, J. F. 1986. Black fly host location: A review. Canadian Journal of Zoology 64:1041–1053. Sutcliffe, J. F. 1987. Distance orientation of biting flies to their hosts. Insect Science and Its Application 8:611–616. Swenk, M. H., and R. E. Mussehl. 1928. The insects and mites injurious to poultry in Nebraska and their control. University of Nebraska Agricultural Experiment Station Circular 37:1–31. Wirtz, H. P. 1990. Bioamines and proteins in the saliva and salivary glands of Palaearctic blackflies (Diptera: Simuliidae). Tropical Medical Parasitology 41:59–64.
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32 Myiasis in Wild Birds Susan E. Little not associated with any actual infestation (Catts and Mullen 2002). Myiasis may also be described according to the tissue ingested or the body region affected (Zumpt 1965). The most common forms in wild birds are hematophagous myiasis, which occurs when larvae of obligatory myiatic species ingest blood of the infested host, and subcutaneous myiasis, in which the larvae burrow beneath the skin to feed on host tissues. The distinction is not absolute; the larvae of some hematophagous species remain on the surface of birds, while larvae of other species continue to burrow deeply into the host while still feeding on blood (Sabrosky et al. 1989). Traumatic myiasis is the secondary invasion of wounds by dipteran larvae and has occasionally been reported from wild birds (Zumpt 1965; Baumgartner 1988). Finally, larvae involved in myiasis may be referred to as primary, secondary, or tertiary depending on when they enter the host or the wound. Species that cause primary myiasis are able to initiate feeding on an otherwise healthy animal with intact skin. Species that cause secondary myiasis are facultative parasites that infest preexisting wounds, feeding on living tissue only once the supply of decaying, necrotic tissue on that animal has been depleted. Species that cause tertiary myiasis prefer carcasses and are only rarely seen on living animals very late in the course of disease (Kettle 1995).
INTRODUCTION Myiasis is the infestation of healthy or necrotic tissue of living vertebrate animals by dipteran larvae and is very common in birds, particularly in nestlings. The larvae of some species of diptera burrow into tissues, creating wounds, while larvae of other species feed on blood but remain on the surface of their avian hosts. The detrimental effects of fly larvae feeding on nestlings are not always readily apparent, and in many cases, may only become clinically evident when compounded by inadequate nutrition or other factors that cause stress. Myiasis is reported across terrestrial vertebrate taxa and may be associated with virtually any dipteran larvae, although cases are usually associated with infestation by blow flies (Calliphoridae), flesh flies (Sarcophagidae), or bot flies (Oestridae) (Catts and Mullen 2002). Myiasis in wild birds is most frequently associated with infestations of hematophagous dipteran larvae from the families Calliphoridae, Muscidae, and Neottiophilidae (Uhazy and Arendt 1986; Spalding et al. 2002). Cutaneous and subcutaneous infestation of wild birds with dipteran larvae of other families also occurs (Zumpt 1965; Wobeser et al. 1981; Baumgartner 1988; Farkas et al. 2001). Infestation of birds by oestrid bot flies is rare (Artmann 1975; Roberts and Janovy 2005) and will only briefly be mentioned here. Although any warm-blooded animal is susceptible to infestation with larvae of Cochliomyia hominivorax and Chrysomya bezziana, the causes of obligatory myiasis, screwworm infestations are rarely reported on birds (Roberts and Janovy 2005). A wealth of terminology is used to characterize the different types of myiases. Larvae may be referred to as obligatory if they require a living host as part of their life cycle, or facultative if they normally develop in decaying organic matter and only occasionally infest necrotic wounds on living animals. A third form, accidental or pseudomyiasis, describes cases where dipteran larvae are mistakenly ingested by an animal and pass through the gastrointestinal tract, but are
SYNONYMS Synonyms vary according to the type of myiasis being described. Hematophagous species typified by the Protocalliphora have been referred to as bird nest flies, bird blow flies, bloodsucking larval flies, parasitic bird flies, nestling “screwworms,” and bird nest “screwworms” (Sabrosky et al. 1989). The hematophagous larvae of muscid flies, such as Philornis spp., are occasionally referred to as bots, although most authors reserve this term for individual oestrid larvae. Oestrid (=botfly) larvae and their cysts may also be called warbles or
546 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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Myiasis in Wild Birds wolves, but true bot fly infestations are rare on avian hosts. Disease caused by species involved in traumatic or secondary myiasis may be called strike, fly strike, infestation with wound maggots, or blow, and affected animals are sometimes referred to as “blown” or “fly struck” (Catts and Mullen 2002). HISTORY Reports of dipteran larvae feeding on Barn Swallow (Hirundo rustica) nestlings can be found as early as 1845 in what is regarded as the first description of Protocalliphora azurea from France, and in another report of a likely Protocalliphora sp. described in 1866 from North America (Dufour 1845 and Walsh 1866 as reported by Sabrosky et al. 1989). Numerous subsequent descriptions of hematophagous and occasionally subcutaneous maggots that parasitize nestling birds have been made, most involving species of the genus Protocalliphora, Philornis, or Passeromyia (Table 32.1). The literature also contains sporadic reports of traumatic myiasis on birds, although this form of myiasis is considered relatively uncommon in wild birds (Wobeser et al. 1981; Baumgartner 1988; Farkas et al. 2001). Despite the frequent descriptions of affected birds in the literature, myiasis has historically been considered of little consequence to otherwise healthy bird populations. Indeed, in a comprehensive description of North American Protocalliphora spp., Sabrosky et al. (1989) notes that most entomology and avian disease books do not mention or emphasize myiasis-inducing diptera on birds. However, the recent discovery that Philornis downsi is parasitizing nestlings of Darwin’s finches in the Galapagos Islands has reignited interest in the importance of myiasis to the health of wild birds, particularly threatened species (Fessl et al. 2006a, b). Similarly intriguing is the recent finding that myiasis in nestlings increases in disturbed habitats (Gentes et al. 2007). Additional work will likely continue to reveal that in times of limited nutrients, habitat stress, and introduction of new fly species into fragile ecosystems, myiasis can have a significant impact on wild birds. DISTRIBUTION Avian myiasis is cosmopolitan in distribution, although the individual species involved may be restricted in their range, and localities often have a predominance of just a handful of important myiasisinducing diptera (Sabrosky et al. 1989). Flies that produce hematophagous larvae which are obligate feeders on nestling birds include members of the families Calliphoridae (Protocalliphora spp.), Muscidae (Philornis, Passeromyia, Mydaea), and Neottiophili-
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dae (Neottiophilum, Actinoptera) (Uhazy and Arendt 1986; Spalding et al. 2002). The Protocalliphora spp. are found in a Holarctic, predominantly northern distribution, extending in North America as far south as Nearctic Mexico, across Palearctic Europe, North Africa, and temperate Asia. Reports in the southern extent of this range are generally from higher altitudes (Sabrosky et al. 1989). These flies appear to be largely absent from the southeastern US (Sabrosky et al. 1989; Spalding et al. 2002). Excellent maps detailing the known range of Nearctic Protocalliphora spp. can be found in Sabrosky et al. (1989). The muscid genera that comprise the tropical nest flies also have a broad distribution but are usually found in warmer climates than Protocalliphora spp.; Philornis is reported from the New World tropics and Passeromyia and Mydaea occur in Australia and Asia (Pont 1974; Couri 1999). Nest skipper flies (Neottiphilum spp.) have a predominantly Palearctic distribution (Catts and Mullen 2002). Other calliphorids that have been reported associated with wounds on wild birds include Calliphora and Lucilia spp. in Europe (Baumgartner 1988; Farkas et al. 2001). Reports of sarcophagid flesh flies associated with wound myiasis in birds are mostly confined to Wohlfahrtia magnifica in the Old World and Wohlfahrtia opaca in the New World (Zumpt 1965; Wobeser et al. 1981). HOST RANGE Nestlings of any nidocolous (=nest dwelling) bird may be subject to hematophagous myiasis. However, reports of hematophagous myiasis caused by Protocalliphora, Philornis, Passeromyia, and Neottiophilum are by far most common in the passerines (Table 32.1). Myiasis caused by species of Protocalliphora is also described from raptors and, occasionally, Piciformes (woodpeckers and allied species). Sporadic reports are found on Columbiformes and Cuculiformes. In a review of known hosts of Protocalliphora spp. in North America, 139 different species of birds were reported as potential hosts of Protocalliphora spp., but there were no documented reports of natural infestations on water or shore birds, Galliformes, Psittiformes, Caprimulgiformes, Apodiformes, Trogoniformes, or Coraciiformes. Traumatic myiasis can be reported from any vertebrate, including any species of bird. Confusion regarding antemortem and postmortem arrival of the fly larvae found on a bird carcass, and the presence of maggots feeding on decaying organic matter in the nests but not the nestlings themselves, can lead to misinterpretation of the involvement of myiasis in a mortality event. Although obligatory and facultative traumatic myiasis has been frequently reported from domestic
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Table 32.1. Examples of hematophagous dipteran larvae found on wild birds. Family Calliphoridae
Species Protocalliphora aenea Protocalliphora asiovora Protocalliphora avium Protocalliphora azurea Protocalliphora beameri Protocalliphora bennetti Protocalliphora bicolor Protocalliphora braueri Protocalliphora brunneisquama Protocalliphora chrysorrhoea Protocalliphora cuprina Protocalliphora deceptor Protocalliphora falcozi Protocalliphora fallisi Protocalliphora halli Protocalliphora hesperia Protocalliphora hesperioides Protocalliphora hirundo Protocalliphora interrupta Protocalliphora isochroa Protocalliphora lata Protocalliphora lindneri Protocalliphora metallica Protocalliphora occidentalis Protocalliphora parorum Protocalliphora peusi Protocalliphora rugosa Protocalliphora seminuda Protocalliphora shannoni Protocalliphora sialia
Muscidae
Protocalliphora spatulata Protocalliphora spenceri Protocalliphora tundrae Passeromyia heterochaeta Passeromyia indecora Philornis carinatus
Host order Passeriformes Falconiformes Passeriformes Falconiformes Passeriformes Strigiformes Coraciiformes Passeriformes Passeriformes Passeriformes Passeriformes Falconiformes Passeriformes Passeriformes Passeriformes Columbiformes Passeriformes Passeriformes Passeriformes Passeriformes Piciformes Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes Falconiformes Passeriformes Piciformes Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes Passeriformes Falconiformes Passeriformes Piciformes Strigiformes Passeriformes Passeriformes Passeriformes Falconiformes Passeriformes Passeriformes Passeriformes
Example reference Halstead (1988) Sabrosky et al. (1989) Whitworth and Bennett (1992) Bortolotti (1985) Pletsch (1948) Tirrell (1978) Sabrosky et al. (1989) Simon et al. (2004) Sabrosky et al. (1989) Whitworth (2003) Sabrosky et al. (1989) Sabrosky et al. (1989) Howe (1992) Whitworth (2003) Whitworth and Bennett (1992) Sabrosky et al. (1989) Boland et al. (1989) Revels (1996) Simon et al. (2004) Sabrosky et al. (1989) Sabrosky et al. (1989) Sabrosky et al. (1989) Sabrosky et al. (1989) Sabrosky et al. (1989) Shields and Crook (1987) Sabrosky et al. (1989) Sabrosky et al. (1989) Sabrosky et al. (1989) Sabrosky et al. (1989) Wiebe and Swift (2001) Sabrosky et al. (1989) Revels (1996) Whitworth (2003) Johnson and Albrecht (1993) Sabrosky et al. (1989) Whitworth (2003) Sabrosky et al. (1989) Dawson et al. (1999) Sargent (1938) Gentes et al. (2007) Sabrosky et al. (1989) Proudfoot et al. (2005) Miller and Fair (1997) Matsuoka et al. (1997) Sabrosky et al. (1989) Gargett (1975) Lindholm (1998) Pont (1974) Young (1993) (continues)
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Myiasis in Wild Birds Table 32.1. (Continued ) Family
Species Philornis deceptivus Philornis downsi Philornis mimicola Philornis pici
Neottiophilidae
Philornis porteri Philornis seguyi Philornis sp. Neottiophilum praeustum
Host order
Example reference
Passeriformes Passeriformes Strigiformes Passeriformes Psittaciformes Passeriformes Passeriformes Falconiformes Falconiformes Passeriformes
Uhazy and Arendt (1986) Fessl et al. (2006a) Proudfoot et al. (2006) Nores (1995) Snyder et al. (1987) Spalding et al. (2002) Couri et al. (2005) Hector (1982) Zumpt (1965) Zumpt (1965)
Note: There are several hundred reports associated with species of wild birds; only a single recent report of each fly species/bird order association is provided in this table of representative examples. Individuals interested in detailed reports from individual species are urged to consult the primary literature.
and wild birds in Europe, reports in Nearctic wild birds are largely lacking (Baumgartner 1988), leading some to propose that the disease may be overlooked due to the relatively short time frame in which there is an opportunity to make a diagnosis prior to decomposition of the carcass (Wobeser 1997). ETIOLOGY Hematophagous myiasis of nestling birds is caused by members of three families of dipteran flies: Calliphoridae, Muscidae, and Neottiphilidae (Uhazy and Arendt 1986; Spalding et al. 2002). Examples of individual genera and species reported as parasites of different avian hosts are summarized in Table 32.1. Protocalliphora is the most common genus reported; approximately 90 species have been described in this genus worldwide (Catts and Mullen 2002). Traumatic myiasis in wild birds is caused by flies in the families Calliphoridae and Sarcophagidae (Table 32.2). The human skin bot, Dermatobia hominis, may use wild birds as a host for larval development (Roberts and Janovy 2005). Other cases of oestrid bot fly infestation in wild birds are rare, but have been reported (Artmann 1975). EPIZOOTIOLOGY The complete life cycle has not been fully delineated for any species of diptera with obligate hematophagous larval stages. However, a general pattern for infestation and development of the members of the calliphorid and muscid genera is evident based on what is known (Sabrosky et al. 1989; Spalding et al. 2002; Fessl et al. 2006a). Nestlings are exposed when adult flies oviposit
into nests containing newly hatched birds or directly onto the recently hatched nestlings early in the spring. The mechanisms by which adult female flies locate nests containing newly hatched birds in order to deposit their eggs are unknown (Catts and Mullen 2002). The fly eggs develop rapidly to active first-instar larvae within 1–2 days and then migrate to the nestlings. The larvae of most species feed on the nestlings intermittently for a few hours at a time over a period of 1–2 weeks, moving from the nest material to the birds to feed. Some species (e.g., Protocalliphora braueri, Philornis nielseni) burrow beneath the skin for several days, breathing through a respiratory pore in the skin through which they later exit (Sabrosky et al. 1989). Other species (e.g., Philornis downsi) develop through the first and second instar in the nasal cavities and complete development in the nest materials, intermittently moving to the birds to feed and then returning to the nest material (Fessl et al. 2006a). After several blood meals, the third-instar larvae leave the bird to pupate in the nest material. The prepuparial stage is approximately 1–4 days long, while pupation itself may take 36 days or more to complete. Adult flies then emerge to mate and seek new nestlings on which to oviposit. The eggs and pupa of Protocalliphora spp. are not considered cold tolerant, and overwintering of those species in the temperate areas where they occur is thought to be achieved by survival of adults that mate the following spring (Sabrosky et al. 1989). The prevalence of bird nest flies is high among nidicolous birds. In a comprehensive study of Protocalliphora spp. on wild birds in Utah, 48% of the nests of 51 species contained Protocalliphora spp. (Whitworth and Bennett 1992). For some host species and in some locations and years, almost all nests examined
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Table 32.2. Examples of nonhematophagous dipteran larvae found on wild birds. Family Calliphoridae (bottle flies)
Species Calliphora sp. Lucilia sericata
Sarcophagidae (flesh flies)
Wohlfahrtia vigil Wohlfahrtia opaca
Cuterebridae (bot flies)
Wohlfahrtia magnifica Cuterebra buccata Dermatobia hominis
Host
Scientific name
Reference
Peregrine Falcon American Kestrel Common Crane Kestrel White Stork Harrier European Honey-buzzard Eurasian Eagle-Owl Tawny Owl Short-eared Owl American Kestrel Peregrine Falcon Greylag Goose American Robin
Falco peregrinus Falco sparverius Grus grus Not specified Ciconia ciconia not specified Pernis apivorus
Cooper (1978) Cooper (1978) Itamies and Merila (1984) Hinaidy and Frey (1984) Hinaidy and Frey (1984) Hinaidy and Frey (1982) Hinaidy and Frey (1982)
Bubo bubo Strix aluco Asio flammeus Falco sparverius Falco peregrinus Anser anser Turdus migratorius
Blue-winged Teal Northern Shoveler Greylag Goose
Anas discors Anas clypeata Anser anser
Hinaidy and Frey (1982) Hinaidy and Frey (1982) Frey and Hinaidy (1978) Cooper (1978) Cooper (1978) Farkas et al. (2001) Eschele and Defoliart (1965) Wobeser et al. (1981) Wobeser et al. (1981) Farkas et al. (2001)
American Woodco*ck Scolopax minor Macaw, other wild Various birds
are infested (Arendt 1985b; Hurtrez-Bousses et al. 1997a; Wesolowski 2001), leading some to conclude that virtually every bird species in North America with dry nests and altricial (=requiring care at birth) young is susceptible to infestation with Protocalliphora spp. (Sabrosky et al. 1989; Bennett and Whitworth 1991). Indeed, while some species of Protocalliphora are host specific, others appear to exhibit habitat preferences, being found in nests in a given geographic region, rather than consistently associated with a single host species (Sabrosky et al. 1989). Intensity of infestations with Protocalliphora spp. are extremely varied, with per nest numbers ranging from 1 to more than 1,000 (Seguy 1955; Whitworth and Bennett 1992). Large numbers of larvae seem to be associated with larger, sturdier nests, presumably because more adequate habitat for development in nest material exists (Sabrosky et al. 1989). Other calliphorids occasionally associated with wound myiasis in wild birds (e.g. Calliphora, Lucilia) normally deposit eggs onto decaying carcasses. The larvae then hatch and develop as saprophytes, feeding on decomposing tissue. However, these flies have also been reported as facultative parasites capable of invading necrotic tissue in wounds on living birds (Baumgartner 1988; Farkas et al. 2001).
Artmann (1975) Roberts and Janovy (2005)
Unlike the calliphorids, female sarcophagid flesh flies associated with cases of obligatory or facultative traumatic myiasis retain the developing eggs until they hatch. Depending on species, each female fly may deposit 30–200 larvae directly onto the animal (Catts and Mullen 2002). Female flesh flies in the genus Wohlfahrtia larviposit directly onto mucous membranes or fresh wounds; first-instar larvae of this genus are also able to penetrate intact skin and create a wound (Gassner and James 1948). After feeding on the host tissues and developing for approximately 1 week, the larvae drop to the ground to pupate and continue their development to adult flies (Gassner and James 1948). CLINICAL SIGNS Many infested birds do not exhibit any overt clinical disease associated with blood feeding by the larvae. However, anemia has been reported, particularly when infestations are severe (Dudaniec and Kleindorfer 2006; Fessl et al. 2006a). The blood-feeding larvae of some bird nest flies, including P. braueri and some Philornis spp., do not remain on the surface of the birds, but rather burrow beneath the skin to invade the subcutaneous space. Such infestations may result
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Myiasis in Wild Birds in grossly evident subcutaneous cysts containing one or more larvae. Protocalliphora avium can cause very distinct clinical signs in raptors. This species has a strong predilection for the aural cavity in species of Falconiformes, and developing larvae of this species can often be found completely occluding the opening to the ear in affected nestlings (Sargent 1938; Tirrell 1978; Bortolotti 1985). Larvae of P. avium and P. downsi may also be found in the nasal cavity (Hill and Work 1947; Fessl et al. 2006a). An early sign of infestation is the accumulation of dried, reddish-brown excreta from developing, blood-feeding larvae around the opening to infested aural and nasal cavities (Bent 1937; Tirrell 1978). PATHOGENESIS All species of diptera with hematophagous larvae are obligatory bloodsucking parasites in the immature form. In order to obtain a blood meal, the larvae of these bird nest flies migrate to the surface of the host and then use hooks on the oral cavity to penetrate the skin. A prothoracic fringe is used to hold the maggot in place and advance the anterior portion more deeply into the skin as the hooks continue to tear at the tissue. Once firmly embedded, the larva begins to feed. Feeding is a relatively slow process, and each larva requires a few hours to ingest an entire blood meal (Sabrosky et al. 1989). The pathogenic effects of infestations with hematophagous dipteran larvae are difficult to summarize succinctly. Otherwise healthy nestlings often seem to tolerate relatively low numbers of feeding larvae well (Gold and Dahlsten 1983; Eastman et al. 1989). However, when the number of larvae is high, particularly in malnourished nestlings or those with concurrent disease, morbidity and mortality may result (Howe 1992; Whitworth and Bennett 1992; Fessl et al. 2006a). In addition, even subclinical infestations appear to prolong nestling development which itself may increase the risk of loss to predation and thus decrease overall nestling success (Arendt 1985a; Hurtrez-Bousses et al. 1997b). Losses are thought to be particularly likely when severe infestations occur concurrent with inclement weather or nutrient shortages (Sabrosky et al. 1989; Whitworth and Bennett 1992). Occasionally, some larvae that burrow subcutaneously will penetrate through the thoracic, abdominal, or ocular cavity, causing severe damage as they migrate through the tissues. This damage often results in sepsis and ultimately death of affected birds (Arendt 1985a, b; Uhazy and Arendt 1986). Migrating larvae that invade and damage vital organs (e.g., lung, brain, liver) can also directly cause death of affected birds (Spalding et al. 2002).
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Despite the impressive number of larvae found in infestations of species which penetrate the aural or nasal cavities, permanent damage does not apparently occur in most cases (Tirrell 1978). Disfigured aural cavities, damaged tympana, and enlarged nostrils have been reported, however (Bent 1937; Fessl et al. 2006a). Disease caused by bird nest flies is directly related to blood loss from the feeding larvae and the subsequent iron deficiency anemia that develops. Gold and Dahlsten (1983) calculated that in a heavily infested nest, feeding larvae can consume more than 55% of a chick’s blood volume. This chronic blood loss early in life may result in prolonged fledging times and a subsequent increase in the likelihood of predation or disease loss due to other causes (Hurtrez-Bousses et al. 1997a, b). In some cases, parents of nestlings heavily parasitized by Protocalliphora spp. are able to somewhat offset the adverse effects of blood loss by nutritional supplementation of the nestlings (Hurtrez-Bousses et al. 1998; Simon et al. 2004), but the energy cost required to increase the feeding has been shown to reduce the subsequent fitness of those parents (Wesolowski 2001). Among species of diptera with hematophagous larvae that also invade subcutaneously, the larvae continue to advance through the skin until they reach the subcutaneous space. Here, they continue to ingest blood and other host tissue. Detrimental host effects may arise from the location, aggressiveness, or sheer number of the migrating larvae (Arendt 1985a, b; Spalding et al. 2002). PATHOLOGY Blood feeding by larvae would be expected to reduce the hematocrit of affected birds. However, perhaps due to rapid production of red blood cells in response to blood loss in the young birds, nestlings with heavy infestations of bird nest flies may have normal or only slightly decreased packed cell volumes (Johnson and Albrecht 1993; O’Brien et al. 2001). Hemoglobin concentrations may be reduced in the face of normal hematocrit levels, which could result in compromised oxygen transport to tissues and subsequent compromised development and survival (O’Brien et al. 2001). The pathogenic effects of other diptera with hematophagous larvae are more clearly established, particularly for those that migrate through body tissues. For example, parasitism by Philornis spp. is responsible for mortalites as high as 97% in nestling PearlyEyed Thrashers (Margarops fuscatus) in Puerto Rico (Arendt 1985b), and is a significant cause of mortality in several other bird species as well (Fraga 1984; Nores 1995; Spalding et al. 2002; Fessl et al. 2006a). Subcutaneous invasion by other hematophagous dipteran
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larvae has also been associated with significant mortality in affected nestlings, presumably due to increased susceptibility to secondary bacterial infections (Young 1993; Warren 1994). Gross lesions associated with blood-feeding larvae that do not penetrate subcutaneously are limited to the presence of larvae on the birds during feeding and the presence of dried brownish-red larval excreta around feeding sites (Tirrell 1978). The skin surrounding the attachment site of feeding larvae may become swollen and edematous (Tirrell 1978). When larvae penetrate into the subcutaneous space and migrate, lesions are more severe. Uhazy and Arendt (1986) described the lesions associated with subcutaneous Philornis deceptivus in nestlings of Pearly-eyed Thrashers as raised and elongate nodules or as open cavities once larvae had exited. Microscopically, the lesions consisted of a cyst lined by mononuclear cells and surrounded by edematous connective tissue. Basophilic necrotic tissue and mononuclear cell infiltrations were associated with the larval mouthparts in active lesions. After larvae exited the cavity, wound contraction occurred and the cyst contained large numbers of mononuclear cells and a fibrinous exudate (Uhazy and Arendt 1986). Pathology caused by sarcophagid species during traumatic myiasis is similar to that caused by subcutaneously migrating hematophagous dipteran larvae. Maggots may be found within subcutaneous cysts that communicate with the surface of the skin via a small circular opening, or in open cavities (Wobeser et al. 1981). In domestic Greylag Geese (Anser anser) infested with Lucilia sericata and W. magnifica, moderate to severe bleeding is also noted at wound sites and epidermal necrosis seen at the edges of wounds (Farkas et al. 2001).
develop to adults. Information on collecting, rearing, and preserving Protocalliphora spp. can be found in Sabrosky et al. (1989). Care should also be taken in identifying the agents of traumatic or wound myiasis, particularly when larvae are collected from dead birds. Carcass-feeding larvae are commonly found on vertebrate carcasses in the wild and are unlikely to have played a role in the death of the bird (Baumgartner 1988).
DIAGNOSIS Avian myiasis due to hematophagous dipteran larvae is readily diagnosed by observing the presence of bloodfed larvae on or in the nests of live birds, or upon direct examination of affected nestlings. Larvae are found on or around nestlings within the first days to weeks after chicks hatch. Larvae may be seen feeding on the nestlings directly or may be found in the nest material between feedings. However, identification of the exact species of larvae found may be more difficult. Keys are available for identification of dipteran larvae found on nestlings (e.g., Hall 1948; Teskey 1981; Sabrosky et al. 1989; Foote 1991; Whitworth 2002). Use of these keys can be challenging, and guidance from an experienced entomologist familiar with taxonomy and identification of dipteran larvae is critical. Identification of these larvae may also be facilitated by collection of live larvae from the nest and then allowing some specimens to
PUBLIC HEALTH AND DOMESTIC ANIMAL CONCERNS There is no known public health concern associated with hematophagous myiasis flies in birds. Traumatic or wound myiasis does occur in people, but fly infestations on birds are not considered a major source of adult flies which can then deposit their offspring on humans. Similarly, there is no major domestic animal health concern associated with avian myiasis. Traumatic or wound myiasis does occur in domestic animals, including domestic birds such as geese (Farkas et al. 2001), but as with myiasis in people, wild birds are not considered a major source of the adult flies.
IMMUNITY Little is known about immune responses of nestling birds to Protocalliphora spp. or other hematophagous dipteran larvae. Although occasionally some species of maggots may be found feeding on older birds, a strong preference for newly hatched nestlings has been shown in several individual species and is likely true for most of these diptera (Sabrosky et al. 1989). The relatively low numbers of larvae on adult birds may also be due to removal by preening (Arendt 1985b). The immune response to traumatic or wound myiasis in birds is similarly understudied, but inferences can be drawn from what is known about the response to infestation in mammalian hosts, particularly livestock. Following infestation by Lucilia cuprina in sheep, granulocytes infiltrate the wound surface while lymphocytes aggregate in the dermis. High antibody titers occur upon secondary infestation; however, titers do not appear to differ between resistant and susceptible animals, suggesting any resistance to infestation may be an innate response (Otranto 2001). At present, vaccination for myiasis is not available in humans or domestic animals.
WILDLIFE POPULATION IMPACTS The role of blood-feeding larvae of bird nest flies as pathogens that can impact wild bird populations
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Myiasis in Wild Birds remains controversial. Infestations of nestlings with hematophagous dipteran larvae are almost ubiquitous across passerine taxa. The Protocalliphora spp. alone appear capable of parasitizing all species of North American birds with altricial young and dry nests (Sabrosky et al. 1989; Bennett and Whitworth 1991), and do not appear to themselves result in wide-scale mortality. However, when coupled with environmental stressors such as habitat reduction, inclement weather, and limited nutrient sources, the presence of the larvae of Protocalliphora spp. and resultant increased feeding demands on parents of heavily parasitized nestlings may contribute to both fledgling failure and decreased parental survival in future years (Hurtrez-Bousses et al. 1998; Simon et al. 2004). While it seems intuitive that chronic blood loss in young, developing birds is compromising, several studies have not shown any adverse effects on the weight or fledging success of the nestlings themselves. Indeed, infestations of P. braueri on nestling House Wrens (Troglodytes aedon) did not have an effect on either nestling survival or fledgling size as measured by tarsus length at fledging (Eastman et al. 1989). However, in other instances, heavy infestations are associated with delayed development, prolonged time to fledge, and nestling mortality (see summary in Sabrosky et al. 1989). In times of abundant food, increased parental feeding may compensate for the physiologic cost of chronic blood loss associated with the hematophagous larvae (Hurtrez-Bousses et al. 1998; Simon et al. 2004). However, if inclement weather or food shortages coincide with heavy infestations by Protocalliphora spp., these diptera may contribute to nestling mortality (Sabrosky et al. 1989; Howe 1992; Whitworth and Bennett 1992). For example, a study of P. braueri in nestling Sage Thrashers (Oreoscoptes montanus) found no effect on fledgling mortality in a year with consistently mild weather, but the following year, when an 8-day period of severe inclement weather occurred, parasitism by P. braueri was positively associated with nestling mortality (Howe 1992). Habitat disruption can also impact patterns of infestation by Protocalliphora spp. on nestlings. In a recent study of Tree Swallows (Tachycineta bicolor) in Alberta, Canada, 100% of nests examined were infested with Protocalliphora spp., and most of the nests contained more than one species. Interestingly, nests were much more heavily infested on wetlands impacted by oil sands mining than those on an undisturbed site, and nestling growth was adversely affected by the higher parasite load (Gentes et al. 2007). Muscid larvae, such as those of Philornis spp. or Passeromyia spp., can also be significant mortality factors for nidicolous birds in tropical areas (Arendt 1985b; Fessl et al. 2006a). These dipteran larvae have
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been reported as a cause of severe impacts in endangered species of birds when they encounter newly introduced dipteran parasites. Such a problem may have occurred when Pearly-eyed Thrashers invaded the forest reserve of the Puerto Rican Parrot (Amazona vittata), bringing with them Philornis spp. Fatal parasitism of parrot nestlings with Philornis larvae was subsequently reported (Snyder et al. 1987). This situation underscores the potentially catastrophic result of introduction of a new parasite into an already fragile system. Introduction of P. downsi into the avifauna of the Galapagos Islands appears to be having similar negative impact on survival of nestling Darwin’s finches, including the Mangrove Finch (Camarhynchus heliobates) and the Medium-Tree Finch (Camarhynchus pauper), of which there are only an estimated 50 and 300 breeding pairs remaining, respectively (Fessl et al. 2006a). Very little information is available about the degree to which traumatic myiasis affects bird populations, although reports of this condition caused by Wohlfahrtia, Lucilia, and Calliphora spp. and associated with morbidity and mortality in waterfowl, raptors, and passerines can be found in the literature (Eschele and DeFoliart 1965; Cooper 1978; Wobeser et al. 1981; Baumgartner 1988; Farkas et al. 2001) (Table 32.2). Infestations of wounds with fly larvae can cause severe disease in affected individuals, but the impact on populations is considered relatively minor. The larvae of W. opaca have been identified as a cause of mortality in ducklings, but only 0.7% of examined ducklings were found infested (Wobeser et al. 1981). Similarly, a recent survey of geese in Hungary found only ∼0.1% of domestic geese infested with W. magnfica and L. sericata (Farkas et al. 2001).
