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 ...

Author: Heatwole | H. & J. W. Wilkinson (eds.)

88 downloads 890 Views 49MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!

Report copyright / DMCA form

DOWNLOAD PDF

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 <34"C) but not insect cell lines in vitro (Speare and Smith 1992). The Venezuelan Ranavirus replicated readily at temperatures ranging from 18°C to 30°C (Zupanovic et al. 1998b). 3. Clinical Sigm and Pathology

The gross pathology of ranaviruses appears similar to that associated with bacterial septicaemia. Since opportunistic or resident bacteria can often be cultured from dead or sick frogs, it appears that many ranaviral outbreaks have been misdiagnosed as septicaemia (Green 2001; see also "Bacterial septicaemia 'red leg"' in the "Bacterial Diseases7' section below). For a more thorough treatment of ranavirus pathology see Wright and Whitaker (2001). Frog Virus-3: Frog virus-3 (FV3) was found while searching for a viral cause of the Luck6 tumour in Rana pipiens in the United States (Granoff et al. 1969). Subsequently, much work was carried out on the morphology and life cycle of FV3 in the laboratory, and experimentally it was shown to cause oedema, necrosis, haemorrhage, and death in embryos, tadpoles, and recent metamorphs (Granoff 1989). During experimental infections, metamorphic toads developed haemorrhages, accompanied by massive oedema, in the ventral skeletal musculature, stomach, and intestines (Came et al. 1968). In addition, tadpoles of the Italian agile frog (Rana latastei) exposed to FV3 sometimes became emaciated (Pearman et al. 2004). Mortality in embryos can occur 3-12 days post-exposure with 4070% survivorship of tadpoles (Tweedell and Granoff 1968). Clinical signs include depigmentation, skin sloughing, and spinal curvature. Generally, the lesions caused by FV3 appear to be milder than those caused by Tadpole oedema virus (Green 2001). Tadpole Edema Ems: Tadpole Edema Virus (TEV) was first isolated from oedemic Rana catesbeiuna tadpoles from West Virginia, USA in August 1965 (Wolf et al. 1968, 1969). 5iolecular evidence suggests that TEV is a strain of FV3 (Hyatt et al. 2000). Acute fatal disease was reproduced with experimental transmission to tadpoles and adults, although pathogenicity varied among host species (Wolf et al. 1968). Terminal changes in tadpoles are obvious oedema, particularly in the ventral regions, and haemorrhaging along the body

HEMINGWAY ET AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS

2967

and hind limbs (Fig. 1) (Green 2001). Internally, haemorrhages and necrosis typically extend into the stomach, intestines, and skeletal muscles, and may include the mesonephros, bladder, liver, lungs, and spleen (Green 2001). The highest titres of the virus occur in the stomach (Clark et al. 1969; Green 200 1). UK Ranavirus: While molecular studies suggest UK ranaviruses are strains of FV3 and may have originated in North America (Hyatt et al. 2000), theses viruses are not highly pathogenic to embryos and tadpoles (Cunningham et al. 1996, Cunningham et al. 2007a). There are several distinct "syndromes" associated with the UK ranaviruses including an "ulcerative syndrome," "haemorrhagic syndrome," and "ulcerative and haemorrhagic

Fzg. I . Ventral view of a moribund larval California red-legged frog (Rana draytonii) with oedema and petechial haemorrhages in the hind legs and inguinal region typical of TEV infection. Photograph by Valentine Hemingway.

syndrome" (Cunningham et al. 1996). While these syndromes are associated with infection by the bacterium Aeromonas hydrophila, Cunningham et al. (2007a) demonstrated that bacteria were not the cause. Frogs with the "ulcerative syndrome" have skin ulcerations and, to a lesser extent, rlecrosis of the legs, while systemic haemorrhages, particularly in the myoskeletal, digestive, renal, and reproductive organs, lungs, and pancreas characterize the "haemorrhagic syndrome" (Cunningham et al. 1996, Green 2001). Frogs with either or both syndromes were thin and some experienced lethargy (Cunningham et al. 1996). Histological lesions typical of "ulcerative syndrome" include epidermal thickening, and epidermal necrosis, as well as necrosis, granulocytic inflammation, congestion, and haemorrhage in internal organs (Cunningham et al. 1996). Experimentally infected Rana temporaria died six to eight days post-infection (Cunningham et al. 2007b). Bohle Iridovirus: The only virus to be isolated from Australian frogs, Bohle iridovirus (BIV), was first found in Limnodynastes ornatus that died during metamorphosis in captivity (Speare and Smith 1992). The frogs had been collected as tadpoles from a temporary pond at Bohle, a suburb of Townsville, Queensland. BIV and FV3 appear to be closely related (Hengstberger et al. 1993). Clinical signs of the Bohle iridovirus were oedema of subcutaneous tissue, especially around the jaw and head, and a swollen abdomen due to ascites (Fig. 2). Subcutaneous haemorrhages occurred on the ventral abdomen, inguinal areas. and lower jaw (R. Speare, unpublished data). Typical pathology in natural and experimental infections included severe renal, pulmonary, hepatic, splenic, and haemopoietic

AMPHIBIAN BIOLOGY

1968

necroses and haemorrhages. Ranavirus immunoperoxidase stained many cell types in liver, lung, spleen and, in particular, fibrocytes in extensive areas of swollen, necrotic dermis and glomeruli (Cullen et al. 1995, L. Berger and R. Speare, unpublished data). In experimental infections, high mortality rates in juvenile frogs typically occurred within 5 to 25 days, depending on dose and type of exposure, but adults were less susceptible (Cullen and Owens 2002). Chronic cases were detected by PCR (Cullen and Owens 2002). Ambystoma tigrinum firus: Experimental infections of Ambystoma tigrinum virus ( A m in tiger salamanders caused 40 to 100% mortality within two to three weeks (Brunner et al. 2005). Infected animals may show haemorrhaging in internal organs as well as white skin polyps, skin sloughing with mucus and ulcers, bloody mucus from the cloaca, and lethargy, and may refuse food, while others may die without clinical signs (Fig. 3) (Jancovich et al.

Fig. 2. Metamorphs of Limnodynastes ornatw with ascites. This frog died from Bohle Iridovirus during the original outbreak from which the virus was first isolated. Photograph by Rick Speare.

1997; Chinchar 2002). Further, infected lung, skin, and liver cells are larger than are uninfected cells (Jancovich et al. 1997). Infected animals may be able to maintain chronic, sublethal or aclinical infections, likely contributing to maintenance of infection in the population (Brunner et al. 2004). Mortality in wild populations occurs primarily in larvae in the northern part of their range and in larvae and neotonic adults in the southern part Brunner, unpublished data). of the salamander's range

u.

Croatian Ranavirus: A ranavirus-like agent called "viral haemorrhagic septicaemia of frogs" was found in dying captive Rana esculenta from Croatia that had lethargy, oedema, haemorrhages, and skin necrosis (Fijan et al. 1991). Rana catesbeiuna Virus 2: A ranavirus similar to other ranaviruses, such as FV3, was isolated from cultured R . catesbeiuna tadpoles in the USA (Majji et al. 2006). Rana catesbeiana virus Z (RCV-Z) appears to be much more pathogenic than FV3, causing mortality in 50100% of exposed tadpoles (Majji et al. 2006). Similar to other ranaviruses, symptoms

HEMINGWAY ET AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS

2969

included oedema in the abdomen, haemorrhaging in ventral regions, and lethargy (Maji et al. 2006).

Venezuelan Ranavim: Seven ranaviruses with strong similarities to BIV and FV3 were isolated from wild-caught Bufo mamnus and hpodactylus spp. in Venezuela (Zupanovic et al. 1998b). Infected animals had no external lesions or internal symptoms (Zupanovic et al. 1998b). Fig. 3 . Moribund ATV-infected larval tiger salamander (Ambystoma tigrinum) with oedema of the hind legs and with petechial haemorrhages o n the jaw, inguinal region, legs, and tail. Photograph by Jesse Brunner.

4 . Epidemiology

Impacts on Populations: While the first ranaviruses were isolated in the 1960s (Granoff et al. 1965; Wolf et al. 1968), it was not until the 1990s that ranaviruses were recognized as causing mass mortality in wild populations of amphibians ( A m Jancovich et al. 1997; RUK: Drury et al. 1995). Ranaviruses have since been identified as the agents of mass mortality events around the world. Most outbreaks of amphibian ranaviruses have been reported from North America and the United Kingdom, although this may represent a bias in reporting rather than the actual distributions. Ranavirus-associated die-offs are characterized by a rapid onset with massive mortality involving nearly all of the individuals in a pond (Green et al. 2002); entire cohorts can be killed (Petranka et al. 2003; Brunner et al. 2004). Die-offs also tend to recur in the same ponds or wetlands for several years in a row (Green et al. 2002; Cunningham et al. 2007a; Petranka et al. 2007; Greer and Collins 2008). Whereas ranaviruses tend to cause recurrent mortality, they are not associated with catastrophic amphibian declines (Green et al. 2002; Daszak et al. 2003). Green et al. (2002) found that the majority of amphibian die-offs (25 of 44 investigated) in the United States were caused by iridoviruses, presumably ranaviruses. ATV has a wide distribution in North America, having been isolated from tiger salamanders during dieoffs throughout the western cordillera and central plains (Bollinger et al. 1999; Green et al. 2002; Jancovich et al. 2005). Similarly, W3-like viruses have been associated with amphibian die-offs - generally involving anurans but also spotted salamander larvae (Ambystoma maculatum) and adult eastern spotted newts (Notophthalmus viridescens) (Petranka et al. 2003; Duffus et al. 2008) - across North America (Mao et al. 1999; Green et al. 2002; Petranka et al. 2003; Greer et al. 2005; Gray et al. 2007; Dufhs et al. 2008; St. Amour et al. 2008; Schock et al. 2008). FV3-like ranavirus caused recurrent, catastrophic die-offs of larval Ambystoma maculatum and Rana sylvatica in a network of natural and constructed ponds and wetlands in North Carolina from 1997, when they were first detected (although monitoring had begun in 1994), until at least 2006 (Petranka et al. 2003, 2007). In most years well over half of the

29iO

AMPHIBIAN BIOLOGY

ponds surveyed experienced die-offs. Juvenile recruitment was dramatically affected by these die-offs, as well as by droughts, which led to a reduction in breeding activity in later years (Etranka et al. 2007). Populations of both species, however, remained extant in this complex. It is worth noting that die-offs and amphibians with signs consistent with ranavirus infection have not been observed in other ponds and wetlands in the region (Harp and Petranka 2006), suggesting that outbreaks of FV3 can be highly localized. Populations of Ambystom tigrinum nebulosum in northern Arizona and the endangered A. tigrinum stebbinsi in southern Arizona have experienced recurrent, catastrophic die-offs since at least the mid-1980s (Collins et al. 1988); these events were later attributed to the mavirus, ATV (Jancovich et al. 1997), yet these population persist (Greer and Collins 2008). At the level of a pond, however, ranavirus epidemics may cause local extinctions; tiger salamanders were apparently extirpated in some ponds in Saskatchewan after 3 to 4 years of ATV-related die-offs (Carey et al. 2003). Unlike Am, which has only been found in central and western North America (Jancovich et al. 2005), and BW which has been detected in the wild only once, in Queensland, Australia (Speare and Smith 1992; Hengstberger et al. 1993), FV3 is globally distributed. FV3-like viruses have also been reported from dead and moribund amphibians in captive populations in North America (Robert et al. 2007; Miller et al. 2008) and in frog farms in Brazil and Uruguay (Galli et al. 2006), Thailand (Kanachanakhan 1998), and China (Zhang et al. 1999). The most extensive amphibian ranavirus epidemics have occurred in the United Kingdom since the mid 1980s, where FV3-like ranaviruses have caused annual mass dieoffs of common frogs ( R a m temporaria) and common toads (Bufo bufo) in garden-ponds across much of the country, but focused in the south and east (Drury et al. 1995; Cunningham et al. 1996; Cunningham et al. 2007a,b). FV3-like ranaviruses were the putative cause of these die-offs (Drury et al. 1995; Cunningham et al. 2007a) affecting tens of thousands of frogs across the United Kingdom but, again, there is little evidence of population declines (Daszak et al. 2003). Still, as Daszak et al. (2003) argued, the potential for ranaviruses to impact amphibian populations should not be discounted. There are at least two reasons why populations may remain extant, or even abundant, in the face of high ranaviral mortality. First, natural mortality of tadpoles is normally very high due to predation and to drying of ponds but only a small fraction of the individuals need to survive in order for adequate recruitment to occur. Ranaviruses may, therefore, cause only compensatory mortality, with little added effect on long-term population dynamics. Secondly, many affected species are rather common and widespread. Consequently, local populations may be rescued by immigration from other ponds and the metapopulation would persist. Of course habitat loss and alteration of habitat would undermine this effect. L f e stages: Most die-offs caused by ranaviruses involve larvae or recent metamorphs inhabiting permanent water (Speare and Smith 1992; Jancovich et al. 1997; Green et al. 2002; Petranka et al. 2003; Fox et al. 2006). Several studies involving experimental exposures have shown that larvae are much more susceptible to BIV and FV3-like viral infections than are adults (Wolf et al. 1968; Clark et al. 1969; Cullen and Owens 2002; Gantress et al. 2003; but also see Cullen et al. 1995), and the patterns of mortality in the wild bear this out. The ranaviruses in the United Kingdom are the exception: larvae are affected, but most of the mortality is observed in adult frogs and toads (Cunningham et al. 1996; Cunningham et al. 200'7a). Range in Host Taxa: The three species of amphibian ranaviruses have a broad range in host taxa. Overall, the array of hosts of ranaviruses under ecologically realistic conditions remains poorly resolved but, potentially, it is important to the epidemiology of ranaviruses in amphibian communities. Several of the amphibian ranavirus are capable of infecting hosts from three vertebrate classes: Amphibia, Pisces and "Reptilia". BIV was highly pathogenic

HEMINGWAY ET AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS

297 1

for mammalian cells in vitro, but presumably in vivo protection is provided by the host's body temperature being above the maximum for replication of BIV (Speare and Smith 1992). BIV was first isolated from the ornate burrowing frog (Limnodynastes ornatus), but has since been found to infect other captive or experimental hosts, including Bufo marinus, Litoria terraereginae, L. caerulea, L. albogutatta, Cyclorana brevipes, and L. latopalrnata, as well as the native barramundi fish (Lutes calcarifer) and reptiles (Owens 1994; Cullen et al. 1995; Ariel 1997; Cullen and Owens 2002; Moody and Chinchar 2002; Williams et al. 2005). Juvenile L. terraereginae, L. caerulea, and L. latopalmata were highly susceptible to BIV, while larval L. terraereginae and adults of L. caerulea and other species were less susceptible. Mortality varied with dose and route (Cullen et al. 1995). Frog Virus 3-like viruses are also infectious to frogs, salamanders, fishes, and reptiles. Mao et al. (1999) found apparently identical FV3-like viruses in sympatric three-spine sticklebacks (Gasterosteus aculeatus) and a tadpole of the northern red-legged frog (Rana aurora), and Johnson and Jacobson (2004) isolated a ranavirus identical in MCP sequence from a captive Burmese star tortoise (Geochelon latynota) and an ill southern leopard frog (Rana utricularia) in the tortoise's enclosure. FV3-like viruses commonly infect both frogs and salamanders under natural conditions. In the 1960s TEV, an FV3-like virus isolated from Rana catesbeiana, was found in tadpole of Bufo amerLcanus, B. woodhousii fowlen, and Spea intermontana (Wolf et al. 1968, 1969). More recently, Petranka et al. (2003) reported FV3-like viruses from sympatric Ambystoma mnuculaturn and Rana qlvatica in North Carolina, and Duffus et al. (2008) found apparently identical FITS-likeviruses in larvae of Pseudacris spp., Hyla versicolor and ambystomatid salamanders in south-central Ontario. Schock et al. (2008), however, found that while Amtytomnu tigrinum, Ram sylvatica, R. pipiens, and Hyla regilk could be experimentally infected with both FV3 and Am, in at least one pond, strains of ATV and FV3 were co-circulating in distinct host species. Ram temporaria United Kingdom virus (RUK) has been reported in the common frog (Rana temporaria) (Cunningham et al. 2006) and a similar virus, Bufo bufo United Kingdom virus (BUK), has been found to infect both the common toad (Bufo bufo) and (experimentally) the common frog (Cunningham et al. 200'7b). ATV may have the most restricted array of hosts. Based on experimental challenges of the northwestern salamander (Ambystoma gracile) and the eastern newt (Notophthalmz~s viridescens), as well as Rana pipiens and R. catesbeiana and three species of fish (Gambusia afinis, Lepomis afinis, and Oncorhynchus mykiss) with ATV-SRV (an isolate from A. tigrinum stebbinsi in the San Rafael Valley in southern Arizona), Jancovich et al. (2001) concluded that ATV was restricted to salamanders. Recent experiments with ATV-RRV (an ATV isolate from Roussell Pond, Regina, Saskatchewan, Canada), however, demonstrated that anurans can be infected with ATV (Schock et al. 2008). ATV has yet to be detected infecting any species of fish. Seasonality: One hallmark of ranavirus epidemics is their seasonality-mortalities primarily occur in late spring and in summer (Green et al. 2002; Petranka et al. 2003; Brunner et al. 2004; but see Gray et al. 200'7). One potential explanation for this seasonality is that amphibians are particularly vulnerable to ranaviral infection in the late spring and in summer when the larvae of many species begin to metamorphose. Many components of the amphibian immune system are down-regulated just prior to metamorphosis (RollinsSmith 1998; Carey et al. 1999). This does not appear to be the case, however, for ATV in tiger salamanders-Brunner et al. (2004, 2005) found that ATV-infected larvae that metamorphose have higher survival rates. Gray et al. (2003) also found that the prevalence of infections by FV3-like virus in bullfrog larvae (Rana catesbeiana) in eight wetlands in Tennessee, United States, decreased with Gosner stage, up through stage 41, which may reflect increasing immunocompetence during development or, alternatively, increasing disease-induced mortality rates (McCallum and Dobson 1995). Adding to the ambiguity, there was no trend in prevalence with developmental stage in the larvae of sympatric green

2972

AMPHIBIAN BIOLOGY

frogs (Ranaclamitans) (Gray et al. 2007). Moreover, developmental changes in susceptibility would not seem to explain the seasonality of ranavirus-associated mortality of adult R a w temporaria and Bufo bufo in the United Kingdom, which tends to peak in July, August, and September, although mortality was observed for at least 40 weeks at one location (Cunningham et al. 1996). Instead seasonal changes in prevalence and mortality may reflect changes in temperature or other environmental factors (Rojas et al. 2005; Raffel et al. 2006; Gray et al. 2007), or simply the dynamics of recurrent, growing epidemics. For instance, at high elevations in northern Arizona, U.S.A. mass die-offs involving hundreds to thousands of primarily larval Ambystoma tigrinurn nebulosum occur in natural ponds and in earthen stockwatering tanks in the summer or early autumn (Berna 1990; Brunner et al. 2004; Greer and Collins 2008). A similar seasonal pattern has been observed in Saskatchewan and Manitoba, Canada (Bollinger et al. 1999; Schock 1999). Brunner et al. (2004) argued that summer or autumn die-offs were the more conspicuous peaks of epidemics that began in early spring, when the occasional aclinically infected adult introduced the virus into ponds when they returned to breed. These authors showed that while many, or even most, larvae die during the epidemic, some young of the year leave the pond infected. A few of these apparently survive with sublethal infections because a small fraction of adults returning to ponds to breed are infected with ATV (Brunner et al. 2004). The persistence of sublethal or even aclinical infections may explain both the recurrence and seasonality of ATV epidemics, and may be important to the epidemiology of other ranaviruses as well. Interestingly, in southern Arizona, where ponds remain ice-free year-round and the salamanders remain in ponds in both metamorphic and neotenic forms, epidemics have been observed throughout the year (Collins et al., unpublished data). Effect of Habitat: Several studies of ranaviruses in amphibian populations have found that artificial or human-impacted bodies of water are more prone to epidemics than are natural ones. Greer and Collins (2008) found that ponds modified to retain water for livestock were over four-times more likely to experience epidemics than were natural sinkholes. With little emergent vegetation in which to hide, Ambystoma tigrinum larvae aggregated at the edges of modified ponds in a "halo," which may have led to increased contact rates and therefore to viral transmission among larvae (Greer and Collins 2008). Similarly, there was a slight increase in the proportion of FV3 die-offs occurring in constructed ponds of the Tulula wetlands complex in North Carolina, compared to those occurring in the natural reference ponds in the complex (Petranka et al. 2003). Gray et al. (2007) found that the prevalence of an FV3-like virus was higher among Rana clamitans in wetlands with access by cattle compared to those without cattle access, although this trend was not significant among sympatric Rana catesbeiana. The authors suggested that this difference was due to these two species having different tolerances to poor water quality (Gray et al. 2007). Degree of human influence and proximity to industrial activity and to human habitation were all significantly associated with higher FV3 prevalence in R. chmituns (presumably adults) in central and northeastern Ontario, Canada (St. Amour et al. 2008).

A higher position within a catchment basin in Acadia National Park in Maine, United States, seemed to predispose wetlands for ranaviral outbreaks (Gahl and Calhoun 2008), although the reasons for this are not clear.

5. Resistance to Infiction Amphibians may fight ranaviral infections via both acquired and innate immunity. A first line of defence includes antimicrobial peptides of the skin (Erspamer 1994) which have been shown in some species to be effective against ranavirus in nitro (Chinchar et al. 2001; Rollins-Smith et al. 2002). Antibodies specific to ranavirus are also produced, although their efficacy appears to be species specific (Chinchar et al. 1984). There seems to be a distinction between anuran and caudate immune responses to ranaviral infections. Rana catesbeiana tadpoles that recovered after exposure to FV3 or RCV-Z appeared to be protected from hture lethal infection with RCV-Z (Majji et al. 2006). Brunner and Schock (unpublished data) found that surviving an experimental exposure to ATV does not provide protection

HEMINGWAY ET AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS

2973

against later exposures (i.e., no immune memory). Cotter et al. (2008) found that although experimentally infected Ambystoma mxicanum mounted an impressive immune response, they lacked the production of lymphocytes by the spleen that are associated with the ability of adult Xenopus to clear ranaviral infections (Robert et al. 2005; Maniero et al. 2006). This may be an important function in species that are able to clear ranaviral infections. The outcome of infection varies greatly with genetic identity of individuals as well as with life history stage (Brunner et al. 2005; Pearman and Garner 2005). Survivorship when infected with ranavirus also appears to be related to dosage. When groups of tadpoles of the Italian agile frog (Rana latastei) were exposed to different dosages of FV3, the animals that received the lowest dosage survived the longest (Pearman et al. 2004). Duffus et al. (2008) found that when tadpoles of the wood frog (Rana sylvatica) were exposed to increasing dosages of FV3, there was a corresponding increase in the proportion of infected tadpoles. Brunner et al. (2005) demonstrated that the proportion of tiger salamanders (Ambystoma tigrinum) that became infected with ATV increased with dose of inoculum, as did mortality, whereas time to death was reduced. Dose varies with route of exposure and this may play an important role in the outcome of infection (Cullen and Owens 2002; Brunner et al. 2005). 6. Transmission and Spread

Local transmission of ranavirus can occur via several direct and indirect routes. While a variety of laboratory studies have demonstrated that ranaviruses can be transmitted via indirect routes, i.e. fomites, soil, contaminated water (Pearman et al. 2004; Duffus et al. 2005; Harp and Petranka 2006; Brunner et al. 2007), it appears that more direct contact, i.e. touching, biting, cannibalism, necrophagy, may be required for transmission in a natural setting (Jancovich et al. 1997, Brunner et al. 2004, 2007; Pearman et al. 2004; Parris et al. 2005; Harp and Petranka 2006). The rate and outcome of infection seem to vary with the route of exposure. For example, when infected tadpoles of the wood frog (Rana sylvatica) were introduced into a tank with uninfected tadpoles, mortality was upwards of 98%, while tadpoles exposed only to water and sediment from a site with an active ranavirus die-off did not experience a catastrophic die-off and individuals primarily developed aclinical ranaviral infections (Harp and Petranka 2006). Movement of water or sediment via fomites or aclinically infected animals between sites may facilitate the spread of ranaviruses (Harp and Petranka 2006). One of the most important issues in epidemiology and conservation is whether transmission scales with host density (McCallum et al. 2001; de Castro and Bolker 2005). When transmission is density-dependent, pathogens can (at least theoretically) drive the population only so low because at some threshold transmission becomes so rare that the pathogen fades from the population. If, however, transmission continues unabated even as the host population declines, then extinction is possible. The evidence from ranaviruses is mixed. In a controlled environment, when infected Rana sylvatica tadpoles were placed in a pool with healthy tadpoles, more than 98% of the tadpoles died, regardless of initial tadpole density in the pool (Harp and Petranka 2006). In contrast, Greer et al. (2008) found that infection rates in na'ive tiger salamander larvae increased with the density of ATVinfected hosts to which they were exposed, although not in a strictly density-dependent manner. Interestingly, greater densities of infected larvae led to an earlier death for the newly exposed larvae. Necrophagy and cannibalism of infected animals are also potentially important routes of direct transmission. Necrophagy, in particular, seems to be a density-independent form

of mnsmission; infected carcasses are a steady source of infection regardless of host density. Transmission by necrophagy and cannibalism is common in host species such as Ambystoma hgnnum, Rana sylvatica, and R. latastei (Pearman et al. 2004; Harp and Petranka 2006; Brunner et al. 2007) and infections acquired by these routes seem to be more lethal.

2974

AMPHIBIAN BIOLOGY

Tadpoles of R . Zutastei that fed on carcasses of FV3 infected tadpoles became infected and were more likely to die than were tadpoles in close proximity to infected carcasses but without direct contact (Pearman et al. 2004). Ram sylvatica tadpoles allowed to consume infected tadpole carcasses died sooner than did those exposed to infected carcasses but not allowed to eat them (Harp and Petranka 2006). Larvae of Ambystoma tigrinurn nebulosum were also easily infected with ATV when allowed to eat tissue from dead, infected larvae, but they also were easily infected via contact with mucous from infected larvae or with a short "bump" with infected larvae (Brunner et al. 2007). All of these modes of direct contact are common in amphibian larvae and it is likely that all of these behaviours contribute to ranaviral transmission in the wild. Transmission through indirect routes has also been demonstrated in the laboratory. Langdon (1989) showed that ranaviruses could survive in a laboratory for over 90 days. In addition, several studies have found that larvae could become infected with ranavirus when exposed to water that previously housed infected larvae andlor sediment from a ranavirus die-off site or when inoculated with ranavirus (Brunner et al. 2004; Harp and Petranka 2006; Duffus et al. 2008; Brunner et al. 2007), although ranaviruses seem to decay in pond water (Brunner et al. 2007). Harp and Petranka (2006) found that the tadpoles exposed to water and sediment from a die-off site experienced only a low mortality rate, and were instead aclinically infected. They hypothesized that this may be a mode of maintaining infection in the population, as well as potentially moving it to other sites with migrating, infected amphibians. Spread of ranaviruses between sites may be due to movement of water, sediment via fomites, or sublethally or aclinically infected animals (Harp and Petranka 2006). Most understanding of transmission of ranavirus originates, however, from experiments in the laboratory, leaving one to extrapolate how the dynamics of transmission play out in a natural and more complex setting. For a discussion of long-distance, anthropogenic spread of ranaviruses, see Section 2 "Taxonomy and Molecular Epidemiology".

7. Diagnosis The current routine techniques for diagnosing ranaviruses in amphibians include immunohistochemistry, viral isolation from tissues and cell culture, and ELISA and PCR. Sublethal or aclinical infections may only be detectable by cell culture or PCR. Immunohistochemistry: Wax-embedded tissue sections can be labelled with an anti-ranaviral immunohistochemical marker so as to visualize infection within tissues (see Cunningham et al. [2008] for a detailed description). Variations on this technique include immunofluorescence and immunoelectron microscopy (Zupanovic et al. 199813).

Cell Culture: Ranaviruses can be grown from fresh or frozen tissues in various fish (e.g. EPC and FHM), mammalian, or amphibian cell lines (Hengstberger et al. 1993) as long as those cells grow at ectothermic temperatures (<25"C). Often samples from amphibian tissues only show cytopathic effects after two or three passages (Brunner, unpublished data). The species of the cultured ranavirus can be identified by sequencing a portion of the major capsid protein gene using a technique described by Marsh et al. (2002). ELISA: Enzyme-Linked ImmunoSorbent Assay (ELISA) is used for detection of antiranaviral antibodies in amphibian sera. Both Whittington et al. (1997) and Zupanovic et al. (1998b) provided detailed descriptions of this technique. PCR: Polymerase chain reactions (PCR) using primers designed to amplify portions of the major capsid gene have become the most common method of detecting ranaviruses. For instance, the MCP4 and MCP5 primers described by Mao et al. (1996) amplify an approximately 500bp region of the MCP gene in all known ranaviruses. Quantitative realtime PCR reactions have also been developed (Mao et al. 1996; Brunner 2004). Galli et al.

HEMINGWAY ET AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS

2975

(2006) provided a useful description of sample extraction and PCR procedures. Various tissues have been used to detect ranaviral infections in epidemiological studies and laboratory experiments, although the sensitivity of PCR tests seems to vary among tissues. St. Amour et al. (2007) found that toe clips, which can be taken non-lethally, were less sensitive for ranaviral detection in green frogs than were liver samples. Similarly, Greer et al. (2007) found non-lethal tail clips to be less sensitive than were whole-body homogenates of experimentally infected tiger salamander larvae. They also found that the ability to detect ATV infections increased with time from infection. Thus, these non-lethal sampling methods should be viewed as providing conservative estimates of infection. 8. Management

Given recent declines in amphibian populations worldwide and the impact novel pathogens have had on na'ive populations, it is essential to avoid spreading pathogens or artificially increasing rates of infection. Disinfection of equipment used in handling captive and wild amphibians and effective quarantine guidelines are key to effectively managing these risks. Since ranaviral disease has been listed by the OIE as a globally notifiable disease, the international legislative requirements will result in a greater focus on surveillance and quarantine for ranaviruses. Control Options: Control of ranaviruses within a site that is already infected may be particularly difficult given the possibility that aclinically infected animals may maintain infection in a population (Brunner et al. 2004; Robert et al. 2007). There are no vaccines available for ranaviruses and, given the logistical difficulties and expenses associated with developing and distributing a vaccine, it seems unlikely that one will be forthcoming. Culling amphibian populations would also be difficult given the secretive nature of many amphibian species, and likely ineffective if transmission is not density dependent (Greer et al. 2008). Currently there are no known treatments for ranaviruses. The only method for their control, therefore, is to limit human-influenced spread of the pathogen. Disinfection: Whether working with captive or wild amphibians, it is essential to minimize the risk of amplifying transmission of pathogens or of exposing animals to new strains. Much of the risk can be managed through careful disinfection of potential fomites both between sites and between animals. At the level of the site, it is important to disinfect waders, boots, boats, float tubes, nets, seines, traps, vehicle tyres and undercarriage, and any other equipment that comes in contact with the water. Before disinfecting equipment, it must be scrubbed clean to permit all surfaces to contact the decontamination solution. Several common disinfectants (70% ethanol, 70% isopropyl alcohol, 10% household bleach) appear to be effective in inactivating ranaviruses if applied liberally for sufficient time, and when used in conjunction with mechanical scrubbing; for more detail see Brunner and Sesterhen (2001). In particular, a dilute solution of bleach is effective, has broad spectrum, is inexpensive, and oxidizes quickly (Speare et al. 2004; D. Green personal communication). Ethanol (70%) is effective against the fish ranavirus, EHNV (Langdon 1989) and can inactivate ranaviruses if given sufficient time or if used to flame equipment (Brunner and Sesterhen 2001; Brunner unpublished data). Because ethanol is relatively expensive, it is generally used only on forceps, scissors, and other instruments. Quaternary compounds are also effective and have the advantage of being less corrosive than bleach, but they require careful rinsing to remove soapy residues. Any disinfectant must be applied for the specified amount of time (Speare et al. 2004) to be effective. Finally, if using a chemical disinfectant, the equipment can be rinsed with sterile water, thus lowering the potential for release of the chemical into the habitat and decreasing the wear on equipment. All instruments and tools that come into contact with animals should be disinfected between each animal. Each animal should be handled with new, disposable gloves, and

2976

AMPHIBIAN BIOLOGY

gloves should be changed between each animal. A new or disinfected bag or container should be used to hold each animal individually. Collectively, these procedures will help lower contact rates between the animals and avoid a positive bias when testing for prevalence of infection. In the laboratory, several other options are available for decontaminating equipment. Autoclaving or exposure to UV light are effective in killing viruses, again when the equipment is first scrubbed and the method is applied for the specified amount of time (Speare et al. 2004). In addition, any water or other materials contacting captive animals must be disinfected, using bleach or autoclaving, before disposal. In commercial frog enterprizes, if water is circulated by pumps, potential ranaviruses can be killed by use of commercial UV sterilizers (minimum effective dose of UV is 2.6 x 104 uW.sec/cm2) which are effective at high flow rates and even up to 70% turbidity (Miocevic et al. 1992). Refer to Speare et al. (2004) or Kast and Hanna (2008) for more details on disinfecting equipment. Note that ranaviruses are more resistant than is the amphibian chytrid fungus Batrachochytrium dendrobatzdis: guidelines for killing this fungus may not be adequate for ranavirus. Quarantine Guidelines: There are several published quarantine guidelines intended to decrease the risk of introducing pathogens into a captive setting or of releasing captive, infected animals. Detection of ranaviruses poses special consideration since they can be difficult or impossible to detect in aclinically infected animals. A brief synopsis follows, but one should also consult the papers by Lynch (2001) and Ferrell (2008) for a more thorough treatment of this topic. When obtaining animals, it is important to consider their origin and whether captive sources follow strict health guidelines (Lynch 2001). On arrival, animals should be held in a separate quarantine room for a minimum of 60 days and their health status checked regularly; they should be tested for fungal, bacterial, viral, and parasitic infections (Lynch 2001; Ferrell 2008). During this time clinical signs of some pathogens of concern, including ranavirus, may become manifest. Unfortunately, some animals that are infected with ranavirus remain aclinical. Brunner et al. (2004) found that aclinical ATV infections recrudesced in salamander larvae when they were co-housed with others under apparently stressful conditions. This may improve the ability to detect infections in newly obtained animals, but one should not rely on aclinical infections becoming patent. Currently there is no definitive test for individual animals for ranavirus that does not involve sacrificing the animal. There are several options for proxies on groups of animals. An ELISA test on sera can test for antibodies, thereby providing information on whether animals were ever exposed to ranavirus (Whittington et al. 199'7; Zupanovic et al. 1998b). Additionally, PCR tests of toe-clips or tail-clips can provide a rather high confidence level about infection within the group, if a large enough number of animals are tested (Greer and Collins 2007; St. Amour and Lesbarreres 2007). Similarly, sacrificing a proportion of animals in the p u p to test organ homogenate or whole-animal homogenate via PCR will provide definitive testing for those animals and, depending on the proportion of animals tested, give a rather high level of confidence (Mao et al. 1996). Any animals to be released into the wild should be certified disease free (Lynch 2001; Ferrell 2008). During the minimum 60-day holding period, quarantined animals should be held in a separate room from other individuals. It is important to minimize human contact with animals, use dedicated equipment, wear new, disposable gloves at all times, and use a disinfectant foot-bath when entering and exiting the room (Lynch 2001; Ferrell 2008). Additionally, all surfaces should be disinfected regularly; cages and cage materials should be disinfected when removing animals, and all waste water must be treated (Lynch 2001; and Ferrell 2008). Risk of introduction: Anthropogenic movement of animals infected with ranavirus may be one of the most important sources for spread of this pathogen to na'ive populations of

HEMINGWAY ET AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS

2977

amphibians (Daszak et al. 2001). Amphibians are transported worldwide for food, research, and in the pet and bait trade. There is little control over this movement or the husbandry of the animals (Daszak et al. 2001; Schlaepfer et al. 2005). Animals to be transported for these activities should be subjected to the quarantine methods described above, holding animals in confinement, examining and testing animals for pathogens, treating as necessary, and disinfecting all equipment and water used. For a more through treatment of this topic, refer to the paper by Daszak et al. (2001). 9. Discussion

Although ranaviruses have not been linked to catastrophic amphibian declines, they can cause high levels of mortality in affected amphibian populations (Cunningham et al. 1996). Ranavirus can survive for long periods in the environment under some conditions (Langdon 1989), they are multi-host pathogens (Schock et al. 2008), and there is little if any regulation on the movement of amphibians around the globe, all of which increase the ability of ranaviruses to emerge in, and have an impact upon, amphibian populations. Given the myriad of other threats to amphibians and the fact that amphibians exist in increasingly fragmented populations and so may be less able to rebound after catastrophic mortality, the potential population-level impacts of ranaviruses should not be dismissed (Cunningham et al. 2007a). For example, populations may be unable to recover from infection with Batrachochytrium dendrobatzdis (amphibian chytrid fungus), if mortality rates are further increased. Both regulation and enforcement to limit human-initiated movement of amphibian pathogens, including ranaviruses, is imperative if there is to be constraint upon the impact of emerging infectious diseases on these sensitive populations. B. Frog Erythrocytic Virus

Frog Erythrocytic Virus (FEV) was discovered in populations of Rana catesbeiana, R. clamituns and R . septentrionalis in Ontario, Canada (Gruia-Gray et al. 1989; Gruia-Gray and Desser 1992). FEV is a large (up to 450 nm in diameter), enveloped, double-stranded DNA virus of uncertain classification within the Iridoviridae. Viral inclusions in the cytoplasm of red blood cells are seen by light microscopy of blood smears, and a large proportion of cells may be infected. Infected red blood cells change shape from oval to spheroidal, and heavily infected frogs can become anaemic (Gruia-Gray et al. 1989; Gruia-Gray and Desser 1992). A survey showed that infection is more common in juvenile bullfi-ogs (3Wo overall) than in adults (9%), and that it peaked in August/September (Gruia-Gray and Desser 1992). Infected juveniles were slightly less likely to be recaptured (4%) than were uninfected juveniles (9%), suggesting that the virus contributed to the mortality of young bullfrogs. FEV is transmitted between frogs by mosquitoes or midges, and is not transmitted by water, orally, or by leeches. Similar large viruses have been found in red blood cells of amphibians in Costa Rca, Brazil, South Africa, China and the United States (Bernard et al. 1968; de Sousa and Weigl 1976; de Matos et al. 1995; Speare et al. 1991; Alves de Matos and Paperna 1993; Werner 1993). C. Lucke Tumour Herpesvirus

Luck6 tumour herpesvirus (LTHV) has been reported only from the northern leopard frog, Rana pipiens in the United States (McKinnell and Carlson 1997). Recently LTHV has been classified as Rana herpesvirus 1 (RaHV-1) (Davison et al. 1999) in the family Herpesviridae. Genomic studies indicated that R a w - 1 belongs to the fish virus lineage of the herpesvirus family rather than to the lineage populated by mammalian and avian viruses (Davison et al. 1999). This virus induces renal adenocarcinoma in R . pipiens in the United States; the tumour was well described by Luck6 (1934). Its transmissible nature was recognized in 1938 and a virus was initially suggested as the cause due to the intranuclear inclusions (Luck6 1938), a surmise that was later confirmed.

2978

AMPHIBIAN BIOLOGY

Vertical transmission occurs with eggs becoming infected, but tumours are slow to develop. Clinical signs in adults are bloating, lethargy and death, which only occur when the tumour is large or has metastasized (Anver and Pond 1984). Single or multiple white nodules occur in the kidneys and grow into large masses. The tumour is an infiltrating and destructive adenocarcinoma or, less often, it is orderly and adenomatous (LuckC 19.34). Although the gross appearance of the tumour remains relatively unchanged, there are significant seasonal differences in its microscopic appearance. Winter tumours display cytopathic characteristics associated with the presence of virus (enlarged nuclei with eosinophilic inclusions) whereas those in summer lack virus (McKinnell 1973). Metastasis of the cancer also depends on temperature. Studies have shown that above 22°C virus replication does not occur and viral particles are not present in the tumour (Anver and Pond 1984). Surveys of wild Rana pipiens for the LuckC tumour have found prevalences of up to 12.5% (McKinnell 1969). Since the 1960s, however, the prevalence of Luck6 renal adenocarcinoma in Minnesota has decreased. This is thought to be due to the population declines and reduced density of R . pipiens (McKinnell et al. 1980). D. Herpes-like Virus of Skin In Italy, up to 80% of a wild population of Rana dalmatina had epidermal vesicles associated with a herpes-like virus, but dead frogs were not found (Bennati et al. 1994). E. Calicivirus A calicivirus was isolated from two captive Ceratophrys orata found dead. Both had pneumonia, while one also had oedema and the other had lymphoid hyperplasia (Smith et al. 1986).

E Leucocyte Viruses Polyhedral cytoplasmic DNA virus was found in the cytoplasm of white blood cells of a Mexican Rana catesbeiuna that was lethargic and had small exudative ulcers (Briggs and Burton 1973). The large iridovirus found in red blood cells of Bufo marinus in Costa Rica also was found in the cytoplasm of reticular cells in the spleen (Speare et al. 1991). 111. BACTERIAL DISEASES A. Bacterial Septicaemia ("Red Leg") Bacterial septicaemia in amphibians has been termed "red leg" due to the cutaneous reddening that occurs on frogs' ventral thighs (Emerson and Norris 1905). This is an unfortunate name since many non-pathologists and non-veterinary clinicians appear to think that a frog with any erythema (redness) of the legs has a bacterial disease. It is important to realize that this is a very non-specific sign and it cannot be used to diagnose bacterial septicaemia. Bacterial septicaemia must be diagnosed by a combination of histopathology and bacterial culture. Some massive die-offs in the wild have been attributed to bacterial septicaemia but these diagnoses are dubious due to a lack of histopathological confirmation and testing for other agents, particularly for ranaviruses and Batrachochytrium dendrobatzdis, the amphibian chytrid (Green et al. 2002). Bacteria (including Aeromonas hydrophila) were reported in die-offs in Alvtes obstetricans in the Pyrenees mountains in Spain (Marquez et al. 1995), in R a m muscosa k California (Bradford 1991) in Bufo boreus boreas in Colorado (Carey 1993) and in tadpoles of Ram sylvatica in Rhode Island, USA (Nyman 1986). These bacteria can be cultured from hogs with ranaviral disease and chytridiomycosis, particularly when the animals are collected dead (Cunningham et al. 1996; Berger et al. 1998). They can also be isolated from the destines of healthy amphibians and from the environment (Carr et al. 1976; Hird et al. t-). For example, Aeromoms hydrophila was found in the intestines of 46% (102) of 222 k a i t h leopard frogs (Hird et al. 1983). Bacteria may be present in sick frogs as normal

HEMINGWAY ET AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS

2979

residents, contaminants, or secondary infections. In addition, reddening of the legs can occur in some amphibians showing clinical ranaviral disease and chytridiomycosis (Cunningham et al. 1996; Berger et al. 1999). During a die-off of Yosemite toads (Bufo canorus) in the 1970s, three of 21 histologically examined toads had evidence of acute septicaemia, one of which also had chytridiomycosis, and all of which had been toe-clipped about two weeks before, which might have led to the infection (Green and Sherman 2001). Bacterial septicaemia is, however, a common cause of outbreaks in captive amphibians. A range of bacteria may be involved including Aeromonas hydrophila and other gram-negative bacteria or combinations of bacteria, such as Pseudomonas spp., Proteus spp., Flavobacterium sp. (Hubbard 1981; Taylor et al. 1993; Olson et al. 1992). Recently, a new virulent subspecies, A. hydrophila ranae, was discovered in Rana rugulosa farmed in Thailand and dying from septicaemia (Huys et al. 2003). With bacterial septicaemia, gross pathological signs include pale skin, petechiation, ulcers, lethargy, oedema, ascites, pale livers, and haemorrhages in the internal organs. Histological examination may show degenerative myopathy and multiple foci of coagulative necrosis with clumps of bacteria. Variable results were obtained from transmission experiments - the disease usually required inoculation of the bacteria, or bath exposure and stress (Glorioso et al. 1974; Dusi 1949; Somsiri et al. 1997) although A. hydrophila ranue appeared to be more virulent than were other subspecies (Huys et al. 2003). In most cases, disease probably occurs secondarily to stress caused by poor husbandry such as overcrowding, dirty conditions, trauma, temperature changes, and also after transport (Hubbard 1981; Glorioso et al. 1974). Recent experimental work using A. hydrophila on the host, Xenopus laevis, has shown that the amphibian host's genetics is also important in determining susceptibility to pathogenic bacteria (Barribeau et al. 2008). The survival and growth of X. laevis tadpoles with different major histocompatability complexes (MHC), when exposed to A. hydrophila, depended on their MHC haplotypes with heterozygous tadpoles being intermediate between tadpoles with resistant and susceptible MHC haplotypes. B. Streptococcosis

A non-haemolytic group B Streptococcus caused an outbreak killing 80% of about 100,000 farmed bullfrogs (Rana catesbehna) in Brazil (Amborski et al. 1983). Septicaemia, necrotizing splenitis, and hepatitis with haemorrhages occurred in frogs. Viral cultures were negative. The outbreak was associated with overcrowding and stress. Mortality due to a similar streptococcus occurred in R. catesbeiana being raised for consumption in Uruguay (Mazzoni 2001) and in the United States (Mauel et al. 2002). In the latter instance, the agent was identified as Streptococcus in&, a species that is a pathogen of fish and has zoonotic potential (Lehane and Rawlin 2001). C. Chlamydiosis Chlamydophila pneumoniae caused chronic pneumonia in a wild immunosuppressed frog, Mixophyes iteratus, in Australia (Berger et al. 1999) and in a captive colony of Xenopus tropicalis in the United States (Reed et al. 2000). C. pneumoniae is an important human pathogen. A range of chlamydia1 species infecting healthy amphibians from Switzerland were identified by PCR (Blumer et al. 2007). Outbreaks of chlamydiosis in captive amphibians can result in hlminant, multisystemic infections with pyogranulomatus inflammation, causing moderate to high mortality rates in various species (Wilcke et al. 1983; Howerth 1984; Honeyman et al. 1992). D. Mycobacteriosis

A number of atypical Mycobacterium species infect amphibians; M. tuberculosis or M. bovis have not been reported. The only report of wild amphibians with mycobacteria was M. chelonei

2980

AMPHIBIAN BIOLOGY

subsp abscessus isolated from four of 66 Bufo marinw and two of 86 B. granulosus in a survey of Amazonian amphibians (Mok and Carvalho 1984). None of these animals had histopathological lesions, although experimental intraperitoneal inoculation of 29 toads resulted in the death of five animals from mycobacteriosis. Disease due to mycobacteria has been reported only in captive amphibians and occurs mainly in immunocompromised animals. Several species have been reported. M. marinurn was experimentally shown to cause a chronic granulomatous non-lethal disease in immunocompetent leopard frogs (Rana pipiens) whereas frogs immunocompromised with hydrocortisone developed an acute lethal disease (Ramakrishnan et al. 1997). A M . ulceranslike species and M. 1eiJandii both caused outbreaks in captive Xenopus hevis (Trott et al. 2004; Godfrey et al. 2007) while M. szulgai caused an outbreak in captive X. tropicalis (Chai et al. 2006). Infections primarily involve the skin, respiratory tract, or intestines. Frogs have been found with single, large tumour-like masses or with disseminated nodules throughout the internal organs. Organs such as liver, spleen, kidney, or testes may become almost completely destroyed by the infection before the animal dies, usually with cachexia (Reichenbach-Klinke and Elkan 1965). Early granulomas are composed of mostly epithelioid macrophages, which may progress to form encapsulated foci with dry caseous centres. Granulomas typically contain large numbers of acid-fast bacilli. IV. REFERENCES Alves de Matos, A. P and Paperna, I., 1993. Ultrastructure of erythrocytic virus of the South African anuran Ptychadena anchietae. Diseases of Aquatic Organisms 16: 105-109.

Blumer, C., Zimmermann, D. R., Weilenmanu, R., Vaughan, L. and Pospischil, A., 2007. Chlamydiae in free-ranging and captive frogs in Switzerland. Eterinary Path. 44: 144-150.

Anver M. R. and Pond, C. L., 1984. Biology and diseases of amphibians. Pp 4 2 7 4 4 7 in "Laboratory Animal Medicine", ed by J. G. Fox, B. J. Cohen, and F. M. Loew. Academic Press, New York.

Bollinger, T K., Mao, J., Schock, D., Brigham, R. M. and Chinchar, V. G., 1999. Pathology, isolation, and preliminary molecular characterization of a novel iridovirus from tiger salamanders in Saskatchewan. J. Wildl. Disease 35: 413429.

Ariel E., 1997. Pathology and serological aspects of Bohle iridoviral infections in 6 selected waterassociated reptiles in North Queensland. PhD thesis. James Cook University, Townsville, Australia. Bank, M. S., Crocker, J., Connery, B. and Amirbahman, A., 2007. Mercury bioaccumulation in green frog (Rana clarnitans) and bullfrog (Rana catesbeiana) tadpoles from Acadia National Park, Maine, USA. Environmental Toxicology and Chemistry. 26: 118-1 25. Barribeau S. M., Villinger, J. and Waldman. B., 2008. Major histocompatibility complex based resistance to a common bacterial pathogen of amphibians. PLoS ONE. 3: e2692. Bennati, R., Bonetti, M., Lavazza, A. and Gelmetti, D., 1994. Skin lesions associated with herpesvirus-like particles in frogs (Rana dalmatina). Eterinary Rec. 135: 625-626. Berger, L., 2001. Diseases in Australian Frogs. PhD Thesis. James Cook University, Townsville, Australia. Berger, L., Volp, K., Mathews, S., Speare, R. and Timms, E, 1999. Chlamydia pneumoniae in a free-ranging giant barred frog (Mixophyes iteratw) from Australia. J. Clin. Microbiology. 37: 2378-2380.

Braunwald, J., Tripier, F. and Kirn, A., 1979. Comparison of the properties of enveloped and naked frog virus 3 (FV3) particles. J. General Virology. 45: 673-682. Briggs, R. T and Burton. E R., 1973. Fine structure of an amphibian leukocyte virus. J. Submicroscopic Cytology and Path. 5: 7 1-78. Brunner, J., 2004. Ecology of an amphibian pathogen: Transmission, persistence, and virulence. PhD Thesis. Arizona State University, Tempe, Arizona. Brunner, J. L. and Sesterhenn, T, 2001. Disinfection of Ambystoma tigrinurn virus (ATV). Froglog 48: 2. Brunner, J. L., Richards, K. and Collins J. E, 2005. Dose and host characteristics influence virulence of ranavirus infections. Oecologia 144: 399406. Brunner, J. L., Schock, D. M. and Collins, J. E, 2007. Transmission dynamics of the amphibian ranavirus Ambystoma tigrinurn virus. Diseases o j ~ ~ u a t Organism ic 77: 87-95. Brunner, J. L., Schock, D. M., Collins, J. M. and Davidson, E. W., 2004. The role of an intraspecific reservoir in the persistence of a lethal ranavirus. Ecol. 85: 560-566.

Berna, H. J., 1990. Ecology and life history of the tiger salamander, Ambystom tigrinum nebulosum Hallowell, on the Kaibab Plateau. PhD Thesis. Arizona State University, Tempe, Arizona, USA.

Came, P E., Geerling, G., Old, L. J. and Boyse, E. A., 1968. A serological study of polyhedral cytoplasmic viruses isolated from amphibia. Virology 36: 392-400.

Bernard, G. W, Cooper, E. L. and Mandell, M. L., 1968. Lamellar membrane encircled viruses in the erythmqtes of Rana pipiens. J. Ultrastructure Res. 26: 8-16.

Carey, C., 1993. Hypothesis concerning the causes of the disappearance of boreal toads from the mountains of Colorado. Cons. Biol. 7: 355-362.

HEMINGWAY E T A L : VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS Carey, C., Cohen, N. and Rollins-Smith, L., 1999. Amphibian declines: an immunological perspective. Devel. and Compas Immunology 23: 459-472. Carey, C., Bradford, D. D., Brunner, J. L., Collins, J. P, Davidson, E. W., Longcore, J. E., Ouellet, M., Pessier, A. F! and Schock, D. M., 2003. Biotic factors in amphibian population declines. Pp. 153-208 in "Multiple Stressors and Declining Amphibian Populations: Evaluating Cause and Effect", ed by G. Linder, D. W. Sparling, and S. K. Krest. Society for Environmental Toxicology and Chemistry Press, Pensacola, Florida. Carr, A. H., Amborski, R. L., Culley, D. D. and Amborski, G. F., 1976. Aerobic bacteria in the intestinal tracts of bullfrogs (Rana catesbeiana) maintained at low temperatures. Herpetologica 32: 239-244. Chai, N., Deforges, L., Sougakoff, W., Truffot-Pemot, C., De Luze, A., Demeneix, B., Clement, M. and Bomsel, M.C., 2006. Mycobacterium szulgai infection in a captive population of African clawed frogs (Xenopus tropicalis). J. Zoo and Wldl. Med. 37: 55-58. Chinchar, V G., 2002. Ranaviruses (family Idoviridae): emerging cold-blooded killers. Arch. Krology 147: 447-470. Chinchar VG., Metzger, D. W., Granoff, A. and Goorha R., 1984. Localization of frog virus 3 proteins using monoclonal antibodies. Rrology 137: 21 1-216. Chinchar VG., Wang, J., Carey, C. and Rollins-Smith L., 2001. Inactivation of frog virus 3 and channel catfish virus by esculentin-2P and ranaturin isolated from frog skin. K~ology288: 351-357. Clark, H. F., Gray, C., Fabian, F., Zeigel, R. and Karzon, D. T., 1969. Comparative studies of amphibian cytoplasmic virus strains isolated from the leopard frog, bullfrog, and newt. Pp. 310-326 in "Biology of Amphibian Tumors", ed by M. Mizell. SpringerVerlag, New York.

2981

Cunningham, A. A,, Hyatt, A. D., Russell, P and Bennett, P M., 2007a. Emerging epidemic diseases of frogs in Britain are dependent on the source of ranavirus agent and the route of exposure. Epzdemwlogy and Infection 135: 1200-12 12. Cunningham, A. A,, Hyatt, A. D., Russell, P and Bennett, E M., 2007b. Experimental transmission of a ranavirus disease of common toads (Bufo bufo) to common frogs ( R a m temporaria). Epidemiology and Infection 135: 1213-1216. Cunningham k A, Langton, T E. S., Bennett, P M., Lewin, J. F., Drury, S. E. N., Gough, R. E. and MacGrrgor, S. K., 1995. Investigations into unusual mortalities of the common frog (Rana temporaria) in Britain. Fifth international colloquium o n the Pathology of Reptiles and Amphibians. Alphen aan den Rijn, The Netherlands. Cunningham A. A,, Langton, T. E. S., Bennett, P M., Lewin, J. F., Dmry, S. E. N., Gough, R. E. and MacGregor, S. K., 1996. Pathological and microbiological findings from incidents of unusual mortality of the common frog (Rana temporaria). Philos. Trans. Roy. Soc. Lond. Series B, Biolog. Sci. 351: 1539-1557. Daszak, P, Cunningham, A. A. and Hyatt, A. D., 2001. Draft guidelines for international translocation of amphibians with respect to infectious diseases. Attachment 6 (pp. 150-156) in "Developing Management Strategies to Control Amphibian Diseases: Decreasing the Risks due to Communicable Diseases", ed by R. Speare and the Steering Committee of Getting the Jump on Amphibian Disease. School of Public Health and Tropical Medicine, James Cook University, Townsville, Australia. Daszak, F!, Cunningham, A. A. and Hyatt, A. D., 2003. Infectious disease and amphibian population declines. Divers. and Distrkb. 9: 141-150. Davison A. J., Sauerbier, W., Dolan, A, Addison, C. and McKinnell, R. G., 1999. Genomic studies of the Luck6 tumor herpesvirus (RaHV-1). J. Cancer Res. and Clin. Oncology 125: 132-238.

Collins, J . P., Jones, T. R. and Berna, H. J., 1988. Conserving genetically-distinctive populations: the case of the Huachuca tiger salamander (Ambystoma tigrinum stebbinsi Lowe). Pp. 45-53 in "Management of amphibians, Reptiles, and Small Mammals in North America", ed by R. C. Szaro, K. C. Severson, and D. R. Patton. USDA Forest Service, Rocky Mountain Forest and Range Experiment Station, Fort Collins, Colorado.

de Castro, F. and Boker, B., 1005. Mechanisms of disease-induced extinction. Ecol. LetteTs 8: 117-1 26.

Cullen, B. R. and Owens, L., 2002. Experimental challenge and clinical cases of Bohle iridovirus (BIV) in native Australian anurans. Diseases of Aquatic Organisms 49: 83-92.

Densmore, C. L. and Green, D. E., 2007. Diseases of Amphibians. Institute Lab. Anim. Res. J. 48: 235-253.

Cullen, B. R., Owens, L. and Whittington, R. J., 1995. Experimental infection of Australian anurans (Limnodynastes terraereginae and Litoria latopalmata) with Bohle iridovirus. Diseases of Aquatic Organisms 23: 83-92. Cunningham, A. A. and Langton, T E. S., 1997. Disease risks associated with translocations of amphibians . into, out of, and within Europe - a U.K. perspective. J. Brit. Nterinary Zool. Soc. 2: 37-41. Cunningham, A. A,, Tems, C. A. and Russell, I! H., 2008. Immunohistochemical demonstration of Ranavirus antigen in the tissues of infected frogs (Rana temporaria) with systemic haemorrhagic or cutaneous ulcerative disease.J. Compas Path. 138: 3-1 1.

de Matos, A. E, Papema, I. and Lainson, R., 1995. An erythrocytic virus of the Brazilian tree-frog, Phrynohyas venulosa. Mem6ria.s do Instituto Oswaldo Cruz 90: 653-655.

de Sousa, M. A. and Weigl, D. R., 1976. The viral nature of Eddia Franca, 1912. Mem6rias do Instituto Oswaldo C ~ U74: L 213-230. Drury, S. E. N., Gough, R. E. and Cunningham, A. A., 1995. Isolation of an iridovirus-like agent from common Erogs (Rana temporaria). Veterinary Rec. 137: 72-73. Duffus, A. L. J., Pauli, B. D., Wozney, K., Bmnetti, C. R. and Berrill, M., 2008. Frog virus 3-like infections in aquatic amphibian communities. J. Wildl. Disease 44: 109-120. Dusi, J. L., 1949. The natural occurrence of "red leg", Pseudomonar hydrophila, in a population of American toads, Bufo americanus. Ohio J. Sci. 49: 70-7 1.

2982

AMPHIBIAN BIOLOGY

Emerson, H. and Norris, C., 1905. "Red-leg" - an infectious disease of frogs. J. Experi. Med. 7: 32-58. Erspamer, V., 1994. Bioactive secretion of the amphibian integument. Chapter 9 (pp. 178-350) i n "The Integument", vol. 1 of "Amphibian Biology", ed by H. Heatwole, G. T Barthalmus, and A. Y. Heatwole. Surrey Beatty & Sons, Chipping Norton.

Gray, M. J., Miller, D. L., Schmutzer, A. C. and Baldwin, C. A., 2007. Frog virus 3 prevalence in tadpole populations inhabiting cattle-access and non-access wetlands in Tennessee, USA. Diseases of Aquatic Organism 77: 97-103. Green, D. E., 200 1. Pathology of Amphibia. Pp 4 0 1 4 8 5 in "Amphibian Medicine and Captive Husbandry", ed by K. M. Wright and B. R. Whitaker. Krieger Publishing Company, Malabar, Florida.

Ferrell, S., 2008. Amphibian quarantine guidelines. Pp 63-72 in "Amphibian Husbandry Resource Guide", Edition 1.1, ed by V. Poole and S. Grow. Association of Zoos and Aquariums Amphibian Taxon Advisory Group. http://www.aza.org/ConScience/Documents/ AmphibianHusbandryResourceGuide-v. 1.1.pdf

Green, D. E. and Sherman, C. K., 2001. Diagnostic histological findings in Yosemite Toads (Bufo canorus) from a die-off in the 1970s.J. Herpetol. 35: 92-103.

Fijan, N., Matasin, Z., Petrinec, Z., Valpotic, I. and Zwillenberg, L. O., 1991. Isolation of an iridoviruslike agent from the green frog (Rana esculenta L.). Eterinarski Arhiu Zagreb 61: 151-158.

Green, D. E., Converse, K. A. and Schrader, A. K., 2002. Epizootiology of sixty-four amphibian morbidity and mortality events in the USA, 1996-2001. Ann. New York Acad. of Sci. 969: 323-339.

Fox, S., Greer, A. L., Torres-Cervantes, R. and Collins, J. F!, 2006. First case of ranavirus-associated morbidity and mortality in natural populations of the South American frog Atelognathus patagonicus. Diseases of Aquatic Organisms 72: 87-92. Gahl, M. K. and Calhoun, A. J. K., 2008. Landscape setting and risk of Ranavirus mortality events. Biolog. Cons. 141: 2679-2689. Galli, L., Pereira, A., MBrquez, A. and Mazzoni, R., 2006. Ranavirus detection by PCR in cultured tadpoles (Rana catesbeiana Shaw, 1802) from South America. Aquaculture 257: 78-82. Gantress, J., Maniero, G. D., Cohen, N. and Robert, J., 2003. Development and characterization of a model system to study amphibian immune responses to iridoviruses. Krology 3 11: 254-262. Glorioso, J. C., Amborski, R. L., Amborski, G. F. and Culley, D. C., 1974. Microbiological studies o n septicemic bullfrogs (Rana catesbeiana). Amen J . Eterinary Res. 35: 1241-1245. Godfrey, D., Williamson, H., Silverman, J. and Small, F! L. C., 2007. Newly identified Mycobacterium species in a Xenopus laeuis colony. Compar Med. 57: 97-104. Goorha, R. and Granoff, A., 1974. Macromolecular synthesis in cells infected by frog virus 3:I. Virusspecific protein synthesis and its regulation. Krology 60: 237-250. Goorha, R. and Granoff, A,, 1978. Icosahedral cytoplasmic deoxyriboviruses. Pp 347-399 i n "Comprehensive Virology", vol. 14, ed by H. Fraenkel-Conrat and R. R. Wagner. Plenum, New York. Goorha, R. and Granoff, A., 1999. Frog virus 3 (Iridouirzdae). Pp 582-587 in "Encyclopedia of Virology", ed by R. G. Webster and A. Granoff. Academic Press, San Diego. Granoff, A., 1989. Viruses of Amphibia: an historical perspective. Pp. 3-12 in "Viruses of Lower Vertebrates", ed by W. Ahne and E. Kurstak. Springer-Verlag, Berlin, Germany. Granoff, A,, Came, F! E. and Rafferty, K. A., 1965. The isolation and properties of viruses from Rana pipiens: their possible relationship to the renal adenocarcinoma of the leopard frog. Ann. New York Acad. Sci. 126: 237-255.

Greer, A. L. and Collins, J. F!, 2007. Sensitivity of a diagnostic test for amphibian ranavirus varies with sampling protocol. J. Wildl. Disease 43: 525-532. Greer, A. L. and Collins, J. F!, 2008. Habitat fragmentation as a result of biotic and abiotic factors controls pathogen transmission throughout a host population. J. Anim. Ecol. 77: 364-369. Greer, A. L., Berrill, M. and Wilson, F! J., 2005. Five amphibian mortality events associated with ranavirus infection in south central Ontario, Canada. Diseases of Aquatic Organism 67: 9-14. Greer, A. L., Briggs, C. J. and Collins, J. F!, 2008. Testing a key assumption of host-pathogen theory: density and disease transmission. Oikos 117: 1667-1673. Gruia-Gray, J. and Desser, S. S., 1992. Cytopathological observations and epizootiology of frog erythrocytic virus in bullfrogs (Rana catesbeiana).J. Wildl. Disease 28: 34-41. Gruia-Gray, J., Petric, M. and Desser, S., 1989. Ultrastructural, biochemical and biophysical properties of an erythrocytic virus of frogs from Ontario, Canada. J. Wzldl. Disease 25: 497-506. Harp, E. M. and Petranka, J . W., 2006. Ranavirus in wood frogs (Rana syluatica): Potential sources of transmission within and between ponds. J. Wildl. Disease 42: 307-318. He, J., Lu, L., Deng, M., He, H., Weng, S., Wang, X., Zhou, S., Long, Q., Wang, X. and Chan, S., 2002. Sequence analysis of the complete genome of an iridovirus isolated from the tiger frog. Wology 292: 185-197. Hengstberger, S. G., Hyatt, A. D., Speare, R. and Coupar, B. E. H., 1993. Comparison of epizootic haematopoietic necrosis and Bohle iridoviruses, recently isolated Australian iridoviruses. Diseases of Aquatic Organisms 15: 93- 107. Hird, D. W., Diesch, S. L., McKinnell, R. G., Gorham, E., Martin, F. B., Kurtz, S. W. and Dubrovolny, C., 1981. Aeromonas hydrophila in wild-caught frogs and tadpoles (Rana pipiens) in Minnesota. Lab Anim. Sci. 31: 166-169. Honeyman, V. L., Mehren, K. G., Barker, I. K. and Crawshaw, G. J., 1992. Bordetella septicaemia and chlamydiosis in eyelash leaf frogs (Ceratobatrachus guentheri). Pp 168 i n "Proceedings of the Joint Meeting of tRe American Association of Zoo Veterinarians and American Association of Wildlife Veterinarians". Oakland CA.

HEMINGWAY ET AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS Howerth, E. W., 1984. Pathology of naturally occurring chlamydiosis in African clawed fmgs (Xenopus laevis). Eterinary Path. 21: 28-32. Hubbard, G. B., 1981. Amomonas hydrophila infection in Xenopus laevis. Lab Anim. Sci. 31: 297-300. Huys, G., Pearson, M., Kampfer, P., Denys, R., Cnockaert, M., Inglis, V and Swings, J., 2003. Aeromonas hydrophila subsp. ranae subsp. nov., isolated from septicaemic farmed frogs in Thailand. Internat. J. Syst. and Evol. Microbiology 53: 885-891. Hyatt, A. D., Gould, A. R., Zupanovic, Z., Cunningham, A. A,, Hengstberger, S., Whittington, R. J., Kattenbelt, J. and Coupar, B. E. H., 2000. Comparative studies of piscine and amphibian iridoviruses. Arch. firology 145: 301-33 1. ICTVdB Management, 2006. Index of Viruses Iridoviridae in "ICTVdB - The Universal Virus Database, version 4", ed by C. Biichen-Osmond. Columbia University, New York. Jancovich, J. K., Davidson, E. W., Morado, J. F., Jacobs, B. L. and Collins, J. P., 1997. Isolation of a lethal virus from the endangered tiger salamander Ambystoma tigrinum stebbinsi. Diseases of Aquatic Organism 31: 161-167. Jancovich, J. K., Davidson, E. W., Seiler, A,, Jacobs, B. L. and Collins, J. P., 2001. Transmission of the Ambystoma tignnum virus to alternative hosts. &eases of Aquatic Organisms 46: 159-163. Jancovich, J., Davidson, E. W., Parameswaran, N., Mao, J., Chinchar, V G., Collins, J. P, Jacobs, B. L. and Storfer, A., 2005. Evidence for emergence of an amphibian iridoviral disease because of humanenhanced spread. Molec. Ecol. 14: 213-224. Johnson, A. and Wellehan J., 2005. Amphibian virology. Eterinary Clin. of North America: Exotic Anim. Practice 8: 53-65. Johnson, A. J . and Jacobson, E. R., 2004. Iridovirus infections of turtles and tortoises. 29th Annual Meeting and Symposium of the Desert Tortoise http://www.deserttortoise.org/abstract/ Council. abstracts2004/2004abs27. html Kanachanakhan, S., 1998. An ulcerative disease of the cultured tiger frog, Rana tigrina, in Thailand: virological examination. Aquatic Anim. Health Res. Institute Newsletter 7: 1-2. J. and Hanna N., 2008. Hygiene and Disease Control: Field and Captivity. Pp 53-62 in "Amphibian Husbandry Resource Guide", Edition 1.1, ed by V Poole and S. Grow. Association of Zoos and Aquariums Amphibian Taxon Advisory Group.

kist,

http://www.aza.org/ConScience/Documents/

.4mphibianHusbandryResourceGuide-v.1.1.pdf Langdon, J. S., 1989. Experimental transmission and pathogenicity of epizootic haematopoietic necrosis r i m s (EHNV) in redfin perch, Pmafluuiatilis L., and 11 other teleosts. J. Fish Diseases 12: 295-310.

Lrbane, L. and Rawlin, G. T., 2000. Topically acquired bacterial zoonoses from fish: a review. Med. J. Aust. ' 173: 25G259.

Lrtt. B., 1934. A neoplastic disease of the kidney of the h g , Rana pipiens. Amer: J. Cancer 20: 352-379. -6,

B., 1938. Carcinoma in the leopard frog: Its probable causation by a virus. J. Experi. Med. 63: 457468.

2983

Lynch, M., 2001. Amphibian quarantine protocols Melbourne Zoo. Attachment 6 (pp. 157-161) in "Developing Management Strategies to Control Amphibian Diseases: Decreasing the Risks due to Communicable Diseases", ed by R. Speare and Steering Committee of Getting the Jump on Amphibian Disease. School of Public Health and Tropical Medicine, James Cook University, TownsvdIe, Australia. Maji, S., LaPatra, S., Long, S. M., Sample, R., Bryan, L., Sinning, A. and Chinchar. V G., 2006. Rana catesbeiana virus Z (RCV-Z):a novel pathogenic ranavirus. Diseases ofAquatic Organism 73: 1-1 1. Maniero, G. D., Morales, H., Gantress, J. and Robert, J., 2006. Generation of a long-lasting, protective, and neutralizing antibody response to the ranavirus FV3 by the frog Xenopus. Deuel. and Cornpar: Immunology 30: 649-657. Mao, J., Hedrick, R. P and Chinchar, V. G., 1997. Molecular characterization, sequence analysis, and taxonomic position of newly isolated fish iridoviruses. firology 229: 212-220. Mao, J., Green, D. E., Fellers, G. and Chinchar V G., 1999. Molecular characterization of iridoviruses isolated fro111 sympatric amphibians and fish. Krus Research 63: 45-52. Marquez, R., Olmo, J. L. and Bosch, J., 1995. Recurrent mass mortality of larval midwife toads Alytes obstetricans in a lake in the Pyrenean mountains. Herpetal. J. 5: 287-289. Marsh, I. B., Whittington, R. J., O'Rourke, B., Hyatt, A. D. and Chisholm, O., 2002. Rapid differentiation of Australian, European and American ranaviruses based on variation in major capsid protein gene sequence. Molecular and Cellular Probes 16: 137-1 5 1. Mauel, M. J., Miller, D. L., Frazier, K. S. and Hines, M. E. 2nd., 2002. Bacterial pathogens isolated from cultured bullfrogs (Rana castesbeiuna). J. Eterinary Diagnostic Investigation 14: 431-433. Mazonni, R., 2000. Diseases in farmed American bullfrog (Rana catesbeiana Shaw, 1802) in Uruguay. P. 37 in "Getting the Jump! On Amphibian Disease". Proceedings 26-30 August 2000. McCallum, H. and Dobson, A., 1995. Detecting disease and parasite threats to endangered species and ecosystems. Trends in Ecol. and Evol. 10: 190-194. McCallum, H., Barlow, N. and Hone, J., 2001. How should pathogen transmission be modelled? Trends in Ecol. and Evol. 16: 295-300. McKinnell, R. G., 1969. Luck6 renal adenocarcinoma: Epidemiological aspects. Pp 254-260 in "Recent Results in Cancer Research (Special Supplement), Biology of Amphibian Tumours", ed by M. Mizell. Springer-Verlag, Berlin. McKinnell, R. G., 1973. The Luck6 frog kidney tumor and its herpesvirus. Amer: 2001. 13: 97-1 14. McKinnell, R. G. and Carlson D. L., 1997. Luck6 rrnal adenocarcinoma, an anuran neoplasm: studies at the interface of pathology, virology, and differentiation competence. J. Cellular Physiol. 173: 115-1 18. McKmnell, R. G., Gorham, E. and Martin, F. B., 1980. Continued diminished prevalence of the Luck6 renal adenocarcinoma in Minnesota leopard frogs. Amer: Midland Naturalist 104: 402-404.

2984

AMPHIBIAN BIOLOGY

Miller, D. L., Rajeev, S., Brookins, M., Cook, J., Whittington, L. and Baldwin, C. A., 2008. Concurrent infection with Ranavirus, Batrachochytrium dendrabatidis, and Aeromonas in a captive anuran colony. J . Zoo and Wildl. Med. 39: 445-449. Miocevic, I., Smith, J., Owens, L. and Speare, R., 1993. Ultraviolet sterilisation of model viruses important to finfish aquaculture in Australia. Aust. VeterinaryJ. 70: 25-27. Mok, W. Y. and Carvalho, C. M., 1984. Occurrence and experimental infection of toads (Bufo marinus and B. granulosus) with Mycobacterium chelonei subsp. abscessw. J. Med. Microbiology 18: 327-333. Moody, N. J. G. and Owens, L., 1994. Experimental demonstration of the pathogenicity of a frog virus, Bohle iridovirus, for a fish species, barramundi Lutes calcanir: Dwemes ofAqwltic Organism 18: 95-102. Murti, K. G., Goorha, R. and Chen, M., 1985b. Interaction of frog virus 3 with the cytoskeleton. Cum Epics in Microbiology and Immunology 116: 107-131. Murti, K. G., Goorha, R. and Granoff, A,, 1985a. An unusual replication strategy of an animal iridovirus. Adu. in Krus Res. 30: 1-19. Nyman, S., 1986. Mass mortality in larval Rana syluatica attributable to the bacterium, Aeromonas hydrophila. J. Heretology 20: 196-20 1.

Reed, K. D., Ruth, G. R., Meyer, J. A. and Shukla, S. K. 2000. Chlamydia pneumoniae infection in a breeding colony of African clawed frogs (Xenopus tropicalis). Emerging Infectiow Diseases 6: 196-199. Reichenbach-Klinke, H. and Elkan, E., 1965. Amphibia. Pp. 209-384, in "The Principal Diseases of Lower Vertebrates", Academic Press, New York. Ridenhour, B. J. and Storfer, A. T, 2008. Geographically variable selection in Ambystoma tigrinum virus (Iridouiridae) throughout the western USA. J. Euol. Biol. 21: 1151-1159. Robert, J., Abramowitz, L., Gantress, J. and Morales, H. D., 2007. Xenopus laeuis: a possible vector of ranavirus infection? J. Wildl. Disease 43: 645-652. Robert, J., Morales, H., Buck, W., Cohen, N., Marr, S. and Gantress, J., 2005. Adaptive immunity and histopathology in frog virus 3-infected Xenopus. Eralogy 332: 667-675. Rojas, S., Richards, K., Jancovich, J. K. and Davidson, E. W., 2005. Influence of temperature on ranavirus infection in larval salamanders, Ambystoma tigrinum. Diseases ofAquatic Organism 63: 95-100. Rollins-Smith, L. A,, 1998. Metamorphosis and the amphibian immune system. Immunological Rev. 166: 221-230.

Olson, M. E., Gard, S., Brown, M., Hampton, R. and Morck, D. W., 1992. Flauobacterium indologenes infection in leopard frogs. J. Ame?: Veterinary Med. Assoc. 201: 1766-1770.

Rollins-Smith, L.A., Doersam, J. K., Longcore, J. E., Taylor, S. K., Shamblin, J. C., Carey, C. and Zasloff, M. A., 2002. Antimicrobial peptide defenses against pathogens associated with global amphibian declines. Deuel. and Campal: Immunology 26: 63-72.

Pallister, J., Goldie, S., Coupar, B., Shiell, B., Michalski, W. l?, Siddon, N. and Hyatt, A,, 2007. Bohle iridovirus as a vector for heterologous gene expression. J . ~rologicalMethods 146: 419423.

St. Amour, V and Lesbarreres, D., 2007. Genetic evidence of ranavirus in toe clips: an alternative to lethal sampling methods. Cons. Genet. 8: 12471250.

Parris, M. J., Storfer, A,, Collins, J. E and Davidson, E. W., 2005. Life-history responses to pathogens in Tiger Salamander (Ambystoma tigrinum) larvae. J. Herpetal. 39: 366372.

St. Amour, V., Wong, W. M., Garner, T W. and Lesbarreres, D., 2008. Anthropogenic influence on prevalence of 2 amphibian pathogens. Emerging Znfectzow k e a s e s 14: 1175- 1176.

Pearman, P: B., Garner, T W. J., Straub, M. and Greber, U. F., 2004. Response of the Italian agile frog (Rana latastei) to a Ranavirus, Frog Virus 3: A model for viral emergence in naive populations. J. Wildl Disease 40: 660-669.

Schlaepfer, M.A., Hoover, C. and Dodd, Jr., K., 2005. Challenges in evaluating the impact of the trade in amphibians and reptiles on wild populations. BioScience 55: 256-264.

Petranka, J. W., Murray, S. S. and Kennedy, C. A,, 2003. Responses of amphibians to restoration of a southern Appalachian wetland: Perturbations confound post-restoration assessment. Wetlands 23: 278-290. Petranka, J. W., Harp, E. M., Holbrook, C. T. and Hamel, J. A., 2007. Long-term persistence of amphibian populations in a restored wetland complex. Biolog. Corn. 138: 371-380. Picco, A. M. and Collins, J. P., 2008. Amphibian commerce as a likely source of pathogen pollution. Cons. Biol. 22: 1582-1589. Raffel, T R., Rohr, J. R., Kiesecker, J. M. and Hudson, P: J., 2006. Negative effects of changing temperature on amphibian immunity under field conditions. Functional Ecol. 20: 819-828. Ramakrishnan, L., Valdivia, R.H., McKerrow, J. H. and Falkow, S., 1997. Mycobacterium marinum causes both long-term subclinical infection and acute disease in the leopard frog (Rana pipiens). Infection and Immunity 65: 767-773.

Schock, D. M., 1999. Ecological aspects of the newly identified viral pathogen Regina ranauirus (RRV) in tiger salamanders (Ambystoma tigrinum diaboli). Manuscript. Saskatchewan, Canada. Schock, D. M., Bollinger, T K., Chinchar, T K., Jancovich, J. K. and Collins, J. l?, 2008. Experimental evidence that amphibian ranaviruses are multi-host pathogens. Copeia 2008: 133-143. Smith, A. W, Anderson, M. I?, Skilling, D. E., Barlough, J. E. and Ensley, l? K., 1986. First isolation of calicivirus from reptiles and amphibians. Amer: J. Veterinary Res. 47: 1718-1 72 1. Somsiri. T., Chinabut, S., and Soontornvit, S., 1997. Challenge of cultured frogs (Rana tigerina) with Aeromonas species. Pp 15-18 in "Diseases in Asian Aquaculture 111", Fish Health Section, ed by T W. Flegel, and I. H. MacRae. Asian Fisheries Society, Manila. Speare, R., 1990. Review of the diseases of the Cane Toad, Bufo marinus, with comments on biological control. A w t . Wildl. Res. 17: 387410.

HEMINGWAY E T AL: VIRAL AND BACTERIAL DISEASES OF AMPHIBIANS Speare, R. and Smith, J . R., 1992. An iridovirus-like agent isolated from the ornate burrowing frog Limnodynastes ornatus in northern Australia. Diseases of Aquatic Organisms 14: 5 1-57. Speare, R., Freeland, W. J. and Bolton, S. J., 1991. A possible iridovirus in erythrocytes of Bufo marinus in Costa Rica. J. Wildl. Disease 27: 457462. Speare, R., Berger, L., Skerratt, L.F., Alford, R., Mendez, D., Cashins, S., Kenyon, N., Hauselberger, K. and Rowley, J., 2004. Hygiene Protocol for handling amphibians in field studies. F! 4. Amphibian Diseases Group, James Cook University: Townsville, Australia. www.jcu.edu.aulschooVphtmiPHTM/frogsi field-hygiene.pdf Storfer, A., Alfaro, M. E., Ridenhour, B. J., Jancovich, J. K., Mecb, S. G., Parris, M. J. and Collins, J. l?, 2007. Phylogenetic concordance analysis shows an emerging pathogen is novel and endemic. Ecol. Letters 10: 1075-1083. Taylor, F.R., Simmonds, R.C. and Loeffler, D. G., 1993. Isolation of Flauobacterium meningosepticum in a colony of leopard frogs (Ram pipiens). Lab Anim. Sci. 43: 105.

2985

Williams, T., Barbosa-Solomieu, V. and Chinchar, V. G., 2005. A decade of advances in iridovirus research. Adu. E r w Res. 65: 173-248. Williams T, Chinchar, V. G., Darai, G., Hyatt, A., Kalmakoff, J. and Seligy, V., 2000. Family Iridoviridae. Pp. 167-182 i n "Virus Taxonomy, Seventh Report of the International Committee on Taxonomy of Viruses", ed by M. H. V. Van Regenmortel, C. M. Fauquet, D. H.L. Bishop, E. B. Carsten, M. K. Estes, S. M. Lemon, J. Maniloff, M. A. Mayo, D. J. McGeoch, C. R. Pringle, and R. B. Wickner. Academic Press: San Diego, USA. Willis D. B., Goorha, R. and Chinchar, V. G., 1985. Macromolecular synthesis in cells infected by frog virus 3. Curx Topics in Microbiology and Immunology 116: 77-106. Wolf, K., Bullock, G. L., Dunbar, C. E. and Quimby, M. C., 1968. Tadpole edema virus: a viscerotropic pathogen for anuran amphibians. J . Znfictious Diseases 118: 253-262. Wolf, K., Bullock, G. L., Dunbar, C. E. and Quimby, M. C., 1969. Tadpole edema virus: pathogenesis and growth studies and additional sites of virus infected bullfrog tadpoles. Pp 327-336 in "Biology of Amphibian Tumors", ed by M. Mizell. SpringerVerlag, New York.

Trott, K. A,, Stacy, B. A., Lifland, B. D., Diggs, H. E., Harland, R. M., Khokha, M. K., Grammer, T. C. and Parker, J . M., 2004. Characterization of a Mycobacterium ulcerans-like infection in a colony of African tropical clawed frogs (Xenopw tropicalis). Compar: Med. 54: 309-317.

Wright, K. M. and Whitaker, B. R., 2001. "Amphibian Medicine and Captive Husbandry". Krieger Publishing, Malabar, Florida.

Tweedell, K. and Granoff, A,, 1968. Viruses and renal carcinoma of Rana pipiem. V. Effect of frog virus 3 on developing frog embryos and larvae. J. Nat. Cancer Zmtitute 40: 407410.

Zhang, Q.Y., Li, Z.-Q. and Gui, J.-F., 1999. Studies on morphogenesis and cellular interactions of Rana grylio virus in an infected fish cell line. Aquaculture 175: 185-197.

Weng, S. l?, He, J. G., Wang, X. H., Lu, L., Deng, M. and Chan, S. M., 2002. Outbreaks of an iridovirus disease in cultured tiger frog, Rana tzgrina rugulosa, in southern China. J. Fish Diseases 25: 423-427. Merrier J. K., 1993. Blood parasites of amphibians from Sichuan Province, People's Republic of China. J. Parasitology 79: 356-363.

ltxittington, R. J., Kearns, C. and Speare, R., 1997. Detection of antibodies against iridoviruses in the serum of the amphibian Bufo marinus. J. Virological methods 68: 105-108.

\\'ilcke, B. W., Newcomer, C. E., Anver, M.R., Simmons, J. L. and Nace, G. W., 1983. Isolation of Chlamydia psittaci from naturally infected African clawed toads (Xenopus laeuis). Infection and Immunity 41: 789-794.

Zhang, Q.-Y., Xiao, F., Li, Z.-Q., Gui, J.-F., Mao, J. and Chinchar, V. G., 2001. Characterization of an iridovirus from the cultured pig frog (Rana grylio) ~ + t hlethal syndrome. Diseases of Aqwtic 07gankm 48: 27-36. Zupanovic, Z., Musso, C., Lopez, G., Lounero, C. L., Hyatt, A. D., Hengstberger, S. and Robinson, A. J., 1998b. Isolation and characterization of iridoviruses from the giant toad Bufo marinus in Venezuela. Diseases ofAquatic Organisms 33: 1-9. Zupanovic, Z., Lopez, G., Hyatt, A. D., Green, B., Bartran, G., Parkes, H., Whittington, R. J. and Speare, R., 1998a. Giant toads Bufo marinus in Australia and Venezuela have antibodies against "ranaviruses".Diseases of Aquatic Organisms 32: 1-8.

CHAPTER 2

Fungal Diseases of Amphibians Lee Ber er, Joyce E. Longcore, Rick Speare, Alex Hyatt and Lee F. S erratt

f

I.

Introduction

II.

Chytridiomycosis A. Overview of Chytridiomycosis B. Distribution and Prevalence of Chytridiornycosis in Relation to Amphibian Declines 1. Australasia 2. Latin America 3. North America 4. Europe 5. Asia 6. Africa C. Taxonomy 1. The Chytridiomycota 2. Batrachochytrium dendrobatidis (Amphibian Chytrid) D. Biology 1. Life Cycle 2. Nutrition and Saprobic Growth 3. Temperature Tolerance 4. pH Tolerance 5. Tolerance to Salts 6. Desiccation E. Chytridiomycosis: The Disease 1. Mortality Rates and Incubation Times 2. Clinical Signs 3. Pathology 4. Distribution of Sporangia 5. Chytridiomycosis in Tadpoles F. Epidemiology 1. Seasonality and Thermal Effects 2. Host Range and Effects on Different Species of Amphibians 3. Effect of Chytridiomycosis on Wild Amphibian Populations G. Resistance to Infection 1. Individuals 2. Populations

H. Transmission and Spread of Batrachochytrium dendrobatidis I. Diagnosis of Chytridiomycosis 1. Sampling 2. Identification of Batrachochytrium dendrobatidis J. Management 1. Disinfection 2. Treatment of Chytridiomycosis 3. Quarantine Guidelines 4. Control Options K. Discussion 1. Chytridiomycosis and Amphibian Declines 2. Chytridiomycosis in the Context of Other Introduced Diseases of Wild

Animals and Plants Oomycoses A. Introduction B. Taxonomy C. Biology D. Oomycete Diseases of Larval Amphibians E. Oornycete Diseases of Amphibian Eggs 1. Pathogenesis 2. Epidemiology F. Population Effects of Oomycete Infection of Eggs G. Diagnosis of Oomycoses H. Management IV. Other Fungal Diseases A. Mucormycosis B. lchthyophonosis C. Chromomycosis D. Amphibiocystidium

Ill.

V. VI.

Acknowledgements References Appendices

Abbreviations and acronyms used in the text or references: DDAC = didecyl dimethyl ammonium chloride; MAb = monoclonal antibody; OIE = World Organization for Animal Health; PAb = polyclonal antibody; PAS = periodic acid-Schiff stain; PCR = Polylmerase Chain Reaction; ELlSA = Enzyme-linked immunosofbent assay

BERGER E T AL: FUNGAL DISEASES O F AMPHIBIANS

2987

I. INTRODUCTION

T

HE largest section of this chapter reviews chytridiomycosis, which has caused the decline and extinction of a multitude of amphibian species worldwide, and is the most devastating disease of wildlife on record. Oomycete infection of eggs in the United States has also been linked to localized amphibian declines and is reviewed in detail. Diseases due to Mucor amphibiorum and Zchthyophonus sp. have caused mortality in wild amphibians and are described in the last section although they have not been associated with population declines. Fungal diseases known only from captive amphibians, or fungal infection of wild amphibians not associated with disease, are not described. For a comprehensive description of fungal diseases in amphibians, including pathology and treatment, see Wright and Whitaker (2001). Although systemic fungal infections in mammals are usually associated with immunosuppression, many fungi are common primary pathogens in aquatic animals such as fish, crustaceans and amphibians (Reichenbach-Klinke and Elkan 1965). 11. CHYTRIDIOMYCOSIS A. Overview of Chytridiomycosis

Chytridiomycosis is a fungal skin disease that has caused mass mortality in amphibians at sites worldwide (Berger et al. 1998; Bosch et al. 2001; Muths et al. 2003). It emerged in the 1970s after being introduced to Australia and the Americas. Africa is a possible origin (Weldon et al. 2004). Chytridiomycosis is caused by the chytrid fungus, Batrachochytrium dendrobatzdis, described in 1999 (Longcore et al. 1999). In Australia, chytridiomycosis was the cause of extinction of one species, the sharp-snouted dayfrog (Taudactylus acutirostris) and suspected to have caused the extinction of at least three other species (Speare et al. 2001). Chytridiomycosis was listed as a Key Threatening Process for Australian amphibians in July 2002. Severe population declines associated with outbreaks of chytridiomycosis have also occurred in Central America, South America, the United States and Spain (Berger et al. 1998; Lips et al. 1999; Bosch et al. 2001; Muths et al. 2003). Genetic and epidemiological studies show that chytridiomycosis is an emerging infectious disease that has spread globally (Fig. 1) (Laurance et al. 1996; Daszak et al. 1999; Morehouse et al. 2003). It caused epidemic waves of high mortality as it spread through na'ive populations. Once introduced, it persists and acts as an endemic pathogen. If populations survive, their mortality rate appears to be reduced suggesting there is selection for resistance (Retallick et al. 2004; McDonald et al. 2005). Subsequent mortality appears to be the result of interaction between the frogs and environment, particularly temperature (Berger 2001; Woodhams et al. 2003; Berger et al. 2004; McDonald et al. 2005). Prevalence in healthy rainforest frogs post-decline has been reported to be about 7%-28%, using histologic diagnosis on toe clips (Retallick et al. 2004; McDonald et al. 2005), which is an underestimation of true prevalence given the low sensitivity of this test (Boyle et al. 2004). During many of the catastrophic amphibian population declines that have occurred since the 1970s, mortality was so rapid it was not observed; populations simply disappeared and environmental problems were suspected. It was not until disease studies were initiated in Australia in the 1990s by Rick Speare and Keith McDonald that chytridiomycosis was discovered in wild frogs (Berger et al. 1998); hence, determining chytridiomycosis as a cause of declines has been retrospective. Apart from a few cases where studies of mortality were undertaken as the disease first arrived, linking chytridiomycosis to declines has been achieved by the testing of archived museum specimens for the pathogen, by examining the current effect of chytridiomycosis, and by considering the epidemiology of the declines. Knowledge concerning amphibian chytridiomycosis is increasing rapidly; the present review has attempted to be comprehensive to mid-2005.

2988

AMPHIBIAN BIOLOGY

Fig. 1. Map of the world with countries coloured if chytridiomycosis has been detected in wild amphibians. An update on the global distribution is available at the Amphibian Disease Home Page (http://www.jcu.edu.au/schooU phtmiPHTM/frogs/ampdis.htm).

B. Distribution and Prevalence of Chytridiomycosis in Relation to Amphibian Declines 1 . Australasia

Dramatic extinctions and declines in Australian frogs have occurred since the 1970s, mostly in protected mountainous areas. Although in many lowland and populated areas habitat loss, environmental degradation and introduced fish have reduced the distribution and abundance of amphibians, these factors are not associated with the disappearance of highland frogs (Gillespie and Hero 1999; Hines et al. 1999). Declining frogs are from ten genera and most are from Queensland, New South Wales and Victoria (Tyler 1997). At least four species from Queensland that only occurred at high elevation have become extinct, including the only two species of gastric brooding frog. At least ten other species in Queensland have declined; some have lost upland populations but persist in the lowlands (McDonald and Alford 1999). The declines in Queensland occurred in 1979-1981 in the southern part of the state, in 1985-1986 in the east-central part, and in 1990-1995 in the far north, a spread of about 100 kmlyear (Laurance et al. 1996; McDonald and Alford 1999). The evidence now available indicates that chytridiomycosis was introduced to Australia and has spread through amphibian populations since the 1970s. Chytridiomycosis is a waterborne disease of high virulence in some species in adults (but not in tadpoles) and occurs at higher prevalence at high elevations (McDonald et al. 2005; Woodhams and Alford 2005). A number of events in protected areas of Queensland are consistent with the introduction of Batrachochytrium dendrobatidk as the cause of declines. These include: (1) sudden, severe declines over a few months at individual sites, (2) asynchronous declines that spread as a front along the eastern coast of Australia, (3) mortality in adults and recently metamorphosed froglets but not in tadpoles, (4) absence of any detected recent environmental changes, (5) disappearance only of stream-associated frogs, (6) disappearance of populations from high elevations, and (7) at one intensively monitored site, mass mortality

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

2989

coincident with a significant population decline. This mass mortality occurred in 1993 at the montane rainforest location of Big Tableland in north Queensland (Laurance et al. 1996, 1997; McDonald and Alford 1999) and chytridiomycosis was diagnosed in dead frogs (Berger et al. 1998; McDonald et al. 2005). The affected frogs were of the endangered species Taudactylus acutirostris, Litoria rheocola and L. nannotis (Laurance et al. 1996). At the time of the declines, healthy tadpoles of ?: acutirostris were collected to be raised in captivity at the Royal Melbourne Zoo and other institutions; however, those that metamorphosed died with chytridiornycosis (Banks and McCracken 2002). ?: acutirostris has since been listed as extinct. The first case of chytridiomycosis detected (retrospectively) in Australia occurred in 1978 (Speare and Berger 2005a) in southeastern Queensland, just before the populations of the southern gastric brooding frog (Rheobatrachus silus) and the southern dayfrog (Taudactylus diurnus) declined and disappeared (McDonald and Alford 1999). These species were likely to have been eliminated by chytridiomycosis but dead frogs were not available for examination. Chytridiomycosis has been found in amphibians from four geographic areas: a large eastern coastal zone extending from Big Tableland near Cooktown in the north to central Victoria in the south, an Adelaide zone, a southwestern zone that includes southwestern Western Australia to just north of Perth and a recently reported occurrence in Tasmania (Speare et al. 2001; Retallick 2003; Berger et al. 2004; Obendorf 2005; Speare and Berger 2005a). In eastern Australia cases have occurred at high and low elevations on or between the Great Dividing Range and the coast. To some extent this distribution also matches searching intensity, although surveys for chytridiomycosis in Cape York and in the Northern Territory have been negative (Mendez, Speare, McDonald and Freeland unpubl. data). One hundred and twenty amphibians tested from Northern Territory in 1999 were free of chytridiomycosis (Speare et al. 2001) and 580 amphibians from 15 species in the Ord region of northern Western Australia tested in 2004 were also free (Speare, unpubl. data). Retallick (2003) analysed the distribution of the disease in relation to temperature and r a i n M using the software BIOCLIM (Nix 1986). Results from testing sick and healthy frogs were included. The predicted distribution closely resembles the observed distribution. The model predicted that Tasmania was a suitable habitat for Batrachochytrium infection (Retallick 2003) and a subsequent survey revealed chytridiomycosis there (Obendorf 2005). B. MmbrrAdk occurs in areas that experience diverse rainfall and temperature. The area of Australia where mean summer temperatures were below 26°C matched the area predicted for the occurrence of B. dendrobatidis. Inland regions and coastal areas north of Cooktown appear to be too hot in summer and too dry in winter to support B. dendrobatidis. Heat and desiccation appear to limit the spread of chytridiomycosis, as suggested by studies on sunival of the hngus (see section D). Remnant populations of the declining species Litoria aurea in Kew South Wales persist in coastal areas with high salinity and at sites contaminated with heavy metals (e.g., a gold mine, copper smelter and tannery), suggesting that these metals and salts may have antifungal effects on B. dendrobatzdis (R. Wellington, pers. comm. 2001; Department of Environment and Conservation New South Wales 2004). In Australia, B. dendrobatzdis appeared suddenly and then expanded its geographic range. Based on current records (Speare and Berger 2005a), the Australian epidemic appears to have commenced in southeastern Queensland in the late 1970s and extended north and south along the eastern coast. In Queensland, 110 samples from museum specimens collected prior to the 1978 appearance tested negative (Speare et al. 2001). Chytridiomycosis in Western Australia seems to have commenced south of Perth in 1985 and subsequently spread in all directions, currently occurring over a wide area of southwestern Western Australia (Aplin and Kirkpatrick 2000). Over 4 000 museum and other samples were tested from Western Australia and the earliest infection was identified in museum specimens that

2990

AMPHIBIAN BIOLOGY

were collected in 1985 near the southern coast. The fungus seems to have arrived in Perth in 1989. All 612 samples collected between 1952 and 1984 tested negatively (Caplin and Kirkpatrick 2000). In a survey of ill and dead amphibians from eastern Australia between October 1993 and December 2000, severe chytridiornycosis was the most common disease and was' the cause of death or morbidity for 133 (55.2%) of 241 free-living amphibians and for 66 (58.4%) of 113 captive amphibians (Berger 2001; Berger et al. 2004). Twelve percent of these frogs had concurrent diseases. Few frogs (5.8%) that were diagnosed with other major diseases were lightly infected with B. dendrobatidis, further supporting the observation that chytridiornycosis was a primary disease. The prevalence of infection, as determined by histology, in 1 578 apparently healthy frogs (L. nannotis, L. genimuculata, L. rheocola and Nyctimystes duyi) in the Wet Tropics of northern Queensland between 1998 and 2002 was 7.1% (McDonald et al. 2005). The four species had a similar prevalence. Declines had occurred in this area 5-9 years previously. In winter, the prevalence at different elevations was similar (between 11.2 and 13.1%) but in summer elevation exerted a large effect and prevalence was 2.3% at <300 m, 4.7% at 300-600 m and 6.5% at >600 m (McDonald et al. 2005). A similar survey at Eungella National Park in east-central Queensland between 1994 and 1998 where declines had occurred in 198511986, revealed an overall prevalence of 15% in 474 samples of six species (Retallick et al. 2004). In Taudactylus eungellensis, a species that had declined, the prevalence was 18.4%. Infection levels in males, females and subadults did not differ within a given species, but did vary among species. Infection rates did not vary from year to year, suggesting that chytridiomycosis is now a more stable infection in these populations ten years following the declines (Retallick et al. 2004). In New Zealand, chytridiornycosis was first diagnosed in introduced Australian Litoria raniformis collected in November 1999 (Waldman et al. 2001), and was detected in the rare native species Leiopelmu archeyi in 2001 (Bell et al. 2004). Both species have been declining. Declines in L. archeyi occurred from 1996 in Coromandel and chytridiornycosis is considered a possible cause (Bell et al. 2004). In November and December 1999, sick and dying L. raniformis were collected from a pond near Christchurch. Large numbers of metamorphs died with chytridiornycosis. Although hundreds of adult frogs in the pond appeared healthy, only six returned to the pond the following summer, possibly because chytridiornycosis killed them during overwintering. The pond where chytridiornycosis was first identified was a source of tadpoles and frogs for the pet trade. The fungus may have been introduced to the pond by a collector, and could then have been spread around the country in animals distributed for pets (Waldman et al. 2001). 2. Latin America

Declines in Central and South America occurred in a pattern similar to that in Queensland, with mysterious, rapid disappearances of montane, stream-associated populations of a variety of species from the 1970s onward (Lips 1998, 1999; Young et al. 2001). Amphibians have declined in at least 13 countries and 40 species are thought to have been extirpated in a country where they once occurred, or to have become extinct altogether (Young et al. 2001). Sudden declines, during which populations disappeared within three years, were observed in Costa Rica, Panama, Venezuela and Ecuador (Young et aZ. 2001). Declines occurred above 500 m in Central America and above 1 000 m in the ,hdes. Declines have occurred asynchronously and a possible generalized spread southwards through the United States to Mexico, Costa Rica and Panama has been noted, consistent with movement of a disease (Lips 1998; Lips et al. 2003a, 2004). Tadpoles were abundant during declines in Costa Rica (Lips 1998). Surveys of h g s in southern Mexico in 2000 showed that 31 populations, representing 24 species, had been extirpated and 11 endemic species may have become extinct (Lips et

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

2991

al. 2004). Riparian species of Hylidae, Centrolenidae and Eleutherodactylus above 800 m elevation were missing from relatively intact habitats. One dead and one ill frog, both with chytridiomycosis, were collected and 19% of the tadpoles from three regions had abnormal mouthparts suggestive of chytridiornycosis. Most declines occurred in the mid to late 1970s and early 1980s. This finding supports the hypothesis that B. dendrobatzdis moved southward from western North America (declines in the 1970s and early 1980s) through Mexico to lower Central America (declines in the mid 1980s and 1990s) (Lips et al. 2004). The abrupt disappearance of Bufo periglenes (golden toad) and decline of Atelopus vamus in Costa Rica's Monteverde Cloud Forest in 1986-1987 suggested high adult mortality rather than just a lack of successhl recruitment (Pounds and Crump 1994). The unusually warm, dry conditions in 1987 caused the frogs to cluster in moist habitats. It is suggested that this clustering of the population may have precipitated an epidemic by increasing transmission (Pounds and Crump 1994; Pounds et al. 1999). All four species of Atelopus in Costa Rica appear to be extinct and recent work led to the detection of chytridiornycosis in two preserved specimens of A. varius collected in 1986 and 1992 at sites south of Monteverde (Puschendorf 2003). In 1993-1994 dead and dying anurans were found before a decline was noticed at Las Tablas, 250 km south-east of Monteverde, near the border with Panama, and chytridiornycosis was diagnosed in dead frogs (Lips 1998; Lips et al. 2003a). Between 1993 and 1998 about half the species at Las Tablas disappeared (Lips et al. 2003a). Chytridiomycosis was detected during mass mortality and declines at Fortuna, Panama in 1996-1997 (Berger et al. 1998; Lips 1999). Fifty-four frogs from ten species were found dying. Anurans had been abundant in 1993-1995 but were rare by July 1997 (Lips 1999). Atelopus cruciger in Venezuela declined in the mid 1970s and early 1980s, and the last two specimens were collected in 1986. Environmental changes do not appear related to these declines. Chytridiomycosis was found in one of the last individuals caught and this disease is thought to be a possible cause of the decline (Bonaccorso et al. 2003). Chytridiomycosis occurred in 96% (46148) of apparently healthy Rana catesbeiana collected near a pond at 2 370 m elevation in the state of MCrida in 2002 (Hanselmann et al. 2004). -41 except two of the infected animals had light infections of B. &ndrobatzdis. Rana catesbeiana was introduced in the late 1990s and the extremely high prevalence in healthy animals suggests they could be an effective reservoir host (Hanselmann et al. 2004). The earliest case of chytridiornycosis found in Ecuador is from December 1980, which is before declines were first noted (in the late 1980s) (Ron and Merino 2000). Infected specimens were found throughout the Andes between 3 100 m and 4 000 m and chytridiornycosis is believed to be a factor in the declines. Dramatic population crashes occurred in stream-dwelling frogs in high-elevation rainforest in Brazil in 1979, and although mass mortality was not observed, it was suspected to have occurred (Heyer et al. 1988; Weygoldt 1989). Tadpoles did not appear to survive metamorphosis in the wild and although in 1975 tadpoles were easily reared in captivity, in 1987 most died during metamorphosis (Weygoldt 1989). Suggested causes of these declines included airborne pollution, disease, and climatic change secondary to loss of forest (Heyer et al. 1988; Weygoldt 1989). On bullfrog farms in Uruguay, chytridiomycosis is commonly seen in healthy animals (Mazzoni et al. 2003). Farms often have outdoor ponds and wild frogs may enter, so that nansmission between captive and wild amphibian populations is possible. In Argentina, dead adult hptodactylus ocellatus were detected in July 2002 and four were diagnosed with chytridiornycosis, although in three of them infection was considered to be slight (Herrera et al. 2005).

