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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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(Haemosporidia: Leucocytozoidae). Journal of Protozoology 17:642–658. Khan, R. A., and A. M. Fallis 1971. A note on the sporogony of Parahaemoproteus velans (=Haemoproteus velans Coatney and Roudabush) (Haemosporidia: Haemoproteidae) in species of Culicoides. Canadian Journal of Zoology 49:420–421. Kirkpatrick, C. E., S. K. Robinson, and U. D. Kitron. 1991. Phenotypic correlates of blood parasitism in the common grackle. In Bird–Parasite Interactions: Ecology, Evolution, and Behavior, J. E. Loye and M. Zuk (eds). Oxford University Press, New York, pp. 344–358. Klei, T. R., and D. L. DeGiusti. 1975. Seasonal occurrence of Haemoproteus columbae Kruse and its vector Pseudolynchia canariensis Bequaert. Journal of Wildlife Diseases 11:130–135. Korpim¨aki E., P. Tolonen, and G. F. Bennett. 1995. Blood parasites, sexual selection and reproductive success of European Kestrels. Ecoscience 2:335–343. Kˇucera, J., K. Marj´ankov´a, V. Racha˘c, and J. V´ıtovec. 1982. Haemosporidiosis as a fatal disease in muscovy ducks (Cairina moschata) in South Bohemia. Folia Parasitologia (Prague) 29:193–200. Laird, M. 1960. Migratory birds and the dispersal of avian malaria parasites in the South Pacific. Canadian Journal of Zoology 38:153–155. Lederer, R., R. D. Adlard, and P. J. O’Donoghue. 2002. Severe pathology associated with protozoal schizonts in two pied currawongs (Strepera graculina) from Queensland. Veterinary Record 150:520–522. Levine, N. D., P. D. Beamer, and J. Simon. 1970. A disease of chickens associated with Arthrocystis galli n.g., n.sp., an organism of uncertain taxonomic position. In H. D. Srivastava Commemoration Volume, pp. 429–434. MacCallum, W. G. 1898. On the haematozoan infections of birds. Journal of Experimental Medicine 3:117– 136. Martinsen, E. S., S. L. Perkins, and J. J. Schall. 2008. A three-genome phylogeny of malaria parasites (Plasmodium and closely related genera): Evolution of life-history traits and host switches. Molecular Phylogenetics and Evolution 47:261–273. McCurdy, D. G., D. Shutler, A. Mullie, and M. R. Forbes. 1998. Sex-biased parasitism of avian hosts: Relations to blood parasite taxon and mating system. Oikos 82:303–312. Mendes, L., T. Piersma, M. Lecoq, B. Spaans, and R. E. Ricklefs. 2005. Disease-limited distributions? Contrasts in the prevalence of avian malaria in shorebird species using marine and freshwater habitats. Oikos 109:396–404.