TREATMENT AND CONTROL Natural mechanisms that result in removal of hematophagous dipteran larvae from nests containing young birds have been shown to increase nestling survival. Adult Bay-winged Cowbirds (Agelaioides badius) that removed larvae from their nestlings increased nestling survival (Fraga 1984). Similarly, when various species of oropendola and cacique located nests near parasitic wasp nests, parasitism by Philornis spp. declined and fledging success increased (Smith 1968). However, artificial removal of dipteran larvae from the nests of birds is not practical in the wild and may adversely disturb developing nestlings. If parasitism is quite high, infested nests can be removed and destroyed and replaced with artificial ones. This approach would only be effective for controlling larvae that do not burrow subcutaneously or remain on the birds for long periods of time.
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Application of insecticides to the nest material has also been proposed, although this approach has inherent limitations in terms of both safety and practicality (Sabrosky et al. 1989). Application of insecticide to nests is perhaps best pursued when severe outbreaks occur that involve threatened or endangered birds. Treatment of nests with insecticides was effective during an outbreak of P. downsi on nestlings of Darwin’s finches. Application of 1% pyrethrin solution to the nests cleared the infestations and significantly improved fledgling success relative to untreated controls. When treated soon after eggs hatched, the nests remained free of P. downsi for the remainder of the nestling period (Fessl et al. 2006b). The safety of pesticide treatments and other similar approaches, such as hanging pest strips near the nest, should be explored further as the potential benefit for nestling survival may be quite significant. In the Galapagos finch study, treated nests had a fledgling rate more than twice that of untreated nests (86.6 and 33.9%, respectively) (Fessl et al. 2006b). Treatment of wound myiasis requires removing the larvae from the affected tissue, managing the wound to encourage healing, and providing necessary supportive care. Animals recovering from myiasis must be kept indoors and protected from flies to prevent reinfestation of healing wounds with fresh maggots. Because the flies that cause facultative and obligatory myiasis develop rapidly and can continue to develop in carcasses after death, prevention of wounds, such as occurs during feather plucking of geese, and prompt carcass removal may aid control of these flies during a die-off (Farkas et al. 2001).
MANAGEMENT IMPLICATIONS Although most bird populations appear to tolerate the presence of blood- and tissue-feeding larvae on their young, these parasites may create an added stressor in times of limited nutrients and excess energy requirements. Some ecologists have speculated that increasing edge effects between different habitats will bring previously segregated species closer together, possibliy leading to spread of parasites, including bird nest flies, to new hosts and new populations. Loss of nesting habitat that forces more frequent use of old nest sites may also increase exposure to bird nest flies among nestlings (Loye and Carroll 1995).
LITERATURE CITED Arendt, W. J. 1985a. Philornis ectoparasitism of pearly-eyed thrashers. I. Impact on growth and development of nestlings. The Auk 102:270–280.
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Hinaidy, H. K., and H. Frey. 1984. Further cases of facultative myiasis in vertebrates in Austria. Wiener Tierarztliche Monatsschrift 71:237–238. Howe, F. P. 1992. Effects of Protocalliphora braueri (Diptera: Calliphoridae) parasitism and inclement weather on nestling sage thrashers. Journal of Wildlife Diseases 28:141–143. Hurtrez-Bousses, S., J. Blondel, and P. Perret. 1997a. Relationship between intensity of blowfly infestation and reproductive success in a Corsican population of blue tit*. Journal of Avian Biology 28:267–270. Hurtrez-Bousses, S., P. Perret, F. Renaud, and J. Blondel. 1997b. High blowfly parasitic loads affect breeding success in a Mediterranean population of blue tit*. Oecologia 112:514–517. Hurtrez-Bousses, S., J. Blondel, P. Perret, J. Fabreguettes, and F. Renaud. 1998. Chick parasitism by blowflies affects feeding rates in a Mediterranean population of blue tit*. Ecology 1:17–20. Itamies, J., and E. Merila. 1984. A Lucilia myiasis in a living crane (Grus grus). Luonnon Tutkija 88:138. Johnson, L. S., and D. J. Albrecht. 1993. Effects of haematophagous ectoparasites on house wrens, Troglodytes aedon: Who pays the cost of parasitism? Oikos 66:255–262. Kettle, D. S. 1995. Medical and Veterinary Entomology. Oxford University Press, Oxford, UK. Lindholm, A. K., G. J. Venter, and E. A. Ueckermann. 1998. Persistence of passerine ectoparasites on the diederik cuckoo Chrysococcyx caprius. Journal of Zoology (London) 244(1):145–153. Loye, J., and S. Carroll. 1995. Birds, bugs and blood: Avian parasitism and conservation. Trends in Ecology and Evolution 10:232–235. Matsuoka, S. M., C. M. Handel, and D. D. Roby. 1987. Nesting ecology of Townsend’s warblers in relation to habitat characteristics in a mature boreal forest. Condor 99(2):271–281. Miller, C.K., and J. M. Fair. 1997. Effects of blow fly (Protocalliphora spatulata) (Diptera: Calliphoridae) parasitism on the growth of nestling savannah sparrows in Alaska. Canadian Journal of Zoology/Revue Canadien de Zoologie 75(4):641–644. Nores, A. I. 1995. Botfly ectoparasitism of the brown cacholote and the firewood-gatherer. Wilson Bulletin 107:734–738. O’Brien, E. L., B. L. Morrison, and L. S. Johnson. 2001. Assessing the effects of haematophagous ectoparasites on the health of nestling birds: Hematocrit versus haemoglobin levels in house wrens parasitized by blow fly larvae. Journal of Avian Biology 32:73–76. Otranto, D. 2001. The immunology of myiasis: Parasite survival and host defense strategies. Trends in Parasitology 17:176–182.
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Pletsch, D. D. 1948. Parasitic dipterous larvae from the nasal cavity of a nestling magpie. The Auk 65:296–297. Pont, A. C. 1974. A revision of the genus Passeromyia Rodhain and Villeneuve (Diptera: Muscidae). Bulletin of the British Museum of Natural History (Entomology) 30:339–372. Proudfoot, G. A., J. L. Usener, and P. D. Teel. 2005. Ferruginous pygmy-owls: A new host for Protocalliphora sialia and Hesperocimex sonorensis in Arizona. Wilson Bulletin 117(2):185– 188. Proudfoot, G. A., P. D. Teel, and R. M. Mohr. 2006. Ferruginous pygmy-owl (Glaucidium brasilianum) and Eastern screech-owl (Megascopes asio): New hosts for Philornis mimicola (Diptera: Muscidae) and Ornithodoros concanensis (Acari: Argasidae). Journal of Wildlife Diseases 42(4):873–876. Revels, M. 1996. Eight new host species for the parasitic blow fly genus Protocalliphora. Wilson Bulletin 108(l):189–190. Roberts, L. S., and J. Janovy. 2005. Foundations of Parasitology. McGraw-Hill, New York, 702 pp. Sabrosky, C. W., G. F. Bennett, and T. L. Whitworth. 1989. Bird Blow Flies (Protocalliphora) in North America (Diptera: Calliphoridae) with notes on the Palearctic species. Smithsonian Institution, Washington, DC, 312 pp. Sargent, W. D. 1938. Nest parasitism of hawks. The Auk 55:82–84. Seguy, E. 1955. Introduction a l’etude biologique et morphologique des insectes dipteres. Publicacoes Avulsas Museu Nacional Rio de Janeiro 17:1–260. Shields, W. M., and J. R. Crook. 1987. Barn swallow coloniality: A net cost for group breeding in the Adirondacks. Ecology 68(5):1373–1386. Simon, A., D. Thomas, J. Blondel, P. Perret, and M. M. Lambrechts. 2004. Physiological ecology of Mediterranean blue tit* (Parus caeruleus L.): Effects of ectoparasites (Protocalliphora spp.) and food abundance on metabolic capacity of nestlings. Physiological and Biochemical Zoology 77: 492–501. Smith, N. G. 1968. The advantage of being parasitized. Nature 219:690–694. Snyder, N. R. F., J. W. Wiley, and C. B. Kepler. 1987. The Parrots of Luquillo: Natural History and Conservation of the Puerto Rican Parrot. Western Foundation of Vertebrate Zoology, Los Angeles, CA, 384 pp. Spalding, M. G., J. W. Mertins, P. B. Walsh, K. C. Morin, D. E. Dunmore, and D. J. Forrester. 2002. Burrowing fly larvae (Philornis porteri) associated with mortality of eastern bluebirds in Florida. Journal of Wildlife Diseases 38:776–783.
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Index Acheta domestica, 378 Acipenser gueldenstaedtii (Sturgeon), 301 Aconoidasida, 15, 36 Acrocephalus schoenobaenus (Sedge Warbler), 345, 479 Acrochordus javanicus (Javan wart snake), 302 Acryllium vulturinum (Vulturine Guinea Fowl), 393, 397, 473 Actitis, 465 Actitis hypoleucos (Common Sandpiper), 475 Actitis macularius (Spotted Sandpiper), 332, 475 Actophilornis africanus (African Jacana), 331, 339 Actornithophilus limosae, 521 Acuaria spp. Acuaria (Hamannia) uncinata, 335 Acuaria nasuta, 327 Acuaria spiralis, 327 Acuariasis, 335 Acuarioid nematodes, 326 Acuariosis, 326 Aechmophorus occidentalis (Western Grebe), 328 Aedeomyia squamipennis, 41 Aegithinidae (Ioras), 61, 444 Aegolius acadicus (Northern Saw-whet Owl), 114, 477 Aegothelidae (Owlet-Nightjars), 443 Aegypius monachus (Cinereous Vulture), 205 Aetheobdella hirudoides, 505 Aethia cristatella (Crested Auklet), 394 Aethia pygmaea (Whiskered Auklet), 394 African Black Duck (see Anas sparsa) African Fish-Eagle (see Haliaeetus vocifer) African Goshawk (see Accipiter tachiro) African Green-Pigeon (see Treron calvus) African Jacana, (see Actophilornis africanus) African Mourning Dove (see Streptopelia decipiens) African Spoonbill (see Platalea alba) African Swift (see Apus barbatus) Agap*rnis, 465 Agap*rnis fischeri (Fischer’s Lovebird), 399 Agap*rnis personatus (Yellow-collored Lovebird), 401 Agar gel precipitation, 92 Age, as disease determinant of wild birds, 3 Agelaius phoeniceus (Red-winged Blackbird), 42, 209, 479 Ahanta Francolin (see Francolinus abantensis) Aigamo method, of rice farming, 255 air sacculitis, 527 Aix, 465 Aix galericulata (Mandarin Duck), 68, 329, 358, 364 Aix sponsa (Wood Duck), 22, 67, 83, 208, 236, 270, 329, 360, 365, 390, 468, 486 Akiba caulleryi, 54 Alauda arvensis (Eurasian Skylark), 479 Alaudidae (Larks), 60, 444 albendazole, 406, 424, 491 albumen, 234 Alca torda (Razorbill), 227, 332
Aatinga leucophthalma (White-eyed Parakeet), 401 Abdominal cavity, 449 Acanthocephala clinical signs, 280 diagnosis, 283 domestic animal health concerns, 283 epizootiology, 277–280 etiology, 277 history, 277 host range and distribution, 277 ova of some common acanthocephalans of birds, 284 pathogenesis and pathology, 280–283 proboscides of acanthocephalans of birds, 278 public health concerns, 283 synonym, 277 treatment and control, 285 wild population impacts, 283–285 Acanthocephalans, 277–279, 280, 282 eggs, 284 proboscides of, 278 Acanthocephaliasis, 277 Acanthocephalosis, 277 Acariasis diagnosis, 534 domestic animal health concerns, 534 epizootiology, 529–530 etiology, 527–529 host range and distribution, 527 immunity, 534 pathogenesis and pathology, 530–534 public health concerns, 534 synonyms, 527 treatment and control, 535 wildlife population impacts, 535 Accipiter, 465 Accipiter badius (Shikra), 77, 130 Accipiter brevipes (Levant Sparrowhawk), 78 Accipiter cooperii (Cooper’s Hawk), 76, 130, 144, 205, 237 Accipiter gentilis (Northern Goshawk), 76, 114, 130, 143–144, 205, 215, 346, 348, 470 Accipiter gularis (Japanese Sparrowhawk), 78 Accipiter minullus (Little Sparrowhawk), 78 Accipiter nisus (Eurasian Sparrow Hawk), 77, 84, 94, 346, 470 Accipiter rufiventris (Rufous-chested Sparrowhawk), 130 Accipiter soloensis (Chinese Goshawk), 78 Accipiter striatus (Sharp-shinned Hawk), 77, 395 Accipiter tachiro (African Goshawk), 78, 130 Accipiter trivirgatus (Crested Goshawk), 395, 471 Accipiter virgatus (Besra), 78 Accipitridae (Hawks, Eagles, and Kites), 57, 59, 74, 395, 441, 505 acetic acid, 509
557 Parasitic Diseases of Wild Birds Edited by Carter T. Atkinson, Nancy J. Thomas and D. Bruce Hunter © 2008 John Wiley & Sons, Inc. ISBN: 978-0-813-82081-1
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558 Alcedinidae (Kingfishers), 60, 443 Alcidae (Auks, Murres and Puffins), 442 Alectoris, 465 Alectoris barbara (Barbary Partridge), 331, 390, 396 Alectoris chukar (Chukar), 114, 155, 169, 391, 396 Alectoris graeca (Rock Partridge), 213, 214, 215, 392, 396 fed with Toxoplasma gondii oocysts, 213, 215 Alectoris rufa (Red-legged Partridge), 144, 171, 214, 270, 317, 392, 406 Alectroenas pulcherrima (Seychelles Blue-Pigeon), 125 Alectura lathami (Australian Brush-turkey), 390, 395 Alexandra’s Parrot (see Polytelis alexandrae) Alexandrine Parakeet. (see Psittacula eupatria) Alisterus scapularis (Australian King-Parrot), 399 Alle, 465 Alligator mississippiensis (American alligator), 303 Allobilharzia, 246, 247, 251 Allobilharzia visceralis, 251 Allolobophora caliginosa, 401, 464 Allolobophora parva, 464 allozymes, 425 Alopochen aegyptiaca (Egyption Goose), 69, 329 Alpine Accentor (see Prunella collaris) Alpine Swift (see Tachymarptis melba) Amabiliidae, 264 Amadina fasciata, 378 Amaurornis phoenicurus (White-Breasted Waterhens), 488 Amazona spp. Amazona aestiva (Blue-fronted Parrot), 399 Amazona agilis (Black-billed Parrot), 399 Amazona amazonica (Orange-winged Parrot), 400 Amazona arausiaca (Red-necked Parrot), 400 Amazona autumnalis (Red-lored Parrot), 400 Amazona farinose (Mealy Parrot), 400 Amazona festiva (Festive Parrot), 399 Amazona leucocephala (Cuban Parrot), 399 Amazona ochrocephala (Yellow-crowned Parrot), 401 Amazona vinacea (Vinaceous Parrot), 400 Amazona vittata (Puerto Rican Parrot), 400, 553 American Avocet (see Recurvirostra americana) American Bittern (see Botaurus lentiginosus) American Black Duck (see Anas rubripes) American Coot (see Fulica americana) American Crow (see Corvus brachyrhynchos) American Goldfinch (see Carduelis tristis) American Kestrel (see Falco sporverius) American Redstart (see Setophaga ruticilla) American White Pelican (see Pelecanus esythroshynchos) American Wigeon (see Anas americana) American Woodco*ck (see Scolopax minor) Amblycera, 516, 517 Amherst (AG) strain, 134 Amicobdella nigra, 505 Amidostomatinae, 366 Amidostomiasis, 355 Amidostomum spp., 355 Amidostomum acutum, 355, 356, 357, 358, 361 Amidostomum anseris, 355, 356, 357, 358, 361, 366 Amidostomum auriculatum, 357, 366 Amidostomum biziurae, 366 Amidostomum cygni, 356, 358, 366 Amidostomum fulicae, 356, 357, 358, 361, 366 Amidostomum henryi, 358, 361, 366 Amidostomum quasifulicae, 361 Amidostomum raillieti, 366 Amidostomum ryzhikovi, 366
Index Amidostomum skrjabini, 370 Amidostomum spatulatum, 355, 356, 358, 366 Amidostomum species, 355 Amidostomum tribonix, 366 Amidostomum tribonyx, 361, 366 avian hosts infected with, 362–365 clinical signs, 368 diagnosis, 370 domestic animal health concerns, 370 eggs, 367 epizootiology, 367–368 etiology, 366–367 history, 355 host range and distribution, 355–366 immunity, 370 management implications, 371 pathogenesis and pathology, 368–370 prevalence, intensity and abundance, 367–368 public health concerns, 370 synonyms, 355 treatment and control, 371 wildlife population impacts, 371 Ammodramus humeralis (Grassland Sparrow), 114 ammonia, 509 Ammoperdix griseogularis (See-see Partridge), 393 Amoebotaenia, 263 Ampeliceps coronatus (Golden-crested Myna), 111 Amphibiocapillaria, 463 Amphimerus, 231 Amphimerus elongates, 238 Amphimerus elongatus, 233, 235 Amphimerus heterolecithodes, 233, 237 Amphipoda, 279 Amphiuma tridactylum (Three-toed amphiuma), 303 Amplicaecum Baylis, 413 Amprolium, 172, 177, 192 Anas spp., 236, 336, 465, 488 Anas acuta (Northern Pintail), 63, 173, 205, 227, 235–236, 296, 329, 336, 355, 358, 364, 395 Anas americana (American Wigeon), 65, 115, 329, 356, 362, 368 Anas bahamensis (White-cheeked Pintail), 329 Anas capensis (Cape Teal), 329, 362 Anas carolinensis (Green-winged Teal), 227, 235, 355, 357, 363 Anas castanea (Chestnut Teal), 329 Anas clypeata (Northern Shoveler), 63, 115, 173, 235, 296, 329, 359, 364, 550 Anas creccai (Eurasian Teal), 64, 173, 468 Anas cyanoptera (Cinnamon Teal), 329, 356, 362 Anas discors (Blue-winged Teal), 64, 115, 173, 227, 234, 235, 236, 249, 329, 336, 356, 362, 550 Anas erythrorhyncha (Red-billed Duck), 364 Anas falcata (Falcated Duck), 65, 329, 357, 363 Anas flavirostris (Speckled Teal), 390 Anas formosa (Baikal Teal), 65, 329, 356, 362 Anas fulvigula (Mottled Duck), 83, 235, 358, 364, 468 Anas georgica (Yellow-billed Pintail), 329, 360 Anas gracilis (Gray Teal), 357, 363 Anas laysanensis (Laysan Duck), 339 Anas luzonica (Philippine Duck), 330 Anas penelope (Eurasian Wigeon), 65, 329, 357, 363, 389 Anas platyrhynchos and others (Mallard), 346 Anas platyrhynchos, 62, 205, 217 Anas platyrhynchos (Domestic White Pekin), 296, 502 Anas platyrhynchos (Ducks), 163, 488 Anas platyrhynchos (Juvenile Mallards), 367
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Index Anas platyrhynchos (Mallard), 62, 83, 154, 173, 205, 217, 228, 233, 235, 236, 255, 296, 304, 329, 336, 358, 364, 379, 380, 389, 395, 468, 504 Anas platyrhynchos (Mallard ducklings), 250 Anas poecilorhyncha (Spot-billed Duck), 359, 365 Anas querquedula (Garganey), 63, 296, 330, 357, 363 Anas rubripes (American Black Duck), 65, 83, 227, 234, 235, 236, 249, 250, 330, 356, 362, 445, 451 Anas sibilatrix (Chiloe Wigeon), 329, 356 Anas smithii (Cape Shoveler), 362 Anas sparsa (African Black Duck), 330, 356 Anas specularis (Spectacled Duck), 330, 359 Anas strepera (Gadwall), 65, 173, 205, 330, 336, 338, 357, 363, 389 Anas superciliosa (Pacific Black Duck), 330, 359, 502 Anas undulata (Yellow-billed Duck), 330, 360 Anas versicolor (Silver Teal), 329 Anastomus oscitans (Asian Openbill), 235 Anatidae (Ducks, Geese, and Swans), 59 Anatini (Dabbling Ducks), 367 Andersonfilaria, 441–445, 448, 454 Andersonfilaria africanus, morphology of microfilariae of, 453 anemia, 21, 44, 87, 88–89 Anhinga (see Anhinga anhinga) Anhimidae (screamers), 441 Anhinga anhinga (Anhinga), 293, 416, 466 Anhinga melanogaster (Darter), 293 Anhingidae (Anhingas), 59, 441 Anisakiasis, 413 anisakid infections, 424 Anodorhynchus hyacinthinus (Hyacinth Macaw), 400 anonymity, of wild birds, 3 anopheline mosquitoes, 40 Anoplura, 516 anorexia, 21, 87 Anous minutus (Black Noddy), 416, 423 proventriculus of a, 422 anoxia, 44 Anseranas semipalmata (Magpie Goose), 208, 217 Anseriformes, 14, 18, 22, 23, 54, 59, 121, 163, 165, 166, 173–175, 205, 208, 217, 235, 247, 251, 252, 253, 296, 322, 328, 376, 441, 465, 468, 521 Anser spp., 465 Anser albifrons (Greater White-fronted Goose), 69, 328, 357, 363, 389 Anser anser (Graylag Goose), 69, 173, 174, 205, 296, 358, 363, 367, 389, 395, 550, 552 Anser brachyrhynchus (Pink-footed Goose), 359 Anser cygnoides (Swan Goose), 69, 328, 346, 390 Anser erythropus (Lesser White-fronted Goose), 358, 364, 389 Anser fabalis (Taiga Bean-Goose), 69, 356, 362 Anser indicus (Bar-headed Goose), 356, 362 anthelmintics, 322, 339, 352, 371, 386, 406 anthropogenic alterations, impact of, 7 Anthropoides paradiseus (Blue Crane), 182, 397 Anthropoides virgo (Demoiselle Crane), 181, 182, 397 Anthus spp. Anthus hodgsoni (Olive-backed Pipit), 479 Anthus pratensis (Meadow Pipit), 479 Anthus trivialis (Tree Pipit), 479 anti-Eimeria IgM, IgY, and IgA antibodies, 171 anti-erythrocyte factor, 89 Aonchotheca, 463 aorta, 449 Apapane (see Himatione sanguinea)
559
Apatemon, 231 Aphelocoma coerulescens (Florida Scrub Jay), 333, 434 Apicomplexa, 15, 36, 163, 195, 204 Aploparaksis furcigera, 264, 268, 269 Aploparaksis penetrans, 264, 268 Apodidae (Swifts), 443 Apodiformes, 14, 59, 163, 435, 443, 478, 521 Aporrectodea rosea, 486 Aproctella, 441–445, 448, 454 Aproctella alessandroi, 451 Aproctella stoddardi, morphology of microfilariae, 453 Aproctiana, 441–445, 448, 454 Aproctoidea, 440 Aproctoids, 439 Aprosmictus erythropterus (Red-winged Parrot), 400 Apterygidae (Kiwis), 441 Apterygiformes, 14, 175, 441 Apteryx mantelli (North Island Brown Kiwi), 175 Apus spp. Apus affinis (Little Swift), 446, 451 Apus apus (Common Swift), 478 Apus barbatus (African Swift), 521 Aquila spp Aquila chrysaetos (Golden Eagle), 75, 76, 131, 215 Aquila clanga (Greater Spotted Eagle), 76, 297 Aquila clanga (Spotted Eagle), 298 Aquila fasciata (Bonelli’s Eagle), 131 Aquila pennatus (Booted Eagle), 131 Aquila pomarina (Lesser Spotted Eagle), 76, 297 Aquila rapax (Tawny Eagle), 76 Aquila wahlbergi (Wohlberg’s Eagle), 76 Ara spp. Ara ararauna (Blue-and-yellow Macaw), 233, 399, 427 Ara chloropterus (Red-and-green Macaw), 400 Ara macao (Scarlet Macaw), 400, 427 Aramidae (Limpkins), 360, 442 Aramides saracura (Slaty-breasted Wood-Rail), 114 Aramus guarauna (Limpkin), 360 Aratinga spp. Aratinga acuticaudata (Blue-crowned Parakeet), 399 Aratinga cactorum (Caatinga Parakeet), 399, 476 Aratinga chloroptera (Hispaniolan Parakeet), 399 Aratinga jandaya (Jandaya Parakeet), 400 Aratinga leucophthalma (White-eyed Parakeet), 401 Aratinga pertinax (Brown-throated Parakeet), 399 Aratinga solstitialis (Sun Parakeet), 400 Arborophila torqueola (Hill Partridge), 391 Arctic Loon (see Gavia arctica) Ardea, 465 Ardea alba (Great Egret), 115, 294, 304, 384, 468 Ardea cinerea (Gray Heron), 293, 346 Ardea cocoi (Cocoi Heron), 294 Ardea goliath (Goliath Heron), 294 Ardea herodias (Great Blue Heron), 115, 235, 293, 304, 308, 468 Ardeidae (Herons, Egrets, and Bitterns), 59, 441, 505 Ardeiformes, 376 Ardeola, 251 Ardeola bacchus (Chinese Pond-heron), 294, 488 Ardeotis kori (Kori Bustard), 269 arecoline hydrobromide, 424 Arhythmorhynchus, 277 Armadillidium vulgare, 327 Artamidae (Wood Swallows), 445 Arthrocystis galli, 23 arthropod ectoparasites, effects of, 3 ascariasis, 388
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560 Ascaridia spp., 402 Ascaridia aegyptiaca, 394, 395 Ascaridia amblimoria, 401 Ascaridia australis, 398, 401 Ascaridia bonasae, 396, 397 Ascaridia borealis, 397 Ascaridia brasiliana, 395 Ascaridia catheturina, 395 Ascaridia circularis, 401 Ascaridia columbae, 394, 396, 398, 399, 401, 405 Ascaridia compar, 395, 396, 397 Ascaridia cordata, 395 Ascaridia cristata, 397 Ascaridia cuculina, 401 Ascaridia dissimilis, 397, 401, 402 Ascaridia dolichocerca, 395 Ascaridia fasciata, 397 Ascaridia francolina, 395, 396 Ascaridia galli, 388, 394, 395, 396, 397, 399, 401, 402, 406 Ascaridia geei, 394, 398 Ascaridia hermaphrodita, 398, 399, 400, 401 Ascaridia hermaphroditia, 394 Ascaridia longecirrata, 398 Ascaridia maculosa, 398 Ascaridia magalh˜aesi, 398 Ascaridia magnipapilla, 396 Ascaridia nicobarensis, 400 Ascaridia numidae, 396, 397, 401, 404 Ascaridia ornata, 400 Ascaridia orthocerca, 395 Ascaridia perspicillum, 402 Ascaridia pintoi, 395 Ascaridia platyceri, 394, 398, 399, 400, 401 Ascaridia pterophora, 397 Ascaridia razia, 398 Ascaridia sergiomeirai, 395, 399, 400, 401 Ascaridia serrata, 395 Ascaridia skrjabini, 396 Ascaridia stroma, 397 Ascaridia struthionis, 395 Ascaridia styphlocerca, 395, 396 Ascaridia trilabium, 401 avian hosts infected with, 395–401 clinical signs, 404 diagnosis, 405 distribution and host range, 388–394 epizootiology, 401–404 etiology, 394–401 history, 388 immunity, 405 pathology, 404–405 public and domestic animal health concerns, 405 synonyms, 388 treatment and control, 406–407 wildlife population impacts, 405–406 Ascaridoidea, 413, 414 Ascaridoid infections, 424 Ascaridoid parasites, 339 Ascaris spp., 425 Ascarosis, 388 Ascocotyle, 230 Ascometra choriotidis, 264, 268 asexual reproduction of Isospora species, 110 of Plasmodium gallinaceum, 39 of Plasmodium species, 36