3-*\Tdh America North America has experienced declines caused by environmental degradation and pollution (Bonin et al. 1997; Alford and Richards 1999), but there have also been mysterious

2992

AMPHIBIAN BIOLOGY

crashes of montane populations. Waves of mass mortalities, described as the "postmetamorphic death syndrome", have been reported in various amphibian populations in western North America (Scott 1993; Muths et al. 2003). Deaths often occnrr-ed after cold weather and a novel disease that spread through watersheds was suggested as the cause (Scott 1993). Examples include Rana turahumarae in Arizona (mid 1980s), Bufo canorus (1976 1982) and R. muscosa (1979-1990) in the Sierra Nevada, R. chiricahuensis in New Mexico and B. boreas in Colorado (Kagarise-Sherman and Morton 1993; Scott 1993). In the Sierra Nevada, a range of species declined in relatively undisturbed, protected areas (Drost and Fellers 1996).Windborne pesticides have been linked to these declines (Davidson et al. 2002). The first case of chytridiornycosis detected in the United States was from 1974 in a Rana pipiens from Colorado (Carey et al. 1999). Green et al. (2002) suggested introduction of Batrachochytrium dendrobatidis as a leading candidate for the cause of many unexplained declines in western United States. Chytridiomycosis has been found in 13 mainland states and in two provinces in Canada (Green et al. 2002; Carey et al. 2003; Speare and Berger 2005b). In an extensive survey of diseased amphibians in the United States, chytridiornycosis was found to be the cause of seven of 44 mortality events (Green et al. 2002). Although outbreaks were insidious and easily overlooked because few dead animals were found on any given day, populations were in decline. In many populations numbers of breeding adults were reduced by >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).

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

2993

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.

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

2995

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

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

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).

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

2999

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).

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3001

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

,

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

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3015

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

3016

AMPHIBIAN BIOLOGY

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

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3017

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;

3018

AMPHIBIAN BIOLOGY

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

BERGER E T AL: FUNGAL DISEASES OF AMPHIBIANS

3019

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;

3020

AMPHIBIAN BIOLOGY

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

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3021

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.

3022

AMPHIBIAN BIOLOGY

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 <4"C is preferable (Hyatt, unpubl. data). Great care is needed to prevent cross-contamination between samples because the PCR assay detects tiny amounts of DNA that may remain even after sterilization. New gloves for handling an animal and new instruments for collecting a sample must be used. D. SAMPLING POPULATIONS AND INDMDUALS

Populations: Sampling of animals is a complex procedure. For example, if populations are to be surveyed for the presencelabsence of B. dendrobatzdis then any of the techniques will yield pertinent information provided the correct number of animals is sampled. To determine whether B. dendrobatzdis is present or absent in a population, a sampling protocol must be used that will detect the lowest level of infection expected with a high degree of likelihood. For example, to achieve a 95% probability of detection, 149 animals must be tested to detect one infected animal in a population that has a 2% infection rate, assuming random sampling (DiGiacomo and Koepsell 1986). To process this number of samples the applied assay must have the capacity for accommodating large numbers in a short period of time (high throughput), such as real-time TaqMan PCR.

More than one positive test per frog population may be needed and this will depend on the estimated prevalence of chytridiomycosis and the false positive error rate of the test. To optimize the chance of detecting infection, sampling should be planned for a time when frogs are readily available and temperatures are below 2'7°C. Individuals: Sampling of individual animals invokes a different set of rules. Although the different assays vary in sensitivity and specificity, false negatives may occur with any test if the sampling procedure misses the site of infection or if the epidermis has sloughed just before sampling (Boyle et al. 2004). Therefore, to increase the accuracy of diagnosis of a single animal when there is a high suspicion of chytridiomycosis, samples should be cdlected at least a second time if results from the first sample are negative.

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3023

2. Identzfication of Batrachochytrium dendrobatidis A. SKIN SCRAPINGS AND SMEARS

Examination of skin scrapings or smears by light microscopy is a quick and simple method of diagnosis and can be done on fresh, frozen or fixed samples. With some practice, accuracy of diagnosis in frogs with severe chytridiomycosis is similar to that when using histology (Berger 2001) and suitable samples are easily obtained from such frogs. Shedding skin is lifted or scraped from the frog, using a scalpel or sterile plastic spoon (Berger 2001; Briggs and Burgin 2003), spread out flat on a slide with a drop of water, covered by a cover-slip and the preparation examined under a compound light microscope. Ideally, an even monolayer of keratinized epidermal cells is obtained. Magnification of lOOx is used initially to scan a section, then 400x is used to confirm the presence of sporangia. The refractile walls of the sporangia are more distinctive if the condenser is racked down. The round to oval intracellular sporangia (5-13 pm) occur in clumps. Old empty sporangia are the most prevalent stage in shedding skin although sporangia containing zoospores are commonly found (Figs 39, 40). Discharge tubes usually point perpendicularly to the skin's surface and thus appear as small circles that can be difficult to discern. The observation of internal septa within sporangia increases confidence in the diagnosis. Epidermal cell nuclei are of similar size to sporangia but can be differentiated by their irregular, indistinct membranes and flat, granular, grey appearance. Stains: Diagnosis of chytridiomycosis by staining skin scrapings or smears has also been described. A 1: 1 mixture of cotton blue (Parker ink) and 10% aqueous KOH is an effective stain in wet preparations (Mazzoni et al. 2003). Congo-red dye stains chitinous components of B. dendrobatzdis (Briggs and Burgin 2003). Following 20 to 30 minutes staining with 0.01% Congo-red, the walls of empty sporangia and exposed discharge tubes stain brick-red. After 45-60 minutes the walls of most immature, mature and empty sporangia are stained; zoospores are not stained by this procedure (Briggs and Burgin 2003). Epidermal cell nuclei stain pale orange with Congo-red if cells are damaged. DipQuick (Jorgensen, U. S. A.) was

?

I

Right:

Fig. 40. Shedding skin from an Infected Mucophyes fasczolatus Note colonlal sporangla wlth Internal septa dlv~dlngthe thallus Into two or four compartments (arrows). Bar = 15 pm.

k

I

Fig. 39. Unstained wet mount of shedding slun from an ~nfectedadult Litoria caerulea. Note refractile round and oval sporangia. Most sporangia are empty but one contains developing zoospores (arrow). Bar = 30 pm. E = epidermal cell. From Berger et al. (1999).

3024

AMPHIBIAN BIOLOGY

used to stain dried smears resulting in staining of cytoplasm, zoospores and walls, as well as host nuclei (Nichols et al. 2001). These stains may improve accuracy and ease of diagnosis, but comparisons have not been carried out. B. WET PREPARATION O F WHOLE SKIN

Samples of full-thickness skin, from webbing or elsewhere, can be examined unstained (Longcore, unpubl. data). This technique maintains the skin's anatomy and a large surface area can be examined. The advantage of this technique is that the sample can be orientated and the location of the suspected agent can aid in identification. For example, it can be ascertained whether suspected fungal profiles are within superficial cells and thus indicative of B. dendrobatidis or whether they are in deeper layers and thus profiles normal amphibian morphology. This technique is quick, inexpensive and, when used by skilled observers, is equivalent in sensitivity to staining with haematoxylin and eosin. It is useful in studying healthy frogs from which sheets of shedding skin cannot be obtained. Batrachochytrium dendrobatzdis is routinely identified in fresh mounts before attempting to isolate the fungus into pure culture. Tadpoles - Wet Preparation: Likely infected hosts can often be identified in the field by the loss of colour on the jaw sheaths, which can be seen with the aid of a hand lens at 10X magnification (Fellers et al. 2001; Rachowicz 2002; Rachowicz and Vredenberg 2004). Tadpole mouth-parts can then be examined by cutting off pieces of the tooth-rows or jawsheaths and squashing them under a cover-slip (Fig. 41). C. HISTOLOGY

Histological sections are prepared from tissue preserved in 10% formalin or 70% ethanol, then dehydrated, embedded in paraffin, sectioned at 5 pm and stained with haematoxylin and eosin (Drury and Wallington 1980). A vertical section through the skin is achieved. Digits are examined by sectioning a whole foot ventral-side down or by sectioning a single toe. For toes, the maximum length of stratum corneum is obtained from a longitudinal section rather than from a cross section. Digits are decalcified in EDTA for 48 hours at 37OC or in 10% formic acid for 3-5 days before processing (Berger et al. 2000). Alternatively with larger digits, for example from amphibians with a snout-vent length > 60 mm, it is possible to remove skin from the underlying phalanx and section the skin without bone.

1

-

Fzg 41 Unstained squash preparation of pigmented keratlnlzed tooth rows from an infected tadpole of Mzxophyes fasczolatus. The arrows ind~cate clusters of sporangla of Batrachochytrzum dendrobatzdzs Bar = 80 pm.

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3025

In the stratum comzeum the chytrid is spherical or oval with discharge papillae projecting from the surface (Fig. 42). Discharge papillae can be seen in histological sections, but are not common. Zoospores that develop in the zoosporangium escape through the open discharge tube. The wall of the zoosporangium is smooth, uniform in thickness and usually stains eosinophilic. The contents of the zoosporangia vary with the developmental stage of the chytrid; four stages can be identified: (1) The earliest stage contains a central basophilic, rather homogenous mass. (2) Zoosporangia become multinucleate and then the cytoplasm divides to form zoospores. Zoospores are basophilic and appear in cross-section as round or oval bodies (Fig. 42), usually numbering about 4-10 depending on the plane of the section. (3) Once the zoospores are released via the discharge papilla, the empty zoosporangia remain. In some empty colonial stages, thin septa are visible dividing the sporangium into internal compartments. (4) The empty sporangium may collapse into an irregular shape (Fig. 43). During this terminal stage the empty shell sometimes becomes colonized by bacteria and these are seen in section as basophilic rods or cocci (Fig. 43). Empty sporangia are the most common stage present in the sloughing surface layer (Berger et al. 2000). In histological sections the diameter of zoosporangia varies from 5 to 13 pm. They are a similar size to epidermal cell nuclei. Discharge tubes have a diameter of 2 pm and a variable length, usually between 2 and 4 pm, but sometimes as large as 10 pm. Zoospores are about 2 pm in diameter (Berger et al. 2000). Infection is usually associated with skin pathology and these changes can be used to detect, at low magnification, areas likely to be infected. Focal hyperkeratosis and erosions are common in the area adjacent to the organisms. Irregular thickening of the epidermis (hyperplasia) may be present. In some fatal cases extensive sloughing of the hyperkeratotic layer leaves the epidermis with few organisms (Fig. 44). In these cases, however, chytrids can be detected in low numbers in the slightly keratinized surface layer or may be seen in large numbers in the sloughed skin. Sporangia are not present in areas of extensive ulceration.

Fig. 42. Section of skin from a heavily infected adult Litoria caerulea. Note homogenous immature stage (I), larger multinucleate stages, zoosporangium with discharge tube (D) containing zoospores, and empty zoosporangium after zoospores have discharged (arrow). E = epidermis. Stained with hematoxylin and eosin.

3026

AMPHIBIAN BIOLOGY

Fzg 43

Sectlon of skm from a juvenlle Mzxophyes Note empty collapsing sporanglum (arrow) and one contalnlng bacterla (B) A sporanglum is dlvlded by an ~nternalseptum (S) Stalned wlth hematoxylin and eosm

fasczolatus wlth mostly empty sporangla

w CTM

A

!?

. -.. ,-:,

+, f*;

"4

Occasionally other fungi invade the epidermis of amphibians, for example cutaneous mucormycosis (Taylor et al. 1999b). Most have thread-like hyphae but in cross-section the hyphae can often have a circular appearance, and if the number of planes of sections are few, the inexperienced examiner may confuse hyphae with sporangia.

Ducts from dermal glands often appear as spherical spaces between epidermal cells and may be confused with empty zoosporangia although they lack a distinct and complete wall, and are extracellular (Fig. 45) (Berger et al. 2000). The basophilic immature stages of the chytrid in the subsurface layer can appear similar to epidermal cell nuclei but are often surrounded by a clear halo. I Special stains demonstrate the fungal wall around immature chytrids.

u

-4 .I..

eT .32

c t,.

+

Tadpoles - Histology: In tadpoles, a section through the mouthparts including the dark brown keratinized jaw sheaths or tooth rows is required for diagnosis. Large tadpoles are cut longitudinally through the midline with a scalpel and embedded with the cut surface downwards. Small tadpoles are best embedded whole on their side in the paraffin block, and then serially sectioned to reach the mouthparts. The size of the mouth varies among species, which affects the ease of obtaining a suitable section. The usual range of hngal stages may be present in tadpole mouthparts. They can occur on all surfaces of the jaw sheaths and on the caudal surfaces of the tooth rows (Marantelli et al. 2004). In experimentally infected tadpoles, sporangia were detected by histology before gross mouthpart abnormalities could be seen, at three weeks post-exposure (Rachowicz and Vredenberg 2004). Serial sections are recommended for detecting light infections (Rachowicz and Vredenberg 2004). Special Histological Stains: Special fungal stains, such as periodic acid-Schiff (PAS) or silver stains (Drury and Wallington 1980; Swisher 2002) may be used to highlight sporangia. With optimal silver staining, sporangia (black) are clearly differentiated from frog skin (green). This technique is comparable in ease of diagnosis to immunohistochemistry, although other fungal species are also stained. These stains are useful for confirming infection in cases where only a few indistinct stages are present and sporangia will be easily seen under low magnification. D. IMMUNOHISTOCHEMISTRY

Antibodies can be used in an immunoperoxidase test that specifically highlights sporangia of B. dendrobatidis. The sensitivity and ease of diagnosis is increased and an

BERGER E T AL: FUNGAL DISEASES OF AMPHIBIANS

3027

. . . . . . . .. Fig. 44. Section of skin from a metamorph of Mixophyes fasciolatus with the heavily infected stratum corneum sloughing, leaving few organisms in the epidermis in the left half of the image. Sporangia appear as round spaces or dark circles in the shedding skin. Stained with hematoxylin and eosin. Fzg. 45. Normal toe skin from a Litoria chloris with vesicular structures in the epidermis (arrow). These are probably ducts from dermal glands but appear similar to empty sporangia. Note that the clear spherical space lacks a cell wall and the adjacent epidermal cells are not hyperkeratotic. Stained with hematoxylin and eosin.

untrained investigator can detect the presence of the fungus (Figs 46, 47). Polyclonal (PAbs) and monoclonal (MAbs) antibodies were generated against B. dendrobatidis by inoculating homogenized whole culture into rabbits, sheep or mice (Berger et al. 2002; Hyatt and Olsen, unpubl. data). When used in an immunoperoxidase test hngal sporangia are stained brickred (Berger et al. 2002). Antisera react strongly with all stages of B. dendrobatidis and stain the walls, cytoplasm, rhizoids and zoospores. Immmunostaining kits with other chromagens have also been used successfully. Both the PAbs and MAbs produce some cross-reactivity but this occurs only with other Chytridiomycetes, which are not animal pathogens. The polyclonal antibodies cross-react with two tested chytrids in the order Chytridiales (Berger et al. 2002), and Mab 19G6 also cross-reacts with three tested chytridialeans (Boyle and Hyatt, unpubl. data). Fungi from other phyla are not stained. The only other spherical fungus known to infect amphibian skin is Mucor amphibiorum (Speare et al. 1997) and this fungus is not labelled by the antibodies. The immunoperoxidase stain with polyclonal antibodies against B. dendrobatzdis has greater sensitivity than does staining with hematoxylin and eosin. At 19 days after experimental exposure, 61.8% (n = 55) of toe-clip samples were evaluated as lightly infected

Fig. 46. Immunoperoxidase stain on skin of a Litoria caerulea with a heavy infection of B. dendrobatidis. Fungal walls, cytoplasm, zoospores and septa stain strongly. From Berger et al. (2002). Fig. 47. Immunoperoxidase stain on skin of a Litoria caerulea with a lighter infection of B. dendrobatidis, demonstrating the sensitivity of the test in highlighting a few sporangia. From Berger et al. (2002).

3028

AMPHIBIAN BIOLOGY

with the immunoperoxidase staining protocol, whereas 52.7% of the same toeclip samples were rated as positive by staining with hemotoxylin and eosin (Berger et al. 2002). The immunoperoxidase stain is useful when combined with recognition of the morphology and infection site of B. dendrobatzdis. Immunohistochemistry can be used instead of, or as an adjunct to, conventional histological staining when increased sensitivity of testing is required. It is useful for screening toe-clip samples from healthy frogs in which only a few sporangia may be present; these can be detected at low magnification. The stain is also useful for testing necrotic or autolyzed samples from sick frogs that may contain few distinctive sporangia. The benefits of increased sensitivity and quicker examination of slides stained by the immunoperoxidase test must be weighed against the greater complexity of the staining method. This takes about four hours and requires more expensive and specialized reagents. Many diagnostic laboratories, however, conduct immunohistochemical tests and various methods can easily be adapted to use B. dendrobatidis antibodies. E. DETECTION O F Bah-achochyt~umdendrobatidis WITH IMMUNOHISTOCHEMISTRY AND A KERATINSPECIFIC STAIN

Histological identification of B. dendrobatidis can be complicated by the sloughing of the superficial keratinized layer (stratum corneum) leading to misdiagnosis because sporangia are lost with the sloughed skin. Combining immunostaining for B. dendrobatidis with Hollande's Trichrome keratin stain helps determine whether a negative result could be due to loss of the keratin layer (Olsen et al. 2004). F. SUMMARY O F MICROSCOPY

The advantages of wet preparations are that preparation of the slide is much quicker and cheaper than preparing a histological section, and this method allows a non-invasive ante mortem diagnosis (Table 3). It is the only test used to confirm infection with B. dendrobatidis in a skin sample before attempting to isolate the fungus. A disadvantage is that interpretation is more difficult for an inexperienced worker. Although a larger surface area of skin can be checked by examination of skin scrapings or whole skin, compared to a 5 pm histological section, sporangia may be more difficult to identify. Most frogs sick with chytridiomycosis have heavy infections with B. dendrobatidis that are easily recognized by standard histopathological techniques (Pessier et al. 1999; Berger et al. 2000). Histological diagnosis, however, is less sensitive when dealing with light infections in healthy animals, autolyzed samples, or if the examiner has limited experience in diagnosis of chytridiomycosis. False negatives may occur when prevalence of infection is low and clusters of sporangia are scattered, and hence the sporangia of B. dendrobatidis missed. Immunohistochemical methods increase sensitivity and confidence in the diagnosis, but are also limited by the fact that selected sections may not contain sporangia (Berger et al. 2002; Boyle et al. 2004). As histology is a routine and widely used method, it will continue to be useful for diagnosis. Histology is particularly useful for testing archived museum specimens where PCR is not recommended as multiple specimens are often stored together with potential for cross contamination with DNA. G. POLYMERASE CHAIN REACTION (PCR)

Real-time PCR: Real-time PCR is a method based on conventional PCR, which is a technique for amplifying DNA sequences by separating DNA into two strands and incubating them with oligonucleotide primers and DNA polymerase. The basic technique amplifies a specific sequence of DNA by as many as one billion times to produce a PCR product. Conventional detection of these PCR products uses electrophoresis and ethidium bromide, southern blots or direct sequencing. Advantages of real time TaqMan PCR include increased sensithit): ability to process large numbers of samples (96 well plates) in a short time and the generation of quantitative data.

3029

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS Table 3 . Comparison of the characteristics of each diagnostic test. All four methods of diagnosis are useful, although they have various advantages and disadvantages.

Wet preparations

Histology

Immunostaining

PCR

+ + + ++

++

++ ++ ++

+++ +++ +++ +++

++++ +++++ +++++ +++++

Yes Less useful Smear, scrape, web, other skin

Yes Less useful Toe, web, other skin

Yes Less useful Toe, web, other skin

Use on formalin-fixed specimens

Yes

Yes

Yes

Use Use Use Use

Yes Yes Possible Yes

Yes Yes Possible Yes

Yes Yes Often Yes

Yes

Yes

Yes

Yes Very useful Scrape, toe, web, other skin, swab, filtrate Yes (< 3 months for real-time PCR) Yes Yes Yes No, unless each specimen is stored separately throughout Yes

Complexity and cost of preparation Facilities required Ease of interpretation Sensitivity* Use on live frogs Use for healthy frogs Specimens required

on ethanol- fixed specimens on frozen specimens on decomposed animals in archival surveys

Quantifiable

*

Frogs sick with chytridiomycosis typically have very heavy infections so that highly sensitive tests are not needed.

Real-time PCR differs from conventional PCR in that the PCR product is monitored in real time via the detection of fluorescence emission. The TaqMan probe has two fluorescent tags attached to it. One is a reporter dye that has its emission spectra quenched due to the spatial proximity of a second fluorescent dye (quencher). Degradation of the TaqMan probe, by the Taq DNA polymerase, frees the reporter dye from the quenching activity of the quencher dye and thus the fluorescent activity increases with an increase in cleavage of the probe, which is proportional to the amount of PCR product formed. The TaqMan probe is located between the two PCR primers. The specificity of this technique is attributed to the combined use of specific primers and a specific TaqMan probe. Boyle et al. (2004) developed a real-time TaqMan PCR for the detection of B. okndrobatzdis with a sensitivity of 0.1 zoospore equivalents. PCR amplification and sequencing of chytrid fungal rDNA, which included the 18S, ITS-1 and 5.8s regions, led to the generation of a forward primer (ITS-1 Chytr), a Taqman probe (Chytr MGB2) located within the ITS -1 region, and a reverse primer (5.8s) located partially within the 5.8s region. Four isolates of B. dendrobatzdis and five other Chytridiomycetes were tested and the assay was found to be specific for B. dendrobatidis. Those tested were Rhizophydium sp. (JEL136), Rhizophlyctis-Rhizophydium-like (JEL142), and R. haynaldii (JEL151) from the Order Chytridiales, Gonapodya sp. (JEL183) from the Order Monoblepharidales, and Allo~nyces macrogynus (JEL204) from the Order Blastocladiales. Standards were generated that permitted the quantitation of B. dendrobatidis within a sample. As there is a high copy number of the ITS-1 region, it is possible to detect low numbers of organisms; for example, these authors demonstrated a sensitivity of 0.1 genome equivalents. This test can therefore detect infection at much lower levels than can any other technique. Preliminary data, however, suggest that frog secretions may interfere with the test, leading to false negatives, if they are not initially diluted (Hyatt, unpubl. data). The assay has been used to successfully detect B. dendrobatidis in the mouth parts of tadpoles, in mummified frogs and in formalin-fixed paraffin-embedded samples. With an effective sensitivity of one zoospore, laboratory experiments have demonstrated that B. dendrobatidis can be detected as early as seven days post-infection (Hyatt, Boyle and Olsen, unpubl. data). Although swab samples can be batched, which would have the advantage of reducing costs, preliminary experiments (Hyatt, Boyle and Olsen, unpubl. data) indicate that batching decreases the sensitivity of the assay. Batching five swabs failed to detect very low levels of

3030

AMPHIBIAN BIOLOGY

infection, i.e., when one low positive swab was included with four B. dendrobatidis-negative swabs, the batched sample returned a negative result. When higher numbers of swabs were batched, the sensitivity of the assay decreased markedly. Standard PCR: A conventional DNA-based assay has also been developed for the detection of Batrachochytrium dendrobatidis (Annis et al. 2004). This assay uses primers designed from the ITS1 and ITS2 regions and produces a 300bp product compared with 146bp for the real-time assay (Boyle et al. 2004). The standard assay was used with small amounts of skin and had a sensitivity of 10 zoospores per sample. It detected all 52 isolates of B. dendrobatzdis examined. The assay did, however, produce a faint band with Podochytrium dentatum (a chytrid that grows on chitin and is not known to grow on amphibians). This assay lacks the capacity for accurate quantitation and is less sensitive than the real-time PCR. The equipment required, however, is cheaper and more readily available. Swabs may be used and good results are achieved when these are stored in ethanol and the ethanol included in the DNA extraction process (J. Wood, pers. comm.). Standard PCR has been used to diagnose infection in samples that have been stored in formalin for years (E. Sadic, unpubl. data) and is therefore useful in testing archived samples. In summary, PCR is much more sensitive than microscopy, larger numbers of samples can be tested rapidly, and swabbing is non-invasive and hence suitable for live frogs. It is the best diagnostic test for surveying wild populations. A disadvantage of all PCR tests is that expensive laboratory equipment is required and a high level of care is needed in the collection of samples and in the running of tests in order to ensure that contamination with DNA does not occur. J. Management 1 . Disinfection

Effective disinfection protocols are essential to render equipment used with amphibians non-infectious. This has a major role in the field, in the laboratory and in captive husbandry. B. dendrobatzdzs is susceptible to a broad range of chemical and physical treatments (Johnson et al. 2003). Effective solutions for field use contain the quaternary ammonium compound didecyl dimethyl ammonium chloride (DDAC). For example, Path-X can be used at 1 in 500 dilution for 30 seconds. Sodium hypochlorite (bleach) is used at concentrations of at least 1% for 1 minute. These chemicals are useful for cleaning boots. Bleach has also been used effectively for disinfecting animal tubs (G. Marantelli, pers. comm.). Exposure to 70% ethanol, 0.1% Virkon or 0.1% benzalkonium chloride for 20 seconds is effective. These chemicals can be used for disinfection in the laboratory, in captive facilities and in the field. Alcohol wipes, for example, can be used to disinfect scissors, calipers and other instruments between use for different animals. Although sodium chloride (10% for two minutes or 5% for five minutes) is effective, it may damage equipment. Cultures of B. dendrobatidis do not survive complete drying but persistence in droplets of water allowed survival of the pathogen for up to three hours of drying (Johnson et al. 2003). Heating to above 37°C for four hours also results in death of sporangia (Table 1). Ultraviolet light (1 000 mW/m2 with a wavelength of 254 nm) used routinely for killing bacteria, fungi and viruses, is ineffective at killing B. dendrobatidis in culture (Johnson et al. 2003). A combination of heating and drying is a safe method for disinfection of many objects, such as clothing and some equipment. Care is needed when using chemicals so that bodies of water are not contaminated. 2. Treatment of Chytridiomycosis

Effective treatments of adults and tadpoles are needed to prevent mortality in captive programmes for threatened species and to reduce the risks associated with movement of

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3031

amphibians. Because infected tadpoles survive and remain at sites after adults have died, a method of clearing them of infection could enable an emergency response, such as captive raising, to mortality and declines in threatened species. Successful treatment with antifungals has been reported although trials were small and lacked controls or thorough post-treatment testing. Oral itraconazole was used successfully in Bufo baxteri for treatment of a mycotic dermatitis that was diagnosed as due to Basidiobolus ranarum (Taylor et al. 1999a), but was retrospectively diagnosed as chytridiornycosis (Carey et al. 2003). One micro-bead from the 100 mg capsules (Janssen Pharmaceuticaa) was given daily for nine days to the toads (average weight 30 g) (Taylor et al. 1999a). Itraconazole has been used orally at 2-10 mglkg per day in some amphibians (Taylor 2001). Bathing in 0.01% itraconazole suspension for five minutes per day for 11 days successfully treated chytridiornycosis in Dendrobates tinctorius (Nichols and Lamirande 2000). A commercial solution of 25 ppm formalin and 0.10 mgll malachite green was used for 24 hours every other day four times to successfully treat Xenopus tropicalis (Parker et al. 2002). As malachite green is teratogenic, it is not recommended for endangered frogs. Attempts to treat Australian frogs and tadpoles are underway but have not been successful so far (Marantelli et al. 2000). Although using high temperatures (37°C) to clear infections was effective in Litoria chloris (Woodhams et al. 2003), this treatment has not worked in other species (Marantelli, unpubl. data). Baths of fluconazole and itraconazole for tadpoles were ineffective or toxic (Marantelli et al. 2000). Success of treatment may be affected by innate resistance of hosts, which varies among species. 3. Quarantine Guidelines

Detailed guidelines have been produced for fieldwork (New South Wales National Parks and Wildlife Service 2000), for amphibian husbandry (Lynch 2001) and for translocation (Daszak et al. 2001). Quarantine periods should be at least two months whether moving frogs between countries, sites or captive collections. Frogs in quarantine should be kept between 17" and 24°C to increase the chance of infected frogs showing signs of disease. During the quarantine period they should be examined for signs of disease and tested for chytridiornycosis by PCR before release. Any animals that die must be necropsied (Daszak et al. 2001; Lynch 2001). Using routine quarantine methods, chytridiornycosis has not spread between frogs housed in close proximity and airborne transmission has not been observed (Marantelli et al. 2004). Even small numbers of zoospores, however, are highly infectious and great care is needed to prevent drops of water from contaminating groups of animals. Important quarantine practices include: changing gloves between every enclosure, disinfection of equipment and tubs between use, cleaning and feeding animals in the same order each time starting with those most likely to be disease-free, use of automated husbandry systems, and disinfection of water used in enclosures before disposal (Marantelli and Berger 2000; Lynch 2001). Hygiene protocols for amphibian field work were produced by the New South Wales National Parks and Wildlife Service (2000) and Speare et al. (2004). The protocols include instructions on how to disinfect boots and equipment when moving between sites. Strategies to avoid increasing exposure to disease when handling frogs involve the use of disposable gloves and plastic bags. Frogs must never be held together in the same container and containers must not be reused without sterilization. Tadpoles for release should not be held with batches of tadpoles collected from other sites even if from the same water body. Although chytridiomycosis is already widespread, it is important not to increase transmission rates and exposure to new strains above natural levels. 4. Control Options

In 2002 chytridiornycosis was accepted as a Key Threatening Process in Australia (http:/ twww.ea.gov.aulbiodiversity/ktp/index.html),and a Threat Abatement Plan has been produced

3032

AMPHIBIAN BIOLOGY

(http://www.deh.gov.au/biodiversity/threatened/publications/tap/chytrid/pubs/chytridbackground.pdf) by the Department of Environment and Heritage with Rick Speare as consultant. The following observations are taken from that plan. Countries have international responsibilities to prevent export of the amphibian chytrid to countries currently chytrid-free. Amphibian chytridiomycosis was placed on the wildlife Diseases List of the World Organization for Animal Health (OIE) in 2001. Control activities include legislation and regulation, education and research. Regulation includes extending quarantine protocols into national protocols. Surveillance is required so that all current instances of B. dendrobatidis infection are identified and mapped, and any new occurrences of B. dendrobatidis are identified rapidly. Implementing effective protocols to prevent movement of strains of the pathogen via infected amphibians, either internationally or within countries, is recommended. This will involve improving management and packing of agricultural and nursery produce liable to accidentally translocate amphibians at the points of origin and destination of the produce. It also involves controlling human movements and release of amphibians between areas. These recommendations have implications for the pet trade, food market, provision of experimental animals to scientists, and transportation of agricultural produce and plants. A procedure for not returning frogs to their point of origin is required unless they can be shown to be free of diseases such as chytridiomycosis. Educating the community about basic disease management and the risks of transporting potentially infected amphibians and water is important. Groups at higher risk of spreading the fungus (pet traders, scientists, schools, wildlife carers) should be targeted. Further research is needed on important general topics and on those that apply directly to management. General topics include epidemiology such as factors affecting spread, transmission, incidence, recovery and mortality, origins, and presence of the pathogen pre1970. Areas of applied research include developing treatment and prevention protocols, investigating whether B. dendrobatidis can be eradicated from infected sites, and understanding and influencing mechanisms of resistance. Captive breeding may be useful to (1) produce research animals so that importation is unnecessaly and (2) supplement numbers of individuals of endangered species. By producing large numbers of frogs in captivity, combined with selective breeding, it may be possible to help species survive and evolve immunity. To improve the chances of populations surviving the effects of chytridiomycosis, activities that reduce population size, such as habitat destruction, need to be avoided. K. Discussion 1 . Chytridiomycosis and Amphibian Declines

In some regions the introduction of exotic predators or habitat changes such as logging, wetland drainage, weed invasion, urban development and agriculture have caused amphibian population declines (Alford and Richards 1999; Gillespie and Hero 1999; Hines et al. 1999). There is no evidence, however, that these factors have been responsible for most of the catastrophic declines in protected montane areas. The introduction of chytridiomycosis is the sole cause of declines in many areas of Australia (Australian Government Department of Environment and Heritage 2006). Evidence that similar situations exist in Latin America, North America and New Zealand is increasing. Genetic studies should provide details on the timing and direction of spread, and may confirm Africa as the origin of the disease. Although the timing of some catastrophic declines in protected areas has been correlated with drought (Pounds et al. 1999), the unusual weather was not unprecedented (Alexander and Eischeid 2001). Environmental factors influence the susceptibility of populations to chytridiomycosis; however, the change that caused declines was the arrival of B. dendrobatzdis. Environmental co-factors affect the susceptibility of populations and research is required

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3033

to determine why species and populations in various regions are affected differently. Temperature and a mesic environment are important factors. Batrachochytrium dendrobatzdis has the ability to spread widely, wiping out susceptible amphibian populations when it arrives. Despite being relatively fragile and killed by drying, it has spread to, and survived in, a wide variety of habitats and climates (Berger et al. 2004). Its broad host range and probable ability to survive in the environment (at least for several months and possibly indefinitely), are factors that may have allowed it to persist when numbers of some frog species have crashed. A crucial area for further research is on the growth of B. dendrobatidis in the environment, in particular determining if and where it grows, what conditions stimulate growth and the role that frogs play in its survival. Since eliminating the fungus from infected frogs is much harder than killing it i n vitro using heat or antifungals (Berger 2001; G. Marantelli, unpublished data), it is possible that sporangia may survive in frogs during times or in areas where the environment does not support overt growth of the fungus.

The amphibian chytrid may have a critical impact on any amphibian population that it infects. This will depend on a complex interaction between the innate characteristics of individual species, the pathogen and the environme~lt.An important protective determinant is a distribution that includes environments not favourable to chytridiomycosis, e.g., low elevations or saline environments. Factors affecting mortality rates and/or incubation times experimentally include host species, host age, temperature, fungal dose and fungal strain (see section 11-E-1). The outcome for a species can range from extinction of the species as the worst-case scenario, to a stable population with sporadic or no deaths caused by chytridiomycosis. However, the vulnerability of the apparently stable population to the effect of natural environmental variations and to anthropogenic environmental factors is probably increased. Chytridiomycosis is now a common disease of amphibians. Although it is a frequent cause of death (Berger et al. 2004), the prevalence of infection in populations post-decline may be quite high (McDonald et al. 2005; Retallick et al. 2004). Evidence suggests that in some areas it may now have a host-parasite relationship more typical of an endemic pathogen with outbreaks of mortality occurring when conditions are optimal for the fungus (Berger et al. 2004). In some areas, outbreaks of chytridiornycosis occur on a seasonal cycle with most deaths taking place in winter (Retallick and Dwyer 2000; Bradley et al. 2002; Berger et al. 2004). Higher temperatures - experimentally (such as 27OC) and in the field, - result in lower mortality rates (Woodhams et al. 2003; Berger et al. 2004). Prevalence is higher at lower temperatures at higher elevations and during winter (McDonald et al. 2005; Woodhams and Alford 2005). Effects of temperature explain the distribution of the declines in terms of elevation and latitude in mesic environments. . The long-term prognosis may be good for species that have survived at high enough numbers and are recovering, as long as remaining habitats are protected to allow damaged populations to re-establish. In many areas, however, frogs are much less abundant than previously noted and some species are missing fi-om their previous distributions, in particular many populations from high elevations are gone (McDonald and Alford 1999). Hence, it appears that at some sites the continuing consequence of the disease on anurans is great. Some frog species have not survived, and although the majority of species remain, the broadscale effect of chytridiomycosis is yet to be determined and additional species could become extinct. Some species, especially those with small distributions in environments favourable to 'chytridiomycosis in currently uninfected areas, may be at high risk It is necessary in the short term to continue mapping the distribution of chytridiomycosis to identify countries and areas that are at high risk for introduction of the disease. Preventing the further spread of chytridiomycosis, rather than responding once declines have occurred, is a key to managing the disease. In the long term, sustained

3034

AMPHIBIAN BIOLOGY

research to understand resistance and epidemiology may lead to management involving immunity or biological control. 2. Chytridiomycosis in the Context of Other Introduced Diseases of Wzld Animals and Plants

Chytridiomycosis is not unusual in being an introduced disease that has caused catastrophic declines of wild populations. Infectious disease is important in the population biology of wild animals, as it is in humans and domestic animals (May 1988). A review of infectious disease and animal populations concluded that disease is an important factor affecting survival, reproduction, dispersal, community structure and genetic diversity, and should therefore be considered by ecologists examining host population-dynamics (Scott 1988). Exotic diseases can have effects similar to those of feral predators with susceptible native species facing extinction while the ecosystem readjusts. Chytridiomycosis could be another example of how increased global homogeneity leads to reduced biodiversity. There are many examples of mass mortality and catastrophic population declines occurring when infectious diseases have been introduced. Rinderpest was an ancient disease of livestock in Asia and Europe. It was introduced into Afiica and spread across the continent between 1889 and 1898 (Scott 1981; Plowright 1982). It is fatal to a wide range of hoofed animals and millions of wild animals died, with carcasses littering the plains. The introduction of avian malaria and birdpox is suspected to have been involved in the extinction of low-elevation birds in Hawaii (Warner 1968). Because Australia's fauna and flora have evolved in isolation, some introduced diseases have had severe effects. A pathogenic herpesvirus was apparently introduced to Australasian pilchard populations (possibly in imported pilchards fed to tuna) causing a massive epizootic that spread across more than 5 000 km of Ausn;ilian coastline in four months (Whittington et al. 199'7). Although many observers suggested that the deaths were due to an environmental problem, the dramatic spread of mortalities from a focal origin and the unprecedented losses in a single species over a huge area were consistent with an infectious disease (Whittington et al. 1997). Sarcoptic mange, caused by the mite Sarcoptes scabiei is an introduced disease affecting many wildlife species globally (Iknce and Ueckermann 2002). In wombats it can cause large epidemics and population declines, and can also occur endemically with sporadic disease in individual wombats (Skerratt et al. 1998). Aphanomyces astaci, the cause of crayfish plague, is an introduced fungal-like pathogen that has caused severe declines of crayfish as it moved across Europe (Josefsson and Andersson 2001). Imported h n g i and fungal-like organisms have been responsible for diseases of plants, with devastating effects on populations (Tainter 2003). The whole landscape of eastern North America changed when chestnut blight (Cryphonectria parasitica), which migrated from Asia on imported nursery stock in the early 1900s, spread and killed most trees of a key hardwood species. The American landscape changed again when another fungus (Ophiostoma ulmi which causes Dutch elm disease) migrated on elm wood from Europe in the 1920s and killed most of the American elms. Currently, a zoosporic fungus-like organism (Phytophthora ramorum) is causing sudden oak death on the western coast of North America. This disease, which was first identified in Europe in 1993, is not limited to a single host, persists saprobically and is devastating oaks. Phytophthora cinnamomi has a broad host range and threatens many native Australian plant species (Dawson and Weste 1985; Wills 1993). Some plant species are highly susceptible, whereas others only become diseased after periods of stress such as a drought (Dawson and Weste 1985; Wills 1993). Similar to these Phytophthora species, B. dendrobatidk can live in many host species, may be able to persist saprobically and has differing pathogenicity and virulence to each host. Chytridiomycosis in amphibians is the first well-documented case in which extinction of a species in the wild has occurred due to a disease. Stricter quarantine regulations, and ongoing surveillance for wildlife diseases, are urgently needed.

BERGER ET AL: FUNGAL DISEASES O F AMPHIBIANS

111. OOMYCOSES A. Introduction Oomycetes are secondary invaders of wounds in adult and larval amphibians, and can be pathogens of larvae or adults, but it is as pathogens of amphibian eggs that they are of conservation concern. Saprolegnia is the most common genus encountered as an amphibian pathogen and species of this genus also are well known as pathogens of fish. High levels of infection with Saprolegnia caused mortality of Bufo boreas egg masses in northwestern United States (Blaustein et al. 1994b; Kiesecker et al. 2001a) and were sufficient to affect local populations (see section 111-E-2-B). Saprolegnia differs in its effect among species of amphibians (see section 111-E-2-D).The relatively low infection rate of Hyla regilla eggs was attributed to the frog's habit of not laying eggs communally (Kiesecker and Blaustein 1997). Factors that increase egg mortality include low temperatures, low pH (Strijbosch 1979; Banks and Beebee 1988) and exposure to UV-B (Kiesecker and Blaustein 1995). Oomycete infections of eggs are clearly not a cause of the rapid catastrophic amphibian declines in which adult populations disappear within months (Laurance et al. 1996). B. Taxonomy Members of the Oomycota (oomycetes) are traditionally studied by mycologists and are considered fungi in the nutritional sense. They are, however, unrelated to the true h n g i (Eumycota). Like the true hngi, they obtain nutrients by excreting enzymes that degrade complex substrates; smaller molecules can then be absorbed through the cell wall, which contains a high proportion of beta-glucans, whereas true fungi have chitinous cell walls. The phylum Oomycota, also known as the Peronosporomycota, is classified in the kingdom Stramenopila with other organisms that produce zoospores or life stages that possess an anteriorly directed tinsel flagellum and a posteriorly directed smooth flagellum. Diatoms, kelps and golden algae are algal members of this group. DNA evidence suggests that the oomycete lineage lost chloroplasts (Yoon et al. 2002) whereas algal members of the kingdom retained them. Saprolepia (Saprolegniales; Saprolegniaceae) is the best-known oomycete genus that contains pathogens of amphibian adults, larvae and eggs. Saprolegnia parasitica, S. diclina and S. ferax (Beakes et al. 1994) are pathogens of fish eggs and infections of fish wounds and S. firax is the species identified as parasitizing amphibian eggs in the United States (Blaustein et al. 1994b). Other genera in the same family that have members that may affect fish and amphibians include Achlya and Aphanomyces. Aphanomyces astaci is a virulent parasite of crayfish (Unestam 1973) and A. invadans is a fish pathogen (Willoughby et al. 1995). Many reports in the literature include only the genus without an attempt to identify the species. Taxonomic characters in the Saprolegniaceae must be determined from the morphology of isolates in pure culture but, even when all the pertinent features can be seen, identifying species by following keys is often difficult. Immunologic (Bullis et al. 1990) and DNA characters are being used to clarify the relationships and taxonomy of this group. The entire mitochondria1 genome of Saprolegnia firax is known (Grayburn et al. 2004) and this information may provide insight into variable sites that can aid in determining population genetics of this important species. Keys to genera and species of the Saprolegniaceae are available (Seymour and Fuller 1987; Johnson et al. 2002).

C. Biology Like most other Oomycetes, members of the Saprolegniaceae form tubular hyphae that contain diploid nuclei. Septations within the hyphal filaments are lacking except where they delimit reproductive structures. Members of the Saprolegniaceae usually first reproduce

3036

AMPHIBIAN BIOLOGY

asexually. Zoosporangia (containers in which the asexual zoospores are formed) produced by Saprolegnia species are slightly inflated and usually terminal; zoospores exit through an apical discharge papilla. Characteristic of this genus, some subsequent zoosporangia form at the tip of a hypha that has already produced a zoosporangium and the new zoosporangium grows through the earlier-formed, empty sporangium. Primary zoospores, which are apically biflagellate, are released and swim away from the sporangium. These zoospores encyst and then may release a secondary zoospore, which is laterally biflagellate. Zoosporangia of Aphanomyces do not differ from the hyphae; the contents of extensive areas of the hyphae differentiate into zoospores, which exit from the tip of the hypha-like zoosporangium. These primary zoospores immediately encyst at the tip of the empty hypha, forming a characteristic sphere of spores. Secondary, motile, biflagellate zoospores emerge from the encysted primary spores. Achlya shares features of both Saprolegnia and Aphanomyces. Like Saprolegnia, the zoosporangia are identifiable by being slightly inflated and, like Aphanomyces, the primary zoospores encyst at the tip of the zoosporangium forming a sphere made up of encysted zoospores. Achlya also differs from Saprolegnia in that its later-formed zoosporangia do not grow through earlier formed ones. Features of the zoosporangia and sexual features also distinguish other genera in the Saprolegniaceae (Seymour and Fuller 198'7;Johnson et al. 2002). Sexual reproduction occurs subsequent to the first flush of asexual reproduction in most species in the Saprolegniaceae. Oogonia (egg-containing structures) form, within which are one or more oospheres (eggs). Antheridial (male) cells 011 the tips of antheridial branches grow toward and contact the oogonia; a tube grows through the oogonial wall and the male nucleus migrates into the oosphere. The fertilized oosphere becomes an oospore; during maturation it becomes thick-walled and is the resistant stage of the oomycete. Variations in lengths and origins of oogonial and antheridial branches, ornamentation of oogonial walls, and numbers and the distribution of globules in oospores supply sufficient variation to be useful as taxonomic characters but c o h s i o n about the limits of species persists. D. Oomycete Diseases of Larval Amphibians Saprohgnia has been reported from dead tadpoles of Spea bombifions, Rana berlandieri and Pseuducris streckeri in pools in Oklahoma, United States (Bragg and Bragg 1958; Bragg 1962). Oomycetes, however, frequently infect wounds of larval and adult amphibians, and immediately infest animals that die from other causes, and so whether these tadpoles were killed by the Saprolegnia infection is unknown. Aphanomyces, however, has been shown to be a disease agent of tadpoles (Berger et al. 2001). Tadpoles of the giant toad (Bufo marinus) growing in muddy water in Queensland, Australia were collected in 1989 with tufts of hyphal fungi attached to the nostrils, mouthparts, other parts of the head, and occasionally to the hind legs and tail. The oomycete did not invade past the dermis. A second outbreak occurred in 1995 in a drying pond behind a dam, also in Queensland. Of a random sample of tadpoles in the second outbreak, 3'7 of 100 were infected (Fig. 48). Aphanomyces and other, probably opportunistic, fungi were found in attempts to culture this pathogen on Sabouraud's Dextrose Agar, Glucose Yeast Extract Agar or Fig 48. Head of a tadpole of Bufo marinus with a tuft of mixed fungi attached to the left nostril. The fungus forming the water agar. Direct examination of the dense, central clump is consistent with Aphanomyces sp. infecting hyphae and scanning

BERGER ET AL: FUNGAL DISEASES OF AMPHIBUNS

3037

electron microscopy of samples of infected skin, which showed clusters of primary encysted spores, were both consistent with the pathogen being a species of Aphanomyces. Examined larvae were not infected with other pathogens and Aphanomyces was considered to be the primary disease agent. Although no dead tadpoles were found in the wild, many of the collected diseased tadpoles died during transport to the laboratory. Infected animals were thin and less mature than were healthy ones. The authors concluded that the tadpoles had starved and that the disease, once started, was probably lethal. Because the stramenopile primarily infected the mouth, it may have prevented eating. Tadpoles of Rana boylii from California had oomycete infections with hyphal tufts about the mouth and spiracle (Green 2001). When Lefcort et al. (1997) studied the effects of silt and used motor oil on the growth and mortality of larval mole salamanders (Ambystoma opacum and A. tigrinum tigrinum) they found that in their silt and silt-plus-oil treatments animals became infected with Saprolegnia parasitica. They traced the source of the infection to the water that contained the zooplankton fed to the salamanders. Infections started on toes and then affected entire legs. All the animals that became infected died but no control or oil-alone animals died. Because uninfected animals in silt-alone treatments grew less than did control animals, the authors concluded that silt may have stressed the salamanders, making them more susceptible to infection by Saprolegnia. E. Oomycete Infections of Amphibian Eggs 1 . Pathogenesis

Any source of nutrients in aquatic systems, including dead amphibian eggs, rapidly becomes infested with oomycetes. The ability of fungi to be pathogenic may vary geographically, by species of host and pathogen, and with differing temperatures and pH. Although infection of fish eggs by Saprolegnia is well known and is a problem in fish hatcheries, less has been published on the effect of Saprolegnia and other genera of oomycetes on amphibian eggs. It seems clear, however, that compromised amphibian eggs can become infected by a broad range of Saprolegniaceae (Czeczuga et al. 1998) and that healthy amphibian embryos also become infected, perhaps by more pathogenic strains or species that can spread from infected dead eggs to living eggs (Blaustein et al. 1994b; Green 1999; Robinson et al. 2003). Two unidentified species of fungi that grew on treefrog egg masses in Central America were shown experimentally to be pathogenic and lethal (Villa 1979). Infection of egg masses can be recognized macroscopically and microscopically by the appearance of a white, f l u e mycelium (Fig. 49). Hyphae are abundant in the egg capsule but may also invade the yolk and embryo (Green 2001). Sometimes oomycetes infect only eggs and embryos that are dead or injured, and a high proportion of larvae from egg clumps with infection may still hatch because healthy eggs may remain uninfected (M. Gahl, pers. comm.). Some oomycete species or strains, however, are pathogenic and spread throughout an egg clump by invading living embryos. Blaustein et al. (1994b) observed microscopically the hyphal filaments of Saprolegnia ferax growing from infected embryos into living, normally developing embryos. More recently, Robinson et al. (1999) used electron microscopy to track the progressive invasion of amphibian eggs by several different strains of saprolegniaceous oomycetes, although detailed information has not been published. Green (1999) reported that, in aquaria, eggs of newts (Triturus spp.) frequently became infected with water moulds (Saprolegniaceae)and that hyphae spread and killed neighbouring eggs. 2. Epidemiology A. SAPROLEGNIA INFECTION O F AMPHIBIAN EGGS IN EUROPE

In the early 1970s Strijbosch (1979) collected vegetational and physiochemical data on 15 sand dune fens in The Netherlands and correlated these data with the success of eggs

3038

AMPHIBIAN BIOLOGY

laid by three species of Rana, two species of Bufo and the spadefoot toad, Pelobates fiscus. He observed egg masses daily and reported that, in seven of 14 fens where amphibians laid their eggs, Saprolegnia infected 100% of the eggs. By comparing the physiochemical characteristics of the different fens he concluded that Sap-olegniu infection was particularly prevalent in acidic, oligotrophic fens and that the eggs of some species (e.g. P h c u s ) seemed to be more susceptible to Saprolegnia than were others (e.g. Bufo calamita). Even in fens where the eggs of other amphibians were always infected, at least some B. calamita eggs developed. Banks and Beebee (1988) studied development of eggs of natterjack toads (Bufo calamzta) in pools and in the laboratory. They observed that death of eggs was caused primarily by infestation by "Saprolegnza" at low temperatures and at pH concentrations below 6. Their laboratory experiments confirmed that infection by Saprolegnza decreased at higher temperatures and they concluded that high water temperature promoted faster development of the embryos, thus minimizing infection of eggs by Saprolegnza. They cited desiccation of breeding pools and predation of tadpoles by invertebrates as key causes of mortality during development, instead of infection by Saprolegnza.

Joseph M Kzesecker)

Beattie et al. (1991) also found that infection of h g eggs was correlated with low temperatures and acidic conditions in England. In their field study of embryonic development of the common frog (Rana temporarza) in 36 ponds (areas less than 0.5 hectare) they found that infection by "probably a Saprolegnza sp." increased with decreasing Ca+ + concentrations, and also with decreasing temperature and increasing acidity. They could not determine whether the oomycete caused the death of eggs or grew on eggs that were already dead but, because 83% of the eggs were not infected and developed into normal tadpoles, these oomycete infections were probably not significant at the population level. Robinson et al. (1999) isolated saprolegniaceous fungi from dead and diseased amphibian eggs in southern England. DNA fingerprinting methods revealed that several different strains were involved and the authors believed that the oomycetes were responsible for egg mortality. B. SAPROLEGNZA O N AMPHIBIAN EGGS IN NORTHWESTERN UNITED STATES

Blaustein and colleagues have published the most research on the effects of Saprolegnia and environmental cofactors on hatching of amphibian eggs. The pattern of infection by Saprolegnia in lakes in Oregon, indicated that a pathogenic strain was present (Blaustein et al. 1994b) and mycologists identified it as S. ferax, which is also a fish pathogen (Beakes et al. 1994). In the study lake, infection moved in a wave-like pattern through two large communal egg masses laid by Western toads (Bufo boreas) and the researchers noted that, when eggs were in close proximity, hyphae grew from infected eggs into adjacent uninfected eggs (Fig. 49). Blaustein et al. (1994a,b) reported that, during three years, millions of Western toad embryos died in three high-elevation lakes from a combination of factors, including a synergistic interaction between UV-B and Saprolegnia (see section D). Mortality rates of embryos ranged from 50 to 95%. For one of these lakes (Lost Lake), which was a long-

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3039

term study site, data collected from 1980 to 1989 revealed that mortality of Western toad eggs was not more than 5%. High mortality of embryos continues in this area. From 1990 to 1997 mortality ranged from 60% to 100% in 6 of 7 years (Kiesecker et al. 2001a,b). Brook trout, rainbow trout and Atlantic salmon are stocked in these lakes and the researchers hypothesized that S. firax could be introduced during stocking of fish, if it were not already present. If it were already present, its density may be increased during fish stocking (Blaustein et al. 199413). C. TRANSMISSION

Members of the Saprolegniaceae are prevalent in most natural fresh waters and in soils. The capability of species that are pathogenic to also grow on decaying plant and animal debris permits these organisms to persist and multiply even when amphibian or fish hosts are absent. The production of zoospores allows transfer of the fungus long distances to new clusters of eggs by water movement, and chemotaxis of the zoospores allows oriented movement over shorter distances. Spread of strains that are pathogenic to both fish and amphibians, and that accumulate in fish hatcheries, may be particularly important. Kiesecker et al. (2001b) showed that hatchery fish could spread Saprolegnia firax to the embryos of Bufo boreas. In laboratory experiments they showed that this oomycete is pathogenic to eggs and that, in 38-L aquaria, developing B. boreas larvae became infected with Saprolegnia when placed in the same tank as a trout experimentally infected with S. f i r m . Eggs also became infected when placed with an un-manipulated (neither disinfected nor exposed to S. f i r m ) hatchery trout; this means that the trout was naturally carrying an oomycete infection. They also showed that hatching success was lower when embryos of B. boreas were placed in aquaria with soil that had been exposed to laboratory-infected trout and then dried for 16 days, thus suggesting that S. ferax can remain viable as resting spores in pond mud. D. COFACTORS

Other factors affect the loss of eggs to Saprolegnia. Kiesecker and Blaustein (1997) showed that infection of amphibian eggs is greater when eggs are communally deposited and greater for eggs that are deposited later in the season. Importantly, they found that Saprolegnia differs in its effect among species of amphibians, with never more than 6% egg mortality for eggs of Hyla regilla in the same bodies of water in which Rana cascadae. had 8% to 80% egg mortality and Bufo boreas eggs had greater than 50% mortality (Kiesecker and Blaustein 1997). The relatively low infection rate of H. regilla eggs was attributed to its habit of not laying eggs communally. Egg mortality associated with Saprolegniu ferax infections is also mediated by the amount of UV-B light incident upon the eggs. More eggs produce larvae when deposited at a greater water depth (Kiesecker et al. Zoola), which naturally attenuates ultraviolet light, or when shielded by UV-B filters in experimental systems. This effect varies among species, with UV-B enhancing the pathogenicity of Saprolegniu to eggs of Bufo boreas and Rana cascadae but not to those of Hyla regilla (Kiesecker and Blaustein 1995). Climatic changes that result in reduced water depth can lead to increased infection rates (Kiesecker et al. 2001a). A probable cofactor for infection of eggs by Saprolegniu is low pH. European researchers (Strijbosch 1979; Banks and Beebee 1988; Beattie et al. 1991) found that bodies of water with lower pH had higher incidence of eggs infected by Saprolegnia. Other implicated cofactors include low temperature (Banks and Beebee 1988; Beattie et al. 1991) and trophic level of the water body (Strijbosch 1979). F. Population Effects of Oomycete Infection of Eggs

Does egg mortality reduce adult population size? Information on the effect of oomycetes on amphibian eggs has come from laboratory studies and from different sizes and types of natural water-bodies; from "pools" that may dry up during the breeding season (Banks and

3040

AMPHIBIAN BIOLOGY

Beebee 1988), "ponds" less than 0.5 hectare in area (Aston et al. 1987; Beattie et al. 1991); "fens" of 0.1 to 6.8 hectares near sand dunes (Strijbosch 1979) and lakes containing stocked, salmonid fish (Blaustein et al. 1994b). These differences as well as differences among species of hosts and pathogens result in a variety of effects on populations. Blaustein et al. (199413) suggested that high egg mortality can decrease populations i n d may have contributed to the decrease in numbers of Bufo boreas breeding in lakes in the Pacific Northwest of the United States. Certainly the 100% mortality of eggs in certain dune fens in The Netherlands affected recruitment from those pools (Strijbosch 1979). Vonesh and de la Cruz (2002) modelled the effects of egg mortality on population levels of B. boreas and Ambystoma macrodactylurn. They concluded that, because of species differences in density dependence of larvae, decreases in egg survival would affect the salamanders more than it would the toads. By their calculations, a 75% decrease in egg survival would result in a 40% decrease in the density of Ambystoma adults but in a 30% difference in adult Bufo densities. Their preliminary analysis of the effect of egg mortality on B. boreas populations led them to conclude that "egg mortality alone, of the magnitude currently observed, may be insuff~cientto explain large declines in this species". Even with the calculations of the Vonesh and de la Cruz model, the levels of mortality of B. boreas egg masses (60-100% in 6 of 7 years) reported by Kiesecker et al. (2001a,b) would be sufficient to affect local populations. The effect of mortality attributable to Saprolegnia at larger geographic scales will depend on the reproductive success in neighbouring breeding sites and the vagility of the affected species. The work of Warkentin et al. (2001) with an unidentified, true fungus (Phaeosphaeriaceae; Dothidieales; Ascomycota) shows that fungal infection of eggs can have more subtle effects than death. They noted that infection of a clutch of red-eyed treefrog (Agalychnis callidryas) eggs by this fungus caused death of infected eggs and early hatching of uninfected eggs. Eggs in infected clutches hatched one half to one day earlier than did eggs in uninfected clutches; uninfected eggs thereby escaped being smothered or killed by the fungus. The ability to hatch early is an important defence strategy but may also put the less-mature larvae at increased risk. Kiesecker and Blaustein (1997) suggested that selective evolutionary pressure may once have favoured egg laying in communal masses because eggs in large groups develop faster than do those in smaller clutches. Now, however, because of the higher mortality from fungal infections in eggs in communal egg masses, this strategy may no longer be beneficial. Green (1999) theorized that fungal infection of amphibian eggs has been important enough during the evolutionary history of amphibians to affect amphibian reproductive strategies. For example, in experiments with Triturus vulgaris newts, he found that 2 1 of 3 1 eggs placed 1 cm from eggs infected with an oomycete died whereas 30 of 31 eggs abutting infected eggs died. Single-egg oviposition (separated by as little as 1 cm) or division of eggs into several small clutches could be a strategy that reduces spread of lethal oomycete infections (Green 1999). G . Diagnosis of Oomycoses

The presence of "hairy" eggs or tadpoles is not sufficient to conclude they are being parasitized or that the fungus is Saprolegnia. True fungi as well as different species of oomycetes invade amphibian eggs. Evidence that live hosts became infected is needed. Currently most oomycetes are identified after isolation. Living material is needed for isolations. Hyphal strands from infected eggs or tadpoles are placed on nutrient agar in culture plates; oomycete hyphae grow rapidly and usually can be subcultured from the isolation plate within a day or two. Multiple isolation attempts may be necessary to detect whether two or more different species of oomycetes might be present. Also, care must be taken to make certain that the hyphae of a single organism are subcultured for each isolate so that identifications are not attempted from mixed cultures. To culture oomycetes so that taxonomic features may be observed, split, boiled hemp seeds are placed around the edges

BERGER ET AL: FUNGAL DISEASES O F AMPHIBIANS

3041

of colonies on nutrient agar. Hyphae grow into the seeds, which are then placed into sterile, distilled water where asexual and sexual reproductive stages develop that allow the oomycete to be identified (Seymour and Fuller 1987; Johnson et al. 2002). As larger genetic databases become available, DNA sequencing will more reliably be able to identify species and strains of amphibian pathogens, as is being done now with pathogens of fish (Cunningham 2002). A true fungus, that did not form typical reproductive structures allowing its identification, infected the eggs of the red-eyed tree frog (Agalychnis callidryas) (Warkentin et al. 2001). This hngus was identified as a member of a family in the Ascomycete order Dothidiales based on DNA sequences of the nuclear small subunit of RNA. Histology of eggs is useful for observing the location of hyphae. H. Management

The high infection rate and mortality of eggs of Bufo boreas associated with introduction of fish into lakes in northwestern United States has management implications for lakebreeding amphibians, especially if this phenomenon is found elsewhere. If molecular techniques (e.g., Lilley et al. 2003) confirm that amphibian egg mortality is caused by infection by strains of Saprolegnia carried by hatchery fish, a reasonable case could be made against stocking fish in lakes that are important breeding areas for amphibians. IV. OTHER FUNGAL DISEASES A. Mucormycosis

Mucor amphibiorum is a zygomycete that causes a fatal disseminated disease with fungal sphaerules inciting granuloma formation in most organs (Speare et al. 1997). Although outbreaks of mucormycosis occur in captive amphibians, high mortality rates have not been observed in the wild where it appears to cause only sporadic infections. Possibly the usual inoculating dose in the wild is not high enough to cause epidemic disease (Speare 2002). Mucor amphibiorum in free-ranging amphibians has only been reported from Australia.

Cases of mucormycosis have been found in amphibians in Queensland, Northern Territory, Western Australia and New South Wales (Speare et al. 1994). In Australia, chronic cases were found in 0.7% of the introduced Bufo murinus in the wild (Speare et al. 1994). Mucormycosis has been found in occasional Litoria caerulea, Litoria infrafrenata and Limnodynastes peronii in the wild in Australia (Berger 2001; Speare 2002) and in Litoria adelensis, Litoria caerulea, L. infiafienata and exotic dendrobatid frogs in captivity in Australia (Creeper et al. 1998) and Germany (Frank 1976). Most cases detected have been fulminating and fatal, but two L. caerulea from Queensland that died with chytridiomycosis had mild localized Mucor infections (Berger 2001). Mucormycosis was first reported as a cause of death in captive anurans in Europe (Frank et al. 1974). It appeared that the original source of the fungus in the European collection may have been specimens of Litoria caerulea from Australia. In transmission experiments, Rana temporaria, R. esculenta and Bufo bufo usually died within a month, lizards were mildly infected, and rats, mice and guinea pigs were unaffected (Frank et al. 1974; Frank 1976). Amphibian mucormycosis is a systemic disease and infected frogs and toads have fungi disseminated through their internal organs and skin. The fungi incite formation of granulomas that consist of inflammatory cells and fibrous tissue. At post-mortem, the liver contains small pale nodules, usually in massive numbers (Fig. 50). These nodules can also be seen in other organs such as the kidney, lung, mesentery, urinary bladder, subcutaneous sinuses and ;kin (Speare et al. 1997). Mucor amphibiorum is dimorphic. In the amphibian host it is a yeast-like spherical structure, called a sphaerule. M. amphibzorum has the unusual characteristic of forming daughter spherules inside the mother spherule (Fig. 51). These can be seen in histological sections of organs or on direct microscopic examination of infected tissue (Speare et al. 1997). Sphaerules range in size from 37 to 4.9 microns. When M. arnphibiorum is growing

3042

AMPHIBIAN BIOLOGY

outside the amphibian host, it becomes thread-like, forming hyphae that form a mat or mycelium. Two mating types of M. amphibiorum exist and, when these meet, they form resistant structures called zygospores. Spores are eventually formed and these are infectious to amphibians when ingested. Mucor amphibiorum is also a pathogen of free-living platypus in Tasmania ( ~ b e n d o ret f al. 1993). It is interesting that, in Queensland, the distributions of platypus and infected toads overlap but disease has not been seen in platypus. In northern Tasmania, mucormycosis has not been found in amphibians but occurs frequently in platypus (Connolly et al. 1999). Mucor amphibiorum grows on soil and will sporulate on the soil. Amphibians probably become infected when they inadvertently ingest soil containing spores along with their prey. M. amphibiorum was isolated from soil in a greenhouse where a series of cane toads (Bufo marinus) had died from mucormycosis over a period of two years (Speare et al. 1994). The available evidence suggests that M. amphibiorum is an environmental hngus that occasionally infects amphibians. The hngus can also be isolated from faeces of infected toads suggesting that cane toads may contaminate their environment (Speare 2002). In captivity, mortality rates in outbreaks have been high (Creeper et al. 1998). Control strategies have not been trialled but recommendations include not feeding contaminated food by using only live food raised in captivity, sterilizing soil and other natural substrates before use, and by following routine quarantine and disinfection guidelines (Lynch 2001; Speare 2002). B. Ichthyophonosis

Ichthyophonus is a protistan parasite of ectotherms and genetic work has grouped it with Dermocystidium in a clade near the animal-fungal divergence (Ragan et al. 1996). In amphibians, the spores cause a granulomatous myositis in skeletal muscles that can be fatal in the wild. Myositis caused by Ichthyophonus-like organisms was detected in 13% of wild amphibians in Quebec, Canada, collected between 1959 and 1999 (Mikaelian et al. 2000). Of 35 infected amphibians, the infection was considered fatal in two R a m clamitans, one R. sylvatica and one Notophthalmus virzdescens, and potentially significant in three additional R. clamitans with severe infections. In June and July 1973 more than 50% of newts in a lake in West Virginia, United States, were infected and by late summer the prevalence approached 100% (Herman 1984). Infection was believed to have caused deaths in this population. In Vermont in 1994,

Fig. 50. An adult Limnodynastes peronii with small nodules caused by Mucor amphibiorum throughout the enlarged liver. Bar = 25 mm. Fig. 51. Sphaerules of Mucor amphibiorum within a granuloma. A mother sphaerule has disintegrated to release daughter sphaerules. Inset shows an intact mother sphaerule. Stained with periodic acid-SchifF.

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3043

nine of 11 red-spotted newts (Not@hthalmwv. vidscens) were collected with myositis caused by Ichthyophonus-like h n g i (Green et al. 1995). Clinically affected amphibians are lethargic and newts may be found floating just below the water's surface. In newts, swellings occur typically on the caudal half of the body, and on the proximal part of the tail (Green 2001). Ulceration over the swellings occurs occasionally (Mikaelian et al. 2000). Spores invade striated muscle fibres and incite mainly granulomatous inflammation in chronic stages. Up to 100% of skeletal muscle fibres may be infected. Acute, chronic and resolving stages of infection occur. The large oval or cigarshaped, thin-walled spores (on average 75 x 175 pm) can be seen on wet mounts of muscle tissue (Green 2001). Infections also occur in muscles of the limbs, neck and head. Two goldfish inoculated with infected muscle from a green frog, and three that were fed infected muscle, were free of infection when euthanased after nine days (Mikaelian et al. 2000). C. Chromomycosis Chromomycosis refers to infection by a range of pigmented, septate hyphal h g i from the phylum Ascomycota. Pigmented fungi, including Fonsecaea pedrosi, E h t i t i d i s , Cladosporium sp. Scolecobasidium sp. and Phialophora sp., have been isolated. Lesions of chromomycosis have been reported in a large range of captive amphibians flaylor 2001) and in wild Bufo melanostrictw in Malaysia (Dhahval and G f i t h s 1963). These organisms have also been isolated from tanks housing captive frogs. Clinical signs are of chronic debilitating disease, and papules and ulceration may occur. In captivity, frogs died 1-6 months after first showing signs of infection. Multiple grey nodules occurred in liver, kidney, heart, lung, skeletal muscle, meninges, bone marrow and other organs. These were fibrous granulomas with mononuclear cells, epithelioid cells and multinucleate giant cells around pigmented, septate hngi or spherical chlamydospores. Transmission experiments had various results. Rush et al. (1974) transmitted disease to healthy, unstressed frogs whereas Elkan and Philpot (1973) could not infect healthy frogs by intraperitoneal inoculation. Cicmanec et al. (1973) transmitted the disease to toads by intracoeloemic injection only if the animals were stressed by refrigeration, infrequent feeding or limited water. D. Amphibiocystidium Amphibiocystzdium has been proposed as a new genus containing species previously described as Dermocystidium, Demzospomdium and Dermmycozdes in amphibians (Pascolini et al. 2003). This protistan grows as large spore-filled cysts in subcutaneous tissue or the dermis and can cause inflammation and ulcerations (Broz and Privora 1951; Jay and Pohley 1981). Infections have been found in Europe and America, and can occur at high prevalence in a population (Reichenbach-Klinke and Elkan 1965; fiscolini et al. 2003). Mortality has not been reported.

V. ACKNOWLEDGEMENTS Thanks a lot to Scott Cashins and Andrea Phillott for helpful comments on earlier drafts, to Frank Filippi for assistance with photography, and to Linda and Clive Berger for their abundant support. VI. REFERENCES Alberts, B., Bray, D., Lewis, J., Raff, M., Roberts, K. and Watson, J. D., 1983. Molecular Biology of the Cell. Garland Publishing, New York.

Annis, S. L., Dastoor, F. F!, Ziel, H., ~ a s z a k ,F! and Longcore, J. E., 2004. A DNA-based assay identifies Batrachochytrium dendrabatidis in amphibians. J. Wildl. Dis. 40: 420-428.

.Uexander, M. A. and Eischeid, J. K., 2001. Climate variability in regions of amphibian declines. Cons. Biol. 15: 930-942.

Aplin, K. and Irkpatrick, P, 2000. Chytridiomycosis in southwest Australia: historical sampling documents the date of introduction, rates of spread and seasonal epidemiology, and sheds new light on chytrid ecology. F! 24 in "Getting the Jump! on Amphibian Disease: Conference and Workshop Compendium". Cairns, 26-30 August 2000.

Alford, R. A. and Richards, S. J., 1999. Global amphibian declines: A problem in applied ecology. Ann. Rev. Ecol. Syst. 30: 133-65.

3044

AMPHIBIAN BIOLOGY

Ardipradja, K., 200 1. A study of resistance towards the emerging pathogen Batrachochytrium dendrobatidis in four species of Australian frogs. [Honours thesis]. Melbourne, Australia. University of Melbourne: Melbourne. Aston, R. J., Beattie, R. C. and Milner, A. G. F!, 1987. Characteristics of spawning sites of the common frog (Rana temporaria) with particular reference to acidity. J. Zool. 213: 233-242. Australian Government Department of the Environment and Heritage, 2006. Background Document for the Threat Abatement Plan: Infection of Amphibians with Chytrid Fungus Resulting in Chytridiomycosis. Department of the Environment and Heritage, Commonwealth of Australia, Canberra, Australia, Pp. 70. Available at http://www.deh.gov.au/ biodiversity/threatened/publications/tap/chytrid/

pubs/chytrid-background.pdf Banks, B. and Beebee, T. J. C., 1988. Reproductive success of natterjack toads Bufo calamita in two contrasting habitats. J. An. Ecol. 57: 475-492. Banks, C. and McCracken, H., 2002. Captive management and pathology of sharp snouted dayfrogs, Taudactylw acutirostris, at Melbourne and Taronga zoos. Pp. 94-102 in "Frogs in the Community. Proceedings of the Brisbane Symposium", ed by A. E. 0 Natrass. Queensland Frog Society, East Brisbane. Barr, D. J. S., 1990. Phylum Chytridiomycota. Pp. 454466 in "Handbook of Protoctista", ed by L. Margulis, J. 0 . Corliss, M. Melkonian and D. J. Chapman. Jones and Bartlett, Boston. Barr, D. J. S., 2001. Chytridiomycota. Pp. 93-112. in "The Mycota" 7 4 ed by D. J. McLaughlin, E. G. McLaughlin, and F! A. Lemke. Springer-Verlag, Berlin. Bartnicki-Garria, S. and Sing, YO., 1986. Adhesion of zoospores of Phytophthora to solid surfaces. Pp. 279283. i n "Zoosporic Fungi in Teaching and Research, ed by M. S. Fuller, A. Jaworski. Dept of Botany, University of Georgia, Athens, USA. Beakes, G. W., Wood, S. E. and Burr, A. W., 1994. Features which characterize Saprolegnia isolated from salmonid fish lesions - a review. Pp. 3366. in "Salmon Saprolegniasis", ed by G. J. Mueller. Report Prepared for: U. S. Department of Energy, Bonneville Power Administration, Division of Fish and Wildlife, Portland Oregon (DOEJBP-02836-1) Beattie, R. C., Aston, R. J. and Milner, A. G. F!, 1991. A field study of fertilization and embryonic development in the common frog (Rana temporaria) with particular reference to acidity and temperature. J . Appl. Ecol. 28: 346-357. Bell, B. D., Carver, S., Mitchell, N. J. and Pledger, S., 2004. The recent decline of a New Zealand endemic: how and why did populations of Archey's frog Oiopelma archeyi crash over 1996-2001? Biol. Cons. 120: 189-199. Berger, L. 2001. Diseases in Australian Frogs [PhD thesis]. Townsville, Australia. James Cook University: Townsville. Berger, L., Speare, R., and Kent, A, 2000. Diagnosis of chytridiomycosis in amphibians by histologic examination. Zoos' Print Journal 15: 184-190.

Berger, L., Speare, R. and Humphrey, J., 1997. Mucormycosis in a free-ranging green tree frog from Australia. J. Wildl. Dis. 33: 903-907. Berger, L., Speare, R. and Hyatt, A., 1999. Chytrid fungi and amphibian declines: overview, implications and future directions. Pp. 23-33 in "Declines- and Disappearances of Australian Frogs", ed by A. Campbell. Environment Australia, Canberra. Berger, L., Speare, R. and Skerratt, L., 2005c. Distribution of Batra~hoch~trium dendrobatidis and pathology in the skin of green tree frogs (Litoria caerulea) with severe chytridiomycosis. Dis. Aquat. Org. 68: 65-70. Berger, L., Hyatt, A. D., Speare, R. and Longcore, J. E., 2005a. Life cycle stages of the amphibian chytrid Batrachochyh%im dendmbatidis. Lhs. Aquat. Org. 68: 5 1-63. Berger, L., Maranteh, G., Skerratt, L. F. and Speare, R., 2005b. Virulence of the amphibian chytrid fungus, Batmchochytn'um dendrobatidis, varies with the strain. Drc. Aquat. Org. 68: 47-50. Berger, L., Speare, R. Thomas, A. and Hyatt, A,, 2001. Mucocutaneous fungal disease in tadpoles of Bufo marinus in Australia. J. Herp. 35: 330-335. Berger, L., Hyatt, A., Olsen, V. Boyle, D. Hengstberger, S. Marantelli, G. Humphreys, K. and Longcore, J. E., 2002. Production of polyclonal antibodies to Batrachochytrium dendrobatidis and their use in an immunoperoxidase test for chytridiomycosis in amphibians. fi.Aquat. Org. 48: 213-220. Berger, L., Speare, R., Daszak, P., Green, D. E., Cunningham, A. A., Goggin, C. L., Slocombe, R., Ragan, M. A., Hyatt, A. D., McDonald, K. R., Hines, H. B., Lips, K. R., Marantelli, G. and Parkes, H., 1998. Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proc. Nat. Acad. Sci. USA. 95: 9031-9036. Berger, L., Speare, R., Hines, H., Marantelli, G., Hyatt, A. D., McDonald, K. R., Skerratt, L. F., Olsen, V., Clarke, J. M., Gillespie, G., Mahony, M., Sheppard, N., Williams, C. and Tyler, M., 2004. Effect of season and temperature on mortality in amphibians due to chytridiomycosis. A w t . I4t. J . 82: 31-36. Blaustein, A. R., Hokit, D. G. O'Hara R. K. and Holt, R. A., 1994b. Pathogenic fungus contributes to amphibian losses in the Pacific Northwest. Biol. Cons. 67: 251-254. Blaustein, A. R., Hoffman, F! D., Holut, D. G., hesecker, J. M., Walls S. C. and Hays, J. B., 1994a. UV repair and resistance to solar UV-B in amphibian eggs: a link to population declines? Proc. Nut. Acad. Sci. USA 91: 1791-1795. Blaustein, A. R., Romansic, J. M., Scheessele, E. A, Han, B. A., Pessier, A. I! and Longcore, J. E., Interspecific variation in susceptibility of frog tadpoles to the pathogenic fungus, Batrachochytrium dendrobatidis. Cons. Biol. (in press) Bonaccorso, E., Guayasamin, J. M., Mendez, D. and Speare, R., 2003. Chytridiomycosis as a possible cause of population declines in Atelopus cruciger (Anura: Bufonidae). Herp. Revieul 34: 331-334. Bonin, J., Desgranges, J. L., Rodrigue, J. and Ouellet, M., 1997. Anuran species richness in agricultural landscapes of Quebec: Foreseeing long-term results of road call surveys. H e Cons. 1: 141-148.

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS Bosch, J., Martinez-Solano, I. and Garcia-Paris, M., 2001. Evidence of a chytrid fungus infection involved in the decline of the common midwife toad (Alytes obstetricam) in protected areas of central Spain. Biol. Cons. 97: 331-337. Boyle, D. G., Boyle, D. B., Olsen, V , Morgan, J. A. and Hyatt, A. D., 2004. Rapid quantitative detection of chytridiomycosis (Batrachochytrium dendrobatzdis) in amphibian samples using real-time Taqman PCR assay. Dis. Aquat. Org. 60: 141-8. Boyle, D. G., Hyatt, A. D., Daszak, F!, Berger, L., Longcore, J. E., Porter, D., Hengstberger, S. G. and Olsen, V., 2003. Cryo-archiving of Batrachochytrium dendrobatidis and other Chytridiomycetes. Dis. Aquat. Org. 56: 59-64. Bradley G. A., Rosen, P C., Sredl, M. J., Jones, T R. and Longcore, J. E., 2002. Chytridiomycosis in native Arizona frogs. J. Wildl. Dis. 38: 206-212. Bragg, A. N., 1962. Saprolegnia on tadpoles again in Oklahoma. Southwest Nut. 7: 79-80. Bragg, A. N. and Bragg, W. N., 1958. Parasitism of spadefoot toads by Saprohgnia. Herpetolopa. 14: 34. Briggs, C. and Burgin, S., 2003. A rapid technique to detect chytrid infection in adult frogs. Herp. Reuieui 34: 124-126. Broz, 0 . and Privora, M., 1951. Two skin parasites of Rana temporaria: Dermocystidium ranae Guyenot & Naville and Dermosporidium granulosum n. sp. Parasitology. 42: 65-69. Bullis, R. A., Noga, E. J. and Levy, M. G., 1990. Immunological relationship of the fish-pathogenic oomycete Saprolegnia parasitica to other oomycetes and unrelated fungi. J. Aquat. Anim. Health 2: 223227. Burrowes, F! A., Joglar, R. L. and Green, D. E., 2004. Potential causes for amphibian declines in Puerto Rico. Herpetologica 60: 141- 154. Carey, C., Cohen, N., and Rollins-Smith, L. A., 1999. An immunological Amphibian declines: perspective. Develop. Comp. Immunol. 23: 459-472. Carey, C., Bradford, D. F., Bmnner, J . L, Collins, J. F!, Davidson, E. W., Longcore, J. E., Ouellet, M., Pessier, A. and Schock, D. M., 2003. Biotic factors in amphibian population declines. Pp. 153-208 in "Multiple stressors and declining amphibian populations", ed by G. Lindre, S. K. Krest and D. W. Sparling. Society for Environmental Toxicolog and Chemistry Press, Pensacola, Florida. Cheville, N. F., 1996. Ultrastmctural Pathology. An Introduction to Interpretation. Iowa State University Press, Ames. Cicmanec, J. L., Ringler, D. H. and Beneke, E. S., 1973. Spontaneous occurrence and experimental transmission of the fungus, Fmecaea pedratoi, in the marine toad, Bufo marinus. Leb. Anim. Sci. 23: 43-47. Connolly, J. H., Obendorf, D. L. and Whittington, R. J., 1999. Haematological, serum biochemical and serological features of platypuses with and without . mycotic granulomatous dermatitis. Aust. Vet. J. 77: 809-813. Creeper, J. H., Main, D. C., Berger, L., Huntress, S. and Boardman, W., 1998. An outbreak of mucormycosis in slender tree frogs (Litoria. adelensis) and whitelipped tree frogs (Litmia infiafienata). Awt. %t. J. 76: 761-762.

3045

Cunningham, A. A., Langton, T E. S., Bennett, l? M., Lewin, J. F., Drury, S. E. N., Gough, R. E. and MacGregor S. K., 1996. Pathological and microbiological findings from incidents of unusual mortality of the common frog (Rana temporaria). Phil. Trans. Roy. Soc. of London B. 351: 1539-1557. Cunningham, C. 0 . 2002. Molecular diagnosis of fish and shellfish diseases: present status and potential use in disease control. Aquaculture 206: 19-55. Czeczuga, B., Muszyiiska, E. and Krzemiiiska, A., 1998. Aquatic fungi growing on the spawn of certain amphibians. Amphibia-Reptilia 19: 239-25 1. Daszak, F!, Cunningham, A. A,, and Hyatt, A. D., 2001. Draft guidelines for international translocation of amphibians with respect to infectious diseases. Attachment 6. Pp. 172-178. In "Developing management strategies to control amphibian diseases: Decreasing the risks due to communicable diseases", ed by Speare, R. and Steering Committee of Getting the Jump! on Amphibian Disease. School of Public Health and Tropical Medicine, James Cook University, Townsville. Daszak, F!, Berger, L., Cunningham, A. A., Hyatt, A. D., Green, D. E., and Speare, R., 1999. Emerging infectious diseases and amphibian population declines. Emerg. Znt Dis. 5: 735-748. Daszak, P, Strieby, A., Cunningham, A. A., Longcore, J. E., Brown, C. C. and Porter, D., 2004. Experimental evidence that the bullfrog (Rana catesbeiana) is a an emerging potential carrier of ~h~tridiomycosis, fungal disease of amphibians. Herp. J. 14: 201-207. Davidson, C., Shaffer, H. B., and Jennings, M. R., 2002. Spatial tests of the pesticide drift, habitat destruction, UV-B and climate-change hypotheses for California amphibian declines. Cons. Biol. 16: 1588-1601. Davidson, E. W., Parris, M., Collins, J. E, Longcore, J. E., Pessier, A. P, and Brunner, J., 2003. Pathogenicity and transmission of chymdiomycosis in tiger salamanders (Ambystoma t i e n u r n ) . COP& 2003: 601407. Dawson, P and Weste, G., 1989. Changes in the distribution of Phytophthora cinnamomi in the Brisbane Ranges National Park between 1970 and 1980-81. Aurt. J. Bot. 33: 309-315. Department of Environment and Conservation NSW. 2004. Green and golden bell frog Litoria aurea (Lesson 1829). Draft Recovery Plan. Unpublished report. Dhaliwal, S. S. and Griffiths, D. A,, 1963. Fungal disease in Malayan toads: an acute lethal inflammatory reaction. Nature. 197: 467-569. DiGiacomo, R. F. and Koepsell, T D., 1986. Sampling for detection of infection or disease in animal populations. J. Am. Vet. Med. Assoc. 189: 22-23. Drost, C. A. and Fellers, G. M., 1996. Collapse of a regional frog fauna in the Yosemite area of the Cahfornia Sierra Nevada, USA Cow. Bwl. 10: 414425. Druly, R. B. and Wallington, E. A., 1980. "Carleton's Histological Technique". Oxford University Press, Oxford. Elkan, E. and Philpot, C. M., 1973. Mycotic infections in frogs due to a Phialophora-like fungus. Sabouraudia. 11: 99-105.

3046

AMPHIBIAN BIOLOGY

Erspamer, V 1994. Bioactive secretions of the amphibian integument. Pp. 178-350 in "The Integument", vol. 1 of "Amphibian Biology", ed by H. Heatwole, G. T. Barthalmus and A. Heatwole. Surrey Beatty and Sons, Chipping Norton. Fellers, G. M., Green, D. E. and Longcore, J. E., 2001. Oral chytridiomycosis in the mountain yellowlegged frog (Rana mwcosa). Copcia 4001: 945-953. Ferri, V, 2002. La micosi cutanea degli anfibi anche in Italia. Bufo News 1/2002: 1 4 . Fox, H., 1994. The structure of the integument. Pp. 132, i n "Amphibian Biology, Volume 1: The Integument", ed by H. Heatwole, G. T. Barthalmus. Surrey Beatty and Sons, Chipping Norton, Australia. Frank, W., 1976. Mycotic infections in amphibians and reptiles. Pp. 73-88 in "Proceedings of the Third International Wildlife Disease Conference" ed L. A. Page. Plenum Press, New York. Frank, W., Roester, U. and Scholer, H. J., 1974. Sphaerule formation by a Mucor species in the internal organs of amphibia. Pp. 161-192, in "A Century of Mycology" ed by B. Sutton. Cambridge University Press. Gillespie, G. and Hero, J.-M., 1999. Potential impacts of introduced fish and fish translocations on Australian amphibians. Pp. 131-144, in "Declines and Disappearances of Australian Frogs", ed by A. Campbell. Environment Australia, Canberra. Gillespie, G. and Hines, H., 1999. Status of temperate riverine frogs in south-eastern Australia. Pp. 109130, in "Declines and Disappearances of Australian Frogs", ed by A. Campbell. Environment Australia, Canberra. Gillespie, G. and Marantelli G., 2000. The role of the amphibian chytridiomycete fungus in population dynamics of the spotted tree frog (Litoria spenceri) a declining riverine species in south-eastern Australia. I? 45, in "Getting the Jump! on Amphibian Disease: Conference and Workshop Compendium", Cairns, 26-30 August 2000. Grayburn, W. S., Hudspeth, D. S. S., Gane, M. K. and Hudspeth, M. E. S. 2004. The mitochondria1 genome of Saprolegnia ferax: organization, gene content and nucleotide sequence. Mycologia 96: 981-989. Green, A. J., 1999. Implications of pathogenic fungi for life-history evolution in amphibians. Func. Ecol. 13: 573-575. Green, D. E., 2001. Pathology of Amphibia. Chapter 27 in "Amphibian Medicine and Captive Husbandry", ed by K. M. Wright and B. R. Whitaker. Krieger Publishing, Malabar, Florida. Green, D. E. and Kagarise-Sherman, C., 2001 . Diagnostic histological findings in Yosemite toads (Bufo can-) from a die-off in the 1970's. J. Herp. 35: 92-103. G m n , D. E., h d r e w s , J., Abell, J. M. 1995. Preliminary lnvesugations on a mycotic myositis in red-spotted newts (Notophthulmw v. viridescens) from Vermont. Pp. 31-62, in: "CinquiSme Colloque International de hthologie des Reptiles et des Amphibiens", ed E Zwart and G. Matz. Alphen Aan den Rijn, The Netherlands.

Green, D. E., Converse, K. A. and Schrader, A. K., 2002. Epizootiology of sixty-four amphibian morbidity and mortality events in the USA, 1996-2001. Ann. N.Y Acad. Sci. 969: 323-339. Groff, J. M., Mughannam, A., McDowell. T S., Wong. A., Dykstra. M. J., Frye, F. L. and Hedrick, R. E, 1991. An epizootic of cutaneous zygomycosis in cultured dwarf African clawed frogs (Hymenochirw curtipes) due to Basidiobolus ranarum. J. Med. E t . Mycol. 29: 215-223. Hanselmann, R., Rodriguez, A., Lampo, M., FajardoRamos, L., Aguirre, A. A., Kilpatrick, A. M., Rodriguez, J. El and Daszak, P, 2004. Presence of an emerging pathogen of amphibians in introduced bullfrogs Rana catesbeiana in Venezuela. Biol. Corn. 120: 115-119. Herman, R. L., 1984. Zchthyophonus -like infection in newts (Notophthulmw viridescens Rafinesque).J. wldl. Dir. 20: 55-56. Herrera, R. A., Steciow, M. M. and Natale, G. S., 2005. Chytrid fungus parasitizing the wild amphibian hptodactylus ocellatus (Anura: Leptodactylidae) in Argentina. Dir. Aquat. 0%. 64: 247-252. Heyer, W. R., Rand, A. R., Goncalvez da Cruz, C. A. and Peixoto, 0 . L., 1988. Decimations, extinctions, and colonizations of frog populations in southeast Brazil and their evolutionary implications. Biotropica 20: 230-235. Hines, H., Mahony, M. and McDonald, K., 1999. An assessment of frog declines in wet subtropical i n "Declines and Australia. Pp. 44-63 Disappearances of Australian Frogs", ed by A. Campbell. Environment Australia, Canberra. Hopkins, S. and Channing, A. 2003. Chytrid fungus in northern and western cape frog populations, South Africa. Herp. Review 34: 334-336. James, T Y., Porter, D., Leander, C. A., Vilaglys, R. and Longcore, J . E. 2000. Molecular phylogenetics of the Chytridiomycota supports the utility of ultrastructural data in chytrid systematics. Can. J. Bot. 78: 336-350. Jay, J. M. and Pohley, W. J., 1981. Dermosporzdium penneri sp. n. from the skin of the American toad, Bufo americanus (Amphibia: Bufonidae). J. Parasit. 67: 108-1 10. Johnson, M. and Speare, R., 2003. Survival of Batrachochytrium in water: quarantine and disease control implications. Emerg. Infect. Dir. 9: 922-925. Johnson, M and Speare, R., 2005. Possible modes of dissemination of the chytrid fungus Batrachochytrium dendrobatidis in the environment. Dir. Aquat. Org. 65: 181-186. Johnson, M., Berger, L., Philips, L. and Speare, R., 2003. Fungicidal effects of chemical disinfectants, W light, desiccation and heat on the amphibian chytrid, Batrachochytrium dendrobatidis. Dis. Aquat. Org. 57: 255-260. Johnson, T W. Jr., Seymour, R. L. and Padgett, D. E., 2002. Biology and systematics of the Saprolegniaceae. On-line publication accessible at http://aa.uncw.edu/digilib/biology/fungi/ taxonomy%20and%20systematics/padgett%2Obookl Josefsson, M. and Andersson, B., 2001. The environmental consequences of alien species in the Swedish lakes Malaren, Hjalmaren, Vanern and Vattern. Ambio. 30: 514-2 1 .

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS Kagarise-Sherman, C. and Morton M. L., 1993. Fbpulation declines of Yosemite toads in the eastern Sierra Nevada of California. J. Herp. 27: 186-198. Kiesecker, J. M. and Blaustein, A. R., 1995. Synergism between UV-B radiation and a pathogen magnifies amphibian embryo mortality in nature. Proc. Nut. Acad. Sci. USA 92: 11049-1 1052. Kiesecker, J. M. and Blaustein, A. R., 1997. Influences of egg laying behavior on pathogenic infection of amphibian eggs. Cons. Biol. 11: 214-220. Kiesecker, J. M., Blaustein, A. R. and Beldon, L. K., 2001a. Complex causes of amphibian population declines. Nature 410: 681-684. Kiesecker, J. M., Blaustein, A. R. and Miller, C. L., 2001b. Transfer of a pathogen from fish to amphibians. Cons. Biol. 15: 1064-1070. Lamirande, E. W. and Nichols, D. K., 2002. Effects of host age on susceptibility to cutaneous chytridlomycosis in blue and yellow poison dart frogs (Dendrobates tinctorius). Pp. 3-13 in "Proceedings of the Sixth International Symposium on the Pathology of Reptiles and Amphibians", ed by R. G. McKinnell and D. L. Carlson. Saint Paul, Minnesota. Laurance, W.F., McDonald, K. R. and Speare R., 1996. Epidemic disease and the catastrophic decline of Australian rain forest frogs. Cons. Biol. 10: 406413. Lefcort, H., Hancock, K. A,, Maur, K. M. and Rostal, D. C., 1997. The effects of used motor oil, silt, and the water mold Sapolegnia parasitica on the growth and survival of mole salamanders (genus Ambystoma). Arch. Enuiron. Contam. Toxicol. 32: 383-388. Letcher, I? M. and Powell, M. J. 2004. Kappamyces, a new genus in the Chytridiales (Chytridiomycota). Noua Hedwigia in press.

3047

Longcore, J. E., 1995. Morphology and zoospore ultrastructure of Entophlyctis luteolus sp. nov. (Chytridiales): Implications for chytrid taxonomy. Mycologaa 87: 25-33. Longcore, J. E., Pessier, A. I? and Nichols, D. K., 1999. Ba.trmhochyCrium dmdrobatidis gen. et sp. nov., a chytrid pathogenic to amphibians. Mycolop 91: 219-227. Lutzoni, F., Kauff, F., Cox, C.J., McLaughlin, D., Celio, G., Bentinger, B., et al. (44 authors) 2004. Assembling the fungal tree of life: progress, classification, and evolution of subcellular traits. Am. J. Bot. 91: 1446-1480. Lynch, M. 2001. Amphibian quarantine protocols: Melbourne Zoo. Attachment 6. Pp. 179-183, in "Developing management strategies to control amphibian diseases: Decreasing the risks due to communicable diseases", ed by Speare, R and Steering Committee of Getting the Jump! on Amphibian Disease. School of Public Health and Tropical Medicine, James Cook University, Townsville. Mahony, M. J., 1996. The decline of the green and golden bell frog Litoria aurea viewed in the context of declines and disappearances of other Australian frogs. Awt. Zool. 30: 237-247. Mahony, M., Clulow, J., Browne, R. and Pomering, M., 1999. Declines and disappearances of frogs: risk assessment and contingency strategies. Pp. 202-2 11 in "Declines and Disappearances of Australian Frogs", ed by A. Campbell. Environment Australia, Canberra. Marantelli, G. and Berger, L., 2000. Quarantine methods for captive collections. P 26, in "Getting the Jump on Amphibian Disease: Conference and Workshop Compendium", Cairns, 26-30 August 2000.

Letcher, P. M., Powell, M. J., Chambers, J . G., and Holznagel, U! E., 2004. Phylogenetic relationships among Rhizophydium isolates from North America and Australia. Mycologia 96: 1363-1375.

Marantelli, G. and Hobbs, R., 2000. Disease vectors and the community: from lost frogs to frog friendly gardens - how do we help frogs while containing disease spread? Pp. 49, in Getting the Jump! on Amphibian Disease: Conference and Workshop Compendium. Cairns, 26-30 August 2000.

Lilley, J. H., Hart, D., Panyawachira, V Kanchanakhan, S, Chinabut, S., Soderhall, K. and Cerenius, L., 2003. Molecular characterization of the fishpathogenic fungus Aphanomyces inuadans. J. Fish Dir. 26: 263-275.

Marantelli, G., Berger, L. and McInnes, K., 2000. Investigations into the matment of Batrathochytrium in frogs and tadpoles. Pp. 27, in Getting the Jump! on Amphibian Disease: Conference and Workshop Compendium. Cairns, 26-30 August 2000.

Lips, K. R., 1998. Decline of a tropical montane fauna. Cons. Biol. 12: 106-117.

Maranteh, G., Berger, L., Speare, L. and Keegan, L., 2004. Distribution of Batrachochytrium dendrobatidk and keratin during tadpole development. Pac. Cons. Biol. 10: 173-179.

Lips, K. R., 1999. Mass mortality and population declines of anurans at an upland site in western Panama. Cons. Biol. 13: 117-125. Lips, K. R. Green D. E. and Papendick, R., 2003. Chytridiomycosis in wild frogs from southern Costa Rica.J. Herp. 37: 215-218. Lips, K. R., Reeve, J. D. and Witters, L. R., 2003. Ecological traits predicting amphibian population declines in Central America. Cons. Biol. 17: 107% 1088. Lips, K. R., Mendelson, J. R., Mufioz-Alonso, A,, Canseco-Mgrquez, L. and Mulcahy, D. G., 2004. Amphibian population declines in montane southern Mexico: resurveys of historical localities. Biol. Cons. 119: 555-564. Longcore, J. E., 1993. Morphology and zoospore ultrastructure of Lacwtromyces hiemalis gen. et sp. nou. (Chytridiales). Can. J. Bot. 71: 414425.

May, R., 1988. Conservation and Disease. Cons. Biol. 2: 28-30. Mazzoni, R., Cunningham, A. A., Daszak, I?, Apolo, A., Perdomo, E. and Speranza, G., 2003. Emerging pathogen of wild amphibians in frogs (Rana catesbeiana) farmed for international trade. Emerg. Inj? Dis. 9: 995-998. McDonald, K. R., 1992. Distribution patterns and conservation status of north Queensland rainforest frogs. Conservation Technical Report No. 1. Queensland Department of Environment and Heritage, Brisbane. McDonald, K. and Alford, R., 1999. A review of declining frogs in northern Queensland. Pp. 14-22 in "Declines and Disappearances of Australian Frogs", ed by A. Campbell. Environment Australia, Canberra.

3048

AMPHIBIAN BIOLOGY

McDonald, K. R., Mkndez, D., Miiller, R., Freeman, A. B. and Speare, R., 2005. Decline in the prevalence of chytridiomycosis in upland frog populations in North Queensland, Australia. Pac. Cons. Biol. 11: 114-120. Mikaelian, I., Ouellet, M., Pauli, B., Rodrigue, J., Harshbarger, J. C. and Green, D. M., 2000. Ichthyophonw - like infection in wild amphibians from Quebec, Canada. Dis. Aquat.Org. 40: 195-201. Miller, C. E. and Dylewski, D. F!, 1981. Syngamy and resting body development in Chytriomyces hyalius (Chytridiales). Am. J. Bat. 68: 342-349. Morehouse, E. A,, James, T Y., Ganley, A. R. D., Vilgal~, R., Berger, L., Murphy, F! J. and Longcore, J. E., 2003. Multilocus sequence typing suggests the chytrid pathogen of amphibians is a recently emerged clone. Mol. Ecol. 12: 395403. Muths, E., Corn, F! S., Pessier, A. P and Green, D. E., 2003. Evidence for disease-related amphibian decline in Colorado. Biol. Cons. 110: 357-365.

Parris, M. J., 2004. Hybrid response to pathogen infection in interspecific crosses between two amphibian species (Anura: Ranidae). Euol. Ecol. Res. 6: 457-471. Parris, M. J. and Baud, D. R., 2004. Interactive effects of a heavy metal and chytridiomycosis on^gray treefrog larvae (Hyla chrysoscelis). Copeia 2004: 344-350. Parris, M. J. and Beaudoin, J. G., 2004. Chytridiomycosis impacts predator-prey interactions in larval amphibian communities. Oecologia 140: 626-632. Parris, M. J. and Cornelius, T. O., 2004. Fungal pathogen causes competitive and developmental stress in larval amphibian communities. Ecology 85: 3385-3395. Pascolini, R., Daszak, P , Cunningham, A. A., Tei, S., Vagnetti, D., Bucci, S., Fagotti, A. and Di Rosa, I., 2003. Parasitism by Dermocystidium ranae in a population of Rana esculenta complex in Central Italy and description of Amphibiocystidium n. gen. Dis. Aquat. Org. 56: 65-74.

Mutschmann, F., Berger, L., Zwart, P, and Gaedicke, C., 2000. Chytridiomycosis in amphibians - first report from Europe. Berliner und Miinchener Tierarztliche Wochenschrzft. 113: 380-383.

Pence, D. B. and Ueckermann, E., 2002. Sarcoptic mange in wildlife. Rev Sci Tech. 21: 385-98.

New South Wales National Parks and Wildlife Service, 2000. Threatened Species Management, Information circular No. 6, Hygiene protocol for the control of disease in frogs. NSWNPWS, May 2000.

Pessier, A. l?, Nichols, D. K., Longcore, J. E. and Fuller, M. S., 1999. Cutaneous chytridiomycosis in poison dart fmgs (Dendrobates spp.) and White's tree frogs (Litoria caerulea).J. Kt. Diag. Invest. 11: 194-199.

Nichols, D. K. and Lamirande, E. W., 2000. Treatment of cutaneous chytridiomycosis in blue-and-yellow poison dart frogs (Dendrobates tinctorius). Pp. 51 in "Getting the Jump on Amphibian Disease: Conference and Workshop Compendium", Cairns, 26-30 August 2000.

Piotrowski, J. S., Annis, S. L. and Longcore, J. E., 2004. Physiology of Batrachochytrium dendrobatidis, a chytrid pathogen of amphibians. Mycologia 96: 9-15.

Nichols, D. K., Lamirande, E. W., Pessier, A. P. and Longcore, J. E., 2001. Experimental transmission of cutaneous chymdiomycosis in dendmbatid frogs. J. Wild.!. Dir. 37: 1-11. Nix, H.A., 1986. A biogeogaphic analysis of Australian Elapid snakes. Pp. 4-15 in "Atlas of Australian Elapid Snakes", ed by R. Longmore. Australian Flora and Fauna Series Number 7. Australian Government Publishing Service, Canberra Obendorf, D., 2000. Chytrid fungus Batrachochyrium dendrobatidis detected in free-living Tasmanian frog populations. Australian Wildlife Health Network: Situation Report (Update -January 2005). Obendorf, D. L., Peel, B. F. and Munday, B. L., 1993. Mucor amphibiorum infection in platypus (Ornithorhynchus anatinus) from Tasmania. J . Wildl. Dis. 29: 485487. Oevermann von, A., Schildger, B., Feldman, S. and Robert, N., 2005. Chytridiomycosis in amphibians (Dyscophus antongilii) in Switzerland. Tierarztliche Umchau 60: 21 1-217. Olsen, V., Hyatt, A. D., Boyle, D. G. and Mendez, D., 2004. Co-localisation of Batrachochytrium dendrobatzdis and keratin for enhanced diagnosis of chytridiomycosis in frogs. Dis. Aquat. 0%. 61: 85-88. Parker, J. M., Mikaelian, I., Hahn, N. and Diggs, H. E., 2002. Clinical diagnosis and treatment of epidermal chytridiomycosis in African clawed frogs (Xenopus trqpualis). Comp. Med. 52: 265-268.

Plowright, W., 1982. The effects of rinderpest and rinderpest control on wildlife in Africa. Animal Disease in Relation to Conservation. Pp. 1-27 in "Proceedings of a Symposium at the Zoological Society of London, November 19817', ed by M. A. Edwards and U. McDonnell. Academic Press, London. 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. Pounds, J. A., Fogden, M. F! L. and Campbell, J . H., 1999. Biological response to climate change on a tropical mountain. Nature 398: 61 1-615. Powell, M. J., 1993. Looking at mycology with a Janus face: A glimpse of chytridiomycetes active in the environment. Mycologia 85: 1-20. Puschendorf, R., 2003. Atelopus uarius (Harlequin frog). Fungal infection. Herp. Review 34: 355. Rachowicz, L. J., 2002. Mouthpart pigmentation in R a m muscosa tadpoles: seasonal changes without chytridiomycosis. Herp. Rev. 33: 263-265. Rachowicz, L. J. and Vredenburg, V. T., 2004. Transmission of Batrachochytrium dendrobatzdis within and between amphibian life stages. Dis. Aquat. Org. 61: 75-83. Ragan, M. A., Goggin, C. L., Cawthorn, R. J., Cerenius, L., Jamieson, A. V., Plourde, S. M., Rand, T. G., Soderhall, K. and Gutell, R. R., 1996. A novel clade of protistan parasites near the animal-fungal divergence. Proc. Natl. Acad. Sci. U S A. 93: 1190711912.

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

3049

Reed, K. D., Ruth, G. R., Meyer, J. A. and Shukla, S. K., 2000. Chlamydia pneumoniae infection in a breeding colony of African clawed frogs (Xenopus tropicalis). Emerg. Infect. Dis. 6: 196-199.

Skerratt L. F., Middleton, D. and Beveridge, I., 1999. Distribution of life cycle stages of Sarcoptes scabzei var wombati and effects of severe mange on common wombats in Victoria.J. Wldl. Dis. 35: 633-646.

Reichenbach-Klinke, H. and Elkan, E., 1965. Amphibia. Pp. 209-384, in "The Principal Diseases of Lower Vertebrates", Academic Press, New York.

Sparrow, F. K., 1960. Aquatic Phycomycetes (second revised edition). University of Michigan Press, Ann Arbor.

Retallick, R. and Dwyer, L., 2000. Using translocations to learn about frog declines and disease. Pp. 41 in "Getting the Jump! on Amphibian Disease: Conference and Workshop Compendium", Cairns, 26-30 August 2000.

Speare, R., 1995. Preliminary study on diseases in Australian Wet Tropics amphibians. Deaths of rainforest frogs at 0 ' Keefe Creek, Big Tableland. Final report to Queensland Department of Environment and Heritage. Unpublished report, Queensland Department of Environment and Heritage, Brisbane.

