Amphibian biology. Volume 8. Amphibian decline: Diseases, parasites, maladies and pollution

Amphibian Biology Edited by

Harold Heatwole

Volume 8

Amphibian Decline: Diseases, Parasites, Maladies and Pollution Co-editor for this volume

John W. Wilkinson

Published by

Surrey Beatty & Sons

This book is copyright. Apart from any fair dealing for purposes of private study, research, criticism or review, as permitted under the Copyright Act, no part may be reproduced by any process without written permission. Enquiries should be made to the publisher. Surrey Beatty & Sons Pty Ltd is a member of Copyright Agency Limited (CAL) in Australia and is a respondent to overseas collecting societies.

The National Library of Australia Cataloguing-in-Publication Entry: Amphibian Biology. Volume 8, Amphibian Decline: Diseases, Parasites, Maladies and Pollution Includes Index. ISBN 0 949324 53 1 (set) ISBN 0 949324 54 X (v. 1) ISBN 0 949324 60 4 (v. 2) ISBN 0 949324 72 8 (v. 3) ISBN 0 949324 87 6 (v. 4) ISBN 0 949324 94 9 (v. 5) ISBN 0 949324 95 7 (v. 6) ISBN 978 0 98031 13 10 (v. 7) ISBN 978 0 98031 13 34 (v. 8) 1. Amphibians - Classification. I. Heatwole, Harold.

Published 2009

PRODUCED IN AUSTRALIA BY

SURREY BEATTY & SONS PTY LIMITED PO Box 8159, Baulkham Hills NSW 2 153 Australia

Preface to Series

T

ERE are several outstanding treatises of amphibian biology. Biology of Amphibians (Duellman and Trueb 1986) is an excellent general work, clearly written and well illustrated, and with a remarkable depth and breadth for a single volume. It will be the standard general reference on amphibians for many years to come and the present generation of herpetologists will consider it the "amphibian bible" much as their predecessors regarded G. K. Noble's (1931) The Biology of the Amphibia. No single volume, however, fulfils the need for a sequential, monographic treatment of specialized topics.

A few individual subjects have been treated in considerable depth. These include the three volumes of Physiology of the Amphibia (Moore 1964; Lofts 1974, 1976), Frog Neurobiology, a Handbook (Llinas and Precht 1976) and The Rep-oductive Bwlogy of Amphibzam (Taylor and Guttman 1977), but it has now been many years since these appeared and many topics are in need of updating. Environmental Physiology of the Amnphihm (Feder and Burggren 1992) did that for some aspects and treated other new ones, but it too is now getting out of date. Two books, Pattems of Dish'bution of Amphibians ( D u e b a n 1999) and Tadpoles (McDiarmid and Altig 1999) closed out the 20h century with excellent reviews of amphibian biogeography and of the biology of larval amphibians, respectively. Starting the millennium was a multi-volume treatment of reproductive biology and phylogeny: R w u c t i v e Biology and Phylogeny of Urodela (edited by David M. Sever) and Reproductive Biology and Phylogeny of Anura (edited by Barrie Jamieson), both in 2003, followed by Reproductive Biology and Phylogeny of Gymnophiona (edited by Jean-Marie Exbrayat) in 2004. Reproduction was further treated by Ogielska (2009) in Reproduction of Amphibians. Recent molecular techniques led to a rearrangement of amphibian taxonomy (Frost et al. 2006). Behaviour and ecology of amphibians was given a recent, comprehensive, thoughtfd update (Wells 2007). Collectively, all the above works, excellent though they are, still leave large, unaddressed gaps in amphibian biology. Recognizing that the discipline of amphibian biology had reached sufficient maturity to warrant detailed, multi-volume review and that such a need had been filled only partly, and with no commitment to continuation, the present series, Amphibian Biology, was launched in 1994. The present volume represents the eighth in the series. Amphibian Biology does not compete with the titles mentioned above. Topics recently reviewed elsewhere are not covered in current volumes, but are reserved for such time as hrther update is required. The need for this series was evidenced by the enthusiastic response from potential authors. Of the 64 people contacted with invitations to contribute to the first few volumes, only three declined, and then because of heavy commitments otherwise. Most expressed the view that such a series was long overdue. With this initial encouragement the series was launched and it is continuing to enjoy undiminished support.

Amphibian Biology was inspired by Biology of the Reptilia (1969-2008), edited by Carl Gans, and is intended as a companion to that series. Biology of the Reptilia is a unique, monumental contribution to herpetology and has become the most authoritative single source of information on reptiles that is available. Comprehensive treatments of all aspects of reptilian biology are presented in detail and are exhaustively documented by literature. It has been, and continues to be, invaluable. It is hoped that Amphibian Biolopy will serve herpetologists in the same way and that it will maintain the high standard set by its reptilian counterpart. Harold Heatwole Series Editor

Raleigh, North Carolina, USA January 1993, revised August 2009

REFERENCES Duelhnan, W. E., 1999. "Patterns of Distribution of Amphibians". The John Hopkins University Press, Baltimore. Duellman, W. E. and Tmeb. L., 1986. "Biology of Amphibians". McGraw-Hill Book Company, New York. Eibrayat, J.-M. (ed.), 2004) "Reproductive Biology and Phylogeny of Gymnophiona". Science Publishers, Enfield. M e z M. E. and Burggren, W., (ed.s) 1992. "Environmental Physiology of the Amphibians". University of Chicago Press, Chicago.

VI

AMPHIBIAN BIOLOGY

Frost, D. R., Grant, T, Faivovich, J., Bain, R., Haas, A., Haddad, C. F. B., de S., R., Channing, A., Wilkinson, M., Donnellan, S. C., Raxworthy, C., Campbell, J. A. , Blotto, B. L., Moler, I?, Drewes, R. C., Nussbaum, R. A., Lynch, J. D., Green, D. M. and Wheeler, W., 2006. The amphibian tree of life. Bulletin of the American Museum of Natural History 297: 1-370 Gans, C. (ed.), 1969-1998. "Biology of the Reptilia". 19 vols, Academic Press, New York; Alan R. Liss, Inc., New York; The University of Chicago Press, Chicago; Society for the Study of Amphibians and Reptiles. Jamieskon, B. G. M. (ed.), (ed.), 2003 "Reproductive Biology and Phylogeny of Anura". Science Publishers, Enfield. Llins, R. and Precht, W., 1976. "Frog Neurobiology, a Handbook. Springer-lkrlag, Berlin. Lofts, B. (ed.), 1974 "Physiology of the Amphibia", volume 2. Academic Press, New York. Lofts, B. (ed.), 1976. "Physiology of the Amphibia", volume 3. Academic Press, New York. McDiarmid, R. W. and Altig, R., 1999. "Tadpoles". The University of Chicago Press, Chicago Moore, J. A. (ed.), 1964 "Physiology of the Amphibia". Academic Press, New York. Noble, G. K., 1931. "The Biology of the Amphibia". McGraw-Hill Book Company, New York. Ogielska, M. (ed.), 2009 "Reproduction of Amphibians". Science Publishers, Enfield. Sever, D. M. (ed.), 2003 "Reproductive Biology and Phylogeny of Urodela". Science Publishers, Enfield. Taylor, D. H. and Gutunan, S. I. (eds.), 1977. "The Reproductive Biology of Amphibians". Plenum Press, New York. Wells, K. D. 2007. "The Ecology and Behavior of Amphibians". The University of Chicago Press, Chicago.

Preface to Volume 8

T

E late 20th century and the early 21" century has been characterized by an unprecedented deterioration of the environment of the earth and the throes of one of the major extinction events of all time. New diseases have emerged. Unmitigated deforestation, desertization, erosion and salinization of soil, pollution of water and air, and thinning of the UV-protective ozone layer constitute dire ecological threats for life on the planet. Fossil carbon is being returned to the atmosphere at an accelerated rate with a concomitant change in the earth's climate that is likely to make serious inroads into ecological stability.

The human population now exceeds the long-term carrying capacity of the earth and is able to subsist at its present level only because it is sustained by fossil resources of energy, soil, water and even oxygen. With continuing decrease in biodiversity, progressive destruction of essential habitats, degradation of major ecosystems, and contamination of life-support systems, it is likely that the carrying capacity of the earth will decline below present levels while at the same time the human population continues to rise. The outstripping of even its fossil resources, likely to occur within the present century, presents a bleak outlook for our own species. We may well become a victim ourselves of this most recent mass extinction. While it is indisputable that many aspects of environmental degradation and loss of biodiversity is directly attributable to unwise human activities, other aspects are deemed to result from natural cycles beyond the influence of mankind. It is important to be able to distinguish between the two, so that attention can be focused on mitigating those effects over which humans do have control. It is important to ascertain the causes of particular declines and extinctions as soon as possible, so that steps can be taken to preseIve as much diversity as possible. Amphibians, by virtue of their thin, moist, permeable skins, are poorly protected from harsh environments and are especially susceptible to chemical changes, desiccation, and alteration of habitat. Accordingly, it is not surprising that they manifest proportionately high extinction rates and more severe declines than do most other organisms. They are especially important to study as they serve as an early-warning system portending changes that may soon impinge upon more resistant species, including Homo sapzens. The topic of conservation and decline of amphibians will be the subject of several volumes. The present volume (8) is devoted to diseases, maladies, and parasites of amphibians and how these relate to decline, followed by the impact of pollution, of various kinds and from various sources, on amphibian populations. Finally, a chapter is devoted to the phenomenon of climatic change. Subsequent volumes will treat (1) the roles of anthropogenic influences such as habitat change; introduction of alien species; roadkills; direct harvesting, trade, and use of amphibian species by humans, (2) various ecological, phylogenetic, and geographic correlates of amphibian decline, (3) monitoring programmes and conservation practices such as establishment of rehgia; captive breeding and re-introduction; and mitigation; as well as the application of education. The last volumes will assess the global status of conservation and decline on a region-by-region basis to serve as a benchmark for subsequent changes that take place. There is modest overlap in subject matter among chapters. This is deliberate as particular aspects interact and it is more convenient to have complete coverage of a given topic without having to shift back and forth between chapters. Each chapter was designed to stand alone. Many of the chapters of these volumes were in mid-preparation when the monograph by Frost et al. (2006) appeared. The new taxonomic arrangements and nomenclatural changes espoused by Frost et al. are still controversial and there has been no editorial policy demanding authors either to adhere to it, or not to do so. Consequently, some chapters have the older scheme and others the newer one, as the authors' own assessments dictated. Harold Heatwole Raleigh, North Carolina, USA 17 January 2009 Series Editor

VIII

AMPHIBIAN BIOLOGY

Lee Bergel

Dedication

I?

RTUNATELY for amphibians Lee Berger decided to take a year off from her veterinary studies in 1991 to travel around Australia to see what research on wildlife diseases was being conducted. On that trip she met the eclectic scientist, veterinarian and doctor Rick Speare in Townsville, Queensland, and worked for him on an anti-parasitic drug that was causing mortality in wallabies. Little did she know how that would change our lives. Several years later, in 1995, while she was working as a veterinarian in Melbourne, Victoria, I remember her excitedly putting down the telephone to tell me that Rick had offered her a PhD position to determine the spreading agent that he thought was causing frog decline and extinction in Queensland. She moved to the Australian Animal Health Laboratory in Geelong, Victoria, to study samples collected by Keith McDonald during a mass mortality of frogs at Big Tableland, Queensland. This is the premier virology laboratory in Australia and she was given the task of finding a virus in those samples. During six months of negative findings for viruses she discovered a novel organism that we now know as Batrachochytm'um dendrobatidis infecting the skin of these frogs as well as others submitted to the laboratory as dead or sick. The key breakthrough came in 1996 when this pathogen appeared in winter dieoffs and Lee decided to take this organism seriously and started to investigate the disease it caused, chytridiomycosis, and its role in amphibian declines and extinction. Since the early 1980s scientists were unable to explain declines and extinctions of amphibians in protected habitats until Lee's discovery. Her work, and that of her colleagues, was published in the Proceedings of the National Academy of Sciences (Berger et al. 1998) and they were awarded the Australian Commonwealth Scientific and Industrial Research Organisation medal for science. It wasn't all smooth sailing, however. There were numerous difficulties working with a novel pathogen and in the field of amphibian disease where there was little expertise. She spent many late nights at work overcoming these obstacles. There was also the pressure of solving the cause of amphibian declines as well as intense scrutiny of her work and opposition to the hypothesis that a spreading disease was the cause of amphibian decline. Since then, she has continued to make major contributions in understanding the epidemiology and pathogenesis of chytridiomycosis and has investigated other emerging diseases of amphibians. She has also devoted much of her time in assisting others in Australia and overseas in conducting research on amphibian diseases. Significant management recommendations for managing the disease have been adopted based on her work. These include (1) the Amphibian Ark Progx-am that operates an emergency captive-breeding programme to prevent extinctions of populations and species due to outbreaks of chytridiomycosis, (2) the World Animal Health Organisation's (OIE) listing of chytridiomycosis as a notifiable disease in order to prevent its spread, and (3) the Australian Government listing it as a key threatening process in order to prevent its spread and to abate its current impact on threatened species of frogs. The spectacular informative nature of Dr. Berger's work demonstrates the risk of introducing virulent pathogens into na'ive host populations and ultimately will lead to better conservation of amphibians in particular and in the long term to better conservation of wildlife in general. Currently, Dr. Berger holds a Postdoctoral Research Fellowship at James Cook University, balancing work on amphibian diseases with the care of three young children. It has been my pleasure to know and work with Lee, who is not only an outstanding scientist but a lovely human being. I am extremely pleased that this volume on amphibian conservation is dedicated to her in recognition of how much she has contributed already to this field during her career.

co tor Creek, Queensland, Australia

Lee F. Skerratt

Contents . . . . . . . . . . . . . . . . . . . . . . . . . .

v

Preface to Volume 8 . . . . . . . . . . . . . . . . . . . . . . . . . .

vii

. . . . . . . . . . . . . . . . . . . . . . . . . . . .

ix

Contributors to volume 8 . . . . . . . . . . . . . . . . . . . . . . . .

xii

Preface to Series

Dedication

Chapter 1. Viral and Bacterial Diseases of Amphibians. Valentine Hemingway, Jesse . . . . . . . . . . . . . . . . Brunner, Rick Speare, and Lee Berger Chapter 2. Fungal Diseases of Amphibians. Lee Berger, Joyce E. Longcore, Rick Speare, Alex Hyatt and Lee F. Skerratt . . . . . . . . . . . . . . . . Chapter 3. Factors Affecting Interspecific Variation in Susceptibility to Disease in Amphibians. Jodi J. L. Rowley and Ross A. Alford . . . . . . . . . . . . Chapter 4. Digenetic Trematodes and their Relationship to Amphibian Declines and Deformities. Jason Rohr, Thomas Raffel, and Stanley K. Sessions . . . . . . Chapter 5. Amphibian Malformations. Michael J. Lannoo

. . . . . . . . . .

Chapter 6. Ultraviolet-B Radiation and Amphibians. Adolfo Marco, Betsy A. Bancroft, Miguel Lizana and Andrew R. Blaustein . . . . . . . . . . . . . . . . Chapter 7. Pollution: Impact of Reactive Nitrogen on Amphibians (Nitrogen Pollution). Adolfo Marco and Manuel Ortiz-Santaliestra . . . . . . . . . . . . . . Chapter 8, Evaluating the Impact of Pesticides in Amphibian Declines. Michelle D. Boone, . . . . . . . . . . . . Carlos Davidson and Christine Bridges-Britton Chapter 9. Endocrine Disrupting Chemicals. Krista A. McCoy and Louis J. Guillette Jr Chapter 10. Role of Petrochemicals and Heavy Metals in Amphibian Declines. Luca M. Luiselli and Jerry Lea . . . . . . . . . . . . . . . . . . . . . . Chapter 11. Acidification and its Effects on Amphibian Populations. Katja E s a n e n and David M. Green . . . . . . . . . . . . . . . . . . . . . . Chapter 12. Climatic Change and Amphibian Declines. Patricia Burrowes Subject Index

. . . .

. . . . . . . . . . . . . . . . . . . . . . . . . . . .

Index to Scientific Names

. . . . . . . . . . . . . . . . . . . . . .

Contributors ALFORD, Ross A., School of Marine and Tropical Biology, James Cook University, Townsville, Qld. 48 11, Australia. ross [email protected] BANCROFT, Betsy A., Department of Zoology, Oregon State University, 3029 Cordley Hall, Corvallis Oregon 9733 1-2914, USA. [email protected] BERGER, Lee, Amphibian Disease Ecology Group, School of Public Health, Tropical Medicine and Rehabilitation Science, James Cook University, Townsville, Queensland 481 1, Australia. 1ee.bergerajcu.edu.a~ BLAUSTEIN, Andrew R., Department of Zoology, Oregon State University, 3029 Cordley Hall, Corvallis Oregon 97331-2914, USA. [email protected] BOONE, Michelle D., Department of Zoology, 212 Pearson Hall, Miami University, Oxford, Ohio 45056, USA. [email protected] BRIDGES-BRITTON, Christine, United States Geological Survey, Columbia Envronmental Research Centger, 4200 New Haven Road, Columbia, MO 65201, USA. [email protected] BRUNNER, Jessee, Department of Environmental and Forest Biology, SUNY-ESF, 248 Illick Hall, 1 Forestry Drive, Syracuse, New York 13210, USA. [email protected] http:// www.esf.edu/efb/bmnner/ BURROWS, Patricia, Department of Biology, University of Puerto Rico, PO. Box 23360, San Juan, Puerto Rico, 00936, USA. [email protected] GREEN, David M., Redpath Museum, 859 Sherbrooke St., McGill University. W. Montreal, Quebec H3A 2K6, Canada. [email protected] GUILLEITE, Louis J. Jr., Department of Zoology, 223 Bartram Hall, Box 118525, University of Florida, Gainesille, Florida 3261 1-8525, USA. [email protected] HEATWOLE, H. (Editor), Department of Biology, North Carolina State University, Raleigh, NC 27695-76 17, USA. [email protected] HEMINGWAY, Valentine, Department of Ecology and Evolutionary Biology, A316 EMS CA 95064, USA. Building, University of California, Santa Cmz, [email protected] HYATT, Alex, Australian Animal Health Laboratory, CSIRO Livestock Industries, Private Bag 24, Geelong, Victoria 3220, Australia. [email protected] LANNOO, Michael J., Holmstedt Hall room 135, Terre Haute Center for Medical Education, Indiana University School of Medicine, Terre Haute, IN 47809, USA. [email protected] LEA, Jerry, Department of Biological Sciences, Delta State University, P.M.B. 01, Abraka, Delta State, Nigeria. [email protected] LIZANA, Miguel, Dept. ~ i o l o ~Animal. ia Univ. Salamanca, Campus Miguel de Unamuno 37007, Salamanca, Spain. [email protected] LONGCORE, of Biological Sciences, University of Maine, Orono, Maine 04469-5722, USA. [email protected] LUISELLI, Luca M., F. I. Z. V (Ecology) via Olona 7, 1-00198 Rome, Italy. [email protected] MARCO, Adolfo, ~ o b a n aBiological Station, CSIC, Avda. Spain. [email protected]

aria

Luisa sln, Sevilla, 41013,

McCOY, Krista A., Department of Zoology, 223 Bartram Hall, Box 118525, University of Florida, Gainesville, Florida 3261 1-8525, USA. [email protected]

AMPHIBIAN BIOLOGY

XI11

ORTIZ-SANTALIESTGRA, Manuel, Dept ~ i o l o ~ Animal. ia Univ. Salamanca, Campus Miguel de Unamuno 37007, Salamanca, Spain. [email protected] RAFFEL, Thomas, Division of Integrative Biology, University of South Florida, Tampa, F1 33620, USA. [email protected] (813) 974-3250 F&S~EN Katja, Department of Aquatic Ecology, Institute of Integrative Biology, EAWAG, ~berlandstrasse144, CH-8600, Diibendorf, Switzerland. [email protected] ROHR, Jason R., Biology Department, University of South Florida, 4202 East Fowler Ave., Tampa, FL 33620, USA. [email protected] ROWLEY, Jodi J. L., Herpetology Section, Australian Museum, 6 College Street, Sydney, NSW 20 10, Australia. [email protected] SESSIONS, Stanley K., Department of Biology, Hartwick College, Oneonta, NY 13820, USA. [email protected] SKERRAm, Lee F., Amphibian Disease Ecology Group, School of Veterinary and Biomedical Science, James Cook University, Townsville, Queensland 481 1, Australia. [email protected] SPEARE, Rick, Amphibian Disease Ecology Group, School of Public Health, Tropical Medicine and Rehabilitation Science, James Cook University, Townsville, Queensland, 48 11, Australia. [email protected] WILKINSON, John (Co-editor), Durrell Institute of Conservation and Ecology, Marlowe Building, University of Kent, Canterbury, Kent CT2 7NR, UK. [email protected]

CHAPTER 1

Viral and Bacterial Diseases of Amphibians Valentine Hemingway, Jesse Brunner, Rick Speare, and Lee Berger

I. Introduction II. Viral Diseases A. Ranaviruses 1. Taxonomy and Molecular Epidemiology 2. Biology 3. Clinical Signs and Pathology 4. Epidemiology 5. Resistance to Infection 6. Transmission and Spread 7. Diagnosis 8. Management 9. Discussion

B. Frog Erythrocytic Virus C. Lucke Tumour Herpesvirus D. Herpes-like Virus of Skin E. Calicivirus F. Leucocyte Viruses Ill. Bacterial Diseases A. Bacterial Septicaemia ("Red Leg") B. Streptococcosis C. Chlamydiosis D. Mycobacteriosis IV. References

I. INTRODUCTION

A

T least six groups of viruses have been reported to affect amphibians, including caliciviruses, herpesviruses, and iridoviruses (Johnson and Wellehan 2005). Only ranaviruses are known to cause widespread mass mortality and have been studied in detail; hence a review of this group of viruses forms the majority of this chapter. Various strains of ranavirus are found worldwide and some appear to have spread recently (Hyatt et al. 2000). Indicative of the broad host range of ranaviruses and their potentially devastating effects, ranaviral disease was listed by the World Organization for Animal Health (OIE) as an internationally notifiable disease in 2008. Although their impacts on populations of declining species are a concern, there is currently no evidence that they have caused permanent declines or extinctions (Daszak et al. 2003). Nevertheless, because of their potential impacts on naive populations, as well as on species that are facing multiple threats, it is important that the risk of spreading these pathogens is minimized (Cunningham et al. 2007a).