Index asexual sporogonic cycle, of Haemoproteus, 17–18 Ashy-headed Goose (see Chloephaga poliocephala) Asian Openbill (see Anastomus oscitans) Asio flammeus (Short-eared Owl), 550 Asio otus (Northern Long-eared Owl), 114, 477 aspartate aminotransferase, 42 Aspergillus sp., 140, 434 atebrine, 28, 95 Athene noctua (Little Owl), 54, 133, 206, 477 Athesmia, 229 Athesmia heterolecithodes, 226 Athesmia jolliei, 237 Atoxoplasma, 163, 204, 214. see also Atoxoplasmosis distribution, 109 domestic animal health concerns, 117 history, 108–109 public health concerns, 116 treatment and control, 117 wildlife population impacts, 117 Atoxoplasmosis. see also Atoxoplasma clinical signs, 112 diagnosis, 112–113 epizootiology, 110–112 etiology, 109–110 host range, 109 immunity, 113 pathogenesis and pathology, 112 synonym, 108 Atrichornithidae (Scrub-birds), 444 atropine sulfate, 371 Attila, 465 Austrobilharzia terrigalensis, 251 Australian Brush-turkey (see Alectum lathani) Australian Kestrel (see Falco cenchroides) Australian King Parrot (see Alisterus scapularis) Austral Parakeet (see Enicognathus Ferrugineus) Australasian Magpie (see Gymnoshina tibicen) Australian Masked-Owl (see Tyto novaehollandiae) Austrobilharzia spp., 231, 246, 247, 249, 251 Austrobilharzia terrigalensis, 251 Austrobilharzia variglandis, 249, 251, 255 Australian Pelican (see Pelecanus conspicillatus) Austromenopon phaeopodis, 521 Austrosimulium spp. Austrosimulium australense, 85 Austrosimulium dumbletoni, 85 Austrosimulium ungulatum, 85 avian coccidia, with direct life cycles, 108 avian filarioid genera, in families of birds, 441–445 avian filarioid nematodes, 453 Dirofilariinae, 454 Lemdaninae, 454 Onchocercinae, 454 Splendidofilariinae, 454 avian haemoproteids, 14, 15, 19 host distribution of by avian order, 14 mitochondrial and nuclear gene sequences, 17 role of host migratory behavior in cycle of transmission, 20 avian life history, 3 avian malaria, 13 clinical signs, 42–43 diagnosis, 45–46 distribution, 36 domesticated animal health concerns, 47 epizootiology, 39–42 etiology, 36–39
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Index history, 35–36 host range, 36 immunity, 46–47 intrinsic and extrinsic factors of distribution, 41 management implications, 48–49 mortality, 48 pathology and pathogenesis, 43–45 prevalence of, 41 public health concerns, 47 spatial and seasonal patterns of transmission, 40 stress-mediated changes in the immune system, 40 synonyms, 35 treatment and control, 48 wildlife population impacts, 47–48 avian species, of Plasmodium, 37 Avioserpensosis clinical signs, 385 diagnosis, 385 domestic animal health concerns, 386 epizootiology, 384–385 etiology, 384 history, 384 host range and distribution, 384 immunity, 385 pathology, 385 public health concerns, 385 treatment and control, 386 wildlife population impacts, 386 Avioserpens spp. in domestic birds, 385 goslings infected with, 385 infections in wild birds, 386 outbreaks of disease, 386 Avioserpens mosgovoyi, 384, 386 Avioserpens taiwana, 384 Aythya, 465 Aythya affinis (Lesser Scaup), 7, 66, 165, 166, 171, 173, 227, 225, 235, 236, 330, 338, 358, 368 Aythya americana (Redhead), 66, 173, 330, 359, 364 Aythya australis (White-eyed Duck), 330 Aythya collaris (Ring-necked Duck), 66, 359, 364, 468 Aythya ferina (Common Pochard), 66, 205, 357, 362, 395 Aythya fuligula (Tufted Duck), 66, 165, 205, 227, 296, 330, 359, 365 Aythya marila (Greater Scaup), 66, 227, 235, 236, 330, 357, 363, 469 Aythya nyroca (Ferruginous Pochard), 330, 357, 363, 469 Aythya valisineria (Canvasback), 66, 173, 227, 235, 330, 356, 362 Aythya valisineria (Canvas Dack), 330 Aythyini (Diving Ducks), 367 Azure-winged Magpie (see Cyanopica cyanus) Bacillus subtilis, 157 Bacillus thuringiensis israelensis (Bti), 94, 544 Baikal Teal (see Anas formosa), 65 Balaenicipitidae (Shoebills), 59, 441 Bald Eagle (see Haliaeetus leucocephalus) Balearica, 181 Balearica pavonina (Black Crowned-Crane), 22, 397, 475 Balearica regulorum (Gray Crowned-Crane), 397 Bali Myna (see Leucopsar rothschildi) Band-tailed Pigeon (see Patagioenas fasciata) Bangkok hemorrhagic disease, 54 Barbary Partridge (see Alectoris barbara) Bare-faced Ibis (see Phimosus infuscatus)
561
Barnacle Goose (see Branta leucopsis) Barnardius, 465 Barnardius barnardi (Mallee Ringneck), 400 Barnardius zonarius (Port Lincoln Parrot), 400, 464, 476 Bar-headed Goose (see Anser indicus) Bar-shouldered Dove (see Geopelia humeralis) Bar-tailed Godwit (see Limosa lapponica) Barn Owl (see Tyto alba) Barn Swallow (see Hirundo rustica) Barred Owl (see Strix varia) Barrow’s Goldeneye (see Bucephala islandica) Baruscapillaria spp., 463 Baruscapillaria belopolskaiae, 476 Baruscapillaria calliopsis, 482 Baruscapillaria cincli, 479 Baruscapillaria corvorum, 480, 482 Baruscapillaria emberizae, 479, 481, 484 Baruscapillaria falconis, 470, 471, 472, 477, 487–488 Baruscapillaria grallinae, 481 Baruscapillaria inflexa, 479, 482, 485 Baruscapillaria jaenschi, 467, 476 Baruscapillaria mergi, 466, 467, 468, 469, 470 Baruscapillaria montevidensis, 473 Baruscapillaria obsignata, 463, 465, 466, 470, 487, 488, 491 Baruscapillaria ovopunctata, 485 Baruscapillaria resecta, 480, 481, 490 Baruscapillaria rudolphii, 467 Baruscapillaria ryjikovi, 466 Baruscapillaria spilculata, 466, 467 basophilic cytoplasmic inclusions, 191 basophils, 21 baylisascariasis, 424 Baylisascaris spp. Baylisascaris columnaris, 425 Baylisascaris laevis, 425 Baylisascaris melis, 425 Baylisascaris procyonis, 425, 427 clinical signs, 426 diagnosis, 425–426 domestic animal health concerns, 428 epizootiology, 425 etiology, 424–425 host range and distribution, 425 host range of larvae (larva migrans) of species of, 426 immunity, 427–428 pathology, 427 public health concerns, 428 synonyms, 424 treatment and control, 428 wildlife population impacts, 428 Baylisacaris columnaris larva, cross section of, 427 Baylisacaris procyonis, live specimen (L3 stage) of, 427 Baylisascaris transfuga, 425, 427 Bean Goose (see Anser fabalis) Belted Kingfisher (see Megaceryle alcyon) benign nature, of parasites, 5 Bennettinia, 37, 39 benzimidazole anthelmintics, 352 Besnoitia infections, 26 Besra (see Accipiter virgatus) Bilharziella, 231, 246, 247, 249, 251 Bilharziella polonica, 251 Bilharziella yokogawai, 246 bird–filarioid relationships, 448 bird mange, 527 Bithynia tentaculata (Faucet Snail), 226–227, 240
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Index
Biziura lobata (Musk Duck), 358 Black-billed Parrot (see Amazona agilis) Black-billed Wood-Dove (see Turtur abyssinicus) Black Crowned Crane (see Balearica pavonina) Black Eagle (see Ictinaetus malayensis) black flies (see Simulium spp) clinical signs, 541 diagnosis, 541–542 distribution, 537–538 epizootiology, 539–540 etiology, 538–539 history, 537 host range, 538 immunity, 542 pathogenesis, 540–541 public and domestic animal health concerns, 542 synonyms, 537 treatment and control, 543–544 wildlife population impacts, 542–543 Black Grouse (see Tetrao tetrix) Black Kite (see Milvus migrans) Black Noddy (see Anous minutus) Black Scoter (see Melanitta nigra) Blackhead disease, 154 Black-necked Swan (see Cygnus melancoryphus) Black-shouldered Kite (see Elanus caeruleus) Black-and-white Warbler (see Mniotilta variai) Black-billed Capercaillie (see Tetrao parvirostris) Black-billed Cuckoo-Dove (see Macropygia nigrirostris) Black-billed Magpie (see Pica hudsonia) Black-crowned Night Heron (see Nycticorax nycticorax) Black-fronted Piping-Guan (see Pipile jacutinga) Black-headed Ibis (see Threskiornis melanocephalus) Black-necked Aracari (see Pteroglossus aracari) Black-winged Stilt (see Himantopus himantopus) Black Swan (see Cygnus atratus) Black Vulture (see Coragyps atratus) Black-winged Lorry (see Eos cyanagenia) Blackburnian Warbler (see Dendroica fusca) Blattella germanica (co*ckroaches), 378 Black Francolin (see Francolinus froncolmus) Black Stork (see Ciconia nigra) Black-billed Whistling Duck (see Dendrocygna autumalis) Black-faced Cormorant (see Phalacrocorax fuscescens) Black-faced Cuckoo-shrike (see Coracina novaehollandiae) Black-headed Gull (see Larus ridibundus) Black-tailed Native-hen (see Gallinula chloropus) blood-feeding ectoparasites, 4, 108, 533, 577 Blood Pheasant (see Ithaginis cruentas) blood-stained diarrhea, 228 Blossom-headed Parakeet (see Psittacula roseata) blow fly (Protocalliphora braueri) larvae, 5 Blue-and-yellow Macaw (see Aca ararauna) Bluebonnet (see Northiella haematogaster) Blue-crowned Paraked (see Aratinga acuticaudata) Blue Dacnis (see Dacnis cayana) Blue Crane (see Arithropodes paradiseus) Blue Duck (see Hymenolaimus malacorhynchos) Blue Eared-Pheasant (see Crossoptilon auritum) Blue-and-yellow Macaw (see Ara arauna) Blue-black Grassquit (see Volatina jacarina) Blue-headed Parrot (see Pionus menstruus) Blue-fronted Parrot (see Amazona aestiva) Blue-gray Tanager (see Thraupis episcopus) Blue Grouse (see Dendragapus obscurus) Blue Jay (see Cyanocitta cristata) Blue Rock-thrush (see Monticola solitorius)
Blue-spotted Wood-Dove (see Turtur afer) Blue-winged Macaws (Primolius maracana) Blue-winged Teal (see Anas discors) Blythipicus rubiginosus (Maroon Woodpecker), 478 Boat-Tailed Grackle (see Quicalus major) Bombycillidae (Waxwings), 60, 435, 444 Bonasa, 465 Bonasa bonasia (Hazel Grouse), 360, 391, 396 Bonasa umbellus (Ruffed Grouse), 20, 155, 167, 226, 327, 331, 388, 393, 397, 529 Bonelli’s Eagle (see Aquila fasciata) Boopidae, 516 Booted Eagle (see Aquila pennatus) borrachudo, 537 Botaurus, 465 Botaurus lentiginosus (American Bittern), 295 Botaurus pinnatus (Pinnated Bittern), 295 Botaurus stellaris (Great Bittern), 328 bothridia, 262 Bothrigaster, 229 Bothrigaster variolaris, 236 Bothrigaster variolaris (Cyclocoelidae), 233 Bothrops atrox (Fer-de-lance), 302 Bothrops jararaca (Jararaca), 302 Bourke’s Parrot (see Neaphema bourkii) Brachylaemidae, 229, 237 Brachylecithum mosquensis, 238 Brachypteraciidae (Ground-Rollers), 443 bradyzoites, 113, 210, 211 brain, 450 Brant (see Branta bernicla) Branta, 465 Branta canadensis (Canada Goose), 68, 83, 166, 167, 173, 174, 199, 205, 233, 236, 328, 347–348, 348, 355, 356, 362, 389 annual mortality of, 93 infection with Amidostomum sp. in the gizzard of, 369 Branta bernicla (Brant), 69, 250, 328, 356, 362 Branta hutchinsii (Cackling Goose), 69 Branta leucopsis (Barnacle Goose), 208, 328, 356, 362, 389 Branta ruficollis (Red-breasted Goose), 328, 346, 359 Branta sandvicensis (Hawaiian Goose), 217, 328, 358, 389 Brewer’s Blackbird (see Euphagus cyanocephalus) Broad-winged Hawk (see Buteo platypterus) bronchitis, 350 Brotogeris chrysoptera (Golden-winged Parakeet), 399 Brotogeris sanctithomae (Tui Parakeet), 400 Brown Dipper (see Cinclus pallasii) Brown Eared-Pheasant (see Crossoptilon mantchuricum) Brown Pelican (see Pelecanus occidentalis) Brown Snake-Eagle (see Circaetus cinereus) Brown-headed Cowbird (see Molothrus ater) Brown-throated Parakeet (see Aratinga pertinax) Bruneria brunnea (grasshoppers), 402 Brucella abortus vaccine, 199 B-2 (sodium 2,5-dichloro-4-bromophenol), 255 Bubo spp. Bubo africanus (Spotted Eagle-Owl), 133 Bubo bubo (Eurasian Eagle-Owl), 133, 401, 477, 550 Bubo scandiacus (Snowy Owl), 21, 24, 115, 346, 348 Bubo virginianus (Great Horned Owl), 21, 133, 206, 213, 477, 541 Bubulcus ibis (Cattle Egret), 208, 235, 294 Bucconidae (Puffbirds), 61, 443
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Index Bucephala, 115, 465 Bucephala albeola (Bufflehead), 173, 330, 356 Bucephala clangula (Common Goldeneye), 67, 166, 173, 227, 297, 362, 469 Bucephala islandica (Barrow’s Goldeneye), 330, 356 Bucerotidae (Hornbills), 60, 443 Budgerigar (see Melopsittacus undulates) Bufflehead (see Bucephala albeola) buffalo gnat, 537 bumble foot, 527 buparvaquone, 28 Burhinidae (Thick-Knees), 442 Burnished-buff Tanager (see Tangara cayana) bursa of knee, 450 bursate nematodes, 317 Bush birds, 505 Buteo spp., 465 Buteo brachypterus (Modagascar Buzzard), 74 Buteo buteo (Eurasian Buzzard), 74, 114, 131, 205, 208, 346, 471 Buteo galapagoensis (Galapagos Hawk), 516 Buteo jamaicensis (Red-tailed Hawk), 75, 114, 131, 205, 471, 490 Buteo lagopus (Rough-legged Hawk), 75, 346 Buteo lineatus (Red-shouldered Hawk), 75, 131, 205, 237, 471 Buteo nitidus (Gray Hawk), 131 Buteo platypterus (Broad-winged Hawk), 75, 346 Buteo regalis (Ferruginous Hawk), 75 Buteo rufinus (Long-Yegged Buzzgrd), 74 Buteo rufofuscus (Jackal Buzzard), 74 Buteo swainsoni (Swainson’s Hawk), 75 Butorides striata (Striated Heron), 115 Butorides virescens (Green Heron), 294 Cacatua spp. Cacatua ducorpsii (Ducorps’ co*ckatoo), 398 Cacatua leadbeateri, (Pink co*ckatoo), 394 Cacatua sulphurea (Yellow-crested co*ckatoo), 398 Cacatuidae (co*ckatoos), 59, 442 Cackling Goose (see Branta hatehinsii) Cairina, 465 Cairina moschata (Muscovy Duck), 23, 163, 233, 249, 250, 329, 338, 390, 395 Cairina scutulata (White-winged Duck), 329 calcaneal region, 450 Calidris, 465 Calidris alpina (Dunlin), 476 Calidris canutus (Red Knot), 238, 475 Calidris minutilla (Least Sandpiper), 384 Calidris maritima (Purple Sandpiper), 345 California Quail (see Callipepla californica) Callaeidae (Wattlebirds), 445 Callipepla, 465 Callipepla californica (California Quail), 331, 390, 451 Callipepla squamata (Scaled Quail), 395 Calliphora spp., 550, 553 Calliphoridae (bottle flies), 548, 550 Calliphoridae (Protocalliphora spp.), 547 Callonetta leucophrys (Ringed Teal), 329 Calodium, 463 cambendazole, 371 Camnula pellucida (grasshoppers), 402 Campanulotes compar, 519 Campephagidae (Cuckoo-shrikes), 61, 444 Canada Goose (see Branta canadensis) Canadian Sandhill Crane (see Grus canadensis rowani)
canaries (see Serinus canaria) Candida sp., 140 canker, 120 Capella, 465 Cape Shoueter (see Anas smithii) Cape Teal (see Anas capensis) Canvasback (see Aythya volisineria) Cape Barren Goose (see Cereopsis novaehollandiae) capillariasis, 463, 491 Capillaria spp., 140, 463, 464 Capillaria anatis, 370, 463, 465, 466, 482 Capillaria avellari, 468 Capillaria avicola, 481 Capillaria brasiliana, 468 Capillaria cecumitis, 475, 476 Capillaria contorta, 471, 472 Capillaria corvorum, 480 Capillaria droummondi, 470 Capillaria ellisi, 470 Capillaria freitasi, 482 Capillaria fulicae, 475 Capillaria gigantoteca, 470 Capillaria graucalina, 480 Capillaria gymnorhinae, 481 Capillaria herodiae, 468 Capillaria longistriata, 478 Capillaria newzealandica, 477 Capillaria nyrocinarum, 468, 469, 470 Capillaria ornate, 479 Capillaria parvumspinosa, 466 Capillaria phasianiana, 474, 489 Capillaria phasianina, 473 Capillaria pirangae, 483 Capillaria plagiaticia, 476 Capillaria pomatostomi, 483 Capillaria pudendotecta, 470 Capillaria recurvirostrae, 476 Capillaria resectum, 482 Capillaria rigidula, 482, 483 Capillaria similis, 484, 485 Capillaria spinulosa, 468, 469 Capillaria strigis, 477 Capillaria tenuissima, 470, 471, 472, 477, 487–488 Capillaria uropapillata, 474 Capillaria vasi, 476 Capillaria venusta, 478, 479 capillarid nematodes in avian hosts, 466–485 clinical signs, 488 diagnosis, 489–490 domestic animal health concerns, 491 eggs of, 464 epizootiology, 464–488 etiology, 464 female sex bias of, 464 genera of birds infected with cosmopolitan, 465 history, 463–464 host range and distribution, 464 immunity, 490–491 larvae, 464 pathology, 488–489 public health concerns, 491 synonyms, 463 taxonomy and genera, 463 treatment and control, 491 wildlife population impacts, 491 Capillostrongyloides, 463
563
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564 Capitonidae (Barbets), 60, 443 Caprimulgidae (Nightjars), 59, 443 Caprimulgiformes, 14, 59, 163, 443, 478 Caprimulgus macrurus (Large-tailed Nightjar), 478 Carcharhinus perezi, 466 Cardinalidae (Saltators and Cardinals), 60 Cardinalis cardinalis (Northern Cardinal), 112, 116, 327, 333, 378 Cardiofilaria spp., 440, 441–445, 448, 454 Cardiofilaria nilesi, 446, 451, 455 Cardiofilaria pavlovskyi, 455 Carduelis chloris (European Greenfinch), 111, 121, 207, 479 Carduelis tristis (American Goldfinch), 112 Carduiceps clayae, 521 Cariama cristata (Red-legged Seriema), 397, 406 Cariamidae (Seriemas), 442 carnidazole, 144 carotenoid-based coloration, by tit*, 5 Carrion Crow (see Corvus corone), 207 Caryospora, 163 Caryothraustes poliogaster (Black-faced Grosbeak), 333 Caspian Snowco*ck (see Tetroogallus caspius) Casuariidae (Cassowaries), 441, 505 Casuariiformes, 163, 441 Catatropis, 231 Catatropis verrucosa, 226 Cathaemasia, 229 Cathaemasia hians, 235 Cathaemasiidae, 229, 235 Cathartes aura (Turkey Vulture), 79, 208 Cathartidae (New World Vultures), 59, 441 Catharus minimus (Gray-cheeked Thrush), 479 Catreus wallichi (Cheer Pheasant), 390 Cattle Egret (see Bubulcus ibis), 208 cecal coccidiosis (Eimeria colchici), 172 cecal nematode (Trichostrongylus tenuis), 5 cecal strongyle, 316 cecal threadworm, 316 cell-mediated immunity (CMI), 199 cell necrosis, in liver, 21 centrid, 419 Centrocercus urophasianus (Greater Sage-Grouse), 6, 167, 169, 391 Centropus sinensis (Greater Coucal), 401 Cepphus columba (Pigeon Guillemot), 332 Ceratopogonidae, 451 ceratopogonid (Culicoides) flies, 13, 15, 18–19 ceratopogonid vectors, 14 cercariae, 248 Cercomonas gallinae, 121 Cercomonas hepaticum, 134 cerebrospinal (or cerebellar or cerebral) nematodiasis, 424 Cereopsis novaehollandiae (Cape Barren Goose), 356, 389 Ceriodaphnia spp., 355 Certhia familiaris (Eurasian Treecreeper), 207 Certhiidae (Creepers), 60, 444 Cestodes, 4, 261 causing pathology and mortality in wild birds, 264–265 clinical signs, 267 diagnosis, 270 domestic animal health concerns, 271 epizootiology cyclophyllidean eggs, 265–266 Diphyllobothriid life cycles, 267 Tetrabothriidea, 266 etiology, 262–263
Index geographic distribution of, 263–265 history, 262 host range, 263 immunity to infections, 270 infections in abnormal sites, 269 life cycles, 265–266 management implications, 272 pathology of Cyclophyllidea, 267–269, 269–270 Diphyllobothriidae, 269, 270 larval cestodes, 269–270 Tetrabothriidea, 269 public health concerns, 270–271 reproductive systems, 263 synonyms, 262 transmission with aquatic life cycles, 263 treatment and control, 272 wildlife population impacts, 271–272 Cestodiasis, 262 Ceylon Junglefowl (see Gallus lafayetii) Chaffinch (see Fringilla coelebs) Chaimarrornis leucocephalus (White-capped Redstart), 480 Chandlerella spp., 440, 441–445, 445, 446, 448, 452, 454 Chandlerella alii, 449 Chandlerella apusi, 449 Chandlerella bosei, 449 Chandlerella buckleyi, 449 Chandlerella bushi, 449, 457 Chandlerella chitwoodae, 446, 449, 451 Chandlerella columbae, 449 Chandlerella columbigallinae, 449 Chandlerella hepatica, 449 Chandlerella himalayansis, 449 Chandlerella inversa, 449 Chandlerella lerouxi, 449 Chandlerella lienalis, 449 Chandlerella longicaudata, 449 Chandlerella pelecani, 449 Chandlerella petrowi, 450 Chandlerella quiscali, 446, 450, 451, 458 Chandlerella robinsoni, 450 Chandlerella shaldybini, 449 Chandlerella sinensis, 449 Chandlerella singhi, 449 Chandlerella skrjabini, 449 Chandlerella stantchinski, 449 Chandlerella striatospicula, 445, 446, 449, 451 Chandlerella sultana, 449 Chandlerella thapari, 449 Changeable Hawk-Eagle (see Spizaetus cirrhatus) Channel-billed Toucan (see Ramphastos vitellinus) Charadriidae (Plovers and Lapwings), 59, 361, 442, 505 Charadriiformes, 14, 59, 121, 163, 175, 205, 208, 238, 247, 251, 253, 263, 291, 376, 377, 435, 465, 475, 521 Charadrius, 465 Charadrius alexandrinus (Snowy Plover), 331 Charadrius hiaticula (Common Ringed Plover), 475 Charadrius vociferus (Killdeer), 476 Chaunocephalus, 230 Chaunocephalus ferox, 235 Cheer Pheasant (see Catreus wallichi) Cheilospirura nasuta, 327 chemotherapy, 285 Chen spp., 465 Chen caerulescens (Snow Goose), 69, 167, 173, 174, 348, 355, 359, 365, 390
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Index Chen caesulescens caerulescens (Lesser Snow Goose), 167, 174 Chen canagica (Emperor Goose), 69, 328, 363 Chen rossii (Ross’ Goose), 173, 355, 359, 365, 390 Chenonetta jubata (Maned Duck), 390 Chestnut-cheeked Starling (see Sturnia philippensis) Chestnut Teal (see Anas castanea) chewing lice clinical signs, 518–520 diagnosis, 520 epizootiology, 518 etiology, 516–518 history, 515 host defense and immunity, 520–522 host range and distribution, 515–516 pathogenesis and pathology, 518–520 public and domestic health concerns, 522 synonyms, 515 treatment and control, 522 wildlife population impacts, 522–523 Chinese Francolin (see Francolinus pintadeanus) chicken, domestic, infections, 54 chickens, with infections of Eimeria, 163 Chilean Flamingo (see Phoenicopterus chilensis), 328, 338 Chiloe Wigeon (see Anas sibilatrix) Chimango Caracara (see milvago chimango) Chinese Goshawk, 78 Chinese Pond-heron (Ardeola bacchus) Chionididae, 442 Chlamydotis undulate (Houbara Bustard), 261, 393 Chlidonias, 465 Chlidonias hybrida (Whiskered Tern), 476 Chloebia gouldiae (Gouldian Finch), 195 Chloephaga picta (Upland Goose), 360, 390 Chloephaga poliocephala (Ashy-headed Goose), 356, 389 Chloropseidae (Leafbirds), 60 chloroquine phosphate, 48 chloroquine sulfate, 28 Choanotaenia, 264 Choanotaenia infundibulum, 264, 268, 269, 271 Chrysolophus, 465 Chrysolophus amherstiae (Lady Amherst’s Pheasant), 392 Chrysolophus pictus (Golden Pheasant), 331, 391, 473 Chtonobdella bilineata, 505 Chtonobdella limbata, 505 Chukar (see Alectoris chuckar) Cicinnurus magnificus (Magnificent Bird-of-paradise), 345 Ciconia spp. Ciconia ciconia (white short), 235, 550 Ciconia maguari (Maguori Stork), 295, 389 Ciconia nigra (Black Stork), 235 Ciconiidae (Storks), 59, 441 Ciconiiformes, 14, 35–36, 59, 163, 208, 235, 247, 251, 253, 293, 306, 328, 441, 465, 468 Cinclidae (Dippers), 61, 444 Cinclus pallasii (Brown Dipper), 479 Cinereous Vulture (see Aegypius monachus) Cinnamon Teal (see Anas cyanoptera) cintrin, 371 Circaetus cinereus (Brown Snake-Eagle), 75 Circaetus gallicus (Short-toed Eagle), 74, 471 Circulatory system, 449 Circus spp. Circus aeruginosus (Western Marsh-Harrier), 79, 130, 237, 346, 471 Circus cyaneus (Northern Harrier), 78, 237, 346
565
Circus macrourus (Pallid Harrier), 78, 205 Circus pygargus (Montagu’s Harrier), 78, 471 Circus spilonotus (Eastern Marsh-Harrier), 395 Cladotaenia, 265 Clangula, 465 Clangula hyemalis (Long-tailed Duck), 66, 66 173, 227, 227, 267, 297, 330, 358, 364, 469 Clay-colored Robin (see Turdus grayi), 208 Clelia clelia (Mussurana), 303 Climactreridae (Australian Treecreepers), 61, 444 Clinostomidae, 229 Clinostominae, 235 Clinostomum, 229 Clinostomum attentuatum, 235 Clinostomum complanatum, 226 cloaca (Leucochloridium), 226 Cloacitrema, 231 Cloacotaenia megalops, 264, 269 clopidol, 177 clorsulon, 424 Clostridium perfringens, 157, 170 Cnephia ornithophilia, 83, 85, 543 Coal Tit (see Periparus ater), 376 Coccidea, 57 Coatinga Parakeet (see Aratinga cactorum) Coccidia, 162, 164 renal, 173, 175, 176 coccidian parasite, 204 coccidiasis, 162 coccidiosis, 3, 108, 111, 116, 165, 181 control of, 172 diagnosis, 170 renal, 175 coccidiostat, 192–193 Coccidium truncatum, 162 Coccothraustes vespertinus (Evening Grosbeak) 109, 527 co*ckatiel (see Nymphicus hollandicus) Coco, Heron (see Ardea cocoi) Cochlearius, 465 cocoons of Glossiphoniidae, 503 Coerulea coerulea (“purple sugarbird”), 376 Colaptes auratus (Northern Flicker), 332, 478 Colaptes maximus (see Piculus rubiginosus) Coliidae (Mousebirds), 59, 443 Coliiformes, 14, 59, 163, 443 Colinus, 465 Colinus virginianus (Northern Bobwhite), 13, 21, 23, 36, 155, 156, 163, 167, 169, 195, 213, 280, 316, 320–321, 327, 331, 378, 388, 390, 395, 426, 487 Colius erythromelon (Red-faced Mouse Bird), 114 Collyriclum, 231 Collyriculum faba, 226, 238 Colpocephalum holzenthali, 517 Colpocephalum turbinatum, 516, 521 Coluber constrictor (Blue Racer), 302 Columba spp., 168, 465 Columba arquatrix (Rameron Pigeon), 70, 398 Columba eversmanni (Pale-backed Pigeon), 70 Columba guinea (Speckled Pigeon), 70 Columba larvata (Lemon Dove), 70 Columba livia (Rock Pigeon), 13, 22, 70, 113, 120, 122, 139, 168, 172, 204, 205, 208, 327, 332, 381, 398, 402, 464, 488, 501, 519, 520, 534 Columba oenas (Stock Dove), 70 Columba palumbus (Common Wood Pigeon), 20, 22, 70, 124, 205, 398 Columba vitiensis (Metallic Pigeon), 70
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566
October 16, 2008
9:20
Index
Columbicola columbae, 517, 519 Columbidae (Pigeons and Doves), 57, 59, 361, 435, 442 Columbiformes, 14, 18, 22, 23, 59, 163, 168, 169, 205, 208, 217, 238, 253, 435, 442, 465, 476, 548 Columbiformes, 114 Columbina spp. Columbina picui (Picui Ground-Dove), 398 Columbina inca (Inca Dove), 129, 144 Columbina passerina (Common Ground-Dove), 129 Columbina talpacoti (Ruddy Ground Dove), 71, 206, 208, 238, 398 Comb Duck (see Sarkidiornis melanotas) Common Crane (see Grus grus) Common Cuckoo (see Cuculus canorus) Common Eider (see Somateria mollissima) Common Goldeneye (see Bucephala clangula) Common Grackle (see Quiscalus quiscula) Common Greenshark (see Tringa nebularia) Common Ground-Dove (see Columbina passerina) Common Hill Myna (see Gracub religiosa) Common Loon (see Gavia immer) Common Merganser (see Mergus merganser) Common Moorhen (see Gallinula chlosopus) Common Murre (see Uria aalge) Common Nightingale (see Luscinia megarhynchos) Common Pochard (see Aythya ferina) Common Quail (see Coturnix coturnix) Common Raven (see Corvus corax) Common Redshark (see Tringa totonus) Common Ringed Plover (see Charadrius hiaticula) Common Sandpiper (see Actitis hypoleucos) Common Shelduck (see Tadorna tadorna) Common Snipe (see Gallmago gallinago) Common Swift (see Apus apus) Common Tern (see Sterna hirundo) Common Wood Pigeon (see Columba palumbus) connective tissues, 449 Conopophagidae (Gnateaters), 443 contracaeciasis, 413 Contracaecum spp., 339, 420 caudal features of diagnostic interest in adult male, 420 clinical signs, 421 Contracaecum bioccai, 420 Contracaecum magnipaillatum, 416 Contracaecum magnipapillatum, 416, 419, 420 Contracaecum microcephphalum, 420 Contracaecum micropapillatum, 420 Contracaecum multipapillatum, 416, 418, 420, 423, 424 Contracaecum multipapillatum sensu lato, 423 Contracaecum ogmorhini, 415 Contracaecum ogmorhini sensu lato, 420 Contracaecum osculatum, 413 Contracaecum pelagicum, 416, 420 Contracaecum radiatum, 420 Contracaecum rudolphii, 415, 416, 418 Contracaecum rudolphii sensu lato, 416, 418, 420, 421 Contracaecum septentrionale, 416, 420 Contracaecum spiculigerum, 424 Contracaecum tricuspe, 420 Contracaecum variegatum, 414, 416 Contracaecum variegatum sensu lato, 420 diagnosis, 419–421 domestic animal health concerns, 423 epizootiology, 416–419 etiology, 413–416 host range and distribution, 416