Retallick, R. W., 2003. Bioclimatic investigations into the potential distribution of Batrachochytrium dendrobatzdis in Australia, Draft Report. University of Queensland, Australia. Retallick, R. W., McCallum, H. and Speare, R., 2004. Endemic infection of the amphibian chytrid fungus in a frog community post-decline. PLoS Biol2: e35 1. Richards, S. J., McDonald, K. R. and Alford, R. A., 1993. Declines in populations of Australia's endemic tropical rainforest frogs. Pac. Cons. Biol. 1: 66-77. Robinson, J., Griffiths, R. A. and Jeffries, P, 1999. Infectious disease and amphibian population dynamics: is egg mortality significant? Froglog 33: 1. Robinson, J., Griffiths, R. A. and Jeffries, P, 2003. Susceptibility of frog (Rana temporaria) and toad (Bufo bufo) eggs to invasion by Saprolegniu. AmphibioReptilia 24: 261-268. Rollins-Smith, L. A,, Reinert, L. K., Miera, \'. and Conlon, J . M., 2002b. Antimicrobial peptide defenses of the Tarahumara frog, Rana tarahumarae. Biochem. Biophys. Res. Comm. 297: 361-367. Rollins-Smith, L. A., Carey, C., Longcore, J., Doersam, J. K., Boutte, A., Bruzgal, J. E. and Conlon, J. M., 2002a. Activity of antimicrobial s h n peptides from ranid frogs against Batra~hoch~trium dendrobatzdis, the chytrid fungus associated with global amphibian declines. Dev. Comp. Immunol. 26: 471479. Ron, S. and Merino, A,, 2000. Amphibian declines in Ecuador: overview and first report of chytridiomycosis from South America. Froglog 42: 4. Rush, H. G., Anver, M. R. and Beneke, E. S., 1974. Systemic chromomycosis in Rana pzpiens. Lab. Anim. Sci. 24: 646-655. Scott, G. R., 1981. Rinderpest. Pp. 18-30 in "Infectious Diseases of Wild Mammals", ed by J. W. Davis, L. H. Karstad and D. D. Trainer. Iowa State University Press, Iowa City. Scott, M. E., 1988. The impact of infection and disease on animal populations: Implications for conservation. Cons. Biol. 2: 40-56. Scott, N. J., 1993. Postmetamorphic Death Syndrome. Froglog 7 : 1. Seymour, R. and Fuller, M. S., 1987. Collection and isolation of water molds (Saprolegniaceae) from water and soil. Pp. 125-127 in "Zoosporic Fungi in Teaching and Research", ed by M. S. Fuller and A. Jaworski. Southeastern Publishing Corporation, Athens, Georgia. Skerratt, L. F., Martin, R. W. and Handasyde, K. A., 1998. Saxoptic mange in wombats. Awt. Rt. J. 76: 408-410.

Speare, R., 2000. Understanding the pieces of the amphibian disease puzzle. Pp. 19 in "Getting the Jump! on Amphibian Disease: Conference and workshop compendium", Cairns, 26-30 August 2000. Speare, R., 200 1. Summary of formidable infectious diseases of amphibians. Pp. 123-130 in "Developing management strategies to control amphibian diseases: Decreasing the risks due to communicable diseases", ed by R. Speare and Steering Committee of Getting the Jump! on Amphibian Disease. School of Public Health and Tropical Medicine, James Cook University, Townsville. URL: http://~~~.jcu.edu.aulschooVphtmi PHTM/frogs/fiamp.htm. Speare, R., 2002. Mucormycosis of amphibians. Amphibian Disease Home Page. Accessed at: http:/ /~~w.jcu.edu.au/schooVphtm/PHTM/frogs/mucor/ mucoramphibiorum. hrm S p e a ~R. and Core IVorking Group of Getting the Jump on Amphibian Disease, 1001. Nomination for listing of amphibian chytridiomycosis as a key threatening process under the Environment Protection and Biodiversity Conservation Act 1999. Pp. 186-208 in "Developing Management Strategies to Control Amphibian Diseases: Decreasing the Risks Due to Communicable Diseases", ed by R. Speare and Steering Comm~ttee of Getting the Jump! on Amphibian Disease, School of Public Health and Tropical Medicine, James Cook University, Townsville. Speare, R. and Berger, L., 2005a. Chytridiomycosis in amphibians in Australia. http://www.jcu.edu.au/

schooVphtmlPHTM/frogs/chyspec.htm. Speare, R. and Berger, L., 2005b. Global distribution of chytridiomycosis in amphibians. Amphibian Disease Home Page. Accessed at: http://www.jcu.edu.au/ schooVphtmlPHTM/frogs/chyglob.htm. Speare, R., Thomas, A. D., O'Shea, F! and Shipton, W. A,, 1994. Mucor amphibiorum in the cane toad, Bufo marinw, in Australia. J. Wildl. Dis. 30: 399-407. Speare, R., Berger, L., O'Shea, P, Ladds, P W. and Thomas, A. D., 1997. Pathology of mucormycosis of cane toads in Australia.J. Wldl. Dis. 33: 105-1 13. Speare, R., Berger, L., Skerratt, L. F., Alford, R., Mendez, D., Cashins, S., Kenyon, N., Hauselberger, K. and Rowley, J., 2004. Hygiene protocol for handling amphibians in field studies. Available at: http://www.jcu.edu.au/schooVphtmlPHTM/frogs/fieldhygiene.pdf Strijbosch, H., 1979. Habitat selection of amphibians during their aquatic phase. Oikos 33: 363-372.

3050

AMPHIBIAN BIOLOGY

Swisher, B., 2002. Microorganisms. Chapter 16 in 'Theory and Practice of Histological Techniques" (5th edition). Churchill Livingston, London.

Weldon, C., du Preez, L. H., Miiller, R., Hyatt, A. D. and Speare, R., 2004. Origin of the amphibian chytrid fungus. Emerg. Inf Dir. 10: 2100-2 105.

Tainter, F. H., 2003. Perspectives and challenges from the 20th century. Phytopathology 93: 1056-1061.

Weygoldt, ,!F 1989. Changes in the composition of mountain stream frog communities in the Atlantic mountains of Brazil: Frogs as indicators of environmental deterioration? Studies in Neotropical Fauna and Environment 243: 249-255.

Taylor, S. K., 2001. Mycoses. Pp. 181-191, in "Amphibian Medicine and Captive Husbandry", ed by K. M. Wright and B. R. Whitaker. Krieger Publishing, Malabar, Florida. Taylor, S. K., Williams, E. S., Pier, A. C., Mills, K. W. and Bock, M. D., 1999b. Mucormycotic dermatitis in captive adult Wyoming toads. J. Wildl. Dis. 35: 70-74. Taylor, S. K., Williams, E. S., Thorne, E. T., Mills, K. W., Withers, D. I. and Pier, k c . , 1999a. Causes of mortality of the Wyoming toad.J. Wzldl. Dir. 35: 49-58. Tyler, M. J., 1997. "The Action Plan for Australian Frogs". Environment Australia, Canberra. Unestam, T., 1973. Fungal diseases of Crustacea. Revieul of Medical and Eterinary Mycology 8: 1-20. Villa, J., 1979. Two fungi lethal to frog eggs in Central America. Copeia 1979: 650-655. Vonesh, J. R. and de la Cruz, O., 2002. Complex life cycles and density dependence: assessing the contribution of egg mortality to amphibian declines. Oecologia 133: 325-333. Waldman, B., 2002. Chytridiomycosis and New Zealand frog population declines. In "Ecology 2002", Cairns, 3 December 2002. Waldman, B., van de Wolfshaar, K. E., Klena, J . D., Andjic, V., Bishop, P J. and Norman, J de B. R., 2001. Chytridiomycosis in New Zealand frogs. Surveillance 28: 9-1 1. Warkentin, K. M., Currie, C. R. and Rehner, S. A,, 2001. Egg-killing fungus induces early hatching of RedEyed Treefrog eggs. Ecology 82: 2860-2869. Warner, R. E., 1968. The role of introduced diseases in the extinction of the endemic Hawaiian avifauna. The Condor 70: 101-120. Weldon, C. and du Preez, L. H., 2004. Decline of the Kihansi Spray Toad, Nectophrynoides asperginis, from the Udzungwa Mountains, Tanzania. Froglog 62: 2-3.

Whittington, R. J., Jones, J. B., Hine, l? M. and Hyatt, A. D., 1997. Epizootic mortality in the pilchard Sardinops sagax neopilchardus in Australia and New Zealand in 1995. I. Pathology and epizootiology. Dis. Aquat. Org. 28: 1-16. Williams, S. E. and Hero, J.-M., 1998. Rainforest frogs of the Australian Wet Tropics: guild classification and the ecological similarity of declining species. Proc. R. Soc. Lond. B 265: 597-602. W~lloughby,L. G., Roberts, R. J. and Chinabut, S., 1995. Aphanomyces invaduns sp. nov., the fungal pathogen of freshwater tropical fish affected by epizootic ulcerative syndrome. J. Fish Dis. 18: 273-275. Wills, R. T., 1993. The ecological impact of Phytophthora cinnamomi in the Stirling Range National Park, Western Australia. Awt. J. Ecol. IS: 145-159. Woodhams, D. C. and AIford, R. A., 2005. Ecology of chytridiomycosis in rainforest stream frog assemblages of tropical Queensland. Cons. Biol. 19: 1449-1459. Woodhams, D. C., Alford, R. A., and Marantelli, G., 2003. Emerging disease cured by elevated body temperature. Dis. Aquat. Org. 55: 65-67. Wright, K. M. and Whitaker, B. R., 2001. Amphibian Medicine and Captive Husbandry. Krieger Publishing. Malabar, Florida. Yoon, H. S., Hackett, J. D., Pinto, G. and Bhattacharya, D., 2002. The single, ancient origin of chromist plastids. Proc. Natl. Acad. Sci. USA 99: 15507-15512. Young, B. E., Lips, K. R., Reaser, J. K., Ibiriez, R., Salas, A. W., Cedeiio, J. R., Coloma, L. A., Ron, S. R., Marca, E. L., Meyer, J. A., Muiioz, A., Bolaiios, F., Chaves, G. and Romo, D., 2001. Population declines and priorities for amphibian conservation in Latin America. Cons. Biol. 15: 12 13-1223.

BERGER ET AL: FUNGAL DISEASES OF AMPHIBIANS

APPENDIX 1 Instructions for Isolating and Culturing Batrachochytrium dendrobatzdis Collection of Amphibians

Collect dead (including road-lulled) and moribund amphibians; cool as soon as possible; keep cool and damp but not waterlogged until examination and possible isolation attempts. Batrachochytrium dendrobatidis can most easily be isolated from the mouthparts of tadpoles. Collect tadpoles with abnormal mouthparts, which can be detected in the field with a 10X hand lens. Isolation of B. dendrobatidis should only be attempted from skin samples from live or freshly dead frogs. Recognition of Chytridiomycosis

Before B. dendrobatidis can be isolated it must be microscopically identified in fresh host tissue. Keep amphibian tissue damp, but without free water. While observing with dissecting microscope at 200 X or 400 X, use a sharp, sterile needle to remove loose skin from between digits of foot and elsewhere on the ventral surface of the animal. If skin is not loose, use needle-nosed forceps and tear pieces from the leading edge of the skin between the hind digits or use a single edged razor blade to excise webbing from between toes. If larval animals have focal tooth loss or depigmentation of the jaw sheath, remove these areas with needle-nosed forceps. Place the skin or jaw sheath on a microscope slide in a drop of sterile distilled water and cover with a coverslip. Observe with a compound microscope at 100 X and 400 X. Look for walled, spherical to oval bodies (10-30 pm in diameter) inside of epidermal cells. Some of the bodies may be septate (delicate walls divide the fungal body, or thallus, into two or more sporangia; Figs. 39, 40). Some may have cleaved zoospores, which look like 5-20 spherical bodies within the sporangia. Other sporangia may have discharged their spores and appear empty. Isolation

If suspect organisms are found, place the piece of skin on a 9-cm culture plate containing mTGh nutrient agar and antibiotics, which are added after autoclaving (see recipe below). While observing through a dissecting microscope (40 X magnification with substage lighting), use a sharpened and sterilized needle to draw and push a small ( > 1 X 1 mm) piece of infected skin through nutrient agar in a 9-cm culture plate (If pieces of skin are thick and do not tear into small pieces, use micro-scissors or cuticle scissors to cut skin into small pieces while viewing with 20 X magnification). Every few millimetres take the needle away from the piece of slun and wipe through the agar. The purpose of wiping the skin and needle through agar is to remove bacteria, yeast and fungal spores. Bearing this in mind, reverse the direction of the skin, wipe it back and forth; imagine what you are trying to do even though you cannot see the bacteria. Jaw sheath tissue from recently killed larvae is usually quite free of other fungi and bacteria and requires less cleaning. When the skin is well wiped (at least back and forth across the diameter of the plate one time), use the cleaned needle to rexliove it from the cleaning plate, which has been open to the air, and put it on a fresh plate of mTGh agar. Barely open the fresh plate with one hand and wipe the skin piece into the agar with the needle held in the other hand. This is so that the pieces of slun will be on a plate that has not been exposed to fungal spores from the air. Repeat this process for as many pieces of skin as you have the patience; at a minimum, for each isolation attempt, put six pieces of wiped skin on each of two plates. Seal the nutrient agar plate with a 10 X 2 cm piece of Parafilm@ or other laboratory film stretched around the circumference of the plate. Label the dish with a permanent marker, recording source of skin and date; circle on the bottom of the plate areas where each of the pieces of skin are located. Incubate sealed plates on the laboratory bench or, if laboratory temperatures are above 25"C, in an incubator at 17" to 23°C. During the next one to three weeks, check development by inverting the culture plate on the stage of a compound microscope and observing the small pieces of skin with the low power objective. Also inspect plates for contaminants. If fungal contaminants are found, flame-sterilize a scalpel and remove. Motile B . dendrobatidis zoospores (3-4 pm diameter) may be evident around the cleaned, infected skin within 1-2 days, but sometimes not for several weeks. If chytrid colonies develop on the skin, they can be recognized as spherical bodies, some of which bear one or more nipple-like, discharge papillae. If hyphae are evident instead of round sporangia, use a sterile knife to aseptically remove the hyphal colony from the isolation plate. If B. dendrobatzdis colonies are produced on many pieces of skin, one piece can be removed to examine the fungus with the compound microscope. Open a plate in a way that minimizes the chances of entry of airborne fungal spores, and use a sterilized scalpel to remove a piece of the colony to a drop of water on a microscope slide. Observe with the compound microscope and compare morphology with published photographs of B. dendrobatidir. After the chytrid forms a colony on the isolation plate, aseptically transfer a piece of the colony to a plate of 15% tryptone agar. This fungus grows best in groups, so do not separate sporangia from each other during transfer.

Incubate for 1-2 weeks; if no bacteria develop and the fungus is growing well, spread the colony on the plate and transfer a part of the colony to nutrient broth in a screw-capped 250 ml flask. For back-up, also transfer bits of the colony to fresh plates and seal with Parafilm@. One should work within a laminar-flow hood if possible. For stocks, keep two sets of cultures in 1% liquid tryptone at 5°C. Stock cultures of other chytrids are kept on agar slants in screw-capped test tubes; however, B. dendrobatidis survives longer in a liquid medium. If stocks are kept on nutrient agar, refrigerate sealed Petri plates that contain scattered, small colonies of B. dendrobatidis. Refrigerated cultures in liquid medium have remained viable for more than four months. Maintaining Cultures of B. dendrobatidk

Place 75 ml of broth into 150 ml screw-capped flasks. The size of the flask or tube is not important; use larger or smaller containers depending on what is available. Because of the danger of contamination, use screw-capped vessels

3052

AMPHIBUN BIOLOGY

and keep duplicates. Incubate at 23°C or below. For long-term (at least three months) storage place at 5" C after growth is evident. When growing to produce zoospores for inoculum or other purposes, grow on 1% tryptone agar medium. Cryoarchiving To store isolates without the need to passage every three months, which is tedious and may result in altered genotypes or contamination, a simple method to freeze cultures was developed by Boyle et al. (2004). Production of Zoospores Grow B. dendrabatidis in 1% liquid q-ptone until clumps of thalli are visible to the eye. Use a sterilized Pasteur pipette to add 112 to 314 ml of this broth culture to 1 9 tryptone agar in 9-cm culture dishes. Leave inoculated dishes open in a laminar-flow hood until the added broth is almost dry; replace covers on dishes and place in plastic sleeve. Incubate in 15" C incubator. Plates can be incubated at higher temperatures, up to 23"C, but the potential harvest period is longer if plates are kept in plastic sleeves at 15°C. After 4-10 days, look for active zoospores around the periphery of fungal colonies by inverting dishes on the stage of a compound microscope and examining with the 10X objective. Harvest zoospores by flooding plates with about 2-3 ml of sterile distilled water. Decant after about 30 minutes to collect zoospores. Zoospore concentration can be measured by optical density or by counting with a hemocytometer. Zoospores may stay motile (thus infective) for up to 24 hours; however, most encyst before 24 hrs. Biosafety Although thorough drying can kill B. dendrabatidis, take all precautions. Autoclave all materials that contain or have come into contact with the fungus before disposal. If B. dendrabatidis is used to inoculate amphibians, kill and incinerate or fix all exposed animals after the experiment. Be sure that cages, water, and other material in cages are disinfected at the end of experiments. Do not dump potentially contaminated material in the trash or down the drain without first treating it (heat or bleach) to kill the fungus. Recipes 8 g tryptone 2 g gelatin hydrolysate 10 g agar 1,000 ml distilled water Add 200 mg/L penicillin-G and 200-500 mg/L streptomycin sulfate after autoclaving; if bacteria are still a problem add 1 mg/L ciprofloxacillin.

1%Tryptone agar

10 g tryptone 10 g agar 1000 ml distilled water 1%Tryptone broth

10 g tryptone 1000 ml of distilled water Naming Isolates A standard naming scheme was proposed using Location-Species-Year-Collector's Initials-Isolate Number (Berger It is important to record passage number, as isolates may evolve in culture.

et a1 2005b), for example Rockhampton-Lcaerula-99-L8-1.

CHAPTER 3

Factors affecting interspecific variation in susceptibility to disease in amphibians Jodi J. L. Rowley and Ross A. Alford.

I. Introduction II. Amphibian Behaviour A. Microenvironment selection 1. Normal Thermoregulation 2 . Behavioural Fever B. Opportunities for Transmission 1. Physical Contact 2. Contact with Water and other Environmental Reservoirs 3. Movement Patterns

Ill. Immune Response A. Antimicrobial Peptides B. Microbiota C. Immunity and Environmental Conditions IV. Morphology/Physiology V. Conclusion VI. Acknowledgements

I. INTRODUCTION

I

N recent decades, infectious diseases affecting populations of wildlife and plants have been emerging at unusually high rates; emerging disease currently poses a great threat to the conservation of global biodiversity (Harvell et al. 1999; Ward and Lafferty 2004). Amphibians that have declined or been eliminated due to disease are taxonomically diverse and occur in a variety of systems (Berger et al. 1998; Bosch et al. 2001; Bradley et al. 2002; Weldon and Du Preez 2004; Lips et al. 2006; Rachowicz et al. 2006). In addition to causing declines, pathogens may prevent the recovery of a species by persisting within the host population or within environmental reservoirs or alternate hosts (Lafferty and Gerber 2002; Retallick 2002; DeCastro and Bolker 2005). Furthermore, if the disease itself does not eliminate the host species, the reduction in abundance of the host after a disease outbreak will increase susceptibility to extinction by other factors, such as random stochastic events (Lafferty and Gerber 2002).

In addition to causing the dramatic increases in host morbidity and mortality typical of disease outbreaks, pathogens have great impacts on the structure of hosts' populations by altering population characteristics such as age structure (Van Rensburg et al. 1987), and by affecting the outcome of interspecific interactions such as competition (Hudson and Greenman 1998; Tompkins et al. 2003; Parris and Cornelius 2004) and predation (Mrebber et al. 1987). Pathogens can also lower host reproductive output (Saumier et al. 1986) and alter the movement patterns of hosts (Steck and Wandeler 1980; Anderson and Trewhella 1985). Pathogens can shape the species richness and structure of entire communities,

3054

AMPHIBIAN BIOLOGY

particularly when the pathogen is not host specific. By causing mortality in a range of species, pathogens may convert species-rich communities into depauperate ones dominated by a few resistant species (McDonald and Alford 1999; Lips et al. 2006). Changes directly produced by pathogens can have strong indirect effects that alter the structure of entire food webs. The dramatic declines in amphibian populations that have occurred at some sites are likely to cause long-term reductions in the abundance of predators that prey on amphibians, and alterations to primary production and the structure of algal assemblages (Whiles et al. 2006). Understanding, and ultimately managing, the impacts of infectious disease on amphibians requires understanding the factors determining the susceptibility of individuals to disease, and the susceptibility of populations to epidemic outbreaks. Individual susceptibility to disease is often considered to be largely a function of the immune system (Morales and Robert 2007; Woodhams et al. 2007a), and the susceptibility of populations to epidemic outbreaks is thought to be related to individual susceptibility and to factors affecting transmission rates (DeCastro and Bolker 2005). It has become clear that the physiology, ecology, and behaviour of amphibians cause the factors affecting susceptibility at both levels to be complex (Rowley and Alford 2007a, 2007b; Woodhams et al. 2007b), and that understanding these factors requires integration of a detailed knowledge of amphibian physiology, behaviour, and ecology with information on the biology of pathogens. The disease chytridiomycosis, caused by the fungus Batrachochytrium dendrobatidis, provides a clear example of the complex nature of the factors controlling susceptibility to disease at the individual and population level in amphibians. Chytridiomycosis can cause rapid mortality (Nichols et al., 2001), with frogs of susceptible species dying less than three weeks after experimental infection in the laboratory (Berger et al., 1998, Berger et al., 2004, Berger et al. 2005a). In almost all cases, however, amphibian species that have disappeared or declined due to chytridiomycosis coexist with non-declining species (Lips and Donnelly 2002; Retallick et al. 2004; Puschendorf et al. 2006). Susceptibility at the level of the individual may not be strongly affected by the adaptive immune system (Woodhams et al. 2007b), but can be strongly influenced by the innate immune system, including antimicrobial peptides contained in secretions from the granular glands (Woodhams et al. 2006a,b, 2007a), by microhabitat use and basking behaviour and their effects on body temperature (Woodhams et al. 2003, 2007b), and by interactions between the amphibian chytrid and the assemblage of other microbes that inhabit amphibian skin (Harris et al. 2006; Woodhams et al. 2007b). Susceptibility at the population level can also be strongly affected by behaviour and by choice of microenvironments, and can vary seasonally in response to changes in temperature and availability of moisture and their associated changes in amphibian behaviour (Berger et al. 2004; Woodhams and Alford 2005; Rowley and Alford 2007a). 11. AMPHIBIAN BEHAVIOUR

Amphibian species that persist in the wild when chytridiomycosis is endemic can be highly susceptible to Batrachochytrium dendrobatidis in the laboratory, experiencing 100% mortality there (Woodhams et al. 2003; Berger et al. 2005a). This suggests that laboratory environments somehow promote the development of disease in infected individuals. One of the obvious differences between most laboratory environments and those in the field is the simplicity of the laboratory; amphibians are typically maintained in relatively simple enclosures at constant temperatures. Differences in the susceptibility of amphibians between the laboratory and the field could be at least partially caused by aspects of the hosts' behaviour and microenvironmental use that occur in the field, but cannot occur in a simplified laboratory setting. Microenvironmental selection, frequency of contact with other amphibians and environmental reservoirs of B. dendrobatzdis, as well as the hosts' movement patterns may be particularly important.

ROWLEY and ALFORD: FACTORS AFFECTING LARIATION IN DISEASE

3055

A. Microenvironment Selection In many cases, a pathogen will only cause disease in a host if environmental conditions are conducive to do so. This phenomenon causes seasonally or climatically fluctuating prevalence of disease in a population (e.g. Dowel1 2001, Bruno et al. 1995, Enger et al. 1991, Cowling and Gilchrist 1982), and leads to the commonly observed disparity in host mortality between the laboratory and field-infection trials (Inglis et al. 1996). In general, the most important environmental factors affecting pathogen survival are temperature, moisture and solar radiation (Benz 1987; Carruthers et al. 1992) although pH, the presence of organic matter and exposure to ~a-iouschemicals can also be important (Edwards 2000). Environmental factors do not act independently and the effect of any one environmental factor is typically dependent on other environmental factors (Holmes and Colhoun 1974; Eyal et al. 1977; Monroe et al. 1997). In addition, environmental factors may act on different stages of the host-pathogen interaction independently, making outcomes difficult to predict. Batrachochytrium dendrobatidzs is highly sensitive to environmental conditions. In culture, the growth and survival of B. dendrobatidts are highb- dependent on temperature (Longcore et al. 1999, Piotrowski et al. 2004). Temperature also influences the progress and outcome of chytridiomycosis in infected amphibians in the laborator?; with low andtor fluctuating temperatures capable of retarding development of the disease, and elevated body temperatures clearing infection (Woodhams et al. 2003, Berger et al. 2004). In the field, prevalence of infection and mortality of hosts are also correlated with ambient temperature, and as a result are highest during cooler months (Berger et al. 2004; Retallick et al. 2004; McDonald et al. 2005; Woodhams and Alford 2005) and at higher elevations (McDonald et al. 2005; Woodhams and Alford 2005).

Hydric conditions are also important for B. dendrobatzdis, with sporangia and zoospores killed by desiccation (Berger 2001; Johnson et al. 2003). In addition, progression of the disease in the laboratory is more rapid in mist than in either constant rain or under very dry conditions (Alford and Woodhams, unpublished data). The effects of hydric conditions on disease dynamics in the field are less clear, at least partly due to the difficulv in isolating the effects of rainfall or humidity from those of temperature. 1. Normal Themoregulation In any discussion of host-pathogen interactions and environmental conditions, it is important to remember that it is microenvironmental conditions, and not ambient conditions, that are of most relevance. For example, while the body temperature of amphibians is tied to ambient temperatures, many amphibians sometimes attain body temperatures only loosely related to environmental ones. The disparity between environmental temperature and body temperature is due to the hosts' thermal behaviour. Consequently, thermal behaviour of the host has a large influence on the outcomes of hostpathogen interactions and may even account for interspecific differences in susceptibility to disease. Thermoregulation in ectotherms can be defined as the active process involving behavioural adjustments that maintain body temperatures as close as possible to an optimal temperature range for physiological functions (Hertz et al. 1993; Thomas and Blanford 2003). In ectotherms, thermoregulation is achieved by using behavioural adjustments (such as shuttling between sun and shade) which, at least under optimal conditions, allow individuals to maintain a relatively constant, optimal body temperature, regardless of environmental temperatures (Hertz et al. 1993). Body temperature and behaviour will be affected by species-specific thermal profiles, the distribution and colour of the animal and its size (age) and sex (Thomas and Blanford 2003). The majority of research investigating the interaction between thermoregulation and disease in ectotherms has focused on insects, particularly locusts and grasshoppers. Many

3056

AMPHIBIAN BIOLOGY

insects are good thermoregulators and are able to raise body temperatures up to 15OC above air temperature, reaching body temperatures of over 40°C via habitat selection and basking alone (Carruthers et al. 1992; Thomas and Blanford 2003). Amphibians are also capable of achieving body temperatures that differ greatly from ambient. For example, basking R a m mucosa may achieve an average of 14.4OC higher body temperature than presumed for adjacent, non-basking frogs (Bradford 1984). Due to thermoregulation, the body temperatures of many amphibians may exceed the upper limits for survival of hngal pathogens, even when air temperature does not (e.g., Carruthers et al. 1992). As optimal temperatures for the growth of B. dendrobatzdis in vitro is between 17°C and 25OC (Piotrowski et al. 2004), with upper limits of 29"-30°C (Longcore et al. 1999; Piotrowski et al. 2004), it is quite possible that behavioural thermoregulation may provide a mechanism whereby animals can avoid or eliminate pathogenic infection. This may be entirely incidental to normal thermoregulatory patterns; a species that chooses to spend sufficient time with body temperatures above the optimal growth range for B. dendrobatidis, for example, will have greatly reduced susceptibility to chytridiomycosis, even if individuals do not alter their behaviour in response to infection. 2. Behavioural Fever

Fever is widespread in both endothermic and ectothermic animals, including mammals, birds, reptiles, fish and arthropods (Kluger 1978; Hart 1990). Ectothermic vertebrates, including some neonate mammals, are unable to produce a fever by physiological means, as they cannot physiologically increase their body temperatures much above ambient. Despite this, many ectotherms, including amphibians, exhibit a fever response following infection, whereby thermoregulatory behaviour, rather than physiology, is modified to enable hosts to attain a new, higher preferred body temperature (Thomas and Blanford 2003). Behavioural fever can therefore be described as an elevation, using behavioural means, of preferred or selected body temperatures above normothermic levels (Hutchinson and Erskine 1981). Although behavioural fever is apparently not ubiquitous among ectotherms (Karban 1998), fevers have been reported in a wide range of ectotherms including teleost fish, amphibians, and reptiles (eg. Vaughn et al. 1974; Reynolds et al. 1976; Casterlin and Reynolds 1977a,b, 1979; Myhre et al. 1977; Bronstein and Conner 1984; Watson et al. 1993; Adamo 1998; Blanford et al. 1998; Sherman 2008). Behavioural fever has been shown to increase host survival in a number of host-pathogen associations, including those with reptilian (Kluger et al. 1975), fish (Covert and Reynolds 1977) and insect hosts (Bronstein and Conner 1984; Inglis et al. 1996; Adamo 1998; Karban 1998). Under field conditions, the ability of behavioural fever to suppress disease may be limited. In the field, ectotherms will only be able to elevate their body temperature diurnally, and even then, generally only during sunny conditions. For some diseases such as chytridiomycosis, even short-term elevation of body temperature may eliminate the infection, as demonstrated by Woodhams at al. (2003), who showed that two days with body temperatures of 37°C completely cured infected Litoria chloris. However, in other systems, behavioural fever may cause reduced pathogenicity and prolonged incubation of the disease, but may not allow recovery of the host or complete suppression of the disease (Blanford et al. 1998; Elliot et al. 2002), as has commonly been demonstrated in the laboratory (Blanford and Thomas 1999). The mechanisms underlying how behavioural fever contributes to a host overcoming infection are unknown. Behavioural fever may act by directly inhibiting growth and development of the pathogen, by increasing the effectiveness of the host's immune system, or by a combination of these two mechanisms (Covert and Reynolds 1977; Kluger and Rothenburg 197'9; Adamo 1998; Ouedraogo et al. 2003).

ROWLEY and ALFORD: FACTORS AFFECTING VARIATION IN DISEASE

3057

B. Opportunities for Transmission

The transfer of pathogens from the environment to the host, or from host to host, is an important factor in the development of epizootics, particularly when host density or contact rates between hosts are low. The long-term spread and persistence of many diseases depends largely on the contact rate between susceptible hosts and infectious pathogens in a population (Swinton 1998). Therefore, differences among amphibian species in the number of opportunities for the transmission of B. dendrobatzdis may influence their susceptibility to chytridiomycosis. The only known mechanism of B. dendrobatidis transmission among hosts, and of intrahost increase in pathogen load, is via infection and reinfection by motile, waterborne zoospores (Longcore et al. 1999). To date, transmission of B. dendrobatidis has been experimentally demonstrated to occur via contact with water that was previously in contact with either infected tadpoles or adult amphibians (Berger et al. 1998; Retallick 2002; Parris and Cornelius 2004; Rachowicz and Vredenburg 2004). However, infective zoospores occur on the skin surfaces of infected animals (Berger et al. 1998; Pessier et al. 1999). B. dendrobatidis' DNA has also been detected on wet rocks at the site of an epidemic (Lips et al. 2006). Transmission of B. dendrobatzdis is therefore also likely to occur via contact with infected individuals or contaminated environmental reservoirs. Transmission rates in the field, and therefore potential susceptibility to chyuidiomycosis, are likely to vary interspecifically in amphibians depending on the degree of physical contact among frogs, or contact between frogs and infected water or other environmental resenloirs. Patterns of movement of the host may also affect the transmission of B. dendrobatidir. 1 . Physical Contact

The contact rate between infected and susceptible hosts varies widely according to characteristics of the host population (Fromont et al. 1998). The concept of a "threshold population size" or density below which infection cannot persist is common in most models of host-pathogen interaction (Anderson et al. 1981; Swinton et al. 2001). Theory predicts that, beyond this threshold, and with typical density-dependent transmission, the prevalence of infection should increase with increasing density (Begon et al. 2003). This is because high host densities increase the contact between infected and uninfected hosts and between host and pathogen (Watanabe 1987). As more hosts become infected, it follows that more pathogens are produced and epizootics are more likely to take place (Watanabe 1987). At other times, however, contact rates may be almost entirely independent of population size simply because of hosts' behaviour (Loehle 1995; McCallum et al. 2001; Ezenwa 2004). The formation of aggregations by individual hosts promotes contact between individuals, even at low population densities, and in many host-pathogen systems formation of aggregations is positively correlated with both the prevalence and intensity of contacttransmitted parasites (Anderson and May 1979; Hoogland 19'19; Brown and Brown 1986; Cot6 and Poulin 1995; Ezenwa 2004). Outbreaks of disease are most commonly observed in aggregations of individuals (Vermeer 1969; Wobester et al. 1979). Amphibian species differ in their frequency and duration of physical contact. For example, in three species of sympatric rainforest stream frogs in northern Australia, the frequency of frog-to-frog contact differed significantly among species and was correlated with the conservation status of the frogs; the highest rates of contact occurred in Litoria nunnotis, the species most susceptible to chytridiomycosis-related declines, and the lowest rates occurred in L. lesueuri, the species least susceptible to chytridiomycosis-related declines (Rowley and Alford 2007a). Disease transmission can also result from physical contact during territorial defense (Loehle 1995) or in breeding aggregations, and amphibian species that frequently engage in male-male combat, or form large aggregations at breeding sites may be more likely to become infected with B. dendrobatidis during the breeding season. Such behaviour has been

3058

AMPHIBIAN BIOLOGY

implicated in the lower survival of Bufo boreas males compared to females during B. dendrobatidis outbreaks, as adult males spend several weeks in frequent physical contact with other males (and with water) during the breeding season, compared to females which spend less than one day at breeding sites (Carey et al. 2006). 2. Contact with Water and other Environmental Reservoirs In the field, pathogen survival is influenced by a complex array of factors and varies spatially according to habitat features, topography and microenvironmental conditions (Dinnik and Dinnik 1961; Levine 1963; Carruthers and Haynes; 1986; Hochberg 1989; Roland and Kaupp 1995; Blanford and Thomas 2000; Arthurs and Thomas 2001). Habitats with conditions optimal for the persistence of pathogens may form "reservoirs", whereby pathogens can be maintained for long periods and transmitted to the host population. Species-specific patterns of habitat use will therefore influence transmission rates of pathogens among species (Holt et al. 2003) and the prevalence of disease or the infection rate may vary predictably with type of habitat or even of microhabitat (Carruthers and Haynes 1986). Opportunities for transmission are likely to be higher in host species that spend the greatest amount of time in habitats that are conducive to the survival of a pathogen (Burdon et al. 1989). As a result, the abundance and species composition of parasites affecting host species differs between habitats, even those in close proximity (Grutter 1998; Krasnov et al. 1998), and with frequency of contact with environmental reservoirs (Brooks et al. 2006). For example, the species richness of platyhelminth parasites in amphibians is highly correlated with the amount of time the host spends in association with aquatic habitats (Brooks et al. 2006). For amphibians, the likelihood of B. dendrobatidis transmission is likely to vary interspecifically with the frequency of contact with water or other, as yet unquantified, environmental reservoirs. Indeed, amphibian species with strong associations with streams have suffered the most dramatic population declines, and even when in sympatry with rapidly declining species, terrestrial species do not typically experience population declines, or decline to a lesser degree (Williams and Hero 1998; McDonald and Alford 1999; Lips et al. 2003; Hero et al. 2005; Lips et al., 2006). The frequency of contact with the stream was correlated with conservation status in three sympatric stream-breeding frogs in northern Queensland, with contact with stream water being more frequent in the species that had declined most due to chytridiomycosis, and least frequent in the species that did not noticeably decline (Rowley and Alford 2003a). Such correlations are not surprising, given the aquatic nature of B. dendrobatidis' zoospores, their sensitivity to desiccation (Johnson et al. 2003), and their ability to remain infective in the laboratory for at least seven weeks in sterile lake water (Johnson and Speare 2003). In addition, B. dendrobatidis has been experimentally transmitted among frogs via water in the laboratory (Berger et al. 1998; Parris and Cornelius 2004; Rachowicz and Vredenburg 2004) and in field enclosures (Retallick 2002). Tadpoles also often have high prevalences of infection, may not be susceptible to the pathogen, and can remain in the stream environment for several years (Woodhams et al. 2006; Rachowicz et al. 2006), potentially providing a continual source of zoospores to the stream. The risk of transmission is likely to vary depending on the nature of a body of water, \slth some areas being more conducive to transmission of pathogens. For example, there is generally a negative correlation between stream flow and fish parasite populations in both k h w a t e r and marine environments (Stables and Chappell 1986; Janovy et al. 1997; Swearer and Robertson 1999; Barker and Cone 2000). Amphibians inhabiting areas of high stream flm- may therefore experience reduced risk of coming into contact with B. dendrobatidis, although to date there is no experimental evidence for this. Son-aquatic reservoirs for B. dendrobatidis are poorly known. Because the abundance and composition of other chytrids differ among microhabitats (Letcher and Powell 2002),

ROWLEY and ALFORD: FACTORS AFFECTING VARIATION IN DISEASE

3059

it is highly probable that the abundance of B. dendrobatzdis zoospores also differs among environmental substrates. The DNA of B. dendrobatidis has been detected in samples of environmental substrate during epidemics (Lips et al. 2006); however, it is not known whether these samples contained viable zoospores. B. dendrobatidis DNA was not detected from 81 environmental substrate samples taken at L. lesueuri and L. nannotis retreat sites in an area where B. dendrobatidis had been endemic for at least ten years (Rowley et al. 2007). Duration of contact with water or environmental substrates is also likely to be important. In the laboratory, the duration of exposure to B. dendrobatzdis can influence the probability of successhl transmission and the speed of progression of the disease, with longer exposures to B. dendrobatzdis resulting in shorter average survival times in Bufo boreas (Carey et al. 2006). Species that return repeatedly to the same retreats, especially communal ones, are also likely to have increased opportunities for transmission, as parasites and pathogens may accumulate at these sites (Altizer et al. 2000). 3. Movement Patterns

Species-specific patterns of movement may affect disease dynamics. Less mobile hosts tend to have higher burdens of contagious parasites as pathogens and parasites often accumulate in a hosts' environment over time (Altizer et al. 2000; Ezenwa 2004). Hosts' movements and migration patterns may also serve to disperse, or contain, the spread of pathogens (Watanabe 1987). Movement patterns may be important in determining susceptibility to decline from B. dendrobatidis; mobility is negatively correlated with susceptibility to decline in three rainforest stream frogs from northeastern Queensland, Australia (Rowley and Alford 2007b). 111. IMMUNE RESPONSE

Amphibians have a well-developed immune system including both adaptive and innate components (Carey et al., 1999). However, they appear to have a poor adaptive immune response towards B. dendrobatidis, displaying a low degree of lymphocytic infiltration in the skin, even when severely infected with this fungus (Berger et al. 1998, 2005; Pessier et al. 1999), and there is little evidence for a humoral defense in amphibians infected with B. dendrobatidis (Woodhams et al. 2006; Woodhams et al. 2007b). The most important defences against infection by B. dendrobatdzdis in amphibians are likely to involve antimicrobial peptides or symbiotic microbiota associated with the skin surface. Together, they may provide an immediate host response to infection by B. dendrobatidis, well before any adaptive immune response is activated. A. Antimicrobial Peptides

Antimicrobial peptides are small (10-50 amino acid residues), cationic, hydrophobic molecules produced in the dermal granular skin glands of amphibians (Andreu and Rivas 1998; Simmaco et al. 1998; Conlon 2004). Amphibian species vary significantly in the composition and antimicrobial activity of their associated antimicrobial peptides (Conlon 2004). Isolated amphibian antimicrobial peptides and natural mixtures of skin peptides are capable of acting against B. dendrobatzdis in &TO (Rollins-Smith and Conlon 2005; Woodhams et al. 2006; Sheafor et al. 2008). It has been suggested that antimicrobial peptides may protect amphibians from the initial infection by B. dendrobatidis zoospores or slow the progression of the disease (Rollins-Smith et al. 2002a,b; Rollins-Smith and Conlon 2005). Antimicrobial peptides in amphibian skin are highly diverse (Erspamer 1994) and although families of peptides can be shared by related species, individual peptides are almost always unique to a single species (Conlon 2004). As a result, interspecific differences in the composition and effectiveness of antimicrobial peptides against B. dendrobatzdis may at least partially explain why some species are more susceptible to chytridiomycosis than others.

3060

AMPHIBIAN BIOLOGY

Indeed, the effectiveness of amphibian antimicrobial peptides against B. dendrobatzdis in vitro is negatively correlated with the hosts' susceptibility to decline from chytridiomycosis in some (Woodhams et al. 2005; Woodhams et al. 2006), but not all (Rollins-Smith et al. 2006; Woodhams et al. 2006) amphibian assemblages.

B. Microbiota A diverse array of bacteria inhabit the skin of amphibians. It has been known for some time that the cutaneous bacteria of some brooding salamanders can inhibit pathogenic fungus capable of infecting embryos (Austin 2000). More recently, it has been discovered that bacteria from the skins of several species of salamanders inhibit the growth of B. dendrobatzdis in vitro (Harris et al., 2006; Lauer et al. 2007). Some of these bacteria produce known compounds with antimicrobial properties (Brucker et al. 2008). Correlations (Woodhams et al. 2007b; 2007c) indicate that antimicrobial bacteria occur more frequently on Rana muscosa in populations that are coexisting with B. dendrobatzdis than they do in populations that have declined due to chytridiomycosis. Very recent evidence (Harris et al. 2008) indicates that exposing Rana muscosa to bacteria with anti-chytrid activity can significantly reduce subsequent frog mortality due to chytridiomycosis. Harris et al. (2008) also demonstrated that compounds with known antifungal properties that are synthesized by bacteria can be present on frogs' skin at concentrations sufficient to completely inhibit the growth of cultures of B. dendrobatidis in vitro. This area of research is in its infancy, but the results obtained to date suggest strongly that interspecific differences in the bacterial fauna associated with amphibian skin could play an important role in determining susceptibility to chytridiomycosis. Because bacteria can be inhibited by amphibian antimicrobial peptides (Woodhams et al. 2007b), it is likely that the cumulative effects of innate immune defenses on the microbial community of pathogens such as B. dendrobatidis are very complex and context-dependent. C. Immunity and Environmental Conditions Changes in environmental conditions may affect host immunity, and therefore the susceptiblity of amphibians to chytridiornycosis. The most powerful factor affecting hosts' immune responses is temperature. The immune systems of amphibians are temperature dependent, with low temperatures reducing, or eliminating, immune system activity, including the production of antimicrobial in some species (Carey et al. 1996; Carey 2000; Matutte et al. 2000). Temperature changes need not be large in order to alter immune responses, with relatively small differences in temperature eliciting huge differences in immune response. The time taken to reject a skin graft in adult and larval bullfrogs (Rana catesbeiana) at 15OC is three times as long as at 25OC (Hildemann and Haas 1959) and an increase in only 3OC can decrease the time taken to reject a graft (Cohen 1966). Aside from low temperatures, environmental factors such as pesticides, environmental endocrine disruptors, and UV-B radiation may suppress aspects of amphibian immune systems (Taylor et al. 1999; Carey 2000; Rollins-Smith et al. 2002b,). For example, exposure to the pesticide carbaryl significantly reduces the cutaneous peptide defences of Rana boylii (Davidson et al. 2007).

Interspecific differences in the morphology and physiology of amphibians may also be important in determining susceptibility to chytridiornycosis, but so far have not been explored. Important determinants of susceptibility may be the abundance and distribution of granular glands that secrete antimicrobial peptides, or the frequency of skin shedding, which may influence the ability of B. dendrobatidis' zoospores to reinvade a host (Woodhams et al. 2007a). Voyles et al. (200'7) demonstrated that Litoria caerulea in the late stages of chytridiomycosis experienced large decreases in the concentrations of plasma electrolytes, which may have been the proximate cause of mortality. These appeared to be caused by

ROWLEY and ALFORD: FACTORS AFFECTING VARIATION I N DISEASE

3061

loss of electrolytes rather than by increased water uptake. This suggests that differences in the structure and function of regions of the skin that are involved in osmoregulation may also be important, as well as differences in tolerance to osmotic imbalance. V. CONCLUSION

It is clear that the interactions of amphibians with pathogenic organisms are extremely complex. Because the body temperatures of amphibians fluctuate, the rates of physiological and biochemical processes within them also fluctuate, potentially affecting the rate at which their defenses can respond to infection. Many amphibians select microenvironments based on their requirements for temperature and moisture. This means that the environments in which host-pathogen interactions involving amphibians occur cannot be characterized using simple measurements of macroenvironmental variables; it is necessary to understand their physiology and behaviour in detail. In addition to complex adaptive immune systems, amphibians possess innate immune defenses in the form of antimicrobial peptides and symbiotic microbiota; all of these may play a role in determining their susceptibility to any pathogen. Although laboratory studies are necessary in determining how amphibians may interact with pathogens, it is important to realize that the results may not generalize to field conditions. For example, the Australian green tree frog Litoria caeruEea is highly vulnerable to chytridiomycosis in the laboratory (Berger et al. 2005a), but has never experienced effects at the population level in the field, although they extensively co-occur with B. dendmbahdk. This is probably caused at least in part by the oversimplification of environmental conditions. Laboratory experiments often maintain environmental conditions at constant levels (high humidity, temperature between 17" and 23C) that are optimal for the growth and development of B. dendrobatdis, and may be optimal or suboptimal for other amphibian pathogens. Because laboratory experiments are usually conducted on animals that are either captive-bred or have been maintained for extensive periods in captivity, it is likely that the results may be affected by changes in the assemblage of microbes inhabiting the host. The complex and poorly understood nature of the factors that determine the susceptibility of amphibians to disease, and the fact that emerging diseases appear to be a major threat to amphibian diversity, indicate that there is an urgent need to gain a much greater understanding of how these processes operate in a wide variety of taxa and environments, so that the present threat of chytridiomycosis and hture threats that may emerge can be managed. VI. ACKNOWLEDGEMENTS

The ideas presented in this chapter could not have been developed without the support of a series of grants, in particular subcontracts to RAA from US National Science Foundation IRCEB grants (IBN-99063 and DEB-02138, J. I? Collins, I? I.) and a research contract (RFT 4312004) to RAA f b m the Australian Department of the Environment and Heritage. The chapter also benefited greatly from discussions of the authors with many students and collaborators. VII. REFERENCES Adamo, S. A., 1998. The specificity of behavioural fever in the cricket Acheta domesticus. J. Parasitology 84: 529-533.

Anderson, R. M. and Trewhella, W., 1985. Population dynamics of the badger (Meles meles) and the epidemiology of bovine tuberculosis (Mycobacterium bowis). Philos. Trans. Roy. Sac. Lond. Series B, Biol. Sci. 310: 327-381.

Altizer, S. M., Oberhauser, K. S. and Brower, L. P 2000. Associations between host migration and the prevalence of a protozoan parasite in natural populations of adult monarch butterflies. Ecol. Entomology 25: 125-139.

Anderson, R. M., Jackson, H. C., May, R. M. and Smith, A. M., 1981. Population dynamics of fox rabies in Europe. Nut. 289: 765-777.

Anderson, R. M. and May, R. M., 1979. Population biology of infectious diseases: Part I. Nat. 280: 361-367.

Andreu, D. and Rivas, L., 1998 Animal antimicrobial peptides: an overview. Biopolymers (Peptde Science) 47: 415433.

3062

AMPHIBIAN BIOLOGY

Arthurs, S. and Thomas, M. B., 2001. Effect of dose, pre-mortem host incubation temperature and thermal behaviour on host mortality, mycosis and sporulation of Metarhitium anisopliae var. acridum in Schistocm regaria. Biocatml Sei. Technology 11: 41 1-420.

Bosch, I., Martinez-Solano, I. and Garcia-Paris, M., 2001. Evidence of a chytrid fungus infection involved in the decline of the common midwife toad (Alytes obstetricans) in protected areas of central Spain. Biolog. Cons. 97: 331-337.

Austin, R. M., 2000. Cutaneous microbial flora and antibiosis in Plethodon ventralis. Pp. 127-136 in 'The Biology of Plethodontid Salamanders", ed by R. C. Bruce, R. G. Jaeger and L. D. Houck. Kluwer AcademidPlenum, New York.

Bradford, D. F., 1984. Temperature modulation in a high-elevation amphibian, Rana mucosa. Copeia 1984: 966-976.

Barker, D. E. and Cone, D. K., 2000. Occurrence of Ergmilus celestis (Copepoda) and Pseudoductylogryrw anguillae (Monogena) among wild eels (Anguilla rostrata) in relation to stream flow, pH and temperature and recommendations for controlling their transmission among captive eels. Aquaculture 187: 261-274. Begon, M., Hazel, S.-M., Telfer, S., Bown, K., Carslake, D., Cavanagh, R., Chantrey, J., Jones, T and Bennett, M., 2003. Rodents, cowpox virus and islands: densities, numbers and thresholds. J. Anim. Ecol. 72: 343-355. Benz, G. 1987. Environment. 177-214 in "Epizootiology of Insect Diseasesp:kl by J R. Fuxa and Y Tanada. John Wiley and Sons Inc, New York. Berger, L., 2001. Diseases in Awtralian Frogs. PhD, James Cook University, Townsville. Berger, L., Marantelli, G., Skerratt, L. F. and Speare, R., 2005a. Virulence of the amphibian chytrid fungus Batrachochytrium dendrobatzdis varies with the strain. Diseases ofAquatic Organisms 68: 47-50. Berger, L., Speare, R. and Skerratt, L. F., 2005b. Distribution of Batrachochytriurn dendrobatidis and pathology in the skin of green tree frogs Litaria caerulea with severe chytridiomycosis. Diseases of Aquatic Organisms 68: 65-70. Berger, L., Speare, R., Daszak, F!, Green, D. E., Cunningham, A. A., Goggin, C. L., Slocombe, R., Ragan, M. A., Hyatt, A. D., McDonald, K. R., Hines, H. B., Lips, K. R., Marantelli, G. and Parkes, H., 1998. Chytridiomycosis causes amphibian mortality associated with population declines in the rainforests of Australia and Central America. Proc. Nut. Acad. Sci. USA 95: 9031-9036. Berger, L., Speare, R., Hines, H. B., Marantelli, G., Hyatt, A. D., McDonald, K. R., Skerratt, L. F., Olsen, V, Clarke, J. M., Gillespie, G., Mahony, M., Sheppard, N., Williams, C. and Tyler, M. J., 2004. Effect of season and temperature on mortality in amphibians due to chytridiomycosis. Aust. Eterinary J. 82: 434-439. Blanford, S. and Thomas, M. B., 1999. Host thermal biology: the key to understanding host-pathogen interactions and microbial pest control? Apicul. For: Entomology 1: 195-202. Blanford, S. and Thomas, M. B., 2000. Thermal behavior of two acridid species: Effects of habitat and season on body temperature and the potential impact on biocontrol with pathogens. Environmental Entomology 29: 1060-1069. Blanford, S., Thomas, M. B. and Langewald, J. 1998. Behavioural fever in the Sengalese grasshopper, Oedaleus senegalensis, and its implications for biological control using pathogens. Ecol. Entomology 23: 9-14.

Bradley, G. A., Rosen, F! C., Sredl, M. J., Jones, T R. and Longcore, J. E., 2002. Chytridiomycosis in native Arizona frogs. J. Wildl. Diseases 38: 206-212. Bronstein, S. M. and Conner, W. E., 1984. Endotoxininduced behavioural fever in the Madagascar cockroach, Gromphadorhina portentosa. J. Insect Physiol. 30: 327-330. Brooks, D. R., Le6n-Regagnon, V, McLennan, D. A. and Zelmer, D., 2006. Ecological fitting as a determinant of the community structure of platyhelminth parasites of anurans. Ecol. 87: S76S85. Brown, C. R. and Brown, M. B., 1986. Ectoparasitism as a Cost of Coloniality in Cliff Swallows (Hirundo pyrrhonota) Ecol. 67: 1206-12 18. Brucker, R. M., Baylor, C. M.,Walters, R. L., Lauer, A, Harris, R. N., and Minbiole, K. P. C. 2008. The identification of 2,4-diacetylphloroglucinol as an antifungal metabolite produced by cutaneous bacteria of the salamander Plethodon cinereus. J. Chemical Ecol. 34: 39-43. Bruno, D. W., Collins, R. 0. and Morrison, C. M,. 1995. The occurrence of Lama salmonae (Protozoa: Microspora) in farmed rainbow trout, Oncorhynchus mykiss Walbaum, in Scotland. Aquaculture 133: 341-344. Burdon, J. J., Jarosz, A. M. and Kirby, G. C., 1989. Pattern and patchiness in plant-pathogen interactions - Causes and Consequences. Ann. Rev. Ecol. Syst. 20: 119-136. Carey, C., 2000. Infectious disease and worldwide declines of amphibian populations, with comments on emerging diseases in coral reef organisms and humans. Environment Health Perspective 108: 143-150. Carev. ,, C.., Cohen. N. W. and Rollins-Smith. L. A.. 1999. Amphibian declines: an immunological perspective. Developmental Campal: Immunology 23: 459-472. Carey, C., Maniero, G. D. and Stinn, J. F., 1996. Effect of cold on immune function and susceptibility to bacterial infection in toads (Bufo marinw). Pp. 123129 in "Adaptations to the Cold: Tenth International Hibernation Symposium", ed by F. Geiser, G. F. Hulbert and S. C. Nicoll. University of New England Press, Armidale. Carey, C., Bruzgul, J. E., Livo, L. J., Walling, M. L., Kuehl, K. A,, Dixon, B. F., Pessier, A. F!, Alford, R. A. and Rogers, K. B., 2006. Experimental exposures of Boreal Toads (Bufo bweas) to a pathogenic chytrid fungus (Batrachochytrium dendrobatzdis). EcoHealth 3: 5-2 1. Carruthers, R. I. and Haynes, D. L., 1986. Temperature, moisture and habitat effects on Entomophthoraceae muscae (Entomophthorales: Entomopthoraceae) conidial germination and survival in the onion agroecosystem. Environmental Entomology 15: 1154-1 160.

ROWLEY and ALFORD: FACTORS AFFECTING VARIATION IN DISEASE Carruthers, R. I., Larkin, T. S. and Firstencel, H., 1992. Influence of thermal ecology on the mycosis of a rangeland grasshopper. Ecol. 73: 190-204. Casterlin, M. E. and Reynolds, W. W., 1977a. Behavioral fever in anuran amphibian la~vae.Li$ Sci. 20: 593-596. Casterlin, M. E. and Reynolds, W. W., 1977b. Behavioral fever in crayfish. Hydrobiologia 56: 99-101. Casterlin, M. E. and Reynolds, W. W., 1979. Fever induced in marine arthropods by prostaglandin E l . L$e Sci. 25: 1601-1604. Cohen, N., 1966. Tissue transplantation immunity in the adult newt, Diemictylus virzdescens 111: The effects of x-irradiation and temperature on the allograft reaction. J. Experi. Zool. 163: 23 1-240. Conlon, J. M., 2004. The therapeutic potential of antimicrobial peptides from frog skin. Rev. i n Medical Microbiology 15: 17-25. Cote, I. and Poulin, R., 1995. Parasitism and group size in social animals: a meta-analysis. Behav. Ecol. 6: 159-165. Covert, J. B. and Reynolds, W. W., 1977. Survival value of fever in fish. Nut. 267: 43-45. Cowling, W. A. and Gilchrist, D. G., 1982. Effect of light and moisture on Stemphylium leaf spot of alfalfa. Plant Disease 66: 291-294. Davidson, C., Benard, M. F., Shaffer, H. B., Parker, J. M., O'Leary, C., Conlon, J. M. and Rollins-Smith, L. A. 2007. Effects of chytrid and carbaryl exposure on survival, growth and skin peptide defenses in foothill yellow-legged frogs. Environmental Sci. & Technology 41: 1771-1776. DeCastro, F., and Bolker, B. 2005. Mechanisms of disease-induced extinction. Ecol. Letters 8: 117-1 26. Dinnik, J. A. and Dinnik, N. N., 1961. Observations on the longevity of Haemonchus contortus larvae on pasture herbage in the Kenyan highlands. Bull. Epizootic Diseases Afiica 9: 193-208. Dowell, S. F., 2001. Seasonal variation in host susceptibility and cycles of certain infectious diseases. Emerging Infectious Diseases 7: 369-374. Edwards, S., 2000. Survival and inactivation of classical swine fever virus. gterinary Microbiology 73: 175-181. Elliot, S. L., Blanford, S. and Thomas, M. B., 2002. Host-pathogen interactions in a varying environment: temperature, behavioural fever and fitness. Proc. Roy. Sac. Lond. Series B 269: 1599-1607. Enger, (d., Husevsg, B. and Goksgyr, J., 1991. Seasonal variation in presence of Rbrzo salmoniczda and total bacterial counts in Norwegian fish-farm water. Canadian J. Microbiology 37: 6 18-623. Erspamer, V., 1994. Bioactive secretions of the amphibian integument. Pp. 178-350 in "Amphibian Biology", vol. 1 "The Integument", ed by H. Heatwole, G. T. Barthalmus and A. Y. Heatwole. Surrey Beatty and Sons, Chipping Norton. Eyal, Z., Brown, J. F., Krupinsky, J. M. and Scharen, A. L. 1977. The effect of postinnoculation periods of leaf wetness on the response of wheat cultivars to infection by Septoria nodorum. Phytopathology 67: 874-878. Ezenwa, V., 2004. Host social behaviour and parasitic infection: a multifactorial approach. Behau. Ecol. 15: 446-454.

3063

Fromont, E., Pontier, D. and Langlais, M., 1998. Dynamics of a feline retrovirus (FeLV) in host populations with variable spatial structure. Proc. Roy. Sac. Lond. Series B-Biolog. Sci. 265: 1097-1104. Grutter, A. S., 1998. Habitat-related differences in the abundance of parasites from a coral reef fish: an indication of the movement patterns of Hemigymnus melapterm. J. Fish Biol. 53: 49-57. Harris, R. N., James, T. Y., Lauer, A,, Simon, M. A. and Patel, A,, 2006. Amphibian Pathogen Batrachochytrium dendrabatidis is inhibited by the cutaneous bacteria of amphibian species. EcoHealth 3: 53-56. Harris, R. N., Lauer, A., Woodhams, D. C., Simon, M. A., Banning, J. L., Becker, M., Brucker, R. M., Lam, B,. Minbiole, K. F! C., and Alford, R. A. 2008. The role of bacteria in treating amphibians exposed to a lethal fungal pathogen: experimental studies of the efficacy of probiotics in conservation biology. Abstract, ISME 12, the lZth international symposium on microbial ecology. http:/l www.kenes.com/isme12/Posters/l468.htm Hart, B., 1990. Behavioral adaptations to pathogens and parasites: five strategies. Neuroscience and Biobehavioral Rev. 14: 273-294. Harvell, C. D., Kim, K., Burkholder, J. M., Colwell, R. R., Epstein, F! R., Grimes, D. J., Hoffman, E., Lipp, E. K., Osterhaus, A. D. M. E., Overstreet, R. M., Porter, J. W , Smith, G. W. and Vasta, G. R., 1999. Emerging marine diseases: climate links and anthropogenic factors Sci. 285: 1505-1510. Hero, J.-M., Williams, S. E. and Magnusson, W E., 2005. Ecological traits of declining amphibians in upland areas of eastern Australia. J. 2001. 267: 221-232. Hertz, F! E., Huey, R. B. and Stevenson, R. D., 1993. Evaluating temperature regulation by field-active ectotherms: the fallacy of the inappropriate question. Ame?: Naturalist 142: 796-818. Hildemann, W. H. and Haas, R., 1959. Homotransoplantation immunity and tolerance in the bullfrog. J. Immunology 83: 478-485. Hochberg, M. E., 1989. The role of pathogens in biocontrol. Nut. 337: 262-265. Holmes, S. J. I. and Colhoun, J., 1974. Infection of wheat by Septoria nodorum and S. tritici in relation to plant age and air temperature and relative humidity. Trans. Brit. Mycological Soc. 63: 329-338. Holt, R. D., Dobson, A. F!, Begon, M., Bowers, R. G. and Schauber, E. M., 2003. Parasite establishment in host communities. Ecol. Letters 6: 837-842. Hoogland, J. L., 1979. Aggression, etoparasitism, and other possible costs of prairie dog (Sciuridae, Cynomys sp..) coloniality. Behav. 69: 1-35. Hudson, P. J., Dobson, A. F! and Newborn, D., 1998. Prevention of population cycles by parasite removal. Sci. 282: 2256-2258. Hutchinson, V. H. and Erskine, D. J., 1981. Thermal selection and prostaglandin E l fever in the salamander Necturus maculosis. Herpetologica 37: 195-198. Inglis, G. D., Johnson, D. L. and Goettel, M. S., 1996. Effects of temperature and thermoregulation on mycosis by Beauveria bassiana in grasshoppers. Biolog. Control 7: 131-139.

3064

AMPHIBIAN BIOLOGY

Janovy, J., Jr., Snyder, D. and Clopton, R. E., 1997. Evolutionary constraints on population structure: the parasites of Fundulw zebrinus (Pisces: Cyprinodontidae) in the South Platte river of Nebraska. J. Parasitology 83: 584-592. Johnson, M., Berger, L., Philips, L. and Speare, R., 2003. Fungicidal effects of chemical disinfectants, W light, desiccation and heat on the amphibian chytrid, Batrachochytrium dendrobatidis. Diseases of Aquatic Organisms 57: 255-260. Johnson, M. L. and Speare, R., 2003. Survival of Batrachochytrium dendrobatzdis in water: quarantine and disease control implications. Emerging Infectious Diseases 9: 922-925. Karban, R., 1998. Caterpillar basking behaviour and nonlethal parasitism by tachinid flies.J. Insect Behau. 11: 714-723. Huger, M. J., 1978. The evolution and adaptive value of fever. Amel: Scientist 66: 3 8 4 3 . Huger, M. J. and Rothenburg, B. A., 1979. Fever and reduced iron: Their interaction as a host defense response to bacterial infection. Sci. 203: 374-376. Kluger, M. J., Ringler, D. H. and Anver, M. R., 1975. Fever and survival. Sci. 188: 166-168. Krasnov, B., Shenbrot, G., Khokhlova, I., Medvedev, S. and Vatschenok, V., 1998. Habitat dependence of a parasite-host relationship: flea (Siphonaptera) assemblages in two gerbil species of the Negev Desert. J. Medical Entomology 35: 303-3 13.

Longcore, J. E., Fkssier, A. F! and Nichols, D. K., 1999. Batrachochytrium dendrobatidis gen. et sp. nov., a chytrid pathogenic to amphibians. Mycologia 91: 219-227. Matutte, B., Storey, K. B., Knoop, F. C. and Conlon, J. M., 2000. Induction of synthesis of an antimicrobial peptide in the skin of the freeze-tolerant frog, Rana syluatica, in response to environmental stimuli. FEBS Letters 483: 135-138. McDonald, K. and Alford, R. A,, 1999. A review of declining frogs in northern Queensland. Pp i n "Declines and disappearances of Australian Frogs", ed by A. Campbell. Environment Australia, Canberra. McDonald, K. R., Mendez, D., Miiller, R., Freeman, A. B. and Speare, R., 2005. Decline in the prevalence of chytridiomycosis in frog populations in North Queensland, Australia. Pac. Cons. Biol. 11: 114-120. Monroe, J. S., Santini, J. B. and Latin, R., 1997. A model defining the relationship between temperature and leaf wetness duration and infection of watermelon by Colletotrichum orbiculare. Plant Disease 81: 739-742. Morales, H. D., and Robert, J. 2007. Characterization of primary and memory CD8 T-cell responses against ranavirus (FV3) in Xenopw laeuis. J. Krology 81: 2240-2248. Myhre, K., Cabanac, M. and Myrhe, G., 1977. Fever and temperature regulation in the frog, Rana esculenta. Acta Physiologica Scandinauica 101: 219-229.

Lafferty, K. D. and Gerber, L. R., 2002. Good medicine for conservation biology: The intersection of epidemiology and conservation theory. Cons. Bwl. 16: 593-604.

Nichols, D. K., Lamirande, E. W., Pessier, A. P. and Longcore, J. E., 2001. Experimental transmission of cutaneous chytridiomycosis in dendrobatid frogs. J. Wildl. Diseases 37: 1-1 1.

Lauer, A., Simon, M. A., Banning, J. L., Andr, Emily, Duncan, K. and Harris, R. N., 2007. Common cutaneous bacteria from the eastern red-backed salamander can inhibit pathogenic fungi. Copeia, 2007: 630-640.

Ouedraogo, R. M., Cusson, M., Goettel, M. S. and Brodeur, J., 2003. Inhibition of fungal growth in thennoregulating locusts, Locwta mig-ratoria, infected by the fungus Metarhirium anisopliae var acridium. J . Invertebrate Path. 82: 103-109.

Letcher, F! M. and Powell, M. J., 2002. Frequency and distribution patterns of zoosporic fungi from mosscovered and exposed forest soils. Mycologia 94: 761-771.

Parris, K. M. and Cornelius, T. O., 2004. Fungal pathogen causes competitive and developmental stress in larval amphibian communities. Ecol. 85: 3385-3395.

Levine, N. D., 1963. Weather, climate and the bionomics of ruminant nematode larvae. Adu. Veterinary Sci. 8: 215-261.

Pessier, A. P., Nichols, D. K., Longcore, J. E. and Fuller, M. S., 1999. Cutaneous chytridiomycosis in poison dart frogs (Dendrobates spp.) and White's tree frogs (Litoria caerulea). J. Veterinary Diagnostic Investigation 11: 194-199.

Lips, K. R. and Donnelly, M. A., 2002. What the tropics can tell us about declining amphibian populations: current patterns and future prospects. Pp. 2 8 8 4 0 6 in "North American Amphibians: Status and Conservation", ed by M. J. Lannoo. University of California Press, Berkeley, California. Lips, K. R., Reeve, J. D. and Witters, L. R., 2003. Ecological traits predicting amphibian population declines in Central America. Cons. Bwl. 17: 1078-1088. Lips, K. R., Brem, F., Brenes, R., Reeve, J. D., Alford, R. A,, Voyles, J., Carey, C., Livo, L., Pessier, A. F! and Collins, J. F!, 2006. Emerging infectious disease and the loss of biodiversity in a Neotropical amphibian community. Proc. Nut. Acad. Sci. USA 103: 3165-3170. Loehle, C., 1995. Social barriers to pathogen transmission in wild animal populations. Ecol. 76: 326-335.

Piotrowski, J. S., Annis, S. L. and Longcore, J. E., 2004. Physiology of Batrachochytrium dendrobatidis, a chytrid pathogen of amphibians. Mycologia 96: 9-15. Puschendorf, R., Bolafios, F. and Chaves, G., 2006. The amphibian chytrid fungus along an altitudinal transect before the first reported declines in Costa Rica. Biol. Cons. 132: 136-142. Rachowicz, L. J. and Vredenburg, V. T., 2004. Transmission of Batrachochytrium dendrobatzdis within and between amphibian life stages. Diseases of Aquatic Organisms 61: 75-82. Rachowicz, L. J., Knapp, R. A., Morgan, J. A. T, Stice, M. J., Vredenburg, V. T, Parker, J. M. and Briggs, C. J., 2006. Emerging infectious disease as a proximate cause of amphibian mass mortality. Ecol. 87: 1671-1683.

ROWLEY and ALFORD: FACTORS AFFECTING VARIATION IN DISEASE Retallick, R. W., 2002. Using experimental translocations to learn about declines in Queensland's frog populations. Implementation of Queensland's threatened frog recovery plans- Experimental Ecology. Queensland Parks and Wildlife Service, Atherton, Queensland. 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.

3063

Sherman, E., 2008. Thermal biology of newts (Notophthalmw viridescens) chronically infected with a naturally occurring pathogen. J. Thermal Biol. 33: 27-3 1. Simmaco, M., Mignogna, G. and Barra, D., 1998. Antimicrobial peptides from amphibian skin: what do they tell us? Biopolymers (Peptide Sci.) 47: 435-450. Stables, J. N. and Chappell, L. H., 1986. Diplostomum spathaceum (Rud. 1819): effects of physical factors on the infection of rainbow trout (Salmo gairdneri) by cercara. Parasitology 93: 71-79.

Reynolds, W. W., Casterlin, M. E. and Covert, J. B., 1976. Behavioural fever in teleost fishes. Nut. 259: 41-42.

Steck, F. and Wandeler, A. (1980) The epidemiology of fox rabies in Europe. Epidemiological Rev. 2: 71-96.

Roland, J. and Kaupp, W. J., 1995. Reduced transmission of forest tent caterpillar (Lepidoptera: Lasiocampidae) nuclear polyhedrosis virus at the forest edge. Environmental Entomology 24: 1175-1 178.

Swearer, S. E. and Robertson, D. R., 1999. Life histov, pathology and description of Kudoa oviuora N. sp. (Myxozoa, Myxosporea): an ovarian parasite of Carribean labroid fishes. J. Parasitology 85: 337-353.

Rollins-Smith, L. A. and Conlon, J. M., 2005. Antimicrobial peptide defenses against chytridiomycosis, an emerging infectious disease of amphibian populations. Developmental and Compar. Immunology 29: 589-598.

Swinton, J., 1998. Extinction times and phase transitions for spatially structured closed epidemics. Bull. Mathematical Biol. 60: 2 15-230.

Rollins-Smith, L. A., Reinert, L. K., Miera, V. and Conlon, J. M., 2002b. Antimicrobial peptide defenses of the Tarahumara frog, Rana tarahumarae. Biochemical ti+ Biophysical Res. Communicatirms 297: 361. Rollins-Smith, L. A,, Carey, C., Longcore, J., Doersam, J. K., Boutte, A., Bruzgal, J. E. and Conlon, J. M., 2002a. Activity of antimicrobial skin peptides from ranid frogs against Batrachochytrium dendrobatidis, the chytrid fungus associated with global amphibian declines. Developmental and Compar. Immunology 26: 471-479. Rollins-Smith, L. A, Doersam, J. K., Longcore, J., Taylor, S. K., Shamblin, J . C., Carey, C. and Zasloff, M. A., 2002b. Antimicrobial peptide defenses against pathogens associated with global amphibian declines. Developmental and Compar. Immunology 26: 63-72. Rollins-Smith, L. A,, Woodhams, D. C., Reinert, L. K., Vredenburg, V. T., Briggs, C. J., Nielsen, P F. and Michael Conlon, J., 2006. Antimicrobial peptide defenses of the mountain yellow-legged frog (Rana mwcosa). Developmental and Compar. Immunology 30: 831-842. Rowley, J. J. L. and Alford, R. A., 2007a. Behaviour of Australian rainforest stream frogs may affect the transmission of chytridiomycosis. Diseases of Aquatic Organisms 77: 1-9. Rowley, J. J. L. and Alford, R. A., 2007b. Movement patterns and habitat use of rainforest stream frogs in northern Queensland, Australia: implications for extinction vulnerability. Wildl. Res. 34: 371-378. Rowley, J. J. L., Skerratt, L. F., Alford, R. A. and Campbell, R., 2007. Retreat sites of rain forest stream frogs are not a reservoir for Batrachochytraum dendrobatidis in northern Queensland, Australia. Diseases of Aquatic Organisms 74: 7-1 2. Saumier, M. D., Rau, M. E. and Bird, D. M., 1986. The effect of Trichinella pseudospiralis infection on the reproductive success of captive American kestrels (Falco spaveriw). Canadian J . Zool. 64: 2 123-2 125. Sheafor, B., Davidson, E. W., Parr, L. and Rollins-Smith, L., 2008. Antimicrobial peptile defenses in the salamander, Ambystoma tigrinum, against emerging amphibian pathogens. J. W5ldl. Dveases 44: 226-236.

Swinton, J., Woolhouse, M. E. J., Begon, M. E., Dobson, A. P, Ferroglio, E., Grenfell, B. T., Guberti, V., Hails, R. S., Heesterbeek, J. A. P, Lavazza. A., Roberts, M. G., White, P J. and Wdson, K., 2001. Microparasite transmission and persistence. Pp. 83101 in "The Ecology of Wildlife Diseases", ed by P J. Hudson, A. Rizzoli, B. T Grenfell, H. Heesterbeek & A. P Dobson.. Oxford University Press, Oxford. Taylor, S. K., Williams, E. S. and Mills, K. W , 1999. Effects of malathion on disease susceptibility in Woodhouse's toads. J. Wildl. Diseases 35: 536-541. Thomas, M. B. and Blanford, S., 2003. Thermal biology in insect-parasite interactions. Trends in Ecol. and Euol. 18: 344-350. Tompkins, D. M., White, A. R. and Boots, M., 2003. Ecological replacement of native red squirrels by invasive greys driven by disease. Ecol. Letters 6: 189-196. Van Rensburg, P J. J., Skinner, J. D. and Van Aarde, R. J., 1987. Effects of feline Panleucopaenia on the population characteristics of feral cats on Marion Island. J. Applied Ecol. 24: 63-73. Vaughn, L. K., Bernheim, H. M. and Kluger, M. J., 1974. Fever in the lizard Dipsosaurw dorsalis. Nut. 252: 473474. Vermeer, K., 1969. Endoparasitic variation between California Gulls and Ring-billed Gulls. Ibis 111: 393-395. Voyles, J., Berger, L., Young, S., Speare, R., Webb, R., Warner, J., Rudd, D., Campbell, R., and Skerratt, L. F, 2007. Electrolyte depletion and osmotic imbalance in amphibians with chytridiomycosis. Diseases of Aquatic Organism 77: 113-1 18. Ward, J. R. and Lafferty, K. D., 2004. The elusive baseline of marine disease: are diseases in ocean ecosystems increasing? PLoS Biol. 2: e120. Watanabe, H., 1987. The host population. Pp. 71-1 12 in "Epizootiology of insect diseases", ed by J. R. Fuxa and Y. Tanada. John Wiley and Sons, New York. Watson, D. W., Mullens, B. A, and Petersen, J. J., 1993. Behavioural fever response of Musca domestica (Diptera: Muscidae) to infection by Entomophthora muscae (Zygomycetes: Entomophthorales). J . Invertebrate Path. 61: 10-16.

3066

AMPHIBIAN BIOLOGY

Webber, R. A., Rau, M. E. and Lewis, D. J., 1987. The effects of Plagimchis noblei (T'matoda: Plagiorchiidae) metacercariae on the susceptibility of Aedes aegypti larvae to predation by guppies (Poecilia reticulata) and meadow voles (Microtus pennsyluanicus). Canadian J. Zool. 65: 2346-2348. Weldon, C. and Du Preez, L., 2004. Decline of the Kihansi Spray Toad, Nectophrynoides asperginis, from the Udzungwa Mountains, Tanzania. Froglog 62: 2-3. Whiles, M. R., Lips, K. R., Pringle, C. M., klham, S. S., Bixby, R. J., Brenes, R., Connelly, S., ColonGaud, J. C., Hunte-Brown, M., Huryn, A. D., Montgomery, C. and Peterson, S., 2006. The effects of amphibian population declines on the structure and function of Neotropical stream ecosystems. Frontiers Ecol. Environment 4: 27-34. Williams, S. E. and Hero, J.-M., 1998. Rainforest frogs of the Australian wet tropics: guild classification and the ecological similarity of declining species. Proc. Roy. Soc. Lond. B 265: 597-602. Wobester, G., Hunter, D. B., Wright, B., Nieman, D. J. and Isbister, R., 1979. Avian cholera in waterfowl in Saskatchewan in spring. J. Wldl. Diseases 15: 19-24. Woodhams, D. C. and Alford, R. A. 2005. The Ecology of chytridiomycosis in rainforest stream frog assemblages of tropical Queensland. Cons. Biol. 19: 1449-1459.

Woodhams, D. C., Alford, R. A. and Marantelli, G., 2003. Emerging disease of amphibians cured by elevated body temperature. Dweases of Aquatic Organisms 55: 65-67. Woodhams, D. C., Rollins-Smith, L. A,, Alford, R. A,, Simon, M. A. and Harris, R. N., 2007b. Innate immune defenses of amphibian skin: antimicrobial peptides and more. Anim. Cons. 10: 425428. Woodhams, D. C., Voyles, J., Lips, K. R., Carey, C. and Rollins-Smith, L. A., 2006. Predicted disease susceptibility in a Panamanian amphibian assemblage based on skin peptide defenses. J. Wildl. Diseases 42: 207-218. Woodhams, D. C., Ardipradja, K., Alford, R. A,, Marantelli, G., Reinert, L. K. and Rollins-Smith, L. A., 2007a. Resistance to chytridiomycosis varies among amphibian species and is correlated with slun peptide defenses. Anim. Cons. 10: 409-417. Woodhams, D. C., Rollins-Smith, L. A,, Carey, C., Reinert, L., Tyler, M. J. and Alford, R. A,, 2005. Population trends associated with skin peptide defenses against chytridiomycosis in Australian frogs. Oecologia 146: 531-540. Woodhams, D. C., Vredenburg, V. T, Stice, M. J., Simon, M. A.,Billheimer, D., Shakkhtour, B., Shyr, Y , Briggs, C. J., Rollins-Smith, L. A., and Harris, R. N. 2007c. Symbiotic bacteria contribute to innate immune defenses of the threatened mountain yellow-legged frog, Rana rnuscosa. Biolog. Cons. 138: 390-398.

CHAPTER 4

Digenetic Trematodes and their Relationship to Amphibian Declines and Deformities Jason Rohr, Thomas Raffel, and Stanley K. Sessions

I. Introduction II. Digenetic Trematode Biology Ill. Trematodes and Deformed Amphibians A. The "Handicapped Frog Hypothesis" B. Co-evolution and Limb Deformities IV. Amphibian Immunity Against Helminths V. Trematodiasis as an Emerging Disease of Amphibians VI. Variation in Trematode Virulence and Amphibian Susceptibility A. What Makes Amphibian Species Particularly Susceptible to Trematodes? B. What Makes Trematode Species Particularly Deadly to Amphibians?

VII. Natural Factors Affecting Amphibian Trematode Infections VIII. Anthropogenic Factors Affecting Amphibian Trematode Infections A. Nutrient Inputs and Trematode Infections B. Pesticides and Trematode Infections C. Contrariwise Views on Amphibian Trematode Infections and Deformities IX. Conclusions and Future Research Directions X. Acknowledgements XI. References

I. INTRODUCTION

E

MERGING diseases in amphibians have recently become the focus of intense research, especially in connection with anthropogenic habitat change and amphibian declines. Of particular concern are apparently newly emerging diseases sometimes involving widespread pathogens previously thought to be rare and/or harmless (Daszak et al. 2003; Stuart et al. 2004). One such case is chytridiomycosis, thought to be emerging and directly involved in amphibian declines, especially in the Neotropics (Berger et al. 1998; Daszak et al. 2003; Lips et al. 2006). The purpose of the present chapter, however, is to focus on another example of an apparently emerging amphibian disease, digenetic trematode infections (Johnson et al. 2003; Skelly et al. 2006). Certain trematode infections can cause grotesque limb deformities, kidney damage, debility, and mortality (Fried et al. 1997; Johnson and Sutherland 2003; Schotthoefer et al. 2003a). After a brief review of the biology and diversity of digenetic trematodes, environmental factors that affect their interaction with amphibians are examined, especially focusing on anthropogenic environmental perturbations. Then a particular trematode (Ribeioria ondatrae) is treated and an evaluation made of evidence that the deformities produced by this trematode are part of an adaptive component of the parasite's life history (the "handicapped frog hypothesis"; Sessions 2003). Also discussed is the co-evolution of the parasite with its several hosts in a complex life cycle. This provides an opportunity to review what is known about amphibian immune

3068

AMPHIBIAN BIOLOGY

defence mechanisms and corresponding infective adaptations utilized by the parasites. Evaluation is made of the evidence that trematodiasis is an emerging disease, followed by an assessment of what makes certain amphibian species particularly susceptible to trematodes and what makes certain trematode species particularly deadly to amphibians. The possible connection between parasite disease and amphibian decline is addressed by examining the incidence of trematode disease and its relationship to other environmental factors, including natural factors and anthropogenic influences such as habitat fragmentation, nutrient input, and pesticide pollution, within a population context. Finally comments are offered concerning future research directions for the study of amphibian trematodes. 11. DIGENETIC TREMATODE BIOLOGY

Trematodes (or flukes) represent a class in the phylum Platyhelminthes (flatworms), characterized by a dorso-ventrally flattened body in which the internal organs are embedded in parenchyma and the animal has one or two holdfast organs ("suckers").The class includes at least 200 families and 18,000 - 28,000 species (Cribb et al. 2001; Poulin and Morand 2004). The digenetic trematodes (subclass Digenea), characterized by the presence of both an oral and a ventral sucker or acetabulum (Fig. I), constitute the vast majority of trematode diversity. Digenetic trematodes are obligate parasites of molluscs and vertebrates. The digestive system of trematodes includes an anterior mouth, muscular pharynx, oesophagus, and usually two intestinal ceca (Fig. I). Most digenetic trematodes are hermaphroditic (monoecious) but some (e.g. schistosomatids) are dioecious and sexually dimorphic. Various organs easily visible inside the adult trematode include the digestive system (oesophagus and ceca), the uterus, ovary, and ovarian follicle, the testes and seminal vesicles, and the excretory vesicle (Fig. 1).

&

oral sucker

Seminal vesicle Ventral sucker Cecum Testis

-

Uterus 0vri-y I

Seminal receptacle Excretory vesicle

Fig. 1. Structure of a generalized adult digenetic trematode. After Barnes (1968).

Digenetic trematodes usually have complex life cycles, often involving two or more intermediate hosts in addition to a definitive host (i.e., the host for the reproductive adult parasite) (Noble and Noble 1982; Fig. 2). All trematodes have a mollusc, usually a gastropod, as the first intermediate host, but secondary intermediate hosts can include amphibians, fish, insects, or gastropods. The definitive host is generally a vertebrate, often a bird or a mammal. Adult trematodes usually reside in the digestive tract of the definitive host.

Adult

First intermediate host niett~cere,~~.ial cysts

Fig. 2. Generalized life cycle of a digenetic trematode (see text). The definitive ("primary") host is usually a vertebrate predator, and the first intermediate host is usually a mollusc (snail). The second intermedmte host is prey for the primary host.

ROHR, RAFFEL ET AL: DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3069

Typically trematode eggs are released from the definitive host into the environment, often in the host's faeces, although the life cycle varies among species. The eggs then hatch and release miracidia, a swimming infectious stage that seeks out a molluscan first intermediate host or, alternatively, are eaten by and hatch inside a mollusc. Once inside a mollusc, the trematode undergoes "polyemb~yonicamplification". This is a kind of asexual reproduction resembling Russian dolls, in which each stage produces and gives birth to numerous embryos of the subsequent stage (Fig. 3). First, a miracidium develops into a sporocyst. This stage produces either cercariae (the final infective stage) or an additional amplifying stage called a redia, which in turn produces cercariae. Sporocysts and rediae can also give birth to additional sporocysts or rediae, so that hundreds or thousands of

Fig. 3. Embryonic amplification in a digenetic trematode. A: miracidium containing germ cells. B: sporocyst containing immature rediae (germ balls). C: Redia containing immature cercariae. D: mature cercariae.

Fig. 4. Sporocyst (removed from snail) giving birth to rediae

cercariae can be produced from a single infection. It is an amazing and educational spectacle to open up one of these infected snails under a dissecting microscope and view the various stages in the trematode's life cycle (Fig. 4). Mature cercariae emerge from the molluscan first intermediate host into the environment, a process that can be harmful or even lethal to the mollusc (Noble and Noble 1982). Once outside, the trematodes are challenged by the problem of getting to the intestine of a definitive host. Most trematodes resolve this issue by utilizing a second intermediate host that is prey for the definitive host. If the definitive host is a bird, for example, the second intermediate host might be a fish or amphibian that is a prey species

3070

AMPHIBIAN BIOLOGY

Fig. 5. Ribeiroia cercaria (hematoxylin).

Fig. 6. Stylet on the oral sucker of an armatae cercaria (phase contrast).

of the bird. Trematodes with amphibian definitive hosts frequently infect invertebrate intermediate hosts. Cercariae actively swim to their second intermediate host using a muscular tail (Fig. 5). Once they infect a second intermediate host, the cercariae drop their tails and form cysts. Some cercariae form cysts on the surface of the second intermediate host, while others penetrate the skin and form cysts in the host's tissues. Cercariae that penetrate the tissues of their hosts are usually equipped with a sharp stylet on the oral sucker (Fig. 6) that allows them to poke a hole in the integument through which the worm then squeezes. Cercariae can form a cyst within a minute or two of contacting the host; this encysted stage is called a metacercaria (Fig. 7). Metacercariae usually induce a reaction in which the host forms a connective tissue capsule around the cyst, and then remains relatively inactive. The trematode's life cycle is completed if, and when, the infected secondary host is eaten by the definitive host, at which time the metacercariae excyst and mature inside the definitive host. 111. TREMATODES AND DEFORMED AMPHIBIANS

fig. 7. Top: Ribeiroia cercariae immediately after encysting in the tail and hind limb region of a tadpole. Their dropped tails can b e seen below the tadpole's tail fin. Bottom: Metacercarial cysts of Echinostorna triuoluis embedded in the kidney of a Rana tadpole.

The occurrence of "deformed" (or "malformed") amphibians became a major environmental issue by the late 1990s, fueled by the fear that the abnormalities were caused by W-irradiation, chemical pollution, or a combination of environmental factors. The causal role of trematodes, identified as Ribeiroia sp. . by, Sessions et al. (1999), in the development of frog and salamander limb deformities has since been confirmed by numerous studies (Sessions and Ruth 1990; Johnson et al. 1999; 2001a,b; Sessions et al. 1999; Stopper et al. 2002). The genus

ROHR, RAFFEL ET AL: DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3071

Ribeiroia is identifiable by unique projections from the oesophagus known as diverticulae (Fig. 8). Like other digenetic trematodes, Ribeiroia, identified as R. odontrae by Johnson et al. (1999), has a complex life cycle involving several hosts (Fig. 9). The preferred definitive host is thought to be a predatory aquatic bird (e.g., heron) that feeds on fish and amphibians. The first intermediate hosts are planorbid snails; amphibians serve as the second intermediate hosts. Experiments have revealed that Ribeiroia infections can cause tadpoles to grow extra limbs and a wide variety of other debilitating deformities or malformations (Fig. 10). These deformities are now believed to represent an adaptive hostmodification strategy of the parasite whereby its chances of survival, i.e. to be captured and eaten by a primary host, are increased. This idea is known as the "handicapped frog hypothesis" (Sessions 2003).

Fig. 8. Ribeiroia anatomy, showing diagnostic oesophageal diverticulae (arrow).

.

?l"..

%'

4, A

;,\

Fig. 9. Life cycle of Ribeiroia ondatrae (drawing by B. Ballengee).

F -:

.A

Fig. 10. Some examples of cleared and stained parasite-induced deformities in Pacific treefrogs (Hyla [Pseudacrir] regilla).

3072

AMPHIBIAN BIOLOGY

A. The "Handicapped Frog Hypothesis" Survival of encysted trematodes depends on the secondary host being eaten by a primary host, so any mechanism that "handicaps" the secondary host and increases its probability of being consumed by a definitive host will be adaptive for the parasite. The process by which Ribeiroia cercariae attack their amphibian hosts and cause them to develop deformed limbs has been observed (Stopper et al., 2002). When Ribeiroiu cercariae encounter a frog tadpole, they attach and crawl along the surface of the body. A few individuals migrate into the mouth or the spiracle, but the majority crawl towards the cloaca1 vent and form cysts in the crease between the body and tail, where the hind-limb buds are located (Fig. 7). The cercariae do not penetrate the skin at first but instead form mectacercarial cysts on the surface of the skin. Over the next few hours, however, the cysts gradually sink into the tissues, which become badly inflamed and swollen, and the cysts become completely embedded in the tissues in and around the limb buds within one day of infection. This process can completely scramble the cells of the developing hind-limb buds (Stopper et al.

Fzg I I Histological preparations of the hlnd limb region of young Rana pzpzens tadpoles showing (a) normal llmb buds and (b) limb buds Infected with Rzbeirosa cysts BrdUlabelled cells are visualized wlth immunocytochemlstry to show patterns of dividlng Ilmb-bud cells Fig. 12. Trematode (presumably Rzbezroza) cysts (C) and supernumerary llmb (S) In Hyla (Pseudacrw) regilla. LHL and RHL = left and right hlnd limbs, respectively; P = pelvts.

ROHR, RAFFEL ET AL: DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3073

2002) (Fig. 1l), resulting in a dizzying array of deformities or malformations (Fig. 10). The cysts remain closely associated with the deformed structures throughout the rest of development (Fig. 12). Consistent with this hypothesis, Riberoia-induced deformities are concentrated in the hind limbs (Fig. lo), interfering with locomotion without causing unnecessary damage to the rest of the frog's body. Presumably this interference makes b g s more vulnerable to predation. Once the deformed tadpoles have resorbed their tails they are quite helpless and seldom live long, even in captivity. Something is now known about the developmental mechanism by which the deformities are produced. Virtually all cases of polymelia (extra limbs) show mirror-image morphologies in which a limb with right-handed morphology grows adjacent to a limb with left-handed

Fig. I?. Some examples of mirror-image limb duplications among parasite-induced limb deformities in Pacrfic treefrogs: a = anterior mirror-image duplication (AMID); b = posterior mirror-image duplication (PMID); c = mirrorimage triplication (MIT).

morphology, and a series of three limbs in a row are always right-left-right or left-rightleft (Fig. 13). The evidence of mirror-image symmetry is apparent even in partial duplications. These results are the hallmark of a well-known developmental mechanism called intercalation, a mitotic cellular growth response to the disruption of normal positional relationships of differentiating limb-bud cells (French et al. 1976; Sessions and Ruth 1990; Stopper et al. 2002). Intercalation resulting in identical deformities to those caused by trematode cysts, including mirror-image duplications and bony triangles (also called "bony bridges") (Meteyer 2000), can be induced simply by surgically removing the distal piece of an early tadpole hind-limb bud, rotating it 180 degrees and grafting it back on the stump (Stopper et al. 2002). This is significant because bony triangles were previously thought to be an indicator of retinoid teratogenicity (Gardiner and Hoppe 1999). Thus, intercalation can readily account for most of the observed parasite-induced limb deformities seen in natural populations of amphibians. The role of intercalation is thus supported by both theory and by empirical evidence. B. Co-evolution and Limb Deformities

Parasites must accomplish two things to success~llyproduce handicapped frogs: (1) they must attack tadpoles at a stage in the tadpole's development when the limbs are capable of being deformed, and (2) they must be able to thwart, or at least survive, attack by the host's immune system, especially in the intermediate host, where there is direct contact between parasite and deep tissues of the host. Limb development in tadpoles exhibits

3074

AMPHIBIAN BIOLOGY

"regenerative decline" (Muneoka et al. 1986) whereby early limb buds can respond to perturbation by regenerating new limbs or duplicate limbs via intercalation. As the tadpole approaches metamorphosis, however, this ability gradually declines and is completely lost at metamorphosis. Thus, regenerative decline creates a critical "window of opportunity" for the parasites during early limb-bud stages, after which handicapped frogs will not- be produced. The existence of these windows of opportunity/vulnerability involving the developmental response of limb buds and immunological competence of tadpoles suggests that co-evolutionary dynamics between Ribeiroia and their second intermediate amphibian hosts might feature adaptive changes in defence mechanisms in both host and parasite. IV. AMPHIBIAN IMMUNITY AGAINST HELMINTHS

Host immune systems play an integral role in the life-cycles of parasites and thus must also be important in regulating trematode interactions with their hosts. Research on amphibian immune responses against trematodes has been limited. Nevertheless, amphibian immune systems are largely similar to those of other vertebrates (Rollins-Smith and Cohen 2005), so comparison with mammalian immunity can provide useful insights into amphibian responses (Charlemagne and Tournefier 1998). In mammals, the primary response to larval helminths is granuloma formation (Anthony et al. 2007), similar to the encapsulation response of invertebrates (Oksov 1991). Shortly after a trematode cercaria invades host tissue, neutrophils, macrophages and eosinophils surround the parasite, effectively separating the developing metacercaria from the surrounding tissue and sometimes killing the parasite using a cocktail of chemical weapons, including hydrogen peroxide (Anthony et al. 2007). This process, involving antibody-dependent cell-mediated cytotoxicity (ADCC) (Roitt et al. 1996), is facilitated by the action of CD4+ Th2 cells, which act to recruit more leukocytes to the site of infection, particularly during acquired immune responses to repeated infection by the same type of parasite (Anthony et al., 2007). Increases in basophils and mast cells are also associated with infection by parasitic helminthes (Anthony et al. 2007). Recent evidence suggests that basophils act to recruit other effector cells, such as eosinophils and neutrophils, to sites of helminth infection (Min and Paul 2008). Like mammals, amphibians possess all these leukocyte types; these probably play similar roles in combating trematode infections (Charlemagne and Tournefier 1998). One potential difference from mammals is that amphibian tissue macrophages often contain melanin (Gallone et al. 2002). These melanomacrophages have been associated with trematode infections in fish (Dezfuli et al. 2007) and probably play a role in amphibian granulomatous responses to larval trematode infections. Melanized granulomas are frequently observed surrounding trematode metacercariae, for example in the muscles and connective tissues of adult frogs (Johnson et al. 2004b) and in the livers of adult newts (T. R. Raffel, personal observation). Pesticide-induced reductions in eosinophils and melanomacrophages have been associated with increased susceptibility to trematode infection in tadpoles (Kiesecker 2002; Rohr et al. 2008b), suggesting a protective role for these cell types against trematode infections. Abundances of both eosinophils and melanomacrophages have been shown to be negatively associated with trematode load in amphibians (Rohr et al. 2008b). Furthermore, resistance to larval trematode infection is often higher in older tadpoles (Schotthoefer et al. 2003b; Dare et al. 2006; Holland et al. 2007; Raffel et al. in preparation a,b), perhaps corresponding to improved immune responses as tadpoles develop (Flajnik et al. 1987; Rollins-Smith 1998). Ribeiroia cysts do disappear soon after metamorphosis in some species of frogs (Stopper et al. 2002; Rajakaruna et al. 2008), but it is not known whether or not this is due to immunological attack. There is also evidence that cysts in surviving, peri-metamorphic frogs can excyst in their frog host (Sessions, unpublished) but the fate of the excysted flatworm is not known (Sessions et al., unpublished observation).

ROHR, RAFFEL ET AL: DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3075

Amphibians possess all the fundamental components for mounting an acquired immune memory response to parasites (Maniero et al. 2006), but little is known about their capacity to mount memory responses to helminths. Improved resistance to the monogenean Protopolystorna xenopodis has been demonstrated in Xenopus Zumis following multiple exposures, indicating a significant immune memory response to this gastrointestinal parasite (Jackson and Tinsley 2001). There were, however, no apparent population-level effects of immune memory on any of four trematode taxa in a study of parasite age-intensity curves in redspotted newts, despite evidence for memory responses to bacteria and a protist parasite (Raffel et al. 2009). Tadpoles exhibit immune memor). responses to antigens, but acquired immune responses to actual parasites have not been documented (Flajnik et al. 1987; RollinsSmith 1998). Such responses might be limited in tadpoles, because antibody-mediated eosoinophil activation requires the action of mature helper T-cells, which in turn requires a mature thymus. In the frog Xenopus lamis, however, the thymus does not complete development until near metamorphosis, so tadpoles are unable to express appropriate Tcell helper function (Roitt et al. 1998). Up to that point, the tadpoles might have a much more limited immunological defence. It is not known whether thymus development in X. laevis accurately reflects thymus development in other amphibians, although in Rana pzpms, the thymus does not complete its development until soon after tail resorption (stage XXV; Rugh 1951). Clearly more research is needed to determine the importance of acquired immunity to trematode infections in amphibians. V. TREMATODIASIS AS AN EMERGING DISEASE OF AMPHIBIANS

Johnson and colleagues (Johnson et al. 2003; Johnson and Sutherland 2003; Johnson et al. 2004a), Beasley et al. (2004), and Skelly et al. (2006) have argued that certain amphibian trematodes are more common now than they have been historically. Although the arguments for the emergence of these trematodes are circumstantial, they are often quite convincing. Considering that historical data on trematode abundance and prevalence is scant, convincing circumstantial evidence of emergence might now be as good as it will ever be. Johnson et al. (2002, 2003) surveyed numerous ponds and amphibians for malformations and R. ondatrae and compared these data to historical accounts and museum specimens to provide qualitative evidence that malformations caused by R. ondatrae have increased in prevalence. Johnson et al. (2003) could only identify seven historical (pre-1990) records of mass malformations in amphibians associated with R. ondatrae. In contrast, there are presently over 25 sites in the northwest and dozens of sites in Minnesota, Wisconsin, and Illinois that are presumably associated with mass malformations and R. ondatrae. Furthermore, it seems unlikely that most of these recent discoveries of sites where malformations occur are due to heightened surveillance, given that over 12,000 Pacific treefrogs were examined between 1950 and 1980 and few malformations relative to present levels were found (Johnson et al. 2003). Beasley et a1.(2004) argued that Echinostornu trivolvis infections of cricket frogs may be increasing in midwestern United States and might be associated with declines of these frogs. Frogs at sites with detectable levels of herbicides had higher E. trivolvis loads than at sites without detectable levels of herbicides. Beasley et al. (2004) proposed that agrochemical use and habitat modification might be promoting the emergence of E. trivolvis infections of amphibians. Skelly et al. (2006) argued that urbanization was partly driving the increase in amphibian E. trivolvis infections. They showed a positive relationship between human densities surrounding ponds and both snail densities and the abundance of E. trivolvis infections in Rana clamitans. Skelly et al. (2006) proposed that humans might be inadvertently elevating snails' resources and densities by increasing inputs of nitrogen, phosphorus, and calcium into wetlands.

AMPHIBIAN BIOLOGY

VI. VARIATION IN TREMATODE VIRULENCE AND AMPHIBIAN SUSCEPTIBILITY In this section two questions are asked: (1) Which amphibian species are most susceptible to trematodes? and (2) Which species or morphotypes of trematodes are most deadly to amphibians? These questions are important because their answers assist in targeting conservation and mitigation efforts for populations and species of amphibians that are at the greatest risk from trematode infections. Most of this section, however, is hypothetical because little is presently known about variation in susceptibility among amphibians or variation in virulence among trematodes. A. What Makes Certain Amphibian Species Particularly Susceptible to Trematodes? It is proposed here that the frequency and duration of exposure to a given trematode species, or perhaps to trematodes in general, will affect the strength of selection for resistance to trematode infection. More specifically, amphibian species with long larval periods, such as many ranid species that overwinter in ponds and thus have two years of exposure to trematode cercariae, have greater exposure to cercariae and should have evolved stronger resistance to infection than have species with short larval periods. This hypothesis was supported by a recent comparison of two species of frogs. Tadpoles of the American toad, Bufo americanus (which has a larval period of about four to five weeks), were significantly more susceptible (as measured by mortality) to three species of trematodes than were green frog (Rana clamitans) tadpoles, which over-winter in ponds (Hall et al., in preparation). Although the duration of the larval period will undoubtedly influence net exposure to trematodes, other factors will also affect exposure. For instance, species of frogs vary in the amount of time adults spend in the water where they are exposed to cercariae. The phenology of snails, trematodes, and amphibians could also be an important determinant of exposure levels. For example, in northern North America the abundance of snails and trematodes seems to peak in mid summer (Lemly and Esch 1984), so amphibians that breed early (e.g. Rana sylvatica, Ambystoma maculatum) probably have low exposure to trematodes and might be more susceptible if exposed. Finally, different species of amphibians, snails, and trematodes utilize different habitats, for example lotic versus lentic systems. Amphibian species found predominantly in lentic systems should have less exposure and be more susceptible to trematodes found predominantly in lotic systems, and vice versa. Species-level variation in susceptibility to trematodes might also be a function of phylogenetic constraints. For instance, salamanders appear to be less capable of mounting an acquired immune response to parasites than are anurans (but see Raffel et al. 2009) and might be more susceptible to trematodes. It is not clear, however, whether this apparent difference between salamanders and anurans is generally true or merely due to unique characteristics of those few species of amphibians and parasites that have been examined so far (Raffel et al. 2009). The age and size of a given amphibian are also likely to affect its susceptibility to trematodes. Recent evidence suggests that older tadpoles had lower proportions of cercariae that successfully encysted and were less likely to die, than was true for younger tadpoles, even when controlling for tadpole size (Schotthoefer et al. 2003b; Holland et al. 2007; Raffel et al.,in preparation a,b). This suggests that tadpole immunity improves with development. Amphibian size, independent of age, should also affect susceptibility to trematodes. Smaller amphibians have less space for trematodes and thus are more likely to succumb to lower trematode loads. In recent work, larger tadpoles exposed to a set number of cercariae had fewer trematodes and were less likely to die than was the case for smaller tadpoles, even after controlling for tadpole developmental stage (Hall et al., in preparation).

B. What Makes Trematode Species Particularly Deadly to A Amphibians? The mode of entry and the type and extent of migration through host tissues (if any) should affect trematode virulence to amphibian hosts. Certain trematode taxa enter through

ROHR, RAFFEL ET AL: DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3077

amphibian orifices such as the cloaca (e.g. Echinostoma spp.), or are obtained by consuming infected intermediate or paratenic hosts followed by innocuous migrations through the host digestive tract (Halipegus spp.). In contrast, some trematode taxa cause substantial tissue damage by using stylets (plagiorchzd and telorchid cercariae) or enzymes (Ribeiroia ondatrae) to burrow into the skin and muscle tissue. Others undergo damaging migrations within the host following infection. Trematodes that enter through the skin can cause substantial injury, with risk of subsequent bacterial infections. The size of the trematode should also affect its virulence. Larger cercariae probably cause more tissue damage and elicit stronger immune responses than do smaller ones. Larger adult trematodes are likely to consume more host resources than are smaller trematode species. Trematode size is also likely to interact with mode of entry. For instance, R. ondatrae, with large cercariae that cause substantial tissue damage upon entry into the host, was more deadly to tadpoles of the American toad and green frog than were plagiorichid and Echinostoma trivolvis cercariae (Hall et al., in preparation). Plagiorchid cercariae cause tissue damage but are smaller than R. ondahue. E. trivolvis cercariae are similar in size to those of R. ondatrae but enter tadpoles through the cloaca, rather than through the skin. Indeed, many small tadpoles exposed to R. ondahue cercariae seemed to consist more of "holes" than of tadpole tissue (Rohr, personal obse~ation). Infracommunity dynamics might also affect trematode virulence. Poor trematode competitors will be kept in check by other parasite species and thus might reach lethal levels less frequently (Kuris and Lafferty 1994; Sousa 1994). Certain combinations of parasites might also have synergistic positive effects on one another by adversely affecting different components of the host immune system. Thus, trematode virulence is likely to be dependent on co-infections. As discussed above, trematodes might also be indirectly lethal by making the host more susceptible to predation. Consequently, how deadly a trematode is will also depend on the strength of trematode-induced behavioural, physiological, and morphological modifications of its amphibian host (Dobson 1988; Moore and Gotelli 1996). Finally, all the factors mentioned above are unlikely to be independent. It is therefore likely that these factors interact in interesting synergistic ways that have yet to be revealed. VII. NATURAL FACTORS AFFECTING AMPHIBIAN TREMATODE INFECTIONS Despite the complexity of trematode life cycles, most seem to have one thing in common: snails are the kingpin. Nearly all trematode species utilize gastropods as first intermediate hosts, and they tend to specialize on a particular snail species more so than they do on any particular definitive or second intermediate host species. Furthermore, snails are particularly important for parasite dynamics due to embryonic amplification of trematodes within snails. Because each miracidium infecting a snail can potentially generate hundreds, thousands, or even hundreds of thousands of infectious cercariae, small fluctuations in snail population sizes might generate large fluctuations in cercarial production in a given pond. As a consequence, gastropod abundance and richness are often positive predictors of amphibian trematode infections (Johnson et al. 2002; Skelly et al. 2006; Rohr et al. 2008b). For example, ponds with the greatest snail densities had amphibians with the highest incidence of R. ondutrae and E. trivolvk infections (Johnson et al. 2002; Skelly et al. 2006), and gastropod richness was a significant positive predictor of total larval trematode loads (all trematode species combined) in Rana pipiens (Rohr et al. 2008b). Except for the abundance and diversity of first intermediate hosts, amphibian age and the seasonality of cercarial shedding from snails might be the most important natural factors affecting larval trematode infections of amphibians. As mentioned above, susceptibility to larval trematode infections has been shown repeatedly to be a function of tadpole development (Schotthoefer et al. 2003b; Dare et al. 2006; Holland et al. 2007; Raffel et al. in preparation a,b). Fine-scale partitioning of tadpole ages revealed that the relationship between tadpole developmental stage and susceptibility to E. trivovlk cercariae is non-linear with the most susceptible tadpoles being of intermediate Gosner stages (Holland et al. 2007;

3078

AMPHIBIAN BIOLOGY

Raffel et al. in preparation a,b). Raffel et al. (in preparation b) suggested that the nonlinearity could be explained by a general improvement in immunity with tadpole age, coupled with the smallest tadpoles having limited kidney space, making them less susceptible to heavy infections. Despite the general reduction in tadpole susceptibility with age, fieldcaught Rana clamitam tadpoles exhibited a curvilinear, asymptotic increase in E. trivolvis infections with age (Raffel et al. in preparation b). Because different mechanisms can generate similar age-parasite-intensity relationships, field data, experiments and model fitting were used to test among plausible drivers of this curvilinear age-intensity relationship for R. clamitam; these drivers included seasonal exposure, age-dependent susceptibility, density-dependent establishment, parasite-induced host mortality, acquired immunity, heterogeneity in susceptibility, and heterogeneity in exposure. The parsimonious explanation for the age-intensity relationship was seasonal exposure to trematodes (Raffel et al. in preparation b). That is, R. clamitam individuals that had the greatest overlap with the peak of seasonal shedding of E. trivolvzli cercariae had the greatest E. trivolvis loads. Amphibian behaviour also seems to affect the risk of trematode infection (Fig. 14). Gray treefrogs (Hyla versicolor) oviposited less often in pools containing snails shedding cercariae than in pools with uninfected snails or no snails, presumably thereby reducing infection risk for their offspring (Kiesecker and Skelly 2000). American toad (Bufo americanus) tadpoles avoided E. trivolvis cercariae and the strength of this avoidance was qualitatively similar to the strength of tadpole avoidance of predation-related cues (Rohr et al. 2009). Certain types of activity can also reduce the risk of infection (Taylor et al. 2004; Koprivnikar et al. 2006~). Thrashing of the body can remove cercariae from the body surface, and activity in general

-m 0

Behavior Morphology Growth/size Lifehistory OsmoImmunoregulation competence

E

cn

E

f

I Pesticides

Drying/moisture

Disease

$ 3 G' 0

Competition

a.