There have been few investigations into other amphibian viruses. Apart from Frog Erythrocytic Virus (FEV), their impact on wild populations has not been studied. Luck6 nunor herpesvirus has been well described, but the other viruses found associated with disease in amphibians have been reported in single papers with little or no experimental work. Their significance as pathogens of amphibians is therefore largely unknown. In addition other viruses, not reviewed here, such as arboviruses (including West Nile Virus), mmvimses and an adenovirus, can infect frogs but their pathogenicity to amphibians is km- or unknown (Densmore and Green 2007).

AMPHIBIAN BIOLOGY

There are no substantiated reports of bacteria causing outbreaks in wild frogs, and cases

of bacterial disease were rare during large surveys for disease in the United States and Ausualia (Berger 2001; Green et al. 2002). Bacterial diseases, including septicaemia, are associated with significant mortality in captivity and are associated with poor husbandry. For details on prevention, management and treatment of bacterial diseases in captivity see Wright and Whitaker (2001). Zoonotic bacteria carried by wild and captive amphibians with minimal effect on themselves, but which are potential risk to humans (e.g., Salmonella and Leptospira), are not included in the present review.

.

11. VIRAL DISEASES

A. Ranaviruses A recent spate of research on ranaviruses was stimulated by the interest in global amphibian declines (Chinchar 2002). It built on earlier work that initially arose out of the discovery of ranaviruses in Luck6 tumor research (Granoff et al. 1969), then fish mortalities (Langdon 1989), and subsequently their possible use as a biological control agent for the cane toad (Bufo marinus) in Australia (Speare 1990; Pallister et al. 2007). Ranaviruses are significant infectious agents that can cause great mortality in free-living amphibian populations (Cunningham et al. 1996) although they are not currently believed to be responsible for amphibian declines (Daszak et al. 2003). Amphibian ranaviruses are enveloped icosahedral DNA viruses in the family Iridoviridae (Hengstberger et al. 1993). Isolates causing disease have been found in wild and cultured amphibians in Australia (Speare and Smith 1992), the Americas (Wolf et al. 1968; Zupanovic et al. 1998b; Majji et al. 2006), Asia (He et al. 2001; Zhang et al. 2001; Weng et al. 2002), and Europe (Cunningham et al. 1996). These include Tadpole edema virus, Frog virus 3, Rana catesbeiana virus Z , Ambystoma tigrinum virus, Bohle iridovirus, and UK ranavirus. In North America, ranaviruses are responsible for massive mortality in amphibian larvae and recent metamorphs, while die-offs rarely occur in adults (Green et al. 2002). These ranavirus-induced mortality events often occur during summer and involve hundreds to thousands of moribund and dead larvae within a few days (Green et al. 2002). Additionally, adult amphibians can be chronically infected carriers, maintaining infection in a population (Wolf et al. 1969; Brunner et al. 2004; Robert et al. 2007). In contrast, ranaviral disease in the United Rngdom typically causes massive, synchronous annual mortality of adult frogs (Drury et al. 1995). Molecular work suggests there has been recent spread of ranaviruses both locally and globally. For instance, the strains that occur in the United Kingdom and in captive-breeding facilities worldwide may have originated from North America (Hyatt et al. 2000) 1 . Taxonomy and Molecular Epidemiology

Currently there are six commonly recognized species of ranavirus (ICTVdB Management 2006), three of which were isolated from amphibians and the others from fish. Species of ranavirus are differentiated on the basis of genetic sequences, particularly the sequencing regions of the major capsid protein (Mao et al. 1997, Hyatt et al. 2000), but also by restriction fragment length polymorphism (RFLP) profiles and by DNA hybridization (Hyatt et al. 2000), virus protein profiles, and the range and specificity of the host (Hyatt et al. 2000, ICTVdB Management 2006). However, while isolates of the same species are largely homologous, or even identical in MCP sequence (2 95% sequence identity), they can be quite distinct in RFLP profiles (Hyatt et al. 2000; Maji et al. 2006; Schock et al. ZOOS), host range, and virulence (Brunner, unpublished data), as well as in tissue tropisms (Cunningham et al. 2007).

HEMINGWAY ET AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBLANS

2965

Frog Virus 3 (FV3), the type ranavirus, was isolated from aclinically infected leopard frogs (Rana pipiens) collected in the United States in 1962 (Granoff et al. 1965). Since then, FV3 or FV3-like viruses, such as Tadpole edema virus (TEV), Rana catesbeiana virus Z (RCVZ), and UK Ranaviruses (RUK, BUK), have been associated with amphibian mortality in North America (Wolf et al. 1968, Wolf et al. 1969, Petranka et al. 2003, Greer et al. 2005, Bank et al. 2007, Majji et al. 2006, Schock et al. 2008), South America (Zupanovic et al. 1998a, 1998b), the United Kingdom (Drury et al. 1995), and Southeast Asia (Kanachanakhan 1998, Zhang et al. 2001). The second distinct amphibian ranavirus species to be discovered was Bohle Iridovirus (BIV), isolated from newly metamorphosed ornate burrowing frogs (Limnodynastes ornatus) in Queensland, Australia (Speare and Smith 1992; Hengstberger et al. 1993). It remains the only isolate of this species although subsequently ill wild Litoria caerulea had positive PCR for BW but the virus could not be isolated (Cullen and Owens 2002). Antibodies against ranaviruses were detected in cane toads throughout most of their range in Australia at an overall prevalence of 2.7% (range 0-18%).The identity of the ranavirus that induced the antibodies is unknown since the test is not species specific and no viruses were isolated from any cane toads (Zupanovic et al. 199813). Lastly, the Ambystoma tigrinum virus (ATV) was isolated from a threatened sub-species of tiger salamander (Ambystoma tigrinum stebbinsi) in southern Arizona, USA in 1995 (Jancovich et al. 1997). Similar ATV-like viruses have been isolated from many locations in western North America (Bollinger et al. 1999; Jancovich et al. 2005; Ridenhour and Storfer 2008), but not elsewhere. Ranaviruses can cause aclinical infections in resistant animals, which may facilitate the spread of disease via the inadvertent movement of infected animals (Robert et al. 200'7). The trade of amphibians for food, research, and as pets has likely played a role in the movement of pathogens such as ranaviruses both within and among continents (Cunningham and Langton 1997; Jancovich et al. 2005; Galli et al. 2006; Picco and Collins 2008). Phylogenetic analyses of ATV isolates based on sequence from the MCP and DNA methyltransferase genes, as well as two non-coding regions, suggest a single introduction and radiation of these salamander ranaviruses, with little genetic divergence among isolates (< 1.1%), but a rather complex phylogeography (Jancovich et al. 2005). Nested-clades analyses suggest long-distance dispersal. One clade encompassed isolates from southern Arizona to Saskatchewan, as well as isolates from the bait trade and one from the axolotl (Ambystoma mexicanum) colony at Indiana University (Jancovich et al. 2005). ATV has been frequently isolated from tiger salamander larvae used as fishing bait (Picco and Collins 2008) and so at least some of these dispersals are probably due to moving infected bait. An analysis of concordance between the phylogenies of ATV and the tiger salamander host showed an overall lack of congruence, but when three isolates of presumably anthropogenic origins were excluded, the trees of the host and virus were identical, indicating co-evolution between host and parasite (Storfer et al. 2007). Interestingly, the UK ranavirus from common frogs (Rana temporaria), RUK, is phylogenetically similar to the FV3-like viruses from North America which, in addition to the fact that animals with signs of ranavirus infection were not observed before the mid1980s, suggests a recent introduction of the virus into the UK (Hyatt et al. 2000; Cunningham et al. 2007a). Recently an FV3-like virus was identified during mortality events in natural populations of the endemic Atelognathus patagonicus in Argentina (Fox et al. 2006). It showed 100% identity with the original FV3 isolate across 500bp of the MCP gene, which is consistent with a recent introduction from North America.

2 Biology Ranaviruses are composed of linear, double-stranded DNA genome encoding - 100 porkns (Wiliams et al. 2000) encapsulated in an icosahedral particle -130 nm in diameter

AMPHIBIAN BIOLOGY

with an internal lipid membrane. The capsid may or may not be enveloped (Braunwald et al. 1979) but both forms are infective (Chinchar 2002). Ranaviruses cause disease involving multiple organs and tissues in fish, amphibians, and reptiles (Chinchar 2002). The virus attaches to an as yet unidentified, but apparently widely distributed, receptor on host cells. Enveloped virions enter via receptor-mediated endocytosis, losing their envelope and releasing their nucleoprotein core. If the virus is naked, it enters the cell via fusion between the internal lipid membrane and the plasma membrane (Chinchar 2002). Early events in virus replication occur in the cell's nucleus, including transcription of early viral mRNAs and replication of copies of one-unit to two-unit lengths of the viral genome. Late viral mRNAs seem to be transcribed in the cytoplasm (Chinchar 2002). Viral assembly occurs at distinct assembly sites in the cell and the newly formed virions either bud off from the cell or, more commonly, accumulate in the cell until the cell lyses (Goorha and Granoff 1978). The machinery and metabolism of infected cells are rapidly diverted into viral replication (Murti et al. 1985a,b; Willis et al. 1985; Goorha and Granoff 1999; Chinchar 2002). Ranavirus particles are environmentally resistant and can remain infectious for long periods in certain environments. Studies of a fish ranavirus (EHNV) showed that it remained infective after drying for over 100 days at 15°C (Langdon 1989). Heating to 60°C for 15 minutes or 40°C for 24 hours inactivated the virus. Brunner et al. (2007), however, found that ATV was rendered non-infectious when dried in pond substrate and viral titres declined dramatically in ATV-spiked pond water. Ultraviolet radiation from aquaculture UV watersterilizers killed BIV rapidly at high flow rates (Miocevic et al. 1993). FV3 replicates optimally in culture between 12°C and 32"C, although it can replicate to some extent both below and above this range (Goorha and Granoff 1974). BIV had a thermal limit of 33"C, above which it would not grow. It was capable of infecting fish and mammalian cell lines (if maintained at 90% within a year after an outbreak (Green et a2. 2002). Chytridiomycosis has been implicated as a cause of declines of Bufo boreas in Colorado, Rana muscosa in California and B. baxteri in Wyoming (Green et al. 2002; Muths et al. 2003). Although B. boreas has declined since the 1970s, an isolated population in the Rocky Mountains had been stable until 1996 when a sudden decline occurred and chytridiornycosis was diagnosed in specimens collected in 1998 and 2000 (Muths et al. 2003). Only 3% survival was recorded between 1998 and 1999 and the population may not persist (Muths et al. 2003). There were no deaths in tadpoles (Muths et al. 2003). Male B. boreas disappeared faster than did females. Males visit breeding sites every season but females may only breed every second to fourth year. Consequently males have more opportunity for exposure to B. dendrobatidis in bodies of water (Muths et al. 2003). All seven native ranid frogs in Arizona have declined and chytridiomycosis may be important (Bradley et al. 2002). Bufo canorus found dying in California during declines in the 1970s, however, were diagnosed with a range of diseases and only 217 had chytridiomycosis (Green and Kagarise Sherman 2001). The toads were suspected to be immunosuppressed. Chytridiomycosis occurs in the northeastern United States, Quebec and British Columbia but declines have not occurred there (Carey et al. 2003). The epidemiology of chytridiomycosis in the United States has been complicated by the early establishment of B. dendrobatzdis in dwarf African clawed frogs (Hymenochim curtipes) which are widely sold in the ornamental pet trade. Groff et al. (1991) identified a pathogen of these frogs as Basidiobolus ranarum. The pathology was atypical for a filamentous fungus, however, and Carey et al. (2003) retrospectively identified the disease as chytridiornycosis. Its presence in such a widely distributed amphibian in the United States means that chytridiornycosis may have been unknowingly spread over an extensive area. Frogs distributed through scientific supply companies in the United States also have been found to be infected, e.g., Rana pipiens (E. Davidson, pers. comm.), R. catesbeiana (Daszak et al. 2004), Xenopus laeuis and X. tropicalis (Parker et al. 2002), thereby further complicating the epidemiology of chytridiornycosis. In Puerto Rico, populations of frogs in streams and bromeliads at high elevations have declined (Burrowes et al. 2004). Three species of Eleutherodactylus are presumed extinct, each last seen in 1976, 1981 and 1990. Chytridiomycosis has been detected in frogs collected in 1976 (in the last known specimen of E. karlschmidti) and in 1978. Declines occurred during droughts and it was suggested that clumping of the distribution of frogs, combined with stress, made them more vulnerable to chytridiomycosis (Burrowes et al. 2004).

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4. Europe

Environmental changes such as habitat destruction have obviously caused declines in amphibian populations in many countries (Alford and Richards 1999). Cases of inexplicable declines are unusual and no species have recently become extinct in Europe (Bosch et al. 2001). Outbreaks of chytridiomycosis in Alytes obsktricans in the alpine Pefialara Natural Park in central Spain, however, caused a severe population decline (Bosch et al. 2001). This area is at about 2 000 m elevation and contains bogs and alpine grasslands. Alytes obstetricans was one of the most abundant frogs. Mass mortality occurred in post-metamorphic animals in the summers of 1997, 1998 and 1999, and frogs disappeared from 86% of the ponds. Lower levels of Ca2+and Mg" and higher levels of H+ characterized ponds where tadpoles were observed compared with those where populations had disappeared. It is possible the disease has recently arrived and could spread to other populations. Introduced amphibians were found in the park, providing a potential transport mechanism for B. dendrobatzdis (Bosch et al. 2001). Numerous imported captive frogs in Germany were diagnosed with chytridiornycosis (Mutschmann et al. 2000). In 1999 an outbreak also was detected in wild frogs (Ram arvalis) near Berlin (F. Mutschmann, unpubl. data). In Italy chytridiornycosis was diagnosed in 2001 in a population of Bombina pachypus (Ferri 2002). An outbreak occurred in captive Dyscophus antongilii in Switzerland (Oevermann et al. 2005). 5. Asia

There have been no reports of chytridiomycosis in Asia. Approximately 100 specimens from Indonesia were negative (Mendez, unpubl. data) but comprehensive surveys from Asia have not been reported.

Epidemiological data support the hypothesis that B. dendrobatzdis originated in Africa (Weldon et al. 2004). The earliest case of chytridiomycosis worldwide is from 1938 froni Xenopus laevis in South Africa. The overall prevalence in 697 specimens of three species (X. laevis, X. gilli and X. muelleri) collected from 1890 to 1999 in South Africa was 2.7% and remained stable over time (Weldon et al. 2004). Diagnosis was made via histology. High prevalences (up to 100%) were found in Afiana fuscigola at some sites in the Western Cape and Northern Cape in South Africa, at sites ranging from 120 m to 1 194 m elevation (Hopkins and Channing 2003). Of five species tested, 36/85 (42%) of the frogs were positive. At Western Cape sites no mass die-offs were seen and frogs appeared healthy except for two of the infected frogs that were collected dead. Chytridiomycosis also was detected in frogs from Kenya (Afiana angolensis) and in wildcaught frogs (X. tropicalis) from Western Africa that had been imported into the United States (Reed et al. 2000; Speare and Berger 2005b). In Tanzania, the endemic Kihansi spray toad (Nectophrynoides asperginis) lived in the spray of the Kihansi River until a dam was built to produce hydropower. When the spray from the falls in the Kihansi Gorge ceased after the dam was installed, the number of spray toads decreased. A sprinkler system was installed to mimic the spray from the river and the toad population rebounded until 2003 when numbers dropped rapidly, and spray toads disappeared. Dead animals were collected and B. dendrobatidis was identified in the skin (Weldon and du Preez 2004). Xenopus laeuis shows minimal clinical effects and because this species has been exported globally since the 1930s for scie~itificresearch and pregnancy testing, it could have transported chytridiornycosis (Weldon et al. 2004). The presence of feral populZtions of X.

AMPHIBIAN BIOLOGY

2994

laevis in Chile, the United Kingdom, and the United States indicate there was an opportunity for the spread of chytridiomycosis (Weldon et al. 2004). To confirm Africa as the origin, genetic studies are needed that show greater genetic diversity in B. dendrobatzdis in populations of the fungus from its putative site of origin. Genetic markers are currently being sought for the fine-scale work on population genetics needed to trace .the intercontinental and intracontinental movement of this fungus.

In contrast to Xenopus laevis, X. tropicalis is susceptible to severe chytridiomycosis. Frogs in several groups of X. tropicalis died with chytridiomycosis shortly after being imported into the United States from Ghana and probably were infected while still in Africa (Reed et al. 2000; Carey et al. 2003). C. Taxonomy 1. The Chytridiomycota

Batrachochytrium dendrobatidis is a fungus in the Phylum Chytridiomycota (chytrids), Class Chytridiomycetes and Order Chytridiales (Berger et al. 1998; Longcore et al. 1999). The Chytridiomycota is one of five phyla of true fungi (Lutzoni et al. 2004) and contains about 1,000 species. The name is based on the flask-shaped appearance of sporangia; "chytr" means "earthen pot" in Greek. The phylum has one class, Chytridiomycetes, which is divided into five orders: Chytridiales, Blastocladiales, Monoblepharidales, Spizellomycetales and Neocallimastigales (Barr 1990, 2000). Chytridiomycetes are typified by the presence of chitin in the cell wall and the production of unwalled motile zoospores, each with a single posteriorly directed flagellum. Ultrastructure of the zoospore is more conserved among phylogenetic groups than is the morphology of the thallus (i.e., the entire organism) and is consequently useful for classification (Barr 1990). Molecular methods for classification are now common, with phylogenetic hypotheses based on analyses of gene sequences. Because chytrids are microscopic and not recovered using routine mycological culturing techniques they have received little attention despite being ubiquitous. Only a few people in the world specialize in studying them and it is probable that many species are yet to be discovered.

2. Batrachochytrium dendrobatidis (Amphibian Chytrzd) Batrachochytrium was erected as a new genus by Longcore et al. (1999); its morphology and the ultrastructure of its zoospore are unique and it is the only chytrid that is a pathogen of vertebrates. The important taxonomic features of its morphology include inoperculate discharge of zoospores, thread-like rhizoids and either monocentric or, occasionally, colonial growth (Longcore et al. 1999). Important ultrastructural features of the zoospore include a nucleus and kinetosome that are not associated with each other, aggregated ribosomes, a microbody that partially surrounds numerous small lipid globules, and a nonflagellated centriole that is parallel with the kinetosome and connected to it by overlapping fibres, as well as other details of the kinetosomal root (Figs. 2-4) (Longcore et al. 1999). Although most members of the Chytridiales have a rumposome (a fenestrated membrane cisterna) along the edge of the lipids and many have a transition-zone plug, these features are not present in B. dendrobatzdis (Berger et al. 1998; Longcore et al. 1999). The zoospore of B. &ndrobatzd& is also unusual for a member of the Chytridiales in having numerous lipid globules (Longcore et al. 1999).

Electron microscopy of zoospores from isolates collected in Australia and the United States did not reveal significant differences (Berger et al. 1998; Longcore et al. 1999). Genetic

studies also indicated that the population is homogeneous. Multilocus sequence-typing has been used to examine genetic diversity among fungal strains from North America (25 strains), Panama (three strains), Australia (four strains) and three strains isolated from frogs imported h m Africa; only five variable nucleotide positions were detected among ten loci (5,918 base pairs) (Morehouse et al. 2003). These results suggest that B. dendrobatidis is a widespread, recently emerged clone and has been spread since continental break up.

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Fig. 2. Transmission electron micrograph of zoospores within a zoosporangium in the skin of Bufo marinw. Zoospores are being released and contain numerous lipid globules that are partially surrounded by the microbody and occur at the edge of the ribosomal mass. Bar = 2 pm. F = flagellum, N = nucleus, R = ribosomes, Mb = microbody, L = lipid droplet, NFC = nonflagellated centriole, K = hnetosome, M = mitochondria, TP = terminal plate, V = vacuole, ER = endoplasmic reticulum, MT = microtubules. From Berger et al. (2005a).