host range of species of, 417 immunity, 421–423 lesions caused by, 421 pathogenesis and pathology, 421 public health concerns, 423 synonyms, 413 treatment and control, 424 wildlife population impacts, 424 Cooper’s Hawk (see Accipites cooperii) copepods, 385 Copperhead (Agkistrodon contortrix), 302 Copper Pheasant (see Syrmaticus soemmerringii) copper sulfate, 255, 340 Coracias garrulus (European Roller), 332 Coraciidae (Rollers), 60, 443 Coraciiformes (Kingfishers), 14, 60, 163, 238, 252, 332, 377, 443, 548 Coracina novaehollandiae (Black-faced Cuckoo-shrike), 480 Coragyps atratus (Block Vulture), 79 Corcoracidae (Choughs and Apostlebirds), 61 cormorants (see Phalacrocorax spp.) Cortrema, 229 Cortrematidae, 229 Corvidae (Crows, Jays, and Magpies), 60, 435, 445, 505 Corvus spp., 263, 465 Corvus brachyrhynchos (American Crow), 207, 209, 327, 333, 450, 451, 454, 480 Corvus corax (Common Raven), 480 Corvus cornix (Hooded Crow), 501 Corvus corone (Carrion Crow), 207, 226, 345, 480 Corvus frugilegus (Rook), 207, 345, 346, 347, 480 Corvus hawaiiensis (Hawaiian Crow), 42–43, 204, 218 Corvus monedula (Eurasian Jackdaw), 207, 345, 346, 480, 490 Corvus nasicus (Cuban Crow), 333 Corvus ossifragus (Fish Crow), 110 Corynosoma constrictum, life cycle of, 279, 280 Corynosoma species, 277, 279 costs to host, against parasitism, 4, 6 Cotingidae (Cotingas), 443 Cotton Pygmy-goose (see Nettapus coromandelianus) Coturnix, 465 Coturnix coturnix (Common Quail), 331, 391, 396 Coturnix japonica (Japanese Quail), 155, 156, 169, 196, 213 Cotylurus, 231 coughing, 197 counter-immunoelectrophoresis, 92 Cracidae (Guans, Chachalacas, and Curassows), 59, 442 Cracticidae (Bellmagpies), 61, 445, 505 Crax blumenbachii (Red-billed Curassow), 390 Creatophora cinerea (Wattled starling), 111 Crested Auklet, (see Aethia cristatella) Crested Fireback (see Lophura ignita) Crested Flycatcher (Myiarchus crinitus), 528 Crested Goshawk (see Accipiter trivirgatus) Crested Guineafowl (see Guttera pucherani) Crested Pigeon (Geophaps lophotes), 70 Crested Partridge (see Rollulus rouloul) Crickets (Acheta domestica), 378 Crimson-backed Tanager (see Rampholcelus dimidiatus), 209 Crimson Rosella (see Platycercus elegans) crisis period, 40 Crocodylocapillaria, 463 cropworm, 463
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Index Crossoptilon, 465 Crossoptilon auritum (Blue Eared-Pheasant), 390 Crossoptilon mantchuricum (Brown Eared-Pheasant), 390 Crotophaga ani (Smooth-billed Ani), 332 crows (see Corvus spp) crustaceans, 340 Cryptocotyle, 230 Cryptocotyle lingua, 237 cryptosporidiosis, 197, 198, 200 Cryptosporidium spp., 163 clinical signs, 197–198 Cryptosporidium andersoni, 195 Cryptosporidium anserinum, 195 Cryptosporidium baileyi, 195 in chickens, 199 infections, in humans, 200 life cycles of, 196 Cryptosporidium blagburni, 195, 199 Cryptosporidium galli, 195 Cryptosporidium meleagridis, 195, 199 infections, in humans, 200 life cycles of, 196 Cryptosporidium muris, 195 Cryptosporidium parvum, 195 Cryptosporidium tyzzeri, 195 diagnosis, 198–199 domestic animal health concerns, 200 epizootiology, 196–197 etiology and host range, 195–196 history, 195 immunity, 199 pathology, 198 public health concerns, 199–200 synonym, 195 treatment and control, 200 wildlife population implications, 200 cryptozoites, 39 Crypturellus spp. Crypturellus noctivagus (Yellow-legged Tinamou), 389 Crypturellus parvirostris (Small-billed Tinamou), 389 Crypturellus tataupa (Totaupa Tinamou), 389 Crypturellus undulates (Undulated Tinamou), 389 Crypturellus variegates (Variegated Tinamou), 389 Crytocotyle lingua, 239 Ctenodactylus gundi, 204 Cuban Parrot (see Amazona leucacephala) Cuclotogaster heterographus, 518 Cuculidae (Cuckoos), 59, 442 Cuculiformes, 14, 59, 114, 163, 332, 442 Cuculus canorus (Common Cuckoo), 401 Culex spp. Culex annulus, 41 Culex nigripalpus, 40, 41 Culex pipiens, 41 Culex quinquefasciatus, 40, 41 Culex restuans, 41 Culex saltanensis, 41 Culex sitiens, 41 Culex stigmatasoma, 40 Culex stimatosoma, 41 Culex tarsalis, 40, 41 Culicidae, 451 culicine species, 40 Culicoides spp. Culicoides arakawae, 85 Culicoides arboricola, 18
567
Culicoides bottimeri, 17, 18 Culicoides circ*mscriptus, 85 Culicoides crepuscularis, 18 Culicoides downesi, 18 Culicoides edeni, 18 Culicoides haematopotus, 18 Culicoides hinmani, 18 Culicoides impunctatus, 18 Culicoides knowltoni, 18 Culicoides nubeculosus, 18 Culicoides odibilis, 85 Culicoides-proof facilities, 28 Culicoides schultzei, 85 Culicoides species, 17, 82 Culicoides sphagnumensis, 18 Culicoides stilobezziodes, 18 Culiseta morsitans, 41 Curatrem, 424 Curonian Spit, 86 Cuterebra buccata, 550 Cuterebridae (bot flies), 550 Cyanerpes caeruleus (Purple Honeycreeper), 111, 377 Cyanerpes cyaneus (Red-legged Horeycreeper), 111 Cyanocitta, 465 Cyanocitta cristata (Blue Jay), 19, 21, 22, 327, 333, 454, 489 Cyanocorax yncas (Green Jay), 24 Cyanopica cyanus (Azure-winged Magpie), 474, 481 Cyanoramphus auriceps (Yellow-fronted Parakeet), 218, 401 Cyanoramphus novaezelandiae (Red-fronted Parakeet), 218 Cyathocotyle, 229 Cyathocotyle bushiensis (Cyclocoelidae), 225, 226, 227, 234, 235, 240 Cyathocotylidae, 229, 235 Cyathostoma spp., 351 Cyathostoma americanum, 351 Cyathostoma bronchialis, 344, 347, 349, 350, 351 Cyathostoma (cf. Syngamus spp.), 343, 346 Cyathostoma (Cyathostoma) spp., 343 Cyathostoma (Hovorkonema) spp., 343 Cyathostoma (Hovorkonema) variegatum, 343, 351 Cyathostoma infections, 348 Cyathostoma lari, 343 cyathostome infections, 352 cyathostomes, 344, 349–351 cyathostomiasis, 343 Cyclocoelidae, 229, 235, 236, 238 Cyclocoelum, 229 Cyclocoelum mutabile, 225, 238 Cyclophyllidea, 262, 269–270 cyclophyllidean and diphyllobothriid cestodes, life cycles of, 264 cyclophyllideans, 265–266 cyclophyllidean scolices, 262 Cyclophyllidean sp., 264 cyclophyllidean species, 262 Cyclops sternuus, 384 Cygnus spp. Cygnus atratus (Black Swan), 328, 356, 389, 470 Cygnus buccinator (Trumpeter Swan), 68, 328, 336, 359, 368 Cygnus columbianus (Tundra Swan), 68, 227, 236, 328, 360, 365, 451, 452, 470, 521 Cygnus cygnus (Whooper Swan), 227, 236, 328, 365, 360, 390
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568 Cygnus spp. (Cont.) Cygnus melancoryphus (Black-necked Swan), 328, 362, 470 Cygnus olor (Mute Swan), 68, 227, 233, 236, 282, 328, 358, 364, 470, 521 Cypsiurus parvus (Palm-Swift), 528 cystacanth, 279 cystic and nodular mite disease, 527 cytochrome c oxidase I gene, 239 cytostome, 39 Dabbling Ducks (Anas spp.), 115, 236, 367 Dacnis cayana (Blue Dacnis), 111 Daphnia species, 335–336 Dark Chanting Goshawk (see Meliera metabates) Dark-throoted Thrush (see Turdus rificollis) Darter (see Anhinga melanogaster) Davaineidae, 262, 264 decapods, 279 Deegeriella regalis, 516, 521 defense cost, to the host against parasites, 4 Delichon urbicum (House Martin), 7 Demoiselle Crane (see Anthropoides virgo) Dendragapus fuliginosus (Sooty Grouse), 393 Dendragapus obscurus (Dusky Grouse), 331, 390, 396 Dendritobilharzia, 246, 247, 251, 255 Dendritobilharzia larvae, 247 Dendritobilharzia pulverrulenta, 247, 249, 250, 251 Dendrobaena adaiensis, 486 Dendrobaena octaedra, 486 Dendrocolaptidae (woodcreepers), 435, 443 Dendrocopos major (Great Spotted Woodpecker), 478 Dendrocygna spp. Dendrocygna arborea (West Indian Whistling-Duck), 328, 390 Dendrocygna autumnalis (Black-bellied Whistling-Duck), 356, 362 Dendrocygna bicolor (Fulvous Whistling-Duck), 83, 328, 363 Dendrocygna viduata (White-faced Whistling-Duck), 328 Dendrodrilus rubidus, 486 Dendroica spp. Dendroica dominica (Yellow-throated Warbler), 481 Dendroica discolor (Prairie Warbler), 333 Dendroica fusca (Blackburnian Warbler), 333, 481 Dennyus hirundinis, 521 dermanyssid mites, 534 Dermatobia hominis, 550 Dessetfilaria, 441–445, 448, 454 Dessetfilaria guianensis, 456 diarrhea, 87, 198 Dicaeidae (Flowerpickers), 61, 444 Dicheilonematinae, 435 diclazuril, 200, 219 Dicranotaenia coronula, 264 Dicrocoeliidae, 226, 229, 237, 238 Dicruridae (Drongos), 61, 445 digenetic trematode, life cycle of, 228 Dilepididae, 262, 264 dimetridazole, 144–145 Dinobdella ferox, 505 Dioctophyma renale, 308 Diomedeidae (Albatrosses), 441 Diphyllobothriidae, 262, 264, 270 Diphyllobothriid cestodes, 271 Diphyllobothriid life cycles, 267
Index Diphyllobothrium species, 267, 271 Diplostomatidae, 229 Diplostomum, 229 Diplotriaena, 435 clinical signs, 436 diagnosis, 437 Diplotriaena ozouzi, 435 epizootiology, 435–436 etiology, 435 history, 434 host range and distribution, 434–435 pathology, 436 public and domestic animal health concerns, 437 treatment and control, 437 wildlife population impacts, 437 Diplotriaenidae, 435 Diplotriaenoidea, 440 diplotriaenoids, 439 Dirofilariinae, 448 dispharagosis, 326 Dispharagus spp. Dispharagus crassicauda, 337 Dispharagus nasutus, 327 Dispharagus spiralis, 327 Dispharagus spiralis columbae, 327 Dispharagus tentaculatus, 327 Dispharagus uncinatus, 335 dispharyngosis, 326 Dispharynx, 326 clinical signs, 327–334 diagnosis, 334–335 Dispharynx nasuta, 326, 327–328, 334–335, 339, 380 Dispharynx spiralis, 326 Dispharynx stonae, 327 domestic animal health concerns, 329 epizootiology, 327 etiology, 327 history, 326–327 host range and distribution, 326, 328–333 immunity, 335 pathogenesis and pathology, 334 synonym, 326 treatment and control, 339–340 dispharynxiasis, 326, 339 Disseminated visceral coccidiosis (DVC) clinical signs, 190 diagnosis, 191–192 domestic animal health concerns, 192 epizootiology, 187–190 etiology, 181–187 history, 181 host range, 181 immunity, 192 management impacts, 193 pathogenesis and pathology, 190–191 public health concerns, 192 species of Eimeria associated with, 181 synonyms, 181 treatment and control, 192–193 wildlife population impacts, 192 diving ducks (Aythyini), 367 DNA vaccines, based on the circ*msporozoite protein, 48 domestic goose (see Anser anser) doves, infected with Leucocytozoon marchouxi, 87, 89, 93 Double-crested Cormosant (see Phalacrocosax auritus) Double-spurred Francolin (see Francolinus bicalcoratus) Dracunculus, 384
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Index Drepanididae (Hawaiian honeycreepers), 444 drepanidotaeniasis, 262 Dromadidae (Crab Plovers), 442 Dromaiidae (Emus), 441 Dromaius novaehollandiae (Emu), 427, 459, 452 droppings, of birds, post malarial infection, 43 Dryadophis bifossatus (Cazadora brown-lined snake), 303 Drymarchon corais (Indigo snake), 302 Dryocopus pileatus (Pileated Woodpecker), 478 Ducorp’s co*ckatoo (see Cacatua ducorpsii) Ducula spp. Ducula bicolor (Pied Imperial-Pigeon), 217 Ducula carola (Spotted Imperial-Pigeon), 71 Ducula spilorrhoa (Torresian Imperial-Pigeon), 217 dulfaquinoxaline, 177 Dulidae (Palmchats), 444 Dumetella carolinensis (Gray Catbird), 332 Dunlin (see Calidris alpina) Dunnock (see Prunella modularis) duokvin, 200 Duplicaecum, 413 Dusky Parrot (see Pionus fuscus) Dusky Turtle-Dove (see Streptopelia lugens) dusting, of birds, 28 dwarf mallards, 252 dyspnea, 137, 197, 349 Eared Dove (see Zenaida auriculata) Eared Grebe (see Podiceps nigricollis) earthworms, 352 Eastern Bluebird (see Sialia sialis) Eastern equine encephalitis, 521 Eastern Marsh-Harrier (see Circus spilonotus) Eastern Meadowlark (see Sturnella magna) Eastern mosquitofish (Gambusia holbrooki), 290 Eastern Rosella (see Platycercus eximius) Eastern Screech-Owl (Megascops asio), 213 Eastern Towhee (see Pipilo esythrophthalmus) Echinochasmus, 230 Echinocoleus, 463 Echinocoleus cyanopicae, 474, 481 Echinoparyphium, 230 Echinoparyphium recurvatum, 236 Echinosotoma spp., 230, 234, 236 Echinosotoma trivolvis, 236 Echinostoma revolutum, 226 Echinostoma trivolvis, 234 Echinostomatidae, 230, 235, 236 Echinostoma trivolvis, 234 echinostomiasis (Echinostoma spp.), 239 echinuriasis, 337 Echinuria spp. clinical signs, 336 diagnosis, 337 domestic animal health concerns, 339 Echinuria jugadornata, 335 Echinuria uncinata, 326, 327, 328, 335, 336, 337 epizootiology, 335–336 etiology, 335 history, 335 host range and distribution, 326, 328–333 pathogenesis and pathology, 336–337 synonyms, 335 treatment and control, 339–340 ectoparasitic hippoboscid flies, 13, 17 ectoparasitic mites, 534
Egretta spp. Egretta tricolor (Tricolored Heron), 294, 305 Egretta caerulea (Little Blue Heron), 115, 468 Egretta garzetta (Little Egret), 114, 294, 395 Egretta thula (Snowy Egret), 294, 304, 305, 384 Egyptian Goose (Alapochen aegyptiaca) Egyptian Vulture (see Neaphron percnoptesus) Eiberg strains, 135 Eimeria spp., 108, 166, 168, 174 common pathogenic intestinal species of, 166–168 in cranes, life cycle of, 187 developmental stages of, 164 Eimeria acervulina, 169 Eimeria adenoeides, 167, 169, 171 Eimeria angusta, 167, 169 Eimeria anseris, 171 Eimeria auritusi, 173, 174 Eimeria avium, 162 Eimeria aythyae, 165, 166 Eimeria boschadis, 173 Eimeria bosquei, 187 Eimeria brunetti, 169 Eimeria bucephalae, 166 Eimeria colchici, 168, 169, 170 Eimeria columbarum, 168 Eimeria dispersa, 163, 167, 171 Eimeria duodenalis, 168, 169 Eimeria dusingi, 168 Eimeria fraterculae, 175 Eimeria gallopavonis, 167, 169, 171 Eimeria goelandi, 175 Eimeria gruis, 162, 181, 186 Eimeria gruis, oocysts of, 186 Eimeria innocua, 169 Eimeria kotlani, 171 Eimeria labbeana, 168 Eimeria lettyae, 167, 169 Eimeria maxima, 169 Eimeria meleagridis, 169 Eimeria meleagrimitis, 167, 169, 171 Eimeria mitis, 169 Eimeria mivati, 169 Eimeria mulardi, 163 Eimeria necatrix, 169 Eimeria nocens, 171 Eimeria phasiani, 168, 169, 170 Eimeria praecox, 169 Eimeria procera, 168 Eimeria reichenowi, 162, 181, 186 Eimeria reichenowi, oocysts of, 186 Eimeria saitamae (Domestic Ducks), 171 Eimeria serventyi, 174 Eimeria somateriae, 165, 173, 174 Eimeria stigmosa, 167 Eimeria subrotunda, 169 Eimeria taldykurganica, 169 Eimeria tenella, 169 Eimeria tropicalis, 187 Eimeria truncata, 174, 175, 176, 177 Eimeria tsunodai, 169 Eimeria uzura, 169 Eimeria wobeseri, 175 eported from avian hosts, 163 hepatic, 177 infection development, 162 intestinal clinical signs of, 170
569
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570 Eimeria spp. (Cont.) diagnosis of, 170–171 distribution and host range of, 162–163 domestic animal concerns of, 171–172 epizootiology of, 164–170 etiology of, 163–164 immunity of, 170–171 management implications, 172 pathology of, 170 public health concerns of, 171 treatment and control, 172 wildlife population impacts, 172 renal clinical signs and pathology, 175 diagnosis, 176 domestic animal health concerns, 177 epizootology, 173 etiology of, 172 host range and prevalence, 173–175 immunity, 176 management implications, 177 public health concerns, 176 treatment and control, 177 wildlife population impacts, 177 sporulated oocyst of, 164 Eimeriidae, 163 Eisenia foetida, 344, 401, 486 Elanus caeruleus (Black-shouldered Kite), 74 Elegant Parrot (see Neophema elegans) Eleonora’s Falcon (see Falco eleonorae) Elimia virginica, 227 ELISA test, 26, 46, 92 Emberiza spp. Emberiza citrinella (Yellowhammer), 206, 401, 481 Emberiza schoeniclus (Reed Bunting), 481 Emberiza sulphurata (Yellow Bunting), 481 Emberiza variabilis (Gray Bunting), 481 Emberizidae (Buntings and Sparrows), 60, 444 embryonation, 401 Emerald-spotted Wood-Dove (see Turtur chalcospilos), 73 Emperor Goose (see Chen canagica) Empidonax minimus (Least Flycatcher), 332 Emu (see Dranaius nevaehoilandiae), 459 Emys blandingi (Blandings turtle), 303 Encephalitis, 521 endopolygony, 113 energy, as a parameter to parasitism, 4 Enicognathus ferrugineus (Austral Parakeet), 399 enrofloxacin, 200 Eomenacanthus stramineus, 521 Eos bornea (Red Lorry), 217 Eos cyanogenia (Black-winged Lorry), 218 eosinophils, 21 epithelial hyperplasia, 21 epizootics, 27 from avian malaria, 35 of eustrongylidosis, 289 at lower elevations, 48 of trematodes, 225 epomidiostomiasis, 355 Epomidiostomum spp., 369 avian hosts infected with, 362–365 clinical signs, 368 diagnosis, 370 domestic animal health concerns, 370 eggs, 367 epizootiology, 367–368
Index Epomidiostomum alii, 364, 366 Epomidiostomum asymmetricum, 362, 366 Epomidiostomum crami, 355, 362, 363, 364, 365, 366, 368, 369 Epomidiostomum cygni, 364, 366 Epomidiostomum orispinum, 362, 363, 364, 365, 366 Epomidiostomum penelopi, 362, 366 Epomidiostomum petalum, 364, 365, 366 Epomidiostomum querquedulae, 363, 364, 365, 366 Epomidiostomum sarkidiorni, 362, 366 Epomidiostomum serratum, 363, 364, 365, 366 Epomidiostomum skrjabini, 362, 363, 364 Epomidiostomum subquadratum, 363, 365, 366 Epomidiostomum sultanai, 362, 363, 366 Epomidiostomum uncinatum, 362, 363, 364, 365, 366, 368 Epomidiostomum vogelsangi, 362, 366 etiology, 366–367 history, 355 host range and distribution, 355–366 immunity, 370 management implications, 371 pathogenesis and pathology, 368–370 prevalence, intensity and abundance, 367–368 public health concerns, 370 treatment and control, 371 wildlife population impacts, 371 Erckel’s Francolin (see Francolinus erckelii) Erithacus, 465 Erithacus rubecula (European Robin), 22, 207 erythrocytes invasion, 15, 42 erythrocytic meronts, 46 erythrocytic parasitemias, 21 erythrocytic stages, of the parasites, 21 Escherichia coli, 157, 521 Escherichia intermedia, 157 Estrildidae (Waxbills), 60, 444 Eucoccidiorida, 163 Eucoleus spp., 463 Eucoleus annulatus, 463, 464, 465, 466, 487, 488 Eucoleus contortus, 463, 464, 465, 466, 472, 486–487, 487–488, 489 eggs of, 487 stichosome, 486 Eucoleus crypturi, 466 Eucoleus dispar, 470, 471, 472, 477, 479, 480, 482, 484, 487–488, 489 Eucoleus frugilegi, 464 Eucoleus obtusiuscula, 475, 476 Eucoleus penidoi, 466 Eucoleus perforans, 464, 468, 473, 474, 488 Eucoleus suppereri, 472 Eucotyle, 230 Eucotyle nephritica (Eucotylidae), 226 Eucotylidae, 230, 237, 238 Eucyclops serratulus, 384, 385 Eudocimus, 465 Eudocimus albus (White Ibis), 115, 285, 290, 296 Eudyptula minor (Little Penguin), 174, 175, 217, 235, 267 Eufilaria spp., 441–445, 448, 454 Eufilaria bartlettae, 451 Eufilaria kalifai, 451 Eufilaria longicaudata, 214, 445, 446, 456 Eufilari longicaudata, 451 Eulimdana spp., 441–445, 448, 454 Eulimdana bainae, 448, 451, 521
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Index Eulimdana clava, 450, 452 Eulimdana cypseli, 446, 451 Eulimdana metcalforum, 456 Eulimdana wongae, 448, 451, 521 Eumegacetes, 230 Eumegacetidae, 230 Eunectes murinus (Green anaconda), 302 Euphagus cyanocephalus (Brewer’s Blackbird), 209 Euphonia violacea (Violaceous Euphonia), 111 Eupodotis ruficrista (Red-crested Bustard), 269 Eurasian Bullfinch (see Pyrrhula pyrrhula) Eurasian Buzzard (see Buteo buteo) Eurasian Capercaillie (see Tetrao urogallus) Eurasian Collared Dove (see Streptopelia decaocto) Eurasian Coots (see Fulica atra) Eurasian Eagle-Owl (see Bubo bubo) Eurasian Hobby (see Falco subbuteo) Eurasian Jackdaw (see Corvus monedula) Eurasian Jay (see Garrulus glandarius) Eurasian Kestrel (see Falco tinnunculus) Eurasian Magpie (see Pica pica pica) Eurasian Nutcracker (see Nucifraga caryocatactes) Eurasian Nuthatch (see Sitta europaea) Eurasian Oystercatcher (see Haematopotus ostralegus) Eurasian Pygmy-Owl (see Glaucidium passerinum) Eurasian Skylark, (see Alauda arvensis) Eurasian Sparrowhawk (see Accipiter nisus) Eurasian Spoonbill (see Platalea leucorodia) Eurasian Teal (see Anas creccai) Eurasian Treecreeper (see Certhia familioris) Eurasian Tree Sparrow (see Passer montanus) Eurasian Turtle-Dove (see Streptopelia turtur) Eurasian Wigeon (see Anas penelope) European Greenfinch (see Carduelis chloris) European Herring Gull (see Larus argentatus) European Honey-Buzzard (see Pernis apivorus) European Robin (see Erithacus rubecula) European Roller (see Coracias garrulus) European Scops-Owl (see Otus scops) European Shag (see Phalacrocorax aristotelis) European Starling (see Sturais vulgaris) Eurylaimidae (Broadbills), 61, 443 Eurypygidae (Sunbittern), 442 Eusimulium geneculare, 85 Eustrongylides spp., 289, 296, 297 avian hosts of, 291–297 diagnostic features of the caudal end of male, 299 diagnostic features of the cranial end of, 299 distribution of, 290 eggs of, 304 environmental factors affecting transmission of infections, 301–304 Eustrongylides acrochordi, 298 Eustrongylides africanus, 298 Eustrongylides excisus, 289, 292, 296, 298, 300, 301, 302, 304, 306, 309 Eustrongylides gadopsis, 298 Eustrongylides ignotus, 289–290, 292–295, 296, 298, 300, 302, 303, 304, 309 egg of, 309 with oligochaete, 301 without oligochaete, 301 Eustrongylides indicus, 298 Eustrongylides mergorum, 298 Eustrongylides papillosus, 298 Eustrongylides phalacrocoracis, 298 Eustrongylides plotinus, 298
571
Eustrongylides rubrum, 298 Eustrongylides sinicus, 298 Eustrongylides spinispiculum, 298 Eustrongylides tricolor, 298 Eustrongylides tubifex, 289–290, 291–293, 297, 298, 300, 301, 303, 306, 307, 309 Eustrongylides wenrichi, 298 larval of, 301 lesions of infections, 298, 305–307 life cycle, 300 life cycle of, 300 life history of, 298–301 paratenic and accidental hosts of, 302–303 prevalence in infected fish, 304, 308 range of development times in days, 301 range of fish intermediate hosts of, 304 Eustrongylidiasis, 289 Eustrongylidosis, 289, 308 clinical signs, 304–305 diagnosis, 306–308 domestic animal health concerns, 308 epizootiology, 298–304 etiology, 298 history, 289 host range and distribution, 289–298 management implications, 309–310 pathology, 305–306 public health concerns, 308 synonyms, 289 treatment and control, 309 wildlife population impacts, 308–309 Evening Grosbeak (see Coccothraustes vespertinus) evolutionary trap, 7 exflagellation process, 40 exoerythrocytic merogony, of Plasmodium elongatum, 35 exoerythrocytic meronts, 39, 40 exoerythrocytic parasites, 42 exsheathment process, 320 extrinsic factors, of parasites, 21 eye infection, with Philophthalmus sp., 239 Fabricii (Prosthogonimus), 226 Falcated Duck (see Anas falcata) Falco spp. Falco biarmicus (Lanner Falcon), 132 Falco cenchroides (Australian Kestrel), 80 Falco cherrug (Saker Falcon), 80, 132, 139 Falco chicquera (Red-necked Falcon), 132, 488 Falco columbarius (Merlin), 79, 132, 148 Falco eleonorae (Eleonora’s Falcon), 80 Falco mexicanus (Prairie Falcon), 236, 434 Falco naumanni (Lesser Kestrel), 80, 131 Falco peregrinus (Peregrine Falcon), 79, 132, 143, 472, 488, 550 Falco rupicoloides (Greater Kestrel), 80 Falco rusticolus (Gyrfalcon), 132, 237, 488 Falco sparverius (American Kestrel), 28, 80, 205, 472, 550 Falco subbuteo (Eurasian Hobby), 80, 472 Falco tinnunculus (Eurasian Kestrel), 28, 79, 114, 131, 205, 346, 424, 434, 472 Falconidae (Falcons and Caracaras), 57, 59, 441 Falconiformes, 14, 59, 130, 163, 205, 208, 236, 297, 441, 465, 470, 548, 549 Fallisia neotropicalis, 36 Fasciola, 225 Fasciola trachea, 343
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572 faucet snail, 226 feather lice, 517 febantel, 491 febrile paroxysms, in birds, 43 fecal flotation visualization and measurement method, 112 f*ckled Duck (see Stictonetta noevosa) female gametocyte. see macrogametocytes fenbendazole, 272, 322, 340, 352, 437, 459, 491 Ferruginous Hawk (see Buteo regalis) Ferruginous Pochard (see Aythya nyroca) Ferruginous Pygmy-Owl (see Glaucidium brasilianum) Festive Parrot (see Amazovi feativa) fertilization, of Haemoproteus species, 17 fibrosis, 370, 421 Fieldfare (see Turdus pilaris) Filaria spp. Filaria cypseli, 521 Filaria nasuta, 327 Filaria uncinata, 335 Filarioidea, 434, 439, 440 filarioid nematodes clinical signs, 450 diagnosis, 454–455 epizootiology, 445–450 etiology, 440 history, 439–440 host range and distribution, 440–445 immunity, 455–458 pathology, 450–454 public and domestic animal health concerns, 458 sites in the avian host occupied by adult avian filarioids, by subfamily and genus, 448–450 treatment and control, 459 wildlife population impacts, 458–459 filarioids, 439 Filicollis anatis, 285 Fimbriaria fasciolaris, 263, 264, 269 fimbriariasis, 262 finches, 111, 121, 204 Fish Crow (see Corvus ossifragus) Fisher’s Lovebird (see Agap*rnis fisheri) fish parasites (see Eustrongylides spp.) flamingo (see Phoenicopterus spp.) Florida Sandhill Crane (see Grus canadensis pratensis) Florida Scrub Jay (see Aphelocoma coerulescens) Florida Snail Kite (see Rostrhamus sociabilis plumbeus) flubendazole, 352, 371 Fluminicola virens, 227 formalin-killed vaccine, 94 Formicariidae (Antthrushes and Antpittas), 443 Forpus passerinus (Green-rumped Parrotlet), 114, 399 Fox Sparrow (see Passerella iliaca) Francolinus spp. Francolinus afer (Red-necked Francolin), 392 Francolinus ahantensis (Ahanta Francolin), 395 Francolinus bicalcaratus (Double-spurred Francolin), 391, 396 Francolinus erckelii (Erekel’s Froncolin), 217 Francolinus francolinus (Black Francolin), 155 Francolinus gularis (Swamp Francolin), 393 Francolinus pintadeanus (Chinese Francolin), 390 Fratercula corniculata (Horned Puffin), 332 free-ranging species, challenges in studying, 3 Fregatidae (Frigatebirds), 441 Freitascapillaria, 463 Fringilla coelebs (Chaffinch), 121, 206, 527 Fringillidae (Siskins and Crossbills), 60, 108, 444