Predation

;;

8

0a.f

2

A

4 8

rr

Fitness

Populationperformance Fig. 14. Primary relationships among natural and anthropogenic stressors, organismic responses, and population performance. Arrows and lines represent connections among direct and indirect effects, respectively.

creates a moving target for cercariae that infect specific regions of the body (e.g. E. trivolvzli must enter through the cloaca), thereby reducing trematode infections (Taylor et al. 2004; Koprivnikar et al. 2006~).Indeed, the mere smell of E. trivolvis cercariae appears to elevate B. americanus activity (Rohr et al. 2009). Changes in community structure can also influence the spread of disease by changing the abundance of hosts or pathogens (density-mediated effects) or by altering host behaviour, host susceptibility, or parasite infectivity (trait-mediated effects) (Lafferty and Holt 2003). Changes in host or parasite abundance have the most obvious effects on parasite dynamics, but transmission can also be affected by factors influencing other trophic levels that then

ROHR, RAFFEL ET AL: DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3059

have top-down or bottom-up effects on the parasite and/or host (Hudson et al. 2002). Predation, for instance, was recently discovered to be an important factor affecting amphibian trematode infections. In long-toed salamanders (Ambystoma macrodactylum), individuals that were missing limbs due to simulated cannibalism (amputation) were more likely to be infected by R. ondatrae and to exhibit limb deformities than were individuals that experienced R. ondatrae alone (i.e. without limb amputation) (Johnson et al. 2006). Caged predatory fish reduced the activity of Rana clamitans tadpoles and increased their E. trivolvis metacercarial loads (Thiemann and Wassersug 2000). Given the importance of activity as a defence against cercarial infections, it is likely that predator-induced reductions in activity made R. clamitans more susceptible to E. trivovlis. In contrast, Raffel et al. (in preparation a) crossed caged predators (eastern red-spotted newts), tadpole density, and E. trivolvis exposure in a study on Bufo americanus tadpoles and found no effect of caged predators on E. trivolvis infections, despite B. americanus significantly reducing their activity in response to the caged predator. Raffel et al. (in preparation a), however, did detect strong effects of density. They revealed that tadpoles held at higher density were more susceptible to trematode infections when per capita trematode exposure was constant, a result comparable to that shown for a different trematode and amphibian species (Dare et al. 2006). Neither the change in tadpole activity nor the cellular immunity could account for this elevated susceptibility, suggesting an alternative mechanism. When exposure was kept constant at the level of the tank, rather than at the level of the individual, doubling density halved per capita exposure for the tadpoles, but resulted in no change in the number of metacercariae per tadpole. This was because the increased susceptibility associated with exposure to high tadpole density was offset by the reduced exposure to cercariae (Raffel et al. in preparation a). Koprivnikar et al. (2008) examined the impact of E. trivolvis infections on the competitive ability of Rana pipiens tadpoles. E. trivolvis infection had no effect on the competitive ability of R. pipiens but it did reduce the growth rates of the tadpoles (Koprivnikar et al. 2008). Similarly, E. trivolvis infections slowed the growth of B. americanus (Raffel et al. in preparation a) and R. clamitans (Fried et al. 1997), but did not seem to affect the growth of Rana sylvatica (Belden 2006). The hydroperiod of the pond also appears to influence trematode infections in amphibians. In ponds that were drying, both tadpoles and trematode-infected snails had decreased survivorship and the tadpoles had higher incidences of trematode infections, possibly because drying concentrated the amphibians and cercariae, thereby facilitating successful transmission (Kiesecker and Skelly 2001). With widespread climatic change, drying of ponds might be considered both a natural and an anthropogenic stressor (Rohr and Madison 2003). Stress, in general, seems capable of elevating the incidence of trematode infections in larval amphibians. Stressors stimulate the release of glucocorticosteroid hormones that can be immunosuppressive (Belden and Kiesecker 2005). Gray treefrogs (Hyla versicolor) exposed to exogenous corticosterone had fewer circulating eosinophils and more Alaria sp. mesocercariae than did treefrogs receiving sham injections (Belden and Kiesecker 2005). These findings provide a mechanism by which both natural and anthropogenic stressors can elevate risk of disease. VIII. ANTHROPOGENIC FACTORS AFFECTING AMPHIBIAN TREMATODE INFECTIONS

Anthropogenic changes to the environment can modify interactions between amphibians and their parasites. For instance, elevated trematode loads have been linked to nutrient and pesticide inputs into wetlands. The evidence for and against these relationships are reviewed and other potentially important anthropogenic factors influencing amphibian trematodes are discussed in the following sections.

3080

AMPHIBIAN BIOLOGY

A. Nutrient Inputs and Trematode Infections Numerous studies have identified links between agriculture, amphibian helminth infections, and deformities (Johnson et al. 2002; Christin et al. 2003; Gendron et al. 2003; Beasley et al. 2004; Christin et al. 2004; Taylor et al. 2005; King et al. 2007; Koprivnikar et al. 2007a; McKenzie 2007), but only recently have the mechanisms underpinning this association been revealed. Johnson et al. (2003) suggested that an increase in artificial impoundments associated with agriculture partly drove increases in amphibian trematodes. These impoundments often receive heavy inputs of fertilizer and cattle waste, and these nutrient inputs were proposed to support a higher biomass of snails as intermediate hosts than in unaffected ponds (Johnson et al. 2003; Johnson and Chase 2004). Indeed, 44 of 59 wetlands associated with R. ondatrue infections were also artificial impoundments or human-altered wetlands (Johnson et al. 2002). "Created" wetlands also represented malformation hotspots in the midwestern United States (Lannoo et al. 2003) and higher helminth loads in frogs from agricultural wetlands (relative to frogs from forested wetlands) were attributed to elevated nutrient inputs (McKenzie 2007). To test the hypothesis that nutrients derived from agriculture could increase the level of R. ondatrae infections in frogs, Johnson et al. (2007) established mesocosms containing snail and amphibian hosts and conducted an experiment crossing fertilizer and additions of R. ondatrae eggs to these mesocosms. Fertilizer additions increased snail abundance, cercarial shedding rates, number of R. ondatrae infections in amphibians, and limb deformities, thereby verifying the postulated causal relationship (Johnson et al. 2007). Belden (2006) showed that nutrient inputs do not increase the pathogenicity or virulence of E. trivolvis, suggesting that the primary effect of eutrophication is to elevate exposure to trematodes. Skelly et al. (2006) suggested that the positive relationship between larval trematode infections of R . clamituns and human population density was due to humans releasing nutrients into wetlands, thereby causing increases in snail populations. B. Pesticides and Trematode Infections

Pesticides common in agricultural landscapes have also been implicated as causes of elevated infections in amphibians. Various pesticides and pesticide mixtures have been shown to be immunosuppressive to both amphibians and snails (Taylor et al. 1999; Kiesecker 2002; Christin et al. 2003; Gilbertson et al. 2003; Christin et al. 2004; Russo and Lagadic 2004; Forson and Storfer 2006a,b; Hayes et al. 2006; Sandland and Carmosini 2006; Brodkin et al. 2007; Davidson et al. 2007; Rohr et al. 2008a,b) and this immune suppression has been linked to elevated amphibian infections (Taylor et al. 1999; Kiesecker 2002; Christin et al. 2003; Forson and Storfer 2006b; Hayes et al. 2006; Davidson et al. 2007; Rohr et al. 2008a,b). For example, Kiesecker (2002) demonstrated that three common pesticides, atrazine, malathion, and esfenvalerate, all suppressed immunity in Rana sylvatica and increased its larval trematode loads. Likewise, relatively high levels of atrazine negatively impacted the immune system of adult Rana pipiens (Houck and Sessions 2006). In a series of studies, Rohr, Raffel, and colleagues examined the lethal and sublethal effects of expected environmental concentrations of atrazine, glyphosate (herbicides), carbaryl, and malathion (insecticides) on E. trivolvis free-living stages and on its first and second intermediate hosts. These chemicals had no detectable effects on the survival of E. trivolvis miricidia, on the infectivity of E. trivolvis cercaria, on the growth, fecundity, or survival of snail first intermediate hosts, or on the survival of tadpole second intermediate hosts (Rohr et al. 2008a, Raffel et al. in press). The pesticides did, however, reduce cercarial survival and increase tadpole susceptibility to E. trivolvis infections (Rohr et al. 2008a). Most importantly, the reduction in exposure to trematodes due to pesticide-induced cercarial mortality (density-mediated effect) was 2.5 times smaller than the pesticide-induced increase in amphibian susceptibility (a trait-mediated effect), suggesting that the net effect of exposure to environmentally realistic levels of pesticides is to elevate the incidence of amphibian trematode infections (Rohr et al. 2008a).

ROHR. RAFFEL ET AL: DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3081

In a follow-up study, Rohr et al. (2008b) surveyed wetlands in Minnesota where they quantified over 240 plausible predictors of larval trematode abundance in the declining northern leopard frog, Ram pipiem. They discovered that the widely used herbicide, atrazine, was the best predictor of larval trematode infections in R. pipimu and that this positive effect was consistent across different taxa of trematodes. Furthermore, they showed that phosphate, a primary ingredient in fertilizers, plays a complementary role, lending further support for the link between pond eutrophication and amphibian trematode infections. The combination of atrazine and phosphates accounted for 74% of the variation in larval trematode abundance (Rohr et al. 2008b). Path and regression analyses suggested that these agrochemicals caused an increase in snail abundance and diversity as well as immunosuppression in R. pipiens. To test whether the detected correlation between atrazine and amphibian trematode loads was causal, Rohr et al. (200%) conducted a community-level mesocosm experiment in which tanks were dosed with atrazine. Tanks with atrazine had significantly less phytoplankton, greater water clarity, and more attached algae and snails than did control tanks (without atrazine). Tanks with atrazine also had immuno-suppressed tadpoles and tadpoles with elevated trematode loads when controlling for eosinophil abundance, hrther supporting a causal relationship between atrazine and elevated trematode infections in amphibians (Rohr et al. 2008b). An increased shedding rate of snails in tanks with atrazine (Johnson et al. 2007) was suggested as the potential mechanism by which atrazine increased trematode loads beyond its affect on immunity. The study of Rohr et al. (2008b) linked the work on pesticides and nutrient inputs by revealing that atrazine and phosphate, two common components of worldwide corn and sorghum production, seem to elevate larval trematode loads in a similar manner; both appear to increase exposure and susceptibility to trematodes by augmenting snail intermediate hosts and suppressing amphibian immune responses. Likewise, in an extensive study of Bufo marinus in Bermuda, Linzey et al. (2003) concluded that agrochemicals and environmental pollutants appear to cause immune suppression, increased susceptibility to trematode infections, limb malformations and possibly declines of amphibian populations. C. Contrariwise Views on Amphibian Trematode Infections and Deformities Not all studies have found a positive relationship between agricultural activities and larval trematode loads of amphibians or between R. ondatrae infections and deformities. Koprivnikar et al. (2006b) showed that 200 pg/L of atrazine reduced E. trivolvis cercarial survival and infectivity and that exposure of R. sylvatica only (cercarie were not exposed) to 30 pg/L of atrazine increased their E. trivolvis infections (2007b), similar to the findings of Kiesecker (2002) and Rohr et al. (2008a, 2008b). Furthermore, the increase in susceptibility of R. sylvatica could not be accounted for by an atrazine-induced change in activity (Koprivnikar et al. 2007b). Similarly, a mixture of metolachlor and atrazine also reduced E. trivolvis cercarial survival (Griggs and Belden 2008). Simultaneous exposure of R. sylvatica tadpoles and E. trivolvis cercariae to 30 pg/L of atrazine, however, caused no increase in tadpole cercarial infections relative to the control treatment (Koprivnikar et al. 2007b).

These results suggest that the primary mechanism by which atrazine elevates infection loads is by increasing periphyton, snails, and amphibian exposure to trematodes. Elevated susceptibility as a contributor cannot be ruled out, however, especially considering that it is not known how long after exposure to pollutants amphibians remain more susceptible to infections. For instance, atrazine exposure early in amphibian development (embryo and larval stages) had seemingly permanent effects on amphibian activity and water conservation, and had delayed effects on their survival (Rohr and Palmer 2005; Rohr et al. 2006b). In addition, Budischak et al. (2008) made a truly remarkable discovery. Just four days of exposure of ranid embryos to an environmentally realistic concentration of the insecticide malathion increased tadpole susceptibility to E. trivolvis infections seven weeks later, indicating that agrochemical exposure can have long-term effects on susceptibility.

3082

AMPHIBIAN BIOLOGY

Three recent field surveys examining the effects of agricultural activities and atrazine on parasite loads in frogs also cast doubt on the effects of pesticides on amphibian trematode infections (Johnson et al. 2002; Koprivnikar et al. 2006a; King et al., 2007). An observational field study that examined over 100 sites and over 60 pesticides revealed that the frequency of frog deformities and R. ondatrae infections was not correlated with any of the measured pesticides; however, only three sites had detectable pesticide concentrations (Johnson et al. 2002). Koprivnikar et al. (2006a) stated that "we found no associations between combined trematode infection a n d . . . the presence of the herbicide atrazine". Only one out of 12 of their sampled wetlands, however, had concentrations of atrazine above the limit of detection; thus, there was insufficient power to test for a relationship. King et al. (2007) revealed that both the amount of agricultural and urban area around wetlands was negatively related to the abundance and diversity of helminth parasites in R. pipiens. Wetlands within each of their treatments (low, medium and high pesticide levels) were clustered spatially, thus making it impossible to ascertain whether the observed patterns were due to pesticide levels or spatial autocorrelation with other important factors (King et al. 2007). King et al. (200'7) suggested that agriculture and urbanization reduce parasite transmission to frogs by reducing the abundance of definitive vertebrate hosts, a suggestion similar to the findings of Rohr et al. (2008b), who showed that the effect of atazine on risk of trematode infection was dependent upon the number of habitat patches "suitable" for definitive hosts (forest, wetlands, and open water) around each wetland, a proxy for visitations by, and the abundance of, definitive hosts. The important message here is that chemical inputs into wetlands cannot increase trematode infections unless there is an ample supply of trematodes initially, a context-dependency that likely explains many conflicting results regarding amphibian trematode infections and anthropogenic factors. Hence, one cannot fully understand the effects of pollution or other anthropogenic factors without knowing something about proxies for the input of trematode eggs into wetlands (a natural factor), which depends upon wetland visitations by infected definitive hosts. For instance, agricultural or urban wetlands surrounded by nothing but unsuitable habitat for definitive hosts could have copious inputs of nutrients and agrochemicals, but are unlikely to have frogs with high levels of trematode infections because these wetlands are unlikely to have consistent inputs of trematode eggs. It was proposed recently that a 'healthy' environment is one rich in parasite species (Marcogliese 2005; Hudson et al. 2006) because the abundance and diversity of parasites often reflects the abundance and diversity of host species on which parasites depend (Hechinger and Lafferty 2005). Although there is certainly merit to this hypothesis, the results presented here have revealed that the effects of anthropogenic factors on risk of disease are extremely complicated. Because of context-dependencies and non-linearities, one cannot categorically assume that high diversity of parasites means a healthy ecosystem. For instance, the effects of pollution are dependent upon the composition of the community (Relyea 2003; Rohr and Crumrine 2005), which can vary substantially across the landscape. While the free-living stages of parasites can be very sensitive to pollution (Lafferty 1997; Lafferty and Kuris 1999; Morley et al. 2003), the net effects of disease risk will depend on the concentration and duration of the exposure, the timing of exposure, delayed and longterm effects of anthropogenic factors, and impacts of environmental stressors on non-host species that compete with, or prey upon, the hosts (Rohr et al. 2006a; Rohr et al. 2008a). Likewise, the effects of other anthropogenic changes, such as modification of land use or change in climate, might also depend on the type and degree of change, the duration of the change, and the impact of any change on the traits and densities of parasites, hosts, and species that interact strongly with hosts. The precise role of trematodes in amphibian limb deformities is also controversial. R. appears to be common in the western United States and is responsible for many amphibian deformities there (Johnson et al. 2002). Substantial evidence, however, is building that, in other parts of the United States, many amphibian deformities cannot be explained o&ae

ROHR, RAFFEL ET AL: DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3083

by R. ondatrae or by limb predation (Skelly et al. 2007). In fact, R. ondutrae is only rarely found in the northeast (Raffel, Rohr, Sessions personal observations) and therefore it does not account for the amphibian deformities found in a Vermont study (Taylor et al. 2005); it was also the least prevalent larval trematode infecting R. piplens in Minnesota (found in only 5 of 18 ponds) (Rohr et al., 2008b). A survey of 5,264 hylid and ranid metamorphs in 42 Vermont wetlands revealed that proximity to agriculture was the best predictor of nontraumatic limb malformations, providing support for the role of chemical toxicants, rather than of trematodes, as the cause of amphibian limb malformations (Taylor et al. 2005). Chemicals were also concluded as a likely cause of amphibian limb deformities in a recently released book on the topic (Lannoo 2008) (but see Sessions, in press). The identification of these missing limb malformations as "nontraumatic" has, however, recently come into question with evidence that limb trauma from selective predation can induce this type of deformity. (Sessions 2009; BallengCe and Sessions 2009). Ultimately, there is much to learn about the effects of environmental change on amphibian deformities and trematode infections, as well as about parasite dynamics in general. The impacts of land use, pollution, invasive species, and climatic change on amphibian development and risk of disease will be important avenues of research for years to come.

IX. CONCLUSIONS AND FUTURE RESEARCH DIRECTIONS Like many parasites, digenetic trematodes are important components of amphibian ecology. Although their role in amphibian population declines has yet to be elucidated, they have the potential to contribute to declines because they can cause substantial mortality in amphibians. As for many parasitic taxa of amphibians, there is much to learn about trematodes. Future research should focus on improving the understanding of amphibian immunity, the population-level effects of helminths, the impacts of reservoir hosts on the severity of amphibian trematode infections, which trematode species are highly virulent to particular amphibian species (and why), which natural and anthropogenic factors affect trematode abundance and virulence, the role of trematodes as drivers of host community dynamics, alternative explanations for amphibian deformities, and the impacts of trematode manipulation of amphibian hosts. Advances on these fronts will reveal the impact that trematodes have on the persistence of amphibian populations and on community dynamics generally. Much progress has been made in understanding the effects of trematodes on amphibians. There are, however, undoubtedly more questions than answers. Following are suggested directions for research into the ecology of trematodes that parasitize amphibians. Immunity of amphibians: Very little is known about the immunity of amphibians, such as the particular defences used against specific parasites, and amphibians' responses to trematodes are no exception. There is still much to learn about the contributions of cellular and hummoral immunity, and physical, biochemical, and behavioural responses to trematodes. Population-level effects: It seems likely that trematodes can regulate amphibian populations given that many trematodes cause mortality in amphibians (Johnson et al. 1999; Johnson et al. 2001a; Schotthoefer et al. 2003a,b; Rohr et al. 2008a). Also, density-mediated increases in survival could compensate for trematode-induced mortality (Vonesh and De la Cruz 2002; Rohr et al. 2006b). Consequently, to assess the potential role, if any, that trematodes play in amphibian declines, one must determine whether they can regulate amphibian populations. Trematode removal studies seem to be the best approach for answering this question (Hudson et al. 1998). Furthermore, they might provide insight into potential antihelmintics that could be effective in remedying infections contributing to population declines.

3084

AMPHIBIAN BIOLOGY

Reservoir hosts: Reservoir hosts that allow highly virulent parasites to persist after they reduce the populations of their primary hosts, are often important factors in declines of a host (de Castro and Bolker 2005). Are there reservoir hosts for the trematodes that are parasitic on amphibians, and if so, which species are they? Natural factors: Despite a long history of ecological research on competition, predation, and parasitism, there is a paucity of knowledge on the effects of competition or risk of predation on the susceptibility and exposure of amphibians to parasites (Raffel et al. 2008). Are competition and predation stressors that can be immunosuppressive? How much of an impact does predation upon free-living stages of trematodes have on trematode infections of amphibians and snails (Schotthoefer et al. 2007)? Can predation on snails reduce the risk of infection of amphibians by trematodes? Anthropogenic factors: There is now strong evidence that various agrochemicals can increase trematode infections of amphibians (Kiesecker 2002; Johnson and Chase 2004; Johnson et al. 2007; Rohr et al. 2008a; Rohr et al. 2008b), and evidence that climatic change might also increase risk of infection (Raffel et al. 2006). There is only a rudimentary understanding of the role of anthropogenic factors in amphibian declines and risk of disease, and little is known about the mechanisms by which anthropogenic factors might affect immunity and subsequent risk of infection. Facilitation: Although the convention in ecology has been to focus on negative interactions among organisms (competition, predation, and parasitism), appreciation for positive interactions is growing (Raffel et al. 2008). Can trematodes have positive indirect effects on amphibians? Other amphibian species appear to be sinks for trematodes (dead end hosts), having positive effects on susceptible amphibian species (Johnson et al. 2008; Johnson and Hartson 2009). How common is parasite-mediated facilitation (Ostfeld and Keesing 2000; Dobson et al. 2006; Raffel et al. 2008b)? Is there selection for susceptible species to mimic species that are resistant to infections (Raffel et al. ZOOS)? Alternative drivers of deformities: While many hot spots for malformed frogs in western United States and perhaps in midwestern United States are driven by exposure to Ribeiroia ondatrae, many hot spots cannot be explained by trematodes, especially on the eastern coast (Taylor et al. 2005; Skelly et al. 2007; Lannoo 2008). Furthermore, a monostome trematode has been discovered recently that also causes amphibian deformities (Rajakaruna et al. ZOOS), suggesting that R . ondatrae might not be the only culprit. Non-trematode drivers of deformities and alternative trematode species should be more thoroughly considered as factors contributing to amphibian malformations. Trematodes' rhythms: Trematodes exhibit diurnal and circannual rhythms that remain understudied. What drives these rhythms, first intermediate, second intermediate, definitive hosts, or something else? What cues are used to entrain these rhythms? Trematodes' manipulation of hosts: Helminths are well known for manipulating their hosts in ways that facilitate transmission of the parasite (Dobson 1988; Moore and Gotelli 1996). Limb deformities induced by R . ondatrae and oedema induced by echinostomes likely increase the incidence of predation by amphibians and thus transmission of the trematode (Sessions and Ruth 1990; Stopper et al. 2002; Johnson and Sutherland 2003; Johnson et al. 2004a; Holland et al. 2007), but this has yet to be demonstrated. Manipulations induced by other trematodes should also be explored. X. ACKNOWLEDGEMENTS

Funds came from a National Science Foundation (NSF: DEB-0809487) to J.R.R. and S.K.S, U.S. Department of Agriculture (USDA: NRI 2008-00622 & 2008-01785) grants to J.R.R., and a U.S. Environmental Protection Agency STAR grant to J.R.R and T.R.R.

ROHR, RAFFEL ET AL: DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3085

XI. REFERENCES Anthony, R. M., Rutitzky, L. I., Urban, J. F., Stadecker, M. J. and Gause, W. C., 2007. Protective immune mechanisms in helminth infection. Nut. Rev. Immunology 7: 975-987. Balleng6e, B. and Sessions, S. K., 2009. Explanation for missing limbs in deformed amphibians. J. Exp. 2001. 10.1002ijez.b.21296

Beasley, V R., Faeh, S. A., Wikoff, B., Staehle, C., Eisold, J., Nichols, D., Cole, R., A. Schotthoefer, A. M., Greenwell, M. and Brown, L. E., 2004. Risk factors and the decline of the northern cricket frog, Acris crepitans: evidence for involvement of herbicides, parasitism, and habitat modifications. Pp. [NEED PAGE NUMBERS HERE] i n "The Status and Conservation of United States Amphibians" ed by M. J. Lannoo. University of California Press, Berkeley. Belden, L. K., 2006. Impact of eutrophication on wood frog, Rana sylvatica, tadpoles infected with Echinostoma trivolvis cercariae. Canadian J . 2001.Revue Canadienne De Zoologie 84: 13 15-1 32 1. Belden, L. K., and Kiesecker, J. M., 2005. Glucocorticosteroid hormone treatment of larval treefrogs increases infection by Ahria sp. trematode cercariae. J. Parasitology 91: 686-688. Berger, L., Speare, R., Daszak, l?, Green, D. E., Cunningham, A. A., Goggin, C. L., Slocombe, R., Ragan, M. A., Hyatt, A. D., McDonald, K. R., Hines, H. B., Lips, K. R., Marantelli, G. and Parkes, H., 1998. Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proc. Nut. Acad. Sci. United States of America 95: 9031-9036.

Daszak, F!, Cunningham, A. A. and Hyatt, A. D., 2003. Infectious disease and amphibian population declines. Divers. Dithib. 9: 141-150. Davidson, C., Benard. M. F., Shaffer, H. B., Parker, J. hi., O'Leary. C., Conlon, J. M. and Rollins-Smith, L A, 2007. Effects of chynid and carbalyl exposure on sunival, growth, and skin peptide defenses in foothill )-ellow-legged frogs. Envisonmental Sci. Technology 41: 17'71-1776. de Castro, F. and Bolker, B., 2009. Mechanisms of disease-induced extinction. Ecol. Letten 8: 117-126. Dezfuli, B. S., Pironi, F., Shinn, k I?, Manera, M. and Giari, L., 2007. Histopathology and ultrastructure of Platichthys Jesus naturally infected with Anisakis simplex s.1. larvae (Nematoda : Anisakidae). J. Parasitology 93: 14 16-1423. Dobson, A., Cattadori, I., Holt, R. D., Ostfeld, R. S., Keesing, F., Krichbaum, K., Rohr, J. R., Perkins, S. E. and Hudson, F! J., 2006. Sacred cows and sympathetic squirrels: The importance of biological diversity to human health. Plos Med. 3: 714-718. Dobson, A. I?, 1988. The population biology of parasiteinduced changes in host behavior. Quart. Rev. Biol. 63: 139-165.

Flajnik, M. F., Hsu, E., Kaufman, J. F. and Dupasquier, L., 1987. Changes in the immune system during metamorphosis of Xenopus. Immunology Today 8: 58-64.

Forson, D. and Storfer, A,, 2006a. Effects of atrazine and iridovirus infection on survival and life-history traits of the long-toed salamander (Ambystoma macrodactylum). Environmental Toxicology and Chemistry 25: 168-173.

Brodkin, M. A,, Madhoun, H., Rameswaran, M. and Vatnick, I., 2007. Atrazine is an immune disruptor in adult northern leopard frogs (Rana pipiens). Environmental Toxicology and Chemistry 26: 80-84. Budischak, S. A., Belden, L. K. and Hopkins, W. A,, 2008. Effects of malathion o n embryonic development and latent susceptibility to trematode parasites in ranid tadpoles. Environmental Toxicology and Chemistry, in press. Charlemagne, J., and Touruefier, A, 1998. Immunology of amphibians. Pp. 63-73 in "Handbook of Vertebrate Immunology", ed by l? I? Pastoret, l? J . Griebel, H. Baszin and A. Govaerts. Academic Press, London. Christin, M. S., Gendron, A. D., Brousseau, E, Menard, L., Marcogliese, D. J., Cyr, D., Ruby, S. and Fournier, M., 2003. Effects of agricultural pesticides on the immune system of Rana pipiens and on its resistance to parasitic infection. Environmental Toxicology and Chemistry 22: 1127-1 133. Christin, M. S., Menard, L., Gendron, A. D., Ruby, S., Cyr, D., Marcogliese, D. J., Rollins-Smith, L. and Fournier, M., 2004. Effects of agricultural pesticides on the immune system of Xenopw laeuis and Rana p$iens. Aquatic Toxicology 67: 33-43. Dare, 0. K., Rutherford, I! L. and Forbes, M. R., 2006. Rearing density and susceptibility of Rana pipiens metamorphs to cercariae of a digenetic trematode. J. Parasitology 92: 543-547.

Forson, D. D. and Storfer, A., 2006b. Atrazine increases ranavirus susceptibility in the tiger salamander, Ambystom tigrinurn. Ecological Applications 16: 23252332.

French, V , Bryant, l? J. and Bryant, S. V, 1976. Pattern Regulation in Epimorphic Fields. Sci. 193: 969-981. Fried, B., Pane, I? L. and Reddy, A., 1997. Experimental infection of Rana pipiens tadpoles with Echinostoma trivoluis cercariae. Parasitology Research 83: 666-669. Gallone, A., Guida, G., Maida, I. and Cicero, R., 2002. Spleen and liver pigmented macrophages of Rana esculenta L. - A new melanogenic system? Pigment Cell Research 15: 32-40. Gardiner, D. M. and Hoppe, D. M., 1999. Environmentally induced limb malformations in mink frogs ( ~ a n septentrionalis). a J. Experi. Zool. 284: 207-216.

Gendron, A. D., Marcogliese, D. J., Barbeau, S., Chrsitin, M. S., Brousseau, l?, Ruby, S., Cyr, D. and Fournier, M., 2003. Exposure of leopard frogs to a pesticide mixture affects life history characteristics of the lungworm Rhabdias ranae. Oecologia 135: 469-476. Gilbertson, M. K., Haffner, G. D., Drouillard, K. G., Albert, A. and Dixon, B., 2003. Immunosuppression in the northern leopard frog (Rana p$kns) induced by pesticide exposure. Environmental Toxicology and Chemistry 22: 101-1 10.

3086

AMPHIBIAN BIOLOGY

Griggs, J. L. and Belden, L. K., 2008. Effects of atrazine and metolachlor on the survivorship and infectivity of Echinostoma trivolvis trematode cercariae. Arch. Environmental Contamination Toxicology 54: 195-202.

Johnson, P T J., Lunde, K. B., Ritchie, E. G. and Launer, A. E., 1999. The effect of trematode infection o n amphibian limb development and survivorship. Sci. 284: 802-804.

Hall, C. A,, Raffel, T R. and Rohr, J. R., Virulence and infectivity of three trematode species to American toads and green frogs. In preparation.

Johnson, P T. J., Lunde, K. B., Ritchie, E. G., Reaser, J. K. and Launer, A. E., 2001b. Morphological abnormality patterns in a California amphibian community. Herpetologica 57: 336-352.

Hayes, T. B., Case, P, Chui, S., Chung, D., Haeffele, C., Haston, K., Lee, M., Mai, V. P, Marjuoa, Y , Parker, J. and Tsui, M., 2006. Pesticide mixtures, endocrine disruption, and amphibian declines: are we understanding the impact? Environmental Health Perspectives 114 (supplement 1): 40-50. Hechinger, R. F. and Lafferty, K. D., 2009. Host diversity begets parasite diversity: bird final hosts and trematodes in snail intermediate hosts. Proc. Rqy. Soc. B. Biolog. Sci. 272: 1059-1066. Holland, M. P, Skelly, D. K., Kashgarian, M., Bolden, S. R., Harrison, L. M. and Cappello, M., 2007. Echinostome infection in green frogs (Rana clamituns) is stage and age dependent. J. Zool. 271: 455-462. Houck, A. and Sessions, S. K., 2006. Could atrazine affect the immune system of the frog, Rana pipiem? Bios 77: 107-112. Hudson, P J., Dobson, A. P, Cattadori, I. M., Newborn, D., Haydon, D. T., Shaw, D. J., Benton, T G. and Grenfell, B. T., 2002. Trophic interactions and population growth rates: describing patterns and identrfying mechanisms. Philos. Trans. Roy. Soc. Lond. Series B. Biolog. Sci. 357: 1259-1271. Hudson, P J., Dobson, A. P. and Lafferty, K. D., 2006. Is a healthy ecosystem one that is rich in parasites? Trends in Ecol. Evol. 21: 381-385. Hudson, P J., Dobson, A. P and Newborn, D., 1998. Prevention of population cycles by parasite removal. Sci. 282: 2256-2258. Jackson, J. A,, and Tinsley, R. C., 2001. Protopolystoma xenopodis (Monogenea) primary and secondary infections in Xenopus laevis. Parasitology 123: 455-463. Johnson, P T. J., and Chase, J. M., 2004. Parasites in the food web: linking amphibian malformations and aquatic eutrophication. Ecol. Letters 7: 52 1-526. Johnson, P T. J., Chase, J. M., Dosch, K. L., Hartson, R. B., Gross, J . A., Larson, D. J., Sutherland, D. R. and Carpenter, S. R., 2007. Aquatic eutrophication promotes pathogenic infection in amphibians. Proc. Nut. Acad. Sci. U.S.A. 104: 15781-15786. Johnson, P T J. and R. B. Hartson, R. B., 2009. All host are not equal: explaining differential patterns of malformations in an amphibian community.J. Anim. Ecol. 78: 191-201 Johnson, P T J., Hartson, R. B., Larson, D. J. and Sutherland, D. R., 2008. Diversity and disease: community structure drives parasite transmission and host fitness. Ecol. Letters 11: 1017-1026. Johnson, P T J., Lunde, K. B., Haight, R. W., Bowerman, J. and Blaustein, A. R., 2001a. Ribeiroia ondutrae (Trematoda : Digenea) infection induces severe lunb malformations in western toads (Bufo bunas). Canadian J . Zoo1.-Revue Canadienne De Zodo@e 79: 370-379.

Johnson, P T. J., Lunde, K. B., Thurman, E. M., Ritchie, E. G., Wray, S. N., Sutherland, D. R., Kapfer, J. M., Frest, T. J., Bowerman, J. and Blaustein, A. R., 2002. Parasite (Ribeiroia ondatrae) infection linked to amphibian malformations in the western United States. Ecological Monographs 72: 151-168. Johnson, P. T. J., Lunde, K. B., Zelmer, D. A. and Werner, J . K., 2003. Limb deformities as a n emerging parasitic disease in amphibians: Evidence from museum specimens and resurvey data. Cons. Biol. 17: 1724-1737. Johnson, P. T. J., Preu, E. R., Sutherland, D. R., Romansic, J . M., Han, B. and Blaustein, A. R., 2006. Adding infection to injury: Synergistic effects of predation and parasitism on amphibian malformations. Ecol. 87: 2227-2235. Johnson, P. T. J . and Sutherland, D. R., 2003. Amphibian deformities and Ribeiroia infection: an emerging helminthiasis. Trends i n Parasitology 19: 332-335. Johnson, P T J., Sutherland, D. R., Kinsella, J. M. and Lunde, K. B., 2004a. Review of the trematode genus Ribeiroia (Psilostomidae): Ecology, life history and pathogenesis with special emphasis on the amphibian malformation problem. Adu in Parasitology 57: 191-253. Kiesecker, J. M., 2002. Synergism between trematode infection and pesticide exposure: A link to amphibian limb deformities in nature? Proc. Nat. Acad. Sci. U.S.A. 99: 9900-9904. Kiesecker, J. M. and Skelly, D. K., 2000. Choice of oviposition site by gray treefrogs: The role of potential parasitic infection. Ecol. 81: 2939-2943. Kiesecker, J. M. and Skelly, D. K., 2001. Effects of disease and pond drying on gray tree frog growth, development, and survival. Ecol. 82: 1956-1963. King, K. C., McLaughlin, J. D., Gendron, A. D., Pauli, B. D., Giroux, I., Rondeau, B., Boily, M., Juneau, P and Marcogliese. D. J., 2007. Impacts of agriculture o n the parasite communities of northern leopard frogs (Rana pipiens) in southern Quebec, Canada. Parasitology 134: 2063-2080. Koprivnikar, J., Baker, R. L. and Forbes, M. R., 2006a. Environmental factors influencing trematode prevalence in grey tree frog (Hyla uersicolor) tadpoles in southern Ontario. J. Parasitology 92: 997-1001. Koprivnikar, J., Baker, R. L. and Forbes, M. R., 2007a. Environmental factors influencing community composition of gastropods and their trematode parasites in southern Ontario. J. Parasitology 93: 992-998. Koprivnikar, J., Forbes, M. R. and Baker, R. L., 2006b. Effects of atrazine on cercarial longevity, activity, and infectivity.J. Parasitology 92: 306-31 1.

ROHR, RAFFEL E T A L : DIGENETIC TREMATODES' RELATIONSHIP: DECLINES & DEFORMITIES 3087 Koprivnikar, J., M. R. Forbes, M. R. and Baker, R. L., 2006c. On the efficacy of anti-parasite behaviour: a case study of tadpole susceptibility to cercariae of Echinostoma trivolvis. Canadian J. Zoo1.-Revue Canadienne De Zoologie 84: 1623-1629. Koprivnikar, J., Forbes, M. R. and Baker, R. L., 2007b. Contaminant effects on host-parasite interactions: Atrazine, frogs, and trematodes. Env. Toxicology Chemistry 26: 2 166-2 170. Koprivnikar, J., Forbes, M. R. and Baker, R. L., 2008. Larval amphibian growth and development under varying density: are parasitized individuals poor competitors? Oecologza 155: 641-649. Kuris, A. M. and Lafferty, K. D., 1994. Community structure - larval trematodes in snail hosts. Ann. Rev. Ecol. Syst. 25: 189-217. Lafferty, K. D., 1997. Environmental parasitology: What can parasites tell us about human impacts on the environment? Parasitology Today 13: 251-255.

Meteyer, C. U., 2000. "Field Guide to Malformations of Frogs and Toads with Radiographic Interpretations. USGS/BRD/BSR-2000-0005,U.S. Geological Survey, Madison, WI. Min, B. and Paul, W. E., 2008. Basophils: in the spotlight at last. Nut. Immunology 9: 223-225. Moore, J. and Gotelli N. J., 1996. Evolutionary patterns of altered behavior and susceptibility in parasitized hosts. Evol. 50: 807-819. Morley, N. J., Irwin, S. W. and Lewis, J. W., 2003. Pollution toxicity to the transmission of larval digeneans through their molluscan hosts. Parasitology 126: S5-S26. Muneoka, K., Hollerdinsmore, G. and Bryant, S. V., 1986. Intrinsic Control of Regenerative Loss in Xenopus laevis limbs. J. Experi. Zool. 240: 47-54. Oksov, I. V., 1991. Tissue level of organization of the host-parasite system. Parazitologzia 25: 3-12.

Lafferty, K. D. and Holt, R. D., 2003. How should environmental stress affects the population dynamics of disease? Ecol. Letters 6: 654-664.

Ostfeld, R. S. and Keesing, F., 2000. Biodiversity and disease risk: The case of lyme disease. Cons. Biol. 14: 722-728.

Lafferty, K. D. and Kuris, A. M., 1999. How environmental stress affects the impacts of parasites. Limnology Oceanography 44: 925-93 1.

Raffel, T. R., Hoverman, J. T., Halstead, N. T., Michel, P. J. and Rohr, J. R., Parasitism in a community context: trait-mediated interactions with competition and predation. In preparation a.

Lannoo, M. J., 2008. "Malformed Frogs: The Collapse of Aquatic Ecosystems". University of California Press, Berkeley. Lannoo, M. J., Sutherland, D. R., Jones, P., Ronsenberv, D., Klaver, R. W., Hoppe, D. M., Johnson, P. T. J., Lunde, K. B., Facemire, C. and Kapfer, J. M., 2003. Multiple causes of the malformed frog phenomenon. Pages 233-262 in "Multiple Stressor Effects in Relation to Declining Amphibian Populations", ed by G. Linder, S. K. Krest, and D. W. Sparling. American Society for Testing and Materials, Conshohocken, PA. Lemly, A. D. and Esch, G. W., 1984. Effects of the trematode Uvulz~erambloplitis on juvenile bluegill sunfish, Lepomis macrochirus - Ecological Implications. J. Parasitology 70: 475492. Linzey, D. W., Burroughs, J., Hudson, L., Marini, M., Robertson, J., Bacon, J. P., Nagarkatti, M. and Nagarkatti, F! S., 2003. Role of environmental pollutants on immune functions, parasitic infections and limb malformations in marine toads and whistling frogs from Bermuda. Internat. J. Env. Health Res. 13: 125-148. Lips, K. R., Brem, F., Brenes, R., Reeve, J. D., Alford, R. A., Voyles, J., Carey, C., Livo, L., Pessier, A. P. and Collins, J. P., 2006. Emerging infectious disease and the loss of biodiversity in a Neotropical amphibian community. Proc. Nut. Acad. Sci. U.S.A. 103: 3165-3170.

Raffel, T. R., Le Gros, R. J., Love, B. C., Rohr, J. R. and Hudson, I? J., 1009. Parasite age-intensity relationships in red-spotted newts: Does immune memory influence salamander disease dynamics? Internat. J. Porasitolog 39: 23 1-241. Raffel, T. R., Martin, L. B. and Rohr, J. R., 2008b. Parasites as predators: unifying natural enemy ecology. Trends Ecol. Evol. 23: 61-618. Raffel, T R., Rohr, J. R., Kiesecker, J. M. and Hudson, P. J., 2006. Negative effects of changing temperature on amphibian immunity under field conditions. Functional Ecol. 20: 819-828. Raffel, 7: R., Rohr, J. R., Sessions, S. K., Lloyd-Smith, J. and Hudson, P. J., The early frog gets the worm: Disentangling the drivers of age-intensity and agedispersion. In preparation b. Raffel, T R., Sheingold, J. L. and Rohr, J . R. Lack of pesticide toxicity in Echinostoma trivolvis eggs and miricidia. J. Parasitology, in press. Rajakaruna, R. S., Piyatissa, P., ayawardena, U. A,, Navaratne, A. N. and Amerasinghe, F! H., 2008. Trematode infection induced malformations in the common hourglass treefrogs. J. Zool. 275: 89-95. Relyea, R. A., 2003. Predator cues and pesticides: A double dose of danger for amphibians. Ecological Applications 13: 1515-1521.

Maniero, G. D., Morales, H., Gantress, J. and Robert, J., 2006. Generation of a long-lasting, protective, and neutralizing antibody response to the ranavirus FV3 by the frog Xenopus. Develop. Compar. Immunology 30: 649-657.

Rohr, J. R. and Crumrine, P. W., 2005. Effects of an herbicide and an insecticide on pond community structure and processes. Ecological Applications 15: 1135-1 147.

Marcogliese, D. J., 2005. Parasites of the superorganism: Are they indicators of ecosystem health? Internat. J. Parasitology 35: 705-7 16.

Rohr, J. R., Kerby, J. L. and Sih, A., 2006a. Community ecology as a framework for predicting contaminant effects. Trends Ecol. Evol. 21: 606-613.

McKenzie, V. J., 2007. Human land use and patterns of parasitism in tropical amphibian hosts. Biolog. Cons. 137: 102-116.

Rohr, J. R. and Madison, D. M., 2003. Dryness increases predation risk in efts: support for an amphibian decline hypothesis. Oecologia 135: 657-664.

3088

AMPHIBIAN BIOLOGY

Rohr, J. R. and Palmer, B. D., 2005. Aquatic herbicide exposure increases salamander desiccation risk eight months later in a terrestrial environment. Environ. Toxicology Chemistry 24: 1253-1 258.

Sessions, S. K., 2003. What is causing deformed amphibians? Pp. 168-186 in "Amphibian Conservation", ed by R. D. Semlitsch. Smithsonian Press, Washington D.C.

Rohr, J. R., Raffel, T R., Sessions, S. K. and Hudson, F! J., 2008a. Understanding the net effects of pesticides on amphibian trematode infections. Ecological Applications 18: 1743-1753.

Sesssions, S. K.,'2009. Book review of: Malformed Frogs, the Collapse of Aquatic Ecosystems" by Michael Lannoo 2008. Herp. Review 40: 121-124.

Rohr, J. R., Sager, T, Sesterhenn, T M. and Palmer, B. D., 2006b. Exposure, postexposure, and densitymediated effects of atrazine on amphibians: Breaking down net effects into their parts. Environmental Health Perspectives 114: 4C50. Rohr, J. R., Schotthoefer, A. M., Raffel, T. R., Carrick, H. J., Halstead, N., Hoverman, J. T, Johnson, C. M., Johnson, L. B., Lieske, C., Piwoni, M. D., Schoff, F! K. and Beasley, V. R., 2008b. Agrochemicals increase trematode infections in a declining amphibian species. Nut. 455: 1235-1239.

Sessions, S. K., Franssen, R. A. and Horner, V. L., 1999. Morphological clues from multilegged frogs: Are retinoids to blame? Sci. 284: 800-802. Sessions, S. K. and Ruth, S. B., 1990. Explanation for naturally-occurring supernumerary limbs in amphibians. J. Experi. 2001. 254: 3 8 4 7 . Skelly, D. K., Bolden, S. R., Freidenburg, L. K., Freidenfelds, N. A. and Levey, R., 2007. Ribeiroia infection is not responsible for Vermont amphibian deformities. Ecohealth 4: 156-163.

Roitt, I., Brostoff, J. and Male, D., 1998. "Immunology". 4th edition. Mosby, London.

Skelly, D. K., Bolden, S. R., Holland, M. F!, Freidenburg, L. K., Freidendfelds, N. A. and Malcolm, T R., 2006. Urbanization and disease in amphibians. Pages 153167 in "Disease Ecology: Community Structure and Pathogen Dynamics", ed by S. K. Collinge and C. Ray. Oxford University Press, New York.

Rollins-Smith, L. A,, 1998. Metamorphosis and the amphibian immune system. Zmmun. Rev. 166: 221-230.

Sousa, W F!, 1994. Patterns and processes in communities of helminth parasites. Trends Ecol. Evol. 9: 52-57.

Rollins-Smith, L. A. and Cohen, N., 2005. Hormones and the immune system of amphibians. Chapter 10 (pp. 2377-2391) i n "Endocrinology", vol. 6 in "Amphibian Biology", ed by H. Heatwole. Surrey Beatty & Sons, Chipping Norton.

Stopper, G. F., Hecker, L., Franssen, R. A. and Sessions, S. K., 2002. How trematodes cause limb deformities in amphibians. J. Experi. Zool. 294: 252-263.

Rohr, J. R., Swan, A,, T Raffel, T R. and Hudson, P J., 2009. Parasites, info-disruption and the ecology of fear. Oecologia 159: 447-454.

Russo, J . and Lagadic. L., 2004. Effects of environmental concentrations of atrazine o n hemocyte density and phagocytic activity in the pond snail Lymnaea stagnalis (Gastropoda, Pulmonata). Env. Pollution 127: 303-311. Sandland, G. J. and Carmosini, N., 2006. Combined effects of a herbicide (atrazine) and predation on the life history of a pond snail, Physa gyrina. Env. Toxicology and Chemistry 25: 2216-2220. Schotthoefer, A. M., Cole, R. A. and Beasley, V. R., 2003a. Relationship of tadpole stage to location of echinostome cercariae encystment and the consequences for tadpole survival.J. Parasitology 89: 475-482.

Stuart, S. N., Chanson, J. S., Cox, N. A., Young, B. E., Rodrigues, A. S. L., Fischman, D. L. and Waller, R. W., 2004. Status and trends of amphibian declines and extinctions worldwide. Sci. 306: 1783-1786. Taylor, B., Skelly, D., Demarchis, L. K., Slade, M. D., Galusha, D. and Rabinowitz, F! M., 2005. Proximity to pollution sources and risk of amphibian limb malformation. Env.Health Perspectives 113: 1497-150 1. Taylor, C. N., Oseen, K. L. and Wassersug, R. J., 2004. On the behavioural response of Rana and Bufo tadpoles to echinostomatoid cercariae: implications to synergistic factors influencing trematode infections in anurans. Canadian J. 2001.-Revue Canadienne De Zoologie 82: 701-706. Taylor, S. K., Williams, E. S. and Mills, K. W., 1999. Effects of malathion o n disease susceptibility in Woodhouse's toads. J. Wildl. Dis. 35: 536-541.

Schotthoefer, A. M., Koehler, A. V., Meteyer, C. U. and Cole, R. A,, 2003b. Influence of Ribeiroia ondatrae (Trematoda : Digenea) infection on limb development and survival of northern leopard frogs (Rana pipiens): effects of host stage and parasiteexposure level. Canadian J. Zoo1.-Revue Canadienne De Zoologie 81: 1144-1153.

Thiemann, G. W. and Wassersug R. J., 2000. Patterns and consequences of behavioural responses to predators and parasites in Rana tadpoles. Biolog. J. Linnean Soc. 71: 513-528.

Schotthoefer, A. M., Labak, K. M. and Beasley, V. R., 2007. Ribeiroia ondatrae cercariae are consumed by aquatic invertebrate predators. J. Parasitology 93: 1240-1243.

Vonesh, J. R. and De la Cruz, O., 2002. Complex life cycles and density dependence: assessing the contribution of egg mortality to amphibian declines. Oecologia 133: 325-333.

CHAPTER 5

Amphibian Malformations Michael J. Lannoo

V. Taxonomic Distribution

I. Introduction II. Historical Perspective Ill. The Nature of the Data on Malformations IV. Types of Malformations A. Missing Limbs B. Missing Limb Segments C. Micromely D. Phocomely E. Taumelia F. Polyextension G. Skin-Webbing H. Arthrogryposis I. Polymelia J. Multiple Limb Segments with Hygroma K. Maxillary Malformations L. Bloating M. Abnormalities of Pigmentation

VI. Causes of Malformations A. Natural Causes B. Anthropogenic Causes VII. Identifying Causes A. Retinoids B. Parasites C. Pollution, Pesticides, and other Chemical Causes VIII. Solutions IX. Acknowledgments X. References

Abbreviations and acromyms used in the text or references: CWB=malformed-frog hotspot in Crow Wing County, Minnesota; EPA=Environmental Protection Agency; JOF, NLA, OSP, WIN=various malformed-frog hotspots in rural Minnesota; LMS, ROI=two malformed-frog hotspots in Meeker County, Minnesota; NEY=malformed-frog hotspot in LeSueur County, Minnesota; SUN= malformedfrog hotspot in Otter Tail County, Minnesota; SVL=snout to vent length; USGS-BRD=United States Geological Survey; UV-B=Ultraviolet-B radiation

I. INTRODUCTION

1

T began in the summer of 1995 in south central Minnesota, with school kids on a field trip (Souder 2000, Lannoo 2008). While exploring a rural wetland, Cindy Reinartz and her junior high school students discovered a large number of malformed northern leopard frogs (Rana pipiens). As word of their discovery spread, many similar sites were discovered and as the State of Minnesota began distributing bottled water to families perceived to be in harm's way, the malformed frog problem, smouldering since the early 1950s, blew up and became a cultural firestorm. The word malformation literally means "bad form." Bad form in most animals means an unintended lack of symmetry, or an imbalance in structure, colour or some other quality. A lack of symmetry can arise through one of three mechanisms. (1) Genetic mechanism: Genes are flawed, o r the expression of genes during development is flawed. Albinism (animals with white bodies and pink eyes) is a familiar

AMPHIBIAN BIOLOGY

3090

genetically-determined malformation with an incidence of between 1:10,000 and 1:40,000 (e.g. Bechtel 1995). (2) Epigenetic mechanism: Genes and gene expression are normal but at the time and site that genes are being expressed some deviation from expected environmental circumstances occurs, e.g. lack of nutrition, toxins that disturb cell-to-cell interactions or disrupt the endocrine system, or presence of parasitic cysts. Both genetic and epigenetic problems are apparent before development is complete and, when present at birth in humans, are called congenital abnormalities.

(3) Post-developmental trauma such as injuries or disease: Injuries can be inflicted by wouldbe predators, sustained during defence of mates or territories, or during amplexus, or arise from environmental exposure (desiccation or frostbite). Infections secondary to injury can also occur and have permanent effects. It can be difficult to determine the cause of any particular malformation but a sound knowledge of natural history offers perspective. For example, in Minnesota and other areas of the Upper Midwest and New England of the United States, most malformed-frog hotspots are isolated wetlands. Frogs are mobile; they travel from wetland to wetland and genes may spread among populations. If genetic mechanisms are the cause of malformations one would expect frogs in wetland clusters to exhibit similar malformations. Furthermore, within any wetland hotspot, more than one species can be affected and many of the malformations found in these species are of the same type. The chance of the same genetic mutation arising in several species at a single site without any sign of mutations in populations from nearby wetlands (and such wetlands are frequently well within the home range of individual frogs) must be astonishingly low. These observations taken together suggest a site-based (epigenetic, or trauma-based) cause. In deciding between epigenetic and trauma-based causes, trauma often seems the least likely. Trauma can and does happen and perhaps most trauma in amphibian larvae comes from failed predation attempts. One would expect trauma to produce scarring, however, and most malformed animals show no signs of scarring, or other evidence of healed (or healing) wounds. Many frogs also have malformations that simply cannot be due to trauma (e.g., misplaced eyes, multiple limbs, multiple bent bones, multiple fluid-filled sacs, abnormal pigment patterns) or, if they were due to trauma, the injuries sustained (missing pelvic and spinal components) would be inconsistent with life. Forelimb malformations are also unlikely to arise from trauma. For most of a tadpole's life, forelimbs are tucked under enclosed gill coverings and therefore hidden from visually-oriented predators, which are unlikely to selectively eat what they cannot see. Given this evidence, the amphibian malformations of most concern here probably arise from epigenetic mechanisms-consequences of the environment in which genetic expression is occurring. Table I. Summary of Martin Ouellet's data (Ouellet, 2000) showing that most'described amphibian malformations have been frog and toad malformations.

Number of Malformed Frogs and Toads Collected Compared to Salamanders Species Affected Sites where Found Numbers Collected

Ratio of Malformed Frogs and Toads to Salamanders

67~26 159~43 11,687~2,512

Ouellet (2000) recently completed a comprehensive review of the literature on amphibian malformations. From that study it can be concluded that most amphibian malformations occur in frogs and toads. He reported malformations in 6'7 species of frogs and toads but in just 26 species of salamanders (a ratio of frog and toad to salamander species of 2.6:l) (Table 1). This discrepancy between frogsltoads and salamanders is even ~ l d e when r considering sites and specimens. Malformed frogs were found at 159 sites but

LANNOO: AMPHIBIAN MALFORMATIONS

3091

salamanders at only 43 (a ratio of frog to salamander sites of 3.7:l). Ouellet also found a total of 11,687 reported malformed frog specimens compared with 2,512 malformed salamanders (a ratio of frog to salamander specimens of 4.5: 1). 11. HISTORICAL PERSPECTrVE

Malformed frogs have been observed throughout the world, although the largest number of reports has come from North America and Europe (Fig. 1). It seems safe to conclude that, wherever there are aquatic frogs and curious people looking at them, malformations will be found. Literature reports of malformed h g s pre-date the use of modern agricultural practices such as the widespread use of pesticides (see below). Literature reports also predate the beginning of the industrial revolution. One implication of this long history of reports of malformed frogs is that natural phenomena, or perhaps mild intervention by humans into natural processes, can cause amphibian malformations. Reports of Clalfonnations

Fig. 1. Reports of amphibian malformations by continent. Data are derived from Ouellet (2000). Note that reports of malformations are not uniformly distributed across the globe, but rather are strongly bimodal, predominating in Europe and North America. Used with permission of the Regents of the University of California and the University of California Press.

Ouellet (2000) noted several important trends in the literature on malformed frogs. The first is that historical reports of amphibian malfomations were primarily of frogs with multiple limbs or limb segments (polymely; this pattern may be due to collecting biases of human beings [see below]). Secondly, the number of individuals reported is strongly bimodal, with reports of either one malformed animal, or of > 10 malformed animals predominating (that is, there is a pattern of occurrence perhaps best described as nearly all or nearly nothing). Thirdly, and building on the second observation, historical accounts of malformed frogs consist mainly of single individuals while more recent accounts tend to describe large numbers of malformed individuals (see also Hoppe 2000, 2005). To illustrate this third point, the number of malformed frogs from Ouellet's (2000) data was plotted against the date of the publication reporting them (Fig. 2A). This graph shows that all sightings of large numbers of malformed frogs have been made since about 1950, suggesting that this problem is historically recent. A second way to examine Ouellet's

3092

AMPHIBIAN BIOLOGY

(2000) third conclusion is to code all reports of malformed frogs with fewer than 10 individuals as "0" and reports of > 10 individuals as "1" and plot them against publication date (Fig. 2B). One finds zeros scattered through time and continuing to the present. Beginning about 1950, however, " 1"s begin to appear. These " 1"s illustrate the beginning of the malformed-frog problem. Note that there is a long lag time between the earliest reports of multiple malformed frogs per site and 1995, the time when society became aware and alarmed, and the problem became a phenomenon. Global MalformationsIS~te

Global Severity Index

Fig. 2. Numbers of malformed frogs. A.) A plot of global data on numbers of malformed frogs per site against the date of the publication (from Ouellet 2000). This graph shows that all sightings of large numbers of malformed frogs have been made since the late 1940s, suggesting that this phenomenon is recent. B.) A plot of the same dataset shown in (A) with all reports of malformed frogs with fewer than 10 individuals coded as a "0" and reports of 10 or more individuals as a "l", graphed against publication date. Historically we find scattered "0"s that continue to the present day. Beginning in about 1950, "1"s appear and also continue to the present day. These reports of "1"s can be said to constitute the malformed frog phenomenon. Used with permission of the Regents of the University of California and the University of California Press.

111. THE NATURE OF THE DATA ON MALFORMATIONS

Historical data on malformed frogs are largely anecdotal. Field biologists collected and preserved curiosities, mainly multi-legged frogs, then wrote a short note in a natural history journal. These historical accounts make little mention of other types of malformed animals. One wonders whether other types of malformed frogs existed historically or whether collections were biased by observers' interests. Perhaps animals with missing legs, for example, stirred little curiosity. Hoppe (2005) concluded: "recent findings of anuran abnormalities in Minnesota. . . represent a new phenomenon. Frog abnormalities were more frequent, more varied, more severe, and more widely distributed in 1996-1999 than in 1958-1992 [Merrill 19691." Modern datasets are equally flawed. Malformed frogs are typically sampled by a biologist, or crew of biologists, at a wetland at about the time of year when frogs are

LANNOO: AMPHIBIAN MALFORMATIONS

3093

metamorphosing. The investigators walk or wade around the shallow margin and collect frogs individually with dip nets. Some astonishing numbers and percentages of malformed frogs can be collected in this way at the hottest of the hot spots, and these are the numbers one normally hears (e.g. "the CWB site has a malformation rate of 60%"). Does this technique represent a true (i.e. unbiased) survey? Some questions arise: (1) Would the numbers be the same if the wetland had been sampled at a different time, say a day or two earlier or later, or a week or two earlier or later? Meteyer et al. (2000) wrote: "At one of our study sites in Minnesota (CWB) frogs with polymelia were found in August of one year, but not until October of the second year." Similarly, Helgen et al. (2000) wrote: " . . . overall malformation frequencies o b s e ~ e dcan change dramatically in some sites, up or down, from July to late September or early October." Johnson et al. (2001b) noted: " . . . dramatic differences in the patterns of morphological abnormalities in amphibians between seasons, ponds, and among seasons and life history stages." Limited sampling times create sampling bias. (2) Do the animals being sampled truly represent the animals that were in the process of metamorphosing, or were numbers reduced by predators? If the latter is true, even though biologists may be sampling in a representative way, the animals remaining might not be representative, since - as Sessions and Ruth (1990) pointed out - the least mobile (i.e. most severely malformed) frogs would be most vulnerable to predation, and the dataset would be biased in favour of healthy frogs. (3) Do the animals being sampled truly represent all the animals that were in the process of metamorphosing or did the healthy animals move on, beyond the wetland fringe and into uplands, while the malformed frogs, being less mobile, stayed behind? If so, the data set would overestimate the proportion of malformed frogs. (4) Do malformed and normal frogs have an equal chance of being captured by biologists using dip nets? If malformed frogs are less mobile than normal frogs, they undoubtedly have a greater chance of being captured using one-on-one sampling techniques (in fact, Helgen et al. [1998] noted "malformed frogs . . . were much easier to capture than the normal young frogs"). This creates a bias towards collecting malformed frogs.

(5) Does every animal metamorphose? Schotthoefer et al. (2003) wrote: "R. ondatrae . . . infections acquired at the pre-limb bud stage negatively affected [Ram pipiens] tadpole survivorship. In addition . . . the proportions of tadpoles that died at this stage increased with the number of cercariae to which tadpoles were exposed." If some animals are so severely malformed that they cannot metamorphose, or if their development is so delayed that they metamorphose far later in the year than biologists would consider sampling, this creates a bias towards underestimating the proportion of malformations. (6) Animals with malformations that affect vital organs likely do not live past embryonic stages, or perhaps past larval stages that require feeding. These animals never get sampled and this creates a bias favouring normal frogs.

('7) Malformed animals do not appear to be as tolerant of stress as normal animals (for example they often do not survive the usually harmless experience of being put into moist containers in a cooler and transported back to a laboratory). They are therefore likely to be more vulnerable than are normal frogs to naturally stressful conditions, such as high water temperatures in the afternoon or low nocturnal concentrations of dissolved oxygen. Stressed frogs that succumb are never sampled and this creates a bias toward normal frogs. . Knowing that these potential biases exist allows a better understanding of data on malformed frogs. For example, when one collects 25 multi-legged animals from wetland X, about the only thing known for sure is that wetland X produced at least 25 multi-legged animals; in truth that's about all one can say. Wetland X may have produced 250 multilegged animals but the others never reached metamorphic stages, or reached metamorphosis but were eaten, or metamorphosed but on a day that was not sampled.

AMPHIBIAN BIOLOGY

3094

Wetland Y may also have produced frogs with missing legs but for the same reasons listed above, they were under-represented in samples. Similarly, when one reads that 60% of the animals at wetland X are malformed, what the data really mean is that 60% of the animals captured at that particular time on that particular day by those particular people using that particular technique were malformed. There has never been a field survey of malformed frogs that continuously sampled embryos, tadpoles and newly-metamorphosed individuals. This would be an enormously time-consuming task, yet it is the only true way to know what percentage of frogs at any particular site is malformed; every other method is prone to error. TV. TYPES OF MALFORMATIONS

In an attempt both to organize the data and to correlate effect with cause, malformations (whether human, frog, or otherwise) are usually placed into one of three categories: (1) structures absent or reduced, (2) structures present but otherwise abnormal (e.g., eye position, jaw shape, skin colour, pigment pattern), and (3) structures duplicated (or multiplied). Meteyer et al. (2000) provided definitions of malformed frogs, which are presented here in slightly modified form (Table 2).

Table 2. Classification and definitions of frog abnormalities, following Meteyer et al. (2000).

Type and Location of Abnormality Craniofacial Anophthalmia Brachygnathia Eye Discoloration Eye Displacement Microcephaly Microphthalrnia Forelimb and Hind limb Abnormal Pigment Amelia Taumelia Bony Expansions Brachydactyly

Curved Long Bone Ectrodactyly Ectromelia Hemimelia Micromelia Polydactyly Polymelia Skin Webbing Syndactyly Whole Body Bloated Body

Description missing eye abnormal shortness of lower jaw iris pigment discoloured or missing eye displaced laterally, medially, cranially, or caudally blunt nose; shortened upper jaw small eye pigment pattern missing or abnormal missing limb long bone bent back on itself forming > 90" angle distal end of a bone expands into spongy balloon normal number of metatarsals but abnormal number of phalanges all long bones bend at the site of artery penetration complete absence of digit including metatarsal bone missing limb segments (i.e.. femur present but rest of limb missing) shortened bone entire limb present but all limb elements shortened complete extra digit including metatarsal bone complete extra limb band of skin crossing a joint fusion of digits swelling in the torso and limbs of the animal

A. Missing Limbs

Missing limbs (ectromelia) are a common malformation found in field-collected animals. The northern leopard frog featured in figure 3A is not only missing its right hind limb but is also missing the ilium - the lateral pelvic element - on the same side ("agenesis" in the terminology of Meteyer et al. [2000]). This animal shows no signs of scarring, which is not surprising. A bite that removes an entire leg would also tear the large-bore femoral artery supplying blood to the leg, and likely result in the animal bleeding to death. The malformation shown in fi.wre 3A was not due to failed predation but instead must be due to developmental error. Rote that on the opposite siie of the malformation, the bony

3096

AMPHIBIAN BIOLOGY

Examples of malformed frogs as seen by X-ray (From Lannoo 2008). Used with permission of the Regents of the University of California and the University of California Press. Fig. 3a. Rana pipiens. 36 mm snout to vent (SVL). Collected on 11 September 1998 at the ROI site in Meeker County by Minnesota Pollution Control Agency Note that in this animal not onlv is the hind limb absent, but field biolo~ists. " the ilium - the lateral pelvic element - on the same side is also missing. Also note that on the opposite side of the malformation, the bony complex of the ilium, ischium, and femur is displaced laterally, towards the side of the missing limb, making the articulation with the intact leg positioned on the opposite side of the midline medial urostyle. These abnormalities of alignment are likely secondary effects, reflecting a remodelling of skeletal elements following use and the placing of the intact hind limb in a position that is more effective at singlelimbed propulsion.Fig. 3b. Rana pipiens. 38 mm SVL. Collected on 11 August 1997 from the OSP site by Minnesota Pollution Control Agency field biologists. Note that not only is this animal missing a forelimb, but it is also missing its entire shoulder girdle, strongly suggesting that this malformation represents a developmental problem. Fig. 3 c . Rana pipiens. 40 mm SVL. Collected 24 September 1997 at the SUN site, in Ottertail County, Minnesota, by Minnesota Pollution Control Agency field biologists. At first glance the femur of this northern leopard frog simply ends - an abrupt termination in the morphology of Meteyer et al. (2000) - and a morphology reminiscent of amputations. However, a close examination of the distal end of this bone reveals-a curvature not present in the femur of the intact contralateral side, suggesting that this malformation is not due to predation but instead has a developmental origin. Fig. 3d. Rana pipiem. 40 mm SVL. Collected on 5 August 1997 from the WIN site by Minnesota Pollution Control Agency field biologists. This micromelic animal also has a kinked femur in the micromelic limb and an extra bony element. Pelvic bones are disarticulated at the level of the junction of the hind limb junction. Scoliosis, associated with a shift in the position of the normal ilium, is also present. Fig. 3 e . Rana pipiens. 32 mm SVL. Collected by Dr. Daniel Sutherland from Trempealeau County, Wisconsin. According to Sutherland, these frogs did not have metacercariae of the trematode Ribeiroia ondatrae. This northern leopard frog exhibits a foreshortened limb with a small foot; the long bones of this limb exhibit taumelia (bony triangles). Meteyer et al. (2000) described this malformation type as phocomelia, or as "an abnormal foot attached to a short limb composed of small, disorganized, and unrecognizable bones.". Note also the opposite limb, which exhibits a foreshortening of the femur with a focal thickening. On this leg it is not possible to determine whether this abnormality is a developmental malformation or a healed broken bone. Fig. 3 f Rana pipiens. 32 mm SVL. Collected on 27 July 1998 at the CWB site in Crow Wing County, Minnesota by Minnesota Pollution Control Agency field biologists. This animal shows hemimely of the tibiofibula with an associated bony triangle (taumely). It is unclear whether this abnormal tibiofibula was duplicated or whether the tibia and fibula did not fuse and are thickened. Distal to this region, the tibiofibulare and foot bones of the foot appear roughly normal in size and proportion. Fig. 3g. Ram pipiem. 36 mm SVL. Collected 29 September 1997 at the NEY site in LeSueur County, Minnesota by Minnesota Pollution Control Agency field biologists. This animal presented with an immobile, straight leg, which I term (polyextension).The hip joint is also immobile and radiographs show a grossly abnormal pelvis.

LANNOO: AMPHIBIAN MALFORMATIONS

3097

Fig. 3h. Rana septentrionalis. 44 mm SVL. Collected on 2 July 1998 from the CWB site in Crow Wing County, Minnesota by Minnesota Pollution Control Agency field biologists. This animal demonstrates the failure of the skin at the knee, and perhaps the ankle and the hip joints, to separate. In addition, and perhaps secondary to this skin webbing, the long bones - femur, tibiofibula, and tibiofibulare - of the aeffected limb are foreshortened. Pelvic elements appear to have shifted, again in a way that favours forward thmst by the normal limb, producing an unusual juxtaposition of the femoral heads. Fig. 3 i . Rana catesbeiana. 44 mm SVL. Collected by Pieter Johnson fmm a site in California, where bullfrogs are nonnative and published by Lannoo et al. (2003). In these animals, all long bones, as well as the ilia, exhibit a kink. Kinks occur at the position where we one expects perforating arteries to penetrate. Bones that were not affected included the coccyx, vertebrae, scapulae, coracoids, and those composing the cranial skull. In the jaws, mandibles have no kinks but are not fully formed (the forward portions remain cartilaginouse). In addition to the skeletal problems, the tails of these animals are not fully resorbed. Fig. 3j. Rana septentrionalis. 38 mm SVL. Collected on 19 August 1997 from the NLA site by Minnesota Pollution Control Agency field biologists. This polymelic mink frog has exaa limb segments that appear at a point that is anatomically inappropriate. The primary limb o n the affected side is the same size as the contralateral limb and in life worked about as well. Pieter Johnson during the 199711998 Fig. 3k. Pseudacris regilla. 20 mm SVL. Collected from a California wetland field season. Note the patterned duplication of hind-limb segments. Grossly, this animal was described accurately as having a thick thigh, a duplication at the knee, and a duplication at one ankle, producing a small foot The radiograph shows that the duplication actually occurs at the hip - that the thick thigh is the xrsult of two femora that share a knee joint. The pelvis appears to support nvo acetabula, one for each femur, with the assist of a thickened ilium. The femur that articulates at the level of the contnlateral femur appears to be the least normal - it is short, kinked and appears to join the lower limb segment that divides at the ankle. Fig. 31. Rana pipiens. 41 mm SVL. Collected on 19 September 1995 at the NEY site in LeSueur County, Minnesota, by Minnesota Pollution Control Agency field biologists. This animal exhibits hemimely with taumely, and demonstrates limb multiplication; there are thrre hind limbs in this complex in addition to two bones near the pelvis - at least one of these is taumelic - which may represent additional femora. Fig. 3 m . Rana septentrionalis. 40 mm SVL. Collected by Dave Hoppe at the CWB site in central Minnesota (specimen # CWB7168012M). Dave Hoppe has taken good photos of this animal both alive and as a cleared and stained specimen. It exhibits four hind-limb fields that arise on one side of the pelvis. These limb fields produce one normal-sized hind limb and three smaller hind limbs. The smaller hind limbs are about equal in size. On both sides of the pelvis, the ilia appear thickened and abnormally shaped. There may be extra ilia1 elements present. Fig. 3 n . Rana catesbeiana. 26 mm SVL. Collected by Charles Facemire from a wetland in Switzerland County, Indiana. This animal is from a series of similar-appearing animals that exhibited bilaterally duplicated hind limbs in combination with subcutaneous swellings causes by accumulations of serous fluid (original description by Lannoo et al. [2003]). These swellings, which we called hygromas, extend from joint to joint - hip to knee, knee to ankle - and extend around each foot. Fig. 30. Rana pipiens. 45 mm SVL. Collected 5 August 1997 at the JOF site by Minnesota Pollution Control Agency biologists. This northern leopard frog has an incompletely formed upper jaw (maxilla) and superficially resembles, at least superficially, the cleft lips (frogs have no palates) seen in humans. Fig. 3 p . Rana pipiens. 52 mm SVL. Collected on 11 August 1997 at the LMS site in Meeker County by Minnesota Pollution Control Agency field biologists. This northern leopard frog exhibits bloating. In general, bloating can be due to build-up of either fluid or air. In this animal, it appears as though air (less dense) is responsible on the left side of the radiograph, fluid (more dense) on the right side.

complex of the ilium, ischium and femur are displaced laterally, towards the side of the missing limb, making the articulation with the intact leg positioned on the opposite side of the midline urostyle. The articulation between the ilium and the transverse processes of the sacrum (the sacroiliac joint) appears to be intact, so skeletal disarticulation, in the formal sense, did not occur. Note also the curvature (scoliosis) of the vertebral column towards the unaffected side. This combination of displaced pelvis and spinal curvature places the intact hind limb in a position that is more behind than beside the animal, where singlelimbed propulsion is more effective. Unlike the missing limb and ilium on the opposite side, these abnormalities of alignment are not the result of development gone wrong but instead are secondary effects, reflecting a remodelling of the position of the skeletal elements following use. Forelimb malformations, while not as common as those of the hind limb, are frequently found. For example, a population of Rana ornativentralis from japan was discovered that included individuals with supernumerary forelimbs (Takeishi 1996; Gardiner and Hoppe 1999). The northern leopard frog illustrated in figure 3B is not only missing a forelimb, but also its entire shoulder girdle, strongly suggesting that this malformation represents a

3098

AMPHIBIAN BIOLOGY

developmental problem. It is unknown why either hind limbs or forelimbs are selectively affected, but it is known that genes such as Hoxc-6 and Tbx-5 are differentially expressed in forelimbs, while Hoxc-10 and Tbx-4 are differentially expressed in hind limbs. Alterations to particular genes, therefore, could selectively affect either forelimbs or hind limbs (Gardiner and Hoppe 1999). B. Missing Limb Segments Missing limb segments, another form of ectromelia, are the most common malformation found in field-collected animals. Figure 3C demonstrates a northern leopard frog missing its right hind limb. At first glance, the femur of this frog just ends - an "abrupt termination" in the terminology of Meteyer et al. (2000) - a morphology reminiscent of amputations. A close examination of the distal end of this bone, however, reveals a curvature not present in the femur of the intact contralateral side. How can amputation of a leg at the knee produce a curvature in the femur above the knee? The morphology of this femur suggests a developmental origin for this malformation.

C. Micromely Micromely is a commonly observed malformation. A field examination of the northern leopard frog shown in figure 3D characterized this animal as having hind-limb micromely. Radiography revealed this diagnosis to be correct but incomplete; there are several additional problems in this interesting animal. Primary defects include a kinked femur in the micromelic limb and an extra bony element near the articulation of the affected limb with the pelvis. This extra bony element is either a femur or an ilium and is also kinked. Pelvic elements are disarticulated. In this animal, secondary effects include scoliosis and a shift in the position of the normal ilium towards the affected side; again, a morphology that favours forward thrust by the normal limb. D. Phocomely The northern leopard frog shown in figure 3E exhibits a foreshortened limb with a small foot; the long bones of this limb exhibit taumelia (bony triangles; see below). Meteyer et al. (2000) called this malformation phocomelia, defined as "an abnormal foot attached to a short limb composed of small, disorganized, and unrecognizable bones". The use of the term phocomelia immediately makes one think of the effects of thalidomide, which is not a fair association. Note also the opposite limb which exhibits a foreshortening of the femur with a focal thickening. It is not possible by looking at this radiograph to determine whether this abnormality is a developmental malformation or a healed broken bone. E. Taumelia Taumelia (bony triangles) are an interesting malformation type. As described by Gardiner and Hoppe (1999) bony triangles occur when a given limb segment is " . . . bent back upon itself, such that the proximal and distal ends of the bone are adjacent to one another and the midpoint of the shaft forms the apex of a triangle.. . T h e degree of angulation varies from bowing to complete folding". This definition varies from the one given by Meteyer et al. (2000) in which to be a taumelia a long bone must be bent back on itself forming an angle > 90". Gardiner and Hoppe (1999) also noted that despite being called "bony," these dysplasias are variably ossifiedlchondrified. The animal shown in figure 3F demonstrates hemimely of the tibiofibula with an associated taumely. It is unclear whether this abnormal tibiofibula was duplicated or whether the tibia and fibula did not fuse and are thickened. Distal to this region, the tibiofibulare and foot bones appear roughly normal in size and proportion.

E Polyextension Animals presenting with an immobile, straight leg (Fig. 3G) constitute a surprisingly common type of malformation. The key presentation is a knee locked in extension. The hip joint is also immobile and radiographs show that an abnormal pelvis may, (as in Fig.

LANNOO: AMPHIBIAN MALFORMATIONS

3099

3H) or may not, be present. Also, a disarticulation between the femur and the pelvic girdle may (as in Fig. 3G), or may not, be present. It is not known whether disarticulations are due to primary developmental effects, are secondary to having extended limb joints, or represent some combination of primary and secondary effects. The ankle joint of every animal expressing this malformation is also in extension (conhsingly called dorsiflexion, as opposed to plantar flexion or true flexion) as are foot and toe joints. This malformation also has been termed "hyperextension", a term that should not be confused with a condition in human sports injuries called by that name. With the exception of some disarticulated hip joints, no joint in these animals is literally hyperextended, although all joints in the affected limb are in extension. A better term is "polyextension".

A few other facts deserve mention. While limbs affected by polyextension present a characteristic appearance, the condition of the pelvis is variable and can appear normal. Measurements show that long bones tend to be shorter in straight limbs. Forelimb hyperextension is rare. For example, it was never seen in over 2,300 radiographs assembled over the past 10 years by the present author - perhaps because of the physical constraints developing limbs experience in the tadpole gill chamber. G. Skin Webbing Developing limbs achieve their proper length and proportion by a combination of addition and selective subtraction (Noden and de Lahunta 1985; Moore 1988). Skin-webbing is a common malformation that likely represents the subtraction process gone awry, although Gardiner and Hoppe (1999) noted: "At this point we do not know how skin webbings develop, and thus do not understand the developmental mechanisms involved. We also do not know if there is a functional relationship between abnormal skin development and abnormal skeletal development. . . "

The mink frog (Rana septentrionalis) shown in figure 3H is from the CWB site in central Minnesota and demonstrates the failure of the skin at the knee, and perhaps the ankle and the hip joints, to separate. In addition, and perhaps secondary to this skin-webbing, the long bones of the effected limb - femur, tibiofibula and tibiofibulare - are foreshortened. Pelvic elements appear to have shifted, producing an unusual juxtaposition of the femoral heads. Skin-webbing is associated with a variety of other malformations, including bony triangles (taumelia), polymelia, crooked limbs and pigment abnormalities. Skin-webbing is typically not found in limbs displaying missing elements, or with malformations in other body regions such as jaws or eyes. Skin webbing is found only rarely in forelimbs, which is surprising, given that forelimbs develop in the confined space of the gill chambers.

H. Arthrogryposis A population of bullfrogs (Rana catesbeianu) from California (where they were introduced) exhibits an unusual form of malformation (Fig. 31) (Lannoo et al. [2003]). In these animals, long bones exhibit a kink. Kinks occur at the position where one would expect perforating arteries to penetrate. Affected bones include the ilium and all limb bones - forelimbs and hind limbs, bilaterally. Bones not affected included the coccyx, vertebrae, scapulae, coracoids and those composing the cranial skull. In the jaws, the mandibles have no kinks but are not fully formed (the forward portions remain cartilaginous). In addition to these skeletal problems, the tails of these animals are not fully resorbed. All animals from this population exhibited these malformations; no malformation pattern like this has been seen in any other bullfrog population or in populations of any other amphibian species. These animals were infected by Ribeiroia ondantrae metacercariae, although the known mechanism (physical disruption) (Sessions and Ruth 1990; Sessions et al. 1999; Stopper et al. 2002) by which these parasites are thought to exert their effects cannot hold here. The chances of metacercariae of this trematode species occurring at exactly the midpoint of each long bone in the body, as well as at the midpoint of each ilium are

3100

AMPHIBIAN BIOLOGY

extremely low. Instead, some form of systemic effect must be occurring; the defect in each bone is at the point where its perforating artery penetrates. While the accepted term for "bent bones" is arthrogryposis, this morphology actually falls within the definition of bony triangles (or taumelia) as defined by Gardiner and Hoppe (1999) when they note: " . . . the degree of angulation varies from bowing to complete folding." I. Polymelia

Polymelia is the replication of limb segments. Polymelic animals serve as the iconic image of malformed frogs but, compared to ectromelic animals, they are rare. What is so different, and therefore so interesting, about the polymelic mink frog (R. septentrionalis) shown in figure 35 is that the extra limb appears at a point that is anatomically inappropriate. It is as if the femur has two knees, a normal one that forms a joint that is mobile and acts normally, and a second one about halfway down its length that does not appear to be a joint at all but supports a tibiofibula and its attached foot. The normal limb is the same size as the contralateral limb and in life worked about as well; the second limb is less robust and exhibits a shorter tibiofibula and smaller foot. The Pacific treefrog (Pseudacris regilla) collected in 199'711998 and shown in figure 3K, exhibits a patterned duplication of hind limb segments. Grossly, this animal was described accurately as having a thick thigh, duplication at the knee and duplication at one ankle, producing a supernumerary small foot. The radiograph shows that bone duplication actually occurs at the hip; the thick thigh is the result of two femurs that share a knee joint. The pelvis appears to support two acetabulums, one for each femur, with an associated thickened ilium. The femur that articulates at the level of the contralateral femur appears to be the least normal - it is short, kinked, and appears to join the lower limb segment that divides at the ankle. The second femur articulates immediately caudally; it is shorter than the contralateral femur but supports lower limb segments comparable in length and girth to the normal side. As unusual as this malformation type would seem to be, northern leopard frogs collected from several Minnesotan hotspots, as well as a wood frog (Rana sylvatica) from an Alaskan wildlife refuge (Reeves 2006) exhibit a similar pattern. The northern leopard frog shown in figure 3L also exhibits hemimely with taumely but, in addition, demonstrates limb multiplication; there are three hind limbs in this complex in addition to two bones near the pelvis - at least one of these is taumelic which may represent additional femurs. It is interesting that taumely may or may not be associated with duplicated limb elements, something that Gardiner and Hoppe (1999) noted only tangentially: "We have observed bony triangles at all levels along the proximal-distal axis of both primary and secondary limbs at equal frequency in each." Typically, the more complicated the malformation, the more complicated the description but the example of a polymelic frog in figure 3M is rather simple. This animal is relatively normal except that four hind limb fields arise on one side of the pelvis. These limb fields produce one normal-sized hind limb and three smaller ones. The smaller hindlimbs are about equal in size. On both sides of the pelvis, the ilia appear thickened and abnormally shaped. There may be extra ilia1 elements present. It would be interesting to examine the (likely diminished) nature of locomotory performance arising from the presence of multiple smaller hind limbs and ask whether they affect the survival of these animals. J. Multiple Limb Segments with Hygroma

Some of the most unusual malformations involve multiple effects. The bullfrog tadpole shown in figure 3N was from a population in Switzerland County, Indiana. This animal exhibited bilaterally duplicated hind limbs in combination with subcutaneous swellings caused by accumulations of serous fluid (Lannoo et al. 2003). These swellings extend from joint to joint - hip to knee, knee to ankle - and extend around each foot. Such abnormalities are called hygromas. Four animals from this population were all examined by the present author and all exhibited some form of hygroma; two animals, including the

LANNOO: AMPHIBIAN MALFORMATIONS

3101

one shown in figure 3N, had multiple hind limbs. The year following the collection of these animals, the original collector, Dr. Charles Facemire, returned to obtain additional animals. Not only did he not find any malformed frogs at this site, he found none at all. The population had become extirpated.

K. Maxillary Malformations The northern leopard frog shown in figure 3 0 has an incompletely formed upper jaw (maxilla).This malformation at least superficially, resembles cleft lips in humans (frogs have no palates). The JOF site was considered a marginal hotspot; no other animals exhibiting this malformation type were collected there.

L. Bloating The northern leopard frog shown in figure 3P exhibits bloating. Bloating can be due to build-up of either fluid or air. In this animal, it appears that air (less dense) is responsible on the left side of the radiograph and fluid (more dense) on the right side. It is possible that the build-up of air followed from accumulation of fluid. M. Abnormalities of Pigmentation

Disruptions of pigment on the proximal portions of limbs with missing or malformed distal segments offer powerful proof that these malformations are not due to failed attempts of predators. No instance is known of trauma inducing disruptions in pigmentation away from the site of the alleged attack. Over 70% of the ectromelic animals received following the 1997 and 1998 field seasons by the Minnesota Pollution Control Agency showed disruptions in pigmentation. These disruptions on the thighs of northern leopard frogs included absence of spotting, spots of different size (usually smaller) compared to the contralateral limb and spots of a different orientation. The ectromelic hind limb of the animal shown in figure 4 exhibits smaller, orthogonally-oriented spots, compared to the normal, contralateral hind limb.

Fig. 4. Rana pipiens. 50 mm SVL. Collected on 24 September 1997 at the SUN site in Ottertail County by Minnesota Pollution Control Agency field biologists. Note that the normal barred pattern along the dorsal hind limb is disrupted in the affected limb. In particular, when compared with the contralateral, normal limb, spots are smaller and oriented along the long axis of the limb. Used with permission of the Regents of the University of California and the University of California Press.

V. TAXONOMIC DISTRIBUTION

Amphibian malformations are not evenly distributed across taxa. As mentioned above, amphibian malformations tend to be mainly in frogs. A survey of amphibians from the United States showed that 51 of 104 United States' frog species (49%) had documented malformations compared with 19 of 187 salamander species (10%) (Lannoo 2008). The frog genera with the highest rates of documented malformations include Ascaphus (1 of 2 species; 50%), Bufo (10 of 23 species; 43%), Ac* (2 of 2 species; loo%), Hyh (7 of 10 species; 70%), Osteopilzu (1 of 1 species; loo%), Pseudarrzs (8 of 14 species; 57%), Gastrqhryne (2 of 2 species; loo%), Scaphiopus (2 of 3 species; 67%), Spea (1 of 4 species; 25%), Xenopus (1 of 1 species; 100%) and Rana (16 of 30 species; 53%). The salamander genera with the highest rates of documented malformations include Ambystoma (8 of 16 species; 50%, plus unisexual hybrids), Notophthalmus (1 of 3 species; 33%) and Taricha (2 of 3 species; 67%) (Lannoo, 2008).

3102

AMPHIBIAN BIOLOGY

The genera Dendrobates, Pternohyla, Smilisca, Eleutherodactylus, Leptodactylus, Hypopachus and Rhynophrynus from the United States had no documented malformations. Likewise, the salamander genera Amphiuma, Cyptobranchus, Dicamptodon, Aneides, Batrachoseps, Ensatina, Gyrinophilus, Hazdeotriton, Hemidactyium, Hydromantes, Phaeognathus, Pseudotriton, Stereochilus, Typhlotriton, Necturus, Rhyacotriton, Pseudobranchus and Siren had no documented malformations. VI. CAUSES OF MALFORMATIONS

In his treatise on the history of malformed amphibians, Ouellet (2000) listed the natural and anthropogenic causes of malformations, as follows. A. Natural Causes

(1) Wounding: Wounds from failed predation attempts ("bites and mutilations") and leech (Erpobdella octoculata) attacks can lead to either (a) missing limbs or parts of limbs, or (b) polymely through "hyper-regeneration following wounding."

(2) Fish Excrement: In the European frog Ram esculenta "anomalie P", consisting of bilateral brachymely, polymely and other types of limb malformations can be induced by rearing tadpoles in the presence of excrement from eels (Anguilla sp.) and/or minnows (Tinca sp.). Other amphibian species appear to be resistant to these influences. It is unknown whether the cause is chemical(s) contained in fish excrement or viruses transmitted through fish excrement. (3) High Densities of Tdpoles: Ram esculenta raised in high densities (3.5 to 11.1 tadpoles1 litre) exhibited forelimb ectromelia and hind limb hyperextension thought to be caused by teratogenic properties of chemical(s) released by the crowded tadpoles. Extremely high tadpole densities may also produce wounding (see above). (4) Lathyrogens: Extracts of sweet pea (Lathyrus odoratus) seeds have teratogenic effects on both salamander larvae and frog tadpoles. When young embryos are exposed to lathyrogens, notochord and tail malformations are produced; when older tadpoles are exposed dislocations of the joints and limbs occur.

(5) Nutritional Deficiencies: Malformations produced by nutritional deficiencies are often skeletal malformations, including decreased bone density, scoliosis, dislocations of joints, mandibular malformations and folding fractures of long (limb) bones. Paralysis may also occur. Nutritional deficiencies are most often seen in captive animals. Because most frog tadpoles are herbivorous and aquatic plants are abundant in healthy, established wetlands, it is difficult to envision food being limiting in these ecosystems. (6) Ultraviolet-B Radiation: The important work by Worrest and Kimeldorf (1975), Blaustein et al. (1994a,b, 1997) and Hays et al. (1996) demonstrates that UV-B can cause severe structural malformations in early-stage embryos of native amphibian species. In the laboratory, Ankley et al. (1998) and their colleagues at the EPA found UV-B effects that included bilaterally truncated hind limbs. The effects of UV-B acting either alone or in association with other factors are interesting and potentially important.

('7) Disease: Ouellet (2000) cited evidence for disease-producing, bilateral, posterior ectromelia and hemimely. The potential role of disease in producing malformations needs to be pursued more ambitiously. (8) Temperature Extremes: Amphibian malformations have been induced by artificially high temperatures (30°C) in the laboratory and by naturally low temperatures (cold spring water) in nature. It is unlikely that severe extremes in temperature are experienced by tadpoles in most natural wetlands. By contrast, however, surface pockets of hot water are present at the KEY pond in Minnesota, and several of the Minnesota wetlands sampled by Lannoo et al. (2003) were spring fed and noticeably cold (S°C below ambient air temperature).

LANNOO: AMPHIBIAN MALFORMATIONS

3103

(9) Hereditary Factors: Ouellet (2000) noted that some types of malformations are due to genetic mechanisms. As Ouellet acknowledged, however, similar genetic mutations can arise simultaneously across large geographic regions, o r arise simultaneously in several species from the same wetland, even when the same species in adjacent wetlands are normal. (10) Parasitic Cysts: Johnson et al. (1999, 2001a,b, 2002, 2007) and Johnson and Lunde (2005) reported a range of malformations induced experimentally by the trematode parasite Ribeiroia ondatrae and noted malformations associated with encysted Ribeiroia metacercariae in field-collected animals. Understanding the role of parasites in inducing malformations (Lannoo 2000; Sutherland 2005) is critical to understanding the recent apparent increase in malformations in frogs (see below). In addition to the natural causes of amphibian malformations, there are some that are induced by humans (Ouellet 2000).

B. Anthropogenic Causes (1) Acidijcation: Ouellet (2000) noted that skeletal malformations such as brachymely and hyperextension may occur when tadpoles are raised in acidified waters. In addition to these potential direct effects, there can be indirect effects in that p H levels influence the toxicity of some chemical contaminants. (2) Radioactive Pollution: Frog limb malformations have been associated with radioactive waste sites in the Netherlands and in Germany. Malformations have also been induced in the laboratory in tadpoles reared in rainwater contaminated with radioactive dust. (3) Ozone Depletion: Depletion of the ozone layer leads to increased atmospheric ultraviolet-B penetration and therefore to increased UV-B intensity at the Earth's surface (see above). ( 4 ) Heavy Metals: An assortment of abnormalities including eye, pigment and jaw malformations are associated with metal contamination from coal ash and coal-combustion wastes.

( 5 ) Retinoids: Several authors including Gardiner and Hoppe (1999) and Gardiner et al. (2003) have proposed that amphibian malformations are a consequence of developmental pathways regulated by retinoids (vitamin A and its derivatives). This topic is discussed in more detail below. ( 6 ) Agricultural Pesticides and Fertilizers: As Ouellet (2000) pointed out: "Agricultural herbicides, insecticides, fungicides, and fertilizers are often toxic to non-target organisms, and can cause deformities and mortality in amphibians." Exposure to agricultural chemicals produces many types of amphibian malformations, including hind limb brachymelia, ectromelia, ectrodactyly, polymelia and polydactyly, as well as scoliosis.

( 7 )Xenobiotics: Tadpoles of various species show several types of malformations including ectromelia, ectrodactyly and polydactyly when exposed to municipal and industrial waste. In addition, some animals develop tumors or tumor-like cysts. VII. IDENTIFYING CAUSES

While it has been generally accepted in Europe and Asia that there are multiple causes for malformed amphibians (Ouellet 2000), and while it is generally accepted that there are multiple causes for congenital anomalies in humans (23 general causes in a recently compiled list [Lannoo 2008]), the discussion in the United States about causes of malformed frogs has largely come down to two (Souder 2000): (1) disruption of retinoic acid pathways and (2) mechanical disruption of developing limb-bud tissue by metacercarial cysts of the trematode Ribeiroia ondatrae. Both hypotheses have supporting data and both hypotheses have serious problems as general explanations.

3104

AMPHIBIAN BIOLOGY

A. Retinoids While it is true that the effects of retinoic acid offer the only single explanation for the range of malformations observed in the field, field data in support of this hypothesis are lacking. While Gardiner et a l . (2003) showed the presence of biologically-active retinoids from the CWB site in Minnesota and from a site in Mission Viejo, California, there have been no published datasets showing a regional relationship between field-levels of retinoids, or chemicals demonstrated to affect retinoic-acid pathways, and malformations; nor has a dose/response curve been generated. In the absence of adequate field samples, the notions of parsimony or strong inference invoked by Gardiner and Hoppe (1999) and Gardiner et al. (2003), as logically appealing as they are, are insufficient. Gardiner and Hoppe (1999) realized this and concluded "frog malformations arise as a consequence of acute exposures - either a single exposure, or more likely a series of exposures to a teratogenic agent - rather than to a continuous, chronic exposure. Sampling protocols that attempt to identify the agent(s) responsible for these malformations need to accommodate this likelihood". B. Parasites

There can be no doubt that metacercariae of the trematode parasite Ribeiroia ondatrae cause malformations, but the statement " . . . the ability of cysts to induce the entire range [italics added] of deformities found in the wild, speak [sic.] to the great role that trematode cysts may be playing7' (Stopper et al. 2002) is certainly hyperbole. Examining a nearly identical dataset, Schotthoefer et al. (2003) wrote: "Notably absent from the malformations observed in R. pipiens in this study were missing or partially missing limbs . . . ". Several problems arise when attempting to generalize the current parasite hypothesis. For example, Ribeiroia ondatrae is the only parasite species known to elicit malformations Uohnson et al. 2003) and Ribeiroia metacercariae are not found in association with many malformations, especially in eastern North America and outside North America (see below). Workers examining malformed frogs have noted the presence or absence of Ribeiroia as follows: Eaton et al. (2004) working in Alberta and Saskatchewan, wrote: "A small number of the defomities we documented are consistent with descr$tim and images of trematode-induced deformities. However; we found no evidence to link Ribeiroia with these deformities." Helgen et al. (1998), working in Minnesota, wrote: "Most of the Le Sueur County malformed frogs had what appears to be parasitic cysts i n the thigh muscles. The frequency of parasitic cysts in normal frogs is unknown because these frogs were released at the site. Two large adults collected from the Le Sueur County site had heavy loads of cysts but appeared normal i n external and internal morphology. The Meeker County malformed frogs showed no visible appearance of parasite cysts in the legs." Gardiner and Hoppe (1999), also working in Minnesota, wrote: " . . . no trematode cysts were observed in association with either extra distal structures, hypomorphic limbs, or bony triangles." Meteyer et al. (2000), working both in Minnesota and Vermont, wrote: " I t should be emphasized again, however; that although polymelia was a predominant malformation i n some limited circumscribed studies, they [sic.] were infrequent in the wide geographic area of our study and Canada and metacercariae were not found i n the connective tissue of some of the malformed frogs." Converse et al. (2000), working on federal lands in the Midwest and New England, wrote: "the trematode theoy does not provule a n answer fm all abnomlities that have been observed." Ouellet et al. (1997), working in southern Quebec, wrote: "InJummatoy changes, parasites, and neoplastic alterations were not encountered in relation to l i ~ n bstructures." Taylor et al. (2005), working in Vermont, wrote: "Examination of a subsample of individual specimens revealed no evidence of Ribeiroia infection. Similarly, representative samples of host snails from the sample wetlands dul not demonstrate evidence of Ribeiroia infection."

LANNOO: AMPHIBIAN MALFORMATIONS

3105

Based on a report of Vermont samples sent to the USGS-BRD National Wildlife Health Center, Levey (2003) wrote: "No correlation has been found between incidence of abnormalities and parasite burden in newly-metamorphosed R. pipiem. Both normal and abnownal frogs can have heavy or light burdens of cysts." One notion that has been featured in recent literature is that Ribeiroia metacercarial cysts are not observed because they clear. Kiesecker (2002) wrote: " . . . tadpoles possess the ability to prevent the formation o j or to shed, metacercarial cysts . . . it will be important to determine whether metacercariae can be shed ajer they induce limb &fmmities. Such a pattern could explain the occurrence of deformities in wild collected fi-ogs that are not associated with metacercariae." Stopper et al. (2002) follow this up by stating: "In past studies, the role of cysts may have been underestimated because of the possibility that fiogs may be able to clear q s k (probably due to immunological rejection of the cysts). . . " It is difficult tb imagine that Ribeiroia cysts could create the sort of massive tissue disruptions described bv Stopper et al. (2002) and, in the case of American toads (Bufo americanus) and other species with short tadpole stages, leave absolutely no trace of their existence one or two weeks later. It also seems hard to believe that there is so much site-specific variation in the clearing process that the results of collections at a large number of sites over a restricted period of time (see Lannoo et al. 2003) are due to differential clearing of parasites, not to parasites being present or absent. Indeed, the comments by Kiesecker (2002) and Stopper et al. (2002) are in direct opposition to those of Johnson et al. (2004), who wrote: " . . . numerous [Ribeiroia] metacercariae may be recovered from mature frogs that are several years old, suggesting that amphibians may be less effective at eliminating Ribeiroia than are fish. These metacercariae are often brown in colour, however, possibly indicative of the colonization of eosinophils." When a group such as Eaton et al. (2004)writes - "One of the preserved specimens had no helminth parasites, whereas the other had two metacercarial cysts of the family Echinostomatidae . . .we are confident that no metacercariae were missed" it appears that no Ribeiroia were present in these animals. Here, dissent is taken to recent, high-profile publications and it is assumed that "Ribeiroia present" means that Ribeiroia is indeed present and that "Ribeiroia absent" means just that. It would seem to be the only scientifically defensible conclusion to draw. A second problem with the current parasite hypothesis is that the only demonstrated mechanism through which Ribeiroia metacercariae induce malformations is mechanical disruption of the developing limb bud (Sessions and Ruth 1990; Sessions et al. 1999; Stopper et al. 2002). For example, Sessions and Ruth (1990) noted:

"The results of the bead implantation experiment indicate that mechanical disruption of developing limb tissues by inert objects similar in size and shape to trematode cysts is sufficient to stimulate the outgrowth of supernumerary limb structures." More recently, Stopper et al. (2002)wrote: "These results indicate that the probable mechanism by which trematode cyst infestation causes limb deformities in frogs is perturbation of the spatial organization of cells in the developing limb buds followed by intercalation." One can add to mechanical perturbation the tendency for Riberioia to cluster in the inguinal region of tadpoles. Johnson et al. (2002) wrote: "Within infected anurans, the parasite exhibited a non-random distribution, with the majority of metacercariae embedded around the base of the limbs and the tail resorption area". Stopper et al. (2002) described how this pattern of localization happens, as follows: "We observed that the trematode cercariae are released from the snail and, once they contact a tadpole, most of them actively target the hind limb bud regions, especially preferring folds and indentations around the base of the limb buds and tail. Rarely, a cercaria was observed crawling into the mouth, spiracle, or cloaca. Multiple cercariae form cysts on the surface of the skin and penetrate into the tissues in and around the limb buds over the next few days."

3106

AMPHIBIAN BIOLOGY

Given (1) mechanical perturbation and (2) inguinal localization, one is forced to conclude that Ribeiroia infection does a poor job of explaining cranial and axial malformations, forelimb malformations, as well as distal hind limb malformations. In fact, the literature bears this out (see tables in papers by Johnson et al. [1999, 2001a1, Stopper et al. [2002] and Schotthoefer et al. [ZOO31 and summarized by Lannoo [2008]). A third problem for the current parasite hypothesis is that even when Ribeiroia are associated with malformations, the malformations that are produced do not match those found in the field. For example, Table 3 compares just the hind limb data (which is much more favourable to the parasite hypothesis than either the craniavaxial or forelimb data) from Pseudacris regzlla, Bufo boreas, and ranids (mostly Rana pipiens). From this table one can see that the relatively tight congruence in I! regillla between field samples and malformation types induced experimentally by exposure to Ribeiroia in the laboratory is notably absent in both B. boreas and ranids. In particular, both B. boreas and ranids produce extra limbs or limb Table 3. A comparison of the hind limb malformations induced experimentally by Ribeiroia and malformations found in field populations of Pseudacris regilla (from Johnson et al. 1999), Bufo boreas (from Johnson et al. 2001a) and Rana pipiens (from Lannoo, 2008). The data for Rana are taken from Schothoeffer et al. (2003; left column), Stopper et al. (2002; second column from left), Meteyer et al. (2000; center column), Converse et al. (2000; second column from right) and Levey (2003; right column). Note the interspecific variation in these data and how Bufo boreas and Rana are much more similar to each other than either is to Pseudacris regilla. Typestyles are used to emphasize great differences between experimental (bold) and field (italicized) data. Pseudacris reeilla

Malformation Type Hindlimb

Experimental (Light)

Experimental (Intermediate)

Experimental (Heavy)

Field

Experimental (Light)

Experimental (Intermediate)

Experimental (Heavy

Fleld

Missing limb Missing digit Cutaneous hsion Bony triangle Extra digit Extra limb Femoral projection Other

Malformation Type Hindhmb

Missing limb Missing foot Missing digit Cutaneous fusion Bony triangle Extra digit Extra foot Extra limb Femoral projection Other Rana sp.

Malformation Type Hindlimb Missing limb Missing foot Missinglshort digit Cutaneous fusion Bony triangle Extra digit Extra foot Extra limb Femoral projection Other

Experimental

Experimental

Field

Field

Field

0 0 0 8.3 0

0 0 0

5.4 34.0 19.2

4.9 31.0 33.5

12.7

? 8.8

16.7 3.9 2.0 2.5 4.9 0 6.4

0 0.8 0 0 0 0 3.2

0 0 0 1.8 0 0

0

0 0

22.2

44

5.6 33.3

0

25

?

1.8 ?

?