In a phylogenetic hypothesis of the Chytridiales based on 18s rDNA sequences, B. dendrobatidis lies outside the four major clades in the order games et al. 2000); in more recent analyses based on additional gene sequences, however, Batrachochytrium affiliates with members of the "Rhizophydium-clade" (T. Y. James, pers. comm.). Some features of its zoospore ultrastructure also are characteristic of the Rhizophydium-clade (Longcore et al. 1999; Letcher et al. 2004). The Rhizophydium-clade consists primarily of chytrids classified in the genus Rhizophydium plus unidentified or misidentified isolates that are associated with this clade on the basis of their zoospore ultrastructure or because of similar gene sequences Uames et al. 2000; Letcher and Powell 2004; Letcher et al. 2004). Members of the genus Rhizophydium form simple, frequently spherical, inoperculate zoosporangia that develop directly from the enlargement of the encysted zoospore. Some isolates that are now affiliated with this clade because of their ultrastructure or DNA sequence develop exogenously; that is, a germ tube forms from the encysted zoospore through which the nucleus migrates to a swelling that becomes the zoosporangium. This is the type of development that results in some chytrids being able to insert their nuclear material into plant cells (e.g., Longcore 1995) and is how Batrachochytrium may gain access to the inside of epidermal cells of amphibians. D. Biology

Chytrids, with a few exceptions, reproduce by forming asexual reproductive zoospores. The life cycle of Batrachochytrium dendrobatzdis has two main stages: (1) the motile, waterborne,

2996

AMPHIBIAN BIOLOGY

Fig. 3. Transmission electron micrograph of a cultured zoospore. The nonflagellated centriole is parallel to the kinetosome. The microtubule root runs parallel to the kinetosome and is embedded in a cone of ribosomes. Bar = 0.6 p m Abbreviations as in figure 2. From Berger et al. (2005a).

unwalled, short-lived zoospore that functions in dispersal and (2) the stationary zoosporangium (or sporangium) that engages in asexual amplification (Figs 5-21). This fungus does not produce hyphae. In pure culture on nutrient agar the morphology of B. &mirobat& is like that of most members of the Rhizophydzum-clade. It produces simple thalli that are anchored to the substrate by root-like rhizoids that also serve to increase the surface area that absorbs nutrients. As nutrients are absorbed, the body of the fungus increases in diameter ( 1 0 4 0 pm), becomes multinucleate by mitotic divisions, and at maturity the entire contents of the thallus, which is now called a zoosporangium (= zoospore container), cleaves into zoospores. The number of zoospores per sporangium varies (several to 100s) depending on the size of the sporangium. Sporangia are larger in culture than in frog skin. During growth, one or more discharge papillae form on the zoosporangium; up to six have been

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2997

NFC

Fig. 4. Transmission electron micrograph of a cultured zoospore. The nucleus is not associated with the kinetosome and is nested in the ribosomal mass, which is surrounded by endoplasmic reticulum. Mitochondria are adjacent to the ribosomal mass. Bar = 1 pm. Abbreviations as in figure 2. From Berger et al. (2005a).

observed. They vary in length, depending on microhabitat, from negligible up to 10 pm. After zoospores have been released, only the walls of the empty, clear sporangia remain. In culture, the cycle from zoospore to production of new zoospores takes about 4 5 days at 23°C (Longcore et al. 1999). Mature zoosporangia may stay dormant if the environment is too dry for zoospore release. If zoospores attach to a suitable substrate, they are capable of starting a new cycle. Zoospores of B. dendrobatidis are about 3 4 pm in diameter with a 19-20 pm flagellum. They are frequently spherical but can be elongate and amoeboid when first released from the zoosporangium (Longcore et al. 1999). After a period of motility and dispersal, the zoospore encysts. Although zoospores are motile, their dispersal distance is short ( < 2 cm under experimental conditions) (Piotrowski et al. 2004). Dispersal on a larger scale is probably via passive transport of zoospores by water or by some agency moving substrate containing thalli. Zoospores of some species of chytrids display chemotaxis towards their particular substrate, thus enabling them to reach new substrates that are not abundant in the vicinity. Chytrid zoospores probably do not require an exogenous energy source and

AMPHIBIAN BIOLOGY

Fq. 3. T r a n ~ s i o nelectron micrograph of an encysted zoospore. The resorbed flagellum is visible and a cell wall has formed. Ribosomes are d i r m i t e d throughout the cytoplasm. Bar = 2 pm. F = flagellum, N = nucleus, M = mitochondria. From Berger et al. (2005a). Fy. 6. P k a n n k g electmn micrograph of a germling showing fine rhizoids spreading out along the substrate. The culture was grown on a p k e cox-ershp and prepared by freeze-drying. The crumpled surface is an artifact of freeze-drying. Bar = 10 pm. From Berger et aL ( 3 0 0 5 a ~ FK. 7. Live immanm sporangium with rhizoids spreading out. Bar = 10 pm. From Berger et al. (2005a). fig. 8. Tmmmkion electron micrograph of an immature colonial sporangium in the skin of Litoria gracilenta. A septum ( S ) divides the thallus into two comparnnents. Bar = 5 pm. V = vacuole, G = golgi, M = mitochondria. From Berger et al. (2005a). fig. 9. Transmission electron micrograph of an immature sporangium with a discharge papilla. The cell is multinucleate after mitotic divisions, but tbe q ~ o p k r mhas not yet divided. The plug bloclung the discharge papilla is clearly seen (arrowhead). The wall over the tip of the plug has dissoh-ed, demonstrating that B. dendrobatzdis is inoperculate. Early stages often have large vacuoles (V). Transverse sections of rhizoids occur in spaces between sporangia. Bar = 5 pm. N = nucleus, M = mitochondria. From Berger et al. (2005a). fig. 10. Transmission electron micrograph of a multinucleate sporangium that is beginning to cleave into zoospores. The arrow indicates a cleavage h e . Bar = 4 pm. N = nucleus, F = flagellum. From Berger et al. (2005a).

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Fig. I I . Transmission electron micrograph of a sporangium in skin of Litoria gracilentu with a cytoplasm that has divided into incompletely formed flagellated zoospores. Bar = 5 pm. N = nucleus, M = mitochondria, F = flagellum, V = vacuole. From Berger et al. (2005a). Fig.12. Scanning electron micrograph of a bulk-frozen hydrated sporangium that has been freeze-fractured. The image is a three-dimensional representation of the similar staged sporangium shown in figure 11. Bar = 5 pm. A = agar. From Berger et al. (2005a). Fig. 13. Live sporangia with discharge papillae. Internal structures of the sporangia are at various stages of zoospore development. Bar = 20 pm. From Berger et al. (2005a). Fig. 14. Scanning electron micrograph of a large zoosporangium on agar with five papillae visible. Zoospores are congregating - - - and encysting . - around the base. Bar = 10 pm. From Berger et al. (2005a). Fig. 15. Transmission electron micrograph of a mature zoosporangium with discharge papilla and plug. The sporangium is packed with flagellated zoospores. Bar = 10 pm. F = flagellum, M = mitochondria, R = ribosomal mass, N = nucleus. From Berger et al. (2005a). Fig. 16. Scanning electron micrograph of a zoosporangium on agar with a long discharge tube. Bar = 10 m. From Berger et al. (2005a).

3000

'

AMPHIBIAN BIOLOGY

Fig. 17. Sporangia that have released most of their zoospores. Bar = 10 pm. From Berger et al. (2005a). Fig. 18. Culture on agar plate. Colonies appear as granular, cream coloured mounds. Fig. 19. Scanning electron micrograph of a cluster of sporangia grown on a plastic coverslip and freeze-dried. Some sporangia have LWO or more open discharge tubes. The threadlike rhizoids hold sporangia together. Bar = 10 pm. From Berger et al. (2005a). Fig. 20. Scanning electron micrograph of thalli with two discharge tubes demonstrating the aptness of the name "chytrid (i.e., earthen pot). Rhizoids from adjacent sporangia are growing over the surface. Bar = 10 pm. From Berger et al. (2005a). Fig. 21. Diagram of the lifecycle of Batracho~h~trium dendrobatidis in culture. After a period of motility, zoospores encyst, resorb their flagella and form germlings. Rhizoids appear from one or more areas. Sporangia grow larger and mature over 4-5 days. The sporangia become multinucleate by mitotic divisions and the entire contents cleave into zoospores while the discharge tubes form. The discharge tube is closed by a plug that absorbs water and deliquesces when zoospores are ready for release. Some thalli develop colonially with thin septa dividing the contents into multiple sporangia each with its own discharge tube. A= zoospore, B = germling, C = immature sporangium, D = monocentric zoosporangium, E = colonial thallus. From Berger et al. (2005a).

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their metabolism is directed towards production of energy used in flagellar movement and in maintenance of homeostasis (Fuller 1996). In the laboratory about 50% of zoospores of B. dendrobatzdis remained motile after 18 hours and 5% were motile after 24 hours (Piotrowski et al. 2004). Zoospores survive longer at low temperatures (Berger 2001). During encystment the flagellum is rapidly resorbed and a chitinous cell wall forms. Zoospores of many fungi produce an adhesive as they encyst on their host (Bartnicki-Garcia and Sing 1986). Zoospores of B. dendrobatdis have not been observed in the act of infecting skin, so the method of penetration remains uncertain. Longcore et al. (1999) suggested that the zoospore could encyst on the surface then inject the nucleus and contents through a germ tube. Batrachochytrium differs from other members of the Rhizophydium-clade in that early in development one or more walls form within some thalli (Fig. 8). Instead of a single zoospore forming a single zoosporangium, several abutting zoosporangia, each with separate discharge papillae, form from a single zoospore. These are called "colonial sporangia" (Longcore et al. 1999). The occasional occurrence of colonial sporangia is one of the diagnostic features that aids in identifying B. dendrobatidis in fresh skin or in stained sections.

Sexual reproduction has not been observed for most chytrid species but within the group diverse methods have been reported (Sparrow 1960). Sexual reproduction may occur by zoospores fusing with each other, zoospores fusing with sporangia, rhizoids hsing, or production of motile gametes of unequal size (Ban- 1990). Sexual reproduction usually results in the formation of a thick-walled resting spore (e.g., Miller and Dylewski 1981). Resting spores are long-lived and resistant to extremes of heat and temperature. They may survive for decades and can then become reanimated by rainfall and grow rapidly (Powell 1993). A few species of Rhizophydium have been reported to reproduce sexually (Sparrow 1960) but sexual reproduction in this genus has not been confirmed in pure culture. Asexually produced resting sporangia (resting spores) are formed by many Rhizophydium species but during the time that B. dendrobatidis has been studied in culture and in amphibian skin, no resting spores have been found. Studies of multilocus sequence-typing indicate that B. dendrobatidis reproduces clonally; this supports the lack, or uncommon occurrence, of a sexually produced resting stage (Morehouse et al. 2003). Batrachochytrium dendrobatidis is well adapted to living in the dynamic tissue of the stratified epidermis of amphibians. Sporangia live initially inside deeper epidermal cells and have a rate of development that coincides with the maturing of the cell as it moves outwards (Berger et al. 2005a). They grow initially in living cells but complete their development in dead keratinized cells that are soon shed from the surface. Discharge tubes push through the epidermal cell membranes and open onto the surface of the skin. These specialized adaptations suggest that the association of B. dendrobatidis with a cutaneous habitat has had a long evolutionary history. 2. Nutm'tion and Saprobic Growth

Some species within the Chytridiomycota are saprobic on various organic substrates in water and soil, such as pollen, cellulose and other plant material, chitin from insect cadavers, or keratin from hair and skin. These species are important primary biodegraders. Others are parasites, including pathogens, of plants, algae, protists, crustaceans, nematodes and insects (Sparrow 1960; Powell 1993). Batrachochytrium dendrobatidis is unique in the phylum in that it is pathogenic to a vertebrate. Thus far, it is known only from the superficial keratinized epidermis of amphibians. Zoospores appear to infect deeper cells of the stratum granulosum and the developing sporangia are carried to the skin surface as the cells cornify. The changes in distribution of sporangia in tadpoles during development and metamorphosis tracked changes in the distribution of keratin (Berger 2001; Marantelli et al. 2004), confirming the requirement of B. dendrobatzdis for a stratified, keratinized epidermis when occurring as a parasite. Immature sporangia grow within the deeper cells that contain prekeratin, not the dense keratin of the outer, dead, cornified layer where mature

3002

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AMPHIBIAN BIOLOGY

zoosporangia are found. Keratin has not been studied in detail in amphibians, but in mammals it is formed when the inner surface of the plasma membrane thickens and microfilaments, keratohyaline and lysed material are deposited into an amorphousfilamentous complex (Fox 1994). Some keratin is present in all layers of the epidermis, but there are many different types encoded by a large family of genes (Alberts et al. 1983). As cells mature, different keratins are expressed. It is not yet known which nutrients are absorbed by sporangia in frog skin. Azocasein and gelatin were degraded by proteases in supernatant from B. dendrobatzdis broth cultures, but keratin azure was not broken down (F'iotrowski et al. 2004). The ability of B. dendrobatidis to grow in pure culture in 0.5-2% tryptone broth (Piotrowski et al. 2004), in sterile lake water for up to seven weeks (Johnson and Speare 2003), and in sterile moist river sand for up to three months Uohnson and Speare 2005) suggests that this fungus also may exist in the environment apart from the skin of amphibians. Piotrowski et al. tested the growth of B. dendrobatdis in pure culture on various substrates and at various temperatures and pHs. Of the nitrogen sources tested, it grew best in 1% tryptone broth; it also grows well, however, in 1% peptonized milk, and produces a protease that breaks down skimmed milk (Piotrowski et al. 2004). Growth was sparse in snake skin autoclaved with distilled water compared with growth in a medium containing tryptone. Different carbon sources added to 1% tryptone liquid media did not increase the growth of the hngus and glucose at levels of 3.6% inhibited growth. Although 1% liquid tryptone medium has provided the nutrition needed for an isolate of B. dendrobatidis to remain alive in culture as long as seven years (transferred at 4-5 month intervals and storage at 5"C), the inability of zoospores to begin growth on 1% tryptone or mTGh agar, unless they are in groups, suggests that these media are not optimal. 3. Temperature Tolerance

Chytridiomycetes have been found in almost every type of environment, including rainforests, deserts, arctic tundra and fresh waters; a few grow in the sea (Barr 1990). Temperature is critical for chytrids that cannot tolerate ambient temperatures at some times of the year without a resting stage. Many chytrids grow at temperatures of 30°C and above (Longcore 1995; Barr 2000); but some require lower temperatures, e.g., 23°C for Laucstromyces hiemalis (Longcore 1993). Batrachochytrium dendrobatidis also requires lower temperatures (Piotrowski et al. 2004). It grows rapidly at temperatures between 17" and 25°C and slower at 10" and 5°C. At 28°C it does not grow but survives, as shown by resuming growth after being placed at optimum temperatures. After eight days at 30°C half of tested cultures failed to revive when placed at optimum temperature. This upper thermal limit for B. dendrobatidis falls within ambient summer temperatures in many parts of the world. This suggests that the severity or persistence of infection is likely to differ by climatic zones and from year to year depending on the temperature peculiarities of individual years. Cultures died within 4 hours at 37"C, within 30 minutes at 47°C and within 5 minutes at 60°C (Table 1) (Berger 2001; Johnson et al. 2003). Without a resting stage, Batrachochytrium may be unable to persist outside of amphibians when water and soil temperatures exceed 25°C for an extended time. When in amphibian skin, however, the temperature regimes experienced by the fungus will be affected by its hosts, which may behaviourally raise or lower their body temperatures compared with ambient levels. The present authors were unable to isolate B. dendrobatzdis from tissue samples that had been frozen, but whether the infection survives in frogs that spend winters in a h z e n state (e.g., R a m sylvatica) is unknown. Cultures survive freezing only when specialized freezing methods and cryoprotectants are used (Boyle et al. 2004). -1- pH Tolerance

Isolates of B. dendrobatidis that were tested for growth in 1% liquid tryptone medium at pHs of 6 and 7, with less growth at pH 8 (F'iotrowski et al. 2004). Although

Fbest

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS Table 1. Times at which all sporangia were killed at different temperatures.

Temperature (iC)

Time at which all sporangia were killed

100 60 47 37 32 26 23

1 min 5 min 30 min

4 hr 96 hr No death No death

3003

growth was minimal at pHs of 4 and 5, after two weeks incubation zoospores were present in these acidic cultures. This means that some growth and shedding of zoospores was taking place. B. dendrobatidis did not survive, however, in an acidic (pH 4.1) sterile potting mix (Johnson and Speare 2005). In the wild pH tolerance may be lowered if the hngus lives saprobically and unprotected by the epidermis of amphibians.

5. Tolerance to Salt In culture, B. dendrobatidis tolerates low salinity and zoospores will encyst and grow (although stunted) in 6.25 mglml NaCl (0.6%)but not in 12.5 mglml (Berger 2001). A 50 mg/ml NaCl solution killed cultures in five minutes Uohnson et d.2003). Salinity of seawater is about 35 mglml (3.5%). Hyla chrysoscelis tadpoles experimentally exposed to B. dendrobatidis zoospores in water with copper chloride, up to 3.18 pgnitre, did not differ in infection levels from those in water without copper (Parris and Baud 2004). 6. Desiccation

Because the delicate zoospores are un-walled and can easily succumb to desiccation, chytrids require free water for reproduction. Most chytrids occur in aquatic habitats or are active when terrestrial habitats are wet. Although species that produce resting spores can survive desiccation, cultures of zoospores and zoosporangia of B. dendrobatidis were killed by complete drying within an hour (Berger 2001; Johnson et al. 2003). E. Chytridiomycosis: The Disease 1. Mortality Rates and Incubation Times

Mortality rates of 100% occurred during natural outbreaks in captivity and in the initial transmission experiments in captivity that used frogs known to be from highly susceptible species (Berger et al. 1998; Longcore et al. 1999; Berger 2001; Nichols et al. 2001) (Table 2). Incubation times during experimental exposures with susceptible species varied from 9 to 83 days, with most frogs dying between 18 and 70 days post-exposure (Berger 2001; Nichols et al. 2001; Woodhams et al. 2003). The time until death varied with fungal dose and fungal strain (Berger 2001; Carey et al. 2003). Laboratory conditions with constant temperatures and small volumes of still water may result in higher mortality rates than those occurring in the wild. Infection is not fatal in all species of amphibians, and apparently healthy amphibians may frequently carry light infections (Mazzoni et al. 2003; Retallick et al. 2004; McDonald et al. 2005). Surveys of healthy frogs have revealed quite high prevalences (see section B. 1). Even heavy doses of inoculum failed to infect all bullfrogs (Rana catesbeiana) and no experimental animals became heavily infected (Daszak et al. 2004). Experimental infections of R a m yavapaiensis, R. boylii and Ambystoma tigrinum resulted in variable infection rates and almost all infected animals survived (Davidson et al. 2003) (Table 2). Mortality rate varies greatly among species, even when infected and housed under the same conditions. Mortality rate was 95% in Litoria caerulea, 65% in L. chloris, 95% in Mixophyes fasciolatus and nil in Limnodynastes tmmniensis (Ardipradja 2001). Time until death also varied greatly among species (Table 2). Young frogs are more susceptible than are older ones. In experimental infections in Dendrobates tinctorius only recently metamorphosed frogs died (313) while three sub-adults

3004

,

AMPHIBIAN BIOLOGY

and three adults became infected but did not die (Lamirande and Nichols 2002). The adults lost infection by 45 days. Similarly, mortalities in captive Bufo marinus occurred mostly in metamorphs, with few deaths in juveniles and none in adults kept in the same room (H. Parkes, unpubl. data). In many other species, however, such as Litoria caerulea, adults are highly susceptible (Berger et al. 2004). High temperatures can reduce mortality rates. In a transmission experiment with Mixophyes fasciolatus, 818 died at 17"C, 818 died at 23°C but 418 survived at 27°C (Berger 2001; Berger et al. 2004) (Table 2). In experimental infections in Litoria chloris, four groups of ten frogs each were held at (1) naturally fluctuating temperatures (range 13.5 to 23.2"C; (2) constant 20°C, and (3; 4) naturally fluctuating temperatures except for two eight-hour periods when they were subjected to either 8°C or 37°C (Woodhams et al. 2003). All ten frogs subjected to. 37°C survived for at least five months and infections were not found in those examined histologically at nine months. All other frogs died except for one in the 8°C group. Tadpoles of Dendrobates tinctorius that were infected at 23°C died sooner (0-15 days) after metamorphosis than those held at 25°C (1-23 days) (Lamirande and Nichols 2002). Low temperatures delayed the onset of disease, however, when postmetamorphic frogs were infected. In the experiment on temperature with Mixophyes fasciolatus, frogs at 27°C died on average at 21.5 days, while groups at 17" and 23°C both died on average at 40 days. In Litoria chloris, frogs at 20°C died after shorter times than did those in the fluctuating group or in the 8°C group. The higher mortality from, and prevalence of, chytridiomycosis at high elevations (Retallick and Dwyer 2000; McDonald et al. 2005; Woodhams and Alford 2005), are probably enhanced by sustained periods with temperatures at the optimum for the fungus and without any high enough to deter the fungus. The average temperature of some high-elevation tropical streams is 23"C, which is the optimum for Batrachochytrium in culture. 2. Clinical S i p

Frogs may have subclinical chytridiomycosis with no obvious changes. Body condition and fluctuating asymmetry of hind limbs in infected rainforest frogs did not differ from those attributes in uninfected frogs (Woodhams and Alford 2005). Clinical signs of severe chytridiomycosis are non-specific and a clinical diagnosis of chytridiomycosis can be provisional only. Chytridiomycosis can be confirmed only by laboratory tests. Typical clinical signs in frogs with severe chytridiomycosis are manifest in three ways: behavioural changes, neurological signs and skin lesions. The behavioural and neurological signs include lethargy, inappetence and sitting unprotected during the day with hind legs slightly abducted (Fig. 22) (Berger et al. 1999; Lips 1999). Burrowing frogs are found uncovered and arboreal frogs are seen sitting on the ground. Frogs in early stages of becoming symptomatic display some escape activity and have a rather rapid righting reflex but, if turned over two or three times, they rapidly tire and they respond slowly. Some species become rigid and tremble with extension of the hind limbs and flexion of the forelimbs, particularly when handled (Speare 1995). Frogs usually become moribund within two to five days of exhibiting lethargy (Berger 2001; Nichols et al. 2001). Skin lesions range from subtle to more obvious changes and include darkening and patchy discoloration of skin, reddened toe tips, presence of excessive sloughed skin, erosions and, less commonly, ulcerations (Figs 22-25) (Berger et al. 1999; Pessier et al. 1999). Excessive amounts of shedding skin were noted 12 to 15 days after experimental exposure (Nichols et al. 2001). Some species such as Litoria caerulea often become intensely hyperaemic on the ventrum, legs and feet. Although disease induced experimentally in Mixophyes fasciolatus had a moderately long incubation period, frogs were active and ate normally until they showed sudden behavioural changes and died within a few days (Berger 2001; Berger et al. 2004). Even frogs with heavy

Table 2. Suiniliary of selected chytridion~~cosis infection experiments. Cultured fungi were used for infections unless noted otherwise.

Species1 Life stage

Dose per frog (Strain)

Mortality rate due to chytridiomycosis (additional mortality)

Mean days until death from chytridiomycosis (Range)

24°C

616

15.2 (10-18)

Housed individually

Berger et al. (1998)

013 313 313

-

Housed individually

Berger et al. (1999)

39.3 (35-47) 28.7 (23-38)

Temperature

Comments

Reference

Mixophyes fasciolatus Juveniles

3 000 sporaugia in skin scraping (M. fasciolatw)

Mixophyes fasciolatus Juveniles

10 zoospores 100 zoospores 1 000 zoospores (Melbourne-Ldumerilii98-LB-1)

20-22°C

Dendrobates auratus

2 0 0 ~ 1zoospores and zoosporangia

20-25°C

616

22.5 (16-26)

Housed individually or in pairs

Nichols et al. (2001)

Litoria caerulea Juveniles

50 000 zoospores (Gibbo River-Llesueuri00-LB-1) (RockhamptonLcaerulea-99-LB-1) (Melbourne-Ldumerilii98-LB-1)

16-20°C

15115

19.4 (9-28)

Housed individually

Berger (2001); Berger et al. (2005b)

14114

37.9 (30-67)

15115

32.7 (24-52)

5 000 zoospores (Gibbo RiverLlesueuri-00-LB- 1)

15-25°C

19/20

28.8 (18-60)

~Ionsetliilclividually

Ardipradja (200 1)

19/20 13/20

52.1 (28-84) 65.1 (29-106)

0118

-

Litoria caerulea Mixophyes fasciolatus Litoria chloris Limnodynastes tasmaniensis Juveniles Dendrobates tinctorius Adults

20-25°C

013

-

Subadults recent metamorphs

1 x 10" 4 days Zoospores 1 x lo6 x 4 days 1 x lo6 x 4 days

20-25°C 20-25°C

013 313

< 32

Tadpoles

1 x lo6 x 4 days

20-25°C

10110

Tadpoles

1 x lo6 x 4 days (D. aureus)

20-23°C

10110

1-23 post metamorphosis 0-15 post metamorphosis

Experiiiierit terillinated at 108 clays

Housed in groups. Adults and subadults diagnosed with irlfections fro111 9-1 1 days. Adults had none after 45 days. All survived to be treated with itraconazole from day 66.