Index Fuadias madagascariensis, 435 Fulica spp., 465 Fulica americana (American Coot), 208, 226, 227, 233, 238, 336, 361, 378, 445, 446, 451, 475 Fulica atra (Eurasian Coot), 205, 226, 227, 297, 331, 361, 365, 385, 331 Fulvous Whistling-Duck (see Dendrocygna bicolor) Furnariidae (Ovenbirds), 61, 435, 443 Gabar Goshawk (see Micronisus gabar) Gadwall (see Anas strepera) Galapagos Dove (see Zenaida galapagoensis) Galapagos hawk (see Buteo galapagoensis) Galbulidae (Jacamars), 443 Gallicolumba luzonica (Luzon Bleeding-heart), 19, 23, 217, 332, 398 Galliformes, 14, 18, 23, 59, 121, 136, 163, 165–169, 167, 205, 208, 217, 252, 322, 360, 376, 435, 441, 465, 473, 521 Gallinago, 465 Gallinago gallinago (Common Snipe), 361, 476 Gallinula spp., 465 Gallinula chloropus (Common Moorhen), 225, 227, 361, 378, 475, 488 Gallinula ventralis (Black-tailed Native-hen), 361 r , 352 Gallomec Plus Galloperdix spadicea (Red Spurfowl), 392 Gallus spp., 465 Gallus gallus (domestic chicken), 23, 113, 114–115, 169, 195, 327, 455, 473 Gallus gallus (Red Jungle Fowl), 155, 331, 392, 396, 521 Gallus lafayetii (Ceylon Junglefowl), 390 Gallus sonneratii (Gray Junglefowl), 391 Gambusia holbrooki (Eastern mosquitofish), 310 game-farm pheasants, 134 gametes, 110 gametocytes, 19 circulating, 36 development in species of Leucocytozoon, 57–58 of Haemoproteus meleagridis, 15 of Leucocytozoon simondi, 83 mature, morphological forms of the, 16 morphology of, 15 of Plasmodium species, 39–40 gametogenesis, of Haemoproteus species, 17 gametogony, 110, 165, 188 Gammarus lacustris, 335, 337, 378 Ganguleterakis, 394 gapes, 343 gapeworm infection, 343, 347–348 gapeworms, 344, 349–350, 351 Garganey (see Anas querquedula) Garniidae (genus Fallisia), 36 Garrulus, 465 Garrulus glandarius (Eurasian Jay), 206, 481 Garter snake (Thamnophis sirtalis), 302 Gastrotaenia spp., 261 Gastrotaenia cygni, 262, 264 Gastrotaenia dogieli, 262, 264 gastrotaeniasis, 262 Gavia spp. Gavia arctica (Arctic Loon), 261, 291, 328 Gavia immer, 56 Gavia immer (Common Loon), 228, 291, 466, 538, 542 Gavia stellata (Red-throated Loon), 291, 328, 360, 466 Gaviidae (Loons), 59, 360, 441, 505 Gaviiformes, 14, 59, 163, 291, 328, 360, 441, 466
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Index Gelochelidon, 465 Geopelia humeralis (Bar-shoulderdd Dove), 73 Geopelia striata (Zebra Dove), 73, 126 Geopetitia aspiculata, 376, 377, 378, 380, 381 Geophaps lophotes (see Crested Pigeon) Geothlypis poliocephala (Gray-crowned Yellowthroat), 333 Geotrygon montana (Ruddy Quail-Dove), 398 giemsa stain, 13, 39, 45, 112, 458 Gigantobilharzia spp., 231, 246, 247, 249, 251 acotylea, 251 adami, 251 ardeola, 251 gyrauli, 251 huronensis, 251 huttoni, 251 lawayi, 251 nettapi, 251 plectropteri, 251 sturniae, 251 tantali, 251 Giovannolaia, 37, 39 gizzard infections, 267 gizzard nematodes, 367 gizzard worm, 326, 355, 371 Glareolidae (Pratincdes and Coursers), 442 Glaucidium brasilianum (Ferruginous Pygmy-Owl), 206 Glaucidium passerinum (Eurasian Pygmy-Owl), 394 2 -globulin, 42 1-and 2 -globulin, 42 Glossiphoniidae, 505, 506 glossiphoniid leeches, 508 glossiphoniids, 506 Glossy Ibis (see Plegadis falcinellus) glutamate dehydrogenase, 42 -glutamyltransferase, 42 gobies (Neogobius kessleri and Neogobius melanostomus), 298 Golden-crested Myna (see Ampeliceps coronatus) Golden Eagle (see Aquila chrysaetos) Golden-olive Woodpecker (see Piculus rubiginosus) Golden Pheasant (see Chrysolophus pictus) Golden-winged Parakeet (see Brotogeris chrysoptera) Goldeneye (see Bucephala spp) Goliath Heron (see Ardea goliath) Gouldian Finch (see Chloebia gouldiae) Goura spp. Goura cristata (Western Crowned-Pigeon), 217 Goura scheepmakeri (Southern Crowned-Pigeon), 217 Goura victoria (Victoria Crowned-Pigeon), 217 Gracula religiosa (Common Hill Myna), 114, 170 Grallina cyanoleuca (Magpie-lark), 162, 481 Grallinidae (Mudnest Builders), 61, 445 granulomas, 191 grasshoppers (Bruneria brunnea, Camnula pellucida, and Melanoplus spp.), 402 Grass snake (Natrix tessellata), 302 Gray Bunting (see Embesiza variabilis) Gray-cheeked Thrush (see Catharus minimus) Gray Crowned Crane (see Balearica regulorum) Gray-faced Woodpecker (see Picus canus) Gray Hawk (see Buteo nitidus) Gray Heron (see Ardeo cinerea) Gray Junglefowl (see Gallus Sonneratii) Gray partridge (see Perdix perdix) Gray Peaco*ck-Pheasant (see Polyplectron bicalcaratum) Gray Teal (see Anas gracilis) Gray Tinamou (see Tinamus tao)
573
Graylag Goose (see Anser anser) Great Bittern (see Botaurus stellaris) Great Black-headed Gull (see Larus ichthyaetus) Great Blue Heron (see Ardea hercdias) Great Bustard (see Otistarda) Great Crested Grebe (see Podiceps cristatus) Great Egret (see Ardeo alba) Great Horned Owl (see Bubo virginianus) Great Painted-Snipe (see Rostratula benghalensis) Great Spotted Woodpecker (see Dendrocopos major) Great-tailed Grackle (see Quiscalus mexicanus) Great Tit (see Parus major) Greater Coucal (see Centropus sinensis) Greater Kestrel (see Falco rupicoloides) Greater Prairie-Chicken (see Tympanuchus cupido) Greater Rhea (see Rhea americana) Greater Sage-Grouse (see Centrocercus urophasianus) Greater Sandhill Crane (see Grus canadensis tabida) Greater Scaup (see Aythya marila) Greater Spotted Eagle (see Aquila clonga) Greater white-fronted Goose (see Anser albifrons) Greater Yellow legs (see Tringa melanoleuca) Green-and-gold Tanager (see Tangara schrankii) Green-backed Tit (see Pasus monticolus) Green-cheeked Parakeet (see Pyrrhula molinae) Green Heron (see Butorides virescens) Green Jay (see Cyanocorax yncas) Green Peafowl (see Pavo muticus) Green Pheasant (see Phasianus versicolor) Green-rumped Parrotlet (see Forpus passerinus) Green-winged Teal (see Anas carolinensis) Green-Woodpecker (see Picus viridis) grouse disease, 316, 326 Gruidae (Cranes), 59, 442 Gruiformes, 14, 22, 59, 163, 169, 205, 208, 238, 297, 360, 361, 376, 465, 475 gruiforms, 121 Grus spp. Grus americana (Whooping Crane), 181, 185, 188, 285, 331, 339, 346, 399, 406, 475 Grus canadensis canadensis (Lesser Sandhill Crane), 183 Grus canadensis pratensis (Florida Sandhill Crane), 184 Grus canadensis pulla (Mississippi Sandhill Crane), 184 Grus canadensis rowani (Canadian Sandhill Crane), 183 Grus canadensis (Sandhill Crane), 182, 331, 475 Grus canadensis tabida (Greater Sandhill Crane), 183, 191 Grus grus (Common Crane), 346, 397, 475, 550 Grus japonicus (Red-crowned Crane), 181, 185 Grus monacha (Hooded Crane), 185, 188 Grus antigone (Sarus Crane), 184, 397 Grus vipio (white-naped Crane), 181, 182, 184 Gryporhynchidae, 264, 266 Guanay Cormorant (see Phalacrocorax bougainvillii) Guttera pucherani (Crested Guineafowl), 396 Gymnophallus (Microphallidae), 227, 230 Gymnorhina tibicen (Australasian Magpie), 481 Gymnostinops montezuma (Montezuma Oropendola), 24 Gynaecophila, 381 Gyps africanus (White-backed Vulture), 74, 208 Gyrfalcon (see Falco rusticolus) Gyropidae, 516 Haemadipsa zeylanica, 505 Haemadipsidae, 502, 505, 506, 508 Haemamoeba, 37, 39
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574 Haematomyzidae, 516 Haematopodidae (Oystercatchers), 442 Haematopus ostralegus (Eurasian Oystercatcher), 331, 516 Haementeria depressa, 505 Haementeria medicinalis, 502 Haemogregarina, 108, 109 haemoproteid infections, among the avian orders, 14 Haemoproteids, 14 haemoproteosis, 35 Haemoproteus spp., 6, 23, 24, 35, 40, 46, 54 clinical signs, 21 complex life cycle of, 17 diagnosis, 25–26 distribution, 14 domesticated animal health concerns, 27 epizootiology, 17–21 etiology, 15–17 gametocytes of, 16 Haemoproteus attenuatus, 22 Haemoproteus balearicae, 22 Haemoproteus balmorali, 18 Haemoproteus belopolskyi, 18 of Blackcaps (Sylvia atricapilla), 20 Haemoproteus coatneyi, 20, 22 Haemoproteus columbae, 13, 18–19, 22, 23 of pigeons and doves, 13 of Rock Pigeons, 26 transmission by hippoboscid flies, 20 Haemoproteus danilewskii, 18–19, 22 Blue Jays with infection of, 21 of ducks, 20 Haemoproteus dolniki, 18 Haemoproteus fringillae, 18 Haemoproteus halcyonis, 24 Haemoproteus handai, 18, 23 Haemoproteus-infected mates, 28 Haemoproteus lanii, 18 Haemoproteus lophortyx, 17, 18, 21, 23 bobwhites with natural infections of, 21 Haemoproteus maccallumi, 18, 22 Haemoproteus majoris, 27 Haemoproteus mansoni, 18 of Ruffed Grouse (Bonasa umbellus), 20 Haemoproteus meleagridis, 15, 18–19, 21, 23, 25, 26, 27 gametocytes of, 15 megalomeront of, 19 of wild turkeys, 27 Haemoproteus nettionis, 18, 22 of ducks, 20 Haemoproteus noctuae, 21 Haemoproteus palumbis, 18, 20, 22 Haemoproteus passeris, 22, 24 Haemoproteus prognei, 7, 28 Haemoproteus ptilotis, 22 Haemoproteus sacharovi, 17, 18, 22, 23 Haemoproteus syrnii, 21, 24 Haemoproteus tartakovskyi, 18 Haemoproteus tinnunculi, 28 Haemoproteus turtur, 18 Haemoproteus velans, 18 of woodpeckers, 20 history, 13–14 host range, 14–15 immunity, 26–27 infection in pigeons and doves, 27 infections, subclinical impacts of, 28 management implications, 28
Index as models for testing evolutionary theories, 13 morphological forms of the mature gametocytes, 16 pathogenesis and pathology, 21–25 preerythrocytic tissue stages of, 13 public health concerns, 27 sexual and asexual reproduction, 17 sexual stages of, 13 synonyms, 13 and temperate climates, 20 treatment and control, 28 wildlife population impacts, 27–28 haemosporidian disease, 54 Haemosporidiosis, 13 Haemospororida order, 36 hairworms, 316, 463 Haliaeetus spp., 465 Haliaeetus albicilla (White-tailed Eagle), 75, 237, 239, 346, 472 Haliaeetus leucocephalus (Bald Eagle), 75, 215, 233, 237, 297, 298, 472 Haliaeetus vocifer (African Fish Eagle), 75 halofuginone, 48 Harrier, 550 Harlequin Duck (see Histrionicus histrionicus) Hartlaub’s Duck (see Pteronetta hartlaubii) Hawaii Amakihi (see Hemignathus virens) Hawaiian Crow (see Corvus hawaiiensis) Hawaiian Goose (see Branta sandvicensis) Hawks (Falconiformes), 277 Hazel Grouse (see Bonasa bonasia) Heliornithidae (Finfoots), 442 Helmeted Guineafowl (see Numida meleagris) Helodrilus caliginosus, 486 hematazoa, 7 hematocrits, 42, 43 hematophagous dipteran larvae, 548–549 hematozoan disease, 54 infections, in birds, 35 hematozoans, 21 hematozoin, 39 Hemignathus virens (Hawaii Amakihi), 46, 48, 111 Hemiprocnidae (Treeswifts), 443 hemorrhagic streaks, 21 hemosiderin, 89 heparinized Mallard blood (Anas platyrhynchos domesticus), 502 hepatic megalomeronts, 26 Hepaticola, 463 hepatic trematodiasis, 233 Hepatozoon, 108, 109 Heptapsogaster sp., 517 Heraldakis, 394 heterakiasis, 388 heterakid eggs, 156 Heterakis spp., 394, 402 avian hosts infected with, 389–394 clinical signs, 404 diagnosis, 405 distribution and host range, 388–394 epizootiology, 401–404 etiology, 394–401 Heterakis beramporia, 391, 392, 393 Heterakis bosia, 393 Heterakis brevispiculum, 391, 392 Heterakis dispar, 389, 390, 392, 393, 394 Heterakis gallinarum, 6, 156, 388, 389, 390, 391, 392, 393, 394, 401, 402, 403–404, 404, 405, 407, 413, 491
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Index Heterakis hamulus, 391, 392 Heterakis indica, 392 Heterakis inglisi, 389 Heterakis interlabiata, 392 Heterakis isolonche, 388, 390, 391, 392, 393, 401, 405 Heterakis kumaoni, 392 Heterakis kurilensis, 394 Heterakis macroura, 390, 392 Heterakis meleagris, 393 Heterakis nainitalensis, 392 Heterakis nattereri, 389, 390 Heterakis oscari, 390 Heterakis parva, 392, 393 Heterakis pavonis, 389, 391, 392, 393, 394 Heterakis pedioecetes, 393 Heterakis psophiae, 393 Heterakis pusilla, 390, 392 Heterakis silindae, 391, 392 Heterakis skrjabini, 393 Heterakis spiculata, 389 Heterakis tenuicauda, 390, 392, 393 Heterakis tragopanis, 393 Heterakis valdemucronata, 389 Heterakis vulvolabiata, 391 history, 388 immunity, 405 pathology, 404–405 public and domestic animal health concerns, 405 synonyms, 388 treatment and control, 406–407 wildlife population impacts, 405–406 heterakosis, 388 Heterocypris incongruens, 335 heteromorphic deutonymphs (hypopi), 529 heterophils, 21 Heterophyidae, 230, 235, 237 Hexametra sp., 425 Hill Partridge (see Arborophila torqueda) Himalayan Monal (see Lophophorus impejanus) Himalayan Snowco*ck (see Tetraogallus himalayensis) Himantopus leucocephalus (Pied Stilt), 361 Himantopus, 465 Himantopus himantopus (Black-winged Stilt), 361 Himasthla, 230 Himatione sanguinea (Apapane), 111 hippoboscid flies, 15, 18 columbiform parasites from, 15 as vectors, 15 hippoboscid transmission, of H. lophortyx, 17 hippoboscid vectors, 13–14 Hirudinidae, 502, 505, 506, 508 hirudinids, 502, 507 Hirudobdella antipodum, 505 Hirudo medicinalis, 505 Hirudo nipponia, 505 Hirundinidae (Swallows), 60, 435, 444 Hirundoecus, 519 Hirundo rustica (Barn Swallow), 481, 518 Hispaniolan Parakeet (see Aratinga chloroptera) Hispaniolepis falcata, 264, 268 histomonads, 156, 157 Histomonas spp. Histomonas bonasae, 156 Histomonas isolonche, 156 Histomonas meleagridis, 140, 154, 158, 405 epizootiology of, 156 infected liver, gross lesions of, 157
575
within lesions, 158 species of the order galliformes reported to be susceptible to infection, 155 histomoniasis clinical signs, 156–157 diagnosis, 157–158 distribution, 154 domestic animal and public health concerns, 158 epizootiology of, 156 etiology, 154 from junglefowl, 159 history, 154 host range, 154 immunity, 158 pathogenesis and pathology, 157 prevention, control, and management implications, 158–159 synonyms, 154 wildlife population impacts, 158 Histrionicus histrionicus (Harlequin Duck), 330, 358 holostomes, 226 hom*o sapiens (Human), 302 honeycreepers (Drepanidinae), 48 Hooded Crane (see Grus monacha) Hooded Crow (see Corvus cornix) Hooded Merganser (see Lophodytes cucullatus) Hooded Vulture (see Necrosyrtes monachus) Hooded Warbler (see Wilsonia citrina) Horned Grebe (see Podiceps auritus) Horned Puffin (see Fratercula corniculata) host foraging or nesting behavior, 21 host–parasite relationship, 3, 13, 256 host responses, from avian hosts, 22–24 host specificity, of the parasites, 21 Houbara Bustard (see Chlamydotis undulata) House Martin (see Delichon urbicum) House Sparrow (see Passer domesticus) House Wren (see Troglodytes aedon) Houttuynia struthionis, 272 Huffia, 37, 39 Huffmanela, 463 Humboldt Penguin (see Spheniscus humboldti) Hyacinth Macaw (see Anodorhynchus hyacinthinus ) Hyalella, 337 Hyalella azteca, 337 Hydrobatidae (Storm Petrels), 441 Hylocichla mustelina (Wood Thrush), 238 Hymenolaimus malacorhynchos (Blue Duck), 329 Hymenolepididae, 262 hymenolepididiasis, 262 Hymenolepis falsata, 269 Hymenolepis microps, 271 hyperplasia, of white pulp arteriolar endothelium, 21 hyperplasia of the proventriculus, 237 Hypodectes, 529 Hypodectes nudus, dorsal and ventral views of adult female, 532 Hypoderaeum, 230 Hypoderaeum conoideum, 226 hypoderatid mites (Hypoderatidae), 529 Hyptiasmus, 229 Hysterothylacium, 413, 414, 423 Ibidorhynchidae, 442 Icteridae (Troupials), 60, 435, 444 Ictinaetus malayensis (Black Eagle), 74 IgA and IgG1 pathogen-specific antibodies, 142
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576 Iiwi (see Vestiaria coccinea) immunoblot analysis, 92 immunocompromised condition, of the bird, 56–57 immunofluorescence, 92 Inca Dove (see Columbina inca) Indian Peafowl (see Pavo cristatus) Indicatoridae (Honeyguides), 61, 443 indirect hemagglutination test (IHAT), 214 Indocapillaria, 463 inflammatory infiltrates, 25 inflammatory myopathy, 44 InPouch culture kits, 136, 139 insecticides, treatment with, 554 intestinal cryptosporidiosis, 198 intestinal flagellates, 3 intestinal infection, in cranes, 187 Intestine, 449 intraerthrocytic development of Haemoproteus species, 15 of Plasmodium species, 36 intraerythrocytic gametocytes, morphology of, 25 intraerythrocytic hemosporidian parasites, 13 intraerythrocytic parasites, of wild birds, 35 intravascular agglutinations, of erythrocytes, 44 intrinsic factors, of parasites, 21 Irenidae (Fairy-bluebirds), 61, 444 Ischnocera, 516, 517 Island Canary (see Serinus canaria) Island Collared-Dove (see Streptopelia bitorquota) Isopoda, 279 isopod intermediate hosts, elimination of the, 339 Isospora spp., 163 clinical signs, 112 diagnosis, 112–113 distribution, 109 domestic animal health concerns, 117 epizootiology, 110–112 etiology that cause atoxoplasmosis, 109–110 history, 108–109 host range, 109 immunity, 113 Isospora buteonis, 424 Isospora canaria, 110 Isospora rothschildi, 109 oocyst of, 109–110 pathogenesis and pathology, 112 public health concerns, 116 synonym, 108 that cause atoxoplasmosis, 108 treatment and control, 117 wildlife population impacts, 117 Israeli House Sparrow (see Passer domesticus biblicus) Ithaginis cruentus (Blood Pheasant), 390, 396 ivermectin, 340, 352, 381, 406, 437, 459, 491, 535 Ixobrychus sinensis (Yellow Bittern), 488 Ixoreus, 465 Ixoreus naevius (Varied Thrush), 481 Jacanidae (Jacanas), 59, 442 Jackal Buzzard (see Butea rufofuscus) Jackass Penguin (see Spheniscus demersus) Japanese Quail (see Coturnix japonica) Japanese Sparrowhawk (see Accipiter gularis) Japanese White-eye (see Zosterops japonicus) Jararaca (see Bothrops jararaca) Jardaya Parakeet (see Aratinga jandaya) Java Sparrow (see Padda oryzivora)
Index Jilinobilharzia, 246, 247, 251 Jilinobilharzia crecci, 251 Jones’ Barn (JB) strain, 134–135 Kalij Pheasant (see Lophura leucomelanos) Kaplan–Meier survivorship curves, 144 Kaupifalco monogrammicus (Lizard Buzzard), 74, 78 Kea (see Nestor notabilis) Keel-billed Toucan (see Ramphastos sulfuratus) Killdeer (see Charadrius vociferus) killifish (Fundulus heterocl*tus), 290 King Eider (see sanaena spectabilis) kingfishers (Megaceryle spp.), 226 King Vulture, (see Sarcoramphus papa) Kinkajou (Potos flavus), 425 Kitodites, 529 Klebsiella spp., 434 Knemidokoptes spp. Knemidokoptes derooi, 528 Knemidokoptes jamaicensis, 528 Knemidokoptes pilae, 528 knemidokoptic podoacariasis, 535 knemidokoptids, 531 koilin lining, 369 Kori Bustard (Ardeotis kori) Kowalewskiella, 263 Kupffer cells, 89 Kytodites nudus, dorsal and ventral views of adult female, 531 labored breathing, 87 lactic acid, 509 Lady Amherst’s Pheasant (see Chrysolophus amherstiae) Laemobothriidae, 516 Lagochilascaris, 425 Lagopus, 465 Lagopus lagopus scotica (Red Grouse), 5, 6, 7, 316, 322 Lagopus lagopus (Willow Ptarmigan), 261, 268, 271, 320, 345, 393, 397, 403, 405, 459, 464 Lagopus leucura (White-tailed Ptarmigan), 397 Lagopus muta (Rock Ptarmigan), 360, 393, 397 Laminosioptes cysticola (Laminosioptidae), 528, 532 laminosioptids, 534 Lamprotornis superbus (Superb Starling), 111 Laniidae (Shrikes), 60, 444 Lanius excubitor (Northern Shrike), 206 Lanius schach (Long-tailed Shrike), 482 Lankesterella, 108, 109 Lanner Falcon (see Falco biarmicus) Lappet-faced Vulture (see Torgos tracheliotus) Large-tailed Nightjar (see Caprimulgus macrurus) Laridae (Gulls), 442, 505 Larus spp. Larus, 465 Larus argentatus (European Herring Gull), 332, 238, 346 Larus atricilla (Laughing Gull), 208, 238 Larus canus (Mew Gull), 291, 332 Larus delawarensis (Ring-billed Gull), 208, 255 Larus ichthyaetus (Great Black-headed Gull), 332 Larus novaehollandiae (Silver Gull), 46, 476 Larus ridibundus (Black-headed Gull), 205, 291, 343, 346 Larus thayeri (Thayer’s Gull), 332 larval Austrobilharzia, 246 larval Bilharziella, 247 larval diphyllobothriids, 263 larval Gigantobilharzia, 247
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Index larval Mesocestoides (Cyclophyllidea), 263 larval Ornithobilharzia, 246 larval Trichobilharzia, 247 larva migrans, 424 Lateriporus sp., 267 Laterotrema, 231 latex agglutination test (LAT), 92, 214 Laughing Dove (see Streptopelia senegalensis) Laughing Gull (see Larus atricilla) Laysan Albatross (see Phoebastria immutabilis) Laysan Duck (see Anas laysanensis) Least Sandpiper (see Calidris minutilla) Lecithodendriidae, 230, 238 leech parasites, of birds clinical signs, 506 diagnosis, 506–508 epizootiology, 503–506 etiology, 502–503 freshwater, 501 history, 501 host range and distribution, 501–502, 505 immunity, 508–509 life cycle of, 503 pathogenesis and pathology, 506 public and domestic animal health concerns, 509 sanguivorous, 503–504 Leipoa ocellata (Malleefowl), 517 Lemdana, 441–445, 448, 454 Lemdana wernaarti, 457 Lemdaninae, 448 Lemon Dove (see Columba larvata) Leptoptilos crumeniferus (Macabou Stork), 269, 295, 389, 395, 454 Leptosomatidae (Cuckoo Rollers), 443 Leptotila verreauxi (White-tipped Dove), 129 lesions by acute infections with Plasmodium, 44–45 associated with atoxoplasmosis, 116 associated with blood-feeding larvae, 552 associated with Trichom*onas gallinae, 137–138 Baylisascaris species, 427 caused by Contracaecum, 421 caused by fungi and nematods, 140 of Cryptosporidium, 198 from Dispharynx nasuta, 334 in ducks infected with Cloacotoenia megalops, 269 due to trematodes, 233 in DVC, 181, 190–191 of Eimeria species, 182–185 Eustrongylides spp. infections, 298, 305–307 of Histomonas meleagridis, 157 of renal Eimeria, 175 from schistosomes, 246, 249, 255 Serratospiculum, 436 of Toxoplasma, 212–213 in waterfowl with fatal leucocytozoonosis, 89, 91 Lesser Flamingo (see Phoenicopterus minor) Lesser Kestrel (see Falco naumanni) Lesser Scaup (see Aythya affinis) Lesser Spotted Eagle (see Aquila pomarina) Lesser Snow Goose (see Chen caerulescens caerulescens) Lesser White-fronted Goose (see Anser erythropus) Lesser Sandhill Crane (see Grus canadensis canadensis) lethargy, 87 Leucochloridiidae, 230 Leucochloridiomorpha, 229 Leucochloridium, 230
577
Leucocytoon, by orders and families of birds, 59–61 Leucocytozoidae, 54, 57 leucocytozoids, 54 Leucocytozoon spp., 13, 17, 27, 35, 46, 54, 59, 60, 61, 82–87 clinical signs, 87 diagnosis, 91–92 disease, 54 distribution, 56 domesticated animal health concerns, 93 epizootiology, 82–87 etiology, 57–82 in a gaviiform, 56 geographic distribution of eight species of, 57, 58 history, 54–55 host range, 56–57 immunity, 92–93 infections, 25 Leucocytozoon balmorali, 60, 61, 85 Leucocytozoon bennetti, 60, 85 Leucocytozoon berestneffi, 60, 81, 85 Leucocytozoon bonasae, 58 Leucocytozoon caprimulgi, 59, 85 Leucocytozoon caulleryi, 54, 55, 57, 59, 81, 82, 85, 89, 92, 93 Leucocytozoon centropi, 59, 85 Leucocytozoon cheissini, 59, 85 Leucocytozoon colius, 59, 85 Leucocytozoon communis, 60, 85 Leucocytozoon danilewskyi, 59, 81, 84, 85, 88 Leucocytozoon dizini, 59, 85 Leucocytozoon dubreuili, 60, 61, 81, 82, 84, 85 Leucocytozoon eurystomi, 60, 85 Leucocytozoon fringillinarum, 60, 61, 81, 85 Leucocytozoon grusi, 59, 85 Leucocytozoon hamiltoni, 61, 85 Leucocytozoon leboeufi, 59, 85 Leucocytozoon legeri, 59, 85 Leucocytozoon lovati, 59, 81, 84, 85 Leucocytozoon maccluri, 85 Leucocytozoon macleani, 57, 59, 85, 93 Leucocytozoon majoris, 27, 60, 61, 81, 85 Leucocytozoon marchouxi, 55, 56, 57, 59, 70–73, 85, 87, 89, 91, 93 Leucocytozoon neavei, 58, 59, 85 Leucocytozoon nycticoraxi, 59, 85 Leucocytozoon nyctyornis, 60, 85 Leucocytozoon polyoon, 226, 234 Leucocytozoon sakharoffi, 60, 85 Leucocytozoon sakharoffi, of crows, 81 Leucocytozoon schoutedeni, 57, 58, 59, 85, 93 Leucocytozoon simondi, 55, 57, 58, 59, 82, 83, 84, 85, 88, 89, 90, 537 gametocytes of, 83 infections in waterfowl, 92 infections in wood ducks, 84 life cycle of, 82 megalomeronts of, 90 in waterfowl, 81, 86, 87, 93 in wild waterfowl, 56, 62–69 Leucocytozoon smithi, 54, 55, 57, 59, 84, 85, 86, 87, 537 from turkeys, 81 in Wild Turkeys (Meleagris gallopavo), 86, 87 Leucocytozoon sousadiasi, 59, 85 Leucocytozoon squamatus, 60, 85 Leucocytozoon struthionis, 57, 59, 85 Leucocytozoon tawaki, 85