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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Ricklefs, R. E., and S. M. Fallon. 2002. Diversification and host switching in avian malaria parasites. Proceedings of the Royal Society of London, Series B 269:885–892. Sanz, J. J., E. Arriero, J. Moreno, and S. Merino. 2001. Female hematozoan infection reduces hatching success but not fledging success in Pied Flycatchers Ficedula hypoleuca. The Auk 118:750–755. Schrader, M. S., E. L. Walters, F. C. James, and E. C. Greiner. 2003. Seasonal prevalence of a haematozoan parasite of Red-Bellied Woodpeckers (Melanerpes carolinus) and its association with host condition and overwinter survival. The Auk 120:130–137. Sergent, E., and M. B´equet. 1914. De l’immunit´e dan le paludisme des oiseaux. Les pigeons gu´eris de l’infection a Haemoproteus columbae ne sont pas immunizes contre elle. Comptes Rendu des Seances de la Societe de Biologie et de ses Filiales 77:21– 23. Sergent, Ed., and Et. Sergent. 1906. Sur le second hˆote de l’Haemoproteus (Halteridium) du pigeon (Note pr´eliminaire). Comptes Rendus des Seances de la Societe de Biologi´e 61:494–496. Sibley, L. D., and J. K. Werner. 1984. Susceptibility of pekin and muscovy ducks to Haemoproteus nettionis. Journal of Wildlife Diseases 20:108–113. Siikam¨aki, P., O. R¨atti, M. Hovi, and G. F. Bennett. 1997. Association between haematozoan infections and reproduction in the Pied Flycatcher. Functional Ecology 11:176–183. Simpson, V. R. 1991. Leucocytozoon-like infection in parakeets, budgerigars and a common buzzard. Veterinary Record 129:30–32. Smith, G. A. 1972. Aberrant Leucocytozoon infection in parakeets. Veterinary Record 91:106. Smith, R. B., E. C. Greiner, and B. O. Wolf. 2004. Migratory movements of Sharp-Shinned Hawks (Accipiter striatus) captured in New Mexico in relation to prevalence, intensity, and biogeography of avian hematozoa. The Auk 121:837–846. Sol, D., R. Jovani, and J. Torres. 2000. Geographical variation in blood parasites in feral pigeons: The role of vectors. Ecography 23:307–314. Sol, D., R. Jovani, and J. Torres. 2003. Parasite mediated mortality and host immune response explain age-related differences in blood parasitism in birds. Oecologia 135:542–547. Tarshis, I. B. 1955. Transmission of Haemoproteus lophortyx O’Roke of the California valley quail by hippoboscid flies of the species Stilbometopa impressa (Bigot) and Lynchia hirsuta Ferris. Experimental Parasitology 4:464–492. Tella, J. L., G. Blanco, M. G. Forero, A. Gaj´on, J. A. Don´azar, and F. Hiraldo. 1999. Habitat, world geographic range, and embryonic development of

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hosts explain the prevalence of avian hematozoa at small spatial and phylogenetic scales. Proceedings of the National Academy of Sciences of the United States of America 96:1785–1789. Valki¯unas, G. 1993. The role of seasonal migrations in the distribution of Haemosporidia of birds in North Palaearctic. Ekologija 2:57–66. Valki¯unas, G. 1997. Bird Haemosporida. Vilnius: Institute of Ecology. Valki¯unas, G. 2005. Avian Malaria Parasites and Other Haemosporidia. CRC Press, New York. Valki¯unas, G., and T. A. Iezhova. 2001. A comparison of the blood parasites in three subspecies of the Yellow Wagtail Motacilla flava. Journal of Parasitology 87: 930–934. Valki¯unas, G., and T. A. Iezhova. 2004. The transmission of Haemoproteus belopolskyi (Haemosporida: Haemoproteidae) of Blackcap by Culicoides impunctatus (Diptera: Ceratopogonidae). Journal of Parasitology 90:196–198. Valki¯unas, G., G. Liutkeviˇcius, and T. A. Iezhova. 2002. Complete development of three species of Haemoproteus (Haemsporida, Haemoproteidae) in the biting midge Culicoides impunctatus (Diptera, Ceratopogonidae). Journal of Parasitology 88:864–868. Valki¯unas, G., F. Bairlein, T. A. Iezhova, and O. V. Dolnik. 2004. Factors affecting the relapse of Haemoproteus belopolskyi infections and the parasitaemia of Trypanosma spp. in a naturally infected European songbird, the blackcap, Sylvia atricapilla. Parasitology Research 93:218–222. Valki¯unas, G., A. M. Anwar, C. T. Atkinson, E. C. Greiner, I. Paperna, and M. A. Peirce. 2005. What distinguishes malaria parasites from other pigmented haemosporidians? Trends in Parasitology 21:357–358. Valki¯unas, G., A. Krizanauskiene, T. A. Iezhova, O. Hellgren and S. Bensch. 2007. Molecular phylogenetic analysis of circumnuclear

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)

September 29, 2008

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

September 29, 2008

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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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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75

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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77

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 — —

September 29, 2008

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).

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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

September 11, 2008

— 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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132

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

September 11, 2008

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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133

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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Trichom*onosis

(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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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.

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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.

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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

September 11, 2008

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

September 11, 2008

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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Section III: Helminths

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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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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

+

+

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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

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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

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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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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

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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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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 f