Lamirande and Nichols (2002)

Table 2

- continued

w 0

Species1 Life stage

Dose per frog (Strain)

Ambystoma tigrinurn Juvenile and adult

9 000 zoosporeslml (Abystorna tigrinum)

Temperature

Mortality rate clue to chytriclio~~iycosis (additional mortality)

Mean days until death from chytridiomycosis (Range)

18°C

Rana boylii metamorphosed

Litoria chloris Juvenile

8 500 zoosporeslml (A. tigrinurn) 8 500 zoosporeslml (R. yauapaeinsis) 850 o 8 500 zoosporeslml (A. tigrinurn) 850 or 8 500 zoosporeslml (R. yavapaeinsis) 15 000 zoospores (Gibbo RiverLlesueuri-00-LB- 1)

Rana catesbeiana

5 infected R. catesbeiana

Mixophyes fasciolatus Juveniles

1 000 zoospores (Gibbo RiverLlesueuri-00-LB-1)

*Two 8-hr periods at 8°C or 37°C.

22°C

Comments Housed individually Infections detected in all salamanders at 9 days, all survived 60 days. Only one frog that died had obvious chvtridiomvcosis.

6 000 zoosporeslml (Rana yavapaeinsis)

Rana yavapaiensis metamorphosed

0 01

013

-

Reference Davidson et al. (2003)

Mortality rate was not different to controls.

(1)

22"

014 (3)

-

22°C

O/ 15 (5)

-

22°C

1/15 ((5)

51

20°C 13.5-23.2 13.5-23.3 (37OC)* 13.5-23.2 (8"C)*

lO/lO 10/10 0110 9/10

(28-55) (40-83) (34-72)

Housed individually

-

515

1 8 days

Housed together

17°C 23°C

818 718 (1) 418

40.0 (25-59) 40.0 (29-76)

27°C

21.5 (18-27)

Housed individually Infections that were detected in 314 survivors at 27C were eliminated by 98 days

Woodhams et al. (2003)

Mazzoni et al. (2003) Berger et al. (2004)

BERGER E T AL: FUNGAL DISEASES OF AMPHIBIANS

3007

Fzg. 22. Live adult of Litoria caeruha in the terminal stages of chytridiomycosis. The frog is weak a n 2 is sitting with legs abducted, andu the skin is severely reddened due to congestion.

............................................ I 1 ' vI Fzg 23 Capt~vemetamorph of Murophyes fmczolutus w t h termlnal chytnd~omycosls Note depressed I

1

I

Q/

-

atatude, pamally closed eyes and accumulanons of sloughed skm over the body. Bar = 5 mm From Berger et a1 (1999) Fzg 24 Formalm-futed adult of Lztwm caeruka wwlth d~scolourat~on and ulcers on dorsal skm. Bar = 25 mm Fzg 25 Dead adult of Mzxophyes jleayz w ~ t h chytr~d~omycosisThe thighs are swollen and there is reddening and dllat~onof blood vessels in the ventral skln of the h ~ n dl ~ m b s Bar = 24 mm (Photo by Harry B Hznes)

infections are able to appear and behave normally until some threshold is reached and signs of disease appear. Frogs with severe clinical signs invariably die. Most wild frogs in Australia found ill or dead with chytridiomycosis were in reasonable body condition with medium-sized o r large fat bodies (Berger 2001; Berger et al. 2004). Many females were gravid. Gross pathology of internal organs is generally unremarkable. 3. Pathology A. HISTOLOGY

On histological examination, sporangia of B. dendrobatidis are seen in the superficial epidermis and occur within cells in the stratum granulosum and stratum corneum. Immature, dark sporangia occur in the more viable cells deeper in the epidermis whereas mature zoosporangia and empty sporangia are niore prevalent in the outer keratinized layers, including layers sloughing from the surface (Berger et al. 1998; Pessier et al. 1999).

3008

AMPHIBIAN BIOLOGY

Skin lesions vary in severity (Figs 26-30). They are often mild, with hyperkeratosis over an intact epidermis being a common change. In hyperkeratotic areas, many keratinized cell layers build up to form a thick stratum corneum. Cell junctions to underlying skin appear to break down intermittently and then lifting of the thickened, infected stratum corneum occurs. Large numbers of sporangia are removed when skin is shed so the intensity of infection varies depending on the stage of this process (Berger et al. 2000). The shedding layers often remain on the skin of debilitated, terminal frogs and are seen grossly. Bacteria may colonize the layers of sloughing keratin and grow within "empty" sporangia.

-

'"

la, 3-

I*,

a -b

+?"*\*

-s

.

"w ' -i

*

i:

2

* .

Fzg 26 H~stolog~cal sectlon of healthy dorsal s k ~ nfrom an adult Lztorza caerulea The e p ~ d e r m ~1ss smooth and isr of even thickness P~gmentcells occur beneath the basement membrane The large serous glands (S) and smaller mucous glands (M) occur m the dermis and their aducts penetrate the e p ~ d e r m ~ sBar = 150 pm Stained -, , w ~ t hhematoxyl~nand eosm AM = substantza amorpha ** --+'" -CFzg 27 Sect~on of ~nfected s k ~ n from a captlve metamorph of Mzxophyes fasczolatus with mild ep~dermal les~ons The surface 1s eroded and there are some pyknot~ccells In the basal layer There 1s neglig~bleinflammation, but cap~llariesin the tela subcutanea are congested Bar = 80 pm Sta~nedw ~ t hhematoxyl~nand eosln Fig. 28. Infected skin from the toe of an adult Litoria chloris. The epidermis is mildly hyperplastic and there is loss of normal stratification. Many epidermal cells are pyknotic and vacuolated. Bar = 60 pm. Stained with hematoxylin and eosin. Fig. 29. Edge of an ulcer from the Litoria caerulea portrayed in figure 24. The remaining epidermis is thin (2-5 cells thick). Stained with hematoxylin and eosin. Fig. 30. Section of skin from a Mixophyes fasciolatus with focal hyperkeratosis associated with infection. There are inflammatory infiltrates in the dermis and epidermis. Stained with hematoxylin and eosin.

.,

.

2,-

-

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3009

Other lesions include irregular multifocal hyperplasia, disordered epidermal cell layers, spongiosis, erosions and occasional ulcerations of the skin (Fig. 29) (Berger et al. 1998; Pessier et al. 1999). An increase in numbers of epidermal mitotic figures may be seen (Davidson et al. 2003). The usually smooth surface becomes roughened and irregular. Width of the epidermis is highly variable with difhse or focal thickening in some regions as well as large areas of thinning. In some frogs the epidermis appears eroded and only a thin layer of one or two cells remains. Clusters of sporangia sometimes occur in deep pockets resulting from missing epithelium. Swollen or hypertrophic epidermal cells are evident in some frogs. Individual epidermal cell necrosis is commonly seen in scattered cells in the stratum basale or stratum spznosum. These cells have pyknotic nuclei and pale swollen cytoplasm (Fig. 28). Occasionally, vacuolated degenerate cells appear to coalesce into vesicles that result in lifting of the epidermis and in ulceration. This ballooning degeneration and cleavage occurs in the suprabasilar layer or between the dermis and epidermis. Some frogs have extensive ulceration leaving the basement membrane exposed. In these frogs, sporangia are lost with the epidermis and only a few remain. Ulceration appears to be more common in Litmia Caerulea. The inflammatory response is mild and may occur as a slight increase in mononuclear cells in the dermis. Foci of lymphocytes, macrophages and a few neutrophils sometimes occur in the superficial dermis, particularly in areas of ulceration. Occasional inflammatory cells are seen in the epidermis (Pessier et al. 1999). Salamanders (which survived infections) had mild cutaneous inflammatory infiltrates and minimal hyperkeratosis (Davidson et al. 2003). On sloughed skin, clusters of sporangia were associated with melanized spots, which were also obvious grossly. B. ELECTRON MICROSCOPY

Studies on infected frog skin (Litoria gracilenta) by electron microscopy (Berger 2001; Berger et al. 2005a) revealed a zone of apparently condensed host cytoplasm, up to 2.5 pm thick, around some sporangia. This zone appears to be mainly fibrils with no organelles (Figs. 3 1-33). The more superficial epidermal cells contain larger sporangia and host nuclei and organelles such as mitochondria are located on one side of the cell. Near the skin surface the epidermal cytoplasm condenses into a thin layer around the fbngal thalli and host organelles are lost as they are during normal epidermal cell maturation. Cell nuclei become dark and condensed but are not as flattened as in normal stratum corneum. Keratinization appears to occur prematurely in infected cells below the skin's surface, compared with uninfected cells in the same epidermal layer (Fig. 3 1) (Berger et al. 2005a). The cell junctions of infected cells usually appear normal. Some infected cells and uninfected cells near foci of infection are acutely swollen, although mitochondria and other organelles in these cells are intact. Nuclei of some infected cells in the stratum granulosum are shrunken and chromatolytic. Pathology in the deeper epidermal cells, as distant as the basal layer, includes focal shrinkage, increased intercellular spaces, vacuolation and dissolution of the cytoplasm (Figs 31-33). The hyperkeratosis appeared to be partly attributable to an increased turnover of epidermal cells. The swelling of epidermal cells near foci of infection suggests an hyperplastic response. Stimulation of the stratum basale leading to hyperplasia is a common response to epidermal injury and occurs with other epidermal infections such as that by the mite Sarcoptes scabiei (Skerratt et al. 1999). Sporangia appear to initiate premature death and keratinization of host cells. Thinning of the epidermis may occur when the germination of epidermal cells does not match the increased rate of sloughing caused by increased cell death. Other infected frogs may have a markedly thickened epidermis because of hyperplasia exceeding sloughing (Berger et al. 2005a). Scanning electron microscopy revealed that surface of skin from a healthy control frog was smooth and intact (Fig. 34), whereas skin from an infected Litoria lesueuri was rough

AMPHIBIAN BIOLOGY

Fig. 31. Transmission electron micrograph of infected epidermis in an adult Litoria gracilenta without clinical signs. Note multiple layers of dark infected keratinized cells, whereas away from the cluster of sporangia the stratum corneum is one cell thick. Infected cells contain between one and three sporangia. Some nuclei of infected cells are degenerate and chromatolytic (arrow head), and cells in the deeper epidermis (arrow) are necrotic with dissolution of cytoplasm. Bar = 18 pm. From Berger et al. (2005a). Fig. 32. Higher magnification of an infected cell taken from figure 3 1. A zone around the sporangium contains no organelles. Mitochondria and apparently normal cell junctions are present. Bar = 4 pm. From Berger et al. (2005a). Fig. 33. Transmission electron micrograph of superficial epidermal cells from an adult Litoria g r a c i h t a that died with chytridiomycosis. A clear fibrillar zone in the cytoplasm sumunds the sporangium (*); the chromatolytic nucleus (N) and necrotic organelles have been displaced. Bar = 6 pm. From Berger et al. (2005a).

because of separation of adjacent cells, irregular rounding of their normally flat surfacelayer, and desquamation (Fig. 35, 36). The above studies did not determine whether death of frogs was caused by a toxin released by B. dendrobatzdis or by direct inhibition of skin functions by sporangia, but toxicity to the skin is suggested by the dissolution of cellular cytoplasm (visible both by histological and transmission electron microscopical methods) in epidermal cells distant to foci of infection. Bacterial overgrowth may contribute to the pathogenesis in terminal stages. Recent studies on biochemical changes in the blood of terminal stage kegs revealed a large decrease in osmolarity and electrolytes (magnesium, sodium, calcium, chloride) (Berger et al., unpubl. data). As levels of protein and urea were unchanged, the decrease in electrolytes was not caused by dilution. Further work is needed to test whether skin is the major excretion pathway. C. OTHER LESIONS

It is rare to see specific internal lesions in frogs sick with chytridiomycosis. This suggests that the ultimate cause of death is metabolic or toxic. Necrosis, vacuolation or cloudy swelling is sometimes apparent in a range of internal organs, including focal or diffuse acute necrosis in renal tubules and possible oedema and vacuolation in the brain.

-

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3011

Histological examination of organs, such as the spleen and bone marrow, that are involved in immunity revealed no evidence of immunosuppression (Berger 2001). It has been suggested that frogs are stressed by environmental changes but, if so, one would expect reproductive and nutritional status to be affected before fatal immunosuppression occurs. In contrast, gravid moribund frogs were found (Mahony 1996) and many had adequate fat reserves (Pessier et al. 1999; Berger 2001; Berger et al. 2004). Also, with severe immunosuppression a range of opportunistic infections is likely to be involved. Although, in an Australian survey, concurrent diseases were diagnosed in 12% of frogs with severe chytridiomycosis, no other abnormalities were found in most frogs (Berger 2001; Berger et al. 2004). Experiments on susceptible species in captivity have demonstrated that B. denhobatidis can cause 100%mortality under conditions in which uninfected animals remained healthy. There is no evidence that immunosuppression is necessary for epidemics of chytridiomycosis to occur. Although fungi are most often secondary pathogens in mammals, in aquatic animals they are often highly pathogenic. 4. Distribution of Sporangia . Heavier infections than elsewhere occur on the ventral skin and feet (Nichols et al. 2001). Quantification of numbers of sporangia in skin from six sites of the body and four toes of ten Litoria caerulea that died with severe chytridiomycosis confirmed that very few . occurred on the back sporangia occurred on dorsal sites (Berger et al. 2 0 0 5 ~ )Sporangia of only one frog but were seen in the dorsal skin over the hind limb in four frogs. Heavy infections of sporangia occurred in all frogs at six ventral sites (mid-abdomen, axilla, pelvic

3012

AMPHIBIAN BIOLOGY

patch, tibia1 skin) and on the toes, but numbers were highly variable and no significant differences were noted. The variation may be related to the stage in the cycle of sloughing of the stratum corneum and because sporangia are not present in ulcerated areas. General differences noted between dorsal and ventral skin were that the dermis was thicker in dorsal skin and the substuntia amorpha (a granular layer containing calcium and polysaccharides believed to prevent dehydration) was thicker and more continuous dorsally. Greater numbers of serous glands occurred on the two dorsal sites and antihngal secretions from these glands may inhibit infection, although sporangia were seen growing at the openings of ducts of serous glands. Mucous glands were more evenly distributed over the body. Many frogs rely on ventral skin for water absorption and the regions where heavier infections occur are areas kept in contact with moist substrates. As zoospores of B. dendrobatidis require water for dispersal, the dryness of dorsal skin in terrestrial frogs could explain the distribution of infection. This conclusion is supported by the fact that in the aquatic Xenopus tropicalis, dorsal skin was infected as heavily as ventral skin (Parker et al. 2002).

5. Chytridiomycosis in Tadpoles A. E F F E C T S

Although B. dendrobatidis may cause fatal disease in post-metamorphic amphibians, it does not seem to kill tadpoles, which may be infected in the mouth parts (Berger et al. 1998). The amphibian chytrid has not been found growing on eggs. Infected tadpoles of Mixophyes fasciolatus and Rana muscosa generally appeared normal and healthy with large fat bodies (Berger 2001; Fellers et al. 2001; Marantelli et al. 2004) but quantitative studies have shown effects in Hyla chrysoscelis, Rana blairi, Bufo fowleri and R. sphenocephala (Parris 2004; Parris and Beaudoin 2004; Parris and Cornelius 2004). When tadpoles of these species were reared with infected frogs in large outdoor mesocosms, they had a reduced body mass at metamorphosis. Exposure to the fungus also resulted in slower development of H. chrysoscelis, but only when predatory newts (Notophthalmus virzdescens) were present (Parris and Beaudoin 2004). Hybrid R . blairi and R . sphenocephala exposed to B. dendrobatzdis were smaller at metamorphosis, had longer larval periods and had higher infection rates than did parental genotypes (Parris 2004). There was also an effect of competition between species. When infected B. fowkri and H. chrysoscelis were reared together they were smaller at metamorphosis compared to their size when reared separately (Parris and Cornelius 2004). That B. dendrobatzdis causes stress in the tadpoles of these two species was also shown by an increase in fluctuating asymmetry of hind-limb length (22% and 37% respectively) (Parris and Cornelius 2004). Hence, the effects of B. dendrobatidis on tadpoles are complex and are affected by community structure. Parris and Beaudoin (2004) suggested that reduced size may lead to lower juvenile survival and reduced reproductive success in animals that reach adulthood. Longer larval periods may lead to decreased survival in ephemeral bodies of water that dry up before metamorphosis occurs. Density of tadpoles of H. chrysoscelis (40, 80, or 120 per 750 litre tank) did not affect infection rates, which were 80% to 92% (Parris and Beaudoin 2004). When groups of captive-spawned tadpoles (Mixophyes fasciolatus and Bufo marinus) are infected, tadpoles appear healthy but mortality approaches 100%within 19 to 25 days after metamorphosis (Berger et al. 1998; Marantelli et al. 2004). Mortality rates in wild metamorphs are not known but survival of significant numbers is suspected (Marantelli et al. 2004). Experimental infections of tadpoles of Dendrobates tinctorius at Gosner stages 25 to 45 resulted in 100% mortality, which occurred from stage 45 till 23 days post-metamorphosis (Table 2) (Lamirande and Nichols 2002). Tadpoles carried infections for prolonged periods. Although effects on the health of tadpoles are not obvious, gross and histological abnormalities of the mouthparts may be severe. In a study on Rana muscosa, all infected

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3013

tadpoles examined had oral disc abnormalities such as misshapen and missing tooth rows, loss of pigmented structures and swollen and pinkish tooth ridges (Fellers et al. 2001). Histological lesions included depigmentation, epithelial hyperplasia and rounding of cutting edges. Rachowicz (2002) reported that loss of pigmented structures occurs naturally in the colder months, even without chytrid infections, but the pattern of temperature-induced pigment loss differs (Rachowicz and Vredenburg 2004). In tadpoles exposed to 4"C, pigment disappeared first from the tooth rows, and then from the jaw sheaths with a continuous decrease in width of the pigmented area. In tadpoles with chytridiomycosis, the jaw sheaths were affected before the tooth rows and had gaps and focal areas of depigmentation. Eventually, however, there was complete loss of all mouthpart pigmentation with both patterns (Rachowicz and Vredenburg 2004). Mouthpart abnormalities may affect feeding efficiency leading to the observed reduction in body size of experimentally infected animals (Parris and Beaudoin 2004). B. DISTRIBUTION O N TADPOLES

On amphibians, B. dendrobatidis occurs only in stratified, keratinized epidermis. As tadpole skin is not generally keratinized, the pathogen has a restricted distribution (Berger et al. 1998; Lamirande and Nichols 2002; Marantelli et al. 2004). In early tadpole stages, B. dendrobatidis occurs in the mouthparts, which are the only sites of keratinized epithelium in the body (Fig. 37'). In later tadpole stages, ventral skin of the feet becomes keratinized and zoosporangia begin to invade. This allows infection to be maintained when mouthparts are lost at metamorphic climax (Berger 2001; Marantelli et al. 2004). Tadpoles of Mixophyes fasciolatus may be infected at Gosner stage 25, which usually is the first stage after hatching. B. dendrobatidis may occur on the side of each tooth row on

Fig. 37. Ayoub-Shklar stained section through the horny beak of an infected tadpole of Mixophyes fasciolatw. Keratin (red) extends caudally towards the mouth and sporangia occur only within the keratinized area. From Marantelli et al. (2004).

AMPHIBIAN BIOLOGY

3014

the surface towards the mouth (Marantelli et al. 2004). Heavier infections occur on the jaw sheaths. Infection also extends caudally from the mouth a short way along the surfaces of the anterior buccal cavity. Infection of the mouthparts in M. fasciolatus was lost when tadpole mouthparts were shed at stage 42 (Berger 200 1; Marantelli et al. 2004). Marantelli et al. (2004) found that the feet of M . fasciolatus were first seen with a light infection of B. dendrobatidis at stage 42. As the tail resorbs, sporangia become established over the body and a thick infection develops by stage 45. The resorbing tails and tail stumps of infected animals contain extremely heavy infections (Berger 2001; Marantelli et al. 2004). Larval caudates had infections only on the tips of digits (Green et al. 2002). C. LARVAL INFECTION RATES

Batrachochytrium dendrobatzdis has been detected at high prevalence in free-living tadpoles. It occurred in 16 of 24 (67%) tadpoles of Rana muscosa in the Sierra Nevada, California (Fellers et al. 2001). All infected tadpoles in that study had abnormalities of the oral disc and all tadpoles with gross abnormalities were infected. Field surveys in the Sierra Nevada detected mouthpart abnormalities in 1581387 (41%) of tadpoles from 16/23 (70%) sites. Abnormal mouthparts consistent with chytridiomycosis were observed in 131106 (12%) and 151133 (11%) of tadpoles during and after a decline in Panama (Lips 1999) and in 601368 (19%) of tadpoles in Mexico (Lips et al. 2004). Infections with B. dendrobatidis were found in the mouthparts of 13 of 15 wild tadpoles of Mixophyes sp. from southeastern Queensland (Berger 2001). Infection rates vary among species from 1% (Litoria lesueuri) to 76% (Mixophyes schevilli) in tadpoles in northeastern Queensland (Woodhams and Alford 2005). The high infection rates observed in some species may be a result of tadpoles having long potential exposure to the water-borne zoospores, combined with survival of infected tadpoles. Surveying of tadpoles has been suggested as a sensitive means of detecting localities containing infections (Berger et al. 1999).