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578 Leucocytozoon spp. (Cont.) Leucocytozoon toddi, 55, 57, 59, 84, 85, 91 in diurnal raptors, 81, 87 in raptors, 56, 89, 93 in wild falconiforms, 74–80 Leucocytozoon vandenbrandeni, 59, 85 management implications, 95 by orders and families of birds, 59–61 pathogenesis, 87–89 pathology, 89–91 prevalence of species of Leucocytozoon in birds in various zoogeographic regions, 56 prevention, treatment and control, 94–95 public health concerns, 93 synonym, 54 wildlife population impacts, 93–94 Leucopsar rothschildi (Bali Myna), 109, 116 merozoite in monocyte of, 110 Leucosarcia melanoleuca (Wonga Pigeon), 217 Leukocytozoon (sic) danilewskyi, 54, 87 levamisole, 339, 340, 381, 406, 459, 491 levamisole hydrochloride, 322, 407, 459 Levant Sparrowhawk (see Accipiter brevipes) Levinseniella, 230 Leyogonimus, 230 Leyogonimus polyoon, 225, 227, 238 life history traits, of a bird, 4 Ligula, 267, 271 Ligula intestinalis, 264, 267, 269, 270 Limnodrilus hoffmeisteri, 300–301 Limosa fedoa (Marbled Godwit), 451, 521 Limosa lapponica (Bar-tailed Godwit), 271–272 Limpkin (see Aranus guarauna) Little Black Cormorant (see Phalacrocorax sulcirostris) Little Blue Heron (see Egretta caesulea) Little Bustard (see Tetrox tetrax) Little Cuckoo-Dove (see Macropyg’s ruficeps) Little Egret (see Egretta garzetta) Little Grebe (see Tachybaptus ruficollis) Little Owl (see Athene noctua) Little Penguin (see Eudyptula minor) Little Pied Cormorant (see Phalacrocorax melanoleucos) Little Sparrowhawk (see Accipiter minullus) Little Swift (see Apus affinis) livebearers (Poeciliidae), 290 liver enlargement, 116 Lizard Buzzard (see Kupifalco monogrammicus) Locusta migratoria, 378 Lofortyx, 465 Lonchura punctulata (Nutmeg Mannikin), 111 Long-billed Partridge (see Rhizothera longirostris) Long-crested Eagle (see Lophaetus occipitalis) Long-legged Buzzard (see Butea rufinus) Long-tailed Cormorant (see Phalacrocorax africanus) Long-tailed Duck (see Clangula hyemalis) Long-tailed Parakeet (see Psittacula longicaudata) Long-tailed Shrike (see Lonias schach) Lophaetus occipitalis (Long-crested Eagle), 76 Lophodytes, 465 Lophodytes cucullatus (Hooded Merganser), 67, 297, 330 Lophophorus impejanus (Himalayan Monal), 391 Lophura spp. Lophura diardi (Siamese Fireback), 114 Lophura ignita (Crested Fireback), 391 Lophura leucomelanos (Kalij Pheasant), 392 Lophusa nycthemera (Silver Pheasant), 393
Index Loriidae (Lories and Lorikeets), 442 Lorinus sp. (Lory), 115 Lucilia sericata, 550, 552 Lucilia spp., 553 Lueheia spp., 277 Lueheia inscripta, 281 Lueheia inscripta, transmission of, 281 Lumbricus terrestris, 486 lumen of blood vessels, 449 Lunaceps numenii, 521 lungs, 449 Luscinia calliopei (Siberian Rubythroat), 482 Luscinia megarhynchos (Common Nightingale), 482 Luzon Bleeding-heart (see Gallicoumba luzonica) lymphatics, 449 lymphocytes, 21 lymphocytic infiltrates, 21 lymphoid hyperplasia, 369 lymphoid–macrophage system capillary endothelium cells, 19 Lynceus brachyurus, 335 Lynchia hirsuta, 17, 18 Lyperosomum, 229 Lyperosomum longicauda, 226 Lyrurus, 465 Macaw, 550 Macrobilharzia, 246, 247, 251 Macrobilharzia pulverulenta, 251 Macrocephalon maleo (Maleo), 390 Macrocyclops fuscus, 385 macrogametocytes, 17, 20, 39–40, 165 fertilization of, 40 Macropygia nigrirostris (Black-billed Cuckoo-Dove), 398 Macropygia ruficeps (Little Cuckoo-Dove), 73 Macyella, 230 Madagascar Buzzard (see Buteo brachypterus) Madagascar Green-Pigeon (see Treron australis) Madagascar Turtle-Dove (see Streptopelia picturata) Magellanic Penguin (see Spheniscus magellanicus) Magnificent Bird-of-paradise (Cicinnurus magnificus) Magpie Goose (see Anseranas semipalmata) Magpie-lark (see Grallina cyanoleuca) magpies, 347 Maguari Stork (see Ciconia maguari) Malaconotidae (Bushshrikes), 60 malaria, human, 35 malarial infection, in avian hosts. see avian malaria malarial parasites, 13 malarial pigment. see hematozoin male gametocyte. see macrogametocytes male microgametocytes, 17 Maleo (see Macrocephalon maleo) Mallard (see Anas platyrhynchos) Malleefowl (see Leipoa ocellata) Mallee Ringneck (see Barnardius barnardi) Mammomanogamus, 351 Mandarin Duck (see Aix galericulata) Maned Duck (see Chenonetta jubata) Mangrove Swallow (see Tachycineta albilinea) Manorina melanocephala (Noisy Miner), 22 Mansonia crassipes, 41 Marabou Stork (see Leptoptilos crumeniferus), 295, 389, 395 Marbled Godwit (see Limosa fedoa) Marbled Teal (see Marmaronetta angustirostris)
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Index Maritrema, 230 Maritrema acadiae, 236, 239 Maritrema bonaerensis n. sp., 240 Maritrema sp., 236 Marmaronetta angustirostris (Marbled Teal), 358, 364 Maroon Woodpecker (see Blythipicus rubiginosus) mass-capture methods, 3 mature gametocytes, 19 Mawsonotrema eudyptulae, 235 Meadow Pipit (see Anthus pratensis) Mealy Parrot (see Amazona farinose) mebendazole, 339, 340, 352, 371, 437, 459 Mediorhynchus spp., 277 Mediorhynchus centurorum, 282 Mediorhynchus centurorum in a Red-bellied Woodpecker, 282 Mediorhynchus gallinarum, 282, 283 Mediorhynchus orientalis, 283 mefloquine, 28, 48 megalomeronts, 19 with associated muscle pathology, 25 of Haemoproteus, 25 hepatic, 26 inflammatory infiltrates affecting, 25 in leucocytozoonosis, 88–89 from pectoral muscle of a domestic turkey with an experimental infection, 27 ruptured, from pectoral muscle of a domestic turkey, 26 Megaceryle alcyon (Belted Kingfisher), 238, 346 Megaceryle torquatus (Ringed Kingfisher), 238 megaloschizonts, 89 Megapodiidae (Megapodes), 441 Megascops asio (Eastern Screech-Owl), 133, 213 Melanerpes spp. Melanerpes carolinus (Red-bellied Woodpecker), 218, 282, 478 Melanerpes erythrocephalus (Red-headed Woodpecker), 332 Melanerpes flavifrons (Yellow-fronted Woodpecker), 376 Melanitta spp. Melanitta fusca (White-winged Scoter), 115, 285, 296, 330, 360, 365, 469 Melanitta nigra (Black Scoter), 67, 356, 362, 395, 469 Melanitta perspicillata (Surf Scoter), 67, 330, 359, 469 Melanopus spp., 378 melarsomine, 95 melatonin, 40 Meleagrididae (Turkeys), 59 Meleagris spp., 465 Meleagris gallopavo (Domestic Turkey, Wild Turkey), 13, 23, 154, 155, 167, 169, 195, 205, 208, 214, 215, 217, 233, 234, 237, 268, 327, 345, 388, 393, 397, 464, 473, 528 with infections of Eimeria, 163 with infections of Plasmodium durae, 42 Meleagris ocellata (Ocellated Turkey), 392, 396 Melierax metabates (Dark Chanting Goshawk), 78 Meliphagidae (Honeyeaters), 61, 444 Melopsittacus, 465 Melopsittacus undulatus (Budgerigar), 113, 137, 168, 206, 210, 213, 399 meningoencephalitis, 424 Menopon gallinae, 521 Menoponidae, 516 Menuridae (Lyrebirds), 444 Mergellus albellus (Smew), 67, 297, 330, 359
579
Mergus merganser (Common Merganser), 66, 227, 255, 297, 306, 331, 469 Mergus serrator (Red-breasted Merganser), 67, 227, 297, 304, 305, 470 Merlin (see Falco columbarius) merogony, 13, 36, 39 meronts, 19, 21, 36, 57, 91, 116 morphology in species of Leucocytozoon, 58 Meropidae (Bee-eaters), 60, 443 merozoites, 82, 165 of Eimeria gruis and Eimeria reichenowi, 187–188 haemoproteids transmitted by hippoboscid flies, 20 of Isospora species, 112 in leucocytozoonosis, 88 released from exoerythrocytic meronts, 39 species of Cryptosporidium, 196–197 of species of Isospora, 110–111 transformation, 15, 19 Mesamidostomum, 366 Mesitornithidae (mesites), 442 Mesocestoides, 271 Mesocestoides sp., 264 Mesocestoididae, 264, 266 Mesocyclops leuckarti, 384, 385 Mesostephanus appendiculatoides, 235 Me-49 strain, 213 metacryptozoites, 39 Metallic Pigeon (see Columba Vitiensis) methanol, 112 Metorchis, 231 Metorchis bilis, 233, 237, 239 Metroliasthes lucida, 264, 268 metronidazole, 144 Mew Gull (see Larus canus) Mhc alleles, 42 mice (Mus musculus), 195 microcrustaceans, 279 microfilariae, 440 of avian filarioids, 453 morphology, 456–458 microfilariae, morphological traits of, 454–458 microfilaria (MF), 439 microgametocytes, 20, 39 Microlynchia pusilla, 18 Micronisus gabar (Gobar Goshawk), 79 Microphallidae, 230, 236 micropyle, 112 Microscaphidiidae, 230 Microsomacanthus collaris, 264, 267 Microsomacanthus parvula, 264, 268, 269 Microtetrameres spp., 376, 377, 381 Microtetrameres centuri, 379 Microtetrameres corax, 378, 379 Microtetrameres egretes, 376 Microtetrameres inermis, 378 Microtetrameres nestoris, 380 Microtetrameres spiralis, 376 Micrurus sp. (Coral snake), 302 midgut basal lamina, 17 midgut epithelium, 40 midgut wall, 17 Milvago chimango (Chimango Caracara), 79, 472 Milvus migrans (Black Kite), 76, 114, 130 Milvus milvus (Red Kite), 76, 472 Mimidae (Mockingbirds and Thrashers), 60, 435, 444 Mimus polyglottos (Northern Mockingbird), 208, 333 miracidium, 226, 248
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580 Mississippi Sandhill Crane (see Grus canadensis pulla) Mistle Thrush (see Turdus viscivorus) mitochondrial gene sequences, of avian haemoproteids, 17 Mitochondrial genes (mtDNA-cox2), 425 Mniotilta variai (Black-and-white Warbler), 482 modified agglutination test (MAT), 214 moist heather plants (Calluna vulgaris), 317 molecular methods, 25, 39 molluscicides, 240 Molothrus ater (Brown-headed Cowbird), 333, 482 Momotidae (Motmots), 60, 443 monensin, 172, 192 Monk Parakeet (see Myiopsitta monachus) Monocercomonadidae, 154 monocytes, 21 Montagu’s Harrier (see Circus pygargus) Montezuma Oropendola (see Gymnostinops montezuma) Monticola solitarius (Blue Rock-thrush), 482 mortality from acanthocephalan infections, 280 among leeches, 509 associated with Haemoproteus, 27 from atoxoplasmosis, 111 in breeding Willow Ptarmigan, 271 in California sea otters (Enhydra lutris), 285 caused by Eimeria truncata, 175 cestodes, 264–265 due to acanthocepalan-induced peritonitis, 285 due to dispharynxiasis, 339 from DVC, 192 of Eimeria spp., 182–185 Eustrongylides spp. infection, 309 leeches, 509 leucocytozoonosis, 94 myiasis, 552 from Plasmodium species, 48 from trematodes, 240 Mosquito Aedeomyia squomipennis, 41 Culex annulus, 41 Culex morsitans, 41 Culex nigripalpus, 41 Culex pipiens, 41 Culex quinque fasciatus, 41 Culex restuans, 41 Culex saltenensis, 41 Culex Sitiens, 41 Culex stimatosoma, 41 Culex tarsalis, 41 Mansonia crassipes, 451 mosquito vectors, 41, 451 Motacilla alba (White Wagtail), 345 Motacillidae (Wagtails and Pipits), 60, 444 motile zygote, 17 Mott cells, 21 Mottled Duck (see Anas fulvigula), 83, 235, 364 Mountain Chickadee (see Poecile gambeli) Mourning Dove (see Zenaida macroura) Mudpuppy (Necturus maculosus), 303 Murre (Uria sp.), 291 Muscicapidae (Old World Flycatchers), 60, 435, 444 Muscicapinae, 505 Muscidae, 547 Muscovy Duck (see Cairina moschata)
Index Musk Duck (see Biziura lobata) Musophagidae (Turacos), 59, 442 Musophagiformes, 59 Mussurana (see Clelia clelia) Mute Swans (see Cygnus olor) Mycteria americana (Wood Stork), 295, 468 Myialges nudus, 527, 530 Myiarchus crinitus (Crested Flycatcher), 529 myiasis clinical signs, 550–551 diagnosis, 552 distribution, 547 epizootiology, 549–550 etiology, 549 history, 547 host range, 547–549 immunity, 552 management implications, 554 pathogenesis and pathology, 551–552 public and domestic health concerns, 552 synonyms, 546–547 treatment and control, 553–554 wildlife population impacts, 552–553 Myiopsitta monachus (Monk Parakeet), 23, 116 myofibroblasts, 19 myositis, 21 in avian hosts, 13 Myrsidea ptilinorhynchi, 518 Myxobdella annandalei, 505 Nacunda Nighthawk (see Podager nacunda) Namaqua Dove (see Oena capensis) narasin, 177 nasal and oral exudation, 137 Nashville Warbler (see Vermivora ruficapilla) Nearctic ecozone, 20 Nearctic region, Leucocytozoon species in, 56 neck, 450 necrosis, 157, 175, 212 Necrosyrtes monachus (Hooded Vulture), 74 Nectariniidae (Sunbirds and Spiderhunters), 60, 444 neguvon, 371 nematodiasis, 368, 413 Neochen jubata (Orinoco Goose), 329 Neochmia modesta (Plum-headed Finch), 195 Neodiplostomum, 229 Neoknemidokoptes laevi, 532 Neoknemidokoptes laevis, 528 Neophema spp. Neophema bourkii (Bourke’s Parrot), 399 Neophema elegans (Elegant Parrot), 399 Neophema pulchella (Turquoise Parrot), 400 Neophema splendida (Scarlet-chested Parrot), 400 Neophron percnopterus (Egyptian Vulture), 130 Neotropic Cormorant (see Phalacrocorax brasilianus) Neottialges, 529 Neottiophilidae (Neottiophilum, Actinoptera), 547, 549 Neottiophilum praeustum, 549 Nerodia sipedon (Northern water snake), 302 Nerodia taxispilota (Brown water snake), 302 Nesoenas mayeri (Pink Pigeon), 70, 125, 134 nestling birds, cost of parasitism, 6 Nestor meridionalis (New Zealand Kaka), 380 Nestor notabilis (Kea), 476 nest site fidelity, 4
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Index Netta, 465 Netta peposaca (Rosy-Billed Pochard), 330 Nettapus auritus (African Pygmy-Goose), 329 Nettapus coromandelianus (Cotton Pygmy-goose), 329, 357 Netta rufina (Red-crested Pochard), 66, 330, 359, 364 neural larva migrans (NLM), 425 New Britain Hawk-Owl (see Ninox odiosa) New Zealand Kaka (see Nestor meridionalis) New Zealand white rabbit (Oryctolagus cuniculus), 302 nicarbazin, 177 niclosamide, 255, 272, 424 Ninox odiosa (New Britain Hawk-Owl), 401 nodular typhlitis, 404 Noisy Friarbird (see Philemon corniculatus), 22 Noisy Miner (see Manorina melanocephala), 22 nonhematophagous dipteran larvae, 550 Northern Bobwhite (see Colinus virginianus) Northern Cardinal (see Cardinalis cardinalis) Northern Flicker (see Colaptes auratus) Northern fowl mite (Ornithonyssus sylviarum), 534 Northern Goshawk (see Accipiter gentilis) Northern Harrier (see Circus cyaneus) Northern Lapwing (see Vanellus vanellus) Northern leopard frog (see Rana pipiens), 303 Northern Long-eared Owl (see Asio otus) Northern Mockingbird (see Mimus polyglottos) Northern Pintail (see Anas acuta) Northern Saw-whet Owl (see Aegolius acadicus) Northern Shoveler (see Anas clypeata) Northern Shrike (see Lanius excubitor) Northern water snake (Nerodia sipedon), 302 Northiella haematogaster (Bluebonnet), 399 North Island Brown Kiwi (see Apteryx mantelli) Nothoprocta, 465 Nothura maculosa (Spotted Nothura), 389, 395, 466 Notocotylidae, 231, 236 Notocotylus, 231 Notocotylus attenuatus, 226, 236, 240 Novyella, 37, 39 Nucifraga caryocatactes (Eurasian Nutcracker), 297, 345, 480 nuclear gene sequences, of avian haemoproteids, 17 Numenius phaeopus (Whimbrel), 451, 521 Numida, 465 Numida meleagris (Helmeted Guineafowl), 113, 155, 168, 331, 388, 391, 396, 473 Numididae (Guineafowl), 59 Nutmeg Mannikin (see Lonchura punctulata) nutrient shortage in early development, effect, 6 Nyctanassa violacea (Yellow-crowned Night-Heron), 115, 238, 295 Nyctibiidae (Potoos), 443 Nycticorax caledonicus (Rufous Night Heron), 424 Nycticorax nycticorax (Black-crowned Night-Heron), 295, 389, 468 Nymphicus hollandicus (co*ckatiel), 398 Ocellated Turkey (see Meleagris ocellata) ocular larva migrans (OLM), 425 Odontophorus capueira (Spot-winged Wood-Quail), 237, 476 Odontoterakis, 394 Oena capensis (Nomaqua Dove), 73 Oligacanthorhynchus species, 279
581
oligochaetes, 301 oligochaete worm (Limnodrilus hoffmeisteri), 300 Olive-backed Pipit (see Anthus hodgsoni) Onchocerca volvulus, 542 Onchocercidae, 440 Onchocercinae, 448 Oncicola canis, 280 Oncicola oncicola, 280 Oniscus species, 327 oocysts, 40, 108 of Cryptosporidium hominis, 199 of Caryospora, Isospora, and Eimeria in feces, 198 contamination of environment, 212 distribution and prevalence of shedding of Eimeria spp., 182–185 of Eimeria reichenowi and Eimeria gruis, 182, 187–189 Eimeria species, 165 of Eimeria species, 176–177 environmental contamination with, 192 from Fish Crow (Corvus ossifragus), 110 of Isospora canaria, 110 of Isospora rothschildi, 109 of Isospora species, 112 species of Cryptosporidium, 196–200 of species of Sarcocystis, 163 sporulated, 212 Toxoplasma, 211–213 Toxoplasma gondii, 211–212 ookinetes, 17 Oosthuizobdella garoui, 505 operculum, 226 Ophidascaris, 425 Ophryocotyle, 263 Ophthalmophagus, 229 Opisthobrachia, 247 Opisthocomidae (Hoatzin), 442 Opisthocomiformes, 59 Opisthorchiidae, 226, 231, 235, 237, 238 Opisthorchis, 231 Opisthorchis felineus, 239 Opisthorchis sp., 233, 237 Opossums (Didelphis spp.), 114 opthaine, 509 oral suckers, 226 Orange Weaver (see Ploceus aurantius) Orange-winged Parrot (see Amazona amazonica) Orchipedidae, 231 Orchipedum, 231 Order Diptera, 451 Order Phthiraptera, 451 Order Tetrabothriidea, 264 Oreoscoptes montanus (Sage Thrasher), 5 Oriolidae (Old World Orioles), 60, 444 Oreortyx, 465 Oriental Turtle-Dove (see Streptopelia orientalis) Orientia tsutsugamushi, 534 Orinoco Goose (see Neochen jubata) ormetoprim-sulfa, 192 Ornithobdella edentula, 505 Ornithobdellidae, 502, 505, 506, 507, 508 Ornithobilharzia spp. Ornithobilharzia, 246, 247, 249, 251 Ornithobilharzia baeri, 251 Ornithobilharzia canalicula, 251 Ornithobilhorzia emberizae, 251
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582 Ornithocapillaria spp., 463 Ornithocapillaria appendiculata, 467 Ornithocapillaria carbonis, 466, 467 Ornithocapillaria cylindrica, 472 Ornithocapillaria ovopunctata, 479, 480, 482, 483, 484 Ornithocapillaria phalacrocoraxi, 467 Ornithocapillaria picorum, 478 Ornithocapillaria quiscali, 481, 483, 485 Ornithomyia aviculria, 18 ornithophilic black flies, bifid claws of, 538 Ornithosis bedsoniae, 521 Ortalis vetula (Plain Chachalaca), 331 Oryctolagus cuniculus (New Zealand white rabbit), 302 Oscelloscia floridana, 327 Osprey (see Pandion haliaetus) Ostrich (see Struthio camelus) Oswaldoia, 229 Otididae (Bustards), 59, 442 Otiditaenia spp., 268 Otiditaenia conoideis, 264, 269 Otiditaenia macqueeni, 264, 269 Otis, 465 Otis tarda (Great Bustard), 393 Otus scops (European Scops-Owl), 133 oviduct ruptures, 234 ovoid merozoites, 39 oxfendazole, 406 oxyclozanide, 240 Oxyruncidae (Cotingas), 444 Oxytrema silicula, 227 Oxyura jamaicensis (Ruddy Duck), 68, 227, 236, 331, 336, 359, 365 Oxyura leucocephala (White-headed Duck), 360, 365 Pachytrema, 231 Pacific Black Duck (see Anas superciliosa) Padda oryzivora (Java Sparrow), 24 Painted turtle (Chrysemys picta), 303 Palearctic ecozone, 20 Pale-backed Pigeon (see Columba eversmanni) Pallid Harrier (see Circus macrourus) Palm Tanager (see Thrgupis palmarum) Palm-Swift (see Cypsiurus parvus) panacur, 437 Pandion haliaetus (Osprey), 80, 236, 472 Pandionidae (Ospreys), 59, 441 Parabasalia, phylum, 154 Paracapillaria, 463 Paracapillaroides, 463 Paradilepis delachauxi, 264 Paradilepis scolecina, 264, 268 Paradilepis sp., 264 Paradisaeidae (Birds of Paradise), 61, 445 Paradise Tanager (see Tangara chilensis) Paradoxornithidae (Parrotbills), 60 Paraguayan caiman (Caiman yacare), 303 Parahaemoproteus, 15 Parakeet (species not reported), 23 Paramonostomum, 231 Paramphistomatidae, 231, 236 Parapronocephalum, 231 parasitism cost of, 4, 6 and distribution of effects of parasites, 6 energy as a cost parameter against, 4 and predation, 5–6
Index study of, in wild birds, 3 in terms of widlife diseases, 3 Paraspidodera, 394 Parastrigea, 231 Parastrigea tulipoides, 237 Paratanaisia, 230 Paratanaisia bragai, 228, 237, 238 Paratrichosoma, 463 Paridae (Chickadees and tit*), 61, 444 paromomycin, 200 Paronchocerca spp., 440, 441–445, 446, 448, 452, 454 Paronchocerca bambusicolae, 449 Paronchocerca bumpae, 449 Paronchocerca ciconarium, 454 Paronchocerca ciconiarum, 449, 455 Paronchocerca francolina, 449 Paronchocerca helicina, 450 Paronchocerca ibanezi, 449 Paronchocerca limboonkengi, 449 Paronchocerca mansoni, 449 Paronchocerca mirzai, 449 Paronchocerca rousseloti, 449, 450 Paronchocerca schelupovi, 449 Paronchocerca sonini, 449 Paronchocerca straeleni, 449 Paronchocerca struthionus, 456 Paronchocerca tonkinensis, 449 Parorchis, 230, 231 Parorchites zederi, 264, 268 Partridge (Perdix spp), 168 Parulidae (New World Warblers), 60, 435, 444 Parus major (Great Tit), 5, 47, 207, 482 Parus monticolus (Green-backed Tit), 482 Paruterina, 265 Paruterinidae, 264 Passer, 465 Passer spp. Passer domesticus biblicus (Israeli House Sparrow), 24 Passer domesticus (House Sparrow), 22, 206, 209, 333, 345, 401, 452, 530 Passer hispaniolensis (Spanish Sparrow), 482 Passer montanus (Eurasian Tree Sparrow), 111, 206, 209 Passerella, 465 Passerella iliaca (Fox Sparrow), 482 Passeridae (Old World Sparrows), 60 Passeriformes (perching birds), 14, 22, 24, 60, 123, 136, 163, 170, 206, 208, 218, 238, 247, 252, 253, 277, 297, 332, 376, 377, 378, 435, 443, 465, 479, 548, 549 Passerini’s Tangara (see Ramphocelus passerinii) Passeromyia heterochaeta, 548 Passeromyia indecora, 548 Pasteurella multocida, 521 Pastor roseus (Rosy Starling), 484 Patagioenas spp. Patagioenas fasciata (Band-tailed Pigeon), 70, 125, 143, 398 Patagioenas leucocephala (White-crowed Pigeon), 125 Patogioenas speciosa (Scaled Pigeon), 398 Pavo cristatus (Indian Peafowl), 155, 331, 392, 396, 402 Pavo muticus (Green Peafowl), 391 Pavoncella, 465 Pearsonema, 463 pediculosis, 515 Pedionomidae (Plains-wanderer), 442
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Index Pegosomum sp., 235 Pelecanidae (Pelicans), 441 Pelecaniformes, 14, 59, 163, 173, 174, 217, 235, 247, 251, 263, 292, 328, 441, 465, 466 Pelecanoididae (Diving-Petrels), 441 Pelecanus spp., 465 Pelecanus conspicillatus (Australian Pelican), 467 Pelecanus erythrorhynchos (American White Pelican), 293, 467 Pelecanus occidentalis (Brown Pelican), 235, 270, 293, 467 Pelecanus rufescens (Pink-backed Pelican), 270, 293 Pelecitus spp., 440, 441–445, 446, 448, 450–452, 454 Pelecitus ceylonensis, 451 Pelecitus fulicaeatrae, 445, 446, 451, 458, 521 Pelecitus tubercauda, 458 penguins (see Spheniscidae) Perch (Perca fluviatilis), 298 Perdix, 465 Perdix perdix (Gray Partridge), 6, 35, 155, 156, 205, 217, 331, 391, 396, 405, 473 Perdix spp., 168 Peregrine Falcon (see Falco peregrinus) Periparus ater (Coal Tit), 376 permethrin spray, 535 Pernis apivorus (European Honey-Buzzard), 79, 550 Phaenicopteridae (Flamingos), 441 Phaethontidae (Tropicbirds), 441 Phagicola, 230 Phagicola longa, 235 phagocytosis, of trichom*onads, 141 Phalacrocoracidae (Cormorants), 59, 441 Phalacrocorax spp., 292, 416, 465 Phalacrocorax africanus (Long-tailed Cormorant), 292 Phalacrocorax aristotelis (European Shag), 227 Phalacrocorax auritus (Double-crested Cormorant), 173, 174, 176, 233, 235, 270, 292, 421, 466 Phalacrocorax bougainvillii (Guanay Cormorant), 270 Phalacrocorax brasilianus (Neotropic Cormorant), 467 Phalacrocorax carbo (Great Cormorants), 264, 292, 300, 328, 467 Phalacrocorax fuscescens (Black-faced Cormorant), 292, 467 Phalacrocorax melanoleucos brevirostris (Pied Cormorant), 290 Phalacrocorax melanoleucos (Little Pied Cormorant), 292, 467 Phalacrocorax pygmaeus (Pygmy Cormorant), 292, 304, 467 Phalacrocorax sulcirostris (Little Black Cormorant), 467 Phalacrodectes, 529 phanerozoites, 39 Phapitreron leucotis (White-eared Dove), 71 Phasianidae (Pheasants and Partridges), 59, 360, 442, 505 Phasianus spp., 154, 465 Phasianus colchicus (Ring-necked Pheasant), 6, 114, 154, 155, 163, 168, 169, 205, 213, 214, 233, 237, 331, 345, 347–350, 388, 392, 396, 405, 474, 488 Phasianus versicolor (Green Pheasant), 270, 391 Pheucticus ludovicianus (Rose-breasted Grosbeak), 333 Philenon corniculatus (Noisy Friarbird), 22 Philepittidae (Asities), 61, 444 Philippine Duck (see Anas luzonica) Philomachus, 465 Philomachus pugnax (Ruff), 332 Philophthalmidae, 231, 236, 238
583
Philophthalmus spp., 231, 234, 236 Philophthalmus gralli, 234, 238, 240 Philophthalmus hegeneri, 238, 239 Philophthalmus megalurus, 239 Philopteridae, 516 Philopterus sp., 517 Philornis spp., 546, 553 Philornis carinatus, 548 Philornis deceptivus, 549 Philornis downsi, 549 Philornis mimicola, 549 Philornis pici, 549 Philornis porteri, 549 Philornis seguyi, 549 Phimosus infuscatus (Bare-faced Ibis), 296 Phoca caspica (Caspian seal), 302 Phocascaris, 415 Phoebastria immutabilis (Laysan Albatross), 527, 530, 535 Phoenicopteriformes (Flamingos), 23, 56, 59, 163, 247, 251, 328 Phoenicopterus chilensis (Chilean Flamingo), 328 Phoenicopterus minor (Lesser Flamingo), 23, 114 Phoeniculidae (Woodhoopoes), 443 Phoenicurus, 465 photoperiod, 7 physiological trade-offs, in relation to parasitism, 5 Phytotomidae (Plantcutters), 444 Pica, 465 Pica hudsonia (Black-billed Magpie), 451, 482 Pica pica pica (Eurasian Magpie), 345, 451, 482 Picathartidae (Rockfowl), 61 Picidae (Woodpeckers), 60, 435, 443 Piciformes (Woodpeckers), 14, 60, 163, 169–170, 218, 332, 435, 443, 465, 478, 548 Picui Ground-Dove (see Columbina picui) Piculus rubiginosus (see Golden-olive Woodpecker) Picus canus (see Gray-faced Woodpecker) Picus viridis (see Green Woodpecker) Pied Avocet (see Recurvirostra avosetta) Pied-billed Grebe (see Podilymbus podiceps) Pied Cormorant (see Phalacrocorax melanoleucos brevirostris) Pied Currawong (see Strepera graculina) Pied Imperial-Pigeon (see Duchula bicolor) Pied Stilt (see Himantopus leucocephalus) Pig frog (Rana grylio), 303 Pigeon Guillemot (see Cepphus columba) Pileated Woodpecker (see Dryocopus pileatus) pillbugs, 327 Pine snake (Pituophis melanoleucus), 302 Pink-backed Pelican (see Pelecanus rufescens) Pink co*ckatoo (see Cacatua leadbeateri) Pink-footed Goose (see Anser brachyrhynchus) Pink-necked Pigeon (see Treron vernans) Pink Pigeon (see Nesoenas mayeri) Pinnated Bittern (see Botaurus pinnatus) Pionus spp. Pionus fuscus (Dusky Parrot), 399 Pionus maximiliani (Scoly-headed Parrot), 400 Pionus menstruus (Blue-headed Parrot), 399 piperazine, 491 piperazine dihydrochloride, 424 piperazine sulfate, 371 piperonyl butoxide, 521–522 Pipile jacutinga (Black-fronted Piping-Guan), 395 Pipilo erythrophthalmus (Eastern Towhee), 483 Pipridae (Manakins), 61, 444