LANNOO: AMPHIBIAN MALFORMATIONS

31 07

elements, skin-webbing and bony triangles in response to laboratory exposure to Ribeiroia, while field collections of malformed frogs yield mostly missing limbs or limb segments. Johnson and his colleagues recognized these interspecific differences in Ribeiroia infection rates and interspecific differences in malformation types. For example, Johnson et al. (2001b) documented malformations in four species of amphibians (Pacific treefrogs, California newts [Taricha torosa],western toads [Bufo boreas] and bullfrogs) from the western United States and noted: "Among species, however, we documented substantial variation in abnormality composition", and on the same page indicated that "The polymelous western toad and bullfrog metamorphs are most likely the result of Ribeiroia infection, given the rarity of mutational events, the high frequency of polymelia caused by Ribeiroia in the laboratory, and the scarcity of other agents shown to cause anuran supernumerary limbs in the field. However, the missing limbs in these species and in the California newt larvae are less easily explained." Given that most data show amphibian malformations to be missing limbs or limb segments (e.g. Ouellet et al. 199'7; Helgen et al. 1998, 2000; Converse et al. 2000; Meteyer et al. 2000; Lannoo et al. 2003; Levey 2003), this conclusion would seem to be inconsistent, at least in emphasis if not in fact, with statements such as: "Limb malformations [constitute] an emerging parasitic disease in amphibians" (Johnson et al. 2003); [amphibian limb malformations constitute an] "emerging helminthiasis" [and suggest that] "the recent outbreak of deformities [might be due to] exogenous agents (e.g. pesticides, nutrient runoff, introduced fishes) ... interacting with Ribeiroiu, resulting in elevated infection levels ..." (Johnson and Sutherland 2003; see also Johnson et al. 2004); "If a spreading epidemic of Ribeiroia accounts for much, or even most, of the increase in h g deformities seen in recent years, what accounts for the epidemic?" (Blaustein and Johnson 2003). A fourth problem for the parasite hypothesis is that, just as the data have not tended to support phylogenetic generalizations (Table 3), the data also do not support geographic generalizations. The reports of malformations (Ouellet 2000) are shown here as figure 1 and all reports of Ribeioria, regardless of host Uohnson et al. 2003) are shown in figure 5. This literature was sorted according to continent (with the obvious exclusion of Antarctica) and while the scale of the literature in these two fields did not necessarily need to be similar (some fields attract much more attention and therefore generate many more papers than others), they, in fact, are. Ouellet (2000) found about 80 papers on malformed frogs from Europe while Johnson and his colleagues found around 80 on Ribeiroia from North America.

More important, though, is the pattern of papers within each discipline (Fig. 5). Reports of malformed frogs are strongly bimodal, with 78 papers from Europe and 68 papers from North America, but only 21 papers from AustraliaINew Zealand, 12 from Asia, 7 from South America and 4 from Africa. In contrast, reports of Ribeiroia infection are strongly unimodal, with 86 papers from North America, while only 5 papers are from South America, 4 papers from Africa, and 1 paper from Europe. At this point it may be tempting to explain away these differences (e.g. perhaps this pattern is due to a diminished interest in parasitology in Europe) but such scenarios do not hold up under close scrutiny. Instead, the data should be taken for what they are, however much one might hope otherwise, and at this point in our understanding accept that Ribeiroia are probably rarely found in Europe and that they cannot be the cause for the large number of reports of malformations found by Ouellet (2000). C. Pollution, Pesticides, and other Chemical Causes There is an enormous literature on the role that chemicals play in producing amphibian malformations-too large to fully review here-yet for some reason, in North America, this literature has been downplayed. Although Sparling et al.'s (2000) book on amphibian and reptilian ecotoxicology features three malformed frogs prominently on its cover, attention has not focused the Norh American literature. For example, the report on wounding from leech parasitism by Vieth and Viertel (1993), despite being written in German, is cited much

AMPHIBIAN BIOLOGY Reports of Malformations

I

90,

-

V)

c

.P m

80

-

706050-

'C

40-

g

30-

0

20-

0

Q

100 , North America

90 m C

.&' m

? 0

Q

South America

Europe

Africa

Asia

Australia1 New Zealand

Asia

Australia1 New Zealand

Reports of Ribeiroia

.

8070-

60504030 20400 , North America

South America

Europe

Africa

Fig. 5 . Reports of amphibian malformations (A) and Ribeiroia (B) by continent. Reports of Ribeiroia indicate its presence in both definitive and intermediate hosts. Data on malformations are taken from Ouellet (2000); data on Ribeiroia are from Johnson et al. (2004; see text for details). Note that while the reports of malformations are strongly bimodal, there are many fewer reports of Ribeiroia from outside North America, and only a single report from Europe, (infection of a Least Bittern, Zxobrychus minutus). Used with permission of the Regents of the University of California and the University of California Press.

more frequently than is the paper written in English by Alvarez et al. (1995) who demonstrated insecticide-induced skeletal malformations. Several workers are currently attempting to restore balance to the investigation of malformed frogs and perhaps currently overlooked literature will once again be featured. It is important to remember that Lowcock et al. (1997) found chromosomal abnormalities in malformed frogs from Quebec. It is also important to remember that, unlike the data on parasites of bufonids and ranids, water samples from affected wetlands produce the same sorts of malformations in the laboratory that were found in the field (Fort 1999a,b; Tietge et al. 2000). Burkhart et al. (1998) stated: "Initial experiments clearly showed that water from affected sites induced mortality and malformation in Xenopus embryos, while water from reference sites had little or no effect. Induction of malformations was dose dependent and highly reproducible . . . Limited evidence from these samples indicates that the causal factor(s) is not an infectious organism nor are ion concentrations or metals responsible for the effects observed. Results do indicate that the water matrix has a significant effect on the severity of toxicity." VIII. SOLUTIONS

Up to the present time, it is both a fair assessment to say, and a surprise for many to learn, that there has been no effective solution to the problem of malformed frogs, despite having spent millions of dollars of public and private h n d s and having published many

LANNOO: AMPHIBIAN MALFORMATIONS

3109

research papers. No single hotspot of malformed frogs has been restored to ecological health due to either direct or indirect action based on scientific enquiry. Why has science failed? It is perhaps because many scientists may have forgotten, or perhaps never knew, that science "dedicates itself above all to fruitful doing, not clever thinking, to claims that can be tested by actual research, not to exciting thoughts that inspire no activity" (Gould 1987). The goal of the inquiry into malformed frogs should be, above all, to fm the problem. Amphibian malformations have several causes and each hotspot should be carefully examined, the cause(s) determined and the source of the cause eliminated. This process is not only costly but time consuming and, given the current funding crises experienced by governments at all levels, it not likely to happen soon. Instead, it might be necessary to emphasize that hotspots tend to be altered wetlands (Lannoo et al. 2003). These alterations grade from what appears to be benign causes (simply having recently established and perhaps not having the buffering capacity of more mature systems), through simple eutrophication that leads to increased plant growth, increased snail populations and increased Ribeiroia levels (see Johnson and Lunde 2005), to serious chemical pollution (Cooke 1972, 1981; Rowe et al. 1996, 1998; Hopkins et al. 2000). The last category is of most concern. Recognizing that hotspots are altered wetlands, probably the quickest and least expensive way to reduce malformations is to ascertain the nature of those alterations and take steps to eliminate them. While the debate on malformations has tended to focus on proximate causes (for example retinoids versus parasites; see above), most runoff potentially contains both nutrients and toxic chemicals. In the case of one notable hotspot in Minnesota (CWB) (Lannoo et al. 2003), both problems are present and both are likely caused by a single factor: the use of the wetland by diary cattle for watering and cooling. Cattle urinate and defecate in the pond and the resulting increase in organic material causes eutrophication which in turn leads to increases in snail populations and the trematodes for which snails are an intermediary host Uohnson and Lunde 2005). On the other hand, retinoids enter the water as non-digested components of feed additives (Gardiner and Hoppe 1999). In either case, parasites or retinoids, the solution is not to minutely evaluate each cause and seek separate resolutions. Instead, the solution is to remove the cattle, thereby removing the source of both inputs, independent of whether causative inputs were nutrients or xenobiotics. Scientists interested in conservation biology must not lose sight of their responsibility. IX. ACKNOWLEDGMENTS

I thank Judy Helgen, David Hoppe and Pieter Johnson for sharing specimens with me. I also thank Joe Eastman for assistance with radiography, and Susan Lannoo and LaKina Curry for their assistance in preparation of the manuscript. This work was h n d e d by the Minnesota Pollution Control Agency, the United States Fish & Wildlife Service and Indiana University School of Medicine. This chapter summarizes a longer argument presented previously (Lannoo 2008) and I thank the Regents of the University of California and the University of California Press for permission to reproduce some of the figures. X. REFERENCES A l v a ~ z ,R., Honrubia, M. I? and Herriez, M. I?, 1995. Skeletal malformations induced by the insecticides zzaphoxm and folidolm during larval development of Rana perai. Arch. Env. Contam. Toxicol. 28: 349-356. Ankely, G. T, Tietge, J. E., DeFoe, D. L., Jensen, K. M., Holcombe, G. W., Durhan, E. J. and Diamond, S. A,, 1998. Effects of ultraviolet light and methovrene on survival and development of Rana fGfiiens: Env. Ex. Chem. 17: 2530-25k. Bechtel, H. B., 1995. "Reptile and Amphibian Variants: Colors, Patterns, and Scales". Krieger Publishing Company, Malabar, Florida.

Blaustein, A. R. and Johnson, I? T. J., 2003. Explaining frog deformities. Sci. Amer. February, pp. 60-65. Blaustein, A. R., Hoffman, I? D., Kiesecker, J . M. and Hays, J. B., 1994b. DNA repair activity and resistance to solar UV-B radiation in eggs of the red-legged frog. Cons. Biol. 10: 1398-1402. Blaustein, A. R., Kiesecker, J. M., Chivers, D. I? and Anthony, R. G., 1997. Ambient W - B radiation causes deformities in amphibian embryos. Proc. Nut. Acad. Sci. 94: 13735-13737.

3110

AMPHIBIAN BIOLOGY

Blaustein, A. R., Hoffman, F! D., Hokit, D. G., Kiesecker, J. M., Walls, S. C. and Hays, J. B., 1994a. UV repair and resistance to solar W - B in amphibian eggs: a link to population declines? Proc. Nut. Acad. Sci. 91: 1791-1795.

Hopkins, W. A., Congdon, J. and Ray, J. K., 2000. Incidence and impact of axial malformations in larval bullfrogs (Rana catesbeiana) developing in sites polluted by a coal-burning power plant. Env. E x . Chem. 19: 862-868.

Burkhart, J. G., Helgen, J. C., Fort, D. J., Gallagher, K., Bowers, D., Propst, T L., Gernes, M., Magner, J., Shelby, M. D. and Lucier, G., 1998. Induction of mortality and malformation in Xenopus laevis embryos by water sources associated with field frog deformities. Env. Health Perspec. 106: 841-848.

Hoppe, D. M., 2000. History of Minnesota -frog abnormalities: do recent findings represent a new phenomenon? Pp. 86-89 in "Investigating Amphibian Declines: Proceedings of the 1998 Midwest Declining Amphibians Conference", ed by H. Kaiser, G. S. Casper and N. Bernstein. Iowa Academy of Science 107, Cedar Falls.

Converse, K. A,, Mattsson, J. and Eaton-Poole, L., 2000. Field surveys of Midwestern and Northeastern Fish and Wildlife Service lands for the presence of abnormal frogs and toads. J. Iowa Acad. Sci. 107: 160-167. Cooke, A. S., 1972. The effects of DDT, dieldrin, and 2,4-D on amphibian spawn and tadpoles. Env. Pollut. 3: 51-68. Cooke, A. S., 1981. Tadpoles as indicators of harmful levels of pollution in the field. Env. Pollut. Sex A 25: 123-133. Eaton, B. R., Eaves, S., Stevens, C., Puchniak, A. and Paszkowski, C. A., 2004. Deformity levels in wild populations of the wood frog (Rana sylvatica) in three ecoregions of western Canada. J. Herpetol. 38: 283-287. Fort, D. J., Propst, T L., Stover, E. L., Helgen, J. C., Levey, R. B., Gallagher, K. and Burkhart, J. G., 1999a. Effects of pond water, sediment, and sediment extracts from Minnesota and Vermont, USA, on early development and metamorphosis of Xenopw. Enu. Tox. Chem. 18: 2305-2315. Fort, D. J., Rogers, R. L., Copley, H. F., Bruning, L. A., Stover, E. L., Helgen, J. C. and Burkhart, J. G., 1999b. Progress toward identifying causes of maldevelopment induced in Xenopw by pond water and sediment extracts from Minnesota, USA. Env. Tox. Chem. 18: 23162324. Gardiner, D. M. and Hoppe, D. M., 1999. Environmentally induced limb malformations in mink frogs (Rana septentrionalis).J. Exp. 2001. 284: 207-216. Gardiner, D., Ndayibagira, A., Griin, F. and Blumberg, B., 2003. Topic 4.6: Deformed frogs and environmental retinoids. Pure Appl. Chem. 75: 2263-2273. Gould, S. J., 1987. "An Urchin in the Storm". Norton, New York. Hays, J. B., Blaustein, A. R., Kiesecker, J. M., Hoffman, P. D., Pandelova, I., Coyle, D. and Richardson, T. 1996. Developmental responses of amphibians to solar and artificial UVB sources: a comparative study. Photochem. Photobiol. 64: 449-456. Helgen, J., McKinnell, R. G. and Gernes, M. C., 1998. Investigation of malformed northern leopard frogs in Minnesota. Pp. 288-297 in "Status and Conservation of Midwestern Amphibians", ed by M. J. Lannoo. University of Iowa Press, Iowa City. Helgen, J. C., Gernes, M. C., Kersten, S. M., Chirhart, J. W., Canfield, J. T., Bowers, D., Haferman, J., Mcknnell, R. G. and Hoppe, D. M., 2000. Field investigations of malformed frogs in Minnesota 1993-97. J. Zowa Acad. Sci. 107: 96-1 12.

Hoppe, D. M., 2005. Malformed frogs in Minnesota: history and interspecific differences. Pp. 103-108 in "Amphibian Declines: The Conservation Status of United States Species", ed by M. J. Lannoo. University of California Press, Berkeley. Johnson, P. T J. and Lunde, K. B., 2005. infection and limb malformations: a problem in amphibian conservation. Pp. in "Amphibian Declines: The Conservation United States Species", ed by M. J. University of California Press, Berkeley.

Parasite growing 124-138 Status of Lannoo.

Johnson, F! T J. and Sutherland, D. R., 2003. Amphibian deformities and Ribeiroia infection: an emerging helminthiasis. Trends Parasitol. 19: 332-335. Johnson, F! T J., Lunde, K. B., Ritchie, E. G. and Launer, A. E., 1999. The effect of trematode infection on amphibian limb development and survivorship. Sci. 284: 802-804. Johnson, F! T. J., Lunde, K. B., Zelmer, D. A. and Werner, J. K., 2003. Limb deformities as an emerging parasitic disease in amphibians: evidence from museum specimens and re-survey data. Cons. BioL 17: 1-14. Johnson, F! T. J., Sutherland, D. R., Kinsella, J. M. and Lunde, K. B., 2004. Review of the trematode genus Ribeiroia (Psilistomatidae): ecology, life history and pathogenesis with special emphasis on the amphibian malformation problem. Adv. Parasitol. 57: 191-253. Johnson, F! T J., Lunde, K. B., Haight, R. W., Bowerman, J. and Blaustein, A. R., 2001a. Ribeiroia ondatrae (Trematoda: Digena) infection induces severe limb malformations in western toads (Bufo boreas). Can. J. 2001. 79: 370-379. Johnson, F! T J., Lunde, K. B., Ritchie, E. G., Reaser, J. K. and Launer, A. E., 2001b. Morphological abnormality patterns in a California amphibian community. Herpetologica 57: 336-352. Johnson, I? T J., Lunde, K. B., Thurman, E. M., Ritchie, E. G., Wray, S. N., Sutherland, D. R., Kapfer, J. M., Frest, T J., Bowerman, J . and Blaustein, A. R., 2002. Parasite (Ribeiroia ondatrae) infection linked to amphibian malformations in the western United States. Ecol. Monogx 72: 151-168. Johnson, F! T J., Chase, J. M., Dosch, K. L., Hartson, R. B., Gross, J. A., Larson, D. J., Sutherland, D. R. and Carpenter, S. R., 2007. Aquatic eutrophication promotes pathogenic infection in amphibians. Proc. Nut. Acad. Sci. 104: 15781-15786. Kiesecker, J., 2002. Synergism between trematode infection and pesticide exposure: a link to amphibian limb deformities in nature? Proc. Nat. Acad. Sci. 99: 9900-9904.

LANNOO: AMPHIBIAN MALFORMATIONS Lannoo, M. J., 2000. Conclusions drawn from the malformity and disease session, Midwest Declining Amphibians Conference, 1998. Pp. 2 12-2 16 in "Investigating Amphibian Declines: Proceedings of the 1998 Midwest Declining Amphibians Conference", ed by H. Kaiser, G. S. Casper and N. Bernstein. Iowa Academy of Science 107, Cedar Falls. Lannoo, M. J., 2008. "Malformed Frogs". University of California Press, Berkeley. Lannoo, M. J., Sutherland, D. R., Jones, E, Rosenberry, D., Klaver, R. W., Hoppe, D. M., Johnson, F! T. J., Lunde, K. B., Facemire, C. and Kapfer, J. M., 2003. Multiple causes for the malformed frog phenomenon. Pp. 233-262 in "Multiple Stressor Effects in Relation to Declining Amphibian Populations", ed by G. Linder, S. Krest, D. Sparling and E. Little. American Society for Testing Materials International, West Conshoshocken, Pennsylvania. Levey, R., 2003. Investigations into the causes of amphibian malformations in the Lake Champlain Basin of New England. Final Report to the Vermont Department of Environmental Conservation, Waterbury, Vermont. Lowcock, L. A., Sharbel, T F., Bonin, J., Ouellet, M., Rodrigue, J. and Des Granges, J. -L., 1997. Flow cytometric assay for in vivo genotoxic effects of pesticides in green frogs (Ram clamitam).Aquat. Ex. 38: 241-255. Merrell, D. J., 1969. Natural selection in a leopard frog population. J. Minnesota Acad. Sci. 35: €689. Meteyer, C. A., Loefller, K. I., Fallon, J. F., Converse, K. A., Green, E., Helgen, J. C., Kersten, S., Levey, R., Eaton-Poole, L. and Burkhart, J. G., 2000. Hind limb malformations in free-living northern leopard frogs (Rana pipiens) from Maine, Minnesota, and Vermont suggest multiple etiologies. Eratology 62: 151-171. Moore, K. L., 1988. "The Developing Human". Saunders, Philadelphia. Noden, D. M. and de Lahunta, A., 1985. "The Embryology of Domestic Animals: Developmental Mechanisms and Malformations". Williams and Wilkins, Baltimore. Ouellet, M., 2000. Amphibian deformities: current state of knowledge. Pp. 617-661 in "Ecotoxicology of Amphibians and Reptiles", ed by D. W. Sparling, G. Linder and C. A. Bishop. Society for Environmental Toxicology and Contaminants (SETAC) Press, Pensacola, Florida. Ouellet, M., Bonin, J., Rodrigue, J., DesGranges, J. -L. and Lair, S., 1997. Hindlimb deformities (ectromelia, ectrodactyly) in free-living anurans from agricultural habitats.J. Wldl. Lhs. 33: 95-104. Reeves, M. K., 2006. Alaska's abnormal frogs. U.S. Fish and Wildlife Service (no number), published in April, 2006 and available at http:l/www.fws.gov.

3111

Rowe, C. L., Kinney, 0. M. and Congdon, J. D., 1998. Oral deformities in tadpoles of the bullfrog (Rana catesbeiana) caused by conditions in a polluted habitat. Copeia 1998: 244-246. Rowe, C. L., Kinney, 0 . M., Fiori, A. I! and Congdon, J . D., 1996. Oral deformities in tadpoles (Rana catesbeiana) associated with coal ash deposition: effects on grazing ability and growth. Freshwater Biol. 36: 723-730. Schotthoefer, A. M., Koehler, A. V, Meteyer, C. A. and Cole, R. A., 2003. Influence of Ribeiroia ondatrae (Trematoda: Digenea) infection on limb development and survival of northern leopard frogs (Rana pipiens): effects of host stage and parasite-exposure level. Can. J. Zool. 81: 1144-1153. Sessions, S. K. and Ruth, S. B., 1990. Explanation for naturally occurring supernumerary limbs in amphibians. J. Exp. 2001. 254: 38-47. Sessions, S. K., Franssen, R. A. and Homer, B L., 1999. Morphological clues from multilegged frogs: are retinoids to blame? Sci. 284: 800-802. Souder, W., 2000. "A Plague of Frogs: The Homfying Tme Story". Hyperion Press, New York. Sparling, D. IV., Linder, G . and Bishop, C. A. (eds), 2000. "Ecotoxicology of Amphibians and Reptiles". Society for Environmental Toxicology and Contaminants (SETAC) Press, Pensacola, Florida. Stopper, G. F., Hacker, L., Franssen, R. A. and Sessions, S. K., 2002. How trematodes cause limb deformities in amphibians. J. Exp. Zool. Part B: Mol. Dev. Evol. 294: 252-263. Sutherland, D., 2005. Parasites of North American frogs. Pp. 109-123 in "Amphibian Declines: The Conservation Status of United States Species", ed by M. J. Lannoo. University of California Press, Berkeley. Takeishi, M., 1996. On the frog, Rana ornatiuentralis, with supernumerary limbs found at Yamada Greenery are in Kitakyushu City, Fukuoka Prefecture. Japan Bull. Kitakyushu Mus. Nut. Hist. 15: 119-13 1. Taylor, B., Skelly, D., Demarchis, L. K., Slade, M. D., Galusha, D. and Rabinowitz, P M., 2005. Proximity to pollution sources and risk of amphibian limb malformation. Enu. Health Perspect. 113: 1497-1 50 1. Tietge, J. E., Ankle); G. T., DeFoe, D. L., Holcombe, G. \V. and Jensen, K. M., 2000. Effects of water quality on development of Xenopw laeuis: a frog embryo teratogenesis assay-Xenopw assessment of surface water associated with malformations in native anurans. Enu. Tox. Chem. 19: 2 114-2 121. Veith, M. and Viertel, B., 1993. Veranderungen an den extremitaten von larven und jungtieren der erdkrote (Bufo bufo): analyse moglicher ursachen. Salamandra 29: 184-199. Worrest, R. C. and ameldorf, D. J., 1975. Photoreactivation of potentially lethal, UV-induced damage to boreal toad (Bufo boreas boreas) tadpoles. Lije Sci. 17: 1545-1550.

CHAPTER 6

Ultraviolet-B Radiation and Amphibians Adolfo Marco, Betsy A. Bancroft, Miguel Lizana and Andrew R. Blaustein

I. Introduction II. UV Radiation at Amphibian Breeding Sites A. Natural Variability B. Stratospheric Ozone Depletion C. Acidification D. Deforestation and Destruction of Riparian and Aquatic Vegetation E. Global Climatic Change Ill. Effects of UVB Radiation on Amphibian Eggs, Larvae and Adults IV. Ecological and Behavioural Photoprotective Strategies A. Nocturnality B. Viviparity and other Egg-brooding Systems C. Terrestrial Oviposition D. Aquatic Refuges E. Deep Water and Dissolved Organic Content (DOC) Concentration

V. Molecular and Morphological Adaptations A. Enzymatic Repair Systems B. Melanin and other Sunscreens C. Gelatinous Matrices and Foam Nests D. Other Photoprotective Systems VI. Synergism of UV Radiation with other Factors A. UV and Acidification B. UV and Contaminants C. UVB and Pathogens VII. General Conclusions A. Factors Affecting Exposure to Ultraviolet Radiation B. Natural Selection C. Interspecific Differences in Resistance to UVB Radiation as Depicted by the Results of Field Experiments D. Measuring UVB in the Field E. The Role of UVB Radiation in Amphibian Population Declines VIII. References

Abbreviations used: CDOM = chromophoric dissolved organic matter; CPD = cyclobutane pyrimidine dimer; DOC = dissolved organic content; DOM = dissolved organic matter; GIs = geographic information systems; HSP = heat shock protein; PAH = polycyclic aromatic hydrocarbon; UV = ultraviolet; UVA = ultraviolet-A; UVB = ultraviolet-B

u

I. INTRODUCTION

LTRAVIOLET-B radiation (hereafter, UVB) has a wavelength of 280-315 nm. It has been a ubiquitous stressor on living organisms since life began (Cockell 2001; Blaustein and Kiesecker 2002). Moreover, all living organisms have probably been subjected to periodic increases in UVB due to natural events (Cockell and Blaustein 2000). Unprecedented increases in UVB radiation, originating from anthropogenic sources, have also been detected since the 1970s (Madronich et al. 1998). Many amphibians are susceptible to UVB radiation but the degree of sensitivity depends upon the species, the life stage and the ecological context.

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3113

Thus, embryos of some amphibian species are sensitive to current ambient levels of UVB radiation (reviewed by Blaustein et al. 2001a) whereas those of other species are more tolerant to current levels. If exposed to UVB radiation as embryos, however, almost all species show sublethal effects in later life (Table 1). As for any toxic agent, there is a doseeffect relationship and a limit level of tolerance of amphibians to UVB. Thus, many amphibians may be resistant to ambient levels of UVB in certain life history stages but suffer detrimental effects in other stages. If the increase in UVB is systematic and occurs over the entire distributional range of a given species, whole populations or even the entire species may ultimately be affected. There are, however, molecular, behavioural and morphological characteristics that make amphibians less vulnerable to UVB radiation (Blaustein and Belden 2003).

Table 1 . Some examples of the effects of ultraviolet-B radiation on amphibians.

Species Ambystoma gracile Ambystoma laterale Ambystoma macrodactylum

Effects of ambient levels of UV

Synergistic effects with UV

Deformities; Larval mortality Embryonic mortality; Deformities; Larval mortality; Reduced growth Embryonic mortality; Reduced growth

Bufo boreas

Embryonic mortality; Behavioural changes

Bufo bufo

Embryonic mortality; Larval mortality

Crinia signqera

Embryonic mortality; Larval mortality Skin darkening Embryonic mortality

With nitrate, reduced growth

Blaustein et al. (1997); Belden et al. (2000); Belden and Blaustein (2002a, 2002~);Hatch and Blaustein (2003) Lesser et al. (2001); Crump et al. (1999a) Crump et al. (1999a); Grant and Licht (1995)

With Saprolegnia, increased embryonic mortality

Blaustein et al. (1994b); Eesecker and Blaustein (1995); Kats et al. (2000) Lizana and Pedraza (1998); Hakkinen et al. (2001) Broomhall et al. (2000)

Deformities; Larval mortality Larval mortality; Juvenile mortality; Deformities

Langhelle et al. (1999) Anzalone et al. (1998)

Avoid UV-B as adults but not as tadpoles Do not avoid UV-B as adults or as tadpoles

Ovaska et al. (1997); Hatch and Blaustein (2003) Crump et al. (1999a); Grant and Licht (1995); Zaga et al. (1998) van de Mortel and Buttemer (1998) van de Mortel and Buttemer (1998)

Avoid UV-B as adults but not as tadpoles

van de Mortel and Buttemer (1998)

Hyla regilla

Larval mortality

With nitrate, reduced survival

Hyla uersicolor

Larval mortality

With carbaryl, increased embryonic and larval mortality

Litoria aurea Litoria dentata

Lioria peronii

Litoria uerreauxii Embryonic mortality; alpina Larval mortality Pseudacris crucifer Larval mortality Rana arualis

Reference Blaustein et al. (1995); Belden and Blaustein (2002~) Crump et al. (1999a)

Embryonic mortality; Skin darkening; Reduced growth

Ambystoma maculatum Bufo americanus

Hyla arborea Hyla cadauerina

Effects of enhanced levels of UV

Embryonic mortality

Broomhall et al. (2000) With copper, reduced larval survival

Baud and Beck (2005) Hakkinen et al. (200 1)

3114 Table 1

AMPHIBIAN BIOLOGY - continued

S~ecies

Effects of ambient levels of UV

Effects of enhanced levels of W

Synergistic effects with UV

Rana aurora

Slows growth

Embryonic mortality; Larval mortality

Rana blairi

Slows growth

Rana cmcadae

Embryonic mortality; Behavioural changes; Tadpoles do not avoid W-B; Larval mortality

Ovaska et al. (1997); Belden and Blaustein (2002b) With landfill leachate, Smith et al. (2000); W - B increases growth Bruner et al. (2002) and decreases deformities With Saprolegnia, reduced embryonic Blaustein et al. survival; With nitrate (199413); Belden et al. and low pH, reduced (2003); Kats et al. (2000); Kiesecker and larval survival and Blaustein (1995); Hatch behavioural changes and Blaustein (2000) Walker et al. (1998) With fluoranthene, skin damage and increased activity Tietge et al. (2001); Crump et al. (1999a); Grant and Licht (1995) Tietge et al. (2001); With low pH, Ankley et al. (2000); increased embryonic Ankley et al. (2002); mortality Long et al. (1995) Tietge et al. (2001) Bridges and Boone (2003) Crump et al. (1999a) Grant and Licht (1995)

Rana catesbeiana

Rana clamitans

Larval mortality

Rana pipiens

Larval mortality; Deformities

Larval mortality; Juvenile mortality; Deformities

Rana septentrionalis Larval mortality Rana sphenocephala Increased larval survival Rana syluatica

Rana t e m p o r a ~ a

Taricha granulosa

Taricha torosa Triturus alpestris Triturw cristatus

Embryonic mortality; Larval mortality; Juvenile mortality; Deformities Increased frequency Slows growth; Later metamorphosis; of abnormalities; Delayed metamorSmaller size at phosis; Smaller size Metamorphosis; at metamorphosis Increased survival to metamorphosis Behavioural changes; Skin darkening

With low pH, reduced Pahkala et al. (2000); survival and increased Pahkala et al. (2001a); Pahkala et al. (2002a); frequency of abnormalities in some Pahkala et al. (2002b); Pahkala et al. (2003) populations Blaustein et al. (2000); Belden and Blaustein (20024 Anzalone et al. (1998) Nag1 and Hofer (1997)

Embryonic mortality Larval mortality; Behavioural changes Embryonic mortality; Larval mortality

Langhelle et al. (1999) With carbaryl, increased embryonic and larval mortality

Xenopus laevis

Reference

Zaga et al. (1998)

11. W RADIATION AT AMPHIBIAN BREEDING SITES A. Natural Variability The impact of UVB radiation on freshwater ecosystems is a result of complex interactions among a variety of factors (Williamson 1995). Estimates of UVB "doses" (exposure over time), however, that actually impinge upon amphibians are largely unavailable. Moreover, uncertainties concerning measurements of UVB exposure of amphibians are complicated for environmental and technical reasons (Diamond et al. 2002). One major problem in assessing UVB exposure is that there is both temporal and spatial

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBLANS

31 15

variation in UVB (Blaustein et al. 2004). Environmental variation is also important in the field. Because UVB irradiance is highly variable within short distances or time periods, the use of satellite data or empirically derived models should be used with caution (Madronich et al. 1998; Middleton et al. 2001). In an attempt to include local variation of UVB into estimates of doses in amphibian habitats, a recent study incorporated ground-based UVB measurements, solar radiation models, geographic information systems (GIs) tools, vegetative features and dissolved organic compound concentrations (DOC) into estimates of UVB in natural wetlands (Diamond et al. 2005). Adams and colleagues (2005) followed the methods developed by Diamond et al. (2005) to examine the relationship between the distribution of eight amphibian species and UVB dose. This study found quadratic relationships between estimated UVB dose and the distribution of two salamander species (Taricha granulosa and Ambystoma macrodactlyum). No effect of UVB dose on distribution of the six other species was detected. Although these studies represent an advance in estimating UVB doses in amphibian habitats, the authors were still unable to incorporate either temporal and spatial variation in UVB irradiance or temporal changes in DOC into their estimates. Accurately estimating dose at local sites requires intensive ground measurements that are impractical at large spatial scales. Unfortunately, understanding the temporal and spatial variation in UVB at local scales is necessary for a full appreciation of the effects of UVB on amphibian distribution. Under certain conditions, UVB irradiance may increase with elevation. For example, in Central Europe under clear skies, erythema1 effective irradiance during the summer increases an average of 18% every 1 000 m of elevation (Blumthaler et al. 199'7).Amphibians from montane populations not only develop at higher elevations than do those at lower elevations but they also develop later in the season. In certain regions, such as in the Pacific Northwest of the United States, there is greater development of cloud cover at that time (Belden and Blaustein 2002a). Geographic and seasonal variation in UVB irradiance may influence the sensitivity to UVB in certain species (Belden and Blaustein 2002a). For example, Ambystoma macrodactylurn larvae from three lowland populations that were exposed to ambient levels of UVB radiation had higher mortality rates than did those from five high elevation populations (Belden et al. 2002a). Broomhall et al. (2000) found that egg and larval mortality due to UVB exposure increased with elevation in three anurans from Australia. Latitude also has a significant effect on UVB irradiance; levels increase from the poles toward the Equator and is maximal in spring and summer in both hemispheres. The differences between the northern and southern hemispheres in ground measurements of summer UVB irradiances at mid-latitudes are about 40%. By contrast, satellite-based estimates show differences of only about 10-15%. Air pollution in the Northern Hemisphere may influence satellite-based estimates and could be one reason for the discrepancy between ground and satellite estimates of UVB irradiances (Madronich et al. 1998). Nevertheless, under certain conditions and disregarding elevation, amphibians breed later and are exposed to higher UVB levels at higher latitudes than at lower ones (Merila et al. 2000). The prediction of a gradient in amphibian declines from the poles toward the equator (Davidson ct al. 2001, 2002) would not be applicable to species for which the phenology of their sensitive stages varies against the latitudinal gradient. Moreover, interactions between elevation and latitude should be considered. The highest levels of UVB irradiance are probably found in high mountains located in the lower latitudes. Cloudiness may reduce the incidence of UVB (Blumthaler et al. 1994). The risk of o\-erexposure to UVB may be still significant, however, when the sun is obscured by clouds (Diffey 1991). Type of cloud influences the amount of UVB that is transmitted. For example, low-level stratus clouds can reduce of UVB transmittance by about 60-75% while higherlevel cirrus clouds only reduce UVB slightly as compared to clear skies (Xenopoulos and Schindler 200 1).

3116

AMPHIBIAN BIOLOGY

Various aerosols, aerial pollutants or gases from volcanic eruptions may absorb UVB and reduce its level at the earth's surface (Xenopoulos and Schindler 2001). It is difficult, however, to predict how these agents influence irradiance because of temporal and spatial variability as well as variability in the types of aerosols and how they are distributed at specific sites. Surface reflection may also vary considerably among different types of substrates (Diffey 1991). B. Stratospheric Ozone Depletion

Several studies have reported an increase in UVB at the earth's surface during the past three decades (Herman et al. 1996; Middleton et al. 2001). The main cause of this increase appears to be anthropogenic alteration of the stratospheric ozone layer (Kerr and McElroy 1993). Ground-based measurements and those from balloons and satellites have all shown a significant decrease in the amount of total stratospheric ozone over mid-latitude areas of the Northern Hemisphere in all seasons (Randel et al. 1999). As a consequence of this depletion over the past three decades the levels of UVB that reach the earth have significantly increased both in tropical and temperate regions, and in both hemispheres (Herman et al. 1996; Middleton et al. 2001). This increase in the North Temperate zone was about 0.5% per year (Blumthaler and Ambach 1990). Madronich et al. (1998) estimated the increase in UVB reaching the earth's surface since the 19'70s to be about 7% in winter/ spring and 4% in summer/fall at mid-latitudes in the Northern Hemisphere and about 6% at mid-latitudes on a year-round basis in the Southern Hemisphere. Current estimates suggest that the ozone layer will continue to be vulnerable during this century and the recovery phase could be delayed by decades (Madronich et al. 1998). The increase in UV is wavelength-specific with the greatest change at the shorter wavelengths, the very part of the spectrum causing significant molecular damage to living organisms. Photo-repair has not been enhanced by these changes because it is mediated by radiation of longer wave-lengths that did not undergo a concomitant increase (Williamson 1995; Bruggeman et al. 1998). C. Acidification

Acid deposition affects aquatic ecosystems in a number of ways and can lead to greater penetration of UVB radiation into the water column (Wright and Schindler 1995; Donahue et al. 1998; Hader et al. 1998). The main reason for increased penetration is most likely the interaction of acidification with dissolved organic compounds (DOC) (Yan et al. 1996). In an experimentally acidified lake the DOC declined by 80%, thereby increasing UV penetration by 900% (Schindler et al. 1996). D. Deforestation and Destruction of Riparian and Aquatic Vegetation Aquatic vegetation serves as a refuge for the eggs and larvae of many amphibian species (Duellman and Trueb 1994; Stebbins and Cohen 1995). Many pond-breeding amphibians attach their eggs to submerged vegetation and larvae often develop within algae-covered, submerged plant-mats. This association between early life stages and vegetation may help protect amphibians from UVB. Under forest canopies the levels of UVB are lower than in places where the vegetation has been removed or destroyed (Kelly et al. 2003). Riparian shading may thus moderate the negative effects of UVB on freshwater communities. Alteration of the riparian canopy by land-use activities, such as logging and urban development, can result in large increases of exposure of amphibians to sunlight. Any activity that negatively influences the growth of freshwater aquatic vegetation, such as excessive herbivory, may also significantly increase exposure of amphibian eggs and larvae to damaging levels of UVB. An increase in the incidence of wildfire may lead to destruction of vegetation that protects both terrestrial and aquatic amphibians from sunlight (Anzalone et al. 1998). Wildfires have affected natural ecosystems over historical time and there has been an increase in the frequency of intense

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3117

wildfires in modern times (France et al. 2000). Eventually, lakes within clear-cut or burnt watersheds may have higher DOC than do undisturbed catchments. The increased incidence of wildfires coincident with warming climates may partially offset the effects of droughts on catchment export of DOC in the short term and mitigate the harmful effects of damaging UVB in aquatic ecosystems (France et al. 2000). E. Global Climatic Change

Global warming and associated climatic changes may increase mortality in amphibian populations for a number of reasons (Pounds 2001; Pounds et al. 2006). For example, global warming can have a significant impact on dissolved organic matter and its residence time in bodies of water (Hader et al. 1998). This interaction could cause a decrease in the transparency of water to UVB (Schindler et al. 1996; Pienitz and Vincent 2000). Climatic change, however, may also cause a migration of the tree-line that in turn can increase shading and raise levels of DOM (dissolved organic matter) in freshwater ecosystems, thereby reducing the penetration of UVB (Williamson et al. 1996, 2001). A warming climate may also favour plant growth in the lakes and ponds of high mountains (Walther et al. 2002), thus protecting amphibians from UVB. Moreover, amphibians could breed earlier in the season due to increased vernal temperatures and earlier snowmelt in mountains. Earlier breeding may favour embryonic development during periods of lower UVB irradiance (Corn and Muths 2002). Warmer years, however, are usually drier and may result in increased evaporation and lead to lower water levels in ponds and a concomitant increase in exposure of amphibians to UVB (Kiesecker et al. 2001). Although some studies have already demonstrated a trend for amphibians to breed earlier in some areas (Beebee 1995, 2002; Gibbs and Breisch 2001), other long-term studies have found no temporal variation in timing of amphibian breeding (Reading 1998; Blaustein et al. 2001b, 2003b). Six of seven populations reported by Blaustein et al. (2001b) from the United States and Canada did not show a significant trend toward earlier breeding. Most of these amphibian populations, however, displayed a negative relationship between air temperature and breeding time. Comparing the dates Albert Hazen Wright recorded frogs to call around 1900 with records from the same area (New York) for 1990, Gibbs and Breisch (2001) found that four species are now breeding 10-13 days earlier while two species called at similar dates. In Britain, Reading (1998) found a significant relationship between air temperatures prior to breeding and the date of first spawning but did not find a significant trend toward earlier breeding. Global warming may affect the water supply and hydrological cycles of amphibian habitats, thereby altering the water depth and affecting the exposure of amphibians to UVB (Nag1 and Hofer 1997; Kiesecker et al. 2001). Kiesecker et al. (2001) found a significant relationship between precipitation during amphibian breeding and the depth of water at which toad eggs developed. During dry years linked to El Nifio events, eggs of western toads (Bufo boreas) developed in shallower water and embryonic mortality was higher. Western toad eggs can be sensitive to ambient levels of UVB, and dry conditions caused an increase in exposure of eggs to UVB. Kiesecker et al. (2001) suggested that UVB could have contributed to increased egg mortality in these years. There is a complex relationship among climatic change, UVB radiation and outbreaks of disease. Kiesecker et al. (2001) showed that when water levels decrease, UVB exposure increases and eggs become infected with a pathogenic oomycete, Saprolegnia, more readily than when water levels are higher. The increase in frequency and magnitude of El Nifio events as a result of global warming (Meehl and Washington 1996; Timmermann et al. 1999) could increase the exposure of amphibians to detrimental levels of UVB (Kiesecker et al. 2001). Corn and Muths (2002) used surface levels of UVB irradiance to document the decrease of embryonic exposure to UVB caused by earlier breeding. They found an average decrease of 20% in UVB irradiance when embryos develop one month earlier in the season (May instead of June). As Kiesecker and Blaustein (2001) demonstrated, however, climatic warming

3118

AMPHIBIAN BIOLOGY

can result in embryos developing in shallower water. In ponds with high DOC concentrations, embryos can develop earlier in dry years due to climatic warming, but because of lower pond levels they may be in shallower water where they become exposed to surface levels of UVB radiation, essentially counteracting the effect of the elevated DOC concentrations. In cold and wet years, embryos can develop later in the season and in deeper water. In terms of exposure to UVB, depth is often more relevant than time of breeding. In ponds with low DOC concentrations, water depth can greatly influence UVB exposure in a way roughly similar to that of time of breeding. The relative importance of variation in water depth and surface UVB irradiance needs to be addressed before there can be complete understanding of the impact of climatic warming on amphibian reproduction. Using data from Blaustein et al. (2001b), Corn (2003) found a significant relationship between amphibian breeding-date and snowmelt. Corn predicted that in temperate zones snow cover will be reduced in extent and duration in future decades, and that montane populations of amphibians will breed earlier. 111. EFFECTS OF W B RADIATION O N AMPHIBIAN EGGS, LARVAE AND ADULTS

The effects of UVB radiation vary with life history stage and ecological context (see Blaustein and Kiesecker 2002 for detailed discussion). Moreover, there are strong interspecific and intraspecific effects of UVB radiation on amphibians. Investigators at numerous sites worldwide have shown that ambient UVB radiation lowers the hatching success of some amphibian species at natural oviposition sites (reviewed by Blaustein et al. 1998, 2001). Fertilized eggs typically were placed in enclosures with filters that either removed UVB radiation (experimentals) or allowed UVB to penetrate (controls). The hatching success of eggs under the two regimes were compared. In some studies, enclosures with no filters were used as an additional control. These studies demonstrated that the embryos of some species are more susceptible to UVB radiation than are others (Blaustein et al. 1998, 2001) (Table 1). For example, in the Pacific Northwest (USA), the hatching success of cascades frogs (Rana cascadae), western toads (Bufo boreas), long-toed salamanders (Ambystoma macrodactylurn) and northwestern salamanders (A. gracile) was lower when exposed to ambient UVB radiation than when eggs were shielded from UVB (Blaustein et al. 1998). However, the hatching success of spotted frogs (Rana pretiosa and R. ZuteiventG), red-legged frogs (R. aurora) and Pacific tree frogs (Hyla regilla) was not significantly different between the UV-shielded and W-exposed treatments (Blaustein et al. 1998). In California, the hatching success of Pacific treefrogs was not affected by ambient levels of UVB radiation but hatching success was lowered in California treefrogs (Hyla cadaverina) and California newts (Taricha torosa) by exposure to UVB (Anzalone et al. 1998). The hatching success of common toads (Bufo bufo) in Spain was lower in UVB-exposed eggs than in those shielded from UVB whereas there was no effect of UVB on the hatching success of natterjack toads (Bufo calamita) (Lizana and Pedraza 1998). In Finland, the hatching success of moor frogs (Rana arvalis) increased when eggs were shielded from UVB but there was no effect of protection from UVB on hatching success in common toads (Bufo bufo) and common frogs (Rana temporaria) (Hakkinen et al. 2001). As the studies described above illustrate, there are differences in how species are affected by UVB radiation (Table 1). For some species, in field experiments, hatching success is lower when eggs are exposed to UVB radiation compared with shielded controls. In other species, hatching success is not affected by UVB exposure. This is not a contradiction as is erroneously reported by some investigators (e.g., Corn 1998). Rather, these studies illustrate clear interspecific differences in tolerance to UVB radiation at early life stages. In fact, within the same study, conducted at the same time and site, it has been shown that the embryos of some species are sensitive to UVB whereas the embryos of other species are resistant (e.g., Blaustein et al. 1994b; Anzalone et al. 1998; Lizana and Pedraza 1998).

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3119

Importantly, although hatching rates of some species may appear unaffected by ambient UV radiation in field experiments, an increasing number of studies illustrate a variety of sublethal effects due to UV exposure. For example, when exposed to UVB radiation, amphibians may change their behaviour (Nagl and Hofer 1997; Blaustein et al. 2000; Kats et al. 2000), growth and development may be slowed (e.g., Belden et al. 2000; Pahkala et al. 2000, 2001; Smith et al. 2000) or UV can induce developmental and physiological malformations (e.g., Worrest and Kimeldorf 19'76; Hays et al. 1996; Blaustein et al. 1997; Fite et al. 1998). Some specific examples of sublethal effects follow (see also Table 1 ) . In field experiments, long-toed salamander (Ambystoma macrodactylum) embryos exposed to UVB radiation not only hatched at a much lower frequency than did those shielded from W B but also displayed a much higher proportion of deformities (Blaustein et al. 1997). Thus, only 14.5% of the embryos survived when exposed to UVB radiation compared with 95% survival in the shielded regimes. Moreover, more than 90% of the survivors exposed to UVB radiation were deformed, compared with only 0.5% that were deformed under UV blocking shields. Changes in behaviour after exposure to UVB radiation have been reported in a number of studies. For example, exposure of roughskin newts (Taricha granulosa) to UVB radiation in the laboratory caused them to increase their activity (Blaustein et al. 2000). Changes in activity patterns after exposure to UVB radiation have been observed in a number of other species as well (e.g., Nagl and Hofer 1997; Zaga et al. 1998) (Table 1 ) . Anti-predator behaviours may also be affected by exposure to UVB radiation. For example, cascades frog tadpoles (Rana cascadae) and juvenile western toads (Bufo boreas) exposed to low levels of UV radiation did not respond to the chemical cues of predators as quickly as did those that were not exposed to UV radiation (Kats et al. 2000). Low level exposure to UVB radiation in the laboratory causes a number of developmental and physiological deformities in frogs and toads (Worrest and Kimeldorf 1976; Hays et al. 1996) (Table 1). These include oedema, skeletal anomalies and eye damage. Adult Rana cascadae from Oregon have distinctive outer retinal abnormalities in the inferior retina that include the abnormal distribution of retinal pigment, damaged photoreceptors and the presence of large pigment-filled macrophages consistent with damage by solar radiation. Fite et al. (1998) suggested that this damage may significantly impair the vision of R . cascadae that bask in sunlight at relatively high elevations. The impairment of vision or the inability to perceive chemical cues from predators after exposure to relatively small doses of UV radiation may have profound implications for amphibians. Individuals that cannot perceive predators obviously will be at a significant disadvantage compared with those individuals that can. Moreover, if such impairments occur in a number of species, the amphibian component of certain ecological communities may be altered. Sublethal effects may become evident even in species whose embryos appear to be resistant in field experiments. Several experimental studies illustrate that early exposure to UVB radiation causes delayed effects in later stages. For example, UVB radiation did not influence hatching success of plains leopard frogs ( R a m blairi) but growth and development were slower in tadpoles that, as embryos, had been exposed to the highest levels of UVB radiation (Smith et al. 2000). Exposure of embryos of Rana temporaria to UVB radiation had no effect on survival rates, frequency of developmental anomalies or hatching size (Pahkala et al. 2001). Larvae exposed to UVB radiation as embryos, however, displayed an increased frequency of developmental anomalies, metamorphosed later and were smaller than larvae shielded from UVB as embryos (Pahkala et al. 2001). Similarly, UVB radiation had no effects on hatching success in red-legged frogs (Rana aurora) (Blaustein et al. 1996). Larvae exposed to UVB radiation as embryos, however, were smaller and less developed than were those shielded from UVB (Belden and Blaustein 2002b).

3120

AMPHIBIAN BIOLOGY

Delayed growth and development after exposure to UVB radiation may significantly affect populations of amphibians that live in ephemeral habitats. For example, if growth and development is slowed significantly and amphibians cannot metamorphose and move onto land before a pond dries or freezes, significant mortality may occur (Blaustein et al. 2002). Most of the studies that have documented mortality after exposure to UVB radiation have been conducted at the egg or embryonic stage but exposure to UVB radiation can also be fatal at other life stages. For example, salamander larvae and juvenile frogs can also die when exposed to ambient levels of UVB radiation (e.g., Hakkinen et al. 2001; Belden and Blaustein 2002a,b; Blaustein et al. 2005). IV. ECOLOGICAL AND BEHAVIOURAL PHOTOPROTECTIVE STRATEGIES

Adults and, especially, larvae of many amphibian species are sensitive to UVB (Blaustein and Kiesecker 1997) but may easily avoid harmful levels of it by selecting specific shaded microhabitats (Hofer 2000). Mobile stages of some amphibian species can detect harmful levels of UVB and behaviourally avoid them (e.g., van de Mortel and Buttemer 1998; Garcia et al. 2004). Individuals of other species, however, may not avoid areas with high levels of UVB but remain both in UVB-exposed and un-exposed locations with equal frequency (van de Mortel and Buttemer 1998; Adams et al. 2001; Belden et al. 2003). For some species, eggs are less sensitive than are larvae when exposed to UVB in clear water (Tietge et al. 2001). Eggs, however, are not mobile and cannot escape UVB. Females may protect eggs from UVB, either directly or by selecting un-exposed oviposition sites (Blaustein and Belden 2003). Over evolutionary time, selection pressure for seeking thermal regimes and water quality that maximize growth and development were probably important in shaping the behaviour of many amphibian species (Blaustein et al. 2004). Eggs are often laid in shallow water or even floating at the surface where they develop in a more oxygenated environment and may have lower risk of predatory or parasitic attacks. Oxygenation of eggs is critical to their development (Duellman and Trueb 1994). Larvae also often seek shallow water where thermal conditions increase their growth rate (Wollmuth et al. 1987). Many frog species bask in sunlight for prolonged periods of time. Maintenance of higher temperatures or development in more oxygenated water may, however, influence exposure of amphibians to UVB (Belden et al. 2000). In open terrestrial situations or in shallow water they can be exposed to high levels of UVB radiation that can cause significant damage to their skin, eyes and other portions of their body (Blaustein et al. 1994; Fite et al. 1998). Even limited exposure to small doses of UVB radiation can damage amphibians. For example, less than 1% of incident UVB can cause retinal damage that can affect visual acuity (Ham 1983) to varying degrees. Of course, damage by UVB radiation depends upon the species and the defence mechanisms they have to cope with the harmful effects of exposure (Blaustein and Belden 2003). Eggs of caudates seem more sensitive to UVB. Many species select oviposition sites that efficiently hide eggs from UVB (Table 2). Only about 20% of caudate species place eggs where they would be often exposed to damaging levels of UVB (Table 2). There is no information about the sensitivity of caecilian embryos to UVB but it seems that the embryos of all species develop where they are sheltered from it (Table 2). Anurans are more likely than are salamanders to deposit their eggs in sites exposed to UVB (Hofer 2000). About 60% of the anuran species that have been studied oviposit where the eggs are often exposed to high ambient levels (Table 2). Information about nest-site selection is scarce for tropical species where terrestrial nesting, embryonic development in foam nests or aquatic development in densely forested or vegetated areas is common. Amphibian species from temperate zones and from savannah or deforested tropical ecosystems are better candidates for exposure to high levels of ambient UVB. Alternatively, if one only considers species that inhabit temperate zones, the percentage that have embryos often exposed to UVB increases significantly.

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3121

Table 2. Habitats selected for oviposition and embryonic development by various families of amphibians and a preliminary assessment of the degree of exposure of their eggs to UVB radiation. Sources: Amphibia Web (2004); LivingUnderWorld Web (2004); AnimalDiversity Web (2004); Duellman and Trueb (1994); pers. obs.

Family

Number of species with data on Number usual level of known of exposure species to UVB

CAUDATA Sirenidae

520 4

Hynobiidae

44

Cryptobranchidae

3

Proteidae

6

Dicamptodontidae

4

Amphiumidae Rhyacotritonidae

3 4

Salamandridae

63

Ambystomatidae

30

Plethodontidae

359

GYMNOPHIONA 168 Rhinatrematidae 9 Ichthyophiidae 38 Uraeotyp hlidae 5 Scolecomorphidae 6 Caeciliidae 110 ANURA 4 928 Allophrynidae 1 Ascaphidae 2 Bombinatoridae 11 Leiopelmatidae Discoglossidae

4 11

Rhinophrynidae

1

Pipidae

30

Reproductive mode or habitat of the eggs

General exposure to W B radiation

Number of species* and the genera in which most of the eggs are often exposed to high UVBR**

From high to low Attached to submerged vegetation Attached to stones or From high to nil vegetation or in underground springs Water in burrows or Very low or nil beneath stones Very low or nil Water beneath stones, logs or other shelters Under stones, in cracks Very low or nil or gravel or attached to woody debris On mud under shelters Very low or nil Cracks deep in rocks Very low or nil at the head of springs From shallow to deep From high to nil water, attached to or wrapped in vegetation, inside deep cracks, under stones, in terrestrial shelters, or ovoviviparous Terrestrial under debris From very high to and logs or aquatic in very low shallow water, attached to branches, vegetation, under stones or on the pond bottom Nil or very low Aquatic beneath rocks or terrestrial in the soil, below rocks and logs, in crevices deep in rocks or inside large logs Terrestrial shelters Mud near water Probably terrestrial Viviparous ? Viviparous or terrestrial eggs

Nil Very low or nil Very low or nil Nil Very low or nil

Aquatic Aquatic under stones Aquatic attached to vegetation Damp terrestrial sites Shallow water or carried by male Floating

Unknown Very low From high to medium Very low From very high to very low Very high

Aquatic, floating or on From very high to vegetation or embedded very low in skin of female

18 (20%) 4 (100%) Siren, Pseudobranchw 3 (60%) Batrachuperw, Hynobius, Salamandrella 0

6 (20%) Nothophthalmw, Pleurodeles, Taricha, Triturus

5 (56%) Ambystoma

265 (60%) 0 0 9 (100%) Bombina 0 6 (55%) Discoglossw 1 Rhinophrynw 3 (30%) Silurana, Xenopw

3122

AMPHIBIAN BIOLOGY

Table 2 - continued

Family

Number of species with data on Number usual level of known of exposure species to W B

Megophrydae

128

Pelobatidae

4

Pelodytidae

3

Scaphiopodidae

7

Myobatrachidae

124

Heleophrynidae

6

Sooglosidae

4

456

Brachycephalidae

6

Rhinodermatidae

2

Pseudidae

9

General exposure to UVB radiation

From very shallow to deep water or ovoviviparous and viviparous Terrestrial eggs coated with soil particles Guarded by males in vocal sac Aquatic vegetation in shallow water On plants over ponds; aquatic, attached to rocks or vegetation, under stones, in water-filled cavities of plants; terrestrial or carried by parents Under leaves or on moss, plants, or rocks above streams Terrestrial chambers or in leaf litter; aquatic eggs in streams

From very high to nil

From high to low

Centrolenidae

139

Arthroleptidae

78

Dendrobatidae

2 16

Humid terrestrial and arboreal sites

Very low

Ranidae

729

Usually aquatic eggs floating in shallow water, on vegetation or under stones; few cases of terrestrial hidden nests

From very high to very low

Subterranean burrows

Nil

1

0 6 (35%) Ceratophrys, Hylorina, Lepidobatrachus, Leptodactylw 19 (95%) Atelopw, Bufo

Nil

849

9

4 (100%) Pelobates 2 (67%) Pelodytes 7 (100%) Scaphiopus, Spea 17 (25%) Crinia, Mixophyes, Neobatrachw, Notaden, Philoria, Spicospzna, Uperoleia

Very low or nil

Hylidae

Hemisotidae

Number of species* and the genera in which most of the eggs are often exposed to high WBR**

Aquatic, floating or From high to low in shallow water Aquatic, floating or Very high to medium in shallow water Attached to aquatic High to medium stems or on vegetation Aquatic, floating or in Very high to medium shallow water Terrestrial foam nests From nil to very high under debris, tunnels; aquatic in shallow water, in vegetation or under rocks; floating foam nests; brooded in adult stomach Aquatic, attached to Low rocks Terrestrial or carried by Very low or nil adults Unknown Terrestrial or foam nests From high to very low or aquatic eggs

Nasikabatrachidae 1 Leptodactylidae 1 125

Bufonidae

Reproductive mode or habitat of the eggs

From very high to very low

2 (100%) Pseudis, Lysapsw 57 (49%) Acris, Cyclorana, H y h , Litoria, Osteopilw, Pseudacris, Scinax

Low or very low

From low to nil

52 (84%) Amnirana, Fejervarya, Hildebrandtia, Hoplobatrachw, Limnonectes, Paa, Phrynobatrachw, Ptychadena, Pyxicephalw, Rana, Sphaerotheca, Emopterna 0

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3123

Table 2 - continued

Family

Number of species with data on Number usual level of known of exposure species to UVB

Hyperoliidae

253

38

Rhacophoridae

2 17

9

Mantellidae

151

23

Microhylidae

363

17

Reproductive mode or habitat of the eggs Aquatic, on vegetation over ponds, in tree holes, on the ground or terrestrial foam nests Foam nests; on regetation overhanging ponds; in tree holes Stagnant shallow water, attached to rocks or plants or in the bottom of ponds; terrestrial under stones or moist leaf litter; plants above brooks Aquatic at the surface or attached to vegetation; terrestrial under debris, rocks or logs, in waterfilled bamboo trunks; foam nests

General exposure to UI'B radiation

Number of species* and the genera in which most of the eggs are often exposed to high UVBR**

From high to very low

12 (32%) Heterucalw, Hyperolius, Kmsina, Parakassina, Semnodactylw

Very low or nil

From very high to very low

From high to nil

7 (30%) Aglyptodactylus, Boophis, Mantzdactylw

7 (41%) Dyscofhw, Gastrophryze, Hypopachw, Phrynomantis, Uperodon,

*For most of species, especially those from tropical areas, there is no precise available published information about the degree of exposure of their eggs to UV radiation. The number of species exposed to high levels of UV radiation is, thus, underestimated. **Eggs that are nahrally exposed in their undisturbed habitats. Habitat alteration can substantially modify the degree of exposure of eggs to UV radiation.

There are several behavioural mechanisms that amphibians use to avoid exposure of eggs or young to harmhl UVB. Some of these involve the parents carrying their offspring away from sunlight. A. Nocturnality

Nocturnality is the most effective way of avoiding solar radiation. Adults of many terrestrial and aquatic amphibian species are only active during the night or during crepuscular periods. This pattern of activity is also exhibited by larval stages of some species that rest on the bottom or are hidden in refuges during the day but move to open water at night (Stebbins and Cohen 1995). At night, amphibians may be less exposed to predators, and terrestrial amphibians do not lose water from their bodies as readily as during dry diurnal conditions. Additionally, amphibians that are active at night can completely avoid exposure to UVB radiation. An exceptional life history trait is exhibited by the nocturnal Alytes. Males carry strings of eggs over their legs until hatching time (MArquez 1992). During dry periods, egg-carrying males move at night to ponds or streams and stick their legs into the water to hydrate the eggs. This parental behaviour avoids exposure of the eggs to UVB radiation. B. Viviparity and other Egg-brooding Systems

Amphibian eggs are obviously quite a vulnerable developmental stage because they cannot actively avoid exposure to sunlight (Blaustein et al. 1998) and consequently may often be exposed to UVB radiation. Egg-retention by females is one way whereby eggs may be protected from UVB radiation. Ovoviviparity and viviparity have evolved independently in several amphibians and selective pressures for the evolution of these reproductive modes have been diverse (Wake 1993). For example, viviparous females can actively protect eggs from predators or optimize developmental rate by selecting warmer sites. Internal eggs can

3124

AMPHIBIAN BIOLOGY

develop during periods when they would be unlikely to survive if laid directly in the environment (Wake 1993). Protection of eggs from harmful UVB by their retention within the mother may be one of the factors to have driven the evolution of viviparity. Although oviparity seems to be the dominant reproductive mode in caecilians (Wilkinson and Nussbaum 1998), there are many tropical species that retain their eggs within' the oviducts and bear live young. Even eggs of non-viviparous species, however, are not exposed to UVB (Table 2). Viviparity has evolved independently in anurans and caudates (Duellman and Trueb 1994) and in both groups many species that exhibit this reproductive mode inhabit montane areas where exposure to UVB is high during times that embryos develop. Several anurans carry eggs or larvae in special pouches that protect them from UVB. For example, Pipa and Gastrotheca transport their eggs embedded in the dorsal skin of the female, while Assa carries tadpoles inside paired inguinal pouches. Tadpoles of Rhinoderma develop in the vocal sac of the male and eggs and tadpoles of Rheobatrachus grow in the female's stomach (Duellman and Trueb 1994). These modes of parental care protect eggs from predation and/or desiccation but they also serve to shelter developing stages from exposure to harmful UVB radiation. C. Terrestrial Oviposition

'

Many species of salamanders, anurans and caecilians deposit their eggs in subterranean terrestrial nests where embryos develop until metamorphosis (Duellman and Trueb 1994; Amphibia Web 2004). Adults often guard the eggs and may even hydrate them. In some cases females wrap or attach their eggs to leaves of terrestrial plants in shaded habitats. In these microhabitats, eggs are sheltered from UVB. There are also some species that lay their clutches under debris or dead leaves at the margins of partially or totally dry ponds. Embryos develop in this environment until the water level rises, eventually inundating the clutch. Submersion of the eggs usually triggers hatching (Petranka et al. 1982). D. Aquatic Refuges

Riparian and aquatic vegetation may provide efficient protection to eggs, larvae and adults. Newt and salamander larvae may be sensitive to ambient levels of UVB when they are fully exposed to sunlight (Blaustein et al. 1995; Nagl and Hofer 1997; Lesser et al. 2001; Belden and Blaustein 2002a,c). Both larvae and adults, however, are mobile and can swim to shaded areas and thereby avoid the harmful effects of UVB. In very shallow ponds with no vegetation or other type of shelter, as is typical of many high montane areas or of sandy or stony sites, larvae may have no possibility of avoiding UVB. In some high montane lakes, the only refuges from UVB are in deep water but temperatures there are intolerably low for amphibians and they must select very shallow water for oviposition and for larval activity. In these habitats Nagl and Hofer (1997) found sublethal UV-like histological damage in alpine newt (Tm'turus alpestris) larvae. Many aquatic newts and salamanders and some anuran adults can be observed in clear water during the night and also during the day on cloudy and rainy days. When the sun shines brightly, however, they quickly hide in vegetation, in the mud at the bottom of the ponds and streams, or beneath stones, logs or other shelter. Garcia et al. (2004) found that the larvae of two ambystomatid salamanders moved to deeper water when experimentally exposed to UVB radiation. Aquatic anuran tadpoles of many species rarely are observed in exposed open locations. They usually occupy densely-vegetated areas during the daylight or hide under various types of underwater shelters. This behaviour probably protects them from predation and dehydration as well as from harmful levels of UVB. Diamond et al. (2002) suggested that behaviours that limit the exposure time by even 50% would keep the animal above toxic levels. It must be realized, however, that toxic levels differ among species or even among stages within a given species. Thus, it is not easy to generalize about doses and toxic levels. Alpine newts (Triturus alpestris) exposed to artificial UVB swam erratically until they reached protected areas. Within five minutes of exposure, 90% of the larvae had escaped

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3125

from the UVB-irradiated area to a shaded place (Nagl and Hofer 1997), while larvae under only visible light showed a positive phototactism. However, Belden et al. (2003) observed that tadpoles of Rana cascadae did not discriminate among levels of UVB and did not avoid lethal levels when they had the option to do so. They also observed a lack of discrimination and avoidance of harmful UVB levels in Ambystomu macrodactylum. Sunlight-avoiding behaviour is not shared by many species. In fact, many species seek sunlight. Thermoregulatory basking is a well-documented phenomenon in anuran amphibians (Duellman and Trueb 1986; Hutchison and DuprC 1992; Stebbins and Cohen 1995). In a study examining thermoregulatory behaviour of metamorphosed American bullfrogs (Ram catesbeiana), approximately 60-80% of the frogs in study ponds were observed basking on the shore at midday. On cloudy days, the percentage of basking bullfrogs dropped below 35% (Lillywhite 1970). Larvae of many amphibian species seek warm, sunny regions of ponds for extended periods of the day (Beiswenger 1977; Warkentin 1992; Hoff et al. 1999). For example, Salamandra salamandra and Ambystoma macrodactylum larvae are easily observed on sunny days in exposed, shallow, transparent montane ponds (Belden et al. 2000; pers. obs.). Similar behaviour is exhibited by tadpoles of some bufonids (Bufo calamita) and ranids (Rana temporaria) that inhabit montane areas (pers. obs.). The high degree of skin pigmentation in the larvae of these species may protect them from UVB. Many frogs and newts deposit aquatic eggs on submerged vegetation, attaching them to leaves or stems of aquatic plants. Vegetation has substantial impacts on local UVB irradiance levels and dose and, accordingly, protects amphibians from noxious effects (Peterson et al. 2002). In field experiments, Marco et al. (2001) tested the effects of ambient levels of UV radiation on the embryos of marbled newts (Triturus marmoratus) that were exposed to sunlight. Females of this species carefully wrap their unpigmented eggs in leaves of aquatic plants in very shallow water. During embryonic development, some eggs become unwrapped for various reasons. When eggs were artificially unwrapped, 95-100% of these embryos died during the first 14 days of exposure to sun. Such embryos showed skin damage, oedemas and fungal infections and lost their rounded shape. In contrast, during the same period, mortality of embryos protected by WB-blocking filters was only 20%. This high sensitivity of newt eggs to UVB could be shared by the unpigmented eggs of many amphibian species that lay eggs in places not usually subject to high levels of UVB. In laboratory experiments, Marco et al. (2001) exposed marbled newt embryos to an artificially elevated intensity of W B radiation and tested the protective effect of leaves. The mortality of eggs wrapped in leaves and exposed to UVB radiation was low and was similar to that of unwrapped eggs shielded from UVB radiation by a filter. Of 120 unwrapped eggs exposed to W-radiation, 119 died within five days (Marco et al. 2001). Eggs of other newt species are also attached or wrapped in aquatic vegetation and, therefore, are well protected from UVB (Nagl and Hofer 1997). Other species (e.g., rhyacotritonids, dicamptodontids, ascaphids) prefer to hide their eggs under stones, logs or other types of submerged refugia or even select underground springs for oviposition (hynobiids, proteids) (Duellman and Trueb 1994; Amphibia Web 2004). There are many species that deposit their eggs in very shallow, open water. For example, some newt species select shallow water to lay their eggs and egg-frequency decreases as a function of depth (Miaud 1995). Some eggs that are laid in very exposed locations do not show any apparent UVB-induced damage. Exposure to W B during embryonic development can, however, result in morphological or developmental alterations in the larvae, even in those that subsequently hide from UVB in their natural habitats (Smith et al. 2000b; Pahkala et d.2001a; Belden and Blaustein 2002b). Susceptibility of prey to UVB may indirectly influence habitat selection by predators. For example, some invertebrates fed on by newts, such as larval chironomids (Diptera), are sensitive to W B and are not present in areas fully exposed to sunlight (Bothwell et al. 1994). Chironmids feed on algae but are more sensitive to UVB than are the algae. This

3126

AMPHIBIAN BIOLOGY

difference in sensitivity causes a counterintuitive increase in algal density in UVB-exposed areas compared to non-exposed ones because of lower predation rates by chironomids on algae in areas exposed to UVB (Bothwell et al. 1994). Anuran tadpoles that feed on algae may then select areas exposed to higher levels of W B if this allows them to avoid other algal grazers that are more sensitive to UVB. Tropical species inhabit areas with the highest absolute levels of UVB radiation, but most such species select habitats that are protected by a dense arboreal canopy. This is true even for temporal aquatic ponds in tropical regions. There are, however, some exceptions in which amphibians are exposed to high and harmful levels of UVB (Lips 1998; Blaustein and Kiesecker 2002). For example, deforestation of tropical habitats may cause a sudden exposure to high (and potentially harmful) levels of UVB radiation in amphibians that have not developed protective mechanisms against it.

E. Deep Water and Dissolved Organic Content (DOC) Concentration The depth of UV penetration in ponds and the variables that control such penetration are important factors for a thorough understanding of the influence of UV on amphibians. It is important to understand how the concentration of dissolved organic content (DOC) acts to attenuate UVB in water (Morris et al. 1995; Williamson et al. 1996; France et al. 2000; Palen et al. 2002). Several studies have documented DOC concentration in amphibian habitats. These show that the amount of DOC varies both temporally and spatially. Berrill and Lean (1988) pointed out that in some ponds and marshes with high DOC concentrations 90% of UVB is attenuated in the first 10 cm of water. Crump et al. (1999b) found that in some ponds in Canada 99% of UVB radiation was attenuated in the top 1020 cm with a high DOC concentration of around 13 mg/L. Peterson et al. (2002) found similar results in some ponds in the northern United States. In that study attenuation from DOC was even higher than predicted during springtime when amphibians lay their eggs. In Canada, a lake with a low DOC concentration of 0.5 mg/L, showed 34% of the surface UVB at 20 cm depth, whereas in a lake with a DOC level of 7.8 mgIL, only 1% of surface UVB was measured at that depth (Scully and Lean 1994). Palen et al. (2002) found a strong relationship between UVB attenuation and dissolved organic matter concentration in 136 amphibian breeding ponds in the Pacific Northwest of the United States. In that study, at 10 cm depth the levels of UVB were undetectable in some sites while in others they were similar to those at the water's surface. It was estimated that 85% of the sampled sites had irradiance levels lower than 22.4 pWlcm2 (Palen et al. 2002) but the reported levels were within the range that can cause death or sublethal damage to amphibians (Blaustein et al. 2004). Moreover, Palen et al. (2002) did not account for temporal changes in UVB and relied generally on single measurements at specific sites. In certain ponds inhabited by newts, Nagl and Hofer (1997) measured a high concentration of dissolved organic matter. In those ponds most UVB was absorbed in the first 20 centimetres of water. High DOC concentrations may allow more sensitive amphibian larvae to inhabit shallow and unshaded portions of a pond with little or no vegetation. For example, in one study, Rana cascadae selected ponds with high DOC and low UVB penetration in the Olympic National Park in Washington, United States (Adams et al. 2001). Nagl and Hofer (199'7) showed that newts were often absent from shallow, relatively transparent ponds with constant water flow. Severe skin damage and even mortality, apparently caused by exposure to UVB radiation, was observed. Freshwater fishes may spawn deeper in lakes with lower UVB attenuation (Williamson et al. 1997) but there is no information as to whether amphibians modify spawning depth as a function of UVB penetration into the water column. In montane lakes, DOC and phytoplankton, both of which absorb W B , often are low and accordingly penetration of UVB can be great. For example, Tartarotti et al. (1999) found that lakes from the Alps and the Andes had UVB levels 10% of surface values at depths of 9.6 and 12.8 m, respectively. Morris et al. (1995) found that, in clear montane lakes, 1% of surface UVB penetrated deeper than 30 m. Adams et al. (2001) estimated that in amphibian

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3127

habitats from montane areas in the northwestern United States, 45% of surface radiation may reach 50 cm depth. When DOC concentration is higher than 5 mg carbon per litre (CIL) only amphibians that are floating or in shallow water are exposed to detectable levels of UVB (Crump et al. 1999b). When DOC is lower than 5 mg CIL, attenuation of UVB decreases rapidly (Crump et al. 1999b) and amphibians can be exposed to harmful levels of UVB in deeper water. In freshwater ecosystems with DOC concentrations below 1-2 mg CIL, small changes in DOC may cause rapid variation in UV attenuation (Williamson 1996). While in many lowland lakes, DOC can be used to predict UV transparency with reliable accuracy, current models fail to estimate UV in clear alpine lakes. In such lakes, phytoplankton may contribute significantly to UV attenuation, either as particles or as a source of chromophoric dissolved organic matter (CDOM) with distinctive properties. In transparent alpine lakes, phytoplankton production and the dynamics of the CDOM pool have a strong effect on temporal changes in underwater attenuation of UV (Laurion et al. 2000). Many amphibian species lay their eggs in shallow water (e.g., Bufonidae, Discoglossidae, Pipidae). The water is often so shallow that the eggs are above the surface and are open to the air and exposed to high levels of UVB radiation (Blaustein et al. 1994a). After they are laid, the eggs of some anurans may sink to deeper water or to the bottom where they complete development. Females of several species seek deeper water in which to lay their eggs. This behaviour can be effective in protecting eggs from UVB radiation but, very often, ponds dry during embryonic development and eggs become exposed to increasing levels of UVB as water level drops. In extreme cases, some ponds dry completely before amphibians can metamorphose and leave the pond. When this occurs, aquatic amphibians are stranded with no cover and are subjected to intense levels of UVB radiation. For example, in Oregon and in many other parts of its range, A m b y s t o m gracile attaches its clutches to rigid branches or sticks suspended in the water column (Nussbaum et al. 1983). When the water level decreases, clutches become exposed to increasing levels of UVB and can even become suspended above the water or stranded (e.g., Marco and Blaustein 1998). In lowland Mediterranean regions, some amphibians lay eggs in open water with high DOC concentration and at different depths. In these habitats, when water level recedes, eggs that were laid underwater may begin to float or become stranded over aquatic vegetation and exposed to surface UVB levels (pers. obs. of AM). In these cases, the initial photoprotection due to depth, DOC and aquatic vegetation cover decreases and eggs may suffer the effects of full UVB exposure. Kiesecker et al. (2001) documented the relationship between climatic events and the year-to-year variation in the mean water depth at which amphibian eggs developed. Although water depth at which eggs developed was partially dependent on the depth at which the eggs were laid, there was also a significant impact by precipitation or melting of snow during the time the embryos were developing (Kiesecker et al. 2001). V. MOLECULAR AND MORPHOLOGICAL ADAPTATIONS Species that are exposed to UVB at some developmental stage do not necessarily suffer the deleterious effects of UVB exposure. UVB radiation probably had a major impact on the evolution of life from very early times (Cockell 2001) and amphibians, like many other organisms, have been exposed to UVB since their origin. There is evidence of previous short-term episodes of levels of UVB that may have been even greater than those caused by present-day stratospheric ozone depletion (Leavitt et al. 1997; Cockell 2001). Long-term exposure to UVB may have favoured the evolution of different photoprotective strategies permitting amphibians to colonize habitats with high levels of UVB (Licht and Grant 199'1; Hofer 2000; Blaustein and Belden 2003). Moreover, characteristics that serve other biological hnctions may secondarily protect amphibians against UVB. Recent reviews document different methods that amphibians or other aquatic species use in coping with UVB (Epel et al. 1999; Blaustein and Belden 2003). Eggs, larvae and adults of some amphibian species normally exposed to relatively high levels of W B are somewhat tolerant to the full exposure of ambient and even enhanced

3128

AMPHIBIAN BIOLOGY

levels of UVB (Blaustein et al. 1996, 1999; Cummins et al. 1999; Pahkala et al. 2001b). At the other extreme, many species inhabit unexposed sites and usually lack effective photoprotective mechanisms. Along this gradient, some species evolved photoprotection against historical levels of UVB but not to enhanced levels or to synergistic effects with emerging stressors. Some species, therefore, may be able to survive in the face of changing levels of UVB because their photoprotective systems can cope with systematic exposure to increasing levels of UVB, whereas others may not. Different developmental stages of a given species may vary markedly in the level of UVB to which they are exposed as well as in the effectiveness of their photoprotective systems. A. Enzymatic Repair Systems

When UV radiation penetrates a cell, DNA photoproducts form that can lead to mutations or cell death. Embryos of some amphibian species may be more resistant to UVB because they can repair such damage more effectively than can others. One important repair process is enzymatic photoreactivation. One enzyme, CPD-photolyase, uses visible light energy (1 300-500 nm) to repair the most frequent UV-induced lesion in DNA, cyclobutane pyrimidine dimers (CPDs) (Friedberg et al. 1995). A second, related enzyme, [6-41photolyase, similarly uses light energy to reverse pyrimidine-[64']-pyrimidone photoproducts ( [ 6 4 ] photoproducts). Moreover, multi-protein, broad-specificity, excision-repair processes can remove CPDs and [ 6 4 ] photoproducts. Both mechanisms may occur simultaneously but excision repair is typically more effective for [ 6 4 ] photoproducts than for CPDs. Thus, CPD-photolyase appears to be the first level of defence against CPDs for many organisms exposed to sunlight (Pang and Hays 1991; Friedberg et al. 1995). Photolyases do not repair DNA in the dark (Vetter et al. 1999) but can stimulate the nucleotide excision-repair system, the other major DNA repair system for UVB-damage (Ozer et al. 1995). Photolyases are found in many organisms such as fungi (Berrocal-Tito et al. 1999), plants (Pang and Hays 1991), invertebrates (Malloy et al. 1997), fishes (Vetter et al. 1999) and mammals (Yasui et al. 1998). Sublethal exposure to W B usually causes a die1 cycle of dimer concentration and repair activity that tracks sunlight intensity (Vetter et al. 1999). A significant and even high repair rate may not be enough to prevent the accumulation of CPDs if the exposure to UVB is intense and prolonged (Malloy et al. 1997). Photolyase activity is markedly sensitive to environmental temperature. Both high (Pang and Hays 1991) and low (MacFayden et al. 2004) temperatures may significantly shorten life or decrease activity of the photorepair system. Especially relevant for amphibians could be sensitivity to cold. Photolyases are less effective at repairing DNA damage at low temperatures. Amphibians that live or breed in cold water and depend heavily on DNA repair processes may therefore be less able to survive high UV exposure. Licht and Grant (1997) found a higher proportion of abnormal Rana sylvatica embryos exposed to UVB at 12°C than of those that were exposed to UVB at 20°C, possibly because of impaired photolyase activity. Photolyase activity is also affected by pH. Neutral pHs are optimal for the repair of photolyase (Sancar and Sancar 1988) and synergisms between pH and UVB may affect photorepair activities (van de Mortel et al. 1998). Exposure to solar radiation in vivo may induce the activity of photorepair enzymes. Embryos of wood frogs (Rana sylvatica) exposed to ambient solar radiation displayed significantly different photolyase activities compared with embryos reared in the dark. There was a positive relationship between photolyase activity and UVB irradiance (Smith et al. 2000a). These authors found significant differences in photolyase activity between embryos of two frog species exposed to similar UVB intensity. Fish embryos spawned in the dark exhibited a strong photorepair response when moved to the light, indicating that photolyase activity does not necessarily depend on previous exposure to light (Vetter et al. 1999). Levels of photolyase activity in eggs and embryos differ substantially among amphibian species (Blaustein et al. 1994a, 2001; Smith et al. 2002) and tend to be positively correlated

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3129

with expected exposure to UVB radiation in the field (Blaustein et al. 1994a; Hays et al. 1996; van de Mortel et al. 1998; Smith et al. 2002). Interspecific differences in photolyase activity may be also related to UVB-based mortality of embryos. Studying several North America11 amphibian species Blaustein et al. (1994a, 1996, 1999) found a negative relationship between photolyase activity and mortality of embryos exposed to ambient UVB. For example, eggs of anuran species such as Hyla regilla, Rana aurora, R. pretiosa and R. luteiventris were relatively tolerant of UVB, as measured by hatching success. These eggs had relatively high levels of photolyase activity. Eggs of Rana cascadae, Bufo boreas, Ambystoma gracile and A. macrodactylum had low levels of photolyase activity and were more sensitive to ambient UVB. Eggs of caudate species that are not exposed to UVB in their natural incubation sites showed relatively low levels of photolyase activity (Blaustein et al. 1994a). In Australia, Litoria aurea had the lowest photolyase activity of three studied species and showed a non-significant trend of reduced hatching success under UVB while L. peronii and L. dentata had lower UVB-mediated mortality and higher levels of photolyase activity (van de Mortel et al. 1998). B. Melanin and other Sunscreens

The synthesis of UV-screening substances is a common and passive method whereby many types of organisms reduce the harmful effects of UVB (Cockell and Knowland 1999). The best-known, common sunscreen in vertebrates is melanin which, when produced in sufficient quantities, causes darkening of the skin (Kollias et al. 1991; Prota 1992; Wu 1999). Melanin seems to be the main sunscreen product in all developmental stages of amphibians. Eggs of species that oviposit in exposed sites usually have large melanin deposits in the animal hemisphere whereas those of species that oviposit in shaded or less-exposed areas have lighter coloration or even lack pigmentation altogether (Duellman and Trueb 1994). The production of melanin and many other photoprotective and antioxidant substances (flavonoids, carotenoids, melanins, serotonin and hemoxygenase) is activated by sunlight in many species (Fraikin et al. 2000). Beudt (1930) found that dark eggs of Rana temporaria were less affected by UVB than were the lighter-coloured eggs of Rana esculenta. Sergeev and Smirnov (1939) also found evidence suggesting that ambystomatid eggs are protected against ultraviolet radiation by pigmentation. DuShane (1943) demonstrated the accumulation of melanophores over the neural tube in pigmented species. Melanin could potentially afford protection to the developing neural tube during neurulation. Jablonski (1998) suggested that melanin would protect embryos against the photolysis of folate and other metabolites that are essential for neural tube formation. Bufo boreas and B. woodhozlsii tadpoles exposed to simulated solar UVB in excess of the actual solar levels observed at their breeding ponds were tolerant and Little et al. (2003) suggested that photoprotective melanin in their skin may protect them from the negative effects of UVB. The epidermis of newt embryos usually contains only a few, scattered melanin granules (Epperlein and Lofberg 1990) and is very sensitive to UVB (Nag1 and Hofer 1997; Marco et al. 2001). In amphibians, the degree of exposure to the sun can influence production of melanin. Zaga et al. (1998) observed that tadpoles darken with exposure to UVB but they did not find a relationship between initial egg colour and sensitivity to UVB. Langhelle et al. (1999) found a significant darkening of Hyla arborea tadpoles exposed to UV. Belden and Blaustein (2002~) observed that five days of exposure to relatively low levels of UVB caused significant darkening of the skin in larval roughskin newts, Taricha granulosa, and in northwestern salamanders, Ambystoma gracile, compared with controls exposed to full-spectrum lighting without UVB. This light-induced increase in skin pigmentation did not, however, fully protect larvae from UVB; experimental manipulation of egg coloration demonstrated that, after three weeks, larvae exposed to UVB were smaller than controls, regardless of skin colour. Lesser et al. (2001) found significant DNA damage in Ambystoma maculatum embryos exposed to UVB despite a significant increase in UVB-induced melanin production. In mammals, however, there is no doubt about the photoprotective function of melanin. In humans, Barker et al. (1995) clearly demonstrated

3130

AMPHIBIAN BIOLOGY

greater susceptibility to the photodamaging and photocarcinogenic effects of sun exposure of individuals with lightly pigmented skin compared to those with dark skin. Perhaps small increases in skin pigmentation in amphibians provide them with some photoprotection but not enough to protect them from intense exposure to UVB. There is evidence for sunscreens other than melanin in amphibians. Hofer and Mokri (2000) found a UVB-absorbing substance in the skin of Rana temporaria tadpoles that had an absorption peak between 290 and 300 nm and its production was induced by both visible and ultraviolet light. This substance could help R . temporaria tadpoles cope with the high levels of UVB typical of their montane lakes at the time of late larval development. Substances with similar properties have been described in invertebrates and fishes (Fabacher and Little 1995; Sommaruga and Garcia-Pichel 1999). For example, some albino fishes do not suffer the harmful effects of UVB because they have a colourless non-melanin photoprotective substance (Fabacher et a2. 1999). These sunscreens can be extracted with methanol from the outer dorsal layers of skin in some fishes. The primary function of these substances in fishes is probably the protection of exposed skin from UVB radiation (Fabacher and Little 1998) and their production is likely induced by UVB (Fabacher and Little 1995). Similar UVB-absorbing substances have been found in the mucus of coral reef fishes (Zamzow and Losey 2002). Sommaruga and Garcia-Pichel (1999) found mycosporine-like amino acids in planktonic invertebrates from clear, high-mountain lakes. These substances may be produced by planktonic algae and then ingested by crustaceans. In very cold lakes, where the enzymatic activity of photorepair is low, the tolerance of invertebrates to W B was mainly due to the accumulation of porphyra-334, mycosporine-glycine and shinorine (Rocco et al. 2002). C. Gelatinous Matrices and Foam Nests

Several authors have suggested that the gelatinous matrix surrounding amphibian eggs can effectively reduce UVB transmission through the egg mass (Beudt 1930; Gurdon 1960; Grant and Licht 1995; Licht 2003). For example, pieces of jelly 3 mm thick in Bufo americanus, Rana aurora and R . sylvatica absorbed 6-14% of UVB (Grant and Licht 1995). However, there are significant interspecific differences in the composition of the acellular jelly matrix surrounding eggs and in the amount of UVB absorbed by the jellies. The thickness of the jelly mass may influence photoprotection in eggs but it is important to note that the WB-absorbance capacity of some thin jellies can be higher that those of some thicker masses. For example, Hyla regilla egg masses have a relatively thin jelly coat compared to those of Rana aurora but the jelly surrounding H. regilla egg masses is about twice as absorbent of UVB than is that of R. aurora (Ovaska et al. 1997). Some gelatinous coats are sticky and soil particles or other debris attaches to them and covers the surface of the mass, protecting the eggs within from UVB (Licht 2003). Obviously the jelly matrix surrounding eggs cannot fully protect them from the harmful effects of UVB radiation. Field experiments leaving the jelly matrix intact have shown that the eggs and embryos of many amphibian species are vulnerable to UVB radiation (Blaustein et al. 1998, 2001a). The jelly coat is just one of several factors that help protect developing embryos from W B . Rasanen et al. (2003) conducted two independent factorial laboratory experiments employing three different UVB treatments (no UVB, normal and enhanced levels) and degrees of jelly-removal (control, modified and completely removed). They found that modification of the jelly, or even its removal, from Rana temporah eggs did not increase susceptibility of the embryos to UVB radiation. Eggs of this species are, however, relatively tolerant of UVB and it was suggested that sun screening and/or activity of enzymatic photorepair systems efficiently protect embryos of this species from high UVB exposure (also see Crump et al. 1999a). The jelly of most amphibians is transparent but a few salamanders can produce opaque jelly that may more efficiently protect embryos from UVB. For example, some populations

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3131

of Ambystomu muculatum synthesize a glycoprotein that crystallizes in the oviduct wall and gives the aquatic egg masses a milky-white coloration (Hardy and Lucas 1991). Foam nests produced by many tropical anurans are usually milky or opaque and may provide efficient photoprotection to embryos. For many species, however, eggs in the foam nests can be exposed to UVB. Melanophores occur in eggs of species that oviposit in exposed locations while eggs deposited in hidden locations lack melanophores (Heyer 1969). Some species produce transparent foam that apparently does not shield the eggs from UVB. Species that produce foam nests inhabit mostly tropical areas where ambient levels of UVB are relatively high (Duellman and Trueb 1994; Amphibia Web 2004). Most of these species, however, breed in dense tropical forests where the canopy protects nests from intense solar radiation and UVB. Additionally, many species breed in the wet season and at high temperatures that favour quick embryonic development and reduced exposure time to UVB. D. Other Photoprotective Systems

The ability to detect UVB can help sensitive amphibians avoid areas with high levels. Ambystomatid salamanders have an effective photoreceptive mechanism in their eyes. Deutschlander and Phillips (1995) found and characterized UVA-sensitive cones in the eyes of Ambystoma mexicanum. Some freshwater fishes can also detect UVB at several developmental stages (Losey et al. 1999; Flamarique 2000). Nagl and Hofer (1997) suggested that these mechanisms may be shared by other amphibians and are used to detect and escape harmful levels of UVB. Further research is necessary to demonstrate whether amphibians have the ability to detect different levels of UVB and avoid areas with the most harmful intensities. Nagl and Hofer (1997) observed that the larvae of alpine newts (Triturus alpestris) exposed to sublethal levels of UVB significantly increased their skin thickness. This histological response was also observed under lethal levels just prior to death and could be a result of UVB damage to dermal cells. Alpine newt larvae and other amphibians may, however, use skin thickening as a defence against UVB exposure. This modification of the epidermis has also been observed in other vertebrate species as one of the most effective photoprotective mechanisms against UVB (Menter et al. 1992). Epel et al. (1999) described the use of extra-embryonic cells to shield tunicate embryos from potentially harmful ultraviolet-A and UVB radiation. Some species of frogs lay nonfertile eggs over the fertilized ones, thereby increasing hatching success. There is no evidence, however, for a photoprotective capacity for this behaviour. In globular masses the upper layers of eggs could shade the central and lower ones from UVB. For example, Ambystoma gracile embryos are sensitive to UVB (Blaustein et al. 1995) and exposure to it caused nearly total mortality in small portions of the egg mass with jelly completely covering the eggs. However, large clutches of this species can have a diameter of up to 15 cm and it is likely that inner or lower eggs have lower exposure to UVB than do those from the upper part. Marco and Blaustein (1998) found a high survival of Ambystoma gracile eggs in masses that had become naturally stranded in the field and which were fully exposed to the air. In that study, however, when eggs became exposed to the air, symbiotic algal growth inside the egg masses was enhanced and the green algae not only provided oxygen to embryos and helped remove nitrogenous wastes from the eggs, but also protected them from UVB (Marco and Blaustein 2000). In a later study, Marco (2001) exposed intact A. gracile egg masses to the air from the beginning of embryonic development when algal growth was not detectable. In this experiment, most of the eggs in the upper part of the mass died whereas most of the eggs from other parts of the clutches survived until hatching. In this study there was no control for UVB exposure and the death of embryos could not be conclusively attributed to UVB exposure. Many other substances may protect against damage from UVB. For example, the addition of L-cysteine, ascorbic acid, reduced glutathione, L-tryptophan and sodium pyruvate to cell cultures exposed to UVB caused a significant increase in survival and growth

3132

AMPHIBIAN BIOLOGY

(Tyagi et al. 2003). Presence of these chemicals in the natural habitat or inside the cells of living organisms may partially protect from UVB or assist in the repair of damage caused by UVB (Tyagi et al. 2003). UV radiation can damage the proteins necessary for proper metabolic hnction (Buma et al. 2003). Heat shock proteins (HSPs) may be involved in both protein protection and in restructuring after denaturation (Lindquist 1986). Proteins closely related to heat shock proteins are expressed in normal cells at low levels (Lindquist 1986). These proteins, called constitutive heat shock proteins, serve as molecular chaperones for newly synthesized proteins. When a cell or organism undergoes physiological stress caused by extrinsic factors such as increased temperature, disease, chemical contamination or UV radiation, heat shock protein synthesis is up-regulated (Lindquist 1986; Sanders 1993). In particular, heat shock protein 70 (HSP-70) is a highly conserved HSP produced in response to environmental stress in all organisms studied (Lindquist 1986, 1988; Sanders 1993). The constitutive isoform of this protein binds to target proteins to facilitate and guide protein folding, transport and repair. During physiological stress, HSP-70 binds to pre-ribosomes and other protein complexes, thereby helping protect them from denaturation (Lindquist 1988; Gething and Sambrook 1992). HSP-70 may also function in repair of denatured proteins by breaking up aggregates and allowing proteins to refold in their proper conformation (Ellis 1990; Gaitanaris et al. 1990). Mammalian cell lines produce HSP-70 in response to UV irradiation (Zhou et al. 1998). Few data are published on the role of HSP-70 in response to UV radiation in intact organisms (Sanders 1993 and references therein; Kane and Maytin 1995). VI. SYNERGISM OF W RADIATION WITH OTHER FACTORS During the past few decades it has become especially apparent that multiple stressors are affecting amphibians simultaneously in the field and that the effects of individual factors can be additive (Blaustein and Kiesecker 2002). Anthropogenic stressors may add their effects to natural factors and the global impact could affect amphibian recruitment. There is also evidence of the negative impact of combinations of stressors that alone are not usually harmful to amphibians. Numerous studies have shown that UVB radiation interacts synergistically with contaminants and pathogens (Blaustein et al. 2001a, 2003a; Blaustein and Kiesecker 2002; specific examples are Kiesecker and Blaustein 1995; Long et al. 1995; Zaga et al. 1998; Hatch and Blaustein 2000). Complex dynamics among weather patterns, disease and UVB radiation have recently been illustrated (Pounds et al. 1999, 2006; Kiesecker et al. 2001). The negative effects of UVB on amphibians in the field may be greatly influenced by the presence and the concentration or density of factors that may interact synergistically with UVB. The absence of effects in a specific field location may be due to the lack of factors that would ordinarily cause synergistic interactions. Results from laboratory or openair experiments where amphibians develop in tap water or in reconstituted water and where the only stressor is UVB (for example, Cummins et al. 1999) can underestimate the real impact of UVB on these animals. Most studies demonstrating a significant negative effect of ambient UVB on amphibian embryos have been conducted in pond water where the tested amphibians were actually developing (for example, Blaustein et al. 199413; Anzalone et al. 1998; Lizana and Pedraza 1998; Hakkinen et al. 2001; Little et al. 2003). However, most studies that suggest that ambient UVB is not causing a significant effect on amphibians have been conducted using tap water, reconstituted or filtered water with little possibility of synergistic agents (e.g., Grant and Licht 1995; Cummins et al. 1999; Smith et al. 2000b; Starnes et al. 2000; Licht 2003). A. W and Acidification Low pH may negatively influence the activity of repair systems whose DNA has been damaged (Sancar and Sancar 1988). Tadpoles of leopard frogs (Rana pipiens) exposed to a

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3133

combination of UVB and low pH suffered significant larval mortality (Long et al. 1995). In the same experiment, however, tadpoles exposed to the same level of both stressors separately did not suffer any detectable effect (Long et al. 1995). UVB levels were similar to those expected at high elevations from temperate zones during amphibian breeding and the levels of pH (4.5-5.0) were similar to those found in disturbed areas. In field experiments, no evidence was found for reduced hatchability or increased frequency of developmental anomalies in Rana arvalis embryos exposed to ambient levels of UVB compared with embryos shielded from UVB radiation (Pahkala et al. 2001b). Although moderately low pH (5.0) reduced hatchability, increased frequency of developmental anomalies and reduced early embryonic growth in R . arualzri, there was no evidence for synergistic effects of pH and UVB on any of these traits (Pahkala et al. 2001b). Similar lack of synergism was found by Pahkala et al. (2000) studying Rana temporaria embryos. In the laboratory, Hatch and Blaustein (2000) tested potential synergism between moderate acidity and UVB in tadpoles. Rana cascadae tadpoles were sensitive to UVB and pH and suffered increased mortality and reduced larval activity. There was, however, no synergistic interaction between these factors. The addition of nitrate to the water (up to 20 mg/L) significantly increased the impact of the combination of UVB and acidicity (Hatch and Blaustein 2000).

B. UV and Contaminants The toxicity of two contaminants acting together may have an additive effect. In some cases, the toxicity of one contaminant may be enhanced by the presence of the other or, alternatively, one of the contaminants may inhibit the detoxifying capacity of the exposed organisms, thereby multiplying the toxic response. It is also possible that one contaminant may reduce the toxicity of the other and the combined effect of simultaneous exposure to both may have less effect than would exposure to a single stressor. The simultaneous exposure to more than two contaminants can cause even more complex interactions. UVB is an environmental stressor of amphibians that may also be considered as a contaminant due to the enhanced levels that the earth's surface is receiving. There is experimental evidence that suggests synergistic effects between UVB and some environmental contaminants. Polycyclic aromatic hydrocarbons (PAHs) are common contaminants of terrestrial and aquatic ecosystems and are frequently found in shallow water selected by many amphibians. These contaminants may become toxic, or substantially more toxic, upon co-exposure with UV light (Arfsten et al. 1996). Amphibians that are exposed to PAHs, or that bioaccumulate small amounts of them, can suffer a high mortality rate when exposed to UVA. For example, Rana pipiens tadpoles exposed to anthracene alone (0.025 mg/L) did not suffer mortality but, when sunlight was present, anthracene was highly toxic (Kagan et al. 1984). Fernhndez and L'Haridon (1992) found synergism between several PAHs and UVA on larvae of the newt Pleurodeles waltl. In that study, benzanthracene and some derivatives alone were toxic to amphibians but their toxicity increased in some cases when coupled with exposure to UVA. Fernhndez and L'Haridon (1994) also found a synergistic effect of UVA and benzopyrene on embryos and larvae of Pleurodeles waltl. When amphibians at two developmental stages were exposed to these factors alone there were no toxic effects but, when the contaminant and light acted together, there were severe cytotoxic and genotoxic effects. Similar results were found when oil refinery emuent and UVA were tested (Fernhndez and L'Haridon 1994). Walker et al. (1998) found a significant positive interaction between fluoranthene and UVA on Rana catesbeiana tadpoles. Fluoranthene had phototoxic effects on the locomotory performance and activity level of tadpoles. It also caused necrosis and structural alterations of the skin (Walker et al. 1998). Hatch and Burton (1998) demonstrated in the laboratory and in outdoor exposures that there was a positive interaction between UVB and fluoranthene in three amphibian species. Newly hatched larvae were more sensitive than were embryos to the combination of both stressors and Xenopous laevis showed the highest rate of deformities when exposed to photoactivated fluoranthene (Hatch and Burton

3134

AMPHIBIAN BIOLOGY

1998). Monson et al. (1999) exposed Rana pipiens larvae to fluoranthene for two days. Larvae accumulated the contaminant in direct proportion to its concentration and survivors were able to detoxify in a few days. When larvae were exposed to as low as 2.18 ppb (bioaccumulation of 3.53 pglg) of fluoranthene and were also exposed to environmental levels of UVA, however, they suffered significant mortality. At that fluoranthene concentration but with a low exposure to UVA, there was no larval mortality (Monson et al. 1999). The combined exposure of Rana cascadae larvae to non-toxic levels of UVB (9-1 1 CLW/ cm2) and nitrate (5 and 20 mg/L) caused a significant decrease in activity level compared to that of controls or to larvae exposed to each stressor alone (Hatch and Blaustein 2000). When tadpoles were also exposed to water with a pH of 5, mortality increased compared to controls (Hatch and Blaustein 2000). The synergistic effect between nitrate and UVB may have a severe impact on tadpole survival and is exacerbated by slight acidification of the water. These factors alone, and at the levels used, do not cause overt damage to tadpoles. Hatch and Blaustein (2000) suggested that tadpoles had a reduced ability to cope with the simultaneous combination of UVB, low pH and high nitrate. Hyla regilla from populations at both high and low elevations also suffered a significant impact when exposed to a combination of nitrate and UVB (Hatch and Blaustein 2003). Those from highland areas suffered a significant increase in mortality when exposed to the combination of both stressors. In a lowland population, both stressors together altered larval development and produced smaller individuals (Hatch and Blaustein 2003). Crump et al. (2002) exposed newly hatched Rana pipiens tadpoles to subambient levels of UVB radiation and octylphenol, a commonly found estrogenic endocrine-disrupting chemical. They found no significant effects of both stressors alone but tadpoles from one of the octylphenol/UVB combinations had greater body weight and earlier hind limb emergence than did controls. The combination of both stressors altered tadpole growth and the expression of hypothalamic genes (Crump et al. 2002). Embryos (<24 h old) of Rana temporaria exposed to three concentrations (10, 100 or 1 000 micrograms/L) of bisphenol A showed a significant decrease in survival when they were simultaneously exposed to UVB radiation. UVB radiation alone or bisphenol controls alone had no significant effect on frog embryos (Koponen and Kukkonen 2002). Zaga et al. (1998) exposed Xenopus laevis and Hyla versicolor to combinations of sublethal levels of the pesticide carbaryl and ambient levels of UVB and found significant positive interactions between these agents. Very low intensity of UVB was able to photoactivate carbaryl and the toxicity of this compound was increased by 10-fold in the presence of ambient levels of UVB for all species and stages tested. Irradiation of carbaryl with UVB induced 100% mortality in X. laevis embryos after 24 hours of exposure; non-irradiated carbaryl did not cause any mortality after 96 hours. UVB photomodifies the carbaryl molecule in the water, increasing the concentration of highly toxic photoproduct. Embryos were more tolerant to the interaction of UVB and carbaryl than were tadpoles. The ingestion of carbaryl by tadpoles or the protective influence of egg envelopes may explain these ontogenetic differences. The toxicity of this and other similar compounds to amphibians should be tested in the absence of UVB (Zaga et al. 1998). LaClair et al. (1998) investigated the interaction between UVB and methoprene, an insect growth-regulator that causes limb malformations in amphibians. The number of body abnormalities of Xenopus laevls was significantly affected by some of the extracted breakdown products of methoprene exposed to sunlight for four days. Rana pipiens larvae suffered a significant number of limb malformations when exposed to UVB but methoprene did not enhance that effect (Ankley et al. 1998). Calfee and Little (2003) found an interactive effect of UVB and fire-retardant chemicals. UVB levels in their study were within tolerance limits but significantly increased the mortality to six chemicals in boreal toads and southern leopard frogs, even when the optical clarity of the water was low (Calfee and Little 2003). The possibility of photoinduced toxicity of chemicals should be considered when conducting environmental risk assessments for amphibians (Blaustein et al. 2003a).