E Epidemiology 1. Seasonality and Thermal Effects

Low temperatures can tip the balance of the infective process in favour of the pathogen. In an Australian survey of wild ill and dead amphibians the incidence of chytridiomycosis was higher in winter, with 53% of frogs from Queensland and New South Wales dying in July and August (Fig. 38) (Berger 2001; Berger et al. 2004). Other diseases were detected mostly in spring and summer. Similarly, die-offs in frogs in Arizona due to chytridiomycosis occurred only in winter (Bradley et al. 2002). In Wyoming, of 58 free-ranging dead Bufo baxteri for which a cause of death was diagnosed, and that were mostly collected in September and October, 54 (93%) were diagnosed as having infections of Basidiobolus ranarum (Taylor et al. 1999a). These were, however, likely misdiagnosed cases of chytridiomycosis (Carey et al. 2003). Surveys of the prevalence of infection in apparently healthy frogs in Queensland and Western Australia have also shown that infections increase in colder months (Aplin and Kirkpatrick, 2000; Retallick et al. 2004; McDonald et al. 2005; Woodhams and Alford 2005). In the Wet Tropics, between 1998 and 2002, overall prevalence was 12.1% in winter and 3.7% in summer. In winter, the prevalences at different elevations were similar but in summer prevalence was greater at higher elevations (see section B.1.) (McDonald et al. 2005). In east-central Queensland, prevalence of infection in Taudactylus eungellensis was 37.8% in winter and 11.3% in summer (Retallick et al. 2004). In experimental infections in the laboratory, lower temperatures enhanced the virulence of chytridiomycosis in Mixophyes fasciolatus. All frogs exposed to B. dendrobatzdis at 17OC and 23OC died, whereas 50% of those exposed at 27OC survived (Table 2) (Berger et al. 2004). Infections in survivors were eliminated by 98 days. In experimental infections with Litoria chloris, frogs were held at naturally fluctuating temperatures (range 13.5 to 23.Z°C) except

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for two eight-hour periods when they were subjected to 37OC; the latter conditions apparently cleared the infection (Table 2) (Woodhams et al. 2003). An experimental translocation in Queensland involved moving Litoria rheocola from lowland areas to enclosures at three upland streams and three lowland streams. Mortality was higher and its onset more rapid at the higher elevations, and was also higher in winter

W C hytrid H Other

Month Fig. 38. Total numbers of ill or dead wild frogs examined from Queensland and New South Wales per month, comparing frogs diagnosed with severe chytridiomycosis with frogs with other diseases. Includes frogs submitted from October 1993 to December 2000. From Berger e t al. (2004).

than in summer. Chytridiomycosis was diagnosed in the dying frogs (Retallick and Dwyer 2000). The sensitivity of B. dendrobatidis to high temperatures may limit its distribution and the disease may not become established or affect frogs in locations where temperatures are consistently high (Retallick 2003). It is unknown whether the effect of temperature on severity and prevalence of disease is due mainly to effects on growth of B. dendrobatidis, or to a combination of reduced host immunity and increased fungal growth at lower temperatures. The loss of high-elevation populations in the tropics is likely attributable to a combination of susceptibility of the host and sustained optimal temperatures for the fungus.

2. Host Range and Effects on Different Species ofAmphibians The amphibian chytrid appears capable of infecting most species of anuran or caudate that occurs in suitable environments but the intensity of infection and the pathological effects appear to be strongly dependent on the species of host. Globally, infections have been detected in two orders (Anura and Caudata), including 19 families (Ambystomatidae, Amphiumidae, Bombinatoridae, Bufonidae, Centrolenidae, Dendrobatidae, Discoglossidae, Hylidae, Leiopelmatidae, Leptodactylidae, Mantellidae, Microhylidae, Myobatrachidae, Pipidae, Plethodontidae, Proteidae, Ranidae, Salamandridae and Sirenidae) and at least 144 species (Carey et al. 2003; Speare and Berger 2005a,b). Transmission experiments have shown that strains of B. dendrobatidis are not host specific (Davidson et al. 2003; Berger et a1. 2004). The low host specificity could be a factor facilitating the emergence of chytridiomycosis. Experimental and observational evidence show that susceptibility to chytridiomycosis varies widely among species. '

Declines have affected amphibian species to various extents. For example at Big Tableland, Queensland, Litoka genimaculata, a species then sympatric with the now extinct

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frog Taudactyus acutirostris, declined at the time other species disappeared (McDonald and Alford 1999). It apparently did not suffer such a high mortality rate, however, and the population has since recovered (McDonald et al. 2005). While Taudactylus eungellensis declined severely at Eungella National Park, sympatric Litoria lesueuri appeared unaffected (Retallick et al. 2004). Individuals of both L. genimaculata and L. lesueuri have been found dead with chytridiomycosis (Berger et al. 2004) and populations of these species 5-10 years after decline have moderately high infection rates (7.8% and 28% respectively) (Retallick et al. 2004; McDonald et al. 2005). The prevalence of infection was significantly different between species at Doolamai Falls, Eungella, with infection of 0% of 26 L. chloris and 27.1% of 129 T eungellensis (Retallick et al. 2004). Some species that have become established as feral pests are resistant. Healthy, freeranging Rana catesbeiana in Venezuela had a very high rate of infection (96%) and may be effective reservoir hosts (Daszak et al. 2004; Hanselmann et al. 2004). In Uruguay, infections are commonly seen in healthy animals from bullfrog farms without increased mortality (Mazzoni et al. 2003). High mortality rates occurred in captive Xenopus tropicalis but not in X. laevis, although subclinical infections were detected (Parker et al. 2002; Weldon et al. 2004). In captivity, high mortality rates occurred in metamorphic cane toads (Bufo marinus), while moderate mortality occurred in juveniles and no deaths were observed in adults in the same room, suggesting that cane toads are a relatively resistant species (H. Parkes and A. Hyatt, unpubl. data). Their dry skin may not be conducive to growth of B. dendrobatidis. Significant inherent differences exist among four Australian species (Ardipradja 2001). Mortality rates after experimental infections were: Litoria caerulea, 95%; L. chlorzs, 65%; Mixophyes fasciolatus, 95%; and Limnodynastes tasmaniensis, 0% (Ardipradja 2001). Time until death also varied (Table 2). Frogs were of similar ages but of different sizes; L. caerulea and M. fasciolatus are much larger species than the others. The juvenile frogs were infected with the same dose of zoospores inside 100 ml specimen jars that were rolled regularly to ensure exposure was not affected by behaviour of the frogs. At ten days post-exposure the number of frogs with detectable infection (determined by histology of toeclips), and the concentration of sporangia, was greatest in L. caerulea, lower in M. fasciolatus, and not detected in L. chloris. At death, however, M. fasciolatus had the greatest concentration of sporangia. Therefore L. chloris can prevent infection or reproduction of B. dendrobatidis whereas M. fasciolatus was able to withstand a heavier infection before reaching a terminal stage (Ardipradja 2001). Litoria chlorG has a waxy, lipid-based, waterproof coating on its skin that may have inhibited zoospore adhesion. Ardipradja (2001) suggested that a longer incubation time of the fungus could lead to increased survival of frogs in the wild because of more time available for environmental conditions to become unfavourable for fungal growth. Although free-ranging adults of L. tasmaniensis have died from chytridiomycosis (Berger et al. 2004), this species was not affected by the strain and conditions of this experiment (Ardipradja 200 1) and wild populations have not declined. Experiments with American species (Rana yavapaiensis, R. boylii and Ambystoma tigrznum), using both a strain isolated from A. tigrinum stebbinsi and one from R. yavapaiensis, resulted in infections in all salamanders but low infection rates in frogs. Some infections were eliminated during the experiment and, except for one frog, mortality that occurred in experimental animals was not associated with chytridiomycosis (Davidson et al. 2003). Behavioural differences among species of frogs have been suggested as a mechanism for variable susceptibility in the wild. Species that elevate body temperatures by thermoregulatory behaviour seem more likely to survive infections (Woodhams et al. 2003). Other differences among frogs in such things as production of cutaneous peptides, skin physiology or immune functions should be investigated. The susceptibility of some populations to decline may result from a combination of factors. Ecological variables that affect the life cycle of B. dendrobatidis and the robustness of its populations are important, as is inherent vulnerability to chytridiomycosis due to aspects of host physiology and biology. The declining species in Australia and Latin America

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have restricted geographic ranges in the uplands, have aquatic larvae associated with streams, spend a large proportion of their time in or adjacent to streams, and may have significantly smaller clutch sizes or large body sizes (Lips 1998; Williams and Hero 1998; McDonald and Alford 1999; Lips et al. 2003b). Populations of these species are less able to recover from declines from any cause and inhabit environments that are optimal for B. dendrobatzdis, i.e., cooler, wetter habitats. Some species may be highly susceptible, such as Litoria caerulea (Ardipradja 2001; Berger et al. 2004), but their wide distribution in the warmer lowlands may explain why they are still common. 3. Effect of Chytrzdiomycosis on Wild Amphibian Populations

Because the amphibian chytrid can be a highly virulent pathogen it has the potential of changing the population status of some species. Four patterns of response to the appearance of this fungus have been observed; (1) extinction of species, (2) extinction of local populations, but survival of the species, (3) population decline and variable levels of recovery, and (4) sporadic or mass deaths in relatively stable populations. All populations in which B. dendrobatidis has been found and which have been intensively studied appear to exhibit at least the minimal effect of occasional deaths due to chytridiomycosis. A. EXTINCTION O F SPECIES

Extinction is illustrated by the response of the sharp-snouted dayfrog, Taudactylus acutirostris. The status of this frog changed from abundant to endangered in 1992 and to extinct in 1999. This species was localized in Queensland in a range that extended in upland , south of wet tropics 310 km from Mt. Graham to Big Tableland (elevation 620 ~ n )just Cooktown (McDonald 1992). The first population declines were in 1989; declines then spread northward and the last remaining population at Big Tableland fell precipitously in late 1993 (Richards et al. 1993; Laurance et al. 1996, 199'7; McDonald and Alford 1999). Chytridiomycosis was found in wild frogs and was responsible for the death of frogs brought into captivity (Berger et al. 1998). Tadpoles of i? acutirostris that had been collected in September and December 1993 to be raised in captivity died from chytridiomycosis after they nletamorphosed (Mahony et al. 1999; Banks and McCracken 2002). Wild adults that were collected in September and October 1993 were taken to James Cook University where they also died within weeks of collection and were diagnosed with chytridiomycosis (Speare 1995; Berger et al. 1998; Mahony et al. 1999). B. LOCAL EXTINCTION B U T SURVIVAL O F T H E SPECIES

Chytridiomycosis also has caused extinctions of local populations. Examples of this pattern are provided by species from the Australian upland Wet Tropics: Litoria nannotis, L. rheocola and Nyctimystes dayi. These species followed the same pattern of decline as did 7: acutirostris, with extinction of all upland populations. Lowland populations on the same watercourses, however, persisted. In the lowlands, sporadic deaths continue to occur but population sizes appear to be stable (McDonald and Alford 1999; Speare et al. 2001). The spotted treefrog, Litoria spenceri, also showed a similar pattern when a stable and abundant population at Bogong Creek (elevation 1 100 m), Victoria, Australia, suddenly declined in 1996 and the last frog was seen in 1999 (Gillespie and Marantelli 2000). Chytridiomycosis was the cause of death in the one frog autopsied. In a retrospective survey of 95 toe clips from 1994 (16 toes), 1995 (59 toes) and 1996 (20 toes), the first appearance of the fungus was in March 1996 (Berger unpubl. data), the last time that frogs were seen in high numbers. In lowland populations at the other end of the distribution of L. spenceri, chytridiomycosis was found in dead frogs but the population did not decline (Gillespie and Hines 1999). C. POPULATIONS DECLINE B U T VARIABLY RECOVER

At Big Tableland, Australia, Litoria genimaculata, a species sympatric with Taudactylus acutirostris, L. nannotis and L. rheocola, suffered a sudden decline in numbers at the same time that T acutirostris disappeared (Laurance et al. 1996; McDonald and Alford 1999;

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McDonald et al. 2005). The population of L. genimculata, however, recovered to close to its former numbers after five years (McDonald et al. 2005). As chytridiornycosis still occurs in L. genimacu2ata and 8% of healthy frogs at Big Tableland were infected during 19982002, apparently a balance between L. genimculata and B. dendrobatidis is evolving (Speare et al. 2001; McDonald et al. 2005). Litoria nannotis and L. rheocola disappeared above'400 m during the original epidemic but their distribution is now moving upwards and low numbers of frogs have been sighted at 650 m (McDonald et al. 2005). However, recovery of populations has not occurred at high elevations e.g., 1 000 m, where frogs often remain absent. D. SPORADIC AND MASS DEATHS I N RELATIVELY STABLE POPULATIONS

A pattern of sporadic deaths due to chytridiornycosis is seen in many of the species infected by B. dendrobatdis in Australia (Berger et al. 2004). Once B. dendrobatidis has become established in a population of amphibians sporadic death is expected to be the minimum impact. Mass mortality also may occur regularly in common species in winter. Some populations appear to be stable but remain at lower abundance.

G. Resistance to Infection 1. Individuals

Innate resistance to B. dendrobatzdis appears to be present in some individuals or species that have survived exposure or infection while other individuals or populations have died (see section E). The mechanisms for this resistance are still unknown. All published work on immunity to chytridiomycosis treats the dermal antifungal peptides. These are produced in the granular (serous) glands and vary among amphibian species (Erspamer 1994). The peptides are believed to cause cell death by disruption of the cell membrane into peptide-coated vesicles (Rollins-Smith et al. 2002b). A range of peptides from amphibian skin is active against B. dendrobatulis in vitro in a concentrationdependent manner (Rollins-Smith et al. 2002a). These peptides are from the ranatuerin-1, ranatuerin-2, esculentin- 1, esculentin-2, brevinin- 1, brevinin-2, temporin, palustrin-3 and ranalexin families (Rollins-Smith et al. 2002a). The minimum inhibitory concentrations were between 2 and >25 microM. Zoospores are inhibited at lower concentrations than are zoosporangia. Berger (2001) found a minimum inhibitory concentration of 12.5 pglml for three peptides (a citropin and two caerins) from the Australian frog, Lit& caerulea, a species susceptible to fatal chytridiomycosis, but that has not declined. Even frogs that have declined, such as Rana tarahumarae, secrete peptides that inhibit growth of B. dendrobatzdis (Rollins-Smith et al. 2002b). It is not clear why frogs with apparently effective antimicrobial peptide defences are susceptible to chytridiomycosis. Perhaps there may not be enough peptides present on the skin to be effective (Rollins-Smith et al. 2002b). Studies on whether immunity can be acquired in individual frogs have not been reported. Infected frogs usually produce a negligible cellular inflammatory response in the dermis. When antifungal drugs or heat have been used to reduce infections to undetectable levels, but not eliminate them entirely, disease has been delayed but not cured. Once treatment stops, sporangia multiply to pathogenic levels again (G. Marantelli, unpubl. data). In many other diseases, a large reduction of pathogen burden enables the host to mount an effective immune response. 2. Populations A statistically significant decrease in prevalence of chytridiornycosis was monitored in Litoria genimculata in the Wet Tropics of northern Queensland between 1998 and 2002, five to seven years after the decline (McDonald et al. 2005). The decrease in prevalence of the disease was associated with recovery of these populations to near pre-decline levels, suggesting that if a species survives the initial epidemic, selection for innate resistance may

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occur naturally. This is also supported by the re-establishment of.Litoria nunnotis and L. rheocola at higher elevations. Although Taudactylus eungellensis crashed at Eungella National Park, central east Queensland, in 1985-1986, infection levels between 1994 and 1998 remained stable with an overall prevalence of 18% (Retallick et al. 2004). This finding suggests that a hostpathogen equilibrium is evolving (Retallick et al. 2004). These populations have a longer association with B. dendrobatidis compared with L. genirnaculata in the Wet Tropics. Even decades after initial declines, however, populations of many species remain at reduced abundance and inhabit reduced distributions. H. Transmission and Spread of Batrachochytrium dendrobatidis Batrachochytrium dendrobatzdis is most likely spread via water or by contact between frogs. It does not survive drying or immersion in seawater (Berger 2001; Johnson et al. 2003) and probably does not form a resting stage (Longcore et al. 1999). The hngus appears to spread slowly over the landscape by natural methods but movement over long distances, such as across oceans or deserts, is most likely by human-assisted translocation of infected amphibians, contaminated water or soil (Johnson and Speare 2003, 2005). Zoospores of B. dendrobatzdis are capable of swimming 2 cm (Piotrowski et al. 2004) and, although chemotaxis occurs in many chytrid species, water flow disseminates zoospores longer distances. Batrachochytrium dendrobatidis is highly infectious and even low experimental doses can result in fatal disease in susceptible species. The infective stage is the waterborne zoospore. In one experiment a dose of 100 zoospores caused fatal chytridiomycosis in all three Mixophyes fasciolatus exposed to inoculated water (Berger et al. 1999). Zoospores released from an infected amphibian can potentially infect other amphibians in the same body of water. The dynamics of infection in the wild, however, have not been studied. Chytridiomycosis was transmitted between infected and uninfected Rana muscosa tadpoles held together in 10 litres of water for three and four weeks, but not when held together for only one or two weeks (Rachowicz and Vredenburg 2004). Postmetamorphic R. muscosa housed with five infected tadpoles also became infected (Rachowicz and Vredenburg 2004). Parris (2004) transmitted infection to tadpoles by placing severely infected adult frogs in mesh cages in large (750 litre) tanks containing 60 tadpoles and complex environments.

During declines in Bufo boreas caused by chytridiomycosis, males disappeared faster than did females. This suggests that transmission occurred at the breeding ponds because males breed every season whereas females may only breed every second to fourth year (Muths et al. 2003). Consideration of the epidemiology of amphibian population crashes shows that B. dendrobatzdis has apparently spread independently from infected foci into adjacent areas, both along and between bodies of water. In Queensland, Victoria and in Western Australia the apparent rate of spread of chytridiomycosis has been approximately 100 km per year (Laurance et al. 1996; Aplin and Kirkpatrick, unpubl. data; Speare 2000, 2001). The mechanism of spread is unknown but probably involves normal movements of infected amphibians, or of water within bodies of water or of surface water, possibly during rain. Transport by birds has been suggested to explain large-scale movements. Johnson and Speare (2005) found that sporangia survived on feathers dried for 1-3 hours. Further studies using live birds are required. . Deliberate and accidental movements of amphibians are surprisingly large-scale phenomena and have been demonstrated to spread disease. Infected amphibians shipped in the pet trade between or within countries are capable of bringing chytridiomycosis to new areas. In Australia, three of six pet axolotyls were infected when purchased in Townsville and Perth (Speare 2000). In Germany, over 200 amphibians from the international pet trade have been found to have chytridiomycosis over a number of years (Mutschmann et al. 2000;

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Mutschmann, unpubl. data). Hymenochirus curtipes have been exported from Africa (Weldon et al. 2004) and trade in this species may have aided dissemination around the United States (Groff et al. 1991; Carey et al. 2003). Similarly, amphibians sold and moved for scientific studies can be potential infection risks. Xenopus laeuis and X. tropicalis caught in the wild in Africa and moved into scientific institutions in South Africa (Speare and Berger 2005b) and the United States (Reed et al. 2000; Parker et al. 2002) have been shown to be infected with chytridiomycosis. As Africa has been suggested as the origin of B. dendrobatidis, transport of X. laeuzs for science and pregnancy testing may have been how B. dendrobatidis originally escaped from its habitat of origin (Weldon et al. 2004). Transport of bullfrogs for food is a large international industry, with over one million frogs imported each year into the United States from South America and others shipped from Asia (Mazzoni et al. 2003). North American bullfrog stock has been introduced into South America numerous times (Hanselmann et al. 2004). As healthy bullfrogs may be infected with B. dendrobatidis at high prevalence, there is frequent opportunity for spread of the disease (Mazzoni et al. 2003; Hanselmann et al. 2004). Amphibians accidentally transported in agricultural produce also have the potential to move B. dendrobatidis long distances, either within or between countries. Tens of thousands of frogs are moved within Australia each year (Marantelli and Hobbs 2000) and native Queensland frogs rescued from produce in Melbourne were found to be infected with B. dendrobatidis (Marantelli and Hobbs 2000). Translocation and release of frogs for conservation purposes is also a potential risk for spread. Although transmission of B. dendrobatzdis between catchments by scientists working with amphibians is perceived as a risk, particularly where chytrid-infected and chytrid-free populations may be adjacent, there has been no evidence to support any spread via this means. The potential also exists to transmit chytridiomycosis between frogs in infected areas by handling, especially when a group of frogs is held in a common container before weighing or other measurements made. Transport of water, wet soil or other wet material could potentially move B. dendrobatzdis. Active zoospores were observed for up to seven weeks in sterile lake water (Johnson and Speare 2003). B. dendrobatidis survived for up to three months in sterile, moist river sand but did not survive in sterile potting mix, which was too acidic (pH 4.1) (Johnson and Speare 2005). I. Diagnosis of Chytridiomycosis

Chytridiomycosis in sick and subclinical frogs can only be diagnosed by laboratory tests. Healthy infected frogs show no signs of disease, and the clinical signs of severe chytridiomycosis are non-specific with similarities to those caused by other diseases, such as iridoviral infection and bacterial septicaemia ("red leg") (Cunningham et al. 1996). Diagnosis of chytridiomycosis involves the identification of B. dendrobatidis, either by light microscopy to visualize sporangia or by PCR (Polymerase Chain Reaction) to detect DNA. Culture is difficult and is not used for diagnosis (see Appendix I). ELISA protocols have been developed but are comparatively insensitive and non-specific compared to PCR (-A. Hyatt, unpubl. data). Electron microscopy is used mainly for research applications and is not used for diagnosis. Microscopy includes examination of wet skin preparations (scrapings, smears or whole skin), histological sections of skin stained with haematoxylin and eosin, and immunohistochemistry of skin sections. These routine tests have a high positive predictive value, and when used on diseased frogs with heavy infections they also have a high sensitivity. Healthy frogs, however, typically have light infections and only small samples can be