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584 Piranga olivaceai (Scarlet Tanager), 483 Piranga rubra (Summer Tanager), 483 piranhas (Serrasalmus nattereri), 290 Piscicapillaria, 463 Pittidae (Pittas), 60, 444 Placobdella spp. Placobdella costata, 505 Placobdella novabritanniae, 505 Placobdella ornata, 502, 505 Placobdella papillifera, 502, 505 Placobdelloides jaegerskioeldi, 501 Placobdelloides maorica, 502, 505 Plagiorchiidae, 231, 237 Plagiorchis, 231 Plagiorchis elegans, 237 Plagiorhynchus cylindraceus, 280, 281, 282, 283 from an American Robin (Turdus migratorius), 279 in an American Robin (Turdus migratorius), 282 Plagiorhynchus species, 277 Plain Chachalaca (see Ortalis vetula) Plain Wren (see Thryothorus modestus) plasmochin, 28 Plasmodiidae, 36 Plasmodioides, 36 Plasmodium spp., 15, 17, 27, 35, 54 from the Australian region, 36 clinical signs of infection, 42–43 domesticated animal health concerns, 47 gold standard for diagnosis of infection, 45–46 immunity, 46–47 infection relapse, 20 infection through blood inoculation, 46 intrinsic and extrinsic factors of distribution, 41 management implications, 48–49 mortality from, 48 natural vectors of, 41 Plasmodium anasum, 37 Plasmodium ashfordi, 37 Plasmodium bertii, 37 Plasmodium cathemerium, 36, 37 infections with, 44 Plasmodium circumflexum, 36, 37, 38, 41 erythrocytic stages of, 38 Plasmodium columbae, 37 Plasmodium coturnixi, 37 Plasmodium dissanaikei, 37 Plasmodium durae, 37 Plasmodium durae (domestic turkeys), 37, 42, 45, 48 Plasmodium elongatum, 36, 37, 40, 41–42 Plasmodium fallax, 37 Plasmodium formosanum, 37 Plasmodium forresteri, 37 Plasmodium gabaldoni, 37 Plasmodium gallinaceum, 37, 39, 40, 41–43, 43, 48 Plasmodium garnhami, 37 Plasmodium giovannolai, 37 Plasmodium (Giovannolaia) sp., 41 Plasmodium griffithsi, 37 Plasmodium gundersi, 37 Plasmodium hegneri, 37 Plasmodium hermani, 37, 40, 41, 47 Plasmodium hexamerium, 37 Plasmodium huffi, 37 Plasmodium juxtanucleare, 37, 41–42 Plasmodium kempi, 36, 37 Plasmodium leanucleus, 37 Plasmodium lophurae, 37
Index Plasmodium lutzi, 37 Plasmodium matutinum, 37 Plasmodium (Novyella) sp., 41 Plasmodium nucleophilum, 36, 37 Plasmodium octamerium, 37 Plasmodium paranucleophilum, 37 Plasmodium parvulum, 37 Plasmodium pedioecetae, 37 Plasmodium pinottii, 37 Plasmodium polare, 37 Plasmodium relictum, 7, 35, 36, 37, 40, 41–42, 44, 45, 46, 48 Plasmodium rouxi, 36, 37, 41 Plasmodium subpraecox, 37 Plasmodium tejerai, 37 Plasmodium vaughani, 36, 37 prevalence of, 41–42 public health concerns, 47 spatial and seasonal patterns of transmission, 40 and stress-mediated changes in the immune system, 40 subgenera and species of avian, 37 treatment and control, 48 wildlife population impacts, 47–48 Platalea, 465 Platalea ajaja (Roseate Spoonbill), 290, 296, 468 Platalea alba (African Spoonbill), 296 Platalea leucorodia (Eurasian Spoonbill), 296 Platycercus elegans (Crimson Rosella), 218, 399 Platycercus eximius (Eastern Rosella), 399 Platyhelminthes, 262 Platynosomum, 229 Plegadis, 465 Plegadis falcinellus (Glossy Ibis), 296 Ploceidae (Weavers), 60, 444 Ploceus qurantius (Orange Weaver), 378 plumage coloration, 21 Plum-headed Finch (see Neochmia modesta) Pluvialis, 465 pneumonitis, 116 Podager nacunda (Nacunda Nighthawk), 401 Podargidae (Frogmouths), 59, 443 Podiceps, 465 Podiceps auritus (Horned Grebe), 262, 292, 328 Podiceps cristatus (Great Crested Grebe), 291, 328 Podiceps grisegena (Red-necked Grebe), 328, 466, 521 Podiceps nigricollis (Eared grebe), 466 Podicipedidae (Grebes), 360, 441, 505 Podicipediformes (Grebes), 14, 56, 59, 163, 252, 263, 291, 328, 360, 441, 465, 466, 521 Podilymbus podiceps (Pied-billed Grebe), 262, 466 podoacariasis, 527 Poecile gambeli (Mountain Chickadee), 4 polar bodies, 112 Polyangium, 230 Polydelphis, 425 polymerase chain reaction (PCR), 25, 41, 46, 109, 140, 425 for Eimeria species, 190 Polymorphus spp., 277, 279 Polymorphus botulus, 285 Polymorphus magnus, 283 Polymorphus mathevossianae, 285 Polymorphus minutus, 280, 282, 283, 285 Polyplectron bicalcaratum (Gray Peaco*ck-Pheasant), 391 Polytelis spp. Polytelis alexandrae (Alexandra’s Parrot), 332, 398 Polytelis anthopeplus (Regent Parrot), 217, 400 Polytelis swainsonii (Superb Parrot), 217
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Index Pomatostomus superciliosus (White-browed Babbler), 483 Porcellionides pruinosus, 327 Porcellio scaber, 327 Porphyrio martinica (Purple Gallinule), 475 Porrocaecum, 339, 413, 415–416, 419, 423, 425 Porrocaecum, host range of species of, 418 Porrocaecum angusticolle, 416, 419 Porrocaecum crassum, 416, 419 Porrocaecum depressum, 415, 416, 419, 421 Porrocaecum ensicaudatum, 416, 419, 421 Port Lincoln Parrot (Barnardius zonarius) Porzana porzana (Spotted Crake), 361 Postharmostomum gallinum, 237 Posthodiplostomum, 229 Prairie Chicken (see Tympanuchus spp.) Prairie Falcon (see Falco mexicanus) praziquantel, 255, 352 predation, 5 preerythrocytic merogony, 19 preerythrocytic meronts, 15 preerythrocytic meronts, from avian hosts, 22–24 preerythrocytic stages, of the parasite, 21 premunition, 46 prepatent period ranges haemoproteids transmitted by Culicoides, 19–20 haemoproteids transmitted by hippoboscid flies, 20 primaquine, 28 primaquine phosphate, 48 Primolius maracana (Blue-winged Macaw), 233 Prionopidae (Helmetshrikes), 60 Procellariidae (Sheawaters and Petrels), 441, 505 Procellariiformes (Albatrosses and Petrels), 14, 56, 59, 163, 173, 174, 441 Profilicollis altmani, 285 Profilicollis botulus, 282 Progne subis (Purple Martin), 28, 543 Pronocephalidae, 231 Prosimulium spp., 543 Prosimulium decemarticulatum, 85 Prosimulium hirtipes, 85 Prosobrachia, 247 prospective case-control studies, 4 Prosthogonimidae, 231, 235, 237 Prosthogonimus, 231 Prosthogonimus cuneatus, 226 Prosthogonimus macrorchis, 237 Prosthogonimus ovatus, 226 Protocalliphora spp., 547, 549, 550, 553 Protocalliphora aenea, 548 Protocalliphora asiovora, 548 Protocalliphora avium, 548, 551 Protocalliphora azurea, 548 Protocalliphora beameri, 548 Protocalliphora bennetti, 548 Protocalliphora bicolor, 548 Protocalliphora braueri, 548 Protocalliphora braueri (Blow Fly) larvae, 5 Protocalliphora brunneisquama, 548 Protocalliphora chrysorrhoea, 548 Protocalliphora cuprina, 548 Protocalliphora deceptor, 548 Protocalliphora downsi, 551 Protocalliphora falcozi, 548 Protocalliphora fallisi, 548 Protocalliphora halli, 548 Protocalliphora hesperia, 548 Protocalliphora hesperioides, 548
585
Protocalliphora hirundo, 548 Protocalliphora interrupta, 548 Protocalliphora isochroa, 548 Protocalliphora lata, 548 Protocalliphora lindneri, 548 Protocalliphora metallica, 548 Protocalliphora occidentalis, 548 Protocalliphora parorum, 548 Protocalliphora peusi, 548 Protocalliphora rugosa, 548 Protocalliphora seminuda, 548 Protocalliphora shannoni, 548 Protocalliphora sialia, 548 Protocalliphora spatulata, 548 Protocalliphora spenceri, 548 Protocalliphora tundrae, 548 proventricular worms, 326 Prunella collaris (Alpine Accentor), 483 Prunella modularis (Dunnock), 483 Prunellidae (Accentors), 60, 444 Psephotus haematonotus (Red-rumped Parrot), 400 Pseudamphimerus, 231 Pseudlemdana, 441–445, 448, 454 Pseudocapillaria, 463 Pseudocapillarioides, 463 Pseudolynchia brunnea, 18 Pseudolynchia canariensis, 18 Pseudomenopon pilosum, 521 Pseudomenopon sp., 521 Pseudomonas aeruginosa, 139 Pseudomonas spp., 434 Pseudophyllidea, 262 Pseudoterranova, 423 Psilochasmus, 231 Psilochasmus oxyurus, 239 Psilostomatidae, 231, 236, 237, 238 Psilostomum, 231 Psittacidae (Parrots), 59, 442 Psittaciformes, 14, 18, 23, 59, 121, 136, 139, 163, 168, 169, 206, 217, 332, 442, 459, 465, 476, 549 Psittacula spp. Psittacula eupatria (Alexandrine Parakeet), 459 Psittacula krameri (Rose-ringed Parakeet), 400 Psittacula longicauda (Long-tailed Parakeet), 400 Psittacula roseata (Blossom-headed Parakeet), 13, 23 Psophiidae (Trumpeters), 442 Pteroclididae (Sandgrouse), 442 Pterocliformes, 59 Pterocliformes (Sandgrouse), 56 Pteroglossus aracari (Black-necked Aracari), 479 Pteronetta hartlaubii (Hartlaub’s Duck), 329 Pterothominx spp., 463 Pterothominx alpina, 475 Pterothominx blomei, 473, 474 Pterothominx bursata, 463, 465, 466, 486 Pterothominx caudinflata, 463, 464, 465, 466, 473, 474, 486, 488, 491 Pterothominx exilis, 468, 479, 480, 481, 483, 484, 485 Pterothominx longifilla, 479, 483 Pterothominx meleagridis, 473, 474 Pterothominx moraveci, 464, 476 Pterothominx philippinensis, 464, 488, 490–491 Pterothominx totani, 475, 476 Ptilogonatidae (Silky-flycatchers), 61, 435 Ptilonorhynchidae (Bowerbirds), 61, 445
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586 Ptilonorhynchus violaceus (Satin Bowerbird), 218, 518 Puerto Rican Parrot (see Amazona vittata) Puffinus tenuirostris (Short-tailed Shearwater), 173, 174, 267 Pulchrosoma pulchrosoma, 238 pulmonary arteries, 449 pulmonary congestion, 116 pulmonary edema, 116 Pulmonata, 247 Pumpkin-seed (Lepomis gibbosus), 298 Purple Gallinule (see Porphyrio martinica) Purple Honeycreeper (see Cyanerpes caeruleus) Purple Martin (see Progne subis) Purple Sandpiper (see Colidris maritima) Pycnonotidae, 444 Pycnonotus jocosus (Red-whiskered Bulbul), 218 Pygmy Cormorant (see Phalacrocorax pygmaeus) Pynonotidae (Bulbuls), 61 pyrantel, 371 pyrethrin solution, 544, 554 pyrethrum, 521 pyrimethamine, 28 pyrimethamine (Daraprim), 216, 219 pyrimethamine–sulfadoxine combinations, 28, 48 Pyrrhocorax graculus (Yellow-billed Chough), 483 Pyrrhula spp. Pyrrhula pyrrhula (Eurasian Bullfinch), 116 Pyrrhura leucotis (White-eared Parakeet), 233, 400 Pyrrhura molinae (Green-cheeked Parakeet), 399 Pytilia hypogrammica (Red-Faced Pytilia), 195 Quadraceps ridgwayi, 516 quinine derivatives, 95 Quiscalus spp. Quiscalus major (Boat-Tailed Grackle), 113, 327, 333 Quiscalus mexicanus (Great-tailed Grackle), 209 Quiscalus niger (Greater Antillean Grackle), 333 Quiscalus quiscula (Common Grackle), 209, 451, 454, 483 Raccoon Dog (Nyctereutes procyonoides), 302 Raccoon (Procyon lotor), 425 raccoon roundworm encephalitis, 424 Radjah Shelduck (see Tadorna radjah) Raillietina spp., 263, 264, 268 Raillietina echinobothrida, 271 Raillietina laticanalis, 270 Rainbow Lorikeet (see Trichoglossus haematodus moluccanus) Rallidae (Rails, Gallinules, and Coots), 59, 361, 442, 505 Rameron Pigeon (see Columba arquatrix) Ramphastidae (Toucons), 443 Ramphastos, 465, 516 Ramphastos sulfuratus (Keel-billed Toucan), 114–115 Ramphastos toco (Toco Toucan), 169, 479 Ramphastos vitellinus (Channel-billed Toucan), 479 Ramphocaenus melanurus (Long-billed Gnatwren), 332 Ramphocelus spp. Ramphocelus dimidiatus (Crimson-backed Tanager), 209 Ramphocelus passerinii (Passerini’s Tangara), 111 Ramphocelus carbo (Silver-beaked Tanager), 111 Rana spp. Rana catesbeiana (Bullfrog), 303 Rana megapoda (Bigfoot leopard frog), 303 Rana ridibunda (Lake frog), 303 Raphidascaris, 413 Razorbill (see Alca torda)
Index Recurvirostra, 465 Recurvirostra americana (American Avocet), 476 Recurvirostra avosetta (Pied Avocet), 361 Recurvirostridae (Avocets and Stilts), 59, 361, 442, 505 Red-and-green Macaw (see Ara chloropterus) Red-and-yellow Barbet (see Trachyphous erythrocephalus) Red-bellied Woodpecker (see Melanerpes carolinus) Red-billed Curassow (see Crax blumenbachii) Red-billed Duck (see Anas erythrorhyncha) Red-breasted Goose (see Branta ruficollis) Red-breasted Merganser (see Mergus serrator) Red Collared-Dove (see Streptopelia tranquebarica) Red-crested Bustard (see Eupodotis ruficrista) Red-crested Pochard (see Netta rufina) Red-crowned Crane (see Grus japonensis) Red-eyed Dove (see Streptopelia semitorquata) Red-faced Mouse Bird (see Colius erythromelon) Red-Faced Pytilia (see Pytilia hypogrammica) Red-footed Booby (see Sula sula) Red-fronted Parakeet (see Cyanoramphus novaezelandiae) Red Grouse (see Lagopus lagopus scotica) Redhead (see Aythya americana) Red-headed Woodpecker (see Melanerpes erythrocephalus) Red Junglefowl (see Gallus gallus) Red Kite (see Milvus milvus) Red Knot (see Calidris canutus) Red-legged Honeycreeper (see Cyanerpes cyaneus) Red-legged Partridge (see Alectoris rufa) Red-legged Seriema (see Cariama cristata) Red-lored Parrot (see Amazona autumnalis) Red Lorry (see Eos bornea) red mite (Dermanyssus gallinae), 109 Red-necked Falcon (see Falco chicquera) Red-necked Francolin (see Francolinus afer) Red-necked Grebe (see Podiceps grisegena) Red-necked Parrot (see Amazona arausiaca) Red-rumped Parrot (see Psephotus haematonotus) Redshank (see Tringa totanus) Red-shouldered Hawk (see Buteo lineatus) Red Spurfowl (see Galloperdix spadicea) Red-tailed Hawk (see Buteo jamaicensis) Red-throated Loon (see Gavia stellata) Red-whiskered Bulbul (see Pycnonotus jocosus) Redwing (see Turdus iliacus) Red-winged Blackbird (see Agelaius phoeniceus) Red-winged Parrot (see Aprosmictus erythropterus) Red-winged Tinamou (see Rhynchotus rufescens) Reed Bunting (see Embesiza schoeniclus) Reeves’ Pheasant (see Syrmaticus reevesii) Regent Bowerbird (see Sericulus chrysocephalus) Regent Parrot (see Polytelis anthopeplus) Regulidae (Kinglets), 61 Remizidae (Penduline tit*), 444 renal cryptosporidiosis, 198 Renicola lari, 236 Renicola pinguis (Troglotrematidae), 226 Renicolidae, 236 Renicolla, 231 repair or replace costs, to the host, 4 residuum, 112 resource trade-offs, against parasitism, 5 respiratory cryptosporidiosis, 198 reticuloendothelial cells, in bone marrow, 35 retrospective case–control studies, 4 Rhabdornithidae, 444 Rhadinea merremii (Graceful brown snake), 303 Rhea americana (Greater Rhea), 345, 389, 395, 466
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Index Rheidae (Rheas), 441 Rheiformes, 163, 441 Rhinocryptidae (Topaculos), 443 Rhinonyssus rhinolethrum, 528, 533 Rhizothera longirostris (Long-billed Partridge), 392 Rhynchophthirina, 516 Rhynchotus rufescens (Red-winged Tinamou) 389, 395 Rhynochetidae (Kagu), 442 Ribeiroia ondatrae, 236, 237 Ribeiroia sp., 231, 239 Richardsonianus howensis, 505 Ricinidae, 516 Ricinus sp., 517 Ring-billed Gull (see Larus delawarensis) Ringed Kingfisher (see Megaceryle torquatus) Ring-necked Dove (see Streptopelia capricela) Ring-necked Duck (Aythya collaris) Ring-necked Pheasant (see Phasianus colchicus) Ring Turtle-Dove (see Streptopelia risoria) Ringed Teal (see Collonetta leucophrys) Rissa, 465 River otter (Lutra canadensis), 302 robenidin, 177 Rock Partridge (see Alectoris graeca) Rock pigeon (see Columba livia) Rock Ptarmigan (see Lagopus muta) rodents, 425 Rollulus rouloul (Crested Partridge) ronidazole, 144 Rook (see Corvus frugilegus) Roseate Spoonbill (see Platalea ajaja) Rose-ringed Parakeet (see Psittacula krameri) Ross’ Goose (see Chen rossii) Rostratula benghalansis (Greater Painted-Snipe), 488 Rostratulidae (Painted Snipes), 59, 442 Rostrhamus sociabilis plumbeus (Florida Snail Kite), 236 Rostrhamus sociabilis (Snail Kite), 233 Rosy-Billed Pochord (see Netta peposoca) Rosy Starling (see Pastor roseus) Rouen ducks, 249 Rough-legged Hawk (see Buteo lagopus) Royal Tern (see Thalasseus maximus) Ruddy Duck (see Oxyura jamaicensis) Ruddy Ground-Dove (see Columbina talpacoti) Ruddy Quail-Dove (see Geotrygon montana) Ruddy Shelduck (see Tadorna ferruginea) Rufescent Tiger-Heron (see Tigrisoma lineatum) Ruff (see Philomachus pugnax) Ruffed Grouse (see Bonasa umbellus) Rufous-chested Sparrowhawk (see Accipiter rufiventris) Rufous Night Heron (see Nycticorax caledonicus) Rynchopidae (Skimmers), 442 Sacred Kingfisher (see Todiramphus sanctus) Sage Thrasher (see Oreoscoptes montanus) Sagittariidae (Secretary birds), 59, 441 Sagittarius serpentarius (Secretory-bird), 80, 131 Saker Falcon (see Falco cherrug) salinomycin, 177 Salmonella gallinarum, 521 Salmonella typhimurium, 170 Sandhill Crane (see Grus canadensis) Sarcocystis spp., 25, 163, 204, 214 clinical signs, 116 diagnosis, 116 domestic animal health concerns, 117 epizootology, 113–116
etiology of infection, 113 host range and distribution, 113 immunity, 116 pathogenesis and pathology, 116 public health concerns, 116 Sarcocystis accipitris, 114 Sarcocystis alectoributeonis, 114 Sarcocystis alectorivulpes, 114 Sarcocystis ammodrami, 114 Sarcocystis aramidis, 114 Sarcocystis buteonis, 114 Sarcocystis cernae, 114 Sarcocystis cheeli, 114 Sarcocystis citellibuteonis, 114 Sarcocystis colii, 114 Sarcocystis dispersa, 114 Sarcocystis espinosai, 114 Sarcocystis falcatula, 113, 114, 115–116, 214 in Brown-headed Cowbird (Molothrus ater), 115 from feces of Virginia Opossum (Didelphis virginiana), 115 like infections, 214 in Lory (Lorinus sp.) elongate meront of, 115 Sarcocystis garzettae, 114 Sarcocystis glareoli, 114 Sarcocystis horvathi, 114 Sarcocystis jacarinae, 114 Sarcocystis kaiserae, 114 Sarcocystis kirmsei, 114 Sarcocystis nontenella, 114 Sarcocystis oliverioi, 114 Sarcocystis peckai, 114 Sarcocystis phoeniconaii, 114 Sarcocystis ramphastosi, 114 Sarcocystis rauschorum, 115 Sarcocystis rileyi, 115 Sarcocystis scotti, 115 Sarcocystis sebeki, 115 Sarcocystis setophagae, 115 Sarcocystis spaldingae, 115 Sarcocystis sulfuratusi, 115 Sarcocystis wenzeli, 115 synonyms, 113 that parasitize birds, 114–115 treatment and control, 117 wildlife population impacts, 117 sarcocystosis, 116 Sarconema, 441–445, 448, 454 Sarconema eurycera, 520, 521 Sarconema eurycerca, 450, 451, 458 Sarcophagidae (flesh flies), 550 Sarcoramphus papa (King Vulture), 390 Sarkidiornis melanotos (Comb Duck), 329, 362 Sarus Crane (see Grus antigone) Satin Bowerbird (see Ptilonorhynchus violaceus) Satyr Tragopan (see Tragopan satyra) Scaled Pigeon (see Patagioenas speciosa) Scaled Quail (see Callipepla squamata) Scaly-headed Parrot (see Pionus maximiliani) Scarlet-chested Parrot (see Neophema splendida) Scarlet Macaw (see Ara macao) Scarlet Tanager (see Piranga olivaceai) Schistocephalus, 267 Schistocephalus solidus, 264, 267 Schistogonimus, 231 Schistosoma japonicum, 247 Schistosomatidae, 231, 246
587
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Index
Schistosome dermatitis, 255 schistosomes anatomical locations of infection, 249 clinical signs, 249 diagnosis, 254 domesticated animal health concerns, 255 epizootiology, 248–249 etiology, 247–248 history, 246 host range and distribution, 246–247 host responses to eggs, 250 immunity, 254 known intermediate and final hosts of avian, 247 location of the parasite in the host, geographic location, vertebrate host order, and selected references, 251–253 male and female, 248 managment implications, 256 pathogenesis and pathology, 249–254 public health concerns, 255 synonyms, 246 treatment and control, 255–256 wildlife population impacts, 255 schistosomiasis, 246, 254 Schistotaenia spp., 268 Schistotaenia scolopendra, 264, 268 Schistotaenia srivastavi, 264, 268 Schistotaenia tenuicirrus, 262, 264, 268 Schulmanela, 463 scolices, of cyclophyllidean species, 268 Scolopacidae (Sandpipers), 59, 361, 442, 505 Scolopax minor (American Woodco*ck), 464, 550 Scopidae (Hamerkop), 441 Secretary-bird (see Sagittarius serpentarius) Sedge Warbler (see Acrocephalus schoenobaenus) See-see Partridge (see Ammoperdix griseogularis) sentinel birds, 86 Sericulus chrysocephalus (Regent Bowerbird), 218 Serinus canaria (Island Canary), 6, 108, 114, 204, 212, 216, 218 serologic tests, 204 Serratospiculiasis, 436 Serratospiculoides spp. clinical signs, 436 diagnosis, 437 epizootiology, 435–436 etiology, 435 history, 434 host range and distribution, 434–435 pathology, 436 public and domestic animal health concerns, 437 Serratospiculoides alii, 435 Serratospiculoides amaculata, 434 treatment and control, 437 wildlife population impacts, 437 Serratospiculum spp. clinical signs, 436 diagnosis, 437 epizootiology, 435–436 etiology, 435 history, 434 host range and distribution, 434–435 pathology, 436 public and domestic animal health concerns, 437 Serratospiculum amaculata, 434
Serratospiculum amaculatum, 435 treatment and control, 437 wildlife population impacts, 437 Setophaga ruticilla (American Redstart), 115 sexual cycle, of Haemoproteus species, 17 Seychelles Blue-Pigeon (see Alectroenas pulcherrima) Sharp-shinned Hawk (see Accipiter striatus) Sharp-tailed Grouse (see Tympanuchus phasianellus) Sheather’s sugar flotation, 116, 176 Shikra (see Accipiter badius), 130 Short-eared Owl (see Asio flammeus) Short-tailed Shearwater (see Puffinus tenuirostris) Short-toed Eagle (see Circaetus gallicus) Sialia mexicana (Western Bluebird), 282 Sialia sialis (Eastern Bluebird), 333, 483 Siamese Fireback (see Lophura diardi) Siberian Rubythroat (see Luscinia calliopei) Silver-beaked Tanager (see Ramphocelus carbo) Silver Gull (see Larus novaehollandiae) Silver Pheasant (see Lophura nycthemera) Silver Teal (see Anas versicolor) Simocephalus vetulus, 335 Simuliidae, 451 Simulium spp. Simulium adersi, 85 Simulium anatinum, 85 Simulium angustitarse, 85 Simulium annulus, 538, 542 Simulium aureum, 85 Simulium congareenarum, 85 Simulium croxtoni, 85 Simulium equinum, 541 Simulium fallisi, 85 Simulium impukane, 85 Simulium innocens, 85 Simulium jenningsi, 85 Simulium latipes, 85 Simulium lineatum, 541 Simulium meridionale, 85, 543 Simulium metatarsale, 85 Simulium minus, 85 Simulium nyasalandicum, 85 Simulium parnassum, 85 Simulium pictipes, 85 Simulium quebecense, 85 Simulium rendalense, 85 Simulium rugglesi, 85, 539 Simulium slossanae, 85 Simulium slossonae, 87 Simulium venustum, 85 Simulium vittatum, 85, 541 Simulium vittatum, larva of, 539 Simulium vittatum, pupa of, 540 Simulium vorax, 85 Sitta europaea (Eurasian Nuthatch), 207, 483 Sittidae (Nuthatches), 61, 435, 444 Skrjabinocapillaria, 463 Slaty-breasted Wood-Rail (see Aramides saracura) Small-billed Tinamou (see Crypturellus parvirostris) Smew (see Mergellus albellus) Smokey jungle frog (Leptodactylus pentadactylus), 303 Smooth-billed Ani (see Crotophaga ani) Snail Kite (see Rostrhamus sociabilis) Snapping turtle (Chelydra serpentina), 303 sneezing, 197 Snow Goose (see Chen caerulescens)
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Index Snowy Egret (see Egretta thula) Snowy Owl (see Bubo scandiacus) Snowy Plover (see Charadrius alexandrinus) sodium chloride, 509 sodium nitrate, 490 sodium pentachlorophenate, 255 Solitary Tinamou (see Tinamus solitarius) Somateria spp., 465 Somateria fischeri (Spectacled Eider), 67, 359 Somateria mollissima (Common Eider), 5, 67, 165, 166, 174, 261, 282, 330, 336, 356, 362, 367, 470 intestine of, 169 Somateria spectabilis (King Eider), 327, 330, 358, 470 Song Thrush (see Turdus philomelos) Sooty Grouse (see Dendragapus fuliginosus) Southern Crowned-Pigeon (see Goura scherpmakeri) Southwellina dimorpha, 285 Southwellina hispida, 279 sow bugs (Porcellio scaber), 402 Spanish Sparrow (see Passes hispaniolensis) sparganosis, 271 Species Survival Plan, for the Bali Myna, 117 Spectacled Duck (see Anas specularis) Speckled Pigeon (see Columba quinea) Speckled Teal (see Anas flavirestris) Spectacled Eider (see Somateria fischeri) Spelotrema, 230 Sphaeridiotrema, 231 Sphaeridiotrema globulus (Psilostomatidae), 225, 226, 227–228, 236, 238–239, 240 Sphaerirostris, 279 Spheniscidae (Penguins), 59, 441, 505 Sphenisciformes, 14, 59, 163, 174, 175, 217, 235, 291, 441 Spheniscus spp. Spheniscus demersus (Jackass Penguin), 217, 291 Spheniscus humboldti (Humboldt Penguin), 217 Spheniscus magellanicus (Magellenic Penguin), 217, 416 Sphyrapicus varius (Yellow-bellied Sapsucker), 332 Spinicauda, 394 Spiny softshell turtle (Apalone spinifer), 303 Spirometra pleurocercoids, 270, 271 Spirometra sp., 264, 267 Spiroptera spp. Spiroptera crassicauda, 337 Spiroptera nasuta, 327 Spiroptera pectinifera, 337 Spiroptera uncinata, 335 Spirurida, 439 Spizaetus cirrhatus (Changeable Hawk-Eagle), 76 spleen enlargement, 116 Splendidofilaria spp., 440, 441–445, 445, 446, 448, 451, 452, 454 Splendidofilaria algonquinensis, 449, 452 Splendidofilaria alii, 449 Splendidofilaria bohmi, 450 Splendidofilaria brevispiculum, 449 Splendidofilaria californiensis, 446, 449, 451 Splendidofilaria caperata, 449, 452, 457 Splendidofilaria columbensis, 450 Splendidofilaria falconis, 449 Splendidofilaria fallisensis, 445, 446, 450, 451, 540 Splendidofilaria gedoelsti, 450