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3135

C. W B and Pathogens Sublethal levels of UVB can impair or suppress the immune response immediately after exposure to the sun o r for some time after exposure (Vermeer and Hurks 1994). For example, one study in rodents showed that exposure to UVB for 25 minutes at 0.27 W/m2 caused about a 50% immune suppression. Infection by the oomycete Saprolegnia causes significant embryonic mortality in amphibian populations (Blaustein et al. 1994b). Solar UVB significantly increases the lethal effects of the pathogen in nature (Kiesecker and Blaustein 1995; Kiesecker et al. 2001). In field experiments, ambient UVB radiation, together with Saprolegniu, enhances mortality in Bufo boreas and Rana cascadae embryos but not in embryos of Hyla regilk (Kiesecker and Blaustein 1995). Embryos of Hyla cadaverina and Taricha torosa that died under the effects of UVB were engrossed with fungus, although it was not determined whether fungal infection affected live embryos or merely colonized them after death (Anzalone et al. 1998). Egg-laying behaviour may affect the rate of infection of amphibian eggs as well as the synergistic interaction with UVB (Kiesecker and Blaustein 1997; Green 1999). Species that form large communal egg masses are more prone to transmission of disease than are solitary breeders (Kiesecker and Blaustein 1997). Myositis associated with infection by Ichthyophonus-like organisms has recently been diagnosed in wild amphibians (Mikaelian et al. 2000). The fungus Basidiobolus ranarum has also been related to naturally occurring fatal mycotic dermatitis in free-ranging amphibians (Taylor et al. 1999). Chytridiomycosis caused by Batrachochytrium dendrobatid& also is a fatal emerging disease that has been recently related to amphibian declines in Europe, America and Australia (Berger et al. 1998; Bosch et al. 2001; Bradley et al. 2002; Lips et al. 2003; Parris and Beaudoin 2004). Some Iridoviruses (Ranavirus and Lymphocystivirus) have recently become noted as a significant cause of disease in amphibians and fishes and could have an impact on populations (Daszak et al. 1999; Chinchar 2002). Environmental changes, along with increasing exposure to UVB, could lead to immune system suppression in host species that could enhance the impact of these pathogens in amphibians. Field studies combining exposure to UVB and to different pathogens are necessary for understanding the overall impact of such interactions on amphibians. VII. GENERAL CONCLUSIONS Blaustein et al. (1998) and Reaser and Blaustein (2005) discussed some problems inherent in assessing the affects of UV-B radiation on amphibians. They also discussed the importance of employing experiments in the field. Their conclusions are briefly summarized here and some additional information presented. A. Factors Affecting Exposure to Ultraviolet Radiation UVB levels vary temporally and spatially, and change with weather pattern, cloud cover, water flow, and depth and turbidity of water. UV levels may also change with latitude and elevation. Thus, amphibians are exposed to constantly changing levels of UVB radiation. This variability makes it difficult to compare results between different studies, especially those conducted in different regions and employing different methods. Even within a single, well-designed experiment, natural variability is expected. For example, in field experiments assessing the effects of UVB on hatching success, when differences between UV-exposed and non-exposed treatments are sufficiently great, it is apparent that UVB, regardless of the variability of exposure, affects hatching success. B. Natural Selection

Why are the eggs, embryos, larvae and adults of some species affected more significantly by UVB than are those of other species? One possible reason is that, over evolutionary time, there has been strong selection for the evolution of behavioural, ecological and physiological mechanisms that counteracted the harmful effects of UVB. For example, in

3136

AMPHIBIAN BIOLOGY

some species, there may have been selection for oviposition in areas with less exposure to UVB radiation. Morphological and molecular mechanisms may also help amphibians cope with exposure to UVB radiation. For example, the composition and thickness of the jelly coat that surrounds amphibian eggs may limit UVB exposure to developing embryos. Darker egg pigmentation, particles attached to the jelly matrix or algae within the capsular-jelly may also impede UVB transmission. In addition to mechanisms that help limit an amphibian's initial exposure to UVB radiation, there are several ways UV-induced DNA damage can be repaired once it has occurred. Photoreactivation, using the enzyme photolyase, is of prime importance in repairing UVB damage. Of the limited number of species examined, those species with the highest levels of photolyase are the most resistant to UVB radiation in field experiments (e.g., Blaustein et al. 1994a, 1996, 1999). There are, however, other mechanisms of repairing UV-induced damage and more than one mechanism may be employed. It is becoming clear that intraspecific variation in the ability to cope with UVB exposure may be due to intraspecific differences in how individuals repair DNA damage at the molecular level. C. Interspecific Differences in Resistance to UVB Radiation as Depicted by the Results of Field Experiments

Most published field studies on the effects of UV on amphibians have addressed the question: Does ambient UVB radiation damage amphibian embryos? Experimental designs were geared for addressing that particular question. Field experiments can be a rigorous method for assessing environmental damage by specific agents. If properly designed, all factors vary naturally and simultaneously between experimental and control treatments except for the variable(s) of interest. Adequate controls are necessary. Employing an adequate number of replicates for each treatment will help ensure that results are not unique to a particular series of treatments. Unfortunately, these aspects of field experimentation have been misunderstood by some, leading to some confusion in the literature (e.g., Licht 2003). Some investigators have found their species of study to be resistant to ambient UVB radiation (e.g., Blaustein et al. 1996, 1999; Ovaska et al. 1997; Corn 1998). Others, within a single study and using identical methods to examine a number of species, have found that some species are resistant whereas others are highly susceptible to ambient levels of UVB radiation (e.g., Blaustein et al. 1994a; Anzalone et al. 1998; Lizana and Pedraza 1998; Broomhall et al. 1999). There is no contradiction in results between studies that do and do not show effects of UVB radiation on amphibians. There are, of course, interspecific differences in resistance to UVB radiation just as there are interspecific differences in responses to other environmental factors such as temperature and pH. Even populations of the same species may show differences in susceptibility to UVB radiation. D. Measuring UVB in the Field Measurements of UVB radiation are not necessary to document a detrimental effect on amphibian hatching rates. They are not necessary to answer the question: Does ambient UVB radiation damage amphibian embryos? Measuring UV levels is necessary, however, when addressing questions concerning the precise levels of UVB radiation that cause damage to a particular amphibian. This is an important consideration but measuring UVB does not alter the results or interpretation of experiments designed to investigate if UVB radiation decreases hatching success. E. The Role of W B Radiation in Amphibian Population Declines UVB radiation is affecting the eggs and embryos of several species of amphibians in widely scattered locales around the world. Studies have also shown that UVB can kill larval and adult amphibians and cause various kinds of sublethal damage. Continued mortality of early life stages may eventually lead to population decline but the effects of continued

MARC0 ET AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS

3137

mortality at the egg and embryo stage may not be observed for many years, especially in long-lived species (Biek et al. 2002; Vonesh and de la Cruz 2002). Those species that lay their eggs in shallow water exposed to sunlight and which have poor ability to repair UV-induced DNA damage are at most risk but the extent of that risk is unknown. UVB radiation is obviously not the only factor contributing to amphibian population declines. It would be an unlikely factor in the declines of species that lay their eggs under logs, under litter, in crevices, in deep water or under a dense forest canopy. It would be an unlikely factor in larvae or adults that inhabit heavily forested regions or that remain away from sunlight. Moreover, even the eggs, larvae and adult forms of some species that are found in open sunlight may be unaffected by UVB radiation if they have molecular and morphological mechanisms that counteract the harmful effects of UVB. Finally, it should be emphasized that increases in UVB radiation are not necessary to demonstrate that UVB is affecting amphibians in nature. Even if levels of UVB remain constant or are only slightly increasing, synergistic interactions with UV and other agents could be damaging to amphibians in ways not previously experienced. VIII. REFERENCES Adams, M. J., Schindler, D. E. and Bury, R. B., 2001. Association of amphibians with attenuation of ultraviolet-B radiation in montane ponds. Oecologia 128: 519-525. Adams, M. J., Hossack, B. R., Knapp, R. A., Corn, l? S., Diamond, S. A., Trenham, F! C. and Fagre, D. B., 2005. Distribution patterns of lentic-breeding amphibians in relation to ultraviolet radiation exposure in western North America. Ecosystems 8: 488-500. Amphibia Web, 2004 (accessed). Information on amphibian biology and conservation. [web application]. 2004. Berkeley, California: AmphibiaWeb. http:/l AnimalDiversity Web, 2004 (accessed). Interagency Education Research Initiative, the Homeland Foundation and the University of Michigan Museum of Zoology. http://animaldiversity.ummz. umich.edu/sitelindex.html Ankley, G. T., Diamond, S. A,, Tietge, J. E., Holocombe, G. W., Jensen, K. M., DeFoe, D. L. and Peterson, R., 2002. Assessment of the risk of solar ultraviolet radiation to amphibians: I. Dose-dependent induction of hindlimb malformations in the Northern Leopard Frog (Rana pipiens). Enuiron. Sci. Technol. 36: 2853-2858. Ankley, G. T., Tietge, J. E., DeFoe, D. L., Jensen, K. M., Holcombe, G. W., Durhan, E. J. and Diamond, S. A,, 1998. Effects of ultraviolet light and methoprene on survival and development of Rana pipiens. Enuiron. Toxicol. Cham. 17: 2530-2542. Anzalone, C. R., Kats, L. B. and Gordon, M. S., 1998. Effects of solar W - B radiation on embryonic development in three species of lower latitude and lower elevation amphibians. Cons. Biol. 12: 646-653. Arfsten, D. l?, Schaeffer, D. J. and Mulveny, D. C., 1996. The effects of near ultraviolet radiation on the toxic effects of polycyclic aromatic hydrocarbons in animals and plants: A review. Ecotoxicol. Enuiron. Safety 33: 1-24. Barker, D., Dixon, K., Medrano, E. E., Smalara, D., Mitchell, D., Im, S., Babcock, G. and Abdel-Malek, Z. A,, 1995. Comparison of the responses of human melanocytes with different melanin contents to ultraviolet B irradiation. Cancer Res. 55: 4041-4046. Baud, D. R. and Beck, M. L., 2005. Interactive effects of UV-B and copper on Spring Peeper tadpoles (Pseudacris crucqer). Southeast. Nut. 4: 15-22.

Beebee, T. J. C., 1995. Amphibian breeding and climate. Nature 374: 219-220. Beebee, T. J. C., 2002. Amphibian phenology and climate change. Cons. Biol. 16: 1454. Beiswenger, R. E., 1977. Die1 patterns of aggregative behavior in tadpoles of Bufo americanus, in relation to light and temperature. Ecology 58: 98-108. Belden, L. K. and Blaustein, A. R., 2002a. Population differences in sensitivity to W - B Radiation for larval long-toed salamanders. Ecology 83: 1586-1590. Belden, L. K. and Blaustein, A. R., 2002b. Exposure of red-legged frog embryos to ambient UV-B in the field negatively affects larval growth and development. Oecologia 130: 551-554. Belden, L. K. and Blaustein, A. R., 2002c. UV-B induced skin darkening in larval salamanders does not prevent sublethal effects of exposure on growth. Copeia 2002: 748-754. Belden, L. K., Wildy, E. L. and Blaustein, A. R., 2000. Growth, survival, and behaviour of larval long-toed salamanders (Ambystoma rnacrodactylum) exposed to ambient levels of UV-B radiation. J. 2001. (Land.). 251: 473-479. Belden, L. K., Moore, I. T., Mason, R. T., Wingfield, J. C., Blaustein, A. R., 2003. Survival, the hormonal stress response and UV-B avoidance in Cascades Frog tadpoles (Rana cascadae) exposed to W - B radiation. Funct. Ecol. 17: 409-416. Bennett, A. T. D. and Cuthill, I. C., 1994. Ultraviolet vision in birds -what is its function. Viion Res. 34: 1471-1478. Bennett, A. T. D., Cuthill, I. C., Partridge, J. C., Maier, E. J., 1996. Ultraviolet vision and mate choice in zebra finches. Nature 380: 433-435. Berger, L., Speare, R., Daszak, P., Green, D. E., Cunningham, A. A., Goggin, C. L., Slocombe, R., Ragan, M. A,, Hyatt, A. D., McDonald, K. R., Hines, H. B., Lips, K. R., Marantelli, G. and Parkes, H., 1998. Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proc. Nut. Acad. Sci. USA 95: 9031-9036.

3138

AMPHIBIAN BIOLOGY

Berrill, M. and Lean, D. R. S., 1998. Problems in conducting in-situ experiments to evaluate the influence of UV-B radiation on amphibian eggs and larvae. Pp. 2 1-30 in "Environmental Toxicology and Risk Assesment", ed by E. E. Little, A. J . DeLonay and B. M. Greenberg. Vol. 7. Ame?: Sac. Testing Mat. Special Tech. Publ. No. 1333. West Conshohocken. Berrocal-Tito, G., Sametz-Baron, L., Eichenberg, K., Horwitz, B. A,, Herrera-Estrella, A,, 1999. Rapid blue light regulation of a Trichoderma ha~zianum photolyase gene. J. Biol. Chem. 274: 14288-14294. Beudt, E. L., 1930. Der einfluss der lichtes der quartzqueck-silber lampe auf der furschungs-und larvenstadien verschieden Amphibien. Zool. Jahrb. Abt. Syst. Oekol. Geogr: Tiere 47: 623-684. Biek, R., Funk, W. C., Maxell, B. A. and Mdls, L. S., 2002. What is missing in amphibian decline research: Insights from ecological sensitivity analysis. Cons. Biol. 16: 728-734. Blaustein, A. R. and Belden, L. K., 2003. Amphibian defenses against ultraviolet-B radiation. Evol. Devel. 5: 89-97. Blaustein, A. R. and Kiesecker, J. M., 1997. The effects of ultraviolet-B radiation on amphibians in natural ecosystems. Pp. 175-188 in "The Effects of Ozone Depletion on Aquatic Ecosystems", ed by D.-I! Hader. R. G. Landes, Austin. Blaustein, A. R. and Kiesecker, J. M., 2002. Complexity in conservation: Lessons from the global decline of amphibian populations. Ecol. Lett. 5: 597-608. Blaustein, A. R., Romansic, J. M. and Scheessele, E. A., 2005. Ambient levels of ultraviolet-B radiation cause mortality in juvenile western toads, Bufo boreas. Amer: Midl. Nut. 154: 375-382. Blaustein, A. R., Chivers, D. E, Kats, L. B. and Kiesecker, J. M., 2000. Effects of ultraviolet radiation on locomotion and orientation in Roughskin Newts (Taricha granulosa). Ethology 106: 227-234. Blaustein, A. R., Hoffman, E D., Kiesecker, J. M. and Hays, J. B., 1996. DNA repair and resistance to solar UV-B radiation in eggs of the red-legged frog. Cons. Biol. 10: 1398-1402. Blaustein, A. R., Hokit, D. G., O'Hara, R. K. and Holt, R. A,, 1994. Pathogenic fungus contributes to amphibian losses in the Pacific Northwest. Biol. Cons. 67: 251-254. Blaustein, A. R., Kiesecker, J. M., Chivers, D. P and Anthony, R. G., 1997. Ambient UV-B radiation causes deformities in amphibian embryos. Proc. Nat. Acad. Sci. U S A 94: 13735-13737. Blaustein, A. R., Romansic, J. M., Kiesecker, J. M. and Hatch, A. C., 2003a. Ultraviolet radiation, toxic chemicals and amphibian population declines. Diver. Distrib. 9: 123-140.

Blaustein, A. R., Hoffman, E D., Hokit, D. G., Kiesecker, J. M., Walls, S. C. and Hays, J. B., 1994. UV-repair and resistance to solar UV-B in amphibian eggs: A link to amphibian declines? Proc. Nut. Acad. Sci. USA 91: 1791-1795. Blaustein, A. R., Root, T L., Kiesecker, J. M., Belden, L. K., Olson, D. H. and Green, D. M., 2003b. Amphibian breeding and climate change: reply to corn. Cons. Biol. 17: 626-627. Blaustein, A. R., Kiesecker, J . M., Chivers, D. E, Hokit, D. G., Marco, A., Belden, L. K. and Hatch, A,, 1998. Effects of ultraviolet radiation on amphibians: Field experiments. Amer: Zool. 38: 799-812. Blaustein, A. R., Belden, L. K., Hatch, A. C., Kats, L. B., Hoffman, E D., Hays, J. B., Marco, A,, Chivers, D. I? and Kiesecker, J. M., 2001a. Ultraviolet radiation and amphibians. Pp. 63-79 in "Ecosystems, Evolution and Ultraviolet Radiation", ed by C. S. Cockell and A. R. Blaustein. Springer, New York. Blaustein, A. R., Hays, J. B., Hoffman, l? D., Chivers, D. E, Kiesecker, J. M., Leonard, W. E, Marco, A., Olson, D. H., Reaser, J. K. and Anthony, R. G., 1999. DNA repair and resistance to UV-B radiation in the western spotted frog complex, Rana pretiosa and R . luteiventris. Ecol. A@pl. 9: 1100-1105. Blaustein, A. R., Han, B., Fasy, B., Romansic, J., Scheessele, E. A., Anthony, R. G., Marco, A,, Chivers, D. F!, Belden, L. K., Kiesecker, J. M., Garcia, T., Lizana, M. and Kats, L. B., 2004. Variable breeding phenology affects the exposure of amphibian embryos to ultraviolet radiation and optical characteristics of natural waters protect amphibians from UV-B in the U.S. Pacific Northwest: Comment. Ecology 85: 1747-1754. Blum, H. F. and Matthews, M. R., 1950. Photorecovery after ultraviolet radiation in amphibian larvae. Biol. Bull. 99: 330. Blum, H. F. and Matthews, M. R., 1952. Photorecovery from the effects of ultraviolet radiation in salamander larvae. J. Cell. Camp. Physiol. 39: 57-72. Blumthaler, M. and Ambach, W., 1990. Indication of increasing solar ultraviolet-b radiation flux in alpine regions. Science 248: 206-208. Blumthaler, M., Ambach, W. and Ellinger, R., 1997. Increase in solar UV radiation with altitude. J. Photochem. Photobiol. B-Biol. 39: 130-134. Blumthaler, M., Ambach, W. and Salzgeber, M., 1994. Effects of cloudiness on global and diffuse uv irradiance in a high-mountain area. Theoret. Appl. Climat. 50: 23-30. Bosch, J., Martinez-Solano, I. and Garcia-Paris, M., 2001. Evidence of a chytrid fungus infection involved in the decline of the common midwife toad (Alytes obstetricans) in protected areas of central Spain. Biol. Cons. 97: 331-337. Bothwell, M. L., Sherbot, D. M. J. and Pollock, C. M., 1994. Ecosystem response to solar ultraviolet-b radiation - influence of trophic-level interactions. Science 265: 97-100.

Blaustein, A. R., Edmond, B., Kiesecker, J. M., Beatty, J. J. and Hokit, D. J., 1995. Ambient ultraviolet radiation causes mortality in salamander eggs. Ecol. Applic. 5: 740-743.

Bradley, G. A,, Rosen, F! C., Sredl, M. J., Jones, T R. and Longcore, J. E., 2002. Chytridiomycosis in native Arizona frogs. J. Wildl. Dis. 38: 206-212.

Blaustein, A. R., Belden, L. K., Olson, D. H., Green, D. L., Root, T. L. and Kiesecker, J. M., 2001b. Amphibian breeding and climate change. Cons. Biol. 15: 1804-1809.

Bridges, C. M. and Boone, M. D., 2003. The interactive effects of UV-B and insecticide exposure on tadpole survival, growth and development. Biol. Cons. 113: 49-54.

MARC0 ET AL. : ULTRAVIOLET-B RADIATION AND AMPHIBIANS Broomhall, S. D., Osborne, W. and Cunningham, R., 2000. Comparative effects of ambient ultraviolet-B (UV-B) radiation on two sympatric species of Australian frogs. Cons. Biol. 14: 420427. Bruggeman, D. J., Bantle, J. A. and Goad, C., 1998. Linking teratogenesis, growth, and DNA photodamage to artificial ultraviolet B radiation in Xenopus laeuis larvae. Enuiron. Toxicol. Chem. 17: 2 114-2 121.

3139

Cummins, C. P., Greenslade, P. D. and McLeod, A. R., 1999. A test of the effect of supplemental UV-B radiation on the common frog, Rana temporaria L., during embryonic development. Global Change Biol. 5: 471-479. Daszak, P., Berger, L., Cunningham, A. A,, Hyatt, A. D., Green, D. E. and Speare, R., 1999. Emerging infectious diseases and amphibian population declines. Emerg. Infect. Diseas. 5: 735-748.

Buma, A. G. J., Boelen, I? and Jeffrey, W. H., 2003. UVR-induced DNA damage in aquatic organisms. Pp. 293-327 in "UV Effects in Aquatic Organisms and Ecosystems", ed by E. W. Helbling and H. Zagarese. Royal Society of Chemistry, Cambridge.

Davidson, C., Shaffer, H. B. and Jennings, M. R., 2001. Declines of the California red-legged frog: climate, UV-B, habitat, and pesticides hypotheses. Ecol. Appli. 11: 464479.

Bruner, M. A,, Shipman, P. A,, Rao, M. and Bantle, J. A,, 2002. Developmental effects of ambient UV-B light and landfill leachate in Rana blairz and Hyla chrysoscelis. Ecotoxicol. Enuiron. Safety 53: 73-80.

Davidson, C., Shaffer, H. B. and Jennings, M. R., 2002. Spatial tests of the pesticide drift, habitat desuuction, UV-B, and climate-change hypothesis for California amphibian declines. Cons. Biol. 16: 1588-1601.

Calfee, R. D., Little, E. E., 2003. The effects of ultravioletB radiation on the toxicity of fire-fighting chemicals. Enuiron. Toxicol. Chem. 22: 1525-1531.

de Fabo, E., Noonan, F. P. and Frederick, J. E., 1990. Biologically effective doses of sunlight for immune suppression at various latitudes and their relationship to changes in stratospheric ozone. Photochem. Photobiol. 52: 811-817.

Carey, C. and Bryant, C. J., 1995. Possible interrelations among environmental toxicants, amphibian development, and decline of amphibian populations. Enuiron. Health Perspec. 103: 13-17. Carey, C., Cohen, N. and Rollins-Smith, L., 1999. Amphibian declines: an immunological perspective. Develop. Cornpax Zmmun. 23: 459472. Chinchar, V. G., 2002. Ranavimses (family Iridoviridae): emerging cold-blooded killers. Arch. Viral. 147: 447-470. Cockell, C. S., 2001. A photobiological history of earth. Pp. 1-34 in "Ecosystems, Evolution, and Ultraviolet Radiation", ed by C. S. Cockell and A. R. Blaustein. Springer Verlag, New York. Cockell, C. S. and Blaustein, A. R., 2000. "Ultraviolet spring" and the ecological consequences of catastrophic impacts. Ecol. Letters 3: 77-81. A-690. Cockell, C. S. and Knowland, J., 1999. Ultraviolet radiation screening compounds. Biol. Rev. 74: 31 1-435. Cook, J. S. and McGrath, J. R., 1967. Photoreactivatingenzyme activity in metazoa. Proc. Nut. Acad. Sci. USA 58: 1359. Corn, E! S., 1998. Effects of ultraviolet radiation on boreal toads in Colorado. Ecol. Appli. 8: 18-26.

Corn,E! S., 2003. Amphibian breeding and climate change: the importance of snow in the mountains. Cons. Biol. 17: 622-625. Corn, El S. and E. Muths, 2002. Variable breeding phenology affects the exposure of amphibian embryos to ultraviolet radiation. Ecology 83: 2958-2963. Crump, D., Lean, D. and Tmdeau, V. L., 2002. Octylphenol and UV-B radiation alter larval development and hypothalamic gene expression in the leopard frog (Rana pipiens). Enutl. Health Pers. 110: 277-284. Crump, D., Lean, D., Berrill, M., Coulson, D. and Toy, L., 1999b. Spectral irradiance in pond water: Influence of water chemistry. Photochern. Photobiol. 70: 893-90 1. Crump, D., Berrill, M., Coulson, D., Lean, D, McGillvray, L. and Smith, A., 1999a. Sensitivity of amphibian embryos, tadpoles and larvae to enhanced UV-B radiation in natural pond conditions. Can. J. 2001. 77: 1956-1966.

Deutschlander, M. E. and Phdlips, J. B., 1995. Characterization of an ultraviolet photoreception mechanism in the retina of an amphibian, the axolotl (Ambystoma mexicanurn). Neurosci. Lett. 197: 93-96. Diamond, S. A., Peterson, G. S., Tietge, J. E. and Ankley, G. T., 2002. Assessment of the risk of solar ultraviolet radiation to amphibians. Ill. Predictions of impacts in selected northern midwestern wetlands. Enuiron. Sci. Technol. 36: 2866-2874. Diamond, S. A,, Trenham, P. C., Adams, M. J., Hossack, B. R., Knapp, R. A, Stark, S. L., Bradford, D., Corn, E! S., Czarnowski, K., Brooks, E! D., Fagre, D., Breen, B., Detenbeck, N. E. and Tonnessen, K., 2005. Estimated ultraviolet radiation doses in wetlands in six national parks. Ecosystems 8: 462477. Diffey, B. L., 1991. Solar ultraviolet radiation effects on biological systems. Phys. Med. Biol. 36: 299-328. Donahue, W. F., Schindler, D. W., Page, S. J. and Stainton, M. P., 1998. Acid induced changes in DOC quality in an experimental whole-lake manipulation. Enuiron. Sci. Technol. 32: 2954-2960. Duellman, W. E. and Tmeb, L., 1994. "Biology of Amphibians". The Johns Hopkins University Press, Baltimore. DuShane, G. P., 1943. The embryology of vertebrate pigment cells. Part I. Amphibia. Quart. Rev. Biol. 18: 109-127. Elis, R. J., 1990. The molecular chaperon concept. Semin. Cell Biol. 1: 1-9. Epel, D., Hemela, K., Shick, M. and Patton, C., 1999. Development in the floating world: defenses of eggs and embryos against damage from UV radiation. Am. Zool. 39: 271-278. Epperlein, H. H. and Lofberg, J., 1990. The development of the larval pigment patterns in Triturus alpestris and Ambystoma mexicanurn. Adu. Anat. Embryol. Cell Bio2. 118: 1-99. Fabacher, D. L. and Little, E. E., 1995. Skin component may protect fishes from ultraviolet-B radiation. Enuiron. Sci. Poll. Res. 2: 30-32.

3140

AMPHIBIAN BIOLOGY

Fabacher, D. L. and Little, E. E., 1998. Photoprotective substance occurs primarily in outer layers of fish skin. Environ. Sci. Poll. Res. 5: 4-6. Fabacher, D. L., Little, E. E. and Ostrander, G., K., 1999. Tolerance of an albino fish to ultraviolet-B radiation. Enuiron. Sci. Poll. Res. 6: 69-71. Feder, M. E. and Hofmann, G. E., 1999. Heat-shock proteins, molecular chaperones, and the stress response: evolutionary and ecological physiology. Ann. Rev. Physiol. 61: 243-282. Fernsndez, M. and L'Haridon, J., 1992. Influence of lighting conditions on toxicity and genotoxicity of various PAH in the newt invivo.J. Mutat. Res. 298: 31-41. Fernindez, M. and L'Haridon, J., 1994. Effects of light on the cytotoxicity and genotoxicity of benzo(a)pyrene and an oil refinery effluent in the newt. Environ. Molec. Mutagen. 24: 124-136. Fite, K. V., Blaustein, A. R., Bengston, L. and Hewitt, H. E., 1998. Evidence of retinal light damage in R a m cascadae: a declining amphibian species. Copeia 1998: 906-914. Flamarique, I. N., 2000. The ontogeny of ultraviolet sensitivity, cone disappearance and regeneration in the sockeye salmon Oncorhynchus nerka. J. Experim. Biol. 203: 1161-1 172. Forchhammer, M. C., Post, E. and Stenseth, N. C., 1998. Breeding phenology and climate. Nature 391: 29-30. Fraikin, G. Y, Strakhovskaya, M. G. and Rubin, A. B., 2000. Light-induced processes of cell protection against photodamage. Biochem.-Moscow 65: 737-746. France, R., Steedman, R., Lehmann, R. and Peters R., 2000. Landscape modification of DOC concentration in boreal lakes: implications for UV-B sensitivity. Water Air And Soil Poll. 122: 153-162. Friedburg, E. C., Walker, G. C. and Wolfram, S., 1995. "DNA Repair and Mutagenesis". ASM Press, Washington, D.C. Gaitanaris, G. A,, Papavassiliou, A. G., Rubock, F,! Silverstein, S. J. and Gottesman, M. E., 1990. Renaturation of denatured repressor requires heat shock proteins. Cell 61: 1013-1020.

Hakkinen, J., Pasanen, S. and Kukkonen, J. V. K., 2001. The effects of solar UV-B radiation on embryonic mortality and development in three boreal anurans (Rana temporaria, Rana arvalis, and Bufo bufo). Chemosphere 44: 44 1 4 4 6 . Ham, W. T., 1983. Ocular hazards of light-sources review of current knowledge.J. Occup. Environ. Med. 25: 101-103. Hardy, L. M. and Lucas, M. C., 1991. A crystalline protein is responsible for dimorphic egg jellies in the spotted salamander, Ambystoma maculatum (Shaw)(Caudata: Ambystomatidae). Compal: Biochem. Physio. 100: 653-660. Hatch, A. C. and Blaustein, A. R., 2000. Combined effects of UV-B, nitrate, and low pH reduce the survival and activity level of larval Cascades frogs (Rana cascadae). Archives of Enuiron. Contam. Toxicol. 39: 494-499. Hatch, A. C. and Blaustein, A. R., 2003. Combined effects of UV-B radiation and nitrate fertilizer on larval amphibians. Ecol. Appl. 13: 1083-1093. Hatch, A. C. and Burton, G. A. Jr., 1998. Effects of photoinduced toxicity of fluoranthene on amphibian embryos and larvae. Environ. Toxicol. Chem. 17: 1777-1785. Hays, J. B., Blaustein, A. R., hesecker, J. M., Hoffman, F! D., Pandelova, I., Coyle, C. and Richardson, T, 1996. Developmental responses of amphibians to solar and artificial UV-B sources: A comparative study. Photochem. Photobiol. 64: 449-456. Heelis, F! F., h m , S. T, Okamura, T and Sancar, A,, 1993. The photo repair of pyrimidine dimers by DNA photolyase and model systems. J. Photochem. Photobiol. B-Biol. 17: 219-228. Herman, J. R., Bhartia, F! K., Ziemke, J., Ahmad, Z. and Larko, D., 1996. UV-B increases (1979-1992) from decreases in total ozone. Geophys. Res. Lett. 23: 21 17-2120. Heyer, W. R., 1969. Adaptive ecology of species groups of genus Leptodactylus (Amphibia, Leptodactylidae). Evolutim 23: 421-428. Heyer, W. R., 2003. Ultraviolet-B and amphibia. Bioscience 53: 540-541.

Garcia, T. S., Stacy, J. and Sih, A,, 2004. Larval salamander response to UV radiation and predation risk: Color change and microhabitat use. Ecol. Appli. 14: 1055-1064.

Hofer, R., 2000. Vulnerability of fish and amphibians to ultraviolet radiation. Res. Adv. Photochem. Photobiol. 1: 265-282.

Gething, M. J. and Sambrook, J., 1992. Protein folding in the cell. Nature 355: 33-45.

Hofer, R. and Mokri, C., 2000. Photoprotection in tadpoles of the common frog, Rana temporaria. J. Photochem. Photobiol. B-Biol. 59: 48-53.

Gibbs, J. I?, Breisch, A. R., 2001. Climate warming and calling phenology of frogs near Ithaca, New York, 1900-1999. Cons. Biol. 15: 1175-1 178. Grant, K. F!, Licht, L. E., 1995. Effects of ultraviolet radiation on life-history stages of anurans from Ontario, Canada. Can. J. Zool. 73: 2292-2301. Green, A. J., 1999. Implications of pathogenic fungi for life-history evolution in amphibians. Funct. Ecol. 13: 573-575. Gurdon, J. B., 1960. The effects of ultraviolet irradiation on uncleaved eggs of Xenopus laeuis. Quart. J. Micro. Sci. 101: 299-311. Hader, D. F!, Kumar, H. D., Smith, R. C. and Worrest, R. C., 1998. Effects on aquatic ecosystems. J. Photochem. Photobiol. B-Biol. 46: 53-68.

Hoff, K., Blaustein, A. R., McDiarmid, R. W. and Altig, R., 1999. Behavior. Pp. 215-239 in "Tadpoles: The Biology of Anuran Larvae", ed by R. W. McDiarmid and R. Altig. University of Chicago Press, Chicago. Hutchinson, V. H. and Dupr*, R. K., 1992. Thermoregulation. Pp. 206249 in "Environmental Physiology of the Amphibia", ed by M. E. Feder and W. W. Burggren. University of Chicago Press, London. Jablonski, N. G., 1998. Ultraviolet light-induced neural tube defects in amphibian larvae and their implications for the evolution of melanized pigmentation and declines in amphibian populations. J. Herpetol. 32: 455-457. Jagger, J., 1985. Solar UV Actions on Living Cells. Praeger, New York.

MARC0 E T AL. : ULTRAVIOLET-B RADIATION AND AMPHIBIANS Kagan, J., Kagan, I? A. and Buhse, H. E., 1984. Lightdependent toxicity of alpha-terthienyl and anthracene toward late embryonic stages of Rana pipiens. J. Chem. Ecol. 10: 1115-1122. Kane, K. S. and Maytin, E. V., 1995. Ultraviolet Binduced apoptosis of keratinocytes in murine skin is reduced by mild local hyperthermia. J. Invest. Dermatol. 104: 62-67. Kats, L. B., lesecker, J. M., Chivers, D. I? and Blaustein, A. R., 2000. Effects of UV-B radiation on antipredator behavior in three species of amphibians. Ethology 106: 92 1-93 1. Kelly, D. J., Bothwell, M. L. and Schindler, D. W., 2003. Effects of solar ultraviolet radiation on stream benthic communities: An intersite comparison. Ecology 84: 2724-2740. Kerr, J. B. and McElroy, C. T, 1993. Evidence for large upward trends of Ultraviolet-B radiation linked to ozone depletion. Science 262: 1032-1034. A-683.

3141

Licht, L. E., 2003. Shedding light on ultraviolet radiation and amphibian embryos. Bioscience 53: 551-561. Licht, L. E. and Grant, K. I?, 1997. The effects of ultraviolet radiation on the biology of amphibians. Amer. Zool. 37: 137-145. Lillywhite, H. B., Licht, I? and Chelgren, C, 1973. The role of behavioral thermoregulation in the growth energetics of the toad, Bufo boreas. Ecology 54: 375-383. Lindquist, S., 1986. The heat-shock response. Ann. Rev. Biochem. 55: 1151-1 191. Lindquist, S., 1988. The heat-shock proteins. Ann. Reu. Gen. 22: 631-677. Lips, K. R., 1998. Decline of a tropical montane amphibian fauna. Cons. Biol. 12: 106-1 17. Lips, K. R., Green, D. E. and Papendick, R., 2003. Chytridiomycosis in wild frogs from southern Costa Rica. J. Herpetol. 37: 215-218.

Kiesecker, J. M. and Blaustein, A. R., 1995. Synergism between UV-B radiation and a pathogen magnifies amphibian embryo mortality in nature. Proc. Nat. Acad. Sci. USA 92: 11049-1 1052.

Little, E. E., Calfee, R. D., Fabacher, D. L., Carey, C., Blazer, V. S. and Middleton, E. M., 2003. Effects of ultraviolet radiation on toad early life stages. Environ. Sci. Poll. Res. 10: 167-172.

lesecker, J. M. and Blaustein, A. R., 1997. Influences of egg laying behavior on pathogenic infection of amphibian eggs. Cons. Biol. 11: 214-220.

Lizana, M. and Pedraza, E. M., 1998. Different mortality of toad embryos (Bufo bufo and Bufo calamita) caused by UV-B radiation in high mountain areas of the Spanish Central System. Cons. Biol. 12: 703-707.

Kiesecker, J. M., Blaustein, A. R. and Belden, L. K., 200 1. Complex causes of amphibian population declines. Nature 410: 681-684. Kim, S. T and Sancar, A., 1993. Photochemistry, photophysics, and mechanism of pyrimidine dimer repair by DNA photolyase. Photochem. Photobiol. 57: 895-904. Kollias, N., Sayre, R. M., Zeise, L. and Chedekel, M. R., 199 1. Photoprotection by melanin. J. Photochem. Photobiol. B-Biol. 9: 135-160. Koponen, P. S. and Kukkonen, J. V. K., 2002. Effects of bisphenol a and artificial UVB radiation on the early development of Rana temporaria. J . Toxicol. Environ. Health-Part A 65: 947-959. LaClair, J . J., Bantle, J. A. and Dumont, J., 1998. Photoproducts and metabolites of a common insect growth regulator produce developmental deformities in Xenopus. Environ. Sci. Technol. 32: 1453-1461. Langhelle, A, Lindell, M. J. and Nystrom, I?, 1999. Effects of ultraviolet radiation o n amphibian embryonic and larval development. J. Herpetol. 33: 449-456. Laurion, I., Ventura, M., Catalan, J., Psenner, R. and Sommaruga, R., 2000. Attenuation of ultraviolet radiation in mountain lakes: Factors controlling the among- and within-lake variability. Limnol. Oceanog. 45: 1274-1288. Leavitt, I? R., Vinebrooke, R. D., Donald, D. B., Smol, J. I? and Schindler, D. W., 1997. Past ultraviolet radiation environments in lakes derived from fossil pigments. Nature 388: 457-459. Lesser, M. I?, Turtle, S. L., Farrell, J. H. and Walker, C. W., 2001. Exposure to ultraviolet radiation ( 2 9 0 4 0 nm) causes oxidative stress, DNA damage, and expression of p. 53 and p. 73 in laboratory experiments on embryos of the spotted salamander, Ambystoma maculatum. Physiol. Biochem. 2001. 74: 733-741.

Long, L. E., Saylor, L. S. and Soule, M. E., 1995. A pH1 UV-B synergism in amphibians. Cons. Biol. 9: 1301-1303. Losey, G. S., Cronin, T W., Goldsmith, T. H., Hyde, D., Marshall, N. J. and McFarland, W. N., 1999. The W visual world of fishes: a review. J. Fish Biol. 54: 921-943. MacFadyen, E. J., Williamson, C. E., Grad, G., Lowery, M., Jeffrey, W. H. and Mitchell, D. L., 2004. Molecular response to climate change: temperature dependence of UV-induced DNA damage and repair in the freshwater crustacean Daphnia pulicaria. Global Change Biol. 10: 408-416. Madronich, S., McKenzie, R. L., Bjorn, L. 0 . and Caldwell, M. M., 1998. Changes in biologically active ultraviolet radiation reaching the Earth's surface. J. Photochem. Photobiol. B-Biol. 46: 5-19. Malloy, K. D., Holman, M. A., Mitchell, D. and Dietrich, H. W. III., 1997. Solar UVB-induced DNA damage and photoenzymatic DNA repair in antarctic zooplankton. Roc. Nat. Acad. Sci. USA 94: 1258-1263. Marco, A,, 2001. Effects of prolonged terrestrial stranding of aquatic Ambystoma gracile egg masses on embryonic development. J. Herpetol. 35: 5 10-5 13. Marco, A. and Blaustein, A. R., 1998. Egg gelatinous matrix protects Ambystoma gracile embryos from prolonged exposure to air. Herpetol. J. 8: 207-2 11. Marco, A. and Blaustein, A. R., 2000. Symbiosis with green algae affects survival and growth of Ambystoma gracile embryos. J. Herpetol. 34: 6 17-62 1. Marco, A., Lizana, M., Alvarez, A. and Blaustein, A. R., 200 1. Egg-wrapping behaviour protects newt embryos from UV radiation. Anim. Behau. 61: 639-644. Msrquez, R., 1992. Terrestrial pattern care and short breeding seasons: reproductive phenology of the midwife toads, Alytes obstetricam and A. cisternasii. Ecography 15: 279-288.

3142

AMPHIBJAN BIOLOGY

Meehl, G. A. and Washington, W. M., 1996. El Nifio-like climate change in a model with increased atmospheric C 0 2 concentrations. Nature 382: 56-60. Menter, J. M., Agin, I? I?, Sayre, R. M. and Wdlis, I., 1992. Dose-response of chronic ultraviolet exposure on epidermal forward scattering-absorption in sk-l hairless mouse skin. Photochem. Photobiol. 55: 705-7 12. Merila J., Pahkala, M., Johanson, U., 2000. Increased ultraviolet-B radiation, climate change and latitudinal adaptation-a frog perspective. Annales Zoologici Fennici 37: 129-134. Miaud, C., 1995. Oviposition site selection in three species of european newts (Salamandridae) genus T r i t u w . Amphibia-Reptilia 16: 265-272. Middleton, E. M., Herman, J. R., Celarier, E. A,, Wilkinson, J. W., Carey, C. and Rusin, R. J., 2001. Evaluating ultraviolet radiation exDosure with satellite iata at sites of amphibian' declines in Central and South America. Cons. Biol. 15: 914-929. Mikaelian, I., Ouellet, M., Pauli, B., Rodriguez, J., Harshbarger, J. C. and Green, D. M., 2000. Zchthyophonus-like infection in wild amphibians from Quebec, Canada. Dzseas. Aquatic Organ. 40: 195-201. Monson, I? D., Call, D. J., Cox, D. A,, Liber, K. and Ankley, G. T., 1999. Photoinduced toxicity of fluoranthene to northern leopard frogs (Rana pipiens). Enuiron. Toxicol. Chem. 18: 308-3 12. Morris, D. I?, Zagarese, H., Williamson, C. E., Balseiro, E. G., Hargreaves, B. R., Modenutti, B., Moeller, R. and Queimalinos, C., 1995. The attenuation of solar UV radiation in lakes and the role of dissolved organic carbon. Limnol. Oceanog. 40: 1381-1391. Nagl, A. M. and Hofer, R., 1997. Effects of ultraviolet radiation on early larval stages of the alpine newt, Triturus alpestris, under natural and laboratory conditions. Oecologia 110: 5 14-5 19. Nussbaum, R. A,, Brodie, E. D. Jr. and Storm, R. M., 1983. "Amphibians and reptiles of the Pacific Northwest". Univ. Press of Idaho, Moscow, Idaho. Ovaska, K., Davis, T. M., Flamarique, I. N., 1997. Hatching success and larval survival of the frogs Hyla regilla and Rana aurora under ambient and artificially enhanced solar ultraviolet radiation. Can. J. Zool. 75: 1081-1088. Ozer, Z., Reardon, J. T., Hsu, D. S., Malhotra, K. and Sancar, A., 1995. The other function of DNA photolyase: Stimulation of excision repair of chemical damage to DNA. Biochemistry 34: 15886-15889. Pahkala, M., Laurila, A. and Merila, J., 2000. Ambient ultraviolet-B radiation reduces hatchling size in the common frog Rana temporaria. Ecography 23: 53 1-538. Pahkala, M., Laurila, A. and Merila, J., 2001a. Carryover effects of ultraviolet-B radiation on larval fitness in Rana temporaria. Proc. Roy. Sac. Land. B 268: 1699-1706. Pahkala, M., Laurila, A. and Merila, J., 2002b. Effects of ultraviolet-B radiation on common frog Rana temporaria embryos from along a latitudinal gradient. Oecologia 133: 458-465. Pahkala, M., Laurila, A., Bjorn, L. 0 . and Merila, J., 2001b. Effects of ultraviolet-B radiation and pH on early development of the moor frog Rana arvalis. J. Appl. Ecol. 38: 628-636. Pahkala, M., Merila, J., Ots, I. and Laurila, A,, 2003. Effects of ultraviolet-B radiation on metamorphic traits in the common frog Rana temporaria. J. Zool. ( h d . ) 259: 57-62.

Palen, W. J., Schindler, D. E., Adams, M. J., Pearl, C. A., Bury, R. B. and Diamond, S. A., 2002. Optical characteristics of natural waters protect amphibians from UV-B in the U.S. Pacific Northwest. Ecology 83: 295 1-2957. Pang, Q. and Hays, J. B., 1991. UV-inducible -and temperature-sensitivephotoreactivation of cyclobutane pyrimidine dimers in Arabidopsis thaliana. Plant Physiol. 95: 536-543. Parris, M. J. and Beaudoin, J. G., 2004. Chytridiomycosis impacts predator-prey interactions in larval amphibian communities. Oecologia 140: 626-632. Peterson, G. S., Johnson, L. B., Axler, R. P and Diamond, S. A., 2002. Assessment of the risk of solar ultraviolet radiation to amphibians. 11. I n situ characterization of exposure in amphibian habitats. Enuiron. Sci. Technol. 36: 2859-2865. Petranka, J. W., Just, J. J. and Crawford, E. C., 1982. Hatching of amphibian embryos: the physiological trigger. Science 217: 257-259. Pienitz, R. and Vincent, W. F., 2000. Effect of climate change relative to ozone depletion on UV exposure in subartic lakes. Nature 404: 484-487. Pounds, J. A., 2001. Climate and amphibian decline. Nature 410: 639-640. Pounds, J. A., Fogden, M. I? L. and Campbell, J. H., 1999. Biological response to climate change on a tropical mountain. Nature 398: 61 1-615. Pounds, J. A., Consuegra, J. A., Fogden, M. I? L., Foster, I? N., Masters, K. L., Puschendorf, R., Ron, S. R., Sinchez-Asofeifa,A. and Still, C. J., 2005. Widespread amphibian extinctions from epidemic disease driven by global warming. Nature in press. Prota, G., 1992. "Melanins and Melanogenesis". Academic Press, San Diego. Randel, W. J., Stolarski, R. S., Cunnold, D. M., Logan, J. A,, Newchurch, M. J. and Zawodny, J . M., 1999. Atmosphere - trends in the vertical distribution of ozone. Science 285: 1689-1692. Rasanen, K., Pahkala, M., Laurila, A. and Merila, J., 2003. Does jelly envelope protect the common frog Rana temporaria embryos from UV-B radiation? Herpetologica 59: 293-300. Reading, C. J., 1998. The effect of winter temperatures on the timing of breeding activity in the common toad Bufo bufo. Oecologia 117: 469475. Reaser, J. K. and Blaustein, A. R., 2005. Repercussions of global change. Pp. 60-63 in "Amphibian Declines", ed by M. Lannoo. University of California Press, Berkeley. Rocco, V E., Oppezzo, O., Pizarro, R., Sommaruga, R., Ferraro, M. and Zagarese, H. E., 2002. Ultraviolet damage and counteracting mechanisms in the freshwater copepod Boeckella poppei from the Antarctic Peninsula. Limnol. Oceanog. 47: 829-836. Sancar, A. and Sancar, G., 1998. DNA repair enzymes. Ann. Rev. Biochem. 57: 29-67. Schindler, D. W., Curtis, P J., Parker, B. R. and Stainton, M. I?, 1996. Consequences of climate warming and lake acidification for UV-B penetration in North American boreal lakes. Nature 379: 705-708. Scully, N. M. and Lean, D. R. S., 1994. The attenuation of the ultraviolet radiation in temperate lakes. Arch. Hydrobiol. 45: 135-144.

MARC0 E T AL.: ULTRAVIOLET-B RADIATION AND AMPHIBIANS Sergeev, A. M. and Smirnov, K. S., 1939. The color of eggs of Amphibia. Probl. Ecol. Biocenol. 5: 3 19-32 1. Smith, G. R., Waters, M. A. and Rettig, J. E., 2000b. Consequences of embryonic UV-B exposure for embryos and tadpoles of the plains leopard frog. Conseru. Biol. 14: 1903-1907. Smith, M. A,, 2000. Problems using an enzyme sensitive site assay for photorepair of exogenous DNA with cell-free extracts made from amphibian embryos. Can. J. Zool. 78: 1869-1872. Smith, M. A., Berrill, M. and Kapron, C. M., 2002. Photolyase activity of the embryo and the ultraviolet absorbance of embryo jelly for several Ontario amphibian species. Can. J. Zool. 80: 1109-1 116. Smith, M. A,, Kapron, C. M. and Berrill, M., 2000a. Induction of photolyase activity in wood frog (Rana syluatica) embryos. Photochem. Photobiol. 72: 575-578. Sommaruga, R. and Garcia-Piche, F., 1999. UV-absorbing mycosporine-like compounds in planktonic and benthic organisms from a high-mountain lake. Archiv fur Hydrobiologie 144: 255-269. Starnes, S. M., Kennedy, C. A. and Petranka, J. W., 2000. Sensitivity of embryos of southern Appalachian amphibians to ambient solar UV-B radiation. Conrerv. Biol. 14: 277-282. Stebbins, R. C. and Cohen, N. W., 1995. "A Natural History of Amphibians". Princeton University Press, Princeton. Tartarotti, B., Cabrera, S., Psenner, R. and Sommaruga, R., 1999. Survivorship of Cyclops abyssorum tatricw (Cyclopoida, Copepoda) and Boeckella gracilipes (Calanoida, Copepoda) under ambient levels of solar UVB radiation in two high-mountain lakes. J . Plankton Res. 21: 549-560. Taylor, S. K., Williams, E. S. and Mills, K. W., 1999. Experimental exposure of Canadian toads to Baszdiobolus ranarum. J. Wildl. Dis. 35: 58-63. Tevini, M. (editor), 1993. "UV-B Radiation and Ozone Depletion: Effects on Humans, Animals, Plants, Microorganisms, and Materials". Lewis Publishers, Boca Raton. Tietge, J. E., Diamond, S. A,, Ankley, G. T., DeFoe, D. L., Holcombe, G. W., Jensen, K. M., Degitz, S. J., Elonen, G. E. and Hammer, E., 2001. Ambient solar UV radiation causes mortality in larvae of three species of Rana under controlled exposure conditions. Photochem. Photobiol. 74: 261-268. Timmermann, A., Oberhuber, J., Bacher, A., Esch, M., Latif, M. and Roeckner, E., 1999. Increased El Nino frequency in a climate model forced by future greenhouse warming. Nature 398: 694-697. Trautinger, F., Kindas-Mugge, I., Knobler, R. M. and Honigsmann, H., 1996. Stress proteins in the cellular response to ultraviolet radiation. J. Photochem. Photobiol. B 35: 141-148.

3143

van de Mortel, T., Buttemer, W., Hoffman, P, Hays, J. and Blaustein, A,, 1998. A comparison of photolyase activity in three Australian tree frogs. Oecologia 115: 366-369. Vermeer, B. J. and Hurks, M., 1994. The clinical relevance of immunosuppression by UV irradiation. J. Photochem. Photobiol. B-Biol. 24: 149-154. Vetter, R. D., Kurtzman, A,, Mori, T, 1999. Die1 cycles of DNA damage and repair in eggs and larvae of northern anchovy, Engraulis mordax, exposed to solar ultraviolet radiation. Photochem. Photobiol. 69: 27-33. Vonesh, J. R. and de la Cruz, O., 2002. Complex life cycles and density dependence: Assessing the contribution of egg mortality to amphibian declines. Oecologia 133: 325-333. Wake, M. H., 1993. Evolution of oviductal gestation in amphibians. J. Experi. Zool. 266: 3 9 4 4 13. Walker, S. E., Taylor, D. H. and Oris, J. Z, 1998. Behavioral and histopathological effects of fluoranthene on bullfrog larvae (Rana catesbeiana). Environ. Toxicol. Chem. 17: 734-739. Walther, G. R., Post, E., Convey, P, Menzel, A., Parmesan, C., Beebee, T J. C., Fromentin, J. M., Hoegh-Guldberg, 0 . and Bairlein, F., 2002. Ecological responses to recent climate change. Nature 416: 389-395. Warkentin, K. M., 1992. Microhabitat use and feeding rate variation in green frog tadpoles (Rana clamitam). Copeia 1992: 731-740. Wilkinson, M. and Nussbaum, R. A., 1998. Caecilian viviparity and amniote origins. J. Nut. Hist. 32: 1403-1409. Williamson, C. E., 1995. What role does UV-B radiation play in freshwater ecosystems? Limnol. Oceanog. 40: 386-392. Williamson, C. E., 1996. Effects of UV radiation on freshwater ecosystems. Intern. J. Environ. Stud. 51: 245-256. Williamson, C. E., Metzgar, S. L., Lovera, P A. and Moeller, R. E., 1997. Solar ultraviolet radiation and the spawning habitat of yellow perch, Perca flavescem. Ecol. Applicat. 3: 1017-1023. Williamson, C. E., Olson, 0 . G., Lott, S. E., Walker, N. D., Engstrom, D. R. and Hargreaves, B. R., 2001. Ultraviolet radiation and zooplankton community structure following deglaciation in Glacier Bay, Alaska. Ecology 82: 1748-1760. Williamson, C. E., Stemberger, R. S., Morris, D. P, Fmst, T M. and Paulsen, S. G., 1996. Ultraviolet radiation in North American lakes: attenuation estimates from DOC measurements and implications for plankton communities. Limnol. Oceanog. 41: 1024-1034.

Tyagi, R., Kumar, A., Tyagi, M. B., Jha, P N., Kumar, H. D., Sinha, R. P, Hader, D. ,.'F 2003. Protective role of certain chemicals against UV-B-induced damage in the nitrogen-fixing cyanobacterium, Nostoc muscorum. J. Basic Microbiol. 43: 137-147.

Wollmuth, L. P, Crawshaw, L. I., Forbes, R. B. and Grahn, D. A., 1987. Temperature selection during development in a montane anuran speices, Rana cascadae. Physiol. 2001. 60: 472-480.

van de Mortel, T. F. and Buttemer, W. A,, 1998. Avoidance of ultraviolet-B radiation in frogs and tadpoles of the species Litoria aurea, L. dentata and L. peronii. Proc. Linn. Soc. N S W 119: 173-179.

Worrest, R. C. and Kimeldorf, D. J., 1976. Distortions in amphibian development induced by ultravioletB enhancement (290-315 nm) of a simulated solar spectrum. Photochem. Photobiol. 24: 377-382.

3144

AMPHIBIAN BIOLOGY

Wright, R. F. and Schindler, D. W., 1995. Interaction of acid rain and global changes: Effects on terrestrial and aquatic ecosystems. Water Air and Soil Poll. 85: 89-99. Wu, C., 1999. Unraveling the mystery of melanin. Science News 156: 190-191. Xenopoulos, M. A. and Schindler, D. W., 2001. Physical factors determining ultraviolet radiation flux in ecosystems. Pp. 36-62 in "Ecosystems, Evolution and Ultraviolet Radiation", ed by C. S. Cockell and A. R. Blaustein. Springer, New York. Yan, N. D., Keller, W., Scully, N. M., Lean, D. R. S. and Dillon, F! J., 1996. Increased W - B penetration in a lake owing to drought-induced acidification. Nature 381: 141-143.

Yasui, A. and McCready, S. J., 1998. Alternative repair pathways for UV-induced DNA damage. Bioessays 20: 291-297. Zaga, A,, Little, E. E., Rabeni, C. F. and Ellersieck, M. R., 1998. Photoenhanced toxicity of a carbamate insecticide to early life stage anuran amphibians. Enuiron. Toxicol. Chem. 17: 2543-2553. Zamzow, J. P., Losey, G. S., 2002. Ultraviolet radiation absorbance by coral reef fish mucus: photoprotection and visual communication. Environ. Biol. Fishes 63: 41-47. Zhou, X., Tron, V. A, Li, G. and Trotter, M. J., 1998. Heat shock transcription factor-1 regulates heat shock protein-72 expression in human keratinocytes exposed to ultraviolet B light. J. Znuestig. Dermatol. 111: 194-198.

CHAPTER 7

Pollution: Impact of Reactive Nitrogen on Amphibians (Nitrogen Pollution) Adolfo Marco and Manuel Ortiz-Santaliestra

I. Nitrogen Pollution or the Nitrogen Cascade II. Environmental Sources of Reactive Nitrogen (Nr) A. Nitrogen Fertilizer B. Livestock Residues C. Wastewater Effluents D. Deposition E. Other Sources Ill. Ecological Effects of an Excess of Reactive Nitrogen IV. Environmental Exposure of Amphibians to Nitrogen Polllution V Physiological Effects of Reactive Nitrogen on Amphibians A. Ammonia 6. Urea C. Nitrate and Nitrite VI. Dose-Effect Relationships A. Ammonia 1. Effects on Embryos 2. Effects on Larvae B. Ammonium Nitrate 1. Effects on Embryos 2. Effects on Larvae 3. Effects on Adults C. Nitrite 1. Effects on Embryos 2. Efffects on Larvae

D. Nitrate 1. Effects on Embryos 2. Effects on Larvae E Urea 1. Effects on Larvae 2. Effects on Juveniles 3. Effects on Adults VII. Sublethal Effects A. Abmormalities B. Effects on Reproduction C. Effects on Embryonic and Larval Development D. Effects on Metamorphosis E. Behavioural Effects VIII. Variability among Species IX. Variability among Populations X. Importance of Age to Sensitivity XI. Synergism with other Ecological Agents A. Ultraviolet Radiation B. Acidification C. Temperature and Dissolved Oxygen D. Other Chemicals E. Pathogens XII. Role of Nitrogen Pollution in Amphibian Conservation XIII. References

Abbreviations and acronyms used in the text and references: ASTM =American Society for Testing and Materials; BFH = Federal Research Center for Forestry and Forest Products; DOC = dissolved organic carbon; EIFAC = European Inland Advisory Commission; EPA = Environmental Protection Agency; FA0 = Food and Agriculture Organization; FETAX = Frog Embryo Teratogenesys Assay - Xenopus; FMA = Fertiliser Manufacturer's Association; G25 = Gosner Stage 25; HMSO = Her Majesty's Stationery Office; LC50 = Medium Lethal Concentration; MAFF = Ministry of Agriculture, Fisheries and Food; MCL = maximum contaminant level; NATO = North Atlantic Treaty Organization; NOEC = No Observed Effect Concentration; OECD = Organization for Economic Co-operation and Development; PAH = Polycyclic aromatic hydrocarbons; PCB = polychlorobiphenyls; SOAFD = Scottish Office of Agriculture and Fisheries Department; TI = teratogenic index; USGS = United States Geological Survey; UV-B = ultraviolet B.

3146

AMPHIBIAN BIOLOGY

I. NITROGEN POLLUTION OR THE NITROGEN CASCADE

TROGEN (N) is an essential component of proteins, DNA and other basic molecules. It is also a key element controlling species-composition, diversity, dynamics and functioning of many terrestrial, freshwater and marine ecosystems. The growth and dynamics of herbivore populations, and ultimately those of their predators, are affected by nitrigen (Tamm 1991; Jarvis 1996). Alteration of the nitrogen cycle due to human activities is a global environmental problem (Vitousek 1994; Tilman 1999). The Earth's growing human population has increased the demand for food and energy worldwide. Meeting these demands has doubled the rate at which reactive nitrogen is produced, thereby greatly increasing the amount of reactive nitrogen in the environment. Agriculture, combustion of fossil fuels and other human activities have altered the global cycle of N substantially, generally increasing both the availability and the mobility of N over large regions of the Earth (Vitousek et al. 1997; Matson et al. 1999). The mobility of N means that, while most deliberate applications of N occur locally, their influence spreads regionally and even globally. Moreover, many of the reactive forms of N themselves have environmental consequences. Although most nitrogen inputs serve human needs (such as agricultural production), the environmental consequences of an excess of nitrogen are serious and longer lasting (Aber 1992; Vitousek et al. 1997). The same atom of reactive N can cause a cascade effect in the atmosphere and in terrestrial, freshwater and marine ecosystems (Galloway et al. 2003). Given the combination of beneficial and harmful effects, nitrogen pollution in the environment is often referred to as "too much of a good thing" (Driscoll 2003). In recent years, many freshwater systems have experienced increased nitrogenous input resulting in elevated levels of nitrite, nitrate and ammonia (EIFAC 1984). For example, the movement of total dissolved nitrogen into surface waters may have increased by up to 20-fold in the North Atlantic Ocean Basin and in the North Sea region since pre-industrial times. During this time, nitrate levels have risen up to ten-fold in most of the rivers and lakes of developed countries. The form of nitrogen that is most toxic to biota is ammonia, followed by nitrite and nitrate (Camargo and Ward 1992). Because ammonia and nitrite in bodies of water are quickly oxidized to nitrate by bacteria and algae, nitrate occurs at the highest concentrations and is the most stable form of nitrogen in the aquatic environment (Camargo and Ward 1992). The United States Geological Survey (USGS) released a report in 1995 that revealed that nitrate concentration in that nation's groundwater supply is increasing steadily; 9% of wells tested had nitrate concentrations exceeding 10 ppm, the Environmental Protection Agency's (EPA) maximum contaminant level (MCL) for N-nitrate. In many populated areas concentrations of groundwater nitrate exceed the EPA maximum contaminant level for drinking water of 44.27 mg/l (10 mgll NO3--N)(Kross et al. 1993; Hudak 2000). Many public water supplies in the United States contain levels of nitrate that routinely exceed a concentration of 10 mg NIL (U.S. EPA 1986). Nitrate loading beneath and around irrigated crops may easily exceed concentrations of 20 mg NO3--N/l(Kraft and Stites 2003). These authors found during a four-year study in a humid north-central sand plain in the United States that groundwater in the field ranged from <0.2 to 50.5 mg N/1. In Japan, 90% of the water samples showed nitrate concentrations above the level that affects humans (3 mgll NO?), while more than 30% exceeded the maximum acceptable level (44 mg/l NO<) according to Japanese regulations (Babiker et al. 2004). Levels of nitrite and un-ionized ammonia (the most toxic forms of reactive nitrogen) in natural aquatic habitats are usually low but under some circumstances and in specific areas, such as shore sites with high contents of organic matter, NOp- and NH, concentrations can rise to toxic levels (McCoy 1972). Some inshore sites that contain rotting vegetation and algae can have nitrite concentrations as high as 180 mg/L N-NO; (McCoy 1972). High levels of environmental nitrite up to 4 mg N-NO,-/L are often found in fish hatcheries.

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3147

11. ENVIRONMENTAL SOURCES OF REACTIVE NITROGEN (NR)

The accumulation of reactive nitrogen compounds in the environment results mainly from non-point source runoff, from the over-application of nitrogenous fertilizers or the cultivation of crops that host symbiotic nitrogen-fixing bacteria. In addition, nitratecontaining wastes are produced by many industrial processes including paper and munitions manufacturing (Puckett 1995; Boyer et al. 2002). Some reactive nitrogen enters the ecosystem from the atmosphere, which carries nitrogen-containing compounds derived from combustion of fossil fuels and other sources. Ammonia and organic nitrogen can enter ecosystems through sewage effluents and runoff from land where manure has been applied as fertilizer or stored. Some human activities such as burning forests and grasslands, draining wetlands or land-clearing for crops also liberate a significant amount of reactive nitrogen to the environment, forming long-term biological storage pools (Vitousek et al. 1997). The evaporation of water in amphibian habitats may also lead to an increased concentration of reactive nitrogen and other chemicals to toxic levels. All these sources of high levels of reactive nitrogen are common all over the inhabited areas of the world and their impact may be considered global. Reactive nitrogen concentration in agricultural or urban runoff is highly variable and nitrogen peaks can easily exceed recommended limits. There is a strong relationship between the nitrogen load and peak nitrogen concentrations; the slope of the regression is usually higher in sandy and loamy soils (Scholefield et al. 1996). These peak concentrations are not often detected by conventional periodic monitoring of nitrogen pollution. In ecotoxicological studies, reliable data on the concentration of nitrogenated ions in water should be recorded during the most sensitive stages of amphibian development. Scholefield et al. (1996) proposed a model to predict peak nitrate concentration in water draining agricultural land, using the annual load of leachable nitrate in the soil. There are other sources of seasonal variability of nitrate or ammonium concentration. For example, during cold seasons most terrestrial and aquatic plants are dormant or their growth is significantly reduced. For this reason, the uptake of nutrients by plants decreases and the amount of reactive nitrogen in the leachate increases (Rouse et al. 1999). The dormant period of plants includes the breeding season for most species of amphibians in temperate zones. Vitousek et al. (1997) reported how anthropogenic sources of N fixation have approximately doubled the global rate of N fxation characteristic of pre-industrial terrestrial ecosystems. Of the 140 teragrams of nitrogen estimated to be fixed each year by anthropogenic sources, between 25 and 40 originate from intensive cultivation of legumes, 80 from application of synthetic N fertilizers, and 20 from combustion of fossil fuels (Vitousek et al. 1997; Smil 2002). A. Nitrogen Fertilizer Man-made fertilizers became widely available after World War I1 and quickly came into use around the world. Today, nitrogen fertilizers (commonly nitrate or ammonium compounds) are used in large quantities in most agriculture. The recent intensification of agriculture, and the prospects of future intensification, will have major detrimental impacts on the nonagricultural terrestrial and aquatic ecosystems of the world. During the past 35 years agricultural practices were associated with a 6.87-fold increase in nitrogen fertilization. At a global scale, fertilizer consumption has increased from 10.8 million tons N per year in 1960 to 72 million tons N per year in 1991 and 85.6 million tons per year in 2000 (FA0 2002). In the United States, use of nitrogen-based fertilizer increased from approximately 2.5 million tons per year in 1960 to almost 11.9 million tons per year in 1985 (Berry 1994). Based on a simple linear extension of past trends, the anticipated next doubling of global food production would be associated with approximately a three-fold increase in nitrogen fertilization rates. These projected changes would have dramatic impacts on the diversity, composition and functioning of the remaining natural ecosystems of the world, and on their ability to provide society with a variety of essential ecosystem services.

3148

AMPHIBIAN BIOLOGY

There has been a recent shift towards spring as the peak period for application of fertilizer (Chalmers et al. 1990). In 1985, 17% of total compound nitrogen was applied after January; this rose to 77% in 1990. Application of straight nitrates is also now concentrated during the spring months; 67% of ammonium nitrate is applied between March and April. Nitrogen fertilizer is typically applied as ammonium nitrate, urea or ammonium sulphate (FMA, MAFF and SOAFD 1993; Meister 1995) and fertilizer-N applications range from 60 to 260 kg N/Ha depending on the crop and the type of soil (Kraft and Stites 2003). Billions of pounds of artificially-made nutrients are applied to crops every year. Such addition of fertilizers to the soil may directly affect amphibian terrestrial stages. Application of N fertilizers is usually concentrated during the winter and spring months and on many occasions is simultaneous with amphibian breeding migrations, spawning and the aquatic development of the early stages. The persistence of ammonium nitrate in the soil is low, but other fertilizers such as urea may remain in the surface at high concentrations for longer. Granulated urea is widely used in forest fertilization. In the 1980s, fertilizer application was a well-established management practice used in many forested regions of the world to sustain or accelerate production of timber (Gessel and Atkinson 1984). For any individual plot, there are three to six applications within approximately ten years, with a rate of 150450 kg N/ha per application (Gessel and Atkinson 1984). Urea fertilizer is a commercial synthetic amide of carbonic acid widely used in solid and liquid fertilizers for direct application. It is also commonly applied to fields in liquid form combined with other fertilizers such as in urea ammonium nitrate solutions, or broadcast in granulated form as urea ammonium sulphate or urea ammonium phosphate (Meister 1995). Amphibians may temporarily use agricultural lands for feeding or during their nuptial migrations. Moreover, forest or grassland fertilization (Bengtson 1979) may directly affect typical amphibian habitats. Aerial fertilization may also affect terrestrial and aquatic habitats adjacent to the fertilized area. Plants never use more than half of the fertilizer applied by growers. The fertilizer mainly percolates into ground water. It also evaporates into the air or flows from fields to streams and lakes. Leached nitrates can also flow laterally over impermeable soil layers, or via groundwater basins into freshwater lakes or rivers (Crews and Peoples 2004). In some instances, water supplies become so contaminated with excess nitrogen they become unfit for human use. For example, losses of ammonia following fertilizer applications to upland and lowland cropping systems can range from -0 to >50%, while losses from flooded rice can reach as high as 80% (see review by Peoples et al. 1995). Concentration of both ammonium and nitrate ions in wetlands influenced by agricultural runoff can reach levels that are likely to produce deleterious effects on wildlife. For example, McDowell and McGregor (1979) found levels of 2-21 mg NH,+-NIL and 2-78 mg NO,--NIL for over 30 days in store runoff from Mississippi no-till corn fields fertilized with 136 kg N/ha. Peak NO3-N in surface runoff from Ohio pastures ranged from 12.9-77.5 mg/L within the first two days following fertilization (Owens et al. 1983). Runoff from fescue grass plots treated with poultry litter (218 kg N/ha) contained from 42 mg NH,+-NIL immediately after application to 1 mg/L at 36 days post-application (Edwards and Daniel 1994). Nitratenitrogen in seepage from tile drains in Indiana corn crops fertilized with 285 kg N/ha from 1984-1988 was seldom < 10 mg/L and was generally 20-30 mg/L (Kladivko et al. 1991). Mueller et al. (1997) found that where application rates exceeded 80 kg N/ha, the level of surface water NO3--Nin the Des Moines River was as high as 5.9-14.5 mg/L in a ten-year study of agricultural practices. Nitrate concentrations > 10 mg/L have been observed in some small Willamette Basin streams (Bonn et al. 1996). Other studies have reported that concentration of nitrate in aquatic ecosystems affected by agricultural and urban activities around the world can exceed 100 mg/L (Bogardi et al. 1991). The greatest impacts of dissolved inorganic nitrogen are in freshwater, and can cause eutrophication by high rates of release of nitrogen and phosphorus from agricultural fields

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3149

(Matson et al. 1997; Tilman 1999). Additionally, the application of ammonium-based fertilizers (ammonium nitrate, ammonium phosphate or ammonium sulphate), increases the net H+ concentration of soils, and thus directly contributes to soil acidification, even in the absence of nitrate leaching (Kennedy 1992). B. Livestock Residues

Ammonium is released to the environment from sewage and manure. It is also the major nitrogenous excretion product of aquatic animals whereas urea is the main product from terrestrial animals. Animal production inevitably causes a loss of nutrients and the production of waste products. The latter may exceed the carrying capacity of an area and become detrimental to the environment (Tamminga 2003). Improperly managed manure can contaminate both surface water and ground water with nitrogenated nutrients. Storing livestock manure allows producers to spread it when crops can best use the nutrients. Accumulating, and therefore concentrating, manure in one area can, however, be risky to the environment and to both human and animal health unless done properly. An excess of livestock grazing in the field may also contaminate soil and water with an excess of manure and urine. C. Wastewater Effluents

Wastewater effluents, even after conventional treatment in sewage treatment plants, are rich in ammonia and nitrate. These effluents can be released directly to aquatic habitats. The practice of irrigating forested and agricultural lands is also widespread. The spray irrigation of treated wastewater effluents is a common agricultural practice that results in alterations of terrestrial and aquatic habitats (Tamminga 1995) and can also affect amphibian survival. For example, wastewater effluents that were sprayed onto forested and agricultural lands at State Game Lands 176, Pennsylvania, contained an average of 40.75 mg/L nitrate (Sopper 1986). Flooding can also be an important cause of wetland pollution by wastewater. Industrial effluents and discharges from wastewater-treatment plants are a source of nitrogen for aquatic ecosystems in urban areas. Although these point sources contribute to only a small percentage of the total nitrogen released to the environment, long-term direct discharge into a watercourse will have a significant detrimental effect on stream ecosystems downstream of the site of discharge (Rouse et al. 1999). D. Deposition Important sources of nitrogen contamination are emissions of trace gases such as nitrogenous oxides, which contribute to the formation of tropospheric smog and ozone as well as acid rain and N fertilization of downwind ecosystems (Vitousek et al. 1997). The effects of N,O emissions occur on a global scale; N20 is a potent greenhouse gas (Peoples et al. 1995) and can also catalyze the destruction of stratospheric ozone (Crutzen and Ehhalt 1977). In the 1970s and 1980s concerns over nitrogen deficiencies and effects of removal in harvests gave way to concerns over excess nitrogen availability and the potential for forest decline and surface-water pollution. Driving this paradigm shift is the increase in atmospheric deposition of nitrogen to forests due to industrial and agricultural activity. At the core of the new paradigm is the concept of "nitrogen saturation" of forest ecosystems (Aber 1992). Depositions of nitrogen still exceed critical loads over a large proportion of European forests (Federal Research Center for Forestry and Forest Products [BFH] 2003). Atmospheric concentrations and deposition of the major nitrogenous compounds and their biological effects in ecosystems are of great concern. In locations close to sources of photochemical smog, concentrations of oxidized forms of N (NO<, NO<) dominate, while in areas near agricultural activities the importance of reduced N forms (NH,, NH4+) significantly increases. Nitrogen deposition in highly exposed areas has led to N saturation of forests. In N-saturated forests high concentrations of NO3- are found in stream waters (Bytnerowicz and Fenn 1996).

3150

AMPHIBLAN BIOLOGY

In urban areas, rain, snow and fog contain various amounts of nitrogen depending on the geographic location. Motor vehicular and industrial exhausts release nitrogen oxides to the atmosphere that are deposited in aquatic ecosystems via precipitation. In heavily populated industrialized areas the concentration of nitrogen in precipitation can be very high. Atmospheric nitrogen can enter aquatic ecosystems through precipitation directly onto the watercourse or through runoff into the watershed via storm sewers. In forested or heavily vegetated areas that are not artificially fertilized, the land has a natural ability to absorb and utilize the nitrogen from precipitation. Atmospheric deposition may be a problem in watersheds that do not have an extensive ground cover of natural vegetation, such as occurs in urban areas (Rouse et al. 1999). The release to the atmosphere of NO, gas from fossil fuel combustion, NH, gas from agricultural production and particulate N from a range of human activities results in wet and dry deposition of nitrate (NO3-)and ammonium (NH,+) on the surface of the land. The rate of atmospheric deposition has greatly increased since the beginning of the 20th century, and high rates of atmospheric N deposition have been widely documented in Europe as well as in North America (Dise and Wright 1995; Fenn et al. 1998). Atmospheric deposition of pollutants can affect not only modified areas, such as urban or agricultural regions, but also pristine ecosystems through long-distance transport of chemicals. Substances in the atmosphere are transported easily and deposited often on mountains or in polar areas, far from their source. Pounds and Crump (1994) suggested that atmospheric scavenging of contaminants by clouds might concentrate contaminants and release them in remote areas such as in Monteverde, Costa Rica, where numerous amphibian species have declined (Pounds et al. 1999). In the Rocky Mountains in Colorado, the rates of No3- leaching and NO3-concentrations in surface waters have increased since the mid-20th century in response to rates of atmospheric N deposition that increased to 4.7 kgka per year by the early 1990s (Williams et al. 1996; Burns 2004). In Europe, about half the ammonia that is volatilized is deposited within a 50 km radius, while the other half is deposited over a much broader region (Ferm 1998). These effects are likely to become more prominent given current or increased rates of N deposition. Matson et al. (1999) suggested that the direct effects of anthropogenic N deposition on N cycling processes will lead to increased fluxes at the soil-water and soil-air interfaces, with little or no lag time in response. When only nitrogen deposition is the source of anthropogenic nitrogen pollution, the levels of reactive nitrogen are relatively low and far from toxic concentrations. In these cases, acidification caused by nitrogen deposition may be the cause of some environmental impacts on amphibians (Burns 2004).

E. Other Sources Forests, grasslands and wetlands represent large reservoirs of N that may be altered by ecosystem perturbations. Wild or prescribed fires can result in nitrogen pulses in amphibian habitats (Pilliod et al. 2003). During the initial firestorm, phosphorus and nitrogen levels can increase significantly in streams and ponds (Minshall et al. 199'7; Earl and Blinn 2003) with some extreme cases reaching levels of up to 60-fold above background level (Spencer et al. 2003). Nutrients can return to background concentrations within several weeks after a fire. During subsequent years, nutrient concentrations can periodically increase on fireimpacted sites, especially during spring runoff (Spencer et al. 2003). Expanded fire activity can mobilize substantial quantities of ammonium and nitrate in lakes and streams, even as a result of atmospheric fallout from fires beyond the immediate area (Earl and Blinn 2003). The potential for increased nutrient loadings in surface waters could extend the risk of their eutrophication (Spencer et al. 2003). In areas with high pre-fire levels of nitrogen pollution, the risk of toxic effects of nutrients such as ammonia or nitrate can threaten survival of the less-tolerant amphibian species. Pumping groundwater up to the surface can also result in an excess of nitrogen in wetlands. Kraft and Stites (2003) demonstrated how nitrate levels in irrigated crop fields

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3151

in the Wisconsin Central Sand Plain rose in a few years as a consequence of increased nitrate concentrations in groundwater used for irrigation. 111. ECOLOGICAL EFFECTS OF AN EXCESS OF REACTIVE NITROGEN

Freshwater ecosystems are especially sensitive to nitrogen pollution. A small excess of nitrogen in bodies of water may lead to their eutrophication, with loss of available dissolved oxygen. Agroecosystems in the western United States use 80-90% of the water resources of the area, often resulting in eutrophic conditions (elevated pH, higher water temperature and un-ionized ammonia) (Schuytema and Nebeker 1999a). Excess ammonia and nitrate concentrations related to eutrophication in bodies of water in agricultural areas may threaten amphibians (Schuytema and Nebeker 1999b). There is good evidence that anoxia or periods of hypoxia have increased in many estuaries and lakes during the last five decades, resulting in losses of biodiversity. Excessive concentrations of nitrate in lakes and streams greater than approximately 5 mg NIL can cause blooms of toxic algae and pathogenic fungi and bacteria (Sterner 1989; Vitousek et al. 1997; Rouse et al. 1999). Aquatic nutrient eutrophication can lead to loss of biodiversity, outbreaks of nuisance species, shifts in the structure of food chains and impairment of fisheries (Hack-ten-Broekeet al. 1996; Matson et al. 1997; Tilman 1999). Small new supplies of nitrogen may also cause a dramatic shift in the dominant plant species and a marked reduction in overall plant species diversity (Vitousek et al. 1997). Burns (2004) reported the effects that atmospheric deposition of N had on alpine ecosystems in Colorado (see his table 3). These changes may affect trophic chains, thus affecting amphibians. Amphibians may play a critical role in energetic transfers through their intermediate position in forest trophic webs and through their efficient processing and storage of energy (Hariston 1987; Pough et al. 1987). Adverse impacts on amphibian populations could possibly reverberate through adjacent trophic levels and affect ecosystems processes (Laposata and Dunson 2000). Nitrogen pollution is also playing an increasing role in the acidification of lakes and streams. One way that acidification takes place is by the deposition of acid directly by rain, snow, fog, mist or dry matter. Nitrogen-saturated soils have a lower capacity to buffer acid rain before it enters streams (Vitousek et al. 1997). Additionally, nitric acid accumulates in the winter snowpack and much of it is flushed out with the spring meltwater, causing a concentrated injection of acid into amphibian breeding habitats. Rasanen and Green (Chapter 11, this volume) analysed in detail the effects and impacts of acidification on amphibians. IV. ENVIRONMENTAL EXPOSURE OF AMPHIBIANS TO NITROGEN POLLUTION

Many amphibian habitats suffer nitrogen pollution. Nitrate concentrations above 3 mg NIL, for example, reflect anthropogenic contamination (Madison and Brunett 1985) and are often found in ponds and small streams in agricultural or urban areas. In the Great Lakes area of North America, 19.8% of 8 545 water samples had nitrate concentrations that can be harmfbl for amphibians (Rouse et al. 1999). Several field studies have detected negative effects of water pollution on amphibians, mainly due to agricultural runoff. In many of these studies, forms of reactive nitrogen were present at high concentrations. These types of chemicals, however, are rarely the only contaminants present in natural habitats and, consequently, it is difficult to establish a causeeffect relationship between the concentration of a given contaminant and a given impact on amphibians or other animals in a field study. Berger (1989) considered that an excess of nitrogen fertilizers caused the decline of several amphibian species in an agricultural area in Poland. Studying the water quality and frogs at the Klamath Basin National Wildlife Refuge (California), Boyer (1993) concluded

3152

AMPHIBIAN BIOLOGY

that un-ionized ammonia may be contributing to the decline of native frog populations. De Solla et al. (2002) found that the hatching success of Rana aurora and Ambystoma gracile was significantly lower at all agricultural sites compared to reference sites and ammonia was one of the measured substances that showed higher, and toxic, concentrations at the agricultural sites (up to 1.27 mg NIL). Bishop et al. (1999) found a significant negative effect of agricultural runoff on anuran development, density and diversity in Canada. In that study, ammonia and total nitrogen were highest in agricultural zones and may have affected anuran survival. Amphibian densities were significantly higher upstream and downstream than in the agricultural area. Incubating the eggs of the American toad in water from the different areas, Bishop et al. (1999) found a significant risk of abnormality associated with increasing nutrient levels. For every 0.1 unit increase in ammonium concentration, the risk of American toad eggs not hatching rose 48%. Similarly for green frog eggs, the significant risk factor was a 68% higher chance of not hatching for every 0.1 mgll increase in ammonia (Bishop et al. 1999). In an investigation of the relationships between amphibian distribution and wetland characteristics, Weyrauch and Grubb (2004) found that ammonium concentration in water was an important variable for predicting species richness of newts and salamanders. In a field survey of 180 ponds and thirteen amphibian species in southwestern Ontario, Canada, Hecnar and M'Closkey (1996) found that amphibian richness was not affected by nitrate concentration and there was a slight positive relationship between ammonium concentration and amphibian diversity. Maximum levels of nitrate and ammonium measured in the field were respectively 17.5 and 4.0 mg NIL. Brodman et al. (2003) also found a significant effect of the presence of ammonium in the water on the abundance of pond-breeding amphibians in northwestern Indiana. Ensabella et al. (2003) found that the selection of breeding sites by Bufo viridis in Rome was related to nitrate concentration. Nitrate concentration where the toads were breeding was significantly lower than the mean concentration from areas where that species was not found. Spray irrigation with secondarily treated, chlorinated wastewater effluent on Pennsylvania State Game Lands during 14 years negatively affected the reproductive success of three amphibian species, compared to natural ponds. In situ hatching success of eggs and the survival of larval Rana sylvatica, Ambystoma jeffersonianum and A. maculatum were lower in ponds with the addition of wastewater than in natural ponds (Laposata and Dunson 2000). The wastewater effluent had high concentrations of ammonium (11.93 mg NIL) and moderate levels of nitrate (2.26 mg NIL). In the ponds the mean concentration of nitrate was 3.426 mg NIL during amphibian breeding. The levels of nitrate were higher than later in the season. The authors did not believe that the levels of nitrate affected amphibians. However, they did not show data for ammonia in the ponds, but levels were probably high and toxic for eggs and tadpoles. V. PHYSIOLOGICAL EFFECTS OF REACTIVE NITROGEN ON AMPHIBIANS

The effects of environmental levels of reactive nitrogen on amphibians have been studied in several species. Unfortunately, however, in many cases the dynamics of these pollutants, the mechanisms that cause such effects or the environmental factors that influence toxicity are not well studied for amphibians. Studies of other organisms can help to understand these important questions. A. Ammonia Ammonia is the main excretory product that aquatic amphibians use to eliminate nitrogenous wastes (Balinsky 1970). Aquatic stages excrete ammonia both by diffusional exchange across the skin and gills and via urine (Boutilier et al. 1992). Ammonia is a highly diffusible and toxic substance and large volumes of water are necessary for its elimination. For that reason the excretion of this substance on land is more difficult and terrestrial amphibians have developed the ability to produce and excrete urea (Shoemaker et al. 1992).

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3153

Biological membranes are readily permeable to ammonia so this substance equalizes between the intracellular and extracellular compartments (Boutilier et al. 1992). External ammonia may enter an amphibian's body through the skin, gills and lungs (United States EPA 1986). The accumulation of ammonia due to physiological problems can be very toxic for amphibians. Moreover, toxicity of ammonia has been described in captive amphibians as due to the excessive accumulation of nitrogenous wastes by inadequate filtration or insufficient water changes (Crawshaw 2000). The exposure of tadpoles and adult amphibians to moderate, non-toxic levels of ammonia can initially cause a reversion in the ammonia difhsion gradient and animals can take up ammonia from the water (Wright and Wright 1996). In these cases, increased urea synthesis and excretion is a common mechanism of detoxifying ammonia. After 2-3 days of exposure, both ammonotelic and ureotelic bullfrogs respond to environmental ammonia by increasing the rate of urea excretion without increasing the activities of ureagenic enzymes (Wright and Wright 1996). Exposure of adult Xenopus to moderate levels of ammonia also increases urea synthesis and urea excretion rates (Janssens 1972). Schmuck et al. (1994) reported that at high ambient concentrations of ammonium (5 mg N-NH,+/L), early developmental stages of Bufo bufo and Hyperolius uiridiJlauus ommatosticus synthesized and excreted detectable amounts of urea; the urea excretion rate in premetamorphic stages of Hyperolius marmoratus taeniutus, however, were always below the detection limit. These authors also demonstrated that the activity of the ammonia-consuming ornithine cycle in synthesizing urea was regulated by environmental ammonia in some amphibian tadpoles. The rate of ammonium uptake from contaniinated water, however, can be greater than the combined rate of ammonia and urea excretion. Amphibians can accumulate ammonia in liver and muscle (Wright and Wright 1996). Un-ionized ammonia (NH,) is the principal toxic form of ammonia. It is uncharged and lipid soluble, whereas the permeability of plasma membranes to charged, hydrated ammonium ions is relatively low. It has been reported to be acutely toxic to freshwater invertebrates (Hickey and Vickers 1994; Alonso and Camargo 2003), amphibians (Dejours et al. 1989a; Jofre and Karasov 1999) and fish (Alabaster and Lloyd 1982; Tomasso and Carmichael 1986; United States EPA 1986). The predominant route of entry of un-ionized ammonia into organisms is via the gills. Un-ionized ammonia molecules can readily pass through cell membranes at the gill surface (Hampson 1976). Several modes of action have been suggested for ammonia toxicity. These include disturbance of electrochemical gradients (Hawkins et al. 1973; Arillo et al. 1981; Smart 198l), acid-base disturbance (Twitchen and Eddy 1994), gill damage that can directly affect osmoregulation (Cardoso et al. 1996; Rebelo et al. 1999; Lease et al. 2003) and a decrease in glutamate, a potential neurotransmitter, in the brains of some animals Fedel et al. 1998). Acute exposure of fish to ammonia can cause loss of equilibrium, an increase in gill ventilation, hyperexcitability, increased cardiac output and oxygen uptake and, in extreme cases, convulsions, coma and death. Chronic exposures can cause a reduction in hatching success, reduction in growth rate and morphological development, and pathologic changes of gills, liver and kidneys (United States EPA 1986). Some proposed mechanisms for the effect of un-ionized ammonia on growth in fish are reduction of oxygen uptake due to gill damage, imposition of additional energy demand caused by the use of alternative detoxification pathways, increased loss of ions by increased urine flow, inhibition of sodium uptake, and damage to various tissues (Colt and Tchobanoglous 1978). Several factors such as dissolved oxygen concentration, temperature, salinity or carbon dioxide concentration are known to affect ammonia toxicity (Downing and Merkens 1955; Bower and Bidwell 1978) but the best-studied interaction is with pH (Table 1). Basic pH is associated with a larger proportion of un-ionized ammonia (Emerson et al. 1975) and thus with a stronger toxic effect of ammonium on freshwater organisms (Dejours et al. 1989a). Moreover, the acute toxicity of NH3 has been shown to increase as pH decreases. Higher temperatures are also associated with a higher concentration of un-ionized ammonia but, at the same time, the toxicity of a given concentration of ammonia to Xenopus Zaevis increases

3154

AMPHIBIAN BIOLOGY

at lower temperatures (Dejours et al. 198913). Hyperoxia decreased ammonia toxicity to Xenopus laevis (Dejours et al. 198913). Hypoxic conditions increased the negative effects of un-ionized ammonia for many other aquatic species (Merkens and Downing 1957; Allan et al. 1990). Some bacterial infections can reduce the uptake of nitrite and the level of methaemoglobin in fish (Tucker et al. 1984). Table I. EPA Chronic (CCC) and acute (CMC) criterion concentrations of total ammonia (mg NIL) for freshwater life. ELS = early life stages. Modified from United States EPA (1998). -

--

--

CCC

PH 6.5 7.0 7.5 8.0 8.5 9.0

8°C 10.1 9.00 6.64 3.70 1.66 0.740

14°C 6.89 6.1 1 4.51 2.52 1.13 0.503

CMC Salmonids Salmonids present absent

Fish ELS present

Fish ELS absent 14°C 6.67 5.91 4.36 2.43 1.09 0.486

20°C 4.68 4.15 3.06 1.71 0.765 0.342

26°C 3.18 2.82 2.08 1.16 0.520 0.232

32.6 24.1 13.3 5.62 2.14 0.885

48.8 36.1 19.9 8.40 3.20 1.32

B. Urea

The mechanisms of urea toxicity in amphibians are not well understood. Urea is soluble in water and in freshwater fish it penetrates into the tissues and induces alterations of the skin and gastric lining (Srivastava and Srivastava 1979). The exposure of freshwater fish to sublethal levels of urea (80 ppm) also causes increased total free-sugar levels and cholesterol levels in the blood and decreased levels of total sugar in the liver and muscle. As a result of urea exposure, the mucous cells of the gill epithelium degenerate and the hepatic cells and tissues suffer necrosis (Balasubramanian et al. 1999). There are two other possible mechanisms to explain the acute toxicity of urea to amphibians. Urea-fertilized soils may become saline causing osmoregulatory stress (Marco et al. 2001). The skin of amphibians behaves as a semi-permeable membrane and the osmotic flow of water is proportional to the osmotic gradient between the medium and the animal's body fluids (Shoemaker et al. 1992). Relatively few species can survive well in salinities that exceed the normal osmotic concentrations (Balinsky 1981). Licht et al. (1975) found that the salinity tolerance of salamanders naturally exposed to saline conditions (Batrachoseps sp.) was higher than the tolerance of inland salamanders. Salamanders from inland populations were only able to acclimatize to water solutions that were iso-osmotic with their blood. The reduced water potential that results from increased urea concentrations in the substrate and the air may impede the maintainiance of water balance Umgensen 1997). Larval amphibians generally appear to be less tolerant of saline conditions than are adults, because the larvae have a lower capacity for synthesis and retention of urea (Balinsky 1981). Terrestrial amphibians that inhabit arid or hypersaline environments are able to double the sodium and chloride concentrations of their plasma or to produce and store urea as a mechanism preventing or slowing down the cutaneous loss of water from the body to a dry environment (Shoemaker et al. 1992). The rate at which fossorial species produce and store urea is inversely proportional to the soil water potential (McClanahan 1972). Some urea production is also used to detoxify the ammonia produced from protein metabolism (Shoemaker et al. 1992). Alternatively, urea may diffuse directly into the organism and be transformed into ammonium and un-ionized ammonia, substances that are highly toxic. Environmental urea can also be transformed into ammonia in the substrate and in the air. Hydrolysis of urea applied to forests may increase the soil pH in the forest floor, thereby producing toxic ammonia (NH,) that can easily cross cell membranes and disrupt metabolism in bacteria (Kernaghan et al. 1995). An excess of urea can also result in methohemoglutanemia (Rouse et al. 1999).

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3155

C. Nitrate and Nitrite

An excess of nitrate in freshwater ecosystems is known to be toxic not only to amphibians (Baker and Waights 1993; Marco et al. 1999) but also to their prey and predators, such as molluscs (Alonso and Camargo 2003), crustaceans (Muir et al. 1990; Scott and Crunkilton 2000), insects (Camargo and Ward 1992) and fish (Westin 1974). Nevertheless, nitrate in moderate amounts is relatively harmless to animal life, and is generally not considered to present a toxic hazard in surface waters (Russo 1985); it is rapidly excreted in the urine. Nitrates do, however, create health problems when reduced to nitrites (Eddy and Williams 1994). Effects of nitrites on amphibians have not been studied extensively and it is necessary to look to other aquatic vertebrates for indications of potential responses. The biological effects of nitrite are the same whether it enters the body through ingestion, inhalation or cutaneous absorption, or is produced by bacterial conversion from nitrate (Kross et al. 1992). Environmental nitrite is concentrated in the blood of aquatic vertebrates by an active transport system in the gills (Perrone and Meade 1977; Lewis and Morris 1986). Bath and Eddy (1980) suggested that fish can transport nitrite against a concentration gradient via a branchial anion exchange mechanism. Moreover, if nitrate is ingested with water or vegetation, it is rapidly reduced to highly reactive nitrite in the large bowel and then enters the bloodstream. In the blood, nitrite reacts with haemoglobin and oxidizes ferrous iron to ferric iron, thus producing methaemoglobin (United States EPA 1986). Methaemoglobin cannot bind or transport oxygen, thereby causing tissue hypoxia (OECD 1986; United States EPA 1986). As exposure time or exposure concentration increases, the proportion of methaemoglobin relative to hemoglobin in the blood increases (Scarano et al. 1984). If methaemoglobin concentration becomes high, animals can suffer methaernoglobinemia (Dappen 1982), also called "blue baby syndrome". In humans, affected infants may present asymptomatic cyanosis, which can progress to dyspnea and lethargy or coma (Kross et al. 1992). From 1945 to 1992, about 2 000 cases of clinical methaernoglobinemia in young infants were reported, with an estimated death rate of 10%. Nitrite induced methaernoglobinemia can result in tissue anoxia and death in fish (Cameron 1971). Methaemoglobinemia has been reported in larval amphibians (Huey and Beitinger 1980a, 1980b). This physiological alteration imparts a distinctive brown colour to the blood in vertebrates (Bodansky 1951). In mammals, early life stages are more susceptible to nitrate toxicity than are older stages. In humans, fatalities are rare, but subacute methaemoglobinemia can be asymptomatic while affecting development, making the condition particularly insidious. Sublethal methaernoglobinemia leads to reduced vitality, increased stillbirth, low birth weight and slow weight gain in livestock (National Research Council 1972). The pH of the digestive system and the composition of the enteric bacterial assemblage may influence susceptibility to methaernoglobinemia. Treatment for humans consists of administration of oxygen and intravenous and oral methylene blue (Kross et al. 1992). Nitrite-induced methaernoglobinemia has been studied in detail in freshwater fish (Brown and McLeay 1975; Smith and Russo 1975; Lewis and Morris 1986). Margiocco et d- (1983) found up to 60 times magnification of nitrite in blood of rainbow trout exposed to nitrite in fresh water. Seawater fish are also sensitive to nitrite, although chloride partially inhibits nitrite toxicity (Crawford and Allen 1977; Tomasso et al. 1979). In salt water, apparently, there is a competitive inhibition of chloride toward nitrite uptake that may p \ i d e protection for marine fish and crayfish from nitrite (Scarano et al. 1984). The amount of dissolved oxygen and the concentration of calcium may also influence the effects d nitrite on aquatic species (Tucker and Schwedler 1983; Lewis and Morris 1986). Chloride, dissolved oxygen and calcium concentration should be monitored and controlled in toxicological studies of the effects of nitrate or nitrite on amphibians. The overall mean percentage of methaemoglobin in fish with no exposure to nitrite is Amut 5% (Tucker et al. 1989). Channel catfish exposed to 0.92 mg/L N-nitrite did not b m e anemic and developed sublethal methaemoglobinemia (Tucker et al. 1989). Fish

3156

AMPHIBIAN BIOLOGY

mortality may begin when haemoglobin concentration drops to 50% of control values in sea bass (Scarano et al. 1984) alhough some fish can survive higher concentrations (Tucker and Schwedler 1983; Huertas et al. 2002). Some fish suffering methaemoglobinemia show brown blood and gills and these tissues turn pink when fish die, but colour is not a reliable indicator of stress in sea bass (Scarano et al. 1984). Temperature has an effect on methaemoglobin formation during nitrite exposure. Higher temperatures increase the rate of haemoglobin oxidation but also increase the rate of haemoglobin recovery after the exposure to nitrite (Huey et al. 1984). The major methaemoglobin-reducing system in fish and mammals is NADH-methaemoglobin reductase (Huey and Beitinger 1981). Fish can acclimate to sublethal levels of nitrite and laboratory studies can overestimate the methaemoglobin ratios of fish chronically exposed in the field to high levels of nitrite (Tucker and Schwedler 1983). Such ability to acclimate to nitrite could be shared by amphibian aquatic stages. Methaemoglobinemia has also been reported in various reptiles, including crocodilians (Gruca and Grigg 1980) and freshwater turtles (Sullivan and Riggs 1964) and in birds (Board et al. 1977). Nitrite can also oxidize the haemoglobin protein, causing denaturation and erythrocyte haemolysis, resulting in haemolytic anaemia (Kross et al. 1992). Exposure to nitrite causes hemolythic anemia in sea bass (Scarano et al. 1984), rainbow trout (Margiocco et al. 1983) and channel catfish (Tucker et al. 1989). The concentration of nitrite in blood seems to be linked to exposure time more than to physiological conditions, whereas in the liver and brain of trout much greater differences exist between seemingly unstressed animals and torpid, unreactive ones (Margiocco et al. 1983). Death following acute intoxication apparently depends on the toxic action of nitrite in vital organs, rather than on methaemoglobinemia (Margiocco et al. 1983). Nitrite can produce cardiovascular effects, such as reduced heart rate, in fish embryos (Williams and Eddy 1989). Nitrates may reduce feeding efficiency. Amphibian tadpoles are largely herbivorous and they have active symbiotic gut bacteria involved in digestion (Duellman and Trueb 1994). These can be affected by nitrate ingestion (Hecnar 1995). Nitrates in the acidic environment of the gut may ultimately be converted into nitrosamines, which are carcinogenic (Committee on Nitrate Accumulation 19'72). Chronic consumption of high levels of nitrate could cause some cancers and have teratogenic effects (Magee 1971; Lijinsky et al. 1972; Bruning-Fann and Kaneene 1993). NOx substances have a clear genotoxic effect on somatic human chromosomes. Mitotic index, chromosome aberrations, sister chromatid exchanges and satellite associations all show a significant increase when exposed to NOx (Yadav and Seth 1998). There are at least three mechanisms by which intracellular elevated NOx could exert genotoxic affects. These include formation of carcinogenic N-nitroso compounds, direct deamination of DNA bases and oxidation of DNA after formation of peroxynitrite and/or hydroxyl radicals (Liu and Hotchkiss 1995). However, teratogenic effects attributable to nitrate or nitrite ingestion are still controversial. Although it has been difficult to prove conclusively, evidence is mounting that nitrates in drinking water may be linked to the development of certain cancers such as bladder cancer in older women (Weyer et al. 2001). There is no direct evidence of a genotoxic effect on amphibians due to dissolved inorganic nitrogen. Rana clamitans tadpoles, however, showed significant annual variation in DNA damage that was greater in samples of tadpoles collected from agricultural areas than from non-cultivated ones (Ralph and Petras 1997). Hecnar (1995) pointed out a possible effect of nitrates affecting gene expression or developmental processes in Pseudacris triseriata. The possibility that high levels of DNA damage in tadpoles collected from agricultural areas may be caused by an excess of fertilizers should be investigated. Dappen (1982) suggested the possibility that nitrate stress may depress the immune response of amphibians. Amphibian tadpoles and adults exposed to nitrate had lower numbers of circulating white cells and lower haemoglobin values and there was a response in weight of lymphatic tissues (Dappen 1982).

MARC0 and ORTIZ-SANTALIESTRA. NITROGEN POLLUTION

3157

The European Union established limits of 11.3 mg NIL for nitrate and of 0.15 mg NIL for nitrite as the criteria for water to be of drinking quality (European Council 1998). In the United States, the EPA recommends a limit of 10 mg NIL for nitrate and 1 mgN1L for nitrite for drinking water. Levels of nitrate below 90 mg NIL and of nitrite below 5 mg N/ L seem to have no effect on warm-water fish (McCoy 1972; Knepp and Arkin 1973). Levels below 1 mgN1L of nitrate (Westin 1974) and 0.06 mg NIL of nitrite (Russo and Thurston 1977) should not be toxic for salmonids. Lethal concentrations of nitrate for a number of anuran species are in the range of 13-40 mgIL, with chronic effects occurring at concentrations below 10 mg/L. Water-quality data for the agricultural and urban areas sampled in North America show that nitrate concentrations in surface waters exceed these critical toxicity levels for amphibians for extended periods of time and during sensitive times of anuran development (egg and tadpole stages) (Rouse et al. 1999). VI. DOSE-EFFECT RELATIONSHIPS

Amphibians are considered good indicators of environmental quality; specifically, they are sensitive to many kinds of environmental pollution (Boyer and Grue 1995). Some studies have compared sensitivity of amphibians to several types of pollutants with that of both other vertebrates and invertebrates; amphibians sometimes were the most sensitive (Holcombe et al. 1987; McCrary and Heagler 1997). A. Ammonia 1. Effects on Embryos

Ammonia had a negative effect on the embryonic development of green frogs (Rana clamitans) and leopard frogs (Rana pipiens). Green frogs were the most sensitive and suffered a significant mortality at an ammonium ion concentration of 6.1'7 mg/L (Jofre and Karasov 1999). However, neither the survival nor the prevalence of deformities of American toad embryos (Bufi amencanus) was affected by target ammonium ion concentrations of 6.17 mg/L (Jofre and Karasov 1999). Tietge et al. (2000) reported complete mortality of African clawed hog (Xenopus laevis) embryos exposed to concentrations of 53.4-55.7 mg/L total ammonia in different test solutions. Diamond et al. (1993) reported a NOEC of 0.27 mg/l un-ionized ammonia in leopard frog embryos and a LC50 for embryos of this species of 1.9 mg1L NH,. Spring peeper (Hyla crucifer) embryos were also very sensitive to low levels of unionized ammonia (Diamond et al. 1993). Schuytema and Nebeker (1999b) found that concentrations of un-ionized NH, as low as 0.06 mg/L had an effect on Xenopus laevis embryos. Amphibian eggs seem to be much more tolerant of ammonia than are fish. Egg capsules or the gelatinous matrix may protect amphibian embryos from this chemical. For the fish Micropterus treculi, the ammonia-N 96-h LC50 was 12.7 mg1L of total ammonia (Tomasso and Carmichael 1986). In salmonids, LC50 values for NH, ranges between 0.083 and 1.09 mg/L whereas for non-salmonids the range is 0.14-4.60 mg/L (United States EPA 1985). The highest concentration shown not to depress hatching in Pimephales promelas was 0.42 mg NH3/L (Swigert and Spacie 1983) and Salmo gairdnierii embryos exhibited malformations when exposed to NH, concentrations between 0.01 and 0.2 mg1L (Cotta Ramusino 1980). The 19 species of invertebrates for which data have been reported have higher LC50 values (0.53 to 22.8 mg/L) than do fish (US EPA 1989). 2. Effects on Larvae As expected, un-ionized ammonia is much more toxic to amphibians than ionized ammonium (Table 2) and the levels of toxicity of NH, to larvae are similar to those recorded for warm-water fish. There is no available information about the effect on amphibians of a long-term exposure (> 10 days) to ammonium but short-term exposures reflect a significant impact of moderately high levels of ammonium on the survival of amphibian larvae.

3158

AMPHIBIAN BIOLOGY

Table 2. Comparative toxicity of different chemical forms of ammonium to embryonic and larval amphibians.

Species

Stage/Agea

Days of expos.

Ammonium source

Ambystoma gracile

5 week old

10

Xenopus laeuis

26-27

Ammonium sulfate Ammonium sulfate Ammonium chloride Ammonium sulfate Ammonium chloride Ammonium chloride Ammonium sulfate Ammonium sulfate Ammonium sulfate Ammonium sulfate Ammonium sulfate Ammonium chloride Ammonium chloride Ammonium sulfate Ammonium sulfate Un-ionized ammonia Un-ionized ammonia Un-ionized ammonia Un-ionized ammonia Ammonium bicarbonate Un-ionized ammonia Un-ionized ammonia Ammonium sulfate

4

Pseudacris regilla

6 week old

10

9 week old

Hyla crucqer

12 (Rugh)

4

12 (Rugh)

10

12 (Rugh)

4

12 (Rugh)

10

8 days old

4

Bufo americanus Rana pipiens

5-6

5

New

4

25

4

Rana clamitans

Rana aurora (a)

20 days old

110

4 week old

10

Effect Decreased size LC50

Decreased size Decreased size

No lethal effects Significant moratlity LC50 50% mortality Significant mortality Significant mortality Decreased size

Reference Nebeker and Schuytema (2000) Schuytema and Nebeker (1999a) Schuytema and Nebeker (1999a) Schuytema and Nebeker (1999a) Schuytema and Nebeker (1999a) Schuytema and Nebeker (1999b) Schuytema and Nebeker (1999b) Schuytema and Nebeker (1999a) Schuytema and Nebeker (1999a) Nebeker and Schuytema (2000) Nebeker and Schuytema (2000) Schuytema and Nebeker (1999b) Schuytema and Nebeker (1999b) Schuytema and Nebeker (1999b) Schuytema and Nebeker (1999b) Diamond et al. (1993) Jofre and Karasov (1999) Jofre and Karasov (1999) Diamond et al. (1993) Sparling and Harvey (2006) Jofre and Karasov (1999) Jofre and Karasov (1999) Nebeker and Schuytema (2000)

Stage or age of the test animals at the beginning of the exposure

Green frog tadpoles (Rana clamitans) exposed to target total ammonia concentrations of 6.1'7 mg/L suffered a significant mortality compared to controls. A target concentration of 3.09 mg/L, however, had no effect on tadpole survival of this species (Jofre and Karasov 1999). At the highest concentration, deaths began after 20 days of exposure and the sensitivity to ammonia differed among different egg masses. Tadpole growth was also negatively affected by the highest ammonium concentration but there were no malformations (Jofre and Karasov 1999). B. Ammonium Nitrate

Toxicity of ammonium nitrate has been mainly attributed to ammonium cations rather than to nitrate anions (Schuytema and Nebeker 1999a,b,c;Johansson et al. 2001). The levels of NO3--Npresent in solutions containing sufficient NH,+-N to cause adverse effects were

MARC0 and ORTIZ-SANTALIESTRA:NITROGEN POLLUTION

3159

insufficient to have an effect on larval survival or growth (Schuytema and Nebeker 1999~). Thus, in exposures to ammonium nitrate, the ammonium ion would be more toxic than the nitrate ion and would be the primary cause of mortality or sublethal effects. Nevertheless, the vast use of ammonium nitrate as fertilizer around the world has stimulated a great deal of work on the effects of this compound on amphibians (e.g., Berger 1989; Hecnar 1995; Xu and Oldham 1997; Ortiz et al. 2004). 1 . Effects on Embryos

Schuytema and Nebeker (1999b) exposed embryos of Pseuducms reg~llaand Xenopus laevis to ammonium nitrate in different experiments. Ammonium nitrate at levels of 50.9 mg NH4+-NILproduced 80% mortality in El regilla embryos after four days of exposure. At day 10, mortality at this level was 100%. Complete mortality was also observed in X. laevis embryos exposed to 101.2 mg NH4+-NILafter five days of exposure. Levels as low as 6.9 mg NH4+-NILproduced negative effects on size and weight of both species (Schuytema and Nebeker 1999b). Using FETAX solutions, complete mortality of embryos in X. laevis was detected at 50.9 mg NH,+-NIL after four days of exposure, and reducton in size and weight was observed at exposures of 25.1 mg NH,+-NIL. 2. Effects on Larvae

Exposure of newt larvae to high levels of ammonium nitrate causes their death (Watt and Jarvis 1997). Sublethal levels can cause a significant reduction in size at metamorphosis (Watt and Oldham 1995). Ammonium nitrate produced a mortality of over 95% in Pseudacris regilla and Xenopus laevis tadpoles exposed to 99.5 mg N-NH4+/Lfor ten days (Schuytema and Nebeker 1999a). Levels greater than 49 mg/L caused delayed growth in El regilla (Schuytema and Nebeker 1999a). The exposure to ammonium nitrate of Rana temporaria tadpoles at Gosner stage 25 caused a significant mortality at 72 hours at a concentration of 100 mg N-NO,-/L. A concentration of 50 mg N-NO,/L had no acute effect on tadpole survival (Johansson et al. 2001). Hecnar (1995) reported a decrease in survival of larvae of several species with increased levels of exposure to ammonium nitrate. Reduced growth and activity and occurrence of abnormalities were also observed in individuals exposed to the pollutant. Hamer et al. (2004) also detected a strong sensitivity of the endangered frog Litoriu aurea to low levels of ammonium nitrate. Some species seem to be very tolerant of ammonium nitrate while others show toxic responses at normal environmental levels (Table 3). Such differences, sometimes even within the same species but in different experiments, may be due to ontogenic, geographic or genetic variation in sensitivity. Water temperature, pH and chloride concentration can also have a significant effect on sensitivity to ammonium ions and may explain part of the variability in sensitivity detected among and within species. 3.