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obtained without sacrificing the animal. As the PCR tests are more sensitive than microscopy (Annis et al. 2004; Boyle et al. 2004), they are rapidly replacing the use of microscopy, particularly for surveys of large numbers of healthy, live frogs. Real-time PCR is highly sensitive and can detect B. dendrobatidis within one week of experimental infection. PCR on toe samples was eight times more sensitive than histology in early infection (Boyle et al. 2004). It is a quantitative test, giving an indication of the levels of infection (Boyle et al. 2004). Each technique, however, requires different skills and facilities. Different tests may be appropriate under different situations. These techniques may be used for surveying wild and captive amphibians, screening archived specimens, testing animals before translocation, and determining the cause of mortality in the wild and in captivity so that appropriate management can be implemented. Next, the methods used to diagnose the presence or absence of B. dendrobatidis in skin of healthy, sick or dead frogs are described. If diseased animals are found and thorough pathological tests are desired to determine the cause of death, then sampling of all tissues is required (see instructions on the Amphibian Disease Home Page: http://www.jcu.edu.au/ schoollphtm/PHTM/frogs/ampdis.htm). 1. Sampling

Sampling for diagnosis of chytridiomycosis requires knowledge of the anatomical sites most heavily infected and an understanding of the test being performed. The type of test determines how samples should be collected, transported and processed. A. SITES O F INFECTION

Adults: In adult amphibians, B. dendrobatidis is restricted to superficial keratinized epidermis. Infection in frogs with severe chytridiomycosis is heaviest on the ventral surfaces of the feet, abdomen and limbs (Berger et al. 2005b). Sampling can be conducted by collecting skin scrapings or smears, excising pieces of skin, or by swabbing. Toe-webbing and/or toes can be excized from live or dead frogs, and strips of skin from the inguinal area (pelvic patch) can be collected from dead animals. Details on swabbing are contained within the section on PCR. Tadpoles: The mouthparts of apparently healthy tadpoles can be infected with B. dendrobatidis (Berger et al. 1998; Marantelli et al. 2004). Infected tadpoles display abnormalities of the jaw sheath and tooth rows, which may be visible grossly or with a hand lens (see section 5A) (Fellers et al. 2001; Rachowicz and Vredenberg 2004). Gaps in the jaw sheath pigment are more certain indicators of infection than are changes in tooth rows (Rachowicz and Vredenberg 2004). These abnormalities, however, are only indicative of infection and diagnostic tests should be carried out, at least on a sample, to confirrn chytridiomycosis. Changes in mouthparts can also occur due to reduced temperatures and to chemicals (Rachowicz 2002; Rachowicz and Vredenberg 2004) and may be more common in some species. For microscopy, whole tadpoles can be collected and preserved. Tail stumps in metamorphs are a sensitive site to sample (Marantelli et al. 2004). Swabbing, which is non-destructive and therefore useful in monitoring the status of threatened or captive populations held for breeding, can be used to sample tadpole mouthparts for PCR (Hyatt, Boyle and Olsen, unpubl. data). The prevalence of chytridiomycosis in tadpoles may be high in some populations with long-lived tadpoles, and sampling tadpoles may be a sensitive way of assessing whether B. dendrobatidis is in a body of water (Berger et al. 1999). B. COLLECTION O F SAMPLES FOR HISTOLOGY AND IMMUNOHISTOCHEMISTRY

Amphibians degenerate rapidly after death. To preserve morphology the animal must be kept cool if not immediately transferred into an appropriate fixative (10% buffered formalin or 70% ethanol for light microscopy). Tissues should be frozen if bacterial or viral cultures are to be attempted.

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Although the quality of the morphological preservation is reduced in frozen, decomposed or mummified animals, it may still be possible to detect the presence of the fungus by histopathology and histochemistry. These techniques work well on formalinfixed paraffin-embedded samples and on museum specimens. If, however, samples have been stored in formalin for excessive periods of time, histochemical studies may-be unsuccessful (Hyatt, Boyle and Olsen, unpubl. data). The oldest global record, a specimen collected in 1938, was confirmed by an immunoperoxidase technique (Weldon et al. 2004). C. COLLECTION OF SAMPLES FOR POLYMERASE CHAIN REACTION (PCR).

A range of samples can be collected for diagnosis by PCR, including those collected for microscopy such as toe-clips, skin scrapings or excized strips of skin from the webbing or inguinal regions. Fresh, frozen or ethanol-fixed samples are preferable. Fixation in formalin for more than a short time (>three months) will prevent the detection of B. dendrobatidis by real-time PCR (Hyatt and Boyle, unpubl. data).

In addition, swabs and filtrates collected non-destructively can be tested by PCR. The use of swabs (e.g., Medical Wire & Equipment Co [UK] MW 100-100 sourced from Biomirieux Australia) is as sensitive as any other protocol for the detection of B. dendrobatzdis (Hyatt, Boyle and Olsen, unpubl. data). The underside of the feet (especially webbing), limbs and abdomen should be swabbed twice in adults, or the mouthparts swabbed in tadpoles. Swabs appear to be the best method of sampling in the field because fixatives or additional solutions are not required and permits for toe-clipping are becoming difficult to obtain. Samples can be stored at 23°C for one month, although Root, T. L., Price, J. T., Hall, K. R., Schneider, S. H., Rosenmeigk, C. and Pounds, A. J., 2003. Fingerprints of global warming on wild animals and plants. 'Vat. 421: 35-60.

Root, T. L, Slaan~nou&i, D. ,!I Masaandrea, M. D. and Schneider, S. H., 2003. Human-modifid temperams induce spxk changes: Joint tio on Aac. Nat Acod. Sci, USA. 102 746-7469. Shoemaker, 1'. H., Hillman, S. S., Hillyard, S. D., Jackson, D. C., McClanahan, L. L., I\ithen, F! C. and Wygoda, M. Y, 1992. Exchange of water, ions and respiratory gases in terrestrial amphibians. Pp. 125-150 in "Environmental Physiolop of the Amphibians", ed by M. E. Feder and \C IV. Burggren. University of Chicago Press, Chicago. Snyder, G. K. and Weathers, W. W'., 1959. Temperature adaptations in amphibians. Amel: Nut. 109: 98-101. Speare, R., 1990. A review of the diseases of the cane toad, Bufo marinus, with comments on biological control. Awt. Wldl. Res. 17: 387-410.

Pounds, J. A. and Crump, M. L., 1994. Amphibian declines and climate disturbance: the case of the golden toad and the harlequin frog. Cons. Biol. 8: 72-85.

Stallard, R. F., 2001. Possible environmental factors underlying amphibian decline in eastern Puerto Rico: analysis of U.S. government data archives. Cons. Biol. 15: 943-953.

Pounds, J. A,, Fogden, M. l? L. and Campbell, J. H., 1999. Biological response to climate change on a tropical mountain. Nature 398: 6 11-615.

Stewart, M. M., 1995. Climate driven population fluctuations in rain forest frogs. J. Herpetol. 29: 437-446.

Pounds, J . A,, Bustamante, M. R., Coloma, L. A,, Consuegra, J. A,, Fogden, M. l? L., Foster, l? N., La Marca, E., Masters, K. L., Merino-Viteri, A., Puschendorf, R., Ron, S. R., Sinchez-Azofeifa, G. A., Still, C. J. and Young, B., 2006. Widespread amphibian extinctions from epidemic disease driven by global warming. Nut. 439: 161-167.

Stuart, S. N., Chanson, J. S., Cox, N. A., Young, B. E., Rodrigues, k S., Fischman, D. L. and Mller, R. W., 2004. Status and trends of amphibian declines and extinctions worldwide. Sci. 306: 1783-1 786.

Retallick, R. W. R., McCallum, H. and Speare, R., 2004. Endemic infection of the amphibian chytrid fungus in a frog community post-decline. PLoS Biol 2: e35 1. Rome, L. C., Stevens, E. D. and John-Adler, H. B., 1992. The influence of temperature and thermal acclimation on physiological function. Pp. 183-205 in "Environmental Physiology of the Amphibia", ed by M. E. Feder and W. W. Burggren. University of Chicago Press, Chicago. Ron, S. R. and Merino-Viteri, A,, 2000. Declinacidn de anfibios del Ecuador: informacidn general y primer reporte de chytridiomicosis para Sudamerica. Froglog 42: 2-3. Ron, S. R., Duellman, W. E., Coloma, L. A. and Bustamante, M. R., 2003. Population decline of the jambato toad Atelopus ignescens (Anura: Bufonidae) in the Andes of Ecuador. J. Herpetol. 37: 116-126.

Thomas, C. D., Cameron, A,, Green, R. E., Bakkenes, M., Beaumont, L. J., Collingham, Y C., Erasmus, B. F. N., Ferreira de Siqueira, M., Grainger, A., Hannah, L., Hughes, L., Huntley, B., van Jaarsveld, A. S., Midgley, G. F., Miles, L., Ortega-Huerta, M. A., Peterson, A. T, Phillips, 0 . L. and Williams, S. E., 2004. Extinction risk from climate change. Nut. 427: 145-148. Vial, J. L. and Saylor, L., 1993. The Status of amphibian populations. Working document No 1. Declining Amphibian Populations Task Force. World Conservation Union, Workmg Document No. 1. iii + 98 Pp. Wake, D. B. and Morowitz, H. J., 1991. Declining amphibian populations - a global phenomenon? Findings and recommendations. Alytes 9: 33-42. Weygoldt, l?, 1989. Changes in the composition of mountain stream frog communities in the Atlantic mountains of Brazil: frogs as indicators of environmental deteriorations? Stud. Neotrop. Fauna Enuiron. 243: 249-255.

Indices to Amphibian Biology Volume 8 - Amphibian Decline: Dieseases, Parasites, Maladies and Pollution To facilitate usage of the index, it is divided into the following hvo indices: Subject Index

page 3282

Index of Scientific names

page 3287

AMPHIBIAN BIOLOGY

SUBJECT INDEX For major subject headings also see the Table of Contents (p. xi). Entries in boldface indicate that the indexed item, or part thereof, is illustrated in a figure as either a drawing or photograph.

A

abnormalities [also see deformities; malformations] 3067-3084, 30893109, 31 19, 3164-3165, 3208, 3222 Acadia National Park 2972 acclimation 3253-3254, 3261 acetochlor 32 19 acetylcholine 3229 acidification 3 116, 3132-3 134, 3 173, 3244-3261 acid rain 3149, 3245 activin 3210, 3222 ADCC (antibody-dependent cellmediated cytotoxicity 3074 Adelaide 2989 adenocarcinoma 2977-2978 adenoviruses 2963 adrenals 3231 Africa 2987, 2993-2994, 3020, 3032, 3034, 3091, 3107, 3240, 3246 African clawed frog 3157, 3220, 32233225 agenesis 3094 agent orange 3218 AGIP (Anonima Gas Italiana Petroli) 3240 Alaskan Wildlife Refuge 3 100 Alberta 3 104 albinism 3089 algae 3081, 3129-3126, 3130-3131, 3151, 3166, 3191, 3194-3195 algaecide 3 193 algal bloom 3 195 alligators 3216, 3222 alpine newt 3124, 3131 Alps 3126 aluminium 3246, 3251-3252 Amazonia 2980 Ambystoma tigrinum virus (ATV) 2964-2965, 2968, 3230 amelia 3094 America(s) 2964, 2987, 3016, 3043, 3135 American toad 3076-3078, 3105, 3152, 3157, 3190 ammonia(m) 3152-3154, 3157-3158, 3166, 3169-3175, 3177 ammonium nitrate 3158-3 160, 3 164, 3166-3172, 3175-3176 ammonium perchlorate 3220, 3223 ammonium sulphate 3 164 Amphibian Action Plan 3275 Amphibian Ark, Partners in Amphibian and Reptile Conservation (PARC) 3277 Amphibian Specialists Group (ASG) 3277 Andes 3126, 3270, 3272 androgen 3214, 3216, 3222, 3225, 3228, 3231 androstenedione 32 16 anoline lizards 3271 Anonima Gas Italiana Petroli (AGIP) 3240 anophthalmia 3094 Antarctica 3 107

anthracene 3 133 antibody-dependent cell-mediated cytotoxicity (ADCC) 3074 antimicrobial peptides 3059-3061 apoptosis 32 18 Appalachian Mountains 3272 arborviruses 2963 Arctic 3 196 Arctic Ocean 3212 Argentina 2965, 2991, 3241 Arizona 2965, 2970-2972, 2992, 3014 arochlors 3223 aromatase 3215-3216 arthrogryposis 3099-3 100 arthropods 3056 ascorbic acid 3 131 ASG (Amphibian Specialists Group) 3277 Asia 2964-2965, 2993, 3020, 3034, 3091, 3103, 3107, 3246 Atlanta Zoo and Botanical Garden 3277 Atlantic salmon 3039, 3227 Atelopw Initiative 3277 atrazine 3080-3082, 3 170, 3 174, 3 194, 3199, 3202, 3216, 3218, 3222-3224, 3230-3231 atrazine desethyl 3216 atrazine desisopropyl 3216 ATV (Ambystoma tigrinum virus) 2965-2966, 2968-2973, 2975, 3230 Am'-SRX' 297 1 Australasia 2988-2990, 3034 Australia 2964-2965, 2967, 2970, 2979, 2987, 2988, 2994, 3007, 30 16-30 18, 3020, 3031-3032, 3034, 3036, 3041, 3057, 3059, 3061, 3091, 3107, 3115, 3135, 3175, 3191, 3197, 3259, 32703271, 3273-3274 avian malaria 3034 avian viruses 2977 axolotl 2965, 3019 B

bacteria 2964, 2966-2967, 2978-2980, 3008, 3010, 3025-3026, 3030, 305 1, 3060, 3075, 3151, 3154-3156, 3168, 3171, 3214, 3230, 3250 barramundi 297 1 behavioural fever 3056 benzanthracene 3 133 benzopyrene 3 133 Berlin 2993 Bermuda 3081 Big Tableland 2989, 3015, 3017-3018 bioindicators 3 175, 3260-326 1 biological control 2964 birdpox 3034 birds 3019, 3056, 3068-3071, 3187, 3271-3272 bisphenol 3134, 3215, 3223-3224 BIV (Bohle iridovirus) 2965-2967, 296% 2971 blue baby syndrome 3155 bluegill sunfish 3 193 bobbing 3 173

Bogong Creek 3017 Bohle iridovirus (BIV) 2964-2965, 29652968 boreal toads 3134 brachydactyly 3094 brachygnathia 3094 brachymelia, brachymely 3 102-3 103 Brazil 2970, 2977, 2979, 2991, 3270 brevinin- 1 30 18 brevinin-230 18 British Columbia 2992 brook trout 3039 buffering 3245, 3261 Bufo bufo United l n g d o m virus (BUK) 2971 BUK (Bufo bufo United Kingdom virus) 2971 bullfrog 2971, 2977, 2979, 2991, 3003, 3016, 3020, 3060, 3097, Burmese star tortoise 2971 Bush meat 3240 butterflies 3271-3272 butylhydroxyanisol 3223 C Caddis fly 3252 caerin 3018 calcivirus 2963, 2978 California 2978, 2992, 3014, 3097, 3099, 3104, 3118, 3151, 3197-3199, 3229, 3257 California newt 3107, 3 118 California red-legged frog 2967, 327 1 California treefrog 3 118 cancer 3215-3216 Canada 2971-2972, 2977, 2992, 3042, 3104, 3117, 3126, 3152, 3170, 3213, 3224 cane toad 2964-2965, 3042, 3272 cannibalism 2973, 3079 Cape York capsid 2964, 2966 carbamate 3 198, 320 1 CAR (constitutive adrostane receptor) 3215 carbaryl 3134, 3191, 3193-3196, 3226 carcinoma 3216, 3239 Caribbean 3270, 3274 carotenoids 3 129 cascade frogs 3 118-3 119, 3 198 Central America 2987, 2990-2991, 3191, 3197, 3259, 3275 central nervous system (CNS) 3225 Central Valley 3197, 3229 cercaria(e) 3069-3070, 3072, 3074, 3076-3081, 3093, 3105 channel catfish 3155-3156, 3161, 3163 ChE (cholinesterase) 3229 chemotaxis 2997, 3019 chestnut blight 3034 chickens 3221 Chile 2994 China 2970, 2977 chinook salmon 3174

chlamydiosis 2979 chlorpyrifos 3 197-3198 chlorothalonil 3 197 cholesterol 3216 cholinesterase (ChE) 3 198-3 199, 3229 chorus frog 3190 chromomycosis 3043 chytrid fungi 2976-2978, 2987-3034, 3051, 3058, 3060, 3273-3274 chytridialeans 3027 chytridiomycosis 2978-2979, 29873034, 3041, 3051, 3054, 3056-3061; 3274-3275 circannual rhythms 3084 citropin 3018 climatic change 3079, 3083, 3117-31 18, 3268-3277 CNS (central nervous system) 3225 co-evolution 2965, 3073-3074 colonial sporangia 3001 Colorado 2978, 2992, 3150-3151, 3258, 3271 common frog 2965, 2970-2971, 3038, 3118, 3227 common toad 2970-297 1, 3 118 competition 3176, 3191, 3193, 3231, 3253, 3261, 3273 Conservation Breeding Specialist Group 3277 Conservation International 3277 constitutive adrostane receptor (CAR) 3215 contraceptives 32 13 Cooktown 2989, 3017 Coromandel 2990 corticosterone 32 16, 323 1 cortisol 32 16 Costa Rica 2977-2978, 2991, 3150, 3270-3271, 3273, 3275 CPD (cyclobutane pyrimidine dimer) 3128 CPD-photolase 3128 crayfish 3034-3035, 3155, 3195, 3241 crayfish plague 3034 crested newts 3225 cricket frogs 3075, 3219 Croatia 2968 Croatian ranavirus 2968 Crocodilians 3 156 Crow Wing County 3096-3097 crustaceans 2987, 3155, 3212, 3246 cryoarchiving 3052 curling defect 3247, 3259 CWB site 3097. 3099. 3104. 3109 cyclobutane py&nidine d i k e r (CPD) 3128 cypermethrin 3227 cytochrome P450 enzymes 32 15-32 16 cytosine methyltransferase 32 17 Czech Republic 3259 D DAFI'F (Declining Amphibian Populations Task Force) 3269 DDAC (didecyl dimethyl ammonium chloride) 3030 DDE 3213 DDT 3187, 3197, 3197-3198, 3221, 3225. 3227. 3229 Declinkg ~ m i h i b i a nPopulations Task Force (DAPTF) 3269 Department of Environment and Heritage (Australia) 3032

deforestation 31 16, 3240, 3242 deformities [also see abnormalities; malformations] 3067-3084, 30893109, 31 19, 3164-3165, 3188, 3192, 3194, 3199 dehydroepiandrostemne 32 16 dermatitis 3 135 DES (diethylstilbestrol) 3221, 3229 Des Moines River 3I48 diatoms 3035 diazinon 3 197didecyl dimethyl ammonium chloride(DDAC) 3030 dibutyl phthalate 3224 dieldrin 3 198 diethylstilbestrol (DES) 3221, 3225 digenetic trematodes 3067-3084 dioxins 3177, 3218 dissolved organic compounds (DOC) 3116-3117,3126-3127, 3246 dissolved organic matter (DOM) 3 117 DNA 2964-2965, 2977-2978, 3020, 3028, 3030, 3035, 3038, 3041, 3059, 3128-3129, 3132, 3135, 3137, 3146, 3156, 3209, 321 1-3212, 3214, 3216-3217 DOC (dissolved organic compounds) 3116-3118, 3126-3127, 3246, 3251 DOM (dissolved organic matter) 31 17 dragonfly 3249, 325 1-3252 Dutch elm disease 3034 dwarf African clawed frogs 2992 dwarfism 322 1 dysgenesis 3223-3224 dysplasia 322 1 E eastern spotted newt 2969 eastern newt 2971 echinostomes 3084 ectrodactyly 3094, 3103 ectromelia 3094, 3098, 3100, 31023103 Ecuador 2990-2991, 3270, 3272 EDC (endocrine disrupting chemica1,endocrine disruptor) 32083232 EE, (17-alpha-ethinylestradiol) 3224 eels 3102 EHNV (a fish ranavirus) 2966, 2975 ELISA (Enzyme-Linked ImmunoSorbent Assay) 2974, 3020 elevational shifts 3271-3273 El Nifio 3 117, 3268, 3276 El Nifio Southern Oscillation Events (ENSO) 3275 embryonic amplification 3077 endocrine disrupting chemicals;,endocrine disruptors (EDC) 3060, 3134, 3189-3190, 3202, 3208-3232 endometriosis 322 1 endosulfan 3 197, 3227 England 3271 EN1 (Ente Nazionale IdrocarburiI) 3240 ENS0 (El Nifio Southern Oscillation Events) 3275-3276 Ente Nazionale Idrocarburi (ENI) 3240 Environmental Protection Agency (EPA) 3146, 3202 Enzyme-Linked ImmunoSorbent Assay (ELISA) 2974

enzymes 3128-3129, 3 135 EPA (Environmental Protection Agency) 3146, 3153-3154, 3157, 3210, 3223-3224 epidemiology 2969-2973, 30 14-30 19, 3032, 3034, 3037-3039, 3200, 3269, 3277 epigenetics 3216-3217, 3221, 3230 ER (estrogen receptor) 3221 ER alpha receptor 32 15 ER beta receptor 32 15 emdentin-1 3018 esculentin-2 30 18 esfenvalerate 3080, 3 194, 3 199 esnadiol 3213, 3215, 3223, 3225 estrogen 321 1, 3213-3215, 3217, 3220-3222, 3224-3223, 3230-3231 estrogen rrceptor (ER) 3221 estrone 3213, 3216 Eungella National Park 2990, 3016, 3019 Europe 2964, 2993, 3034, 3041, 3043, 3091, 3103, 3107-3108, 3119, 3135, 3149-3150, 3186, 3191, 3246, 3257-3259, 3272 European frog 3102, 3225 European Commission 3209-32 10 European Union 3 157 eutrophication 3150-3151 extinction 3017, 3190, 3228, 3269 F facilitation 3084 fathead minnows 3161, 3193, 3213, 3216 Federal Republic of Nigeria 3240 Federal Research Center for Forestry and Forest Products 3149 feminization 3214, 3216, 3223-3224, 3230 fenarimol 3216, 3228 fenthion 3228 fertilizer 3 103, 3 147-3149, 3159, 3170, 3172, 3174-3176, 3251 FETAX test 3164 FEV (Frog Erythrocytic Virus) 2963, 2977 Finland 31 18, 3259 fish 2964, 2966, 2971, 2977, 2979, 2987-2988, 3035, 3038-3041, 3056, 3058, 3068-3069, 3071, 3074, 3079, 3105, 3107, 3126, 3128, 3130-3131, 3153-3157,3160-3161,3171,3188, 3193, 3212, 3216, 3219, 3222, 3229, 3246, 3249-3250, 3252, 3271, 3273 flatworm 3074 flavobacteria 3230 flavonoids 3 129 Florida 3225, 3228 fluoranthese 3 133-3 134 foam nests 3130-3131 Fortuna 2991 France 3272 Frog Embryo Teratogenesis Assay 3 192 Frog Erythrocytic Virus (FEV) 2963, 2977 Frog Virus 3 (FV3) 29642965 FSH 3210 fungi 3 128 fungicides 3103, 3202, 3214, 32163217, 3228, 3251 FV3 (Frog Virus 3) 2965-2973