Splendidofilaria grettillati, 449 Splendidofilaria hibleri, 450 Splendidofilaria longicaudata, 449 Splendidofilaria kasmirensis, 449 Splendidofilaria mavis, 450 Splendidofilaria osmaniae, 449 Splendidofilaria pachacuteci, 449 Splendidofilaria pavlovskyi, 449 Splendidofilaria pectoralis, 450 Splendidofilaria periarterialis, 449 Splendidofilaria picacardina, 445, 446, 449, 451 Splendidofilaria rotundicephala, 450 Splendidofilaria singhi, 450 Splendidofilaria skrjabini, 449 Splendidofilaria smithi, 449, 450 Splendidofilaria travassosi, 449 Splendidofilaria tuvensis, 450 Splendidofilaria verrucosa, 449 Splendidofilaria wehri, 449 Splendidofilariinae, 448 splenic artery, 449 splenomegaly, 91 sporocyst of avian schistosomes, 248 sporocysts, 108, 164 of Sarcocystis, 198 trematodes, 226 sporogony, 40 of Eimeria gruis and Eimeria reichenowi, 187 Eimeria species, 164–165 of Isospora species, 110 Sporozoasida, 204 sporozoites, 17, 39, 82, 164, 187–188 development of, 19 of Isospora species, 110 species of Cryptosporidium, 196–197 Spot-billed Duck (see Anas poecilorhyncha) Spotless Starling (see Sturnus unicolor) Spotted Crake (see Porzana porzana) Spotted Dove (see Streptopelia chinensis) Spotted Eagle-Owl (see Bubo africanus) Spotted Eagle (see Aquila clanga) Spotted Imperial-Pigeon (see Ducula carole) Spotted Nothura (see Nothura maculosa) Spotted Owl (see Strix occidentalis) Spotted Sandpiper (see Actitis macularius) Spot-winged Wood Quail (see Odontophorus capueira) spring relapse, 40, 83 Stabler-gallinae (SG) strain, 134 Steatornithidae (Oilbirds), 443 Stephanoprora, 230 Stercorariidae (Skuas), 442 Sterna, 465 Sterna hirundo (Common Tern), 205, 332 Sternidae (Terns), 59 Sternostoma trachaecolum, 528, 533 Sternostoma trachaecolum, dorsal and ventral views of adult female, 530 Stictonetta naevosa (f*ckled Duck), 357 Stilbometopa impressa, 17, 18 Stock Dove (see Columba oenas) Stomylotrema, 231 Stomylotrematidae, 231 Straw-necked Ibis (see Threskiornis spinicollis) Strepera graculina (Pied Currawong), 24
589
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590
October 16, 2008
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Index
Streptocara spp., 337 clinical signs, 338 diagnosis, 338–339 domestic animal health concerns, 339 eggs of, 337–338 epizootiology, 337–338 etiology, 337 history, 337 host range and distribution, 326, 328–333 life cycle of S. incognita, 337 pathogenesis and pathology, 338 Streptocara crassicauda, 326, 327, 328, 338 Streptocara crassicauda anseri, 337 Streptocara crassicauda charadrii, 337 Streptocara crassicauda crassicauda, 337 Streptocara crassicauda skrjabini, 337 Streptocara incognita (SI), 328 Streptocara pectinifera, 337 synonyms, 337 treatment and control, 339–340 wildlife population impacts, 339 Streptocariasis, 337 Streptococcus equinus, 521 Streptococcus fecalis, 157 Streptopelia spp., 168, 465 Streptopelia bitorquata (Island Collared-Dove), 73 Streptopelia capicola (Ring-necked Dove), 73 Streptopelia chinensis (Spotted Dove), 71, 126, 208, 398 Streptopelia decaocto (Eurasian Collared Dove), 73, 125, 143, 205, 361, 394, 398 Streptopelia decipiens (African Mourning Dove), 73 Streptopelia lugens (Dusky Turtle-Dove), 73 Streptopelia orientalis (Oriental Turtle-Dove), 72, 398 Streptopelia picturata (Madagascar Turtle-Dove), 126, 134 Streptopelia risoria (Ring Turtle-Dove), 125, 136, 140–142 esophageal junction of a, 140, 141, 142 Streptopelia semitorquata (Red-eyed Dove), 71, 125 Streptopelia senegalensis (Laughing Dove), 72, 114, 126, 206 Streptopelia tranquebarica (Red Collared Dove), 73 Streptopelia turtur (Eurasian Turtle-Dove), 72, 398 stress-mediated changes in the immune system, post malarial infection, 40 Striated Heron (see Butorides striata) Striatofilaria, 441–445, 448, 454 Strigea, 231 Strigea falconis, 237 Strigea spp., 226 Strigeidae, 231, 236, 237 Strigidae (Owls), 59, 443 Strigiformes, 14, 24, 59, 163, 206, 208, 217, 443, 477, 548, 549 Strix spp. Strix aluco (Tawny Owl), 21, 115, 133, 206, 208, 477, 550 Strix occidentalis (Spotted Owl), 477 Strix uralensis (Ural Owl), 477 Strix varia (Barred Owl), 133, 206, 213, 216–217, 346, 477 strobila, 269 Strongyluris, 394 Strongylus spp. Strongylus mucronatus, 355 Strongylus pergracilis, 316 Strongylus serratus, 316 Struthio camelus (Ostrich), 57, 154, 196, 208, 240, 395, 516
Struthiofilaria, 441–445, 448, 454 Struthiofilaria megalocephala, 457 Struthionidae (Ostriches), 59, 441 Struthioniformes, 14, 59, 163, 208, 441, 466 Sturgeon (Acipenser gueldenstaedtii), 301 Sturnella magna (Eastern Meadowlark), 379 Sturnella neglecta (Western Meadowlark), 284, 379 Sturnia philippensis (Chestnut-cheeked Starling), 484 Sturnidae (Starlings), 61, 108, 435, 444 Sturnus, 465 Sturnus unicolor (Spotless Starling), 484 Sturnus vulgaris (European Starling), 206, 209, 281, 333, 345, 454, 484, 518, 534 subclinical impacts, of Haemoproteus infections, 28 subcutaneous tissues, 450 “sucking worm.” see Trematodes Sula sula (Red-footed Booby), 217 sulfachloropyrazine, 48 sulfachlorpyrazine (ESB3), 117 sulfachlorpyridazine (Vetisulid), 117 sulfadiazine, 216, 219 sulfadimethoxine, 192 sulfamethazine, 172 sulfamethoxazole, 95 sulfamonomethoxine, 48 sulfaquinoxaline, 172 sulfathiazole, 509 Sulidae (Boobies and Gonnets), 441 Summer Tanager (see Piranga rubra) sunfish (Centrarchidae), 290 Sun Parakeet (see Aratinga solstitialis) Superb Parrot (see Potytelis swainsenii) Superb Starling (Lamprotornis superbus) Surf Scoter (see Melanitta perspicillata) Swainson’s Hawk (see Buteo Swainsoni) Swamp Francolin (see Francolinus gularis) Swan Goose (see Anser cygnoides) swans (see Cygnus spp.) swimmer’s itch, 246, 255 Sylviidae (Old World Warblers), 61, 435 syncytia, 82 syngamiasis, 343 Syngamidae, 343 Syngamus spp., 345 free-living hosts of, 347 prevalence of S. trachea in young rooks, 348 Syngamus alcyone, 345 Syngamus gibbocephalus, 345 Syngamus microspiculum, 345 Syngamus palustris, 345 Syngamus taiga, 345 Syngamus trachea, 343, 344, 347, 348, 349, 350, 351 Syngamus trachealis, 343 Syrmaticus reevesii (Reeves’ Pheasant), 392 Syrmaticus soemmerringii (Copper Pheasant), 391 Tachybaptus, 465 Tachybaptus ruficollis (Little Grebe), 291, 360, 466 Tachycineta albilinea (Mangrove Swallow), 543 Tachycineta bicolor (Tree Swallow), 553 Tachymarptis melba (Alpine Swift), 6 Tachyphonus rufus (White-lined Tanager), 111, 380 tachyzoite, 210, 211, 212 Tadorna, 465 Tadorna ferruginea (Ruddy Shelduck), 329, 359, 365 Tadorna radjah (Radjah Shelduck), 359 Tadorna tadorna (Common Shelduck), 357, 363, 389
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Index Taeniopygia guttata (Zebra Finch), 113, 378 Taiga Bean-Goose (Anser fabalis), 390 Tambourine Dove (see Turtur typanistria) Tangara spp. Tangara cayana (Burnished-buff Tanager), 111 Tangara chilensis (Paradise Tanager), 111 Tangara mexicana (Turquoise Tanager), 111 Tangara schrankii (Green-and-gold Tanager), 111 tapeworms. see Cestodes Tataupa Tinamou (see Crypturellus tataupa) Tawny Eagle (see Aquila rapax) Tawny Owl (see Strix aluco) T-cell-mediated immune response, 6 temephos, 94 Tenoranema, 463 Terranova sp., 423 Terek Sandpiper (see Xenus cinereus) Tetrabothriidea, 262, 266 Tetrabothriidean scolices, 262 Tetrabothrius skoogi, 264, 269 Tetrabothrius spp., 263, 264, 267 tetracyclines, 28 Tetrameres spp., 337, 376 Tetrameres americana, 379, 381 Tetrameres cardinalis, 378 Tetrameres crami, 381 Tetrameres fissispina, 378, 381 Tetrameres globosa, 378 Tetrameres mohtedai, 379 in fowl, 379 Tetrameres pattersoni, 378, 381 Tetrameres striata, 379 Tetrameres strigiphila, 376 Tetrameres zakharowi, 381 tetrameridosis clinical signs, 379 diagnosis, 380–381 domestic animal health concerns, 381 epizootiology, 377–379 etiology, 377 history, 376 host range and distribution, 376–377 immunity, 381 management implications, 381 pathology, 379–380 public health concerns, 381 treatment and control, 381 wildlife population impacts, 381 tetramisole, 406 Tetrao, 465 Tetrao urogallus (Eurasian Caporcaille), 214, 396, 404, 475 Tetrao parvirostris (Black-billed Capercaillie), 360 Tetrao tetrix (Black Grouse), 390, 396, 475 Tetrao urogallus (Eurasian Capercaillie), 215, 360, 391, 396, 475 Tetraogallus caspius (Caspian Snowco*ck), 390 Tetraogallus himalayensis (Himalayan Snowco*ck), 392, 396 Tetraonidae (Grouse, Ptarmigans, and Prairie Chickens), 59 Tetrathyridia (Mesocestoides), 269 Tetratrichom*onas gallinarum, 140 Tetrax tetrax (Little Bustard), 393 Thalasseus, 465 Thalasseus maximus (Royal Tern), 238 Thapariella, 231 Thapariellidae, 231 Thayer’s Gull (see Larus thayeri)
591
Thermocyclops hyalinus, 384, 385 Thermocyclops taihokuensis, 385 Theromyzon spp. Theromyzon affinis, 505 Theromyzon bifarium, 505, 507 Theromyzon cooperi, 501, 505, 506, 508 Theromyzon garjaewi, 502, 505, 508 Theromyzon (Glossiphoniidae), 503 Theromyzon maculosum, 502, 505, 507 Theromyzon matthaii, 502, 505, 508 Theromyzon mollissimum, 505 Theromyzon pallens, 505 Theromyzon propinquum, 502, 505, 508 Theromyzon rude, 505, 507 Theromyzon sexoculatum, 505 Theromyzon tessellatoides, 505 Theromyzon tessulatum, 501–502, 504, 505, 507, 509 Theromyzon trizonare, 502, 504–507, 505 thiabendazole, 339, 352, 424 thick-walled megalomeronts, 19, 21 Thinocoridae (Seedsnipes), 442 thin-walled branching meronts, 22 thin-walled meronts, of Haemoproteus, 39 Thominx, 463 Thomix tridens, 473, 478, 479, 481, 482, 483, 484, 485 Thraupidae (Tanagers), 60, 435 Thraupis episcopus (Blue-gray Tanager), 111, 209, 333, 451 Thraupis palmarum (Palm Tanager), 206, 209 threadworm, 463 Three-toed amphiuma (Amphiuma tridactylum), 303 Threskiornis melanocephalus (Black-headed Ibis), 296 Threskiornis spinicollis (Straw-necked Ibis), 215 Threskiornithidae (Ibises and Spoonbills), 59, 441 Thryothorus ludovicianus (Carolina Wren), 332 Thryothorus modestus (Plain Wren), 208 Timaliidae (Babblers), 61 Tinamidae (Tinamou), 441 Tinamiformes (Tinamous), 14, 56, 59, 163, 441, 465, 466, 517 Tinamus solitarius (Solitary Tinamou), 389 Tinamus tao (Gray Tinamou), 389, 466 tissue injury cost, to the host, 4 Toco Toucan (see Ramphastos toco) Todidae (Todies), 443 Todiramphus sanctus (Sacred Kingfisher), 24 toltrazuril (Baycox), 117, 200 Torgos tracheliotus (Lappet-faced Vulture), 74 Torresian Imperial-Pigeon (see Ducula spilorrhoa) toucans (Ramphastos spp.), 516 Toxascaris leonina, 425 Toxocara spp., 425 Toxoplasma gondii, 204 clinical signs, 212 developmental stages of, 210–211 diagnosis, 214–215 distribution and host range, 204 DNA of, 214 epizootiology, 211–212 etiology, 204–211 history, 204 isolated strains, 210 isolation of Toxoplasma gondii from tissues of naturally infected wild birds, 205–207 isolation of, from tissues of naturally infected wild birds, 205–207 life cycle of, 211 oocysts, brain of birds fed, 214
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592 Toxoplasma gondii (Cont.) oocysts of, 211–212 pathogenesis and pathology, 212–213 public health and domestic animal concerns, 215 serologic prevalence of antibodies, 208–209 tachyzoites, 216 tissue cysts of, 214 tissue necrosis and, 212 treatment and control, 216–219 wildlife population impacts, 215–216 Toxoplasma gondii-specific antibodies, 211 Toxoplasma-like parasites in sparrows, 204 toxoplasmosis, 204, 215 toxoplasmosis, at necropsy from birds that died in captivity (C) or in the wild (W)., 217–218 trachea/esophagus, 449 tracheal worms clinical signs, 349 diagnosis, 351 distribution and host ranges, 344 domestic animal health concerns, 351–352 eggs of, 347, 350 etiology, 343–344 history, 343 immunity, 351 infection, 343 life cycle and epizootiology, 344–349 pathogenesis and pathology, 349–351 public health concerns, 351 synonyms, 343 treatment and control, 352 wildlife population impacts, 351 Trachyphonus erythrocephalus (Red-and-yellow Barbet), 345 trade-offs, against parasitism, 5, 7, 28 Tragopan satyra (Satyr Tragopan), 393 Travassosascaris, 425 Tree Pipit (see Anthus trivialis) Tree Swallow (see Tachycineta bicolor) trematodes anatomical location in avian hosts, 229–232 air sacs, 233 bile ducts and gall bladder, 233 eyes, 234 gastrointestinal tracts, 233–234 kidneys, 233 oviduct, 234 clinical signs, 228 density-dependent effects, 227 diagnosis, 234–239 digenetic, 225, 226 life cycle of, 228 domestic animal health concerns, 239 epizootiology, 226–228 etiology, 225–226 history, 225 host range and distribution, 226 immunity, 239 larval stages of, 227 management implications, 240 migrations and, 226 parasite families and species that can cause disease in wild birds, 235–238 pathogenicity of a, 228 pathology, 228–234 public health concerns, 239 synonyms, 225
Index treatment and control, 240 wildlife population impacts, 239–240 Treron spp. Treron australis (Madagascar Green-Pigeon), 71 Treron calvus (African Green-Pigeon), 397 Treron phoenicopterus (Yellow-footed Pigeon), 398 Treron sieboldii (White-bellied Pigeon), 71 Treron sphenurus (Wedge-tailed Pigeon), 71 Treron vernans (Pink-necked Pigeon), 71 Trichinella sprialis, 491 Trichobilharzia spp., 247, 249, 250, 251, 255 Trichobilharzia alaskensis, 252 Trichobilharzia anatina, 252 Trichobilharzia adamsi, 251 Trichobilharzia arcuata, 249 Trichobilharzia aureliani, 249 Trichobilharzia australis, 249 Trichobilharzia berghei, 252 Trichobilharzia brantae, 252 Trichobilharzia brevis, 252 Trichobilharzia burnetti, 252 Trichobilharzia cameroni, 252 Trichobilharzia cerylei, 252 Trichobilharzia corvi, 252 Trichobilharzia duboisi, 249 Trichobilharzia elvae, 252 Trichobilharzia filiformis, 252 Trichobilharzia franki, 254 Trichobilharzia horiconensis, 252 Trichobilharzia indica, 252 Trichobilharzia jequitibaensis, 252 Trichobilharzia jianensis, 252 Trichobilharzia kegonsensis, 252 Trichobilharzia kossarewi, 252 Trichobilharzia kowalewskii, 252, 253 Trichobilharzia littlebi, 252, 253 Trichobilharzia maegraithi, 252 Trichobilharzia nasicola, 249 Trichobilharzia ocellata, 248–250, 253, 254–255 Trichobilharzia oregonesis, 252, 253 Trichobilharzia paoi, 253, 255 Trichobilharzia parocellata, 252 Trichobilharzia physellae, 253, 255 Trichobilharzia polonica, 252, 253 Trichobilharzia regenti, 247, 249–250 Trichobilharzia rodhaini, 249 Trichobilharzia salmanticensis, 252 Trichobilharzia schoutedeni, 252 Trichobilharzia spinulata, 249 Trichobilharzia stagnicola, 255 Trichobilharzia szidati, 250, 253 Trichobilharzia waubesensis, 252, 253 Trichodectidae, 516 Trichoglossus haematodus moluccanus (Rainbow Lorikeet), 218 Trichom*onadida Order, 154 Trichom*onas foetus, 139 in cattle, 140, 142 Trichom*onas gallinae, 120–121, 142 in birds, 140 in columbiforms, 141 cultured trophozoites of, 134 in free-ranging and captive columbiforms, 122–129 from free-ranging and captive falconiforms and strigiforms, 130–133 life cycle of, 136 pathogenicity of strains of, 134
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Index prevalences of, 136–137 transmission via contaminated water, 136 Trichom*onas gallinae, distribution of, 121 Trichom*onas gallinae, formation of pseudocysts of a strain of, 135 Trichom*onas gallinae, gross lesion (arrow) of, 138 Trichom*onas phasiani, 134 Trichom*onas vagin*lis, 139 in humans, 134, 142 trichom*oniasis, 490 trichom*onosis, 137, 143 clinical signs, 137 diagnosis, 139–141 distribution, 120 domestic animal health concerns, 142–143 epizootiology, 136–137 etiology, 121–136 history, 120 host range, 121 immunity, 141–142 management implications, 145 pathogenesis and pathology, 137–139 public health concerns, 142 synonym, 120 treatment and control, 144–145 wildlife population impacts, 143–144 Trichosomum contorta, 470 Trichosomum contortum, 471 trichostrongyliasis, 316 trichostrongylosis, 316 Trichostrongylus spp. avian species of, 317 clinical signs, 320 diagnosis, 321 domestic animal health concerns, 321–322 epizootiology, 317–320 etiology, 317 history, 316–317 host infection by species of, 317 immunity, 321 impacts of disease, 321 larvae, 317, 320 life cycle of, 317–318 management implications, 322 pathology, 320–321 synonyms, 316 treatment and control, 322 wildlife population impacts, 322 Trichostrongylus cramae, 316–317, 321 Trichostrongylus cramae caudal bursa, 318 Trichostrongylus pergracilis, 316 Trichostrongylus tenuis, 316, 319 caudal bursa, 319 in Red Grouse, 320 Trichostrongylus tenuis caudal bursa, 318 trichurids, 464 Trichuris vulpis, 491 Tricolored Heron (see Egretta tricolor) Tridentocapillaria, 463 Tridentocapillaria eurycerca, 478 Tridentocapillaria hirundinis, 478, 481 Tridentocapillaria parusi, 482, 483 Trigrisoma lineatum (Rufescent Tiger-Heron), 295 Trimenoponidae, 516 trimethoprim, 95 trimethoprim-sulfadiazine, 117 trimethoprim-sulfamethoxazole, 117, 192
593
Tringa, 465 Tringa glareola (Wood Sand piper), 361 Tringa melanoleuca (Greater Yellowlegs), 238 Tringa nebularia (Common Greenshank), 331 Tringa semipalmata inornata (Willet), 476 Tringa totanus (Redshank), 6 Tringa totanus (Common Redshank), 332 Trinoton anserinum, 521 Trinoton anserium, 520 Tritrichom*onas foetus in cattle, 134 Trochilidae (Hummingbirds), 59, 443 Troglodytes aedon (House Wren), 553 Troglodytidae (Wrens), 61, 444 Troglotrematidae, 231, 238 Trogonidae (Trogons and Quetzals), 59, 443 Trogoniformes, 14, 59, 163, 443 trombidiosis, 531 Tropical fowl mite (Ornithonyssue bursa), 534 Tropocyclops prasinus, 385 Trumpeter Swan (see Cygnus buccinator) Trypanosoma, 27 tsutsugamushi disease (scrub typhus), 534 Tubifex tubifex, 301 tubules, of Eustrongylides spp., 305 Tufted Duck (see Aythya fuligula) Tui Parakeet (see Brotogeris sanctithomae) Tundra Swan (see Cygnus columbianus) Turdidae (Thrushes), 61, 435 Turdus, 465 Turdus grayi (clay-colored Robin), 208 Turdus grayi (Clay-colored Robin), 333 Turdus iliacus (Redwing), 345, 484 Turdus merula (Eurasian Blackbird), 206, 345, 451, 484 Turdus migratorius (American Robin), 208, 238, 280, 333, 345, 484, 532–533, 550 Turdus philomelos (Song Thrush), 207, 345, 485 Turdus pilaris (Fieldfare), 485 Turdus ruficollis (Dark-throated Thrush), 485 Turdus viscivorus (Mistle Thrush), 207, 485, 401 turkey, domestic (see Meleagris gallopavo) Turkey Vulture (see Cathortes aura) Turnicidae (Buttanquail), 442 Turquoise Parrot (see Neophema pulchella) Turquoise Tanager (see Tangara mexicana) Turtur spp. Turtur abyssinicus (Black-billed Wood-Dove), 73 Turtur afer (Blue-spotted Wood-Dove), 73 Turtur chalcospilos (Emerald-spotted Wood-Dove), 73 Turtur tympanistria (Tambourine Dove), 73 Tylotropidus patagiatus, 378 Tympanuchus spp. Tympanuchus cupido attwateri (Attwater’s Prairie Chicken), 321, 322, 543 Tympanuchus cupido (Greater Prairie-Chicken), 155, 391, 396 Tympanuchus pallidicinctus (Lesser Prairie-Chicken), 392 Tympanuchus phasianellus (Sharp-tailed Grouse), 331, 393, 397, 406 Typhlocoelum, 229 Typhlocoelum cucumerium, 235 Tyrannidae (Tyrant Flycatchers), 435, 444 Tyto alba (Barn Owl), 114, 132, 208, 269, 477 Tyto novaehollandiae (Australian Masked-Owl), 477 Tytonidae (Barn Owls), 59, 443 Tyzzeria, 163
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594 UK moors, 317 Undulated Tinamou (see Crypturellus undulates) Upland Goose (see Chloephaga picta) Upupidae (Hoopoes), 60, 443 Ural Owl (see Strix uralensis) Uria, 465 Uria aalge (Common Murre), 291, 332 Urotocus, 230 Uvulifer, 229 Uvulifer ambloplitis, 226 vaccine for coccidiosis, 172 protection, against leucocytozoonosis, 94 protection, against parasitism, 7 protection, against Eimeria, 172 protection, against malaria, 48 Vanellus, 465 Vanellus vanellus (Northern Lapwing), 331, 361, 365, 476 Vangidae (Vangas), 61, 444 Variegated Tinamou (see Crypturellus variegates) Varied Thrush (see Ixoreus naevius) vector–parasite associations, 20 vector proofing, of aviaries, 544 vectors of avian filarioids, 451 distribution, 14–15 of Haemoproteus, 18 of Leucocytozoon, 85 of Plasmodium, 41 Venezillo evergladensis, 327 ventral suckers, 226 ventricular nematodiasis, 355 verminous encephalitis, 424 Verminous peritonitis, 289 Vermivora peregrina (Tennessee Warbler), 333 Vermivora ruficapilla (Nashville Warbler), 116 Vestiaria coccinea (Iiwi), 45, 111 Victoria Crowned-Pigeon (see Goura Victoria), 217 Viduidae (Indigobirds), 60 Vinaceous Parrot (see Amazona vinacea) Violaceous Euphonia (see Euphonia violacea) Vireo griseus (White-eyed Vireo), 485 Vireonidae (Vireos), 61, 435, 444 Virginia opossums (Didelphis virginianus), 115 visceral larva migrans (VLM), 424, 425 vitamin B12 supplements, 117 Viviparus georgianus, 240 Volatinia jacarina (Blue-black Grassquit), 114 Vulturine Guineafowl (see Acryllium vulturinum) Wahlberg’s Eagle (see Aquila wahlbergi) Wardianum, 229 waterfowl, 316, 339, 355 infected with cestodes, 270 infected with gastrointestinal trematodes, 228 infected with Philopthalmus gralli, 234 infected with species of Corynosoma and Polymorphus, 279 infected with various hymenolepidid species, 267–268 infection with Echinuria uncinata, 336 Wattled Starling (see Creatophora cinerea) weak ammonia, 509 weak gastric juice, 509 Wedge-tailed Pigeon (see Treron sphenurus) Western Bluebird (see Sialia mexicana)
Index Western Crowned-Pigeon (see Goura cristala) Western Grebe (see Aechmophorus occidentalis) Western Marsh-Harrier (see Circus aeruginosus) Western Meadowlark (see Sturnella neglecta) West Indian Whistling-Duck (see Dendrocygna arborea) Whimbrel (see Numenius phaeopus) Whip snake (Masticophis flagellum), 302 Whiskered Auklet (see Aethia pygmaea) Whiskered Tern (see Chilidonias hybrida) White-backed Vulture (see Gyps africanus) White-bellied Pigeon (see Treson sieboidii) White-Breasted Waterhen (see Amaurornis phoenicurus) White-browed Babbler (see Pomatostomus superciliosus) White-copped Redstart (see Chaimarrornis leucocephalus) White-crowned Pigeon (see Patagioenas leucocephala) White-eared Dove (see Phapitreron leucotis) White-eared Parakeet (see Pyrrhura leucotis) White-eyed Parakeet (see Aratinga leucophthalma) White-eyed Vireo (see Vireo griseus) White-faced Whistling-Duck (see Dendrocygna viduala) White-headed Duck (see Oxyura leucocephala) White Ibis (see Eudocimus albus) White-lined Tanager (see Tachyphonus rufus) White-naped Crane (see Grus vipio), 182, 184 white pulp arteriolar endothelium, hyperplasia, of, 21 White Stork (see Ciconia ciconia) White-tailed Eagle (see Haliaeetus albicilla) White-tailed Ptarmigan (see Lagopus leucura) White-throated Sparrow (see Zonotrichia albicollis) White-tipped Dove (see Leptotila verreauxi) White Wagtail (see Motacilla alba) White-winged Dove (see Zenaida asiatica) White-winged Duck (see Cairina scutulata) White-winged Scoter (see Melanitta fusca) Whooper Swan (see Cygnus cygnus) Whooping Crane (see Grus americana) Wild Turkey (see Meleagris gallopavo) Willet (see Tringa semipalmata inornata) Willow Ptarmigan (see Lagopus lagopus) Wilsonia citrina (Hooded Warbler), 485 Wohlfahrtia spp Wohlfahrtia magnifica, 550, 552 Wohlfahrtia opaca, 550 Wohlfahrtia spp., 553 Wohlfahrtia vigil, 550 Wonga Pigeon (see Leucosarcia melanoleuca) Wood Duck (see Aix sponsa) Wood Sandpiper (see Tringa glareola) Wood Stork (see Mycteria americana) Wood Thrush (see Hylocichla mustelina) Wormersia midwayensis, 527 Xenicidae, 444 Xenopus laevis (African clawed frog), 303 Xenus cinereus (Terek Sandpiper), 476 Yellow-bellied Sapsucker (see Sphyrapicus varius) Yellow-billed Chough (see Pyrrhocorax graculus) Yellow-billed Duck (see Anas undulata) Yellow-billed Pintail (see Anas georgica) Yellow Bittern (see Ixobrychus sinensis) Yellow Bunting (see Embesiza sulphurata) Yellow-collared Lovebird (see Agap*rnis personatus) Yellow-crested co*ckatoo (see Cacatua sulphurea) Yellow-crowned Night-Heron (see Nyctanassa violacea) Yellow-crowned Parrot (see Amazona ochrocephala) Yellow-footed Pigeon (see Treron phoenicopterus)
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Index Yellow-fronted Parakeet (see Cyanoramphus auriceps) Yellow-fronted woodpecker (see Melanerpes flavifrons) Yellowhammer (see Emberiza citronella) yellowish caseous lesions, 137 Yellow-legged Tinamou (see Crypturellus noctivagus) Yellow perch (Perca flavescens), 298 Yellow-throated Warbler (see Dendroica dominica) Zebra Dove (see Geopelia striata) Zebra Finch (see Taeniopygia guttata) Zeinaida spp. (Doves), 425, 465 Zenaida asiatica (White-winged Dove), 71, 128, 398, 404
595
Zenaida auriculata (Eared Dove), 394 Zenaida galapagoensis (Galapagos Dove), 128 Zenaida macroura (Mourning Dove), 17, 22, 23, 71, 126, 136, 144, 332, 398 zinc sulfate solution, 490 zoalene, 172, 177 Zonotrichia albicollis (White-throated Sparrow), 20, 22, 23, 234 Zosteropidae (White-eyes), 60, 444 Zosterops japonicus (Japanese White-eye), 111 Zygocotyle, 231 Zygocotyle lunata, 236