Effects on Adults

Granular ammonium nitrate fertilizer was acutely toxic to Rana temporaria at concentrations well below those recommended for field application. This fertilizer is very soluble and loses its effect within three hours of being dissolved in the soil. Fertilizers quickly affected frog ventilation rates. In field trials, all the frogs showed symptoms of toxicity after 3-1 10 minutes from the start of exposure. Affected animals were immediately removed and washed but they still usually died. The EC50** of frogs with acute exposures (less than 24 h) was 3.6 g/m2 when over a paper substrate, whereas EC50** on a soil substrate was 6.9 g/m2 (Oldham et al. 1997). Schneeweiss and Schneeweiss (1997) also found lethal and sublethal effects of ammonium nitrate fertilizers on amphibians when they migrated over recently fertilized substrates.

3160

AMPHIBIAN BIOLOGY

Table 3 . Comparative toxicity of ammonium nitrate to amphibian eggs and larvae. LC5,= Medium Lethal Concentration; NOEC = No Observed Effect Concentration.

Species

Stage/Agea

Pleurodeles waltl Triturus helveticus

10-12 42 days 42 days

?: vulgaris Discoglossus galganoi Pelobates cultripes Bufo americanus B. bufo

2? 10- 12 10-12 25 10-12 32 32 ?

B. calamita Xenopus laevis Hyla arborea Pseudacris regilla

I? triseriata Litoria aurea Crinia signqera Lymnodynastes peronii Rana aurora R . clamitans R . esculenta R . lessonae R . pipiens R . ridibumh R. temporaria

10-12 26-27 26-27 10-11 10-12 26-27 26-27 12 (Rugh) 12 (Rugh) 25 25 25 25 11-12 11-12 25 ? ?

23 ?

G25

Days of expos.

Effect

15 6 6 3 15 15 4 15 4

Decreased size Significant mortality 100% mortality Decreased size LC50 Decreased size

4 4 4 10 5 8 4 10 4 10 4 150 21 91 16 16 4 3-4 3-4 4 3-4 3

Significant mortality Decreased size Lcso LC,, LC50 LC50 LC50 LC,, LC50 LC,,

Lc50

LC5, LC50

Lc50

Significant mortality No mortality No mortality LC5,, Decreased size LC50 100% mortality 100% mortality LC50 100% mortality Near 90% mortality

(mg NO;/L) 200 155 387.5 155 193.5 50 60.2-1 74 254.6 1704 1637 15.5 200 446 243.3 193.9 33.1 599.6 244.5 181.9 110.7 75.3 7.75 11.63 11.63 318.3 58.44 143.5 15.5 15.5 100.1 15.5 443

Reference Ortiz et al. (2004) Watt and Jarvis (1997) Watt and Jarvis (1997) Watt and Oldham (1995) Ortiz et al. (2004) Ortiz et al. (2004) Hecnar (1995) Ortiz et al. (2004) Xu and Oldham (1997) Xu and Oldham (1997) Berger (1989) Ortiz et al. (2004) Schuytema and Nebeker (1999a) Schuytema and Nebeker (1999a) Schuytema and Nebeker (1999b) Ortiz et al. (2004) Schuytema and Nebeker (1999a) Schuytema and Nebeker (1999a) Schuytema and Nebeker (1999b) Schuytema and Nebeker (1999b) Hecnar (1995) Hamer et al. (2004) Hamer et al. (2004) Hamer et al. (2004) Schuytema and Nebeker (1999c) Schuytema and Nebeker (1999c) Hecnar (1995) Berger (1989) Berger (1989) Hecnar (1995) Berger (1989) Johansson et al. (200 1)

?)Stage or age of the test animals at the beginning of the exposure.

C. Nitrite 1. Effects on Embryos

Nitrite exposure significantly affected hatching time and developmental stage at hatching for Ambystoma tigrinum, whereas no effects on embryonic development were reported for Rana syluatica (Griffis-Kyle 2007). In fish, eggs seem to be very tolerant of nitrite. The 24-hour LC50 value for Salmo salar eggs was of 3 290 mg/L N- NO,- (Williams and Eddy 1989). Long-term exposure to 14 mg/L N- NO< delayed hatching in Salmo salar and retarded embryonic growth and development (Williams and Eddy 1989). 2. Effects on Larvae

The few experiments conducted to elucidate the direct toxicity of nitrite to amphibians show strong effects at low levels of this chemical (Table 4). In some cases, amphibian larvae were negatively affected by concentrations of nitrite below the recommended limit for drinking water. The 96-hour LC50 for Ambystoma tigrinum in water with low chloride (5.0 mg/ L) was 0.33 mg/L nitrite. Nearly 100% mortality was observed in larvae exposed to 0.76 mg/L. Exposure to high levels of nitrite causes excessive mucus production and gulping of air. However, Ambystoma texanum larvae exposed to 3 mg/L N-nitrite in water with a concentration of chloride of 300 mg/L suffered no mortality (Huey and Beitinger 1980b). This tolerance to nitrite in the presence of high levels of chloride is probably related to lower nitrite uptake. Rana pretiosa, Rana aurora, Bufo boreas, Hyla regilla and Ambystoma gracile tadpoles were very sensitive to nitrite. After 15 days of exposure to 0.44 mg N-NO
MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3161

Table 4 . Comparative toxicity of nitrite to amphibian eggs and larvae. LC,, = Medium Lethal Concentration; LOEC = Lowest Observed Effect Concentration; NOEC = No Observed Effect Concentration.

Species Ambystoma gracile A. texanum

Stage/Agea Newly hatched Newly hatched

Days of expos. Nitrate source 4 15 4 4

A. tigrinurn

Newly hatched

21

Bufo boreas

Newly hatched Newly hatched Newly hatched Newly hatched Newly hatched Newly hatched G 3940 G26 Newly hatched Newly hatched Newly hatched

4 15 4 15 4 15 14 15

Pseudacris regilla Rana aurora R. cascadae R. catesbeiana R.pretiosa R.syluatica

4

15 21

Sodium nitrite Sodium nitrite Sodium nitrite Low chloride Sodium nitrite High chloride Sodium nitrite Sodium Sodium Sodium Sodium Sodium Sodium Sodium Sodium Sodium Sodium Sodium

nitrite nitrite nitrite nitrite nitrite nitrite nitrite nitrite nitrite nitrite nitrite

Effect

(mg NNO;/L)

Reference

LC,, LC,, LC,o

1.90 1.01 0.33

Marco et al. (1999) Marco et al. (1999) Huey and Beitinger (1980b)

No mortality Slowed development LOEC LC,, LC,, LC,, LC,, LC, Lower growth NOEC LC,, LC,, Slowed development

3

Huey and Beitinger (1980b)

0.3-6.1

Griffis-Kyle (2007)

3.5 1.75 5.50 1.23 5.59 1.19 3.5 10 6.82 0.57 0.3-6.1

Marco et al. (1999) Marco et al. (1999) Marco et al. (1999) Marco et al. (1999) Marco et al. (1999) Marco et al. (1999) Marco and Blaustein (1999) Smith et al. 2004 Marco et al. (1999) Marco et al. (1999) Griffis-Kyle (2007)

(") Stage or age of the test animals at the beginning of the exposure

No decrease in total haemoglobin was observed in larval bullfrog tadpoles (Rana catesbeiana) of 15-26 g exposed for 24 hours to 1.0-50 mg/l NO2--N(Huey and Beitinger 1980a). These authors did find, however, a strong positive relationship between methaemoglobinemia and nitrite concentration; the mean proportion of methaemoglobin over total haemoglobin ranged from 5.7% for controls to 51.0% at 50 mg/L of nitrite for experimentals. The percentage of methemoglobin in bullfrog tadpoles is relatively low compared to that of warm-water fish (Huey and Beitinger 1980a). Huey and Beitinger suggested that this low response to nitrite might be related to reduced nitrite uptake and/or to the presence of an efficient metbaemoglobin reductase system in tadpoles. Bullfrog tadpoles show gill ion absorption similar to that of fish (Alvarado and Kirschner 1963) but large tadpoles have limited surface area of the gills, compared to fish, resulting in a lower nitrite uptake and a lower dose response to nitrite. Low levels of chloride provide an efficient protection against methaemoglobimemia in bullfrog tadpoles (Huey and Beitinger 1980a). Smith et al. (2004) also found a significant tolerance of bullfrog tadpoles to nitrite. Fifteen days of exposure to a concentration of 10 mg/l NO2--Nhad no detectable effect. Larvae of many amphibian species seem to be more tolerant of nitrite than are salmonids but less tolerant than are warm-water fish. Lethal levels (96-hour LC50) for 6eshwater fish range from 8.22 mg/L of N-nitrite for channel catfish (Konikoff 1975) to 0.21 mg/L of N-nitrite for rainbow trout (Russo et al. 1974). Sensitivity of rainbow trout to nitrite is the highest reported for vertebrates. The 24-hour LC50 value for Salmo sahr early alevins was of 2 940 mg/L N-NO,, decreasing to 121.8 mg/L N-NO2 in late alevins and 10.4 mg/L N-NO,- in older stages (Williams and Eddy 1989). A significant reduction in total haemoglobin has been reported for rainbow trout in freshwater after 96 hours of exposure to 0.1 to 0.5 mg/l NO2--N(Brown and McLeay 1975; lZ1argiocco et al. 1983). The LC50 of fathead minnows of 0.9 to 3.3 g exposed to nitrite for 96 hours was of 147.4 mg/l NO?-N (Palachek and Tomasso 1984a). The 96-hour LC50 for nitrite-N was 187.6 mg/L for Micropterus treculi (Tomasso and Carmichael 1986) and 23.3 for channel catfish (Palachek and Tomasso 1984b).

AMPHIBIAN BIOLOGY

D. Nitrate 1. Effects on Embryos

Sodium nitrate produced complete mortality in Pseudacris regzlla embryos after four days of exposure at 2 716 mg NO,-IL. Size and weight of embryos after ten days of exposure to 111 mg NO,--NIL was significantly lower than values measured in controls (Schuytema and Nebeker 1999b). Xenopus laevis embryos exposed to nitrate for five days showed 100% mortality at 979.2 mg NO<-NIL and reduced weight at 56.7 mg NO,-NIL (Schuytema and Nebeker 1999b). There was no effect on hatching success of embryos of Rana sylvatica, Ambystoma jeffersonianum, A. maculatum and Bufo americanus at nitrate levels up to 40 mg/L (Laposata and Dunson 1998). Pre-hatch growth of Xenopus laeuis embryos was significantly decreased at nitrate concentrations as low as 56.7 mg/L (Schuytema and Nebeker 199913). 2. Effects on Larvae When nitrate ions were added to the water, some larvae of some species reduced feeding activity, swam less vigorously, showed disequilibrium and paralysis, suffered abnormalities and oedemas and eventually died (Hecnar 1995; Marco et al. 1999; Watt and Oldham 1995). Table 5. Comparative toxicity of nitrate to amphibian eggs and larvae. LC5, = Medium Lethal Concentration; NOEC = No Observed Effect Concentration.

Species Am bysoma jeffersonianum A. gracile A. rnacrodactylum A. maculatum Bufo americanw B. boreas B. bufo B. terrestris Xenopus laevis

Stage/Agea Harrison < 12

Days of expos. Nitrate source 17-25

Newly hatched Newly hatched Harrison < 12 <12 Newly hatched G25 G25 G25 10-1 1 (New)

G25 Newly hatched 15 Newly hatched 2 1 Pseudacris regilla 12 (Rugh) 10

H. regilla

12 (Rugh) Litoria caerulea Rana aurora

4

Xenopus laevis

Reference

NOEC

Potassium nitrate Sodium nitrate Sodium nitrate Sodium nitrate Potassium nitrate Sodium nitrate Sodium nitrate Sodium nitrate Sodium nitrate

Marco et al. (1999) Hatch and Blaustein (2003) Laposata and Dunson (1998) Laposata and Dunson (1998) Marco et al. (1999) Baker and Waights (1993) Baker and Waights (1993) Edwards et al. (2006) Schuytema and Nebeker (199913) NOEC 66 Sullivan and Spence (2003) NOEC 25 Marco et al. (1999) Decreased size 4.5 Hatch and Blaustein (2003) LC50 578 Schuytema and Nebeker (199913) LC50 643 Schuytema and Nebeker (199913) 50% of mortality 9.03 Baker and Waights (1994) 636.3 Schuytema and Nebeker (1999c) Decreased size 235 Schuytema and Nebeker (1999c) 85% inactive 4.5 Hatch and Blaustein (2000) 35% of mortality 20 Smith et al. (2005) 75% of mortality 20 Smith et al. (2005) Increased size 66 Sullivan and Spence (2003) 16.45 Marco et al. (1999) NOEC 9.03 Laposata and Dunson (1998) Decreased size 5 Johansson et al. (200 1) NOEC 1 000 Johansson et al. (2001) LC50 1 750 Schuytema and Nebeker (1999a) LC50 266.2 Schuytema and Nebeker (1999a) 3 666.4 Schuytema and Nebeker (1999a) LC50 1 236.2 Schuytema and Nebeker ( 1999a)

Sodium nitrate Potassium nitrate Sodium nitrate LC50 LC,,

16 16

Sodium nitrate Sodium nitrate

Gosuer 11-12

16

Sodium nitrate Sodium nitrate Sodium nitrate Sodium nitrate Sodium nitrate Potassium nitrate Sodium nitrate Sodium nitrate Sodium nitrate Sodium nitrate

Gosner 12 G26 G26 G25-26 Newly hatched 112 25 25 Pseudacris regilla 26-27

Conc. (mg NIL)

Sodium nitrate

25 Gosner 11-12

R . cascadae R . catesbeiana R . clamitam R . pipiem R . pretiosa R . sylvatica R . temporaria

Effect

26-27

10

Sodium nitrate

26-27

4

Sodium nitrate

26-27

10

Sodium nitrate

Lc50

Increased size NOEC NOEC NOEC 100% mortality 92% mortality Delayed growth LC50

?)Stage or age of the test animals at the beginning of the exposure.

9.03 Laposata and Dunson (1998) 23.39 2.26 9.03 9.03 25 22.6 9.03 30 438.4

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3163

The levels that affect amphibians vary among species (Table 5). Low concentrations may cause an increase in growth rate in some species but similar relatively low levels can cause deleterious effects in other species (Table 5). Some species were tolerant to very high and unrealistic levels of nitrate in the water. The observed effects increased with both time of exposure and concentration (Baker and Waights 1993, 1994; Marco et al. 1999). The 15-day LC50 was 16.45 mg N-NO,-/L for R. pretiosa and 23.39 mg N-NO,-/L for A. gracile (Marco et al. 1999). Rana catesbeiuna, R. pipiens and Xenopus laevis tadpoles and adults exposed for three weeks to 1 0 4 0 ppm of nitrate suffered a reduction of the immune response (Dappen 1982). Using sodium nitrate, 5 mg N-NOJL had no effect on survival but had an effect on growth rate and size at metamorphosis of Rana temporaria tadpoles at Gosner stage 25 (G25) exposed until metamorphosis (Johansson et al. 2001). In acute tests, over a 72-hour period, Rana temporaria tadpoles at G25 did not suffer a significant mortality at 1 000 mg N-NO,-IL (Johansson et al. 2001). Xenopus laevis larvae (G25) exposed to sodium nitrate concentrations of 66 mg N-NOflL did not suffer significant mortality and nitrate had a positive effect on mass at metamorphosis (Sullivan and Spence 2003). Tadpoles of Rana pipiens exposed from the firstfeeding stage through metamorphosis to 0, 5, and 30 mg N-NO3-ILexhibited no significant differences in developmental rate, metamorphic success, time to metamorphosis or size at metamorphosis, but nitrate slowed growth of larvae (Allran and Karasov 2000). Litoria caerulea suffered a mortality of 50%, a decrease in larval size and rate of development at 40 ppm of sodium nitrate (Baker and Waights 1994). Total mortality was recorded in larval Pseudacris regilla and Xenopus laevis exposed for ten days at 2 986.5 mg N-NO3-/Las sodium nitrate. Both species were significantly smaller .than controls when exposed to 259.1 mg NIL (Shuytema and Nebeker 1999a). There is great variability among species (Table 5). The most sensitive species showed sublethal effects at 20 mg NO,-/L and there was a significant mortality for these sensitive species at 40 mg NO,-IL. Several species, however, tolerated very high nitrate levels. The 96-hour LC50 values for nitrate in some warm-water fish are substantially higher than those observed in most amphibians; 8 858 mg1L as sodium nitrate for bluegill (Trama 1954), 6 200 mgIL as sodium nitrate for channel catfish (Ictalurus punctatus) (Colt and Tchobanoglous 1976) and 846 mg/L as potassium nitrate for guppy fry (Poecilia reticulatus) (Rubin and Elmaraghy 1977).

E. Urea I. Effects on Larvae

Aquatic amphibian stages exposed to pure urea show a strong tolerance. Schuytema and Nebeker ( 1 9 9 9 ~ )demonstrated that urea in its pure form affects tadpoles only at extremely high concentrations. Ghate (1985) found no teratogenic effects in embryos of the frog Microhyla ornata exposed to 1 000 mg/L urea (equivalent to 467 mg NIL) for four days. The lowest concentration that caused an effect on the growth of Hyla regilla and Xenopus laevis was 6 000 mg NIL and 2 400 mg NIL respectively (Schuytema and Nebeker 1999~).These authors reported a 10-day LC50 of 8 396 mg NIL for P regilla and 9 108 mg NIL for X. laevis. 2. Effects on Juveniles

Few studies have investigated the effects of environmental contaminants on terrestrial amphibians, but terrestrial toxic responses may have a considerable impact on amphibian conservation (Hatch et al. 2001). Newly metamorphosed amphibians could be especially sensitive to the pollution of their terrestrial habitats. Although the post-metamorphic

3164

AMPHIBIAN BIOLOGY

developmental stage is a very fragile one, ecotoxicological studies in post-metamorphic amphibians are very scarce. Hatch et al. (2001) found exposure to urea fertilizer on the soil substrate (100 Kg N/Ha) for five days to be lethal for two terrestrial metamorphic anurans. Mortality of metamorphic Bufo boreas was 17% while mortality of metamorphic R a m cascadae was 60%. Neither Ambystoma mucrodactylum nor Taricha granulosa, however, died as a result of urea exposure. 3. Effects on Adults

Marco et al. (2001) exposed adults of three terrestrial salamander species to urea at doses of 225 kg N/ha and 450 kg N/ha for four days. The observed effects increased with time and dose, and there were significant differences in sensitivity among the species. Both treatment levels had an acute effect on survival of Plethodon vehiculum and Rhyacotriton variegatus. For the urea doses of 225 Kg N/ha and at 48 hours, l? vehiculum and R. variegatus had a mortality rate of 47%. At 12 hours, the mortality rate at the highest doses of urea was 60% for I? vehiculum and 40% for R . variegatus. At 24 hours, mortality at the highest dose was 67% for I? vehiculum and 47% for R. variegatus. In contrast, there was no mortality for Taricha granulosa at these concentrations (Marco et al. 2001). VII. SUBLETHAL EFFECTS

A. Abnormalities Some abnormalities in amphibian embryos and larvae have been associated with an excess of nitrate or ammonium. The most common deformities are cardiac and abdominal oedemas, bent tails and dorsal tail curvature (lordosis). Few studies, however, have dealt with the relationship between nitrate or ammonium exposure and the quantitative analysis of abnormalities. Xenopus laevis showed deformities, mainly oedemas after a four-day exposure to ammonium nitrate (Schuytema and Nebeker 1999b). Discoglossus galganoi showed a high abnormality rate after two days of exposure to 200 mg NO
MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3165

Some embryos exposed to nitrogenous toxicants display a crescent-shaped body pattern (Laposata and Dunson 1998). Normal larvae move linearly through the water using serpentine tail movements to propel themselves rapidly in a given direction. Curled larvae move in a helical pattern, rather than in a linear one and at greatly reduced speeds. This deformity is hypothesized to be caused by the deactivation of a hatching enzyme responsible for enlarging the perivitelline membrane within which the embryo develops (Dunson and Connell 1982). With this enzyme inactive, the developing embryo curls as it grows larger within the restricted space. Curling sometimes proved fatal to embryos, but some curled embryos were able to hatch (Freda and Dunson 1985). The hatching enzyme is apparently most affected by cations (Laposata and Dunson 1998), so ammonium would be the main toxicant implicated in this deformity. B. Effects on Reproduction

Nearly all anuran species have external fertilization. Both ova and sperm released by adults come into contact with the aquatic environment even before fertilization. Poor water quality may, therefore, have an effect on the efficiency of fertilization by altering the properities of ova or sperm. In a recent study, ammonium nitrate was shown to cause a decrease in fertilization rates in Hyla arborea and Bufo calamita compared to controls (unpubl. data). Fecundity might also be indirectly affected by nitrogen contamination. Some life-history traits, such as fecundity, have been linked to larval fitness (Berven and Gill 1983; Smith 1987; Semlitsch et al. 1988; Berven 1990; Scott 1994; Morey and Reznick 2001) and the alteration of larval fitness may reduce their plastic responses of size and age at metamorphosis and alter reproductive success (Travis 1984; Alford and Harris 1988; Pfennig et al. 1991; Blouin 1992; Leips and Travis 1994; Morey and Reznick 2000; Doughty 2002).

C. Effects on Embryonic and Larval Development Amphibian embryos and especially larvae have a high ecological plasticity in their development that is mainly influenced by environmental conditions, such as temperature and dissolved oxygen, that are critical in this process (Podrabsky et al. 1998). Exposure to sublethal levels of reactive nitrogen in the aquatic habitat during this stage can alter the ecological response of tadpoles to their environment, leading to a failure in larval development, and thus decreasing reproductive success. Exposure to pollutants can cause physiological constraints by reducing the energetic balance because of diversion of resources into detoxification. This can result in smaller larvae with delayed growth rates and, ultimately, changes in time to metamorphosis and size and weight at metamorphosis. For example, Baker and Waights (1993, 1994) recorded significantly lower body lengths of Bufo bufo and Litoria caerulea tadpoles, respectively, when exposed for 16 days to 40 and 100 mg NO,-IL of sodium nitrate. Hecnar (1995) found a lower larval size in Bufo americanus, Pseudacris triseriuta and Rana pipiens exposed for four days at ammonium nitrate levels up to 50 mg NO,--NIL. However, under chronic exposures (100 days at 10 mg N-NO,-IL), three of these five species did not show an effect of the pollutant on larval size. Growth rate, measured in terms of increased mg of weight per day, in larval Rana temporaria exposed up to 5 mg N-NO,-/L decreased with increasing levels of sodium nitrate in one of two populations analyzed (Johansson et al. 2001). There is a strong variability, however, in developmental responses of tadpoles to nitrates and in other cases, reduction in growth rates among exposed individuals was very subtle or absent. A population of Rana temporaria studied by Johansson et al. (2001) showed no negative effects of nitrate on growth and Allran and Karasov (2000) found a very low quantitative effect, which was not significant, on growth rates in tadpoles of the leopard frog (Rana pipiens) exposed to sodium nitrate. The development of embryos or early larval stages has also been found to be adversely affected by nitrates. Pre-hatching growth of Xenopus laevis embryos was significantly decreased at nitrate concentrations as low as 56.7 mg/L (Schuytema and Nebeker 1999b). Embryos of Ram aurora exposed for 16 days to sodium nitrate at 29.1 mg N-NO,-/L and at 13.2 mg N-NH,+/L as ammonium nitrate were smaller than controls at the end of exposure (Schuytema and Nebeker 1999~).

3166

AMPHIBIAN BIOLOGY

Nitrate can be reduced to nitrite in tlie gut of tadpoles, and thus lead to decreased respiratory fitness. The animals respond by increasing their ventilatory rate which in turn elevates energetic costs. Also, alteration of the gut environment can lead to lower feeding rates that result in smaller tadpoles. Ireland (1991) presented evidence to suggest that anions such as nitrate or nitrite could adversely affect larval growth. Ammonium is also likely to affect larval development; un-ionized ammonia is highly toxic to cells and requires a strong detoxifying mechanism; this mechanism may be energetically expensive and divert resources that could otherwise be used in development. Growth of Xenopus laevis embryos was significantly reduced by total ammonia concentrations between 13.7 and 15.4 mg/L in three different test solutions after four days of exposure (Tietge et al. 2000). Jofre and Karasov (1999) exposed tadpoles of Rana clamitans to un-ionized ammonia. Larval size was lower at sublethal levels between 0.5 and 1.0 mg NHJL. After 114 days of exposure at the highest ammonia levels, the percentage of non-metamorphosed tadpoles not reaching Gosner stage 30 was lower than in controls. There are some examples of a positive effect of nitrogenous compounds on larval growth. Xu and Oldham (1997) found that Bufo bufo tadpoles exposed for 30 days to 100 mg NO,/L as ammonium nitrate were smaller than controls; tadpoles exposed for 15 days to 50 mg NO,-/L, however, produced significantly larger metamorphs than did controls, despite metamorphosing at an earlier date. A similar result was obtained by Ortiz et al. (2004), who observed a larger size of larval Pelobates cultripes exposed to 100 mg NO,-/L as ammonium nitrate than that showed by control individuals. Sullivan and Spence (2003) found a significant positive relationship in Xenopus laevis between weight at metamorphosis and the nitrate level to which tadpoles were exposed. Sublethal levels of nitrates can favour algal growth and thus influence food availability for tadpoles. Nevertheless, in the experiments performed by Xu and Oldham (199'7) and Ortiz et al. (2004) water was periodically renewed and algal growth was not allowed during laboratory experiments. The real causes underlying the observed results in these experiments remain unknown. Finally, under certain conditions a selective mortality can occur among smaller individuals. For example, Schmuck et al. (1994) observed a size-biased mortality that selectively eliminated the smallest size classes of reedfrog tadpoles (Hyperolius marmoratus) when reared under high densities that produced ammonia concentrations of up to 7.82 mg NH,+/L. D. Effects on Metamorphosis

Reactive nitrogen can affect metamorphosis in several ways. First, survival to metamorphosis can be reduced by deleterious effects of pollutants on later larval stages or during the metamorphic process itself. Effects on development can result in alterations of time to metamorphosis, and individuals' length and weight at metamorphosis. In addition, metamorphosis can be inhibited by nitrogenous contaminants under some circumstances. Finally, behavioural effects of chemicals on individuals pre-metamorphosis or postmetamorphosis have been recorded. Ammonia target levels of up to 1 mg NHJL reduced the metamorphic rate of larval Rana clamitans (Jofre and Karasov 1999). Moreover, higher ammonia levels reduced the snout-vet length at metamorphosis. On the other hand, ammonia exposure did not affect the time to metamorphosis (Jofre and Karasov 1999). Watt and Oldham (1995) showed that exposure to high concentrations of ammonium nitrate causes a reduction in size of Triturus vulgaris at metamorphosis. Watt and Jarvis (1997) exposed tadpoles of Triturus helveticus to 0-387.5 mg NO,-/L. Survival to metamorphosis was nil at the highest levels, while more than 90% of controls reached metamorphosis. Lower ammonium nitrate levels caused a delay of metamorphosis with respect to controls. Hecnar (1995) found similar effects of ammonium nitrate on the metamorphic success of Pseudacl-is triseriata, which was exposed to levels of up to 10 mg N-NO,-/L. Metamorphic rate was lower at the highest concentration, while the pollutant did not affect time to metamorphosis. A lower weight of metamorphs was detected among individuals exposed to lower ammonium

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3167

nitrate levels; higher concentrations, however, did not produce lighter metamorphs with respect to controls. This result could be due to a size-biased mortality caused by ammonium nitrate. In a comparative study of three Australian species, Hamer et al. (2004) observed how the only declining species, Litoria aurea, was sensitive to 10 mg NO,-/L as ammonium nitrate, in terms of survival to metamorphosis, to 10 mg NO,-IL as ammonium nitrate, whereas similar levels did not produce any effect on the metamorphosis of Crinia signifera or Limnodynastes peronii, both non-declining species. In another comparative study, sodium nitrate of up to 5 mg N-NOyIL produced a smaller size at metamorphosis and a longer time to metamorphosis in one of the two Rana temporaria populations assessed (Johansson et al. 2001). Sullivan and Spence (2003) exposed Xenopus laeuis tadpoles to 40 mg1L nitrate. The only effect observed was a delay in metamorphosis, correlated with heavier metamorphs as a result of the longer developmental period. Nitrate did not produce any effect on snoutvent length at metamorphosis, hematocrit, body condition or developmental rate. The absence of effects of sodium nitrate on metamorphosis was also reported by Allran and Karasov (2000), who exposed larval Rana pipiens to 0, 5 and 30 mg NO<-NIL. No differences were observed among treatments either in helmatocrit, survival to metamorphosis, or snoutvent length or weight at metamorphosis. Exposing tadpoles of Rana cascadae to 3.5 mg NNOp-IL as sodium nitrite produced no effects in terms of survival, time to metamorphosis or size at metamorphosis (Marco and Blaustein 1999). All results suggesting reduced rates of growth in the presence of nitrate are contradicted by data obtained by Xu and Oldham (1997), in which Bufo bufo tadpoles exposed for 15 days to 50 mg NO3-ILproduced metamorphs significantly larger than the controls despite metamorphosing at an earlier date. Freshly metamorphosed Hyperolius marmoratus froglets that were raised under adverse climatic conditions and at high ammonia levels during the larval period exhibited mortality rates, during a three-week period, of less than 15%, whereas froglets that developed under low ammonia levels, but otherwise similar conditions, suffered from a highly elevated mortality rate (40%) (Schmuck et al. 1994). This could indicate that ammonia exposure may be advantageous for relatively tolerant tadpoles, as it may pre-adapt them to adverse terrestrial conditions (Schmuck et al. 1994). Apart from these life history traits, effects of nitrogen pollution on metamorphosis are expressed in different ways. Inhibition or alteration of the metamorphic process itself can be caused by prolonged exposure to pollutants. For example, Marco and Blauste in (1999) observed a lower rate of tail reabsorption in individuals of Rana cascadae exposed to 3.5 mg NOyIL as sodium nitrite. Xu and Oldham (1997) found that 17% of Bufo bufo tadpoles exposed to 100 mg NO3-/Lfailed to reabsorb their tails.

E. Behavioural Effects Aquatic stages of amphibians are generally not able to avoid contamination of their habitats. Terrestrial stages, however, can select their habitats and thus avoid unsuitable sites. Few studies have dealt with this topic in amphibians and avoidance-ability remains unknown in most cases. Ortiz-Santaliestra et al. (2005) demonstrated that adult Lissotriton boscai were not able to detect and avoid substrates with potentially dangerous ammonium nitrate levels. Marco et al. (2001) exposed adult Plethodon uehiculum, Taricha granulosa and Rhyacotriton irariegatus in tanks in which half of each tank was sprayed with granulated urea and recorded which half of the tank the animals occupied. All three species displayed effective avoidance, spending more time in the part of the tank without urea. In another study, new metamorphs of Bufo boreas and Rana cascadae were exposed to urea on two different substrates. When paper towels were used as substrate juveniles could detect, and thus avoid, contaminated substrates (Hatch et al. 2001); when the same levels of urea were added to soil substrates, however, juvenile toads were only able to avoid contaminated soil at low levels of urea. The toxic effects of urea may have altered the activity of these toads. Mortality among exposed individuals of this species occurred within a five-day period, so animals could have been

3168

AMPHIBIAN BIOLOGY

stressed by the exposure to urea and consequently may not have been able to respond optimally (Hatch et al. 2001). Behaviour may be altered as a result of exposure to environmental stressors, including contaminants (e.g., Marco and Blaustein 1999), with the potential for altered biological interactions among species. Moreover, behavioural assays are ecologically relevant because altered behaviour due to stressors may result in an inability of animals to navigate toward breeding or hydration sites, and in an alteration of reproductive patterns that could lead to reduced reproductive potential or individual fitness (Blaustein et al. 2000). Effects of reactive nitrogen on activity and feeding behaviour, especially in larvae, have been assessed mostly in ecotoxicological studies. Activity level is usually measured as swimming performance or response to stimuli. Activity of tadpoles is a response to environmental conditions (such as changes in light and temperature). Exposure to ammonium nitrate seems to affect the central nervous system, which controls both activity and the sensitivity to environmental conditions (Hecnar 1995). The uncoordinated lateral twisting, disequilibrium or paralysis observed in most amphibians exposed to nitrogenous compounds (Hecnar 1995; Xu and Oldham 1997; Marco et al. 1999) also points to an effect on the central nervous system. Hatch and Blaustein (2000) exposed Rana cascadae larvae to three different levels of sodium nitrate. They measured activity by scoring animals as moving or not moving at intervals, and calculated the proportion of time spent active. The level of activity was significantly reduced at initial nitrate doses of 20 mg nitratell. By using ammonium nitrate, Xu and Oldham (1997) observed reduced activity levels in Bufo bufo tadpoles exposed to the pollutant for three days. After exposure, however, animals that were transferred into clean water recovered their normal activity level. Tadpoles chronically exposed to high levels of ammonium nitrate tended to be motionless and exhibit non-directed movement which, in the wild, would be expected to reduce their survival. Hecnar (1995) also observed behavioural effects in tadpoles of four amphibian species exposed to ammonium nitrate; exposed tadpoles swam less vigorously, showed disequilibrium, paralysis and delayed responses to stimuli. Similar effects were observed by Marco et al. (1999) in Ambystoma gracile and Rana pretiosa larvae exposed to up to 25 mg N-NOyIL as potassium nitrate. Both Hecnar (1995) and Marco et al. (1999) observed reduced feeding activity in tadpoles exposed to nitrates. Feeding behaviour may be affected by nitrates or nitrites that alter symbiotic gut bacteria. Reduced feeding activity results in lower tadpole size and, indirectly, in a lower probability of survival. Feeding rates were altered in Bufo bufo tadpoles exposed for three days to 100 mg N0,IL; affected individuals recovered quickly after the end of exposure (Xu and Oldham 1997). Non-lethal levels of nitrite increased feeding activitiy of smooth newt (Trzturus vulgaris) larvae and caused their prey (Daphnza) to spend more time near the top of the water column (Watt and Oldham 1995). Trophic relationships may be altered when feeding activity of predators or prey are reduced or increased by nitrogen exposure. This can affect individuals at the community level, and thus produce ecologically relevant effects. Hatch et al. (2001) exposed juveniles of Ambystom mcrodactylum, Taricha granulosa, Bufo boreas and Rana cascadae to urea at levels of 100 mg NIL for five days. After exposure, individuals were placed in clean tanks with crickets for prey. In the two anuran species, exposure to urea caused a significant decrease in the feeding rate, while feeding was not altered by urea in the two caudates. Several behavioural alterations have been observed in metamorphs. These can affect individuals in a different way than do the alteration of life-history traits reported above. For example, premetamorphic tadpoles exposed to nitrite emerge from the water at an early stage of development, conserving characteristics of the larval stage, such as relatively long tails (Marco and Blaustein 1999) that make them vulnerable to predation in the terrestrial environment. Moreover, exposed tadpoles spent a significantly longer time in

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3169

the shallow part of the tank with their heads out of the water than did controls (Marco and Blaustein 1999). Exposure to pollutants can alter locomotory performance, thus indirectly reducing the effectiveness of their defensive strategies. For example, escape from predators would be reduced in larvae of Ambystoma gracile, Bufo bufo, Rana pretiosa and Rana cascadae, whose swimming performance is altered by nitrate, ammonium and nitrite (Xu and Oldham 1997; Marco et al. 1999; Hatch and Blaustein 2000). Behavioural alteration by nitrite during metamorphosis, as described by Marco and Blaustein (1999) in Rana cascadae, probably would render metamorphs andlor larvae of this species more vulnerable to predation. There is rapid consumption by crayfish of larval Discoglossus galganoi exposed to increased levels of ammonium (unpubl. data). VIII. VARIABILITY AMONG SPECIES Several studies suggested a strong interspecific variability in sensitivity of amphibians to reactive nitrogen (Watt and Oldham 1995; Watt and Jarvis 1997; Smith et al. 2005). For example, under identical exposure conditions, larval Rana clamitans were up to three times more sensitive to sodium nitrate than were Rana catesbeiuna (Smith et al. 2005). In many cases, however, differences in other factors such as the age or size of test animals, the experimental conditions, water quality or the source of the nitrogenated ion may cause or enhance such variability. Authors should be cautious in suggesting the existence of interspecific or intraspecific variability. For example, larval survival of the palmate newt (Triturus helveticus) was highly dependent on the concentration of ammonium nitrate (Watt and Jarvis 1997). The high mortality suffered by these larvae on exposure to ammonium nitrate was in contrast to other work on Triturus vulgaris larvae in which mortality was very low (<5%) (Watt and Oldham 1995). The implication is that the larvae of these two species of newt differ in their sensitivity to ammonium nitrate exposure. However, the smooth newt larvae were tested at a later stage of development and a larger body size than were the palmate newt larvae; such differences could lead to false assessments of interspecific variability in vulnerability. Hatching success of Bufo americanus was not affected by un-ionized ammonia. By contrast, embryos of Rana pipiens and especially of Rana clamitans, showed significantly higher mortality rates in the presence of the pollutant than those observed in controls Uofre and Karasov 1999). Hecnar (1995) observed lethal effects of ammonium nitrate in concentrations up to 10 mg N-NOc/L in larvae of Pseudacris triseriata and Rana pipiens exposed for 100 days whereas in Rana clamitans, the most tolerant species, survival did not differ between exposed individuals and controls. Hecnar (1995) suggested that the resistance to agrochemicals by R . clamitans could be a factor in its success in the agricultural areas of southwestern Ontario. He also reported a threefold range in 96-hour LC50 for four different species subjected to acute exposures of ammonium nitrate. Marco et al. (1999) exposed hatchlings of five amphibian species to nitrate and nitrite under standard experimental conditions in water. They found significant variability in sensitivity among species. Ambystoma gracile displayed the highest acute effects from nitrate and nitrite. Three ranid species had acute effects in water from nitrite but the variability in LC50 and mortality at different nitrite doses was highly significant. In chronic exposures, Rana pretiosa was the species most sensitive to nitrates and nitrites. In nitrite concentrations of 0.88 mg N-NOy/L, Rana pretiosa was seven times more sensitive than Rana aurora and 20 times more sensitive than Hyla regilla. Marco et al. (2001) also found significant interspecific variability in sensitivity to urea using the same methodology and experimental conditions. Rough-skinned newts were very tolerant of high urea concentrations while red-backed salamanders and southern torrent salamanders were very sensitive. This interspecific variability is probably related to differences among species in the ability to withstand dry conditions and to the degree of development

3170

AMPHIBIAN BIOLOGY

of the lungs. Rough-skinned newts are much more terrestrial and consequently better adapted to dry conditions than are the other two species (Nussbaum et al. 1983). The red-backed salamander has no lungs and those of the southern torrent salamander are greatly reduced in size while the rough-skin newt has well-developed lungs (Duellman and Trueb 1994). The advantage of pulmonary exchange, with respect to water balance, is that lungs lose less water than does exposed wet skin. Salamanders that lack or have reduced lungs exchange respiratory gases almost exclusively across cutaneous surfaces. These salamanders are usually more sensitive to water losses in saline environments and have more permeable skins (to water and probably to other substances such as ammonia) than do salamanders that can respire through well developed lungs (Shoemaker et al. 1992). Species that mainly depend on cutaneous respiration and, consequently, have more permeable skins are probably more susceptible to the negative effects of urea and ammonia pollution. IX. VARIABILITY AMONG POPULATIONS

In Canada, Hecnar (1995) found inter-populational differences in the sensitivity of Bufo americanus tadpoles to ammonium nitrate fertilizer. Individuals from an area with heavy agricultural use were more tolerant of nitrate than were individuals from an area with lower agricultural use. Johansson et al. (2001) also found significant variation in sensitivity to nitrate between Rana temporaria tadpoles from two different Scandinavian populations. High concentrations of nitrate affected growth and size at metamorphosis of larvae from populations in northern Sweden while those from southern Sweden were unaffected by similar nitrate concentrations. These results, however, do not demonstrate the existence of genetic variability in tolerance to fertilizers among populations as methods differed between the two studies. In Hecnar's study, eggs were collected in the field after having been exposed to the natural levels of ammonium nitrate in their respective locations of origin. Johansson collected adult males and gravid females with developed ova. Fertilization took place in controlled clean water. However, the ova had developed in the field and one cannot exclude a possible maternal effect on sensitivity of tadpoles to the fertilizers. In these studies there was no conclusive demonstration of the evolution of amphibian tolerance to chemical fertilizers. Sullivan and Spence (2003) performed two series of experiments with a similar methodology to assess the effects of nitrates and atrazine on metamorphosis of Xenopus laevis. The results concerning the effects of nitrate were equivocal because in one of the series a significant positive relationship between weight at metamorphosis and the level of nitrate was found, while in the other series mean weight of tadpoles did not differ among nitrate treatments. The authors suggested that, because each experiment was conducted with a clutch of eggs from a distinct adult pair, some of the variation in the response may be attributable to genetic differences among clutches. The three clutches also displayed variation in mortality (1-14%) among experiments within the control groups, which would support the intraspecific-variability hypothesis. De Solla et al. (2002) found that hatching success in Rana aurora embryos varied among clutches from the same agricultural site exposed to field-water samples, suggesting either genetic or maternal effects on egg development. Other factors can affect intraspecific sensitivity to nitrates. For example, Hatch and Blaustein (2003) found that UV-B and nitrate together reduced the mass of larval Hyla regzlla in two populations at low and high elevations. At the high-elevation experiment, they found that UV-B and nitrate in combination reduced the survival of larval H. regilla. At both elevations, nitrate increased the mass of larval Ambystmu mucrodactylum. In the high-elevation experiment, however, this occurred only when UV-B was blocked. This result indicates that addition of nitrate could depend upon the presence of other factors, such as UV-B. Both genetic and maternal effects have been linked with hatching success and with growth rates of amphibian larvae and eggs exposed to acidic conditions (Pierce et al. 1987; Pierce and Wooten 1992).

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3171

X. IMPORTANCE OF AGE T O SENSITIVITY

Most ecotoxicological studies on anurans are conducted when larvae are close to their maximum body size (Gosner stage 25) (Devillers and Exbrayat 1992). This may not be the most sensitive stage, however, because gill circulation starts in larvae at stage 20 and opercular folds totally cover gills at the beginning of stage 25 (Gosner 1960). Hence, gills are directly exposed between stages 20 to 25 and absorption of pollutants may be more active during these stages. In general, comparison of different ecotoxicological studies carried out on amphibians indicates a trend toward higher sensitivity of younger individuals to some chemicals. For example, several studies have suggested that low pH is most toxic to the early developmental stages of amphibians (Freda 1986; Freda and McDonald 1993). Dial and Dial (1987) found that diquat and paraquat pesticides were more toxic to Rana pipiens embryos than to 15 day-old tadpoles. Nitrogen ions seem to affect the earlier developmental stages of amphibians to a greater extent than later ones. For example, Xu and Oldham (1997) found a greater tolerance of Bufo bufo larvae to ammonium nitrate than did Ortiz et al. (2004). The difference between these two studies may have depended upon the developmental stages used; whereas Xu and Oldham started their experiment with tadpoles at Gosner stage 32, Ortiz et al. began with embryos at Gosner stage 12. Life stage could also be the cause of an extremely high sensitivity showed by Bufo bufo larvae in Poland (Berger 1989). Unfortunately, Berger (1989) did not specify which stage she employed. In a comparison of two R a m temporaria populations, Johansson et al. (2001) found mortality from sodium nitrate (up to 5 mg N-N03/L) to be higher in a population with smaller individuals than in one in which the animals were larger. Larval Triturus helveticus suffered higher mortality on exposure to ammonium nitrate in a study by Watt and Jarvis (1997) than in one on Triturus vulgaris larvae conducted by Watt and Oldham (1995) in which mortality was very low. Differences in sensitivity to exposure to ammonium nitrate between these two species could be related partly to age. 7: vulgaris larvae were at a more advanced stage of development and were larger than were those of 7: helveticus. Watt and Jarvis also indicated that smaller 7: helveticus might have a lower survival rate than larger ones at all concentrations of ammonium nitrate so the difference in sensitivity within that species may be one of size. Schmuck et al. (1994) observed that the smallest size classes of Hyperolius murmoratus tadpoles were lacking when they were raised in environments with high ammonia levels, possibly because of size-biased mortality. Higher probability of survival as a fimction of age has also been reported in fish exposed to nitrite or ammonium. In these cases, however, bigger individuals tended to be more sensitive than were smaller ones (e.g., Palachek and Tomasso 1984a; Lewis and Morris 1986). As mentioned above, the probable mechanism of deleterious effects stemming from nitrates in amphibians is the development of methaemoglobinemia. In humans, this occurs mainly in infants younger than six months of age that are exposed to nitrate-contaminated water (Johnson et al. 1987). Young children have insufficient bacteria for efficiently reducing the available nitrite and are not capable of proper nitrate metabolism. Adults have a more diverse population of gut bacteria and are capable of effectively reducing the amount of available nitrite (Johnson et al. 1987). Higher tolerance of older individuals to nitrite has already been suggested; some animal species may become more tolerant to nitrite as they grow (United States EPA 1986). Marco and Blaustein (1999) suggested that tadpoles in later developmental stages might be more tolerant to nitrite than are those in earlier stages. Tadpoles have a greater functional respiratory surface area than do frogs of the same mass (Bradford 1983) and this difference may affect the uptake of NH3 in tadpoles compared to adults. Dejours et al. (1989a,b) demonstrated that ammonium toxicity was higher in smaller Xenopus laevis frogs and in the sharp-ribbed salamander (Pleurodeles waltl) than in larger ones; these authors suggested that higher body volume reduced the surface-volume ratio for absorption, producing a lower difision of toxicants through the skin.

3172

AMPHIBIAN BIOLOGY

In a recent study, the effects of ammonium nitrate on four amphibian species were evaluated at different stages of their embryonic and larval development (Ortiz-Santaliestra et al. 2006). Newly hatched individuals of Pelobates cultripes were even more sensitive than were embryos; sensitivity then decreased during subsequent larval development. By contrast, Bufo calamita showed a higher sensitivity during the later larval stages. Newly hatched larvae in anurans may be more sensitive than are embryos because of the lack of protection by a gelatinous egg capsule. Moreover, gill circulation starts at Gosner stage 20, just after hatching, when there is direct contact of the gill surfaces with the environment, and thus with possible toxicants diluted in the water. Covering of the gills and progressive thickening of the skin provide a stronger protection against diffusion of pollutants. In the case of Bufo calamita, the higher sensitivity could be the result of hypoxia. Respiraton of larval bufonids is almost exclusively through the skin because they are not able to breathe out of water as are other anuran groups. Hence, thicker skins can lower the rate of diffusion. If nitrates cause a decrease in respiratory efficiency because of methaemoglobinemia, older larvae may be more sensitive due to lack of alternative respiratory mechanisms. Age-related sensitivity has ecological implications. Small larvae suffer a greater risk of predation (Travis et al. 1985) and so poor growth rates of larvae presumably reduce their probability of survival. Maximum levels of nitrogen fertilizers in surface water are related to the timing of irrigation and of the application of fertilization. It has been demonstrated that leaching of nitrate is related to the amount of drainage (Pratt 1984) and that it peaks during the first rains after application of fertilizer (Ritter et al. 1991). Fertilizer is applied in temperate regions mainly during late winter, so the first spring rains cause an increase in the leaching of nitrate into bodies of water used by amphibians. Although this peak in nitrogen levels in water may not last long (Bogardi et al. 1991), it occurs during early spring when most amphibian species are breeding. Thus, although overall nitrate or ammonia levels in bodies of water are not usually high enough to cause high mortality in amphibians, peaks of toxic levels may negatively affect development of the early life stages of some species. XI. SYNERGISM WITH OTHER ECOLOGICAL AGENTS Results of ecotoxicological studies sometimes have scarce ecological relevance because of artificial conditions. For example, effects of pollutants observed in the laboratory may not be representative of events in the field since toxicity to aquatic organisms varies with water quality, temperature and the presence of other contaminants (Mason 1991). Many factors influence sensitivity of amphibians to pollutants in the field, and it is very difficult to assess the specific factor that is causing a particular effect. Multifactorial experiments where two or more factors act together is a better approach toward understanding the real ecological interaction between contaminants and amphibians in the field. A. Ultraviolet Radiation Hatch and Blaustein (2000) exposed larval Rana cascadae to different levels of UV, pH and sodium nitrate in an orthogonal multifactorial experimental design. Two conditons of UV were included: with UV (using acetate filters) and without UV (using Mylarmfilters). After three weeks of exposure, the authors observed that nitrate alone contributed significantly to reduced survival. Initial doses of 20 mg nitratell reduced activity both with and without UV-B. At initial doses of 5 mg nitratell, however, reduction in activity was only observed when UV-B was not blocked. Hatch and Blaustein (2003) found that UV-B and nitrate together reduced the mass of larval Hyh regilla in populations at both low and high elevations. In the high-elevevation experiment, they found that UV-B and nitrate together reduced the survival of larval H. regilla. In both the low-elevation and the high-elevation experiments, nitrate increased the mass of larval Ambystomu mucroductylum. In the high-elevation experiment, however, this result occurred only when UV-B was blocked. This result indicates that the effects of nitrate could

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3173

depend upon the presence of other factors such as UV-B. For a more detailed discussion of the effects of UV, see Chapter 6, this volume.

B. Acidification Variations in pH principally influence the equilibrium between ionized and un-ionized forms of ammonia. In this case, low pH favours the occurrence of the less-toxic form, ionized ammonium (e.g., Jofre and Karasov 1999). Water acidification rather than alkalinity is a widespread problem that can affect amphibians either independently (e.g., Freda and Dunson 1986; Simon et al. 2002) or in combination with other factors such as nitrogen pollutants. Hatch and Blaustein (2000) observed negative combined effects of pH and nitrate on Rana cascadae tadpoles. Nitrate produced significantly high mortality either alone or in combination with low pH (5). Negative effects of nitrates on activity level, however, were not detected when individuals were exposed to both high nitrate levels and low pH. In the same experiment, these two factors were combined with UV radiation; effects of the interaction between nitrates and UV-B have already been discussed in Chapter 6, this volume. Both survival and activity levels were significantly reduced in the treatment group in which all three factors were augmented in comparison to treatment groups involving only one factor. The three-way interaction was not significant either in the models of survival or for activity level. C. Temperature and Dissolved Oxygen

-

Water temperature influences toxicity of ammonium by determining the ratio between ionized and un-ionized forms. Diamond et al. (1993) exposed two amphibian species to un-ionized ammonia at different temperatures; they obtained a 96-hour LC50 of 1.44 mg NH$L for Rana pipiens embryos exposed to un-ionized ammonia at 20°C, while at lZ°C this value was 0.42 mg NHJL. In the case of Hyla crucqer, the 96-hour LC50s were 0.53 mg NH JL at 20°C and 0.46 mg NHJL at lZ°C. Interference in haemoglobin activity is one of the mechanisms of nitratelnitrite toxicity (Johnson et al. 198'1), so low levels of dissolved oxygen can increase the negative effects of these pollutants. Similarly, a reduction in oxygen tension increases the toxicity of un-ionized ammonia, as has been observed in some fish (Allan et al. 1990; Wajsbrot et al. 1991). Some adults and larvae of a number of amphibians overwinter in habitats that are particularly susceptible to oxygen depletion, like shallow lakes covered with ice, and hence they can be affected by ammonia under hypoxic conditions. Moreover, ammonia availability is higher under conditions of low oxygen concentrations in sediments (National Research Council 1979). Behavioural alterations may be caused by respiratory problems induced by reactive nitrogen. For example, "bobbing" behaviour has been described in many amphibians. Such behaviour makes the amphibian more vulnerable to predators (Moore and Townsend 1998). Apparently, the bobbing rate is related to oxygen availability in the water (Wassersug and Seibert 1975). Under hypoxic conditions, tadpoles may increase their frequency of breathing (West and Burggren 1982). Tissue hypoxia, due to exposure to nitrite, forced metamorphosing Ram cascadae tadpoles to shift to shallower water where they could breathe more efficiently (Marco and Blaustein 1999). Given that tadpoles are not suited to terrestrial life, the presence of re-locating near the shore could increase their risk of predation (Marco and Blaustein 1999). High temperatures may also affect the temporality of the bodies of water where amphibians develop. Amphibians in temporary ponds are subjected simultaneously to a variety of abiotic and biotic environmental changes that become more severe as the pond disappears. For example, the drying of ponds can lead to increased predation and competition as resources become limited and temperature and water quality undergo drastic fluctuations (Blaustein et al. 2001). Decreasing water volume as a consequence of drought in temporary habitats then results in a compilation of stressors that can interact with

3174

AMPHIBIAN BIOLOGY

nitrogen pollution. Moreover, Pounds and Crump (1994) speculated that, under hot and dry conditions, contaminants would accumulate to high levels in amphibian breeding ponds. Morey and Reznick (2004) analyzed larval development of three pelobatid species. They reported an increase in ammonium levels in the water as volume decreased. Slower growth during the larval stage, as caused by nitrogen pollution, can therefore result in failure to escape from a deteriorating aquatic environment (Savage 1961). D. Other Chemicals

Exposure of aquatic organisms to agricultural runoff implies the possibility of interaction of pesticides with fertilizers and nitrogenous wastes (Sullivan and Spence 2003). That amphibian numbers have significantly decreased in agricultural regions (Berger 1989) strongly suggests the impact of a variety of agricultural chemicals. While a number of studies have examined acute toxic effects of single chemicals on amphibians (e.g., Devillers and Exbrayat 1992), few have examined the possibility of deleterious interactions between environmentally realistic concentrations of agricultural products such as fertilizers and herbicides. Sullivan and Spence (2003) investigated the interaction of nitrate with atrazine, the most heavily used agricultural pesticide in North America, with more than 50 million kilograms applied to 25 million hectares annually in the United States (Eisler 1989). They exposed larval Xenopus laevis through metamorphosis to different levels of both nitrate and atrazine, both individually and combined. Nitrate alone caused only a delay in metamorphosis that was compensated by a greater metamorph mass. When combined with atrazine, nitrate reduced both length and weight at metamorphosis. Such a decrease in larval growth can have a significant detrimental impact later in the life of a frog by affecting survival and adult size (Berven 1990), rate of sexual maturation (Smith 1987), mate selection (Forester and Czarnowsky 1985) and locomotory ability, and hence ability to evade predators (Goater et al. 1993). Another investigation of the effects of mixtures of atrazine and nitrate on amphibian development found no statistically-significant interaction of these two chemicals in Ram pipiens (Allran and Karasov 2000). Differences in sensitivity among species could be the cause of these disparate results. Levels of both chemicals used by Sullivan and Spence were, however, higher than those tested by Allran and Karasov, thereby confounding comparison between the two studies. Some chemicals can present antagonistic effects when combined with nitrogen pollution. For example, after exposing Ambystoma texanum embryos to nitrite, Huey and Beitinger (1980b) reported a high mortality that was greatly reduced in the presence of chloride. A similar increase in nitrite tolerance in the presence of chlorides was also observed by Crawford and Allen (1977) in a demonstration of reduced nitrite lethality for Chinook salmon (Oncorhynchus tshawytscha) in seawater relative to freshwater. Results of these experiments support a model in which monovalent anions compete with nitrites for ionic uptake sites on respiratory surfaces. Decreased nitrite mortality of Ambystoma texanum in the presence of chlorides was suggested not to be an outcome of increased physiological tolerance but rather related to lower NO,-uptake rates (Huey and Beitinger 1980b). Other chemical properties can also influence the toxicity of reactive nitrogen to amphibians. Increasing salinity decreases the proportion of un-ionized ammonia, thus affecting total ammonium toxicity (Emerson et al. 1975; Whitfield et al. 1985; United States EPA 1989). Dissolved organic carbon (DOC) can bind to pollutants making them biologically inert although, at high concentrations, DOC can also be acutely toxic to amphibians (Horne and Dunson 1995).

E. Pathogens Emergent diseases have been detected in recent years in some amphibian populations around the world (Carey 2000). Occurrence of these diseases has been hypothesized to be related to a change in the hostlpathogen ratio or, more usually, with an immune depression caused by other stressors such as increased ultraviolet radiation (Kiesecker and Blaustein 1995).

MARC0 and ORTIZ-SANTALIESTRA. NITROGEN POLLUTION

3175

Dappen (1982) revealed that nitrate stress may depress immune response and blood haemoglobin in amphibian tadpoles. That author found decreased levels of circulating white cells and decreased haemoglobin values in bullfrogs and leopard frogs exposed to 9-26 mg/L of nitrate for three weeks. In a recent study, Romansic et al. (2006) observed how survival of R a m aurora was reduced in the presence of the water mold Saprolegnia in nitratefree water. When nitrate was added to the water, however, there were no significant differences in mortality between controls and animals treated with Saprolegnia. These authors concluded that nitrate prevented Saprolegnia from causing mortality in R . aurora. XII. ROLE OF NITROGEN POLLUTION IN AMPHIBIAN CONSERVATION Available information about the exposure and sensitivity of amphibians to an excess of reactive nitrogen suggests that this environmental problem may be contributing to the detected global amphibian decline. The risk of the negative impact of nitrogen pollution on amphibians can be especially relevant in agricultural, industrialized and heavily populated areas. Nitrate is by far the least noxious of the nitrogenous ions, but its combination with ammonium is more toxic for amphibians. Un-ionized ammonia and nitrite seem to be the most toxic forms of reactive nitrogen to amphibians. Fortunately, both chemicals are present at low concentration in the environment but under some circumstances peak concentrations of these two substances rise to toxic levels. The predicted global climatic change may increase the rate of evaporation from amphibian habitats, leading to a significant concentration of nitrogenous pollutants and an increase in the exposure of amphibians to these and many other contaminants. It is difficult to assign the decline of a given species directly to nitrogen pollution, as nitrogen often occurs in combinaton with many other contaminants. However, a high sensitivity of a declining species to nitrogenated ions, in addition to the disappearing of that species from areas with actual or past high levels of such ions, is highly indicative that nitrogen pollution is a contributor to the species' decline. For example, Rana pretiosa is very sensitive to nitrate and nitrite (Marco et al. 1999). This species is considered to be one of the most endangered amphibian species in North America. During the past four decades, it has disappeared from 90% of its original distributional range that included heavily fertilized agricultural areas (Nussbaum et al. 1983; Leonard et al. 1993). This species only remains in pristine forests and mountain areas while other sympatric amphibians that are more tolerant to nitrate are still relatively common in agricultural areas (Marco et al. 1999). Similarly, Hamer et al. (2004) found in Australia that the golden bell frog, Litoria aurea, is very sensititve to the presence of low levels of ammonium nitrate in the water; this species has severely declined during the decades following a pulse of fertilizers into freshwater habitats. Sympatric species, however, such as Crinia szgn$era and Lyrnnodynastes peronii, are very tolerant of ammonium nitrate and are still common in areas where L. aurea has disappeared (Hamer et al. 2004). An excess of ambiental reactive nitrogen could be seriously contributing to the decline of this very sensitive species.

Some species show a high mortality at the recommended limits of nitrite (5 mg N-NO,-/L) and nitrate (90 mg N-NO,-/L) for warm-water fish. A significant amphibian larval mortality and other deleterious effects have also been found at the recommended limits of nitrite (1 mg N-NO,-/L) and nitrate (10 mg N-NO,-/L) concentrations for drinking water. These results indicate that water quality criteria elaborated for other purposes do not guarantee the survival of some protected and endangered amphibians. Restrictive water quality criteria for nitrate, nitrite and ammonia in amphibian habitats are necessary to guarantee their survival. Larvae of sensitive species could be used as bioindicators of water quality. The sensitivity to reactive nitrogen has been studied in very few amphibian species. Further studies involving different species would be useful in understanding the role of an excess of ammonia in the water so that the information could be applied to the conservation of amphibians. Static renewal tests (Stephan 19'75) constitute a useful tool for assessing the acute impact of ammonia on amphibians. Freshwater mesocosms would be more appropriate for testing prolonged effects (Arnold et al. 1991).

3176

AMPHIBIAN BIOLOGY

Environmental levels of nitrogen pollution alone do not seem to cause a generalized decline of many amphibian species. Fortunately, the levels of reactive nitrogen that cause generalized damages to amphibians occur only occasionally in ponds or streams from developed countries where environmental and water quality protection is a priority. During previous decades, however, the levels of water pollution were much higher than at present in developed countries and this type of pollution, as well as others, may have contributed in the recent past to amphibian declines. There is very little information about the impact of fertilizers on amphibians in undeveloped countries. Those are now experiencing an intense development of their industry and agriculture, but without the desirable measures of environmental protection. Amphibians have high diversity in some of those countries and may be now suffering the impact of contaminants. Concentration of reactive nitrogen in surface freshwater can be quite variable, depending on many factors. Streams or rivers usually have low concentrations of nitrogenated ions, far below toxic levels for amphibians. Wetlands, ponds and lowland springs in agricultural or populated areas, however, can have nitrogen loads high enough to harm amphibians (Berger 1989; Hecnar 1995; Marco et al. 1999; Schuytema and Nebeker 1999a). Granular chemical fertilizers have had an acute impact on terrestrial amphibians (Oldham et al. 1997; Hatch et al. 2001; Marco et al. 2001). The environmental impact of ammonium nitrate fertilizers on adult terrestrial amphibians is probably very low. Granular ammonium nitrate is very soluble in water and its persistence at the soil surface is ephemeral. Moreover, application of fertilizer usually occurs during daylight, while amphibian migration occurs at night. Further studies are necessary to evaluate the impact of ammonium nitrate fertilizer on juvenile amphibians. Application of fertilizer may have an acute impact on amphibian survival in those habitats such as forests or grasslands where amphibians remain for long periods of time, including daytime. Other chemical fertilizers such as urea, muriate of potash or inorganic growmore, widely used on herbaceous crops and grasslands and in forests, dissolve more slowly and remain for longer and at higher concentrations on the soil surface. Thus, amphibians may be exposed to such fertilizers on the substrate for longer periods of time after application (Marco et al. 2001). Moreover, in forest ecosystems, many forest-dwelling, terrestrial amphibians would be exposed to fertilizers or other chemicals not only when chemicals are on the substrate, but even when they are dissolved in the forest soil. Amphibians of various families (e.g., Plethodontidae, Rhyacotritonidae, Ascaphidae, Dicamptodontidae and Salamandridae) live under surface debris, gravel or stones, in decaying logs and in burrows in soil or crevices in the forest canopy. In these microhabitats, time of exposure to chemicals would also be longer. Further studies are necessary for understanding the impact of fertilizers on amphibians in these habitats. Monitoring of amphibians in the field should be carried out in forests and prairies to which fertilizer has been applied. Terrestrial amphibians adapted to saline or arid environments are probably not very sensitive to fertilizer pollution. Those terrestrial species that have been found to be sensitive to an excess of environmental urea mostly inhabit humid temperate forests or non-arid mountains. Sublethal levels of reactive nitrogen may influence interspecific competition. More tolerant species may have an advantage over competitors in a polluted environment. For example, bullfrogs are very aggressive and very efficiently compete for food and space with other anurans. This ecological interaction may be contributing to the decline of native amphibians in areas where the bullfrog is not autochthonous (Kiesecker and Blaustein 1997). The relatively high tolerance of bullfrogs to nitrite (Huey and Beitinger 1980a; Smith et al. 2004) may favour this exotic species over the native species in habitats polluted by nitrate or nitrite. In the Pacific Northwest of the United States the endangered Rana pretiosa is very sensitive to nitrate and nitrite and has disappeared during recent decades from many agricultural areas where Rana catesbeiuna has been introduced. The interaction between water pollution and the presence of the exotic, resistant bullfrogs may have contributed to the extintion of native species.

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3177

Special attention should be paid to the common exposure of amphibians in the field to mixtures of chemicals. Different nitrogenated ions may interact with other contaminants (e.g., pesticides, PCBs, PAHs, dioxins), potentiating their negative effects on amphibians. Little research has focussed on this problem but preliminary studies indicate that synergism between reactive nitrogen and other contaminants may decisively contribute to amphibian decline. Even combinations of low levels of these chemicals (that alone are not toxic) may be affecting amphibians. Increased attention should be directed toward the study of the combined effects of toxicants on amphibians. It is essential to develop recovery plans for different ecosystems, amphibian assemblages, or species if the conservation of amphibian diversity is to be guaranteed (Semlitsch 2002). These plans should recommend the control of levels of nitrate, nitrite and ammonium, and institute measures to reduce their concentrations when they are close to toxic levels for the most sensitive species. There are several ways to lower the use of compounds that include reactive nitrogen, or to reduce their impact. In comparison with conventional, high-intensity agricultural methods, "organic7' alternatives can improve soil fertility and exert fewer detrimental effects on the environment. These alternatives can also produce equivalent crop yields to those of conventional methods (Tilman 1998). Use of chemical fertilizers can be made more efficient and their use and mobilitiy can be reduced. For example, they can be applied only to those parts of fields where they are needed and only in the doses required. Computerized maps of fields, calibrated to specific soil and water conditions can assist in this process. There are also other methods, such as introducing tertiary treatments or nutrient removal to sewage treatment plants; proper handling of manure and animal waste lagoons can minimize the discharge of animal waste or waste runoff to streams. Large herbivores that reduce vegetation and increase urea and ammonium concentration in the water can be excluded by fences around important amphibian habitats. Establishing riparian zones that can be used to regulate the transport of nitrate in groundwater flow from uplands to streams may be important. The current consensus is that most riparian zones effectively remove nitrate from subsurface water (Hill 1996). Semlitsch and Bodie (2003) noted the importance of defining core habitats used by local breeding populations of amphibians in wetlands. If these terrestrial core areas are vegetated, they may also significantly reduce the reactive nitrogen loads that are entering amphibian aquatic habitats from surrounding areas (Haycock and Burt 1991; Muscutt et al. 1993; Osborne and Kovacic 1993). Alternatively, the installation of tiled drains that divert agricultural runoff to wetlands constructed parallel to streams or ponds can be very effective in controlling non-point-source agricultural inputs (Osborne and Kovacic 1993). XIII. REFERENCES Aber, J. D.,1992.Nitrogen cycling and nitrogen saturation in temperate forest ecosystems. Trenh Ecol. Evol. 7: 220-224.

J. S' and R'* lgg2. "Water Quality for Freshwater Fish, Second edition. Butterworths, London. Alford, R. A. and Harris, R. N., 1988. Effects of larval growth history on anuran metamorphosis. Am. Nat. 131: 91-106. M a n , G. L., Maguire, G. B. and Hopkins, S. J., 1990. Acute and chornic toxicity of ammonia to juvenile Metapenaew macleayi and Penaew monodon and the influence of low dissolved-oxygen levels. Aquaculture 92: 277-288. Allran, J. W., Karasov, W. H., 2000. Effects of atrazine and nitrate on northern leopard frog (Rampipiens) larvae exposed in the laboratory from posthatch through metamorphosis. Envinm. Toxicol. Chem. 19: 2850-2855.

Alonso, A,, Camargo, J. A., 2003. Short-term toxicity of ammonia, nitrite and nitrate to the aquatic snail Potamopyrgus antipodarum (Hydrobiidae, Molluscs). Bull. Environ. Contam. Toxicol. 70: 10061012. Alvarado, R, H, and Kirschner, L, B,, 1963,Osmotic and ionic regulation in Ambystoma tigrinum. Compar. Biochem. Physiol. 10: 55-67. Arillo, A., Margiocco, C., Melodia, F., Mensi, P: and Schenone, G., 1981.Ammonia toxicity mechanism in fish: studies on rainbow trout (Salmo gairdnedi R.). Ecotoxicol. Environ. Saf: 5: 3 16-328. Arnold, D., Hill, I., Matthiessen, P and Stephenson, R., 1991. Guidance document on testing procedures for pesticides in freshwater mesocosms. SETAC-Europe. Brussels. ASTM, 1997. Standard guide for conducting the frog embryo teratogenesis assay -Xenopus (Fetax). E143991,Annual Book of ASTM Standards, Vol. 11.05.E47, Committee on Biological Effects and Envimnmental Fate, West Conshohocken, Philadelphia.

3178

AMPHIBIAN BIOLOGY

Babiker, I. S., Mohamed, M. A. A., Terao, H., Kato, K. and Ohta, K., 2004. Assessment of groundwater contamination by nitrate leaching from intensive vegetable cultivation using geographical information system. Environ. Internat. 29: 1009-10 17. Baker, J. and Waights, V., 1993. The effect of sodium nitrate on the growth and survival of toad tadpoles (Bufo bufo) in the laboratory. Herpetol. J. 3: 147-148. Baker, J. and Waights, V., 1994. The effects of nitrate on tadpoles of the tree frog (Litoria caerulea). Herpetol. J. 4: 106-108. Balasubramanian, P, Saravanan, T S., Palaniappan, M. K., 1999. Biochemical and HistopathologicalChanges in Certain Tissues of Oreochromis mossambicus vrewaves) Under Ambient Urea Stress. Bull. Environ. Contam. Toxicol. 63: 117-124. Balinsky, J. B., 1970. Nitrogen metabolism in amphibians. Pp. 519-637 in "Comparative Biochemistry of Nitrogen Metabolism. Vol. 2. The Vertebrates", ed by J. W. Campbell. Academic Press. Balinsky, J. B., 1981. Adaptation of nitrogen metabolism to hyperosmotic environment in Amphibia. J. Exp. 2001. 215: 335-350. Bath, R. N. and Eddy, F. B., 1980. Transport of nitrite across fish gills. J. Exp. 2001. 214: 119-121. Bengtson, G. W., 1979. Forest fertilization in the United States: progress and outlook. J. For: 78: 222-228. Berger, L., 1989. Disappearance of amphibian larvae in the agricultural landscape. Ecol. Int. Bull. 17: 65-73. Berry, J. T, 1994. "Commercial Fertilizers". Tennessee Valley Authority Environmental Research Center Muscle Shoals. Bewen, K., 1990. Factors affecting population fluctuations in larval and adult stages of the wood frog (Rana sylvatica). Ecology 71: 1599-1608. Berven, K. A. and Gill, D. E., 1983. Interpreting geographic variation in life-history traits. Am. Zool. 23: 85-97. Bishop, C. A, Mahony, N. A, Struger, J. N. G. P and Pettit, K. E., 1999. Anuran development, density and diversity in relation to agricultural activity in the Holland River watershed, Ontario, Canada (19901992). Environ. Monit. Assess. 57: 21-43. Blaustein, A. R., Chivers, D. P, Kats, L. B. and Kiesecker, J. M., 2000. Effects of ultraviolet radiation on locomotion and orientation in roughskin newts (Taricha granulosa). Ethology 106: 227-234. Blaustein, A. R., Wildy, E. L., Belden, L. K. and Hatch, A., 2001. Influence of abiotic and biotic factors on amphibians in ephemeral ponds with special reference to long-toed salamanders (Ambystoma macrodactylum). Israel J. Zool. 47: 333-345. Blouin, M. S., 1992. Comparing bi-variate reaction norms among species - time and size at metamorphosis in 3 species of Hyla (Anura, Hylidae). Oecologia 90: 288-293. Board, P G., Agar, N. S., Gmca, M. and Shine, R., 1977. Methemoglobin and its reduction in nucleated erythrocytes from reptiles and birds. Compar: Biochem. Physiol. B-Biochem. Molec. Biol. 57: 265-267. Bodansky, O., 1951. Methemoglobinemia and methemoglobin producing compounds. Pharmacol. Rev. 3: 144-196.

Bogardi, I., Kuzelka, R. D. and Ennenga, W. G., 1991. "Nitrate Contamination: Exposure, Consequence, and Control". NATO AS1 series G - Ecological Sciences, Vol. 30. Springer-Verlag, New York. Bonn, B. A., Wentz, D. A. and Hinkle, S. R., 1996. "Willamette Basin, Oregon - Nitrogen in Streams and Ground Water 1980-1990". Open File Report 96-227. United States Geological Survey, Portland. Boutilier, R. G., Stiffler, D. F. and Toews, D. P, 1992. Exchange of respiratory gases, ions, and water in amphibious and aquatic amphibians. Pp. 81-124 in "Environmental Physiology of Amphibians", ed by M. E. Feder and W. W. Burggren. The University of Chicago Press, Chicago. Bower, C. E. and Bidwell, J. P, 1978. Ionization of ammonia in seawater: effects of temperature, pH, and salinity.J. Fish. Res. Board Can. 35: 1012-10 16. Boyer, E. W., Goodale, C. L., Jaworski, N. A and Howarth, R. W., 2002. Anthropogenic nitrogen sources and relationships to riverine nitrogen export in the Northeastern USA. Biogeochemistry 57/58: 137-69. Boyer, R., 1993. Evaluation of water quality in relation to frogs at Klamath Basin National Wildlife Refuges. (MS thesis). University of Washington. Seattle. Boyer, R. and Grue, C. E., 1995. The need for water quality criteria for frogs. Environ. Health Perspec. 103: 352-357. Bradford, D. F., 1983. Winterkill, oxygen relations, and energy metabolism of a submerged dormant amphibian, Rana muscosa. Ecology 64: 1171-1 183. Brodman, R., Ogger, J., Bogard, T., Long, A. J., Pulver, R. A., Mancuso, K. and Falk, D., 2003. Multivariate analyses of the influences of water chemistry and habitat parameters on the abundances of pondbreeding amphibians. J. Freshw. Ecol. 18: 425436. Brown, D. A. and McLeay, D. J., 1975. Effect of nitrite on methemoglobin and total hemoglobin of juvenile rainbow trout. Progress. Fish-Culturist 37: 36-38. Bmning-Fann, C. S. and Kaneene, J. B., 1993. The effects of nitrate, nitrite, and N-nitroso compounds on human health: a review. E t . Human Toxicol. 35: 521-538. Burns, D. A., 2004. The effects of atmospheric nitrogen deposition in the Rocky Mountains of Colorado and southern Wyoming, USA - a critical review. Environ. Pollut. 127: 257-269. Bytnerowicz, A. and Fenn, M. E., 1996. Nitrogen deposition in California forests: A review. Environ. Pollut. 92: 127-146. Camargo, J. A. and Ward, J. W., 1992. Short-term toxicity of sodium nitrate (NaNO,) to non-target freshwater invertebrates. Chemosphere 24: 23-28. Cameron, J. N., 1971. Methemoglobin in erythrocytes of rainbow trout. Comp. Biochem. Phyiol. 40: 743-749. Cardoso, E. L., Chiarini-Garcia, H., Ferreira, R. M. A. and Poli, C. R., 1996. Morphological changes in the gills of Lophiosilurus alexandri exposed to un-ionized ammonia. J. Fish. Biol. 49: 778-787. Carey, C., 2000. Infectious disease and worldwide declines of amphibian populations, with comments on emerging diseases in coral reef organisms and in humans. Envircm. Health Perspec. 108 (Suppl.1): 143-150.

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3179

Chalmers, A,, Kershaw, C. and Leech, E, 1990. Fertilizer use on farm crops in Great-Britain - results from the survey of fertilizer practice, 1969-88. Outlook Agric. 19: 269-278.

Downing, K. M. and Merkens, J. C., 1955. The influence of dissolved oxygen concentration on the toxicity of un-ionized ammonia to rainbow trout (Salmo gardnierii Richardson). Ann. Appl. Biol. 43: 243-246.

Colt, J. and Tchobanoglous, G., 1976. Evaluation of the short-term toxicity of nitronenous compounds to channel catfish, ~ctaluruspu%ctatus. ~ ~ u a c u l t u r8:e 209-22 1.

Driscoll, C. T, 2003. Elevated inputs of reactive nitrogen in the environment causes, consequences, and controls. Biosczence 53: 3 11-3 11.

Colt, J. and Tchobanoglous, G., 1978. Chronic exposure of channel catfish, Ictalurus punctatus, to ammonia: Effects on growth and survival. Aquaculture 15: 353-372. Committee on Nitrate Accumulation, 1972. "Accumulation of nitrate". National Academy of Sciences, Washington, DC. Cotta Ramusino, M., 1980. Water pollution and embryonic development. Boll. Zool. 47: 263-265. Crawford, R. E. and Allen, G. H., 1977. Seawater inhibition of nitrite toxicity to Chinook salmon. Trans. Amer Fish. Sac. 106: 105-109. Crawshaw, G. J., 2000. Diseases and pathology of amphibians and reptiles. Pp. 199-232 in "Ecotoxicology of Amphibians and Reptiles", ed by D. W. Sparling, G. Linder and C. A. Bishop. Society of Environmental Toxicology & Chemistry (SETAC), Pensacola.

Duellman, W. E. and Trueb, L., 1994. "Biology of amphibians". The John Hopluns University Press, Baltimore. Dunson, W. A. and Connell, J., 1982. Specific inhibition of hatching in amphibians exposed to low pH. J. Herpetol. 16: 314-316. Earl, S. R. and Blinn, D. W., 2003. Effects of wildfire ash on water chemistry and biota in South-Western USA streams. Freshw. Biol. 48: 1015-1030. Eddy, F. B. and Williams, E. M., 1994. Freshwater fish and nitrite. Pp. 117-143 in "Water Quality for Freshwater Fish", ed by G. Howells. Gordon and Breach Science Publishers, Yverdon. Edwards, D. R. and Daniel, T C., 1994. Quality of runoff from fescue grass plots treated with poultry litter and inorganic fertilizer.J. Enuiron. @l. 25: 579-584.

Crews, T E. and Peoples, M. B., 2004. Legume versus fertilizer sources of nitrogen: ecological tradeoff and human needs. Agricul. Ecosys. Enuiron. 102: 279-297.

Edwards, T M., McCoy, K. A, Barbeau, T, McCoy, M. W., Thro, J. M. and Guillete, L. J. Jr., 2006. Environmental context determines nitrate toxicity in Southern toad (Bufo terrestris) tadpoles. Aquat. Toxicol. 78: 50-58.

Cmtzen, I!J. and Ehhalt, D. D., 1977. Effects of nitrogen fertilizers and combustion on the stratospheric ozone layer. Ambio. 6: 112-1 17.

EIFAC, 1984. "Water Quality Criteria and Criteria for European Freshwater Fish. European Inland Advisoly Commission Tech. Pap. 46. 19 p. FA0 Rome.

Dappen, G. E., 1982. Effects of nitrates upon hemopoietic, lymphoid and vascular tissues of tadpoles and frogs. Proc. Nebraska Acad. Sci. and AfJil. Soc. 92: 23.

Eisler, R., 1989. "Atrazine Hazards to Fish, Wildlife, and Invertebrates: A Synoptic Review". Biological Report 8511.18. U.S. Fish and Wildlife Service, Washington.

Dejours, E, Armand, J. and Beekenkamp, H., 1989a. La toxicit6 de I'ammoniac est fonction du pH de I'eau. Etude chez la salamandre Pleurodeles waltl. C. R . Acad. Sci. Paris 308: 55-60.

Emerson, K., Russo, R. C., Lund, R. E. and Thurston, R. V., 1975. Aqueous ammonia equilibrium calculations: effect of pH and temperature. J. Firh. Res. Board. Can. 32: 2379-2383.

Dejours, E, Armand, J. and Beekenkamp, H., 1989b. Action de la tempkrature et de la taille sur la toxicit6 de I'ammoniac chez l'amphibien anoure Xenopus laeuis. C. R . Acad. Sci. Paris 309: 363-368.

Ensabella, F., Loriga, S., Formichetti, l?, Isotti, R. and Sorace, A., 2003. Breeding site selection of Bufo uiridis in the city of Rome (Italy). Amphibia-Reptilia 24: 396-400.

de Solla, S. R., Pettit, K. E., Bishop, C. A., Cheng, K. M. and Elliott, J. E., 2002. Effects of agricultural runoff on native amphibians in the Lower Fraser River Valley, British Columbia, Canada. Environ. Toxicol. Chem. 21: 353-360.

European Council, 1998. Quality of water intended for human consume. European Council Directive 981831 CE. Official Journal no L 330, 0511211998, Pp. 0032-0054.

Devillers, J. and Exbrayat, J. M., 1992. "Ecotoxicity of chemicals to amphibians". Gordan and Breach Science Publishers, Lyon. Dial, N. A. and Dial, C. A. B., 1987. Lethal effects of diquat and paraquat on developing frog embryos and 15-day-old tadpoles, Rana pipiens. Bull. Enuiron. Contam. Toxicol. 38: 1006-10 11. Diamond, J. M., Mackler, D. G., Rasnake, W. J. and Gmber, D., 1993. Derivation of site-specdic ammonia criteria for an effluent-dominated headwater stream. Enuiron. Toxicol. Chem. 12: 649-658.

FAO, 2002. Fertilizer Yearbook 2000 - Annuaire des engrais 2000 - Anuario de fertilizantes 2000. Food and Agriculture Organization (UN). FA0 Statis. Series 50: 1-272. Federal Research Center for Forestry and Forest Products (BFH), 2003. "The Condition of Forests in Europe. 2003 Executive Report". European Commission and United Nations Economic Commission for Europe, Geneva.

Dise, N. B. and Wright, R. F., 1995. Nitrogen leaching from European forests in relation to nitrogen deposition. Forest Ecol. Man. 71: 153-161.

Fenn, M. E., Poth, M. A., Aber, J. D., Baron, J. S., Bormann, B. T, Johnson, D. W., Lemly, A. D., McNulty, S. G., Ryan, D. F. and Stottlemyer, R., 1998. Nitrogen excess in North American ecosystems: predisposing factors, ecosystem responses, and management strategies. Ecol. Applic. 8: 706-733.

Doughty, E, 2002. Coevolution of developmental plasticity and large egg size in Crinia georgiana tadpoles. Copeia 2002: 928-937.

Ferm, M., 1998. Atmospheric ammonia and ammonium transport in Europe and critical loads: a review. Nutr Cycle Agroecosyst. 51: 5-17.

3180

AMPHIBIAN BIOLOGY

FMA, MAFF and SOAFD [Fertiliser Manufacturer's Association, Ministry of Agriculture; Fisheries and Food; Scottish Office of Agriculture and Fisheries Department], 1993. "The British Survey of Fertiliser Practice: Fertiliser use on Farm Crops 1992". HMSO, London. Forester, D. C. and Czarnowsky, R., 1985. Sexual selection in the spring peeper, Hyla crucfer (Amphibia, Anura): Role of the advertisement call. Behaviour 92: 112-128. punctuation? Freda, J., 1986. The influence of acidic pond water on amphibians: a review. Water Air Soil Pollut. 30: 439-450. Freda, J. and Dunson, W. A., 1985. The influence of external cation concentration on the hatching of amphibian embryos in water of low pH. Can. J. 2001. 63: 2649-2656. Freda, J. and Dunson, W. A,, 1986. Effects of low pH and other chemical variables on the local distribution of amphibians. Copeia 1986: 454-466. Freda, J. and McDonald, D. G., 1993. Toxicity of amphibian breeding ponds in the Sudbury region. Can. J. Fish. Aquat. Sci. 50: 1497-1503. Galloway, J. N., Aber, J. D., Erisman, J. W., Seitzinger, S. E, Howarth, R. W., Cowling, E. B. and Cosby, B. J., 2003. The nitrogen cascade. Bioscience 53: 341-356. Gessel, S. E and Atkinson, W. A., 1984. Use of fertilizers in sustained productivity of Douglas-fir forests. Pp. 67-88 in "Forest Soils and Treatment Impacts", ed by E. L. Stone. The University of Tennessee, Knoxville. Ghate, H. V., 1985. Dithiocarbamate induced teratogenesis in frog embryo. Rib. Biol. 78: 288-291. Goater, C. E, Semlitsch, R. D. and Bernasconi, M. V., 1993. Effects of body size and parasite infection on the locomotory performance of juvenile toads, Bufo bufo. Oikos 66: 129-136. Gosner, K. L., 1960. A simplified table for staging anuran embryos and larvae with notes on identification. Herpetologica 16: 183-190. Griffis-Kyle, K. L., 2007. Sublethal effects of nitrite on eastern tiger salamander (Ambtoma t i p n u m tigrinum) and wood frog (Rana sylvatica) embryos and larvae: implications for field populations. Aquat. Ecol. 41: 119-127. Gruca, M. and Grigg, G. C., 1980. Methemoglobin reduction in crocodile blood - are high-levels of methb typical of healthy reptiles. J. Experi. 2001. 213: 305-308. Hack-Ten-Broeke, M. J. D., De-Groot, W. J. M. and Dijkstra, J. E, 1996. Impact of excreted nitrogen by grazing cattle on nitrate leaching. Soil Use Man. 12: 190-198. Hamer, A. J., Makings, J. A., Lane, S. J. and Mahony, M. J., 2004. Amphibian decline and fertilizers used on agricultural land in south-eastern Australia. Agricul. Ecosys. Environ. 102: 299-305. Hampson, B. L., 1976. Ammonia concentration in relation to ammonia toxicity during a rainbow trout rearing experiment in a closed freshwater-seawater system. Aquaculture 9: 6 1-70. Hariston, N. G., 1987. "Community Ecology and Salamander Guilds". Cambridge Univ. Press, Cambridge.

Hatch, A. C. and Blaustein, A R., 2000. Combined effects of UV-B, nitrate, and low pH reduce the survival and activity level of larval Cascades frogs (Rana cascadae). Arch. Environ. Contamin. Toxicol. 39: 494-499. Hatch, A. C. and Blaustein, A. R., 2003. Combined effects of W - B radiation and nitrate fertilizer on larval amphibians. Ecol. Appl. 13: 1083-1093. Hatch, A. C., Belden, L. K., Scheessele, E. and Blaustein, A. R., 2001. Juvenile amphibians do not avoid potentially lethal levels of urea on soil substrate. Environ. Toxicol. Chem. 20: 2328-2335. Hawkins, R. A., Miller, A. L., Nielsen, R. C. and Veech, R. L., 1973. The acute actions of ammonia on rat brain metabolism in vivo. Bwchem. J. 134: 1001-1008. Haycock, N. E. and Burt, T. E, 1991. The sensitivity of rivers to nitrate leaching: the effectiveness of nearstream land as a nutrient retention zone. Pp. 3 12-319 in "Landscape Sensitivity", ed by R. J. Allison and D. S. Thomas. Willey, London. Hecnar, S. J., 1995. Acute and chronic toxicity of ammonium nitrate fertilizer to amphibians from southern Ontario. Environ. Toxicol. Chem. 14: 2 131-2 137. Hecnar, S. J. and Mcloskey, R. T., 1996. Amphibian species richness and distribution in relation to pond water chemistry in South-western Ontario, Canada. Freshlo. Biol. 36: 7-1 5. Hickey, C. W. and Vickers, M. L., 1994. Toxicity of ammonia to nine native New Zealand freshwater invertebrate species. Arch. Environ. Contam. Toxicol. 26: 292-298. Hill, A. R., 1996. Nitrate removal in stream riparian zones. J. Environ. Qual. 25: 743-755. Holcombe, G. W., Phipps, G. L., Sulaiman, A. H. and Hoffman, A. D., 1987. Simultaneous multiple species testing - acute toxicity of 13 chemicals to 12 diverse ksh-water amphibian, fish, and invertebrate famihes. Arch. Environ. Contam. Toxicol. 16: 697-710. Horne, M. T and Dunson, W. A., 1995. The interactive effects of low pH, toxic metals, and DOC on a simulated temporary pond community. Environ. Pollut. 89: 155-161. Hudak, E F., 2000. Regional trends in nitrate content of Texas groundwater. J. Hydrol. 228: 37-47. Huertas, M., Gisbert, E., Rodriguez, A., Cardona, L., Williot, E and Castello-Orvay, F., 2002. Acute exposure of Siberian sturgeon (Acipemer baeri, Brandt) yearlings to nitrite: median-lethal concentration (LC50) determination, haematological changes and nitrite accumulation in selected tissues. Aquat. Toxicol. 57: 257-266. Huey, D. W. and Beitinger, T L., 1980a. Hematological responses of larval Rana catesbeiana to sublethal n i t h e exposures. Bull. Environ. Contam. Toxicol. 25: 574-577. Huey, D. W. and Beitinger, T. L., 1980b. Toxicity of nitrite to larvae of the salamander Ambystoma texanum. Bull. Environ. Contam. Toxicol. 25: 909-91 2. Huey, D. W. and Beitinger, T L., 1981. A methemoglobin reductase system in channel catfish (Zctalurus punctutus). Can. J. 2001. 60: 1511-15 13. Huey, D. W., Beitinger, T L. and Wooten, M. C., 1984. Nitrite-induced methemoglobin formation and recovery in channel catfish (Zctalurus punctatus) at three acclimation temperatures. Bull. Environ. Contam. Toxicol. 32: 674-68 1.

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION Ireland, P H., 1991. Separate effects of acid-derived anions and cations on growth of larval salamanders of Ambystoma maculatum. Copeia 1991: 132-137. Janssens, P: A., 1972. The influence of ammonia on the transition to ureotelism in Xenopus laeuis. J. Exp. Zool. 182: 357-366. Jarvis, S., C., 1996. Future trends in nitrogen research. Plant Soil 181: 47-56. Jofre, M. B. and Karasov, W. H., 1999. Direct effect of ammonia on three species of north american anuran amphibians. Enuiron. Toxicol. Chem. 18: 1806-1 812. Johansson, M., Rasanen, K. and Merila, J., 2001. Comparison of nitrate tolerance between different populations of the common frog, Rana temporaria. Apuat. Toxicol. 54: 1-14. Johnson, C. J., Bonrud, P A., Dosch, T. L., klness, A. W., Senger, K. A,, Busch, D. C. and Meyer, M. R., 1987. Fatal outcome of methemoglobinemia in an infant. J. Am. Med. Asso. 257: 2796-2797. Jirgensen, C. B., 1997. Urea and amphibain water economy. Comp. Biochem. Physiol. 117: 161-1 70. Kennedy, I. R., 1992. "Acid Soil and Acid Rain". Second edition. Wiley, New York. Kernaghan, G., Berch, S. and Carter, R., 1995. Effect of urea fertilization on ectomycorrhizae of 20-yearold Euga heterophylh. Can. J. For Res. 25: 891-901. kesecker, J. M. and Blaustein, A. R., 1995. Synergism between UV-B radiation and a pathogen magnifies amphibian embryo mortality in nature. Proc. Nut. Acad. Sci. USA 92: 11049-1 1052. kesecker, J. M. and Blaustein, A. R., 1997. Population differences in responses of red-legged frogs (Rana aurora) to introduced bullfrogs. Ecology 78: 17521760. Kladivko, E. J., VanScoyoc, G. E., Monke, E. J., Oates, K. M. and Pask, W., 1991. Pesticide and nutrient movement into subsurface tile drains on a silt loam soil in Indiana. J . Enuiron. Qual. 20: 264-270. Knepp, G. L. and Arkin, G. F., 1973. Ammonia toxicity levels and nitrate tolerance of channel catfish. Prog. Fish-Culturist 35: 221-224. Konlkoff, M., 1975. Toxicity of nitrite to channel catfish. Prog. Fish-Culturist 37: 96-98. Kraft, G. J . and Stites, W., 2003. Nitrate impacts on groundwater from irrigated-vegetable systems in a humid north-central US sand plain. Agr. Ecosyst. Enuiron. 100: 63-74. Kross, B. C., Ayebo, A. D. and Fuortes, L. J., 1992. Methemoglobinemia: nitrate toxicity in rural America. Am. Fam. Phys. 46: 183-188. Kross, B. C., Hallberg, G. R., Bruner, D. R., Cherryholmes, K. and Johnson, J. K., 1993. The Nitrate Contamination of Private Well Water in Iowa. Am. J. Public Health 83: 270-272. Laposata, M. M. and Dunson, W. A., 1998. Effects of boron and nitrate on hatching success of amphibian eggs. Arch. Environ. Contam. Toxicol. 35: 615-619. Laposata, M. M. and Dunson, W. A., 2000. Effects of spray-irrigated wastewater effluent on temporary pond-breeding amphibians. Ecotoxicol. Enuiron. Safety 46: 192-201. Lease, H. M., Hansen, J. A,, Bergman, H. L. and Meyer, J. S., 2003. Structural changes in gills of Lost River suckers exposed to elevated pH and ammonia concentrations. Comp. Biochem. Physiol. C Toxicol. Pharmacol. Endocrinol. 134: 491-500.

3181

Leips, J. and Travis, J., 1994. Metamorphic responses to changing food levels in two species of hylid frogs. Ecology 75: 1345-1 356. Leonard, W. l?, Brown, H. A,, Jones, L. L. C., Mcallister, K. R. and Storm, R. M., 1993. "Amphibians of Washington and Oregon". Seattle Audubon Society, Seattle, Washington. Lewis, W. M. Jr. and Morris, D. P, 1986. Toxicity of nitrite to fish: a review. Trans. Am. Fisher Soc. 115: 183-194. Licht, P, Feder, M. E. and Bledsoe, S., 1975. Salinity tolerance and osmoregulation in the salamander Batrachoseps. J. Comp. Physiol. 102: 123-134. Lijinsky, W., Conrad, E. and de Bogart, R. V., 1972. Carcinogenic nitrosamines formed by druglnitrite interactions. Nature 239: 165-167. Liu, R. H. and Hotchkiss, J. H., 1995. Potential genotoxicity of chronically elevated nitric-oxide a review. Mutat. Research-Reviews Genet. Toxicol. 339: 73-89. Madison, R. J. and Brunett, J. O., 1985. "Overview of the occurrence of nitrate on ground water of the United States". Water supply paper 2275. U.S. Geological Survey, Denver. Magee, P: N., 1971. Toxicity of uitrosamines. There possible human health hazards. Fd.$ Cosmet. Toxicol. 9: 207-218. Marco, A., Blaustein, A. R., 1999. The effects of nitrite on behavior and metamorphosis in cascades frogs (Rana cascadae). Enuiron. Toxicol. Chem. 18: 946-949. Marco, A., Quilchano, C. and Blaustein, A. R., 1999. Sensitivity to nitrate and nitrite in pond-breeding amphibians from the Pacific northwest, USA. Enuiron. Toxicol. Chem. 18: 2836-2839. Marco, A,, Cash, D., Belden, L. K. and Blaustein, A. R., 2001. Sensitivity to urea fertilization in three amphibian species. Arch. Enuiron. Contam. Toxicol. 40: 406-409. Margiocco, C., Arillo, A., Mensi, P: and Schenone, G., 1983. Nitrite bioaccumulation in Salmo gardnierii Rich. and hematological consequences. Apuat. Toxicol. 3: 261-270. Mason, C. F., 1991. "Biology of Freshwater Pollution" Longman Scientific and Technical, Harlow. Matson, P A., McDowell, W. H., Townsend, A. R. and Vitousek, P: M., 1999. The globalization of N deposition: ecosystem consequences in tropical environments. Biogeochemistry 46: 67-83. Matson, P: A,, Parton, W. J., Power, A. G. and Swift, M. J., 1997. Agricultural intensification and ecosystem properties. Science 277: 504-509. McClanahan, L. L., 1972. Adaptations of the spadefoot toad, Scaphiopus couchii, to desert environments. Comp. Biochem. Physiol. 20: 73-99. McCoy, E. F., 1972. "Role of Bacteria in the Nitrogen Cycle in Lakes". 16010, EHR 03/72. Environmental Protection Agency, Water Pollution Control Service, Washington, DC. McCrary, J. E., Heagler, M. G., 1997. The use of a simultaneous multiple species acute toxicity test to compare the relative sensitivities of aquatic organisms to mercury. J. Enuiron. Sci. Health, Part A-Enuiron. Sci. Engin. Toxic and Hazard. Subs. Cont. 32: 73-81.

3182

AMPHIBIAN BIOLOGY

McDowell, L. L., McGregor, K. C., 1979. Nitrogen and phosphorus losses in runoff from no-till soybeans. Trans. ASAE 23: 643-648. Meister, R. T (ed), 1995. "Farm Chemicals Handbook. Meister Publishing Co., Willoughby, Ohio.

Ortiz-Santaliestra, M. E., Marco, A. and Lizana, M., 2005. Sensitivity and behavior of the Iberian newt, Triturus boscai, under terrestrial exposure to ammonium nitrate. Bull. Enuiron. Contam. Toxicol. 75: 662-669.

Merkens, J. C. and Downing, K. M., 1957. The effect of tension of dissolved oxygen on the toxicity of unionized ammonia to several species of fish. Ann. Appl. Biol. 45: 521-527.

Ortiz-Santaliestra, M. E., Marco, A,, Fernandez, M. J. and Lizana, M., 2006. Influence of developmental stage on sensitivity to ammonium nitrate of aquatic stages of amphibians. Enuiron. Toxicol. Chem. 25: 105-1 11.

Minshall, G. W., Robinson, C. T and Lawrence, D. E., 1997. Postfire responses of lotic ecosystems in Yellowstone National Park, USA. Can. J. Fkh. Aquat. Sci. 54: 2509-2525.

Osborne, L. L. and Kovacic, D. A., 1993. Riparian vegetated buffer strips in water-quality restoration and stream management. Freshw. Biol. 29: 243-258.

Moore, M. K. and Townsend, V. R. Jr, 1998. The interaction of temperature, dissolved oxygen and predation pressure in a n aquatic predator-prey system. Oikos 81: 329-339. Morey, S. and Reznick, D., 2000. A comparative analysis of plasticity in larval development in three species of spadefoot toads. Ecology 81: 1736-1749. Morey, S. and Reznick, D., 2001. Effects of larval density on postmetamorphic spadefoot toads (Spea hammondii). Ecology 82: 5 10-522. Morey, S. R. and Reznick, D. N., 2004. The relationship between habitat permanence and larval development in California spadefoot toads: field and laboratory comparisons of developmental plasticity. Oikos 104: 172-190. Mueller, D. K., Ruddy, B. C. and Battaglin, W. A,, 1997. Logistic model of nitrate in streams of the uppermidwestern United States. J. Enuiron. Qual. 26: 1223-1230. Muir, P. R., Sutton, D. C. and Owens, L., 1990. Nitrate toxicity to Penaew monodon protozoea. Mar. Biol. 108: 67-71. Muscutt, A. D., Harris, G. L., Bailey, S. W. and Davies, D. B., 1993. Buffer zones to improve water-quality - a review of their potential use in UK agriculture. Agric. Ecosys. Enuiron. 45: 59-77. National Research Council, 1972. $Hazards of Nitrate, Nitrite, and Nitrosoamines to Man and Livestock. Pp. 46-75 in "Accumulation of Nitrate". National Academy of Sciences, Committee on Nitrate Accumulation, Ag. Board, Div. of Biology & Agriculture, Wash., DC. National Research Council, 1979. "Ammonia". University Park Press, Baltimore. Nebeker, A. V. and Schuytema, G. S., 2000. Effects of ammonium sulfate on growth of larval Northwestern salamanders, red-legged and Pacific treefrog tadpoles, and juvenile fathead minnows. Bull. Enuiron. Cant. Toxicol. 64: 271-278. Nussbaum, R. A., Brodie, E. Jr. and Storm, R. M., 1983. "Amphibian and Reptiles of the Pacific Northwest". University of Idaho Press, Moscow, Idaho. OECD, 1986. "Water Pollution by Fertilizers and Pesticides". Organization for Economic Cooperation and Development, Paris. Oldham, R. S., Lathan, D. M., Hilton-Brown, D., Towns, M., Cooke, A. S. and Burn, A,, 1997. The effect of ammonium nitrate ferulizer on frog (Rana temporaria) survival. Agric. Ecosys. Enuinm. 61: 69-74. Ortiz, M. E., Marco, A., Saiz, N. and Lizana, M., 2004. Impact of ammonium nitrate o n growth and survival of six European amphibians. Arch. Enuiron. Contam. Toxicol. 47: 234-239.

Owens, L. B., VanKeuren, R. W. and Edwards, W. M., 1983. Nitrogen loss from a high-fertility, rotational pasture program. J. Enuiron. Qual. 12: 346-350. Palachek, R. M. and Tomasso, J. R., 1984a. Nitrite toxicity to fathead minnows: effect of fish weight. Bull. Enuiron. Contam. Toxicol. 32: 238-242. Palachek, R. M. and Tomasso, J. R., 1984b. Toxicity of nitrite to channel catfish (Zctalurus punctatus), tilapia (Tilapia aurea), and largemouth bass (Micropterus salmozdes):evidence for a nitrite exclusion mechanism. Can. J. Fkh. Aquat. Sci. 41: 1739-1744. Peoples, M. B., Freney, J. R. and Mosier, A. R., 1995. Minimizing gaseous losses of nitrogen. Pp. 565-602 in "Nitrogen Fertilization in the Environment", ed by P. E. Bacon. Marcel Dekker, New York. Perrone, S. J. and Meade, T. L., 1977. Protective effect of chloride on nitrite toxicity to coho salmon (Oncorhynchus kisutch). J. Fish. Res. Board. Can. 34: 486-492. Pfennig, D. W., Mabry, A. and Orange, D., 1991. Environmental causes of correlations between age and size at metamorphosis in Scaphiopus multi-, plicatw. Ecology 72: 2240-2248. Pierce, B. A. and Wooten, D. K., 1992. Genetic variation in tolerance of amphibians to low pH. J. Herpetol. 26: 422-429. Pierce, B. A,, Margolis, M. A. and Nirtaut, L. J., 1987. The relationship between egg size and acid tolerance in Rana sylvatica. J. Herpetol. 21: 178-1 84. Pilliod, D. S., Bury, R. B., Hyde, E. J., Pearl, C. A. and Corn, P. S., 2003. Fire and amphibians in North America. Fox Ecol. Man. 178: 163-181. Podrabsky, J. E., Hrbek, T. and Hand, S. C., 1998. Physical and chemical characteristics of ephemeral pond habitats in the Maracaibo basin and Llanos region of Venezuela. Hydrobiologia 362: 67-78. Pough, F. H., Smith, E. M., Rhodes, D. H. and Collazo, A., 1987. The abundance of salamanders in forest stands with different histories of disturbance. For. Ecol. Man. 20: 1-9. ?? check pages 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. Pounds, J. A., Fogden, M. P. L. and Campbell, J. H., 1999. Biological response to climate change on a tropical mountain. Nature 398: 61 1-615. Pratt, I! F., 1984. Nitrogen use and nitrate leaching in irrigated agriculture. Pp. 319-333 in "Nitrogen in Crop Production", ed by R. D. Hauck. American Society of Agronomy, Madison. Puckett, L. J., 1995. Identifying the major sources of nutrient water pollution. Enuiron. Sci. Technol. 29: 408-414.