AMPHIBIAN BIOLOGY G gastric brooding frog 2988-2989 gastropod 3068, 3077 Germany 2993, 3019, 3041, 3103, 3258-3259 Ghana 2994 giant toad 3036, 3225 glaucous gulls 3212-3213 global warming 3246, 3269, 3271, 3275 glucocorticoids 323 1 glutathione 3 131 glyphosate 3202 GnRH 3210 golden algae 3035 golden bell frog 3 175 golden toad 2991, 3270 goldfish 3043 gonadotropin 3225 grasshoppers 3055 gray treefrogs 3078-3079 Great Britain 3 117, 3258 Great Dividing Range 2989 Great Lakes 3151, 3196 green frog 2971-2972, 2975, 3043, 3076-3077, 3152, 3157-3158, 3213, 3224 green treefrog 3061 ground beetles 3242 Guatemala 3272 guinea pigs 304 1 WPPY 3 163

H haemoglobin 3155-3156, 3161, 3173, 3175 haemorrhagic wndrome 2967 handicappgd kghypothesis 307 l3073 harlequin frog 3270, 3275 Hawaii 3034 heat shock protein (HSP) 3132 heavy metals 3103, 3239-3242 helminthiasis 3 107 helminths 3074-3075, 3080, 3083 hemimelia. hemimely 3094, 3 100, 3 102 hemoxygenase 3 129 hepatitis 2979 herbicides 3103, 3189, 3191, 3195, 3202, 3218-3219, 3229, 3251 herbs 3271 herpesvirus 2963, 3034 heron 3071 Homeobox genes (Hox genes) 32203222 homeostasis 3209, 3214-3215, 3232 Hox genes (Homeobox genes) 32203222 H I T (hypothalamic-pituitary-thyroid axis) 3218 HSP (beat shock protein) 3132 HSP-70 3132 hydrocarbons 3239 hydrogen sulphide 3242 hygroma 3097, 3100-3101 hyperextension 3099, 3 102-3 103 hypothalamic-pituitary-thyroid axis (HF'T) 3218, 3231 hypothalamus 3210, 3225 hypothyroidism 32 18 I Iberian Peninsula 3272 ichthyophonosis 3042 icosahedral DNA viruses 2964

Illinois 3075 immunoelectron microscopy 2974 immunofluorescence 2974 immunohistochemistry 3021, 30263028 immunoperoxidase 2968, 3022, 30263028 immunosuppression 301 1, 3079, 3081, 3083-3084, 3230 inbreeding depression 3 190 India 3246 Indiana 3097, 3100, 3148, 3152 Indiana University 2965 Indonesia 2993, 3246 inhibin 3210, 3222 insecticides 3103, 3190-3191, 31943195, 3202, 3218, 3225, 3227-3229 insects 2966, 3055-3056, 3068, 3155, 3191-3192, 3195, 3246, 3252 Intergovernmental Panel on Climate Change (IPCC) 3268, 3276 intermediate host 3068-3071, 3074, 3077, 3080, 3109 International Program on Chemical Safety (IPSC) 3209 International Union for Conservation of Nature (IUCN) 3277 Intersexuality 3214, 3216, 3223-3224, 3230 invertebrates 3128, 3130, 3153, 3157. 3191, 31943196, 3271 IPCC (Intergovernmental Panel on Climate Change) 3268, 3276 IPSC (International Program on Chemical Safety) 3209 iridoiviruses 2963, 2978, 3135, 3273 Italian agile frog 2966, 2973 Italy 2993, 3272 IUCN (International Union for Conselvation of Nature) 3277

J

James Cook University 3017 Japan 3097 JOF site 3097 Johannesburg 3268 K kelp 3035 Kenya 2993 Kihansi Gorge 2993 Kihansi River 2993 l h a n s i spray toad 2993 Klamath Basin National Wildlife Refuge 3151

L La Tablas 2991 lathyrogens 3 102 Latin America 2990-2991, 3016, 3032 L-cycteine 3 131 L-tryptophan 3131 leeches 2977, 3107 lead 3241 least bittern 3108 Leghorn rooster 32 17 leopard frogs 2965, 2978, 2980, 3081, 3157, 3175, 3198, 3218, 3224 Le Sueur County 3096-3097, 3104 Liming 325 1 lipophilic hormones 3209, 32 11 lizards 3041, 3271 LMS site 3097

locomotory performance 3 169 locusts 3055 long-toed salamanders 3079, 31 1831 19 lordoscoliosis 3241 LTHV (Luck6 tumor herpesvirus) 2977 Luck6 tumor, Luck6 tumor herpesvirus (LTHV) 2963-2964, 2966, 2977-2978 Luquillo Mountain Range 3270 M MAbs (monoclonal antibodies) 3027 Maine 2972 malathion 3080-3081, 3194, 3197-3199 Malaysia 3043 malformations [also see abnormalities; deformations] 3067-3084, 3089-3109, 3119, 3164-3165, 3221-3222 mammals 2966, 2971, 2977, 2987, 3056, 3068, 3074, 3 128-3129, 3 156, 3218, 3222, 3227-3228, 3271 Manitoba 2972 marbled newts 3125 marbled salamander 3 190 Massachusetts 3260 MCP 2964-2965, 2974 Mediterranean 3 127 Meeker County 3096-3097, 3104 melanin 3 129-5 130 Melbourne 3020 meningitis 3230 MCrida 2991 mesocercariae 3079 mesocosms 3 192, 3194-3196, 3200, 3253 metacercaria(e) 3070, 3072, 3074, 3079, 3096, 3099, 3 103-3105 metamorphosis 3 166-3 167, 3 169-3 170, 3174, 3188-3191, 3195, 3197, 3202, 3218-3220, 3222-3223, 3241, 3253, 3255, 3260 methaehemoglobin 3155-3156, 3161 methaemoglobinemia 3 155-3 156, 3 161, 3171-3172 methohemoglutanemia 3 154 methoprene 3 134 methoxyclor 3217, 3221, 3227-3228, 3230, 3231 methyltransferase 2965 Mexico 2990-2991, 3272 MHC 2979 mice 3027, 3041, 3220-3221, 3227 microcephaly 3094 micromelia, micromely 3094, 3096, 3098 microphthalmia 3094 midges 2977 Midwest 3104, 3229 migration 3253-3254 mink frog 3 100, 3224 Minnesota 3075, 3081, 3083, 30893090, 3093, 3097, 3099-3100, 3 1023104, 3109, 3193, 3221 Minnesota Pollution Control Agency 3096-3097, 3101 minnows 3 102 miracidium (a) 3069, 3077, 3080 Mission Viejo 3 104 Mississippi 3148 mole salamander 3037, 3190 mollusc 3068-3069, 3 155 monoclonal antibodies (MAbs) 3027 monogenean 3074

Monteverde 2991, 3150, 3271, 3273, 3275 Moor frog 3 118 mosquitoes 2977, 3228 mosquitofish 3227-3228 mountain yellow-legged h g 3197, 3229 mRNA 2966, 3219 mucormycosis 3026, 3041-3042 Miillerian inhibiting hormne 3222 mycobacteriosis 2979-2980 mycosporine-glycine 3 130 myositis 3042-3043, 3 135 N NADH-methaemoglobin reductase 3156 natterjack toad 3038, 31 18 natural selection 3 135-3 136 necrophagy 2973 neotenic, neoteny 2968 Neotropics 3067 neuroendocrine disrupters 3225 New Brunswick 3260 New England 3090, 3104 New Mexico 2992 New South Wales 2988-2989, 3015, 3041, 3274 New South Wales National Parks and Wildlife Service 303 1 New Zealand 2990. 3032, 3091, 3107 NEY site 3097, 3102 Niger Delta 3240 Nigeria 3240 nitrate 3155-3157, 3162-3163, 3165, 3168-3177 nitrite 3155-3157, 3160-3161, 31663169, 3171, 3173-3177 nitrogen 3 145-3 177 nitrogen oxides 3245 nitrosamines 3 156 NLA site 3097 N-nitroso compounds 3 156 nocturnality 3123 nonylphenol 3223, 3225 North America 2964-2965, 29692970, 2991-2992, 2994, 3020, 3032, 3034, 3076, 3091, 3104, 3107-3108, 3128, 3150-3151, 3157, 3175, 3202, 3244-3246, 3254, 3257-3259, 3271-3272 North Atlantic Ocean Basin 3 146 North Carolina 2969, 2971-2972 North Cascade Mountains 3275 Northern Cape 2993 Northern Hemisphere 31 15-31 16, 3246 Northern Territory 3041 northern leopard frog 2977, 3089, 3096-3097-3098, 3100-3101, 3270 northern red-legged frog 297 1 Northern Territory 2989 North Pole 3212 North Sea 3 146 North Temperate one 3 116 northwestern salamander 297 1, 3 118, 3129 Norway 3212, 3259 nuclear receptors 32143215, T3 (3,5,3'-triiodothyronine) 3218 T4 (3,5,3,',5'-tetraiodothyronine 3218 0

octylphenol 3134, 3223 Ohio 3148

OIE (World Organization for Animal Health) 2963, 2975, 3032 oil spills 3239-3240 Oklahoma 3036 Olympic National Park 3126 Ontario 2971-2972, 2977, 3152, 3 169, 3188, 3198, 3213, 3224 oomycete 3035, 3117, 3135, 3273, 3275 oomycosis 3035-3043 Oregon 3038, 3119, 3127, 3275 organochlorine 3201, 32 12 organophosphates 3 198, 320 1, 32283229 ornate burrowing frogs 2965, 2971 ornithine cycle 3 153 osmoregulation 3 153-3 154 OSP Site 3096 Ottertail County 3096 ovoviviparity 3 123 ozone 3103, 3116, 3149 P PAbs (polyclonal antibodies) 3027 Pacific Northwest 3040, 31 15, 31 18, 3176, 3273, 3275-3276 Pacific treefrog 3073, 3075, 3100, 3107, 31 18, 3197-3198, 3229 PAH (polycyclic aromatic hydrocarbon) 3133. 3177 Palmate newt 3169 palustrin-3 3018 Panama 2990-2991, 2994, 3014, 3271 parasite(s) 2965, 3058, 3067-3084, 3103-3107, 3109, 3196 paratenic hosts 3077 PARC (Amphibian Ark, Partners in Amphibian and Reptile Conservation) 3277 PBDEs (polybrominated biphenyl ethers) 3214, 3218 PCBs (polychlorinated biphenyls) 3177, 3186, 3212, 3218, 3223 PCR (Polymerase chain reaction) 2965, 2968, 2974, 2979, 3020-3022, 3028-3031 Pefialara Natural Park 2993 Pennsylvania 3 149 Pennsylvania State Game Lands 3152 peppered moth 3230 perchlorate 3219-3220 permethrin 3 193 personal care products 32 18 Perth 2989-2990, 3019 "pesticide puzzle" 3 199-3200 pesticides 3080-3081, 3091, 3107, 3134, 3170, 3174, 3177, 3186-3203, 3192-3203, 3212-3213, 3223, 3228-3231, 3251 petrochemicals 3239-3242 pet trade 3019 phenol 3242 phenology 3271 phocomelia, phocomely 3096, 3098 photolase 3128, 3135 phthalates 3218, 3224 phytoestrogens 32 13 phytoplankton 3 126-3127 pigmentation 3101 pilchards 3034 pituitary 3225 plains leopard frog 3 119 plankton 3 130

plants 3128, 3272 platypus 3042 pneumonia 2979 Poland 3 171 polar bears 32 12-2 13 pollutants, pollution 3 107, 3109, 3 116. 3145-3177, 3186-3203, 3208-3131. 3239-3242 polybrominated biphenyl ethers (PBDEs) 3214, 3218 polychlorinated biphenyls (PCBs) 3212. 3218 polyc~oha~ antibodies (PAbs) 3027 polycyclic aromatic hydrocarbon (PAW 3133 polydactyly 3094, 3 103 polyembryonic amplification 3069 polyextension 3096, 3098-3099 polymelia, polymely 3093-3094, 3099-3100, 3102-3104, 3107 Polymerase Chain Reaction (PCR) 2974-2976, 3020, 3022, 3028-3030 porpyra-334 3130 post-metamorphic death syndrome 2992 pregnane X receptor (PXR) 32 15 pregnenolone 32 16 primary host 3068, 3070-3071 prochloraz 32 16 progesterone 32 14, 32 16, 3222, 323 1 prolactin 3225 propazine 32 16 proteosomes 32 1 4 3 2 15 Puerto Rico 2992, 3269-3273 PXR (pregnane X receptor) 3215 Pyrenees Mountains 2978 Pyrethoids 3229 pyrimidine-[6-4'1-pyrimidone photoproducts ([6-41 photoproducts) 3128

Q

quail 3228 ~ u a r a n t i n e3031, 3034 Quebec 2992, 3042, 3104, 3108 Queensland 2965, 2967, 2970, 29882990, 3014-3015, 3017-3020, 3036, 3042, 3058-3059, 3274 R rabbits 3027 RaHV-1 (Rana herpesvirus 1) 2977 rainbow trout 3039, 3155-3156, 3161, 3193 Rana catesbeiana virus Z (RCV-Z) 2964-2965, 2968 Rana herpesvirus 1 (RaHV-1) 2977 ranalexin 30 18 ranatuerin-1 3018 ranatuerin-2 3018 ranaviruses 2963-2979 RAR (retinoic acid receptor) 3222 rats 3041, 3217, 3221, 3228 RCV-Z (Rana catesbeiana virus Z) 2965, 2968, 2972 red-backed salamander 3 169-3 170, 3273 red-eared lider turtle 3225 red-eyed treefrog 3040-3041 redia(e) 3069 red leg 2966, 2978-2979, 3020 red-legged frog 3 118-3 119 red-spotted newts 3075, 3226-3227

AMPHIBIAN BIOLOGY reductase 3215-3216 reed frog 3166,3225 renal adenocarcinoma 2977 reptiles 2966,2971,3056,3271-3272 resting spore 3001 Restriction Fragment Length Polymorphism (RFLP) 2964 retinonic acid 3103-3104,3214,3221-

3222 retinoic acid receptor (RAR) 3222 retinoids 3073,3103-3104,3109,3209,

3211-3212,3221-3222,3231-3232 retroviruses 2963,3223 RFLP (Restriction Fragment Length Polymorphism) 2964 Rhode Island 2978 rinderpest 3034 ringed seals 3212 RNA 3041 RNA polymerase 3212 Rocky Mountains 2992,3150,3258 ROI site 3096 Rome 3152 roughskinned newts 3119,3129,

3169-3170 Royal Melbourne Zoo 2989 RUK (ranavirus from the UK) 2965,

2969,2971

Spain 2978,2987,2993,3118 spermatogenesis 3222,3232 splenitis 2979 sporangia (see zoosporangia) sporocyst 3069 spotted frog 311 8 spotted salamander 2969,3260 spotted treefrog 3017 spring peeper 3157,3213 State Game Lands 176 3149 steroid and xenobiotic receptor (SXR)

3215 steroidogenesis 3222 steroids, steroid hormones 3209-3212,

3214-3216,3222,3227,3231 steroid dehydrogenase 3215-3216 streptococcosis 2979 sucker 3249 sudden oak death 3034 sulphur dioxide 3245 sunscreen 3129-3130 SUN site 3096 Svalbard 3212 Sweden 3170,3258-3259 Switzerland 2979,2993 Switzerland County 3097,3100 SXR (steroid and xenobiotic receptor)

3215 syndactyly 3094 synergism 3132-3135,3172-3175,

S

Saint Lawrence River 3239 salmon 3228 sarcoptic mange 3034 Saskatchewan 2965,2970-2972,3104 Save a Frog programme 3277 Scandinavia 3170,3244 scoliosis 3096-3097,3103 Scotland 3259 sea bass 3156 septicaemia 2964,2966,2968,2978-

2979,3020 Sequoia Kings Canyon National Park

3197 serotonin 3129 sex determination 3222 sex ratio 3222-3224 sex reversal 3222 sharp-ribbed salamander 3171 sharp-snouted dayfrog 3017 sheep 3027 shinorine 3130 shrubs 3271 Sierra Nevada 2992,3014,3197-3198,

3257 simazine 3216 smog 3149 smooth newt 3168 snails 3069,3071,3075-3078,3080-

3081,3083-3084,3104,3109 sodium perchlorate 3220 sodium pyruvate 3131 South Africa 2977,2993,3020,3223,

3254 South America 2965,2987,2990,

3020,3107,3246,3270,3273,3275 southern day frog 2989 Southern Hemisphere 3116 southern gastric brooding frog 2989 southern leopard frog 2971, 3134,

3193-3194 southern torrent salamanders 3169-

3170 spadefoot toad 3038

3192,3196,3251,3269,3273-3275 T Tadpole Edema Virus (EV) 2964-2966 Tanznia 2993 Tasmania 2989,3042 taumelia, taumely 3094,3096-3100 Tennessee 2971 temporin 3018 TEI' padpole Edema V i s ) 2965,2967, 297l testosterone 3213,3215-3216,3222-

3224,3228 Texas 3219 Thailand 2970,2979 The Netherlands 3037,3040,3103 thermoregulation 3055-3056,3125 Threat Abatement Plan 3031 three-spine stickleback 2971 thymus 3230 thyroid 3218-3220,3225 thyroid hormones 3209,3211-3212, 3214,3218-3220 thyroid stimulating hormone (TSH)

3219 thyroxine 3220 tiger salamander 2965,2968-2970,

2973,3230,3241 Townsville 3019 toxaphene 3196-3197 transcription factors 3217 TR beta 3219 treefrogs 3189,3193 trees 3271-3272 trematodes 3067-3084,3103-3105, 3109,3199 trematodiasis 3075 Trempealeau County 3096 triazine herbicides 3216 triphenyltin 3251 tropisms 2964 trout 3039,3156 TSH (thyroid stimulating hormone)

3219

tuna 3034 turtles 3156,3225,3240 T, (3,5,3'-triiodothyronine) 3218-3219 T, (3,5,3,',5'-tetraiodothyronine3218

u UK (United Kingdom) 2965 UK ranavirus 2964-2967 Ulcerative syndrome 2967 ultraviolet (UV) 2966,3030,3102,

3112-3137,3172-3173,3189, 3193-3194,3196,3250-3251 United Kingdom (UK) 2964-2965, 2969-2972,2994 United States (USA) 2964-2966, 2971-2972,2977,2979,2987,2990, 2992-2994,3020,3035-3036,3038, 3040-3042,3075,3082,3084,3090, 3101-3103,3107,3115,3117,31263127,3146,3151,3153,3157,3174, 3176,3186,3191,3196-3198,32013202,3218,3224,3229,3270,3273, 3275 United States Geological Survey (USGS) 3146 urbanization 3082 urea 3152-3154,3163-3164,3168-

3170,3176-3177 Uruguay 2970,2979,2991,3016 USA (United States) 2968-2969,2972,

2978,3118 USGS (United States Geological Survey) 3146 USGS-BRD National Wildlife Health Center 3105 UV (ultraviolet) 2966,2976,3035,

3038-3039,3060,3102-3103,31123137,3170,3231,3246,3250-3251, 3261,3275-3276

v vaccines 2975 Venezuela 2966,2969,2990-2991,

3016 Venezuelan ranavirus 2969 Vermont 3042,3083,3104-3105,3222 Victoria 2988-2989,3017,3019 vinclozilin 3217,3228 viral diseases, viruses 2963-2980 Vison 3251 vitamin A 3103,3221 vitellogenesis 3223 vitellogenin 3217,3225 viviparity 3123-3124 W

warty newts 3227 Washington (state) 3126 Water boatmen 3252 water mold 3175 Western Australia 2989,3014,3019,

3041,3197 Western Cape 2993 western toad 3038-3039, 3107,3117-

3119 West Nile virus 2963 West Virginia 2966,3042 WHO (World Health Organization)

3209-3210 Wildlife Diseases List 3032 WIN Site 3096 Wisconsin 3075 Wisconsin Central Sand Plain 3151

wombats 3034 wood frogs 3128, 3190, 3199 World Health Organization (WHO) 3209 World Organization for Animal Health (OIE) 2963, 3032 World Summit on Sustainable Development 3268 World War I1 3 147 Wyoming 2992 Wyoming toad 3228 X xenobiotics 3103, 3109, 3215, 3225 XEMA (Xenopus Metamorphosis Assay) 3219 Xenopus Metamorphosis Assay (XEMA) 3219

Y Yosemite toad 2979, 3199

zooplankton 3 191, 3 194-3195, 3271 zoosporangium 2995-3001, 3003, 3005, 3007-3013, 3018-3019, 3023, 3025-3027, 3036 zoospore(s) 2994-3003, 3005-3006, 3012, 3014, 3016, 3019-3020, 3023-3025, 3027, 3031, 3035-3036, 3039, 3051-3052, 3057-3060, 3273, 3275 zygomycete 3041 zygospores 3042 INITIAL NUMERALS 2-chloro-N-(ethoxymethy1)-N-(2-ethyl6-methylphenyl) acetamide 32 19 3 beta-dehydrogenase 32 16 3 beta-hydrogenase 32 16 3,5,3'-triiodothyronine (T,) 3218 3,5,3,',5'-tetraiodothyronine(T,) 3218 4-nonylphenol 3225 5-alpha dehydrotestosterone 3216 5-alpha reductase 3216 [6-41 photoproducts (pyrimidine-[6-4'1pyrimidone photoproducts) 3128

11-beta hydroxylase 32 16 11-deoxycorticosterone 32 16 11-deoxycortisol 32 16 17-alpha-ethinylestradiol (EE,) 3213, 3224 17-alpha hydroxylase 32 16 17-alpha hydroxypregnenolone 32 16 17-alpha hydroxyprogesterone 3216 17-beta dehydrogenase-1 3216 17-beta dehydrogenase-2 3216 17-beta estradiol 3213, 3215-3217, 3221, 3224 17-beta reductrase 32 16 17-desmolase 32 16 20, 22-desmolase 3216 20-alpha, 22 alpha-hydroxycholesterol 3216 20-alpha hydroxylase 2 1 hydroxylase 32 16 22 hydroxylase 32 16

INDEX TO SCIENTIFIC NAMES Entries in boldface indicate that the indexed item, or part thereof, is illustrated in a figure as either a drawing or photograph. A Achlya 3035-3036 Acris 3101, 3122 Acris crepitans 32 19 Acris gryllw 3248, 3252 Aeromoinas hydrophila 2967, 2978-2979 Aereommas hydrophila ranae 2979 Afrana angolensis 2993 Afrana fucigola 2993 Agalychnis callzdryas 3040-3041 Aglyptodactylus 3 123 Alaria 3079 Allomyces macrogynus 3029 AUophrynidae 3 121 Alytes 3 123 Alytes obstetricans 2978, 2993 Ambystoma 3040, 3 101, 3 121 Ambystoma gracile 2971, 31 13, 31 18, 3127, 3129, 3131, 3152, 3158-3163, 3 168-3 169 Ambystoma jeffersonianum 3 152, 3 162, 3248, 3250, 3252-3253, 3258, 3260 Ambystoma laterale 31 13, 3249, 3258 Ambystoma macroductylum 3040, 3079, 3113, 3115, 3119, 3125, 3129, 3162, 3164, 3170, 3172, 3227 Ambystoma maculatum 2969, 297 1, 3076, 3113, 3118, 3129, 3131, 3152, 3162, 3168, 3249-3250, 3252-3254, 3258, 3260

Ambystoma mexicanum 2965, 2973, 3 131 Ambystoma opacum 3037, 3 190 Ambystoma talpozdeum 3 190 Ambystroma texanum 3 160-3 161, 3 174, 3249 Ambystomatidae 2971, 3015, 3121, 3129, 3131, 3248-3249, 3256, 3258 Ambystoma tigrinurn 2964, 2968-2973, 3003, 3005-3006, 3016, 3160-3161, 3230, 3249, 3254, 3258 Ambystoma tipinum nebulasum 2970, 2972. 2974 Ambystoma t i p n u m stebbinsi 2965, 2971, 3016 Ambystoma tigrinurn t i p n u m 3037 Amnirana 3 122 Amphibiocystzdium 3043 Amphiuma 3 102 Amphiumidae 30 15, 3 121 Aneides 3 102 Anguilla 3 102 Anura 3121, 3272 Aphanomyces 3035-3036 Aphanomyces astaci 3034-3035 Aphanomyces inuadans 3035 Arthroleptidae 3 122 Ascaphidae 3121, 3125, 3176 Ascaphus 3 101 Ascomycota 3040, 3043 Assa 3 124

Atelognathus patagonicus 2965 Atelopus 2991, 3122, 3275 Atelopus cruciger 2991 Atelopus uarius 2991, 3270 B

Basidiobolus ranarum 2992, 3014, 3031, 3135 Batrachochytrium 2994, 3004 Batrachochytrium dendrabatidis 29762978, 2987-3034, 3051-3052, 3054-3061, 3135, 3273-3275 Batrachoseps 3 102, 3 154 Batrachuperus 3 121 Biston betularia 3230 Blastocladiales 2994, 3029 Bombina 3 121 Bombina pachypus 2993 Bombinatoridae 3015, 3 121, 3248 Bombina uariegata 3248 Boophis 3 123 Brachycephalidae 3 122 Bufo 3038, 3101, 3122 Bufo americanus 2971, 3076, 30783079, 3105, 3113, 3130, 3157-3158, 3160, 3162, 3165, 3169-3170, 3248, 3250, 3254, 3257 Bufo arenarum 3241 Buqo baxteri 2992, 3014, 3031

AMPHIBIAN BIOLOGY Bufo bmeas 2992, 3019, 3035, 30383041, 3059, 3106-3107, 31 13, 3117-3119, 3129,3135,3160-3162, 3164, 3167-3168, 3248, 3275-3276 Bufo boreas boreas 2978, 3058 Bufo bufo 2970-2972, 2972, 3041, 3113, 3118, 3153, 3160, 3162, 3164-3169, 3171, 3248, 3257, 3259 Bufo calamita 3038, 31 18, 3160, 3165, 3172, 3248, 3257 Bufo canorus 2979, 2992, 3199, 32483249, 3257-3258 Bufo fowlerii 3012, 3249 Bufo granulosw 2980 Bufo hemiophrys baxteri 3228 Bufo marinus 2964, 2969, 2971, 2978, 2980, 2995, 3004, 3012, 3016, 3036, 3041-3042, 3081, 3218, 3225, 32723273 Bufo melanostrictus 3043 Bufonidae 3015, 3108, 3122, 3125, 3127, 3202, 3248, 3256-3257, 3275 Bufo periglenes 2991, 3269-3270 Bufo punctatus 3248 Bufo terrestris 3162, 3257 Bufo ualliceps 3250 Bufo uiridis 3 152 Bufo woodhowii 3129, 3248, 3250 Bufo woodhousii fowleri 297 1 I.

Caeciliidae 3 121 Campostoma anomalum 32 19 Catostomzcs commersoni 3249 Caudata 3121, 3272 Centrolenidae 2991, 3015, 3122 CeratophTys 3 122 Chironomidae 3 125-3 126 Chlamydophila pneumonie 2979 Chytridiales 29942995, 3027, 3029 Chytridiomycetes 2994, 3027 Chytridiomycota 2994, 3001 Cladosporium 3043 Chryphonectria parasitica 3034 Crinia 3122 Crinia sig-n$era 31 13, 3160, 3167, 3175 Chryptobranchidae 3 121 Cryptobranchus 3 102 Cyclorana 3 122 Cyclorana breviceps 297 1 D Daphnia 3168, 3246, 3249 Dendrobates 3 102 Dendrobates auratus 3005 Dendrobates azurew 3005 Dendrobates tinctorius 3004-3005, 3012, 303 1 Dendrobatidae 30 15, 3 122 Dermocystzdium 3042-3043 Dermomycoides 3043 Desmognathus carolinensis 3272 Desmognathus ocoee 3272 Dicamptodon 3 102 Diocamptodontidae 3 121, 3 125, 3 176 Discoglossidae 30 15, 3 121, 3 127 Discoglossw 3 121 Discoglossus galganoi 3 160, 3 164 Digenea 3068 Dothidieales 3040-3041 Dyscophus 3 123 Dyscophus antongilii 2993

E Echinostoma 3077 Echinostomatidae 3 105 Echinostoma triuoluis 3075, 3077-3081 Eleutherodactylus 2991-2992, 3102, 3270, 3272-3273 Eleutherodactylw antillensis 3272 Eleutherodactylus coqui 3269-3270, 3273-3274 Eleutherodactylus enezdae 3270 Eleutherodactylus gryllw 3272 Eleutherodactylw jasperi 3270 Eleutherodactylus karlchmzdti 2992, 3270 Eleutherodactylus unicolor 3272 Ensatina 3 102 Erpobdella octoculata 3 102 Eumycota 3035 F Fejeruarya 3 122 Flauobacterium 2979 Fonsecaea dermatitidis 3043 Fonsecaea pedrosi 3043 G Gambwia affinis 297 1 Gambwia afinis holbrooki 3227 Gasterostew aculeatw 297 1 Gastrophryne 3 101, 3 123 Gastrophryne carolinensis 3248 Gastrotheca 3 124 Geochelon latynota 297 1 Gonapodya 3029 Gymnophiona 3 121 Gyrinophilus 3 102 H Haideotriton 3 102 Halzpegas 3077 Heleoph~ynidae3 122 Hemidactylium 3 102 Hemisotidae 3 122 Herpesviridae 2977 Heterixalus 3 123 Hildebrandtia 3 122 Hoplobatrachus 3 122, 3240 Hydromantes 3 102 Hyla 3101, 3122, 3272 Hyla andersonii 3248-3249, 3254 Hyla arborea 3113, 3129, 3160, 3165, 3248 Hyla cadauerina 31 13, 31 18, 3135 Hyla chrysoscelis 3003, 3012, 3248 Hyla cinerea 3249, 3251, 3257 Hyla cruc$er 3157-3158, 3173, 3248, 3257 Hyla femoralis 3248, 3252 Hyla gratiosa 3248-3249, 3252 Hyla regilla 2971, 3035, 3039, 30713072, 3113, 3118, 3129-3130, 3 1 3 4 3135, 3160, 3162-3164,3169-3170, 3172, 3197, 3229 Hyla squirella 3248 Hyla uersicolor 2971, 3078-3079, 31 13, 3134, 3189, 3193, 3226, 3248 Hylidae 2991, 3015, 3122, 3248, 2563257 Hylmina 3 122 Hymenochirw curtipes 2992, 3020 Hynobiidae 3121, 3125 Hynobius 3 121 Hyperoliidae 3 123 Hyperolius 3 123

Hyperolius argus 3225 Hyperolius marmoratus 3 166-3 167, 3 17 1 Hyperoliw uiridzjlauus ommatosticus 3 153 Hyperolius uirdijlauus taeniatus 3 153 Hypopachw 3 102, 3 123 I Ichthyophidae 3 121 Zchthyophonw 2987, 3042-3043, 3135 Zctalurus punctatw 3 163 Iridoviridae 2964, 2977 Ixobrychus minutus 3108

K Kassina 3 123 L L a r w hyperboreus 32 12 Lathyrus odoratus 3 102 Laucstromyces hiemalis 3002 Leiopelma archeyi 2990 Leiopelmatidae 30 15, 3 12 1 Lepidobatrachus 3 122 Ltpomis affinis 2971 Leptodactylidae 30 15, 3 122 Leptodactylus 2969, 3 102, 3122 Leptodactylus ocellatw 2991 Ltptospira 2964 Limnodynastes ornatw 2965, 2968, 2971 Limnodynastes peronii 3041-3042, 3160, 3167 Limnodynastes tasmaniensis 3003, 3005, 3016 Limnonectes 3 122 Lissotriton boscai 3 167 Litoria 3 122 Litoria adelensis 304 1 Litoria alboguttata 297 1 Litoria aurea 2989, 31 13, 3129, 3 1593160, 3167, 3175 Litoria caerulea 2965, 2971, 3003-3005, 3007-3008, 301 1, 3016-3018, 3023, 3025, 3027, 3041, 3060-3061, 31623163, 3165 Litoria chlmis 3003-3006. 3008, 3014, 3016, 3027, 3031 Litoria dentata 3 113, 3 129 Litoria genimaculata 2990, 3015-30 18 Litoria gracilenta 2999, 3009-30 10 Litoria infiafienata 304 1 Litoria lutopalmata 297 1 Litoria lesueuri 3009, 301 1, 3014, 3016, 3057, 3059 Litoria nannotis 2989-2990, 30173019, 3057, 3059 Litoria peronii 3 113, 3 129, 3 175 Litoria ran$ormis 2990 Litoria rheocola 2989-2990, 3015, 3017-3019 Litoria spenceri 30 17 Litoria terrareginae 297 1 Litoria uerreauxii alpina 3 113 Litoria wilcoxii 3274 Lymphocystiuirw 3 135 Lysapus 3122 M Mantellidae 3015, 3 123 Mantdactylus 3 123 Megophrydae 3122 Microhyla ornata 3163, 3248

Microhylidae 3015, 3123, 3248, 3256 Micropterw treculi 3 157, 3 161 Mixophyes 3014, 3122 Mixophyes fasciolatw 3003-3008, 30123014, 3016, 3019, 3023-3024, 3027 Mixophyes Jleayi 3007 Mixophyes zteratw 2979 Mixophyes scheuilli 30 14 Monoblepharidales 2994, 3029 Mucor 3041 Mucor amphibiorum 2987, 3027, 30413042 Mus musculus 3220 Myobatrachidae 3 122 Mycobacterium 2979 Mycobacterium bouis 2979 Mycobacterium chelonei 2979 Mycobacterium leijlandii 2980 Mycobacterium marinum 2980 Mycobacterium tuberculosis 2979 Mycobacterium ulcerans 2980 Mycobatcerium szulgai 2980 Myobatrachidae 30 15 N Nasikabatrachidae 31 22 Nectophrynoides asperginis 2993 Necturus 3 102 Neobatrachw 3 122 Neocallimastigales 2994 Notaden 3 122 Notophthalmus 3 101, 3 12 1 Notophthalmus uiridescens 2969, 2971, 3012, 3042, 3226-3227, 3247, 3258 Notophthalmus uiridescens uiridescens 3043 Nyctimystes dayi 2990, 3017

0 Oncorhynchw mykiss 297 1 Oncorhynchus tshawytscha 3 174 Oomycota 3035 Ophiostoma ulmi 3034 Osteopilw 3101, 3122 P Paa 3122 Parakassina 3 123 Pelobates 3 122 Pelobates cultripes 3 160, 3 166, 3 172 Pelobatesj&scw 3038 Pelobatidae 3122, 3248 Pelodytes 3 122 Pelodytidae 3 122 Peronosporomycotu 3035 Phialopora 3043 Phaeognathw 3 102 Phaeosphaeriaceae 3040 Philoria 3 122 Phoca hispida 32 12 Phrynobatrachw 3 122 Phrynomantis 3 123 Phytophthora 3034 Phytophthora cinnamomi 3034 Phytophthora ramorum 3034 Pimephales promelas 3157, 3213, 3216 Pipa 3 124 Pipidae 3015, 3121, 3127, 3248 Pisces 2970 Platyhelmintes 3068 Plethodon cinereus 3247, 3249-3250, 3257-3258, 3273

Plethodontidae 3015, 312 1, 3 176, 3258, 3272 Plethodon uehiculum 3 164, 3 167 Pleurodeles 3 121 Pleurodeles waltl 3 133, 3 160, 3 171 Podochytrium dentatum 3030 Poecilia reticulatus 3 163 Proteidae 3 121, 3 125 Proteus 2979 Protopolystoma xenopodis 3075 Pseudacris 2971, 3101, 3122 Pseudacris crucijer 31 13, 3213, 3248, 3254 Pseudacris negrita 3248 Pseudacris ornata 3248 Pseuducris regilla 3071-3072, 3097, 3 100, 3106, 3158-3163, 3257 Pseudacris streckeri 3036 Pseuidacris triseriata 3156, 3160, 3165, 3166, 3169, 3190, 3248, 3252 Pseudidae 3 122 Pseudis 3 122 Pseudobranchus 3 102, 3 121 Pseudomonas 2979 Pseudotriton 3 102 Pternohyla 3 102 Ptychadena 3 122, 3240 Pyxicephalw 3 122

R

Rana 3038, 3070, 3101, 3106, 3122 Rana arualis 2993, 3113, 31 18, 3133, 3247, 3248-3251, 3254-3258, 3260 Rana aurora 297 1, 3 114, 3 118-3 119, 3129-3130, 3152, 3158, 3160-3162, 3165, 3169-3170, 3175, 3271 Rana aurora draytonii 3 199 Rana berlandieri 3036 Rana blairi 3012, 3114, 3119 Rana boylii 3003, 3006, 3016, 3060, 3199 Rana calamita 3125, 3128, 3258 Rana cmcadae 3039, 3 1 14, 3 1 18-3 119, 3125-3126, 3129, 3133-3135, 31613162, 3164, 3167-3169, 3172-3173, 3198-3199, 3251 Rana catesbeiana 2964-2966, 2968, 2971-2972, 2977-2979, 2991-2992, 3003, 3006, 3016, 3060, 3097, 3099, 3114, 3125, 3133, 3161-3163, 3169, 3176, 3219, 3227, 3241, 3248, 3237, 3273 Rana chiricahuensis 2992 Rana clamitans 2972, 2977, 3042, 3076, 3078-3080-3080, 3114, 3156-3158, 3160, 3162, 3166, 3169, 3193, 3213, 3224, 3226-3227, 3248, 3252, 3254, 3257 Rana dalmatina 2978, 3248 Rana draytonii 2967 Rana esculenta 2968, 3041, 3102, 3129, 3160, 3225, 3248, 3251, 3257 Rana grylio 3257 Rana latmtei 2966, 2973-2974 Rana lessonae 3160, 3251 Rana luteiuentris 31 18, 3129, 3241 Rana mucosa 2978, 2992, 3012, 3014, 3019, 3056, 3060, 3197, 3199, 3229, 3248, 3257-3258 Rana ornatiuentralis 3097 Rana palwtris 3248, 3257

Rana pipiens 2965-2966, 2971, 29772978, 2980, 2992, 3072, 3075, 3077, 3079-3083, 3089, 3093, 3096-3097, 3101, 3104, 3106, 3114, 3133-3134, 3157-3158, 3160, 3162-3163, 3165, 3167, 3169, 3171, 3173-3174, 3198, 3218, 3226, 3230, 3247-3248-3252, 3254, 3257, 3270 Rana pretiosa 3 118, 3 129, 3 160-3 163, 3168-3169, 3175-3176 Rana rzdibunda 3160, 3248 Rana rugosa 3224 Rana rugulosa 2979 Rana septentrionalis 2977, 3097, 3100, 31 14, 3224, 3257 Rana sphenocephala 3012, 31 14, 3193, 3248, 3257 Rana sylutaica 2969, 2971, 2973, 2978, 3002, 3042, 3076, 3079, 3081, 3100, 3114, 3128, 3130, 3152, 3160-3162, 3 190, 3 199, 3247, 3248-3249, 325 13255, 3257 Rana tarahumarae 2992, 3018 Ram temporaria 2965, 2967, 2970-2972, 3038, 3041, 3114, 3118-3119, 3125, 3129-3130, 3134, 3159-3160, 3162-3163, 3165, 3167, 3170-3171, 3227, 3247, 3248-3249-3252, 3254-3255, 3257-3259 Rana utricularia 297 1 Rana uirgatipes 3248 Ranavirw 3135, 3273 Rana yauapaiensis 3003, 3005-3006, 3016 Ranidae 3015, 3108, 3122, 3125, 3199, 3222, 3242, 3248, 3256-3257 Rattus rattw 3217 Reptilia 2970 Rhacophoridae 3 123 Rheobatrachus 3 124 Rheobatrachus silw 2989 Rhinatrematidae 3 121 Rhinoderma 3 124 Rhinodermatidae 3 122 Rhinophrynidae 3 121 Rhinophrynus 3 121 Rhizophydium 2993-2996, 3001, 3029 Rhizophydiurn haynaldii 3029 Rhizophlyctis 3029 Rhyacotriton 3102, 3167 Rhyacotritonidae 3121, 3125, 3176 Rhyacotriton uariegatus 3 164 R h y n o p h ~ u s3 102 Ribeiork 3070-3074, 3 104-3 109 Ribeioria ondatme 3067, 3071-3072, 3077, 3079-3080, 3082-3084, 3093, 3096, 3099, 3103-3104 S Salalnandridae 3015, 312 1, 3 176, 3249, 3256, 3258 Salamandra salamandra 3 125 Salamandrella 3 121 Salmo gairdnierii 3 157 Salmonella 2964 Salmonidae 3 157, 3 161 Salmo salar 3 160-3 161, 3227 Saprolegnia 3035-3036, 3038-3041, 3117, 3135, 3175, 3189 Saprolegnia diclina 3035 Saprolegnia ferax 3035, 3037-3039, 3194, 3273, 3275-3276 Saprolegnia parasitica 3035, 3037

AMPHIBLAN BIOLOGY

Sapmlegniaceae 3035-3037, 3039 Saprolegniales 3035 Sarcoptes scabiei 3009, 3034 Scaphiopodidae 3 122 Scaphiopus 3 101, 3 122 Scaphiopus holbrookii 3248 Scaphiow intermontanw 3248 Scinax 3122 Scolecobasidium 3043 Scolecomorphidae 3 121 Semnodactylus 3 123 Silurana 3 121 Siren 3102, 3121 Sirenidae 30 15, 312 1 Smilisca 3 102 Sooglossidae 3 122 Spea 3101, 3122 Spea bombzfions 3036 Spea intermontana 297 1 Sphaerotheca 3 122 Spicospina 3 122 Spizellomycetales 2994 Stereochilw 3 102 Stramenopila 3035 Streptococcus 2979 Streptococcus i n k e 2979 T Taricha 3101, 3121 Taricha granulosa 3 114-3 115, 3 119, 3129, 3164, 3167-3168

Taricha torosa 3107, 3114, 3118, 3135 Taudactylus acutirostris 2987, 2989, 3016-3017 Taudactylus diurnus 2989 Taudactylus eungellensis 2990, 3014, 3016, 3019 Tinca 3102 Tomopterna 3 122 Trachemys scripta 3225 Triturus 3037, 3121 Triturw alpestris 3 114, 3 124, 3 131, 3249, 3258 Triturus carnqex 3225 Triturus cristatus 31 14, 3227, 3249, 3252, 3258 Triturus helveticus 3 160, 3 166, 3 169, 3171, 3249, 3252, 3258 Triturus marmoratus 3 125 Triturus vulgaris 3040, 3 160, 3 166, 3168-3169, 3171, 3247, 3249, 3252, 3258 Tjphlotriton 3 102 U Uperodon 3 123 Uperoleia 3 122 Urae~t~phlidae 3 121 Ursw maritimus 32 12

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Xenopus 2973, 3101, 3108, 3121, 3153, 3192 Xenopus plli 2993, 3248, 3254-3255 Xenopus laevis 2979-2980, 2992-2994, 3016, 3020, 3075, 31 14, 3133-3134, 3153-3154, 3157-3160, 3162-3167, 3170-3171, 3174, 3217, 3219-3220, 3222-3224, 3240-3241, 3247-3248, 3255 X e n o f w muelleri 2993 Xenopus tropicalis 2979-2980, 29922994, 3012, 3016, 3020, 3031