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION Ralph, S. and Petras, M., 1997. Genotoxicity monitoring of small bodies of water using two species of tadpoles and the alkaline single cell gel (comet) assay. Enwirun. Molec. Mutagen. 29: 418430. Rebelo, M. F., Santos, E. A., Monserrat, J. M., 1999. Ammonia exposure of Chasmagnathus granulota (Crustacea, Decapoda) Dana, 1851: Accumulation in haemolymph and effects on osmoregulation. Camp. Biochem. Physiol. A Mol. Integx Physiol. 122: 429-435. Ritter, W. F., Scarborough, R. W. and Chirnside, A E. M., 1991. Nitrate leaching under irrigated corn. J. Irrig. Drain. Eng. ASCE 119: 544-553. Romansic, J. M., Diez, K. A,, Higashi, E. M. and Blaustein, A. R., 2006. Effects of nitrate and the pathogenic water mold Saprolegnio on survival of amphibian larvae. Dis. Aquat. Organ. 68: 235-243. Rouse, J. D., Bishop, C. A. and Struger,J., 1999. Nitrogen pollution: an assessment of its threat to amphibian survival. Environ. Health Perspec. 107: 799-803. Rubin, A. J. and Elmaraghy, G. A., 1977. Studies on the toxicity of ammonia, nitrate and their mixtures to guppy fry. Water Res. 11: 927-935. Russo, R. C., 1985. Ammonia, nitrite and nitrate. Pp. 455-471 in "Fundamentals of Aquatic Toxicology", ed by G. M. Rand and S. R. Petrochelli. Hemisphere Publishing Corporation, Washington. Russo, R. C. and Thurston, R. 17, 1977. The acute toxicity of nitrite to fishes. In "Recent Advances in Fish Toxicology", ed by R. A. Tubb. U.S. Environmental Protection Agency, Ecological Research Series EPA-60013-77-085, Corvallls. Russo, R. C., Smith, C. E. and Thurston, R. V., 1974. Acute toxicity of nitrite to rainbow trout (Salmo gardnierii). J. Fish. Res. Board Can. 31: 1633-1655. Savage, R. M., 1961. "The Ecology and Life History of the Common Frog (Rana temporaria temporaria)". Pitman and Sons Ltd., London. Scarano, G., Saroglia, M. G., Gray, R. H. and Tibaldi, E., 1984. Hematological responses of sea bass Dicentrarchus labrax to sublethal nitrite exposures. Trans. Am. Fish. Sac. 113: 360-364. Schmuck, R., Geise, W. and Linsenmair, K E., 1994. Life cycle strategies and physiological adjustments of reedfrog tadpoles (Amphibia, Anura, Hyperoliidae) in relation to environmental conditions. Copeia 1994: 996-1007. Schneeweiss, N. and Schneeweiss, U., 1997. Mortality of amphibians as a consequence of mineral fertilizing. Salamandra 33: 1-8. Scholefield, D., Lord, E. I., Rodda, H. J. E. and Webb, B., 1996. Estimating peak nitrate concentrations from annual nitrate loads. J. Hydrol. 186: 359-373. Schuytema, G. S. and Nebeker, A. V., 1999a. Comparative toxicity of ammonium and nitrate compounds to Pacific treefrog and African clawed frog tadpoles. Envtl. Toxicol. Chem. 18: 2251-2257. Schuytema, G. S. and Nebeker, A. V., 1999b. Comparative effects of ammonium and nitrate compounds on Pacific treefrog and African clawed frog embryos. Arch. Environ. Con. Toxicol. 36: 200-206. Schuytema, G. S. and Nebeker, A. V , 1999c. Effects of Ammonium Nitrate, Sodium Nitrate and Urea on Red-legged Frogs, Pacific Treefrogs, and African Clawed Frogs. Bull. Environ. Contam. Toxicol. 63: 357-364.

3183

Scott, D. E., 1994., The effect of larval density on adult demographic traits in Ambystomu opacum. Ecology 75: 1383-1396. Scott, G. and Crunkilton, R. L., 2000. Acute [$ word?) an chronic toxicity of nitrate to fathead minnows (Pi*& promelas), Ceriodaphnia dubia, and Daphnia magna. Environ. Toxicol. Chem. 19: 2918-2922. Semlitsch, R. D., 2002. Critical elements for biologically based recovery plans of aquatic-breeding amphibians. Cons. Biol. 16: 61-629. Semlitsch, R. D. and Bodie, J. R., 2003. Biological criteria for buffer zones around wetlands and riparian habitats for amphibians and reptiles. Cons. Biol. 17: 1219-1228. Semlitsch, R. D., Scott, D. E. and Pechmann, J. H. K., 1988. Time and size at metamorphosis related to adult fitness in Ambystoma talpoideum. Ecology 69: 184-192. Shoemaker, V. H., Hillman, S. S., Hdlyard, S. D., Jackson, D. C., McClanahan, L. L., Withers, F! C. and Wygoda, M. L., 1992. Exchange ofwater, ions, and respiratory gases in terrestrial amphibians. Pp. 125-150 in "Environmental Physiology of Amphibians", ed by M. E. Feder and W. W. Burggren. The University of Chicago Press, Chicago. Simon, M. P., Vatnick, I., Hopey, H. A,, Butler, K., Korver, C., Hilton, C., Weimann, R. S. and Brodkin, M. A., 2002. Effects of acid exposure on natural resistance and mortality of adult Rana pipiens. J . Herpetol. 36: 697-699. Smart, W. A., 1981. Investigations of the toxic mechanisms of ammonia to fish gas exchange in rainbow trout (Salmo gairdneri) exposed to acutely lethal concentrations. J. Fish. Biol. 12: 93-104. Smil, V., 2002. Biofixation and nitrogen in the biosphere and in global food production. Pp. 7-9 in "Nitrogen Fixation: Global Perspectives", ed by T. Finan, M. O'Brian. D. Lavzell. , , K. Vessev and W. Newton. CAB lnternaiional, Wallingford. ' Smith, C. E. and Russo, R. C., 1975. Nitrite induced methemoglobinemia in rainbow trout. Progress. Fish Cultux 37: 150-152. Smith, D. C., 1987. Adult recruitment in chorus frogs: effects of size and date at metamorphosis. Ecology 68: 344-350. Smith, G. R., Vaala, D. A,, Dingfelder, H. A. and Temple, K. G., 2004. Effects of nitrite on bullfrog (Rana catesbeiana) tadpoles from Central Ohio, USA. Bull. Environ. Contam. Toxicol. 72: 1012-10 16. Smith, G. R., Temple, K. G., Vaala, D. A. and Dingfelder, H. A,, 2005. Effects of nitrate on the tadpoles of two Ranids (Rana catesbeiana and R . clamitans). Arch. Environ. Contam. Toxicol. 49: 559-562. Sopper, W. E., 1986. Penn State's "Living filter": twentythree years of operation. Pp. 406-419 in "The Forest Alternative for Treatment and Utilization of Municipal and Industrial Wastes", ed by D. W. Cole, C. L. Henry and W. L. Nutter. University of Washington Press, Seattle. Sparling, D. W. and Harvey, G., 2006. Comparative toxicity of ammonium and perchlorate to amphibians. Bull. Environ. Contam. Toxicol. 76: 2 10-2 17. Spencer, C. N., Gabel, K. 0 . and Hauer, F. R., 2003. Wildfire effects on stream food webs and nutrient dynamics in Glacier National Park, USA. Forest Ecol. Man. 178: 141-153.

3184

AMPHIBIAN BIOLOGY

Srivastava, G. J. and Srivastava, 0. F!, 1979. Ureainduced histopathology in skin and gastric lining of teleost Channa punctatus (Bloch). Znd. J. Exptl. Biol. 17: 840.

Tucker, C. S., Francis-Floyd, R. and Beleau, M. H., 1989. Nitrite-induced anemia in channel catfish, Ictalunw punctatus Rafinesque. Bull. Enuiron. Contam. Toxicol. 43: 295-301.

Stephan, C. E., 1975. "Methods for Acute Toxicity Tests with Fish, Macroinvertebrates and Amphibians". EPA-66013-75-009. U.S. Environmental Protection Agency, Corvallis.

Tucker, C. S., MacMillan, J. R. and Schwedler, T. E., 1984. Influence of Edwardsiella ictaluri septicemia on nitrite-induced methemoglobinemia in channel catfish (Zctalurus punctatus). Bull. Enuiron. Contam. Toxicol. 32: 669-673.

Sterner, R. W., 1989. Resource competition during seasonal succession toward dominance by cyanobacteria. Ecology 70: 229-245. Sullivan, B. and Riggs, A., 1964. Haemoglobin reversal of oxidation + polymerization in turtle red cells. Nature 204 (496): 1098-[NEED REST OF PAGINATON, OR WAS IT ONLY ONE PAGE?] Sullivan, K. B. and Spence, K. M., 2003. Effects of sublethal concentrations of atrazine and nitrate on metamorphosis of the African clawed frog. Enuiron. Toxicol. Chem. 22: 627-635. Swigert, J. F! and Spacie, A., 1983. "Survival and Growth of Warm Water Fishes Exposed to Ammonia Under Low Flow Conditions". Water Resources Research Technical Report 157. Purdue University, West Lafayette. Tamm, C. O., 1991. "Nitrogen in Terrestrial Ecosystems". Springer-Verlag, Berlin. Tamminga, K., 1995. Is the living filter sustainable? Assesing the land application of municipal effluent. Enuiron. Pro$ 17: 290-300. Tamminga, S., 2003. Pollution due to nutrient losses and its control in European animal production. Liustck. Prod. Sci. 84: 101-111. Tilman, D., 1998. The greening of the green revolution. Nature 396: 21 1-212. Tilman, D., 1999. Global environmental impacts of agricultural expansion: The need for sustainable and efficient practices. Proc. Nut. Acad. Sci. USA. 96: 5995-6000. Tietge, J. E., Ankley, G. T, DeFoe, D. L., Holcombe, G. W., Jensen, K. M., 2000. Effects of water quality on development of Xenopus laevis. a frog embryo teratogenesis assay - Xenopus assessment of surface water associated with malformations in native anurans. Enuiron. Toxicol. Chem. 19: 2 114-2 12 1. Tomasso, J. R. and Carmichael. G. J., 1986. Acute toxicity of ammonia, nitrite, and nitrate to the guadalupe bass, Micropterus treculi. Bull. Enuiron. Contam. Toxicol. 36: 866-870.

Twitchen, I. D. and Eddy, F. B., 1994. Effects of ammonia on sodium balance in juvenile rainbow trout Oncorhynchus mykiss Walbaum. Aquat. Toxicol. 30: 2 7 4 5 . United States EPA, 1985. "Ambient Quality Criteria for Ammonia". EPA 44015-85-001. Environmental protection agency, Washington, DC. United States EPA, 1986. "Quality Criteria for Water". EPA 44015-86-001. United States Environmental Protection Agency (US EPA). Washington DC. United States EPA, 1989. "Ambient Water Quality Criteria for Ammonia (Saltwater)-1989". EPA 4401 5-88-004 Office of Water Regulations and Standards, Criteria and Standards, Washington, DC. United States EPA, 1998. "1998 Update of Ambient Water Quality Criteria for Ammonia". EPA 822-R98-008. Office of Water Environmental Protection Agency, Washington, DC. Vedel, N. E., Korsgaard, B. and Jensen, F. B., 1998. Isolated and combined exposure to ammonia and nitrite in rainbow trout (Oncorhynchus mykiss): Effects on electrolyte status, blood respiratory properties and brain glutaminelglutamate concentrations. Aquat. Toxicol. 41: 325-342. Vitousek, F! M., 1994. Beyond global warming: ecology and global change. Ecology 75: 1861-1876. Vitousek, F! M., Aber, J. D., Howarth, R. W., Likens, G. E., Matson, F! A., Schindler, D. W., Schlesinger, W. H. and Tilman, D. G., 1997. Human alteration of the global nitrogen cycle: Sources and consequences. Ecol. Applic. 7: 737-750. Wajsbrot, N., Gasit, A., Krom, M. D. and Popper, D. M., 1991. Acute and chronic toxicity of ammonia to juvenile gilthead seabream Sparus aurata under reduced oxygen levels. Aquaculture 92: 277-288. Wassersug R. J. and Seibert, E. A., 1975. Behavioral responses of amphibian larvae to variation in dissolved oxygen. Copeia 1975: 86-103. Watt, F! J. and Jarvis, El, 1997. Survival analysis in palmate newts exposed to ammonium nitrate agricultural fertilizer. Ecotoxicol 6: 355-362.

Tomasso, J. R., Simco, B. A., Davis, K. B., 1979. Chloride inhibition of nitrite-induced methemoglobinemia in channel catfish (Zctalurus punctatw). J. Fish. Res. Board Can. 36: 1141-1 144.

Watt, F! J. and Oldham, R. S., 1995. The effect of ammonium nitrate on the feeding and development of larvae of the smooth newt, Triturus uulgaris (L.), and on the behaviour of its food source. Daphnia. Freshw. Biol. 33: 319-324.

Trama, F. B., 1954. The acute toxicity of some common salts of sodium, potassium and calcium to the common bluegill (Lepomis macrochirus Rafinesque). Proc. Acad. Nut. Sci. Phil. 106: 185-205.

West, N. H. and Burggren, W. W., 1982. Gill and lung ventilatory responses to steady-state aquatic hypoxia and hyperoxia in the bullfrog tadpole. Respi. Physiol. 47: 165-176.

Travis, J., 1984. Anuran size at metamorphosis: experimental test of a model based on intraspecific competition. Ecology 65: 1155-1 160. Tucker, C. S. and Schwedler, 7: E., 1983. Acclimation of channel catfish (Zctalunw punctatus) to nitrite. Bull. Enuiron. Contam. Toxicol. 30: 5 16-52 1.

Westin, D. T., 1974. Nitrate and nitrite toxicity to salmonoid fishes. Prog. Fish-Cult. 36: 86-89. Weyer, F! J., Cerhan, J. R., Kross, B. C., Hallberg, G. R., Kantamneni, J., Breuer, G., Jones, M. F!, Zheng, W. and Lynch, C. F., 2001. Municipal drinlung water nitrate level and cancer risk in older women: the Iowa Women's Health Study Epidemiology 12: 327-338.

MARC0 and ORTIZ-SANTALIESTRA: NITROGEN POLLUTION

3185

Weyrauch, S. L. and Grubb, T C., 2004. Patch and landscape characteristics associated with the distribution of woodland amphibians in an agricultural fragmented landscape: an informationtheoretic approach. Biol. Cons. 115: 443-450.

Wright, P. M. and Wright, P. A,, 1996. Nitrogen metabolism and excretion in bullfrog (Rana catesbeiana) tadpoles and adults exposed to elevated environmental ammonia levels. Ph~siol.2001. 69: 1057-1078.

Whitfield, M., Butler, R. A. and Covington, Q. K., 1985. The determination of pH in estuarine waters. I. Definition of pH scales and the selection of buffers. Oceanologica Acta 8: 423-432.

Xu, Q. and Oldham, R. S., 1997. Lethal and Sublethal Effects of Nitrogen Fertilizer Ammonium Nitrate on Common Toad (Bufo bufo) Tadpoles. Arch. Euiron. Contam. Toxicol. 32: 298-303.

Williams, E. M. and Eddy, F. B., 1989. Effect of nitrite on the embryonic development of Atlantic Salmon (Salmo salar). Can. J. Fish. Aquat. Sci. 46: 1726-1729.

Yadav, J. S. and Seth, N., 1998. Effect of NOx on the somatic chromosomes of goldsmiths. Environ. Health Perspec. 106: 643-647.

Williams, M. W., Baron, J. S., Caine, N., Sommerfeld, R. and Sanford Jr., R., 1996. Nitrogen saturation in the Rocky Mountains. Environ. Sci. Technol. 30: 640-646.

CHAPTER 8

Evaluating the Impact of Pesticides i n ~rnphibianDeclines Michelle D. Boone, Carlos Davidson and Christine Bridges-Britton

I. Introduction to the Problem II. Why Linking Declines with Pesticides is Difficult

V. Assessing Causality VI. Conclusions VI I. Acknowledgements

Ill. How Pesticides Elicit an Effect VIII. References IV. Approaches to Studying Declines A. Laboratory Studies B. Mesocosm Field Studies C. Natural Field Studies

I. INTRODUCTION TO THE PROBLEM

I

N the global ecosystem, elements move through the earth's crust, bodies of water, atmosphere, and plant and animal life via the food web and circulation of air and water. It is then likely, if not inevitable, that the consequences of application of pesticides result in global distribution and dispersion of potentially toxic compounds through these same natural processes; these toxins could compromise plant and animal life across habitats. For instance, organisms in remote arctic marine ecosystems have been found to accumulate high levels of contaminants like PCBs (reviewed by Bard 1999) far from the sites of initial pollution; such results indicate that the site of contamination is not fixed and that contamination can gradually diffuse throughout the ecosystem. Many of the early synthetic organochlorine pesticides still persist in the environment because they are stable and biomagnify in the food web. Although many of these pesticides were banned in the United States and Europe in the 1970s (Peterle 1991), many continue to be used in developing nations. New-generation pesticides break down quickly and are often thought to be less harmful than early synthetic pesticides, although they can also be present chronically at low-levels and be transported aerially, in runoff, and through above-ground and below-ground water sources (e.g., Thurman and Cromwell 2000). Short-term exposure to short-lived contaminants can also, therefore, exert chronic effects on organisms and may contribute to population declines (Cowman and Mazanti 2000). For these reasons, exposure to pesticides is repeatedly presented as a potential contributor to amphibian population declines and a threat to biodiversity in general (Corn 1994; Semlitsch 2003). The role that contaminants may play in declines has been addressed now in a number of volumes (including Sparling et al. 2000; Linder et al. 2003; Semlitsch 2003; Stuart et al. 2004; Lannoo 2005). Amphibian

BOONE ETAL.: IMPACT O F PESTICIDES I N AMPHIBIAN DECLINES

3185

declines continue to be a concern, particularly in areas where population density has decreased at the landscape level and where mass die-offs have occurred. It is not clear if populations that have not experienced declines are free from danger, if trouble has not yet reached them, or if they have already adapted to the hazard. Understanding how the major threats of habitat destruction and alteration, disease and pathogens, introduced species, global change, and contamination may singly and interactively influence populations is essential to pinpoint sources of danger and to construct solutions that remedy or reduce the impact. To address the question of how pesticides may contribute to the risk of amphibian population declines, this chapter evaluates (1) why linking declines with pesticides may be difficult, (2) the role pesticides may play in amphibian populations, (3) the approaches and results of studying declines, and (4) a strategy for assessing the causation and effects of contaminants in natural systems. 11. WHY LINKING DECLINES WITH PESTICIDES IS DIFFICULT

When Rachel Carson published "Silent Spring" in 1962, she was able to pull together large amounts of anecdotal and quantitative data to convincingly demonstrating that a number of chemical toxicants were negatively affecting organisms across taxonomic groups. So, then, if chemical contaminants are in fact linked to amphibian declines, should one not be able to make this connection relatively easily for a single taxonomic group? Or, is it possible that even if contamination by pesticides is a significant contributor to amphibian declines, making these links could still be difficult? Certainly many of the pesticides in use today have some advantages over the early synthetic pesticides: they are less likely to bioaccumulate or magnify, they are less persistent in the environment, and federal regulations often maintain them at concentrations below limits lethal to non-targeted wildlife. Regulations banning the use of the early synthetic pesticides were meant to protect humans and non-targeted wildlife from the ill effects of contaminants, so those used presently should be "more benign," although reports of presentday chemicals disrupting endocrine system function make this proposal questionable (see review by Colburn et al. [1996]). Chemical effects on organisms, therefore, should be expected to be more subtle. Yet, chemicals like DDT also had subtle effects on bird reproduction, and scientists were still able to make cause-and-effect linkages. While it is believed that the role of pesticides in amphibian declines may be determined through experimental and correlational studies, there are a number of reasons why making the link between pesticides and amphibian declines in nature may be difficult to achieve. Species differ in their sensitivity to pesticides. In initial discussions addressing whether declines were occurring, or if declines were just the results of natural variation in time and space (Pechmann et al. 1991; Pechmann and Wilbur 1994), researchers pointed out that while one species in a community may go extinct, other species appear to be unaffected or actually increase in abundance. Research has demonstrated variation within (Bridges and Semlitsch 2001) and among (Bridges and Semlitsch 2000) amphibian species in sensitivity to lethal levels of contamination, which supports observations in nature that some amphibian populations could decline due to pesticides (andlor other factors) while others in the same community appear to be unaffected. Linking declines with pesticides may be difficult because species are differentially affected by pesticides (as well as other stressors) and because a community is unlikely to have a unanimous response to stressors. While differences in species' responses to pesticides may make declines difficult to detect, variation in pesticide concentrations and application rates, and the types of pesticides used fluctuate spatially and temporally, which incorporates another component of variation into the equation. Farmers routinely rotate their crops, which leads to changes in the kinds or amounts of pesticides used on a particular parcel of land. The amount of pesticide that drifts into nearby habitats or that runs off into aquatic habitats will also fluctuate with differences in seasonal weather conditions, such as the amount of wind or rain - even if pesticide application rates are consistent. Furthermore, a chemical's persistence will also

3188

AMPHIBMN BIOLOGY

vary depending on environmental temperatures as well as amount of rainfall in a given year. This means that amphibians living in areas characterized by periodic use of pesticides may experience pulses of exposure in varied contexts. Making clear links between any particular pesticide and any exposure scenario (e.g., timing of exposure, interval of exposure, environmental conditions) with amphibian declines may therefore be extremely diff~cultbased solely on field observations, because field conditions will almost certainly vary every year. It is also conceivable that while pesticide exposure may play a role in declines, the impact may be tied to the presence of other stressors. It is often difficult to ascertain whether population declines are due to compounds applied to crops, or to concomitant ecological alterations such as habitat fragmentation or diversion of water. Many correlated factors must be considered when trying to determine the effects of pesticides on amphibians. Hecnar (1995) found that the amount of woodland surrounding a pond is the most important factor in determining species richness in southern Ontario ponds. Beja and Alcazar (2003) reported that the change from temporary bodies of water to permanent ones was the most important factor dictating amphibian presence or absence on agricultural lands. Knutson et al. (2004) found that ponds near grazed land and row crops experienced increases in turbidity, phosphorus, nitrogen, and low concentrations of agricultural chemicals. Factors such as presence or absence of fish, amount of vegetation, and area of the pond were more important in determining the presence of a species than were landscape variables. In southern Ontario, Bishop et al. (1999) examined species diversity and density, as well as hatching success and deformities, among marshes embedded in an agricultural matrix. They concluded that amphibians were more affected within agricultural areas than either upstream or downstream of agricultural lands; they suggested nutrient run-off to be the culpable agent. Bonin et al. (199'7) also pointed out the difficulties in distinguishing between other agricultural practices and the application of chemicals to crops as the cause of decline of amphibian populations. Determining critical interactions between multiple stressors will take time, and the presence of multiple factors increases the difficulty of directly tying declines to pesticides. Current research and thought suggest that multiple stressors may be the likely cause of amphibian declines (Carey et al. 2001; Linder et al. 2003), and there is some evidence of synergistic interactions among factors (Little et al. 2000; Relyea and Mills 2001; Boone et al. 2004; Bridges-Britton et al., in review). Although post-application levels may be high enough to induce mortality in cases of direct exposure, more often pesticides will be transported aerially or in runoff at sublethal levels (LeNoir et al. 1999) so that direct lethal effects would not be anticipated. Even low levels of contaminants, however, could contribute to declines through interactions with other stressors. Contaminants are also able to select for resistant genotypes that can disguise the role pesticides may be having. Large-scale monocultures now dominate many landscapes where diverse amphibian populations formerly thrived (Burnett 1997; Kupferberg 1997), perhaps due to selection favoring species more tolerant of stress and eliminating those that did not develop resistance. Meanwhile, amphibians that have the genetic ability to adapt to chemical contamination may still be hindered by agriculturally induced fragmentation of their habitats. Furthermore, populations often are not completely isolated from each other but rather are part of a network of populations known as a metapopulation. Among populations within a metapopulation, gene flow can ameliorate the net effect of a stressor on one of the populations. Semlitsch (2000) pointed out that two primary factors govern the dynamics of amphibian metapopulations: (1) the number and density of amphibians moving among ponds and (2) the probability of individuals reaching ponds. Thus, a reduction in number of ponds, the inimical nature of the terrestrial habitat surrounding ponds, and chemical contamination of ponds (which may make some ponds "sinks" [Rowe et al. 20011) are all factors likely to be contributing to the demise of amphibians within an agricultural setting. Other stressors, either natural or anthropogenic, may make it difficult to distinguish subtle effects of contaminants on metamorphosis. Small size of metamorphs, for example,

BOONE ET AL.: IMPACT OF PESTICIDES IN AMPHIBIAN DECLINES

3189

can result either from exposure to contaminants or from high densities of larvae. Therefore, although one knows (in general) that herbicides reduce body mass of anurans at metamorphosis, finding small metamorphs in agricultural areas in ponds known or suspected to be contaminated is not particularly informative, given this could result either from exposure to contaminants or from high larval densities. It is important to demonstrate the general patterns of responses experimentally and to know how particular pesticides or classes of pesticides alter those responses. One can then use an understanding of how the principle components of the community (amphibians, food resources, predators) are affected by contaminants to predict the consequences of contaminants in the field over time. Furthermore, knowing how various natural and anthropogenic factors interact with contaminants is requisite for evaluating how these factors intensify or reduce the contaminant's effect. Relyea and Mills (2001) found that treefrogs (Hyla versicolor) reared in the laboratory were significantly more affected by an interaction of predators and contaminants than by either alone and Boone and Semlitsch (2001, 2002) showed that manipulations of competition, predation, and the drying of ponds influenced the magnitude of the chemical effect. Bridges-Britton et al. (in review) demonstrated that susceptibility to a pesticide was greater in the presence of Saprolegnia or under ultraviolet radiation. Relating declines in the wild to pesticides may be difficult for all the reasons listed above, and there seems to be no experimentation in progress designed to clarift. this link. While laboratory and field mesocosm studies and correlational studies are all important for understanding the effects of pesticides on individual species, and on population and community dynamics, there are some critical pieces missing. Demonstration that contaminants affect hatching success, length of larval period, survival to metamorphosis and body-size at metamorphosis are important, but do not definitively reveal how population persistence will be affected over time. For example, unless a pesticide (or an interaction of a pesticide with another factor) completely and consistently eliminates a species through reproductive failure (poor hatching success or low survival to metamorphosis), current data are insufficient to make accurate predictions about the population trajectory through time. While reduced hatching success or reduced survival to metamorphosis seem like negative impacts (and they may well be), alone they do not indicate what the long-term impact on the population will be as long as some individuals continue to reach metamorphosis and maintain the population (Vonesh and De la Cruz 2002). Many species of amphibian may adapt to periodic bouts of reproductive failure. If exposure to pesticide, therefore, varies in time and space in ways that influence population dynamics, then the outcome will be confounded by natural variation in the population, making precise assessment of the impact of the contaminant difficult. 111. HOW PESTICIDES ELICIT AN EFFECT

One of the difficulties in evaluating the effects of pesticides on amphibians is that the response can vary from no apparent effect on one hand to mortality resulting in population declines and extinction on the other. Occasionally, amphibian populations have been shown to be reduced or eliminated locally as a result of mortality arising either directly or indirectly from contaminants such as heavy metals (Beyer et al. 1985), pesticides (Lambert 1997), and coal ash (Rowe et al. 2001). It is, however, exceptional that die-offs can be linked clearly to a particular contaminant or pesticide (especially at expected field application rates). The more subtle effects of pesticides are likely to be more common, yet are more difficult to diagnose or recognize in the field. Subtle chemical effects may take a number of forms. Even when environmental levels of pesticides are below concentrations that would induce mass mortality, they may still have an impact on the persistence of a population through changes in body-mass at metamorphosis or time to metamorphosis. Pesticides may also have other sublethal effects, such as endocrine disruption (discussed elsewhere in this volume), altered growth rates of metamorphosed animals, and changes in critical behaviours. Time to metamorphosis and body-size at metamorphosis often are used in toxicological studies because they have been correlated

3190

AMPHIBIAN BIOLOGY

with fitness. Pesticides that can influence either of these have the potential to alter amphibian communities. Both of these traits are plastic, meaning that amphibians can trade early time to metamorphosis for smaller size at metamorphosis, or conversely, later time to metamorphosis for larger size at metamorphosis. In other words, these traits are inversely related to one another for many amphibian species. Mark-recapture studies of wood frogs (Ram sylvatica), mole salamanders (Ambystom tulpozdeum), marbled salamanders (Ambystoma opacum), and chorus frogs (Pseudacris trilseriata) have shown that differences in amphibian size at metamorphosis and time to metamorphosis can influence individuals throughout their lifetimes by affecting reproductive success, survival of post-metamorphic animals and lifetime size (i.e., animals that are larger than others at metamorphosis maintain this size-advantage throughout their lifetime) (Smith 1987; Semlitsch et al. 1988; Berven 1990; Scott 1994). Werner (1986) suggested, however, that species might have different strategies for utilizing aquatic and terrestrial environments. Some experimental studies have demonstrated that some species may be able to compensate for small size at metamorphosis (e.g., American toads; Goater [1994], Boone [2005]). In species with short larval periods, such as the American toad, early metamorphosis may foster greater growth of terrestrial stages prior to winter. It may be necessary to take into account both time to metamorphosis and size at metamorphosis in order predict persistence of populations adequately. Even if exposure to pesticides results in reduced mass (without any compensation in the length of the larval period), population density might not change initially. If smaller metamorphs are less likely, therefore, to survive the winter (as evidence suggests) then over a number of years population size may dwindle to extinction. In natural systems, roughly 3-5% of amphibian eggs reach metamorphosis (Semlitsch 1987; Berven 1990; Semlitsch et al. 1996). Even if this value remains unchanged but individuals leaving the ponds are smaller, then the percentage of juveniles suffering winter mortality would increase (assuming exposure and response to exposure remains constant), even when the pesticide affects only the aquatic stages. As a population decreases in size it may become vulnerable to "small population effects" such as inbreeding depression (as found in hgmented populations of anurans; Hitchings and Beebee [1997]; Beebee [2001]) or increased risk of extinction due to stochastic events (Pechmann et al. 1989; Hels 2002), thereby further exacerbating the problem. A pesticide may affect body-mass at metamorphosis, time to metamorphosis, or survival to metamorphosis, either directly by influencing individual physiology or behaviour, or indirectly by altering the food web. For instance, neurotoxic insecticides may affect amphibians directly by disruption of nervous system function. At high environmental levels this may prove lethal but at sub-lethal levels may influence critical swimming, feeding, and defensive behaviours (Berrill et al. 1993; Semlitsch et al. 1995; Bridges 1997; Eroschenko et al. 2002; Greulich and Pflugmacher 2003; Rohr et al. 2003) or predator-prey interactions (Bridges 1999; Ingermann et al. 2002). In ephemeral ponds, even short-term impairment of feeding behaviour may decrease the probability of an organism reaching metamorphosis before the pond dries. Lowering of the feeding rate likewise may reduce size at metamorphosis and/ or lengthen the time to metamorphosis. Additionally, the breaking down of pesticides in the body incurs a metabolic cost in insects (Appel and Martin 1992; Bernard and Lagadic 1993), crayfish (Rowe et al. 2001) and amphibians (Rowe et al. 1998), which may reduce the amount of energy available for growth and development. There may also be important effects of pesticides on the terrestrial life stage of amphibians. Although the effects of contaminants on terrestrial stages of amphibians have been relatively unstudied, a few investigations have indicated that pesticides may have important effects on juvenile and adult anurans (Hopkins et al. 1998; Mann and Bidwell 1999; Gendron et al. 2003; James et al. 2004). Pesticides can disrupt mating behaviour so that breeding migrations are affected or reproductive success lowered, and, over time, populations decline (Park et al. 2001). Accumulated evidence suggests that pesticides have endocrine-disrupting properties (Hayes et al. 2002, 2003; Goulet and Hontella 2003; MacKenzie et al. 2003; Howe et al. 2004) that can directly affect development of testes, ovaries, and secondary sex characteristics (such as pharyngeal size), thereby impairing reproductive behaviours. Furthermore, James et al.

BOONE ET AL.: IMPACT O F PESTICIDES IN AMPHIBIAN DECLINES

3191

(2004) suggested that amphibians might be more susceptible to population reductions if exposed to pesticides when they are less mobile (i.e., hibernating in contaminated areas). Pesticides can indirectly affect amphibians through changes in the food web. Concentrations of contaminants that cause no direct risk to an amphibian species may have serious impacts on its food resources and thus indirectly result in a population decline. Survival to metamorphosis in salamanders has been found to decline with exposure to insecticide, an effect attributed to decreases in their zooplanktonic prey (Boone and James 2003). Additionally, body-mass at metamorphosis can be reduced when herbicides reduce algal food resources (Diana et al. 2000; Boone and James 2003). Mills and Semlitsch (2004) experimentally demonstrated that, for the insecticide carbaryl, indirect effects on the food web had greater influence on amphibians than did direct chemical effects; this appears to be the only study explicitly testing such a relationship. Contaminants may also affect the efficacy or the survival of predators upon amphibians. Insects are significant predators on many larval amphibians (Wilbur and Fauth 1990) and are an important food resource for terrestrial amphibians. Insecticides are intended to be lethal to insect pests, and invertebrates in general will be particularly sensitive to insecticide exposure, which could increase abundance of larval amphibians thereby increasing competition (Boone and Semlitsch 2003; Relyea et al. 2005) and potentially reduce growth andlor survival of terrestrial amphibians. Elimination or reduction of predation could result in high larval survival so that individuals experience high competition for food resources, which in turn may result in death via starvation and reduced size of metamorphs (e.g., the ameliorative effects of predation [Morin 19811). Low exposure to chemicals may result in increased metabolic expenditure that could reduce the energy available for growth and reproduction while having no observable effects on the food web. Contaminants may also suppress function of the immune system thereby making organisms more susceptible to other environmental factors, both anthropogenic and natural (Carey 1993; Carey and Bryant 1995; Carey et al. 1999; Hayes et al. 2006). Sudden die-offs in Australia, Central America, western United States, and parts of Europe have been attributed to a chytrid fungus (Berger et al. 1998; Bosch et al. 2001; Muths et al. 2003), which may be a novel pathogen in these environments. The presence of chemicals in the environment, however, may make organisms more likely to succumb to pathogens or diseases that were already present (Parris and Baud 2004; Parris and Cornelius 2004; Bridges-Britton et al., in review). Christin et al. (2004) found that exposure to mixtures of chemicals commonly found in agricultural settings had the potential to compromise immune system function, leaving individuals more susceptible to disease. Changes in individual immune responses or in metabolism may be markers of significance at the population level and could contribute to an understanding of how contaminants affect amphibians and influence sensitivity to other factors. IV. APPROACHES TO STUDYING DECLINES

Although linking pesticides to amphibian declines may be difficult, there are a number of reasons why amphibians may be susceptible to pesticides and other chemical contaminants. Many researchers have suggested that amphibians may be particularly vulnerable to contaminated environments because they (1) have permeable eggs, skin, and gills that may increase absorption of toxins into the body, (2) often occupy critical aquatic and terrestrial habitats where their chances of encountering contaminants are high, (3) live in ,environments stressful for larvae where the addition of a pesticide may further reduce juvenile recruitment, (4) are generally philopatric to natal sites and thus unlikely to avoid contaminated sites, and (5) can hibernate, which may subject them to pesticides during times when they cannot avoid exposure (Henry 2000 and references therein). Some research suggests, however, that amphibians are not particularly sensitive to chemical contamination (Bridges et al. 2002), at least not more so than species of vertebrates typically used in

3192

AMPHIBIAN BIOLOGY

toxicological testing (i.e., fish). The simple fact that chemical contaminants are widespread, however, means that one should try to understand what role they are playing in amphibian populations and communities. Recent research has focused on assessing the importance of multiple stressors in amphibian declines. Contaminants do not exist in nature in isolation from other environmental variables, and so it becomes necessary to construct more elaborate experimental designs to incorporate multiple factors. These may interact in a manner not predictable merely from an understanding of the effects of the individual factors. Environmental stressors can interact with one another in three ways. They can interact antagonistically, whereby the combined effect of the stressors is less than would be predicted from the sum of their individual effects. They can be additive, in which their collective effect is simply the sum of their individual effects. Finally they can be synergistic, whereby the effect is greater than would be predicted by simply adding the effects of the individual stressors. A multiple-stressor approach has been advocated in many studies that have examined single stressors, and current research suggests that synergistic interactions may contribute to declines (Carey et al. 2001; Linder et al. 2003 and references therein). Researchers have conducted studies in the laboratory, in mesocosm field studies and at the landscape level to assess contaminant effects. Each of these research tiers will help to evaluate the potential outcome that exposure to pesticides will have in the field in the presence and absence of other factors, and will help to evaluate the mechanisms driving the effects in the field. Integrating the information collected in the laboratory and from studies at the level of the mesocosm, field and landscape will help to determine contaminant classes that pose the greatest risk and to ascertain which populations will most likely be affected. Some of the important concepts that research in amphibian toxicology has established are highlighted below. A. Laboratory Studies Laboratory studies represent a first-tier effort in understanding the effects of environmental contaminants on amphibians. It is in the laboratory that controlled efforts can determine how a particular compound affects an organism. It is necessary to observe singular effects of contaminants if one is to predict responses in a natural habitat. Compounds that kill insect pests by disrupting cholinesterase, for instance, may also have similar effects on non-target vertebrates, such as amphibians. Research in the laboratory has helped evaluate: (1) how changes in behaviours can influence a contaminant's effect, (2) the relative sensitivity within amphibians, and compared to other taxa, and (3) how contaminant influences sensitivity to other factors. In the laboratory, traditional experimental studies in amphibian toxicology are usually aimed at understanding the physiological basis for the mode of action of a chemical and its effect on mortality, growth, development, and morphological deformities (e.g., LC6,,s or Frog Embyro Teratogenesis Assay - Xenopus; American Society for Testing and Materials 1991). Laboratory studies have become more sophisticated than the "kill-them-and-countthem" tests conducted in early days. Because contaminants do not generally occur at environmental concentrations great enough to elicit direct mortality, traditional LC5,, studies (determining the concentration of a compound required to kill 50% of a test population) are less useful today than studies examining the subtler effects of lower, sub-lethal concentrations. Examining behaviour, growth and development can provide a better understanding of the impact of low-level exposure of pesticides on amphibian life histories. Small changes in life-history characteristics can alter population dynamics (e.g., juvenile recruitment, fecundity) and cause declines in population numbers over time, making examination of sub-lethal endpoints particularly important. Behavioural responses to chemicals are most easily measured in the laboratory, and represent important endpoints. Toxic compounds can cause indirect mortality and

BOONE ET AL.: IMPACT OF PESTICIDES IN AMPHIBIAN DECLINES

3193

detrimental changes in life history by altering such behaviours as swimming, feeding, or predator avoidance (Little et al. 1990). The presence of an environmental contaminant, for instance, can decrease (Freda and Taylor 1992; Bridges 1997) or increase (Rohr et al. 2003) larval amphibian activity and alter swimming and feeding behaviour (Freda and Taylor 1992; Bridges 1997). Skelly (1994) demonstrated that diminished activity generally leads to decreased predation by visually oriented predators. Alternatively, it is possible for contaminants to increase activity (Rohr et al. 2003) and, therefore, predation rates (Cooke 1971). Behavioural responses such as these are difficult to observe and control in the field, but may explain the mechanisms through which pesticides affect populations. In the laboratory it is also possible to determine how amphibians respond to chemicals relative to other, more commonly used, test species such as rainbow trout, bluegill sunfish, or fathead minnows that are used by governmental agencies to set environmentally acceptable limits. For example, studies have suggested that allowable levels of copper in the environment, as determined by tests using a fish species, may not be protective of amphibians (Bridges et al. 2002). Copper is introduced into the environment through mining and industrial practices and is used as an algaecide. Application of this metal can thus potentially contaminate amphibian habitats, such as ponds, and be injurious to amphibian larvae. Conversely, amphibians are not as sensitive to some pesticides (e.g., permethrin, carbaryl) as are other taxa (Bridges et al. 2002). Laboratory studies are useful in that they allow researchers to understand how single pesticides affect species in the absence of other factors, so that differential sensitivity of species and life stages can be determined (Bridges 2000). This is important when considering how pesticides may influence the evolution of resistance to increasingly contaminated environments. Bridges and Semlitsch (2001) found significant differences in responses to carbaryl exposure among half-sibling families within a population of southern leopard frogs (Rana sphenocephala), indicating the presence of additive genetic variance (i.e., heritability) and likely, the ability to adapt to environmental stressors like carbaryl. Additionally, because responses to pesticides are partly genetic, selection for resistant individuals can occur. Semlitsch et al. (2000) found that treefrog (Hyla versicolor) tadpoles from clutches most sensitive to high levels of carbaryl (as determined in the laboratory) had lower survival under some field conditions (i.e., high density), indicating that resistance may come at the price of reduced competitiveness in natural, "clean" habitats. Furthermore, the timing of exposure influences the effect of a contaminant (Bridges 2000; Greulich and Pflugmacher 2003), which suggests that breeding phenology and management of pesticide application could be co-ordinated to minimize negative effects on amphibian populations. Laboratory studies indicate that the presence of sub-lethal contaminants may be more potent in the presence of other factors. De Solla et al. (2002) found that developing tadpoles in situ had reduced hatching success, lower survival, and more deformities than did individuals reared in the laboratory in water from the same site. This suggests that some factor or factors present in the natural environment influence the toxicity of the compounds present there. Another example involved the photo-enhanced toxicity of a chemical by ultraviolet-B radiation (Zaga et al. 1998; Chapter 6 in the present volume). The results of this two-factor design indicate that ultraviolet-B and the insecticide carbaryl each affected tadpole responses separately, but that in combination the toxicity of carbaryl increased tenfold above that when the tadpoles were subjected to either factor alone. Additionally, higher water temperature increased mortality of Rana clarnitans tadpoles at four of the seven highest concentrations of carbaryl (Boone and Bridges 1999), suggesting that natural stressors can influence the potency of contaminants. When tadpoles were reared with carbaryl and non-lethal (caged) predatory salamander larvae, they had higher mortality than when reared with predators or carbaryl alone (Relyea and Mills 2001). Observing such interactions in the laboratory provides a better understanding of responses in the field, and yields predictions for changes in natural populations. Another example of uncovering responses that are unpredictable from examining singular effects is a study of two natural Minnesotan ponds (one reference and one

3194

AMPHIBIAN BIOLOGY

chemically impacted), each tested in the presence and in the absence of ultraviolet radiation (Bridges et al. 2004). The incidence of embryonic deformities was high in the reference site when ultraviolet radiation was excluded, but low when ultraviolet radiation was present. This suggests the presence of a permanent, naturally occurring, deleterious compound that is broken down by ultraviolet radiation. In the impacted site, deformities were greatest when ultraviolet radiation was present, indicating the presence of a compound(s) potentiated by ultraviolet radiation. Ultraviolet radiation is present in nature and accordingly it can influence the toxicity of some contaminants. This highlights the importance of amalgamating laboratory and field studies to achieve a clear understanding of the impacts of contaminants in natural environments. B. Mesocosm Field Studies

Toxicological studies in the laboratory are critical for determining lethal ranges of contaminants for a large number of species and to understand mechanisms that might drive effects seen in nature. Field studies are often viewed as being more time-consuming, more difficult to assess, and more variable. While to some extent these views may be valid, field studies are necessary for determining how contaminants will affect populations and communities in more complex environments. Mesocosm studies are ideal for combining the benefits of both field and laboratory studies and have been used to examine the role of pesticides in amphibian declines (Rowe and Dunson 1994; Boone and James 2005). Boone et al. (2004) found that large-scale experimental ponds yield similar results to mesocosm studies conducted in cattle-tank ponds, suggesting that mesocosm studies are a powerful tool for evaluating field-level responses of populations and communities. From ecotoxicological studies conducted in mesocosms, it has been learned that (1) environmental conditions in ponds to which pesticides are added can influence the magnitude of the chemicals' effects, (2) indirect effects of other factors on the food web can dramatically alter the response to the contaminant, (3) laboratory studies are often not predictive of fieldlevel responses, and (4) expected environmental concentrations of pesticides could contribute to some amphibian declines. Mesocosm studies have contributed to an understanding of how contaminants may influence populations in complex natural environments, and also how diversity may be influenced by contaminants. When factors, such as larval density, predation, and ponddrying, that are known to be important to natural amphibian communities (Semlitsch et al. 1996) are manipulated with chemical contaminants, natural stressors can influence the magnitude of the chemical effect (reviewed by Boone and Bridges [2003]). Furthermore, the addition of multiple factors is hypothesized to contribute to amphibian declines and exacerbate the chemical effect, a result also supported by laboratory studies (see above). For instance, amphibians appear to be more susceptible to trematode infection in the presence of pesticides such as atrazine, malathion and esfenvalerate (Kiesecker 2002). Additionally, there is some evidence that indicates multiple stressors have greater effects than do single factors alone (e.g., Britson and Threlkeld 2000; Gendron et al. 2003; Hatch and Blaustein 2003; Boone et al. 2005; Boone et al., in press). For instance, Bridges-Britton et al. (in review) demonstrated that the effects of the insecticide carbaryl became more lethal when both ultraviolet radiation-B and a waterborne pathogen (Saprolegnia firax) were present than when just one or two factors were present in a laboratory environment. In a field study conducted simultaneously, they found that the presence of both the pathogen and expected environmental concentrations of carbaryl decreased survival for southern leopard frogs, suggesting that the accumulation of stress under more realistic conditions can have negative consequences for the population. Thus, there was a congruence of laboratory and field studies. Another important conclusion drawn from mesocosm studies is that contaminants can affect amphibian population and community dynamics by modifying the food web. Pesticides and other contaminants often have potent effects on larval amphibians' food base (algae or zooplankton), as well as on their predators, especially aquatic invertebrate ones such as

BOONE ET AL.: IMPACT OF PESTICIDES I N AMPHIBIAN DECLINES

3195

insects and crayfish. Most expected environmental concentrations of pesticides appear to be below a level needed to induce direct mortality, but these concentrations can increase or decrease food resources. Herbicides can reduce the amount of algal resources upon which anuran tadpoles feed (Diana et al. 2000; Boone and James 2003), while insecticides can reduce the abundance of (or eliminate) zooplankton on which carnivorous salamander larvae prey (Hanazato and Yasuno 1990; Boone and James 2003). In this way, one can predict survival, time of metamorphosis and size at metamorphosis by knowing what parts of the food web are most affected by a pesticide (Fig. 1). For instance, the insecticide carbaryl predictably results in trophic cascades. Carbaryl lowers the abundance of zooplankton and predatory invertebrates, which results in an algal bloom. As a result of carbaryl exposure, primary consumers increase (Boone and Semlitsch 2002) and secondary consumers decrease in abundance (Boone and James 2003). Leftcort et al. (1999) examined tadpole and snail communities exposed to heavy metals and predator cues and found that the metals shifted the outcome of competitive interactions and altered species abundance. An empirical measure of indirect and direct chemical effects was conducted by Mills and Semlitsch (2004) with the insecticide carbaryl, indicating that the indirect effect of carbaryl outweighed any direct effects through its effects on the food resources.

-

Herbicide---------------- -b

Algae

anurans

v-'.

.'

'-. .'

L.

Insecticide 0-

0-

.* I,*

-0-

KO-

Vertebrate predators

Invertebrate predators

Fig. 1. Diagrammatic model of the effect of herbicides and insecticides on aquatic communities containing amphibians. The model suggests that in general (1) herbicides that reduce the algal food base of a community will reduce the abundance of all species in the food web and have negative impacts on anuran and salamander survival, mass, and time to metamorphosis and that (2) insecticides that have sublethal effects on amphibians can negatively affect all invertebrates in the community and result in trophic cascades that can positively affect anurans (through increasing food resources) and negatively impact salamanders (through decreased food resources).

Another interesting pattern arising from mesocosm studies is that they suggest laboratory studies may not always accurately reflect field responses or predict population and community-level consequences of exposure to contaminants. This does not preclude laboratory studies usefully addressing other issues, such as understanding underlying mechanisms. Laboratory studies on contaminants can result in overestimating the effect found in the field, while missing other important effects. Relyea and Mills (2001) and Relyea (2003), for instance, found that tadpole mortality increases significantly for some anuran species in the presence of pesticides and predators combined over mortality caused by either factor alone. This suggests that in complex environments even low levels of contamination, if present in conjunction with predators (which is almost always the case), may lead to increased mortality. Field studies, however, suggest that these interactions may not always be significant in the field because other factors (like changes in food resource-base) may be more important (Boone and Semlitsch 2001; Boone and Semlitsch 2003; Mills and

3196

AMPHIBIAN BIOLOGY

Semlitsch 2004). Field studies including both predators and pesticides have not demonstrated synergistic interactions resulting in greater mortality. Additionally, laboratory studies have found that contaminants may have photo-enhanced toxicity with exposure to environmental levels of ultraviolet radiation (Zaga et al. 1998), which suggests that lethal concentrations may be much lower in nature than laboratory studies suggest. Field studies, however, have not shown any negative effects of exposure to sub-lethal levels of contaminants in the presence or absence of ultraviolet-B radiation (Bridges and Boone 2003), presumably because tannins in the water column can effectively filter out ultraviolet light and reduce the potential for phototoxicity. Mesocosm studies do suggest that pesticides could contribute to population declines in two ways. First, for communities exposed to any anthropogenic stressors (which are additional stresses), the likelihood of declines may increase; species at the edge of their ranges or those near their physiological limits (in terms of temperature, pH, or other factors) may be the most susceptible to contaminant stress. Evidence suggesting that amphibians are more susceptible to pathogens (Parris and Cornelius 2004; Bridges-Britton et al., in review), parasites (Gendron et al. 2003), ultraviolet radiation (Hatch and Blaustein 2003), and other contaminants (Boone et al. 2005; Hayes et al. 2006) in the presence of low-level contamination supports the notion that multiple sub-lethal stressors can reduce amphibian abundance over time. The mechanism whereby cumulative stressors affect organisms may be through increased metabolic costs for detoxification or through reduced immunological defense (Carey et al. 1999), although there has been very limited research addressing these questions. Secondly, changes in the food web may be an important contributor to some declines, especially for carnivorous amphibians. Invertebrates are sensitive to low levels of insecticides and reduction or elimination of them will have an impact on carnivorous amphibians such as salamander larvae. Boone and James (2003) found that salamander populations could virtually be eliminated from aquatic environments by exposure to carbaryl. Experimental mesocosm data do suggest that exposure to sub-lethal levels of contaminants could contribute to the problem of declining populations; linking patterns observed in such experiments to declines in the field would help determine the degree to which contaminant stress is important in the natural world. C. Natural Field Studies

Ultimately, only natural field studies can link pesticides with actual declines of amphibian populations and provide powerful associations between a conjectural cause and effect. In situ field studies provide a means of examining broad, landscape-level responses to a factor, and of ascertaining whether aerial, or other, transport of pesticides is important. Such studies should lead to the emergence of landscape-level patterns. Most field research has focused primarily on three aspects: (1) measuring the accumulation of pesticides in individual amphibians across the landscape, (2) physiological responses to exposure, and (3) landscape-level patterns of declines. So to date, what do natural field studies reveal about the possible role of pesticides in enigmatic declines - that is, declines that cannot be explained by simple habitat destruction? The most basic tenet that field studies have revealed is that declines have occurred in "pristine" areas. There are, therefore, population declines that cannot be explained either by gross alteration of habitat or by introduced predators. This finding indicates that some factor or factors, such as disease, ultraviolet-B radiation, or contaminants must be playing a role in enigmatic declines. This is an important starting point. If one could explain declines on the basis of habitat destruction or exotics there would be no need to investigate other factors that are much more difficult to study. Worldwide, amphibian declines appear to be most severe in montane areas, which generally are not subject to runoff from agricultural land. While these locations provide protection from the high concentrations of pesticides found in agricultural runoff waters, aerial transport can bring pesticides to even the most remote areas. For example, atmospheric transport carried the pesticide toxaphene from southern United States to the Great Lakes region (Ryan and Hites 2002) and even to the Arctic (Derek et al. 1990).

BOONE ET AL.: IMPACT OF PESTICIDES IN AMPHIBIAN DECLINES

3197

Although long-range pesticide transport is possible, transport over medium ranges may be more relevant because it can result in deposition of higher concentrations of pesticides. This is especially true for pesticides that break down rapidly and do not bioaccumulate. In California, pesticide transport from the Central Valley into the Sierra Nevada at distances up to 150 km is well documented (Zabik and Seiber 1993; Aston and Seiber 1997; McConnell et al. 1998; LeNoir et al. 1999). In fact, Bridges and Little (2005) report that tadpoles took longer to reach metamorphosis when exposed to extracts from air and water taken from an alpine site in Sequoia Kings Canyon Nationa Park in the Sierra Nevada Mountain range. Three of the regions of the world with the best studied amphibian population declines, the Sierra Nevada of California, Western Australia, and the mountains of Central America, are all downwind from extensive agricultural areas. Lips and Donnelly (2005) pointed out that in Australia and Central America lowland populations that are much closer to agricultural regions (and are likely, therefore, to have greater exposure to pesticides) have not declined as much as have populations at higher elevations. This pattern suggests that if pesticides are contributing to declines, it is not through direct effects, but rather in synergism with some other factor (e.g., cold or disease) that is not present or active in the lowlands. A similar pattern exists in the United States. In the southern and southeastern regions there is extensive use of pesticides, but few amphibian declines, while in California there is heavy pesticide use and widespread amphibian population declines. Like the elevational pattern of declines in Australia and Central America, the pattern in the United States again suggests that pesticide exposure alone is not the sole cause of declines. While it is clear that pesticides are transported into remote regions where declines have occurred, little is known about the resulting geographic and temporal distribution of pesticide residues impinging on amphibians. The temporal distribution is especially difficult to quantify because pesticide applications are extremely "spiky." In California, for example, pesticide applications during a few weeks of the year typically account for a substantial share of total use (Fig. 2). This means that field sampling of pesticides in water is likely to miss peak concentrations and, therefore, will underestimate short-term exposure levels faced by animals. In order to conduct experiments with ecologically relevant doses, one needs to know the levels to which animals are exposed in the field. California has received the most extensive studies of pesticide transport to montane areas in which there are amphibian declines. No studies involving transport and deposition of pesticides into montane areas seem to have been carried out for Central America or Australia. Even in California, less than a dozen montane sites have been examined for residues relevant to amphibian declines. Furthermore, transport of only a few different chemicals, mainly endosulfan, chlorpyrifos, malathion, and diazinon, have received attention (Zabik and Seiber 1993; Aston and Seiber 1997; McConnell et al. 1998; LeNoir et al. 1999). Currently, however, many hundreds of different chemicals are in use. During 2000 in California alone, 781 different pesticides were used (Department of Pesticide Regulation 200 1). A growing number of studies in California are bypassing issues of transport and deposition; instead, researchers are seeking and finding pesticides directly in the bodies of frogs. As early as 1970, DDT residues were found in mountain yellow-legged frogs (Rana muscosa) throughout the Sierra Nevada (Cory et al. 1970). It was almost thirty years before the next study was completed when Datta et al. (1998) examined Pacific treefrogs (Hyla regdla) for DDE, chlorothalonil, and chlorpyrifos at two Sierra Nevadan sites. Sparling et al. (2001) also examined Pacific treefrogs and found higher pesticide levels in the Sierra Xevada, where a number of amphibian species have declined, than at two coastal sites that have experienced few declines. Pesticide residues in mountain yellow-legged frogs were compared between an experimental reintroduction site in the Californian Sierra Nevada where survival was very low and a site with healthy frog populations (Fellers et al. 2004). Of the six chemicals studied, five were found at higher levels in frogs from the reintroduction site (three significantly so) than in those from the comparison site. Finally, Angermann d al. (2002) found that although the pesticide toxaphene had been banned from use for

3198

AMPHIBIAN BIOLOGY

Week

Fig. 2. Timing of chlorpyrifos pesticide applications upwind of historic sites for Cascades frogs (Rana cascadae) in northern California in 1998. Applications are kilograms of active ingredients. The locations of sites are from Jennings and Hayes (1994). Pesticide data are for 1998 (California Department of Pesticide Regulation; annual report on pesticide use [1999]). The methodology for calculating upwind use is described by Davidson (2004).

almost 20 years, it was still widespread in Pacific treefrogs from 21 sites across the Sierra Nevada. It is clear that pesticides are widespread in the bodies of frogs even in rather remote regions. At this time, however, the biological significance of the levels of pesticides that have been found is still unknown. The situation is similar to that in humans; pesticide residues appear to be widespread in the bodies of people in the United States (CDC 2003) but the risks to health of the measured levels of residues are unknown. Understanding the biological effects of field-relevant doses of pesticides is a key challenge for future laboratory experiments and field observational studies. To date, there are only a few studies directly indicating that exposure to sub-lethal levels in the field has biological effects. Sparling et al. (2001) found that pesticide residues were higher and cholinesterase levels lower in Pacific treefrogs in the Californian Sierra Nevada than in treefrogs on the coast. The low cholinesterase levels in Sierra Nevadan frogs may have resulted from exposure to cholinesterase-inhibiting pesticides (most organophosphate and carbamate pesticides). Without further research, however, one cannot rule out alternative explanations for suppressed cholinesterase levels, such as cold or altitude stress. Two studies have combined laboratory experiments with field observations or field experiments to demonstrate that exposures to pesticides in the field can suppress aspects of the anuran immune system. Gilbertson et al. (2003) found that leopard frogs (Rana pipiens) collected in Ontario in areas exposed to pesticides had lower antibody responses and cellular immune responses than did frogs from areas with less exposure to pesticides. In the laboratory, they found that sub-lethal levels of DDT, dieldrin, and malathion all suppressed the activity of the immune system of leopard frogs. Kiesecker (2002) studied the effects of exposure to

BOONE ET AL.: IMPACT OF PESTICIDES IN AMPHIBIAN DECLINES

3199

pesticides on wood frogs' (Rana sylvatica) susceptibility to trematode infection and found that frogs in sites exposed to agricultural runoff had dramatically greater trematode-induced deformities of the limbs than did frogs not exposed to pesticides (26% versus 4%). Companion laboratory experiments found that exposure to atrazine, malathion, and esfenvalerate all suppressed frogs' immune response and increased the success rate of trematodes encysting in frogs. Some of the strongest evidence for a link between pesticides and actual amphibian population declines comes from studies by Davidson et al. (2001, 2002). They examined landscape-scale patterns of decline at almost 1 500 historic locations across California for eight amphibian species. For each site, they calculated the amount of upwind agricultural land-use based on predominate wind patterns, as a rough proxy for sites' exposure to pesticides. For four species of ranid frogs (Ram aurora draytonii, R. boylii, R. cascadae and R. muscosa), population declines were strongly associated with the amount of agricultural land use upwind from a site, suggesting that wind-borne pesticides may be an important factor in declines. In multivariate regression models, the association between declines and upwind agricultural land use was strong, even when other factors such as latitude, elevation, precipitation and local land-use were taken into account. It was striking that the same association held for four species with different ranges. The studies by Davidson et al. (2001, 2002) did not contain actual data on pesticide use, and instead relied upon the amount of upwind agricultural land use as a proxy for actual pesticide use. A follow-up study, however, did find a strong association between declines for five species and upwind pesticide use, based on California state records of actual pesticide applications from 1974 to 1991 (Davidson 2004). In multivariate regression models upwind pesticide use was significant and a strong predictor of site-status for Rana aurora draytonii, R. boylii, R. cascadae, and R. muscosa. Furthermore, for these four species and the Yosemite toad (Bufo canorus) taken together, upwind use of cholinesterase-inhibiting pesticides was more strongly associated with declines than were total pesticides or any of 64 classes of pesticides (Davidson 2004). Although the work by Davidson et al. (2001, 2002) and Davidson (2004) provided some of the best links between actual amphibian population declines and pesticide use, it has several limitations. First, although the association between population declines and upwind pesticide-use holds even when a number of covariates are taken into account, the pattern has not been tested in studies that also consider exotic predators or disease. Second, the strength of association between upwind pesticide-use, predominant wind patterns, and actual exposure levels are currently unknown. Ideally, one would want to know the actual exposures (e.g., concentration of residues in water or in frogs' bodies) for a large number of sites, but this information is not available. Clearly, a key area for future fieldwork is to measure exposure at a sufficiently large number of sites to be able to evaluate the relationship between the geographic pattern of exposure and patterns of species' decline. Due to the large temporal and spatial scale of amphibian declines, they can never be directly subjected to experimentation. Therefore, the role of pesticides in amphibian population declines must be studied by a combination of large-scale observational studies, and manipulative laboratory and field experiments. If pesticides are affecting amphibian populations, it must be through a web of processes and relationships including the geographic patterns of agricultural land use, the pattern of pesticide use, and site-specific interactions with other factors. All of these processes ultimately contribute to large-scale patterns of amphibian population decline. The causal relationships generate patterns that can be called the "pesticide puzzle" of how pesticides may be affecting amphibian populations (Fig. 3). The various pieces in the puzzle (causal relationships and patterns) provide opportunities for many different types of studies, all of which contribute to understanding and evaluating the links between pesticides and amphibian declines. Observational studies can examine many different parts of the pesticide puzzle. For example, Davidson et al. (2001, 2002) examined the association between the geography of

AMPHIBIAN BIOLOGY

3200

Geography of Agricultural

Geographic, Temporal and Chemical Pattern Of Pesticide Use

Wind patterns, Pesticide transport Geography of Pesticide Residues

+

Experim~ntal

I

Zone

Conditions

A

imal expc

Different life stages

+

Site population Response

i Geographic, temporal, and species patterns of Fig. 3. Pieces in the pesticide puzzle: Diagram showing ways pesticides contribute to amphibian declines. All the relationships between the pieces in the puzzle present opportunities for research on the role of pesticides in declines. The gray box labelled "Experimental Zone" indicates relationships that are subject to experimental study. Only half the interacting stressors are included in the experimental zone to indicate that laboratory and mesocosm studies can never be sure of including all the important interactions in natural systems.

agricultural land use, wind patterns, other population stressors, and geographic patterns of decline. Ultimately, observational studies need to be coupled with experiments that can explore mechanisms at the level of individual animals. A variety of laboratory and mesocosm experiments is possible and needs to ascertain the effects of ecologically relevant doses of pesticides on different endpoints. New information about any one piece of the puzzle often opens new opportunities to test presumed relationships (e.g., Davidson 2004). As more information on the actual geographic pattern of pesticide residues becomes available, it will be possible to test the association between pesticide uses, predominate winds, and residues that are assumed in the Davidson study. Similarly, information on pesticide residues would allow a direct test of the relationship between the geographic patterns of residues and the geographic patterns of amphibian population declines.

V. ASSESSING CAUSALITY For complex large-scale phenomena that cannot be experimentally manipulated, causality is not proven by a single study but rather inferred from the weight of evidence from multiple studies, both observational and experimental. Epidemiologists have long struggled with how to infer causality from observational studies and from multiple strands of evidence (Susser 1986; Fox 1991). Furthermore, epidemiologists regularly deal with the problem of assessing the causes of events with multiple factors, operating on both large temporal and spatial scales. To assess causality, epidemiologists have developed multiple criteria. These same techniques may be usehl in evaluating the causes of amphibian declines. Seven criteria are commonly used in assessing causality. The simplest criterion is time order. Does a supposed cause precede the effect? This criterion is useful for rejecting hypothesized causes that occur after the effect. Because there is an infinite number of events that can precede an effect, meeting this criterion cannot confer support for a hypothesized cause. In some cases where a supposed cause precedes an event by a long time, time order

BOONE ET AL.: IMPACT O F PESTICIDES IN AMPHIBIAN DECLINES

320 1

can argue against the cause, unless there is a plausible mechanism for the time-lag. This may be the case for organochlorine pesticides that have been in heavy use since the 1950s (Aspelin 2003). Their use long preceded amphibian declines and, therefore, may not explain recent declines. A number of authors (Stebbins and Cohen 1995; Sparling et al. 2001; Davidson 2004) have suggested that organophosphate and carbamate pesticides may be contributing to declines, in part because the pesticides came into widespread use in the late 1960s and amphibian population declines, at least in western United States, are believed to have started in the early 19'70s. A second criterion for assessing causality is biological plausibility. Is the hypothesis consistent with known biological processes and other scientific facts? For the pesticide hypothesis, probably the most difficult question in this regard is how extremely low levels of pesticides can either cause or contribute to mass mortalities. Work by Hayes (e.g., Hayes et al. 2002) indicates that levels as low as parts per billion may have dramatic effects on amphibian development. Currently the most plausible mechanism whereby low-level exposures could lead to dramatic population declines is that exposures to pesticides impair immune function, leading to mass die-offs from disease or pathogens present in the environment.

Aprobability critem'on is used to determine if the observed associations between a supposed cause and effect are statistically significant. With the strength of association criterion one evaluates how strongly associated a supposed cause is with its effect. Strength of association means looking beyond simple statistical significance, and examining the magnitude of effect or its relative risk compared to other explanatory factors. Because causality is ultimately assessed across multiple studies with widely different approaches, the consistency upon replication criterion appraises whether evidence for a supposed cause is consistent over multiple studies representing different approaches, locations, or species.

The dose-response relationship criterion evaluates whether the incidence of effect increases monotonically with an increase in the dose of a supposed cause. A finding of dose-response relationship is strongly supportive of a causal relationship. Not all causal relationships, however, have a monotonic dose-response relationship; therefore, lack of a dose-response relationship is not strong evidence against causality. Comparisons of an effect at two levels of a supposed cause (e.g., population declines at sites with low versus high pesticide exposures) are less convincing of causality than are comparisons across multiple levels of the supposed cause, in which the effect increases with increased levels of the cause. The final criterion is speczjicity. Does only the supposed cause lead to the effect or are there other factors that also lead to that effect? Unfortunately, in amphibians there are multiple causes of decline, so none of the hypothesized causes for it score high when using these criteria. Ultimately, only by considering multiple studies and applying criteria for causality, such as those described above, will it be possible to begin assessing the role of pesticides in amphibian declines. In this respect, the pesticide hypothesis is no different from other hypothesis for amphibian declines. The criteria described above do not lead to a simple or clear-cut determination of causality. Many of the criteria involve difficult subjective judgments, and there are no rules for how to weight the different criteria (Fox 1991). Nonetheless, these criteria provide a start in organizing a "weight-of-the-evidence"evaluation of an hypothesized cause of an event. These criteria could aid in making conscious and explicit evaluations of the weight of evidence and lead to better assessment of causality of complex phenomenon such as amphibian population declines. VI. CONCLUSIONS

Following the news of worldwide amphibian declines, researchers in amphibian ecotoxicology expected to find links between contaminants and declines. The physiological properties and life history characteristics of amphibians would seemingly make them sensitive to environmental contamination and, accordingly, good bioindicators of the quality of aquatic and terrestrial habitats. After a decade or more of research, however, amphibians do not

3202

AMPHIBIAN BIOLOGY

appear to be any more sensitive to contaminants than are other vertebrate species. It is, therefore, unlikely that pesticides alone will explain worldwide amphibian declines. Studies examining multiple stressors are the most promising avenue of research, and may offer insight into how pesticides influence species diversity in the natural world. It seems logical that the more stressors, natural or a~lthropogenic,to which an organism is exposed, the greater will be the probability that it will not be able to reach the critical bar allowing completion of metamorphosis (Boone et al., in press), survival over winter or successfbl reproduction. Are there characteristics that may make certain amphibian populations more sensitive to the presence of a contaminant (or other) stressor? Among the 27 North American species that Semlitsch (2003) listed as either endangered, threatened, or proposed endangered1 threatened, caudates and anurans were equally represented, although reports of declines in North America are most strongly associated with declines of ranids in the west. Given that there are approximately 1.7 times more North American caudates, one would expect that there should be more caudates experiencing declines all things being equal, which suggests anurans in North America may be more susceptible. Among North American anurans, Crump (2003) reported that ranids are experiencing declines at greater than the expected rates for any of the anuran genera; among tropical anurans, bufonids appear to be experiencing more than expected declines. Comparisons of characteristics of declining amphibians worldwide (as in Crump [2003] for anurans in the tropics) may help anticipate which characteristics make amphibians most susceptible to the probability of experiencing declines, and may also help implicate the cause. For instance, Crump (2003) found that species have a higher probability of declining if they have restricted geographic distributions, occupy running water as eggs and lame, or produce small clutchs. Data such as these suggest that species that are geographically isolated or with few populations should be closely monitored, with particular attention to species with certain life history characteristics (like small clutch size), and also suggests that the problem could be associated with running water which then provides a direction for experimental research and a tangible basis for analysis (e.g., of contaminants and/or pathogens). Conversely, comparisons between areas where declines have occurred (e.g., western United States) and where they have not (e.g., eastern United States), may also help understand the factors at play in declines or which buffer populations from negative effects. In multi-species studies involving a pesticide, species with relatively short larval periods appeared to be most strongly affected by chemical exposure and/or resulting changes in the food web. In species such as toads and treefrogs, however, many of the effects were apparently "positive" due to changes in the food web (Boone and Semlitsch 2001, 2002; Boone et al. 2004). Species with short life spans may also be more likely to experience declines if contaminant exposure (in combination with other factors) reduces juvenile recruitment into the population, especially if metapopulation dynamics are disrupted by habitat destruction. Furthermore, populations at the edge of their range may be more likely be approaching their physiological limits; the addition of a stressor may increase the probability of extinction. It is difficult to say which pesticides pose the greatest risk to any organism, particularly amphibians, given that they are not routinely used in standard toxicological testing. The United States Environmental Protection Agency (USEPA), however, does collect information on the amount of registered pesticides that are sold in the United States and World markets. From this information, it is known that throughout the world, herbicides are the most commonly applied pesticide, followed by insecticides, and fungicides (Kiely et al. 2004). So, based on sheer volume of pesticides applied, herbicides could pose the greatest threat. Herbicides typically have modes of action that are unlikely to affect animals directly, although the herbicide atrazine has been found to disrupt the endocrine system (Hayes et al. 2002, 2003), the carrier of the herbicide glyphosate has been found to have toxic effects on amphibians (Howe et al. 2004), and herbicides can reduce the food base of the community (Diana et al. 2000; Boone and James 2003). Herbicides may, therefore, still have important negative effects on amphibian populations. Insecticides have modes of action that affect amphibians

BOONE ET AL.: IMPACT OF PESTICIDES IN AMPHIBIAN DECLINES

3203

and their food web; so while a lesser amount of active ingredient is applied, the effect could be more (or equally) detrimental. Evaluating the effects of commonly used pesticides that have widespread use may be one way to begin evaluating the effects of pesticides on amphibian populations. The big question regarding amphibian declines is why they seemingly began at roughly the same time across the world. Such a pattern does suggest a global phenomenon and a cause that has global distribution. Pesticides and environmental contamination could have served as a trigger, especially if they do make amphibians more susceptible to stressors already present in the environment (e.g., pathogens or disease). Evaluation of critical interactions in experimental manipulations and examining other factors that may operate in the same manner as pesticides might lead to better understanding of the causes of decline and enable formulation of better management policies for protection of amphibian populations. VII. ACKNOWLEDGEMENTS

David Bradford introduced us to the relevant epidemiological literature and has long advocated using epidemiological criteria to evaluate causes of amphibian decline. VIII. REFERENCES Angermann, J. E., Fellers, G. M. and Matsumura, F., 2002. Polychlorinated biphenyls and toxaphene in Pacific tree frog tadpol& (HYla regilla) from the California Sierra Nevada, USA. Enuiron. Toxicol. Chem. 21: 2209-2215. Appel, H. M. and Martin, M. M., 1992. Significance of metabolic load in the evolution of host specificity of Manduca sexta. Ecolow 73: 216-228. -2

Aspelin, A. L., 2003. Pesticide usage in the United States: Trends during the 20th century. Center for Integrated Pest Management, Nth. Caroline State Uni., Raleigh, NC. CIPM Tech. Bull. 105. Aston, L. S. and Seiber, J. N., 1997. Fate of summertime airborne organophosphate pesticide residues in the Sierra ~ e v a d a~ o u n t a i n s :J. Enuiron. Qual. 26: 1483-1492.

Berven, K. A,, 1990. Factors affecting population fluctuations in larval and adult stages of the wood frog (Rana syluatica). Ecology 71: 1559-1 608. Beyer, W. N., Pattee, 0. H., Sileo, L., Hoffman, D. J and Mulhern, B. M., 1985. Metal contamination in wildlife living near two zinc smelters. Enuiron. Pollut. Series A Ecol. Biolog. 38: 63-86. Bishop, C. A., Mahony, N. A., Struger, J., Ng, I? and Pettit, K. E., 1999. Anuran development, density, and diversity in relation to agricultural activity in the Holland River watershed, Ontario, Canada (1990-1992). Enuiron. Monitor. Assess. 57: 2 1-43. Bonin, J., DesGranges, J-L., Rodrigue, J. and Ouellet, M., 1987. Anuran species richness in agricultural landscapes of Quebec: for seeing long-term results of road call sunreys. Herpetol. Conseru. 1: 141-149.

Bard, S. M., 1999. Global transport of anthropogenic contaminants and the consequences for the &tic marine ecosystem. Mar Pollut. Bull. 38: 356-379. Beebee, T J. C., 2001. The role of population genetics in conservation biology. Rivista di Idmbiolop 40: 231-238.

Boone, M. D., 2005. Juvenile frogs compensate for small metamorph size with terrestrial growth: overcoming the effects of larval density and insecticide exposure. J. Herpetol. 39: 416-423.

Beja, P. and Alcarzar, R., 2003. Conservation of Mediterranean temporary ponds under agricultural intensification: an evaluation using amphibians. Biol. Cons. 114: 317-326.

Boone, M. D. and Bridges, C. M., 1999. The effect of temperature on the potency of carbaryl for survival of tadpoles of the green frog, Rana clamitans. Enuiron. Toxicol. Chem. 18: 1482-1484.

Berger, L., Speare, R., Daszak, P., Green, D. E. A., Cunningham, A., Goggin, C. L., Slocombe, R., Ragan, M. A., Hyatt, A. D., McDonald, K. R., Hines, H. B., Lips, K. R., MaRantelli, G. and Parkes, H., 1998. Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proc. Nut. Acad. Sci. 95: 9031-9036.

Boone, M, D. and Bridges, C. M., 2003. A hierarchical approach in studying the effects of an insecticide on amphibians. Chapter 8D, Pp. 165-170 in "Multiple Stressors and Declining Amphibian Populations: Evaluating Cause and Effect", ed by G. Linder, S. K. &st and D. W. Sparling. SETAC Press, Pensacola.

Bernard, L. and Lagadic, L., 1993. Sublethal effects of dietary cyfluthrin on nutritional performance and gut hydrolase activity in larvae of the Egyptian cotton leafworm, Spodoptera littoralis. Pesticide Biochem. Physiol. 46: 171-180. Berrill, M., Bertram, S., Wilson, A., Louis, S., Brigham, D. and Stromberg, C., 1993. Lethal and sublethal impacts of pyrethroid insecticides on amphibian embryos and tadpoles. Enuiron. Toxicol. Chem. 12: 525-539.

Boone, M. D. and Bridges, C. M., 2003. Impacts of chemical contaminants on amphibian populations. Pp. 152-167 in "Amphibian Conservation", ed by R. D. Semlitsch. Smithsonian Press, Washington, DC. Boone, M. D. and James, S. M., 2003. Interactions of an insecticide, herbicide, and natural stressors in amphibian community mesocosms. Ecol. Applic. 13: 829-841. Boone, M. D. and James, S. M., 2003. Use of aquatic and terrestrial mesocosms in ecotoxicology. App. Herpetol. 2: 23 1-257.

3204

AMPHIBIAN BIOLOGY

Boone, M. D. and Semlitsch, R. D., 2001. Interactions of an insecticide with larval density and predation in experimental amphibian communities. Cons. Biol. 15: 228-238. Boone, M. D. and Semlitsch, R. D., 2002. Interactions of an insecticide with competition and pond drying in amphibian communities. Ecol. Applic. 12: 307-316. Boone, M. D. and Semlitsch, R. D., 2003. Interactions of bullfrog tadpole predators and an insecticide: predation release and facilitation. Oecologia 442: 610-616.

Bridges-Britton, C. M., Boone, M. D. and Hopkins, W. A., in review. The interactive effects of UV radiation, a pesticide, and a fungus (Saprolegnia ferax) on survival, growth, and development of larval amphibians. Oecologia. Britson, C. A. and Threlkeld, S. T, 2000. Interactive effects of anthropogenic, environmental, and biotic stressors on multiple endpoints in Hyla chrysoscelis. J. Iowa Acad. Sci. 107: 61-66. Burnett, S., 1997. Colonizing cane toads cause population declines in native predators: Reliable anecdotal information and management implications. Pac. Cons. Biol. 3: 65-72.

Boone, M. D., Bridges, C. M., Fairchild, J. F. and Little, E. E., in press. Effects of ammonium nitrate fertilizer and insecticide exposure on populations of the green frog, Rana clamitans. Environ. Toxicol. Chem.

Carey, C., 1993. Hypothesis concerning the causes of the disappearance of boreal toads from the mountains of Colorado. Cons. Biol. 7: 355-362.

Boone, M. D., Semlitsch, R. D., Fairchild, J. F. and Rothermel, B. B., 2004. Effects of an insecticide on amphibians in large-scale experimental ponds. Ecol. Applic. 14: 685-691.

Carey, C. and Bryant, C. J., 1995. Possible interrelations among environmental toxicants, amphibian development, and decline of amphibian populations. Envirrm. Health Perspec. 103: 13-17.

Boone, M. D., Semlitsch, R. D., Little, E. E. and Doyle, M, C., in press. Multiple stressors in amphibian communities: Interactive effects of chemical contamination, bullfrog tadpoles, and bluegill sunfish. Ecol. Applic.

Carey, C., Cohen, N. and Rollins-Smith, L., 1999. Amphibian declines: An immunological perspective. Develop. Compar: Immunol. 23: 459-472.

Bosch, J., Martinez-Solano, I. and Garcia-Paris, M., 2001. Evidence of a chytrid fungus infection involved in the decline of the common midwife toad (Alytes obstetricans) in protected areas of central Spain. Biol. Cons. 97: 331-337. Bridges, C. M., 1997. Tadpole swimming performance and activity affected by acute exposure to sublethal levels of carbaryl. Environ. Toxicol. C h m . 16: 1935-1939. Bridges, C. M., 1999. Effects of a pesticide on tadpole activity and predator avoidance behavior. J. Herpetol. 33: 303-306.

Carey, C., Heyer, W. R., Wilkinson, J., Alford, R. A., Arntzen, J. W., Halliday, T., Hungerford, L., Lips, K. R., Middleton, E. M., Orchard, S. A. and Rand, A. S., 200 1. Amphibian declines and environmental change: Use of remote-sensing data to identify environmental correlates. Cons. Biol. 15: 903-913. Carson, R., 1962. "Silent Spring". Riverside Press, Cambridge. CDC, 2003. Second national report on human exposure to environmental chemicals. National Center for Environmental Health, Centers for Disease Control and Prevention, Atlanta. NCEH Pub. No. 02-0716.

Bridges, C. M., 2000. Long-term effects of pesticide exposure at various life stages of the southern leopard frog (Rana sphenocephala). Arch. Environ. Contamin. Toxicol. 39: 91-96.

Christin, M. S., Menard, L., Gendron, A. D., Ruby, S., Cyr, D., Marcogliese, D. J., Rollins-Smith, L. and Fournier, M., 2004. Effects of agricultural pesticides on the immune system of Xenopus laeuis and Rana pipiens. Aquatic Toxicol. 67: 3 3 4 3 .

Bridges, C. M. and Boone, M. D., 1999. Effects of a pesticide on tadpole survival, growth, and development. Biol. Cons. 113: 49-54.

Colburn, T, Dumanoski, D. and Myers, J. E!, 1996. "Our Stolen Future". Dutton, New York.

Bridges, C. M. and Little, E. E., 2005. Toxicity to amphibians of environmental extracts from natural waters in National Parks and Fish and Widlife Regfuges. Alytes 22: 130-145. Bridges, C. M. and Semlitsch, R. D., 2000. Variation in pesticide tolerance of tadpoles among and within species of Ranidae and patterns of amphibian decline. Cons. Biol. 14: 1490-1499. Bridges, C. M. and Semlitsch, R. D., 2001. Genetic variation in insecticide tolerance in a population of southern leopard frogs (Rana sphenocephala): Implications for amphibian conservation. Copeia 2001: 7-13. Bridges, C. M., Dwyer, F. J., Hardesty, D. K. and Whites, D. W., 2002. Comparative contaminant toxicity: Are amphibian larvae more sensitive than fish? Bull. Environ. Contamin. Toxicol. 69: 562-569. Bridges, C. M., Little, E. E., Gardiner, D. M., Petty, J. D. and Huckins, J. N., 2004. Assessing the toxicity and teratogenicity of pond water in Minnesota to native amphibian larvae. Environ. Sci. Pollut. Res. 11: 233-239.

Cooke, A. S., 1971. Selective predation by newts on frog tadpoles treated with DDT Nature 229: 275-276. Corn, P S., 1994. What we know and don't know about amphibian declines in the west. Pp. 59-67 i n "Sustainable Ecological Systems: Implementing an Ecological Approach to Land Management", ed by W. W. Covington and L. F. Debano. U.S. Forest Service. General technical report RM-247. Fort Collins, Colorado, USA. Cory, L., Fjerd, P and Serat, W., 1970. Distribution patterns of DDT residues in the Sierra Nevada mountains. Pesticide Monitor J. 3: 204-2 11. Cowman, D. F. and Mazanti, L. E., 2000. Ecotoxicology of "new generation" pesticides to amphibians. Pp. 233-268 i n "Ecotoxicology of Amphibians and Reptiles", ed by D. W. Sparling, G. Linder and C. A. Bishop. Society of Environmental Toxicology and Chemistry, Pensacola, FL, USA. Crump, M. L., 2003. Conservation of amphibians in the new world tropics. Pp. 53-69 i n "Amphibian Conservation", ed by R. D. Semlitsch. Smithsonian Press, Washington, DC.

BOONE E T AL.: IMPACT OF PESTICIDES IN AMPHIBIAN DECLINES Datta, S., Hansen, L., McConnell, L., Baker, J., LeNoir J. and Seiber, J. N., 1998. Pesticides and PCB contaminants in fish and tadpoles from the Kaweah River Basin, California. Bull. Enuiron. Contamin. Toxicol. 60: 829-836. Davidson, C., 2004. Declining downwind: Amphibian population declines in California and historic pesticide use. Ecol. Applic. 14: 1892-1902. Davidson, C., Shaffer, H. B. and Jennings, M. R., 2001. Declines of the California red-legged frog: climate, UV-B, habitat and pesticides hypotheses. Ecol. Applic. 11: 464-479. Davidson, C., Shaffer, H. B. and Jennings, M., 2002. Spatial tests of the pesticide drift, habitat destruction, UV-B and climate change hypotheses for California amphibian declines. Cons. Biol. 16: 1588-1601. Department of Pesticide Regulation, 1999. "Pesticide Use Report: 1998". Department of Pesticide Regulation, Sacramento. Department of Pesticide Regulation, 2001. "Pesticide Use Report: 2000". Department of Pesticide Regulation, Sacramento. De Solla, S. R., Pettit, K. E., Bishop, C. A., Cheng, K. M. and Elliott, J . E., 2002. Effects of agricultural runoff on native amphibians in the lower Fraser River Valley, British Columbia, Canada. Enuiron. Toxicol. Chem. 21: 353-360. Derek, C. G., Muir, N. F!, Ford, C. A., Reiger, A. W., Hendzel, M. R. and Lockhart, W. L., 1990. Evidence for long-range transport of toxaphene to remote arctic and subarctic waters from monitoring of fish tissues. Pp. 329-346 in "Long Range Transport of Pesticides", ed by D. A. Kurtz. Lewis Publishers, Chelsea. Diana, S. G., Resetarits, W. J., Schaeffer, D. J., Beckmen, K. B. and Beasley, V R., 2000. Effects of atrazine on amphibian growth and survival in artificial aquatic communities. Enuiron. Toxicol. Chem. 19: 2961-2967. Eroschenko, V F,! Amstislavsky, S. Y., Schwabel, H. and Ingermann, R. L., 2002. Altered behaviors in male mice, male quail, and salamander larvae following early exposures to the estrogenic pesticide methoxychlor. Neurotoxicol. Teratol. 24: 29-36. FeUers, G. M., McConnell, L. L., Pratt, D. and Datta, S., 2004. Pesticides in mountain yellow-legged frogs (Rana muscosa) from the Sierra Nevada Mountains of California, USA Enuiron. Toxicol. Chem. 23: 2170-2177. Fox, G. A,, 1991. Practical causal inference for ecoepidemiologists.J. Toxicol. Enuiron. Health 33: 359-373. Freda, J. and Taylor, D. H., 1992. Behavioral response of amphibian larvae to acidic water. J. Herpetol. 26: 429-433. Gendron, A. D., Marcogliese, D. J., Barbeau, S., Christin, M-S., Brousseau, P, Ruby, S., Cyr, D. and M. Eburnier, 2003. Exposure to leopard frogs to a pesticide mixture affects life history characteristics of the lungworm Rhabdias ranae. Oecologia 135: 469-476. Gilbertson, M., Haffner, G. D., Drouillard, K. G., Albert, A. and Dixon, B., 2003. Immunosuppression in the northern leopard frog (Rana pipiens) induced by pesticide exposure. Enuiron. Toxicol. Chem. 22: 101-110.

3205

Goater, C. F!, 1994. Growth and survival of postmetamorphic toads: Interactions among larval history, density, and parasitism. Ecology 75: 2264-2274. Goulet, B. N. and Hontela, A , 2003. Toxicity of cadmium, endosulfan, and atrazine in adrenal steroidogenic cells of two amphibian species, Xenopus lamis and R a m catesbeim. Environ. Toxicol. Chem. 22: 210621 13. Greulich, K. and Pflugmacher, S., 2003. Differences in susceptibility of various life stages of amphibians to pesticide exposure. Aquatic Toxicol. 65: 329-336. Hanazato, T. and Yasuno, M., 1990. Influence of Chaoborw density on t he effects of an insecticide on zooplankton communities in ponds. Hydrobiologia 194: 183-197. Hatch, A. C. and Blaustein, A. R., 2003. Combined effects of UV-B radiation and nitrate fertilizer on larval amphibians. Ecol. Applic. 13: 1083-1093. Hecnar, S. J., 1995. Acute and chronic toxicity of ammonium nitrate fertilizer to amphibians from Southern Ontario. Enuiron. Toxicol. Chem. 14: 2131-2137. Hels, T., 2002. Population dynamics in a Danish metapopulation of Pelobates fuscus. Ecography 25: 303-3 13. Henry, F! F. E , 2000. Aspects of amphibian anatomy and physiology. Pp. 71-1 10 in "Ecotoxicology of Amphibians and Reptiles", ed by D. W. Sparling, G. Linder and C. A. Bishop. SETAC Press, Pensacola. Hitchings, S. F! and Beebee, T J. C., 1997. Genetic substructuring as a result of barriers to gene flow in urban Rana temporaria (common frog) populations: implications for biodiversity conservation. Heredity 79: 117-127. Hopkins, W. A,, Mendonca, M. T, Rowe, C. L. and Congdon, J. D., 1998. Elevated trace element concentrations in southern toads, Bufo terrestris, exposed to coal combustion waste. Arch. Enuiron. Contamin. Toxicol. 35: 325-329. Howe, C. M., Berrill, M., Pauli, B. D., Helbing, C. C., Werry, K. and Veldhoen, N., 2004. Toxicity of glyphosate-based pesticides to four North American frog species. Ennircm. Toxicol. C h m . 23: 1928-1938. James, S. M., Little, E. E. and Semlitsch, R. D., 2004. Effects of multiple routes of cadmium exposure on the hibernation success of the American toad (Bufo americanus). Arch. Enuiron. Contam. Toxicol. 46: 518-527. Jennings, M. R. and Hayes, M. F,! 1994. "Amphibian and Reptile Species of Special Concern in California". California Department of Fish and Game, Inland Fisheries Division, Rancho Cordova. Kiely, T, Donaldson, D. and Grube, A., 2004. "Pesticide Industry Sales and Usage: 2000 and 2001 Market Estimates". US EPA, Office of Prevention, Pesticides, and Toxic Substances, EPA-733-R-04-001. Kiesecker, J. M., 2002. Synergism between trematode infection and pesticide exposure: A link to amphibian limb deformities in nature? Proc. Nut. Acad. Sci., USA 99: 9900-9904. Knutson, M. G., Richardson, W. B., Reineke, D. M., Gray, B. R., Parmelee, J. R. and Weick, S. E., 2004. Agricultural ponds support amphibian populations. E c o ~Applic. . 14: 669-684.

3206

AMPHIBIAN BIOLOGY

Kupferberg, S. J., 1997. Bullfrog (Rana catesbeiana) invasion of a California river: The role of larval competition. Ecology 78: 1736-1 75 1.

Parris, M. J. and Cornelius, T O., 2004. Fungal pathogen causes competitive and developmental stress in lama1 amphibian communities. Ecology 85: 3385-3395.

Lambert, M. R. K., 1997. Environmental effects of heavy spillage from a destroyed pesticide store near Hargeisa (Somaliland) assessed during the dry season, using reptiles and amphibians as bioindicators. Arch. Enuiron. Contam. Toxicol. 32: 80-93.

Pechmann, J. H. K. and Wilbur, H. M., 1994. Putting declining amphibian populations in perspective: natural fluctuations and human impacts. Herpetobgiaa 50: 65-84.

Lannoo, M., 2005. "Amphibian Declines: The Conservation Status of United States Species." Univeristy of California Press, Berkeley. Leftcort, H., Thomson, S. M., Cowles, E. E., Harowicz, H. L., Livaudais, B. M., Roberts, W. E. and Ettinger, W. F., 1999. Ramifications of predator avoidance: predator and heavy-metal-mediated competition between tadpoles and snails. Ecol. Applic. 9: 1477-1489.

Pechmann, J. H. K., Scott, D. E., Gibbons, J. W. and Semlitsch, R. D., 1989. Influence of wetland hydroperiod on diversity and abundance of metamorphosing juvenile amphibians. Wetlands Ecol. Manage. 1 : 1-9. Pechmann, J. H. K., Scott, D. E., Semlitsch, R. D., Caldwell, J. P, Vitt, L. J. and Gibbons, J. W., 1991. Declining amphibian populations: The problem of separating human impacts from natural fluctuations. Science 253: 892-895.

LeNoir, J. S., McConnell, L. L., Fellers, M. G., Cahill, T. M. and Seiber, J. N., 1999. Summertime transport of current-use pesticides from California's Central Valley to the Sierra Nevada mountain range, USA. Enuiron. Toxicol. Chem. 18: 2715-2722.

Peterle, T J., 1991. "Wildlife Toxicology". Van Nostrand Reinhold. New York.

Linder, G., Krest, S. K. and Sparling, D. W., 2003. "Amphibian Decline: An Integrated Analysis of Multiple Stressor Effects". SETAC Press, Pensacola.

Relyea, R. A. and Mills, N. E., 2001. Predator-induced stress makes the pesticide carbaryl more deadly to gray treefrog tadpoles (Hyla versicolor). Proc. Nut. Acad. Sci. 98: 2491-2496.

Lips, R. K. and Donnelly, M. A,, 2005. Lessons from the tropics. In "Amphibian Declines: The Conservation Status of United States Species", ed by M. J. Lannoo. University of California Press, Berkeley. Little, E. E., Calfee, R., Cleveland, L., Skinker, R., ZagaParkhurst, A. and Barron, M. G., 2000. Photoenhanced toxicity in amphibians: Synergistic interactions of solar ultraviolet radiation and aquatic contaminants. J. Ioula Acad. Sci. 107: 67-71. MacKenzie, C. A, Berrill, M., Metcalfe, C. and Pauli, B. D., 2003. Gonadal differentiation in frogs exposed to estrogenic and antiestrogenic compounds. Enuiron. Toxicol. Chem. 22: 24662475. Mann, R. M. and Bidwell, J. R., 1999. The toxicity of glyphosate and several glyphosate formulations to four species of southwestern Australian frogs. Arch. Enuiron. Contamin. Toxicol. 36: 193-199. McConnell, L. L., LeNoir, J. S., Datta, S. and Seiber, J. N., 1998. Wet deposition of current-use pesticides in the Sierra Nevada mountain range, California, USA. Environ. Toxicol. Chem. 17: 1908-1 9 16. Mills, N. E. and Semlitsch, R. D., 2004. Competition and predation mediate indirect effects of an insecticide on southern leopard frogs. Ecol. Applic. 14: 1041-1054. Morin, P J., 1981. Predatory salamanders reverse the outcome of competition among three species of anuran tadpoles. Science 212: 1284-1286. Muths, E., Corn, P S., Pessier, A. P and Green, D. E., 2003. Evidence for disease-related amphibian decline in Colorado. Biol. Cons. 110: 357-365. Park, D., Hempleman, S. C. and Propper, C. R., 2001. Endosulfan exposure disrupts pheromonal systems in the red-spotted newt: a mechanism for subtle effects of environmental chemicals. Enuiron. Health Perspec. 109: 669-673. Parris, M. J. and Baud, D. R., 2004. Interactive effects of a heavy metal and chytridiomycosis on gray treefrog larvae (Hyla chrysoscelis). Copeia 2004: 344350.

Relyea, R. A., 2003. Predator cues and pesticides: A double dose of danger for amphibians. Ecol. Applu. 13: 1515-1521.

Rohr, J. R., Elskus, A. A., Shepherd, B. S., Crowley, P. H., McCarth~,T M., Niedzwiecki, J. H., Sager, T, Sieh, A. and Palmer, B. D., 2003. Lethal and sublethal effects of atrazine, carbaryl, endosulfan, and octylphenol on the streamside salamander (Ambystoma barbouri). Enuiron. Toxicol. Chem. 22: 2385-2392. Rowe, C. L. and Dunson, W. A,, 1994. The value of simulated pond communities in mesocosms for studies of amphibian ecology and ecotoxicology.J. Herpetol. 28: 346-356. Rowe, C. L., Hopkins, W. A. and Coffman, V. R., 2001. Failed recruitment of southern toads (Bufo terrestris) in a trace element-contaminated breeding habitat: direct and indirect effects that may lead to a local population sink. Arch. Enuiron. Contamin. Toxicol. 40: 399-405. Rowe, C. L., Hopkins, W. A. and Congdon, J. D., 2001. Integrating individual-based indices of contaminant effects: How multiple sublethal effects may ultimately reduce amphibian recruitment from a contaminated breeding site. Scientzfic World 1: 703-7 12. Rowe, C. L., Kinney, 0. M., Nagle, R. D. and Congdon, J. D., 1998. Elevated maintenance costs in an anuran (Rana catesbeiana) exposed to a mixture of trace elements during the embryonic and early larval periods. Physiol. 2001.71: 27-35. Ryan, R. J. and Hites, R. A., 2002. Atmospheric transport of toxaphene from the southern United States to the Great Lakes region. Enuiron. Sci. Echnol. 36: 3474-348 1. Scott, D. E., 1994. The effect of larval density on adult demographic traits in Ambystoma opacum. Ecology 75: 1383-1396. Semlitsch, R. D., 1987. Relationship of pond drying to the reproductive success of the salamander Ambystoma talpoideum. Copeia 1987: 61-69. Semlitsch, R. D., 2000. Principles for management of aquatic-breeding amphibians. J. Wildl. Manage. 64: 615-631.

BOONE ET AL.: IMPACT OF PESTICIDES IN AMPHIBIAN DECLINES Semlitsch, R. D., 2003. "Amphibian Conservation". Smithsonian Press, Washington, DC. Semlitsch, R. D., Foglia, M. and Mueller, A., 1995. Short-term exposure to triphenyltin affects the swimming and feeding behavior of tadpoles. Enuiron. Toxicol. Chem. 14: 1419-1423. Semlitsch, R. D., Scott, D. E. and Pechmann, J. H. K., 1988. Time and size at metamorphosis related to adult fitness in Ambystoma talpozdeum. Ecology 69: 184-192. Semlitsch, R. D., Scott, D. E., Pechmann, J. H. K. and Gibbons, J . W., 1996. Structure and dynamics of an amphibian community: evidence from a 16-year study of a natural pond. Pp. 217-248 in "Long-term Studies of Vertebrate Communities", ed by M. L. Ccdy and J. A. Smallwood. Academic Press, San Diego. Skelly, D. K., 1994. Activity level and the susceptibihty of anuran larvae to predation. Anim. Behou. 47: 454-468. Smith, D. C., 1987. Adult recruitment in chorus frogs: effects of size and date at metamorphosis. Ecology 68: 344-350. Sparling, D. W., Bishop, C. A. and Linder, G., 2000. The current status of amphibian and reptile ecotoxicological research. Pp. 1-13 in "Ecotoxicology of Amphibians and Reptiles", ed by D. M'. Sparling, G. Linder and C. A. Bishop. Society of Environmental Toxicology and Chemistry, Pensacola. Sparling, D. W., Fellers, G. M. and McConnell, L. L., 200 1. Pesticides and amphibian population declines in California, USA. Envimn. Toxicol. Chem. 20: 1591-1595. Sparling, D. W., Linder, G. and Bishop, C. A., 2000. "Ecotoxicology of Amphibians and Reptiles". SETAC Press, Pensacola.

3207

Stebbins, R. C. and Cohen, N. W., 1995. "A Natural History of Amphibians". Princeton University Press, Princeton.

Stuart, S. N., Chanson, J. S., Cox, N. A., Young, B. E., Rodrigues, A. S. L., Fischman, D. L. and Waller, R. LV., 2004. Status and trends of amphibian declines and extinctions worldwide. Science 306: 1783-1 786. Susser, M., 1986. The logic of Sir Karl Popper and the practice of epidemiology. Amer J. Epidemiol. 124: 511-718. Thurrnan, E. M. and Cromwell, A. E., 2000. Atmospheric transport, deposition, and fate of triazine herbicides and their metabolites in pristine areas at Isle Royale National Park. Enuzron. Sci. Technol. 34: 3079-3085. Vonesh, J. R. and De la Cruz, O., 2002. Complex life qcles and density dependence: Assessing the contribution of egg mortality to amphibian declines. Oecologia 133: 325-333. Werner, E. E., 1986. Amphibian metamorphosis: growth rate, predation risk and the optimal size at transformation. Arne?: Nut. 128: 3 19-341. Wilbur, H. M. and Fauth, J. E., 1990. Experimental aquatic food webs: interactions between two predators and two prey. Arne?: Nat. 135: 176-204. Zabik, J. M. and Seiber, J. N., 1993. Atmospheric transport of organophosphate pesticides from California's Central Valley to the Sierra Nevada Mountains. J. Enuiron. Qual. 22: 80-90. Zaga, A., Little, E. E., Rabeni, C. F. and Ellersieck, M. R., 1998. Photoenhanced toxicity of a carbamate insecticide to early life stage anuran amphibians. Enuiron. Toxicol. Chem. 17: 2543-2553.

CHAPTER 9

Endocrine Disrupting Chemicals Krista A. McCoy and Louis J. Guillette Jr.

I. lntroduction II. Brief lntroduction to the Endocrine System Ill. Definition of Endocrine Disrupting Chemicals IV. The Nature and Sources of Endocrine Disrupting Chemicals V. Modes of Action - How and Why? A. Nuclear Receptors 6. Hormone Catabolism and Carrier Protein Binding C . Steroid Biosynthesis D. Epigenetics VI. Phenotypic Responses A. Thyroid 1. Effects of EDCs on the Amphibian Thyroid

B. Development - Homeobox Gene Expression C. Reproduction 1. Effects of EDCs on the Amphibian Reproductive System D. Behaviour 1. Effects of EDCs on Amphibian Behaviour 2. Effects of EDCs on Non-amphibian Behaviour VII. Implications A. Conservation and Ecology 0. Evolution VIII. What Needs to be Known IX. References

I. INTRODUCTION

E

NDOCRINE disrupting chemicals (EDCs) are ubiquitous pollutants and their presence in the environment has created an important and difficult issue in wildlife conservation (Colborn and Smolen 1996).Any physiological, behavioural or developmental process that is controlled or influenced by the endocrine system is susceptible to modification by EDCs. In fact, they are known to have dramatic effects on vertebrate development, physiology, and behaviour (Fox 2001; Hayes et al. 2002; Guillette and Iguchi 2003; Guillette 2004; Crews and McLachlan 2006; Milnes et al. 2006). Therefore, EDCs can change the reproductive success of individuals, alter patterns of sexual selection, and reduce the longterm fitness and persistence of affected populations. Indeed, environmental contaminants, such as EDCs, could have fbrther reaching and much more important effects on amphibians and other wildlife than is presently appreciated. Amphibians are thought to be particularly susceptible to the effects of EDCs because they have permeable skin, often reproduce in areas that receive large amounts of run-off and can be in intimate contact with environmental contaminants in all stages of their life history. For example, recent studies have documented reproductive abnormalities in amphibians from agricultural areas where EDCs are used (Russell et al. 1995; Ouellet et al. 1997; Hayes et al. 2002; Hayes et al. 2003; see Chapter 5 this volume). In addition, because EDCs are

McCOY and GUILLETTE Jr.: ENDOCRINE D I S R U R I N G CHEMICALS

3209

distributed globally they have been hypothesized to contribute to the global decline of amphibian populations (Carey and Bryant 1995; Hayes et al. 2003). It is important to note, however, that relatively little is known concerning the role EDCs potentially play in amphibian population declines. This is an important question to be addressed via research at all levels of organization, including the molecular and cellular mechanistic levels, and individual, population, and community levels. Although, to date, relatively little is known about the effects of EDCs on amphibians, there is substantial literature from other species, and a growing understanding of the endocrinology of amphibians from genes to individual physiological performance. Therefore, since many generalities have been observed, literature on these topics fi-om other vertebrates and various ecological systems are used here to better understand the data currently available for amphibians specifically, and examples of endocrine disruption in amphibians are emphasized. 11. BRIEF INTRODUCTION T O THE ENDOCRINE SYSTEM

The endocrine system consists of ductless endocrine glands that secrete hormones directly into the bloodstream or into surrounding tissues. Endogenous hormones transmit information and regulate biological processes (Fig. 1). Lipophilic hormones, such as steroid hormones, are well suited for signaling because they diffuse readily from a source cell (or capillary) to a target cell, where they bind to specific protein receptors (Fig. 2). The binding of a steroid hormone to its receptor induces a structural transformation that enables the hormone-receptor complex to bind to high affinity sites in the DNA and modulate gene transcription flamamoto 1985; Mangelsdorf et al. 1995) (Fig. 3). Steroid hormone receptors belong to a super-family of nuclear receptors that function as transcription factors, capable of altering the gene expression of thousands of genes (Watanabe et al. 2003; Terasaka et al. 2004). Therefore, hormones are secreted from an endocrine gland, and are transported through the body to regulate biological processes in "distant" tissues by modulating gene expression. Lipophilic hormones, including the steroids, retinoids, and thyroid hormones, are important regulators of development, cellular differentiation and organ physiology in all vertebrates studied to date (Mangelsdorf et al. 1995). Indeed, most EDCs identified to date are known to influence gene expression by interacting with the steroid, thyroid, and retinoid super-family of receptors (Crews and McLachlan 2006) (Fig. 2). In addition to interactions with hormone receptors, EDCs alter the function of hormone degradation, carrier proteins, and key steroidogenic enzymes, all of which are crucial for normal gene expression and hormonal homeostasis (Guillette and Gunderson 2001; Sanderson 2006) (Fig. 1). Altered gene expression can lead to a variety of negative effects - including abnormalities in the structure (via development and growth) and function of organs associated with hormonal signaling and regulation, or induction of cancers. EDCs are also known to influence gene expression epigenetically, which can induce transgenerational effects (Crews and McLachlan 2006; Mukerjee 2006). 111. DEFINITION OF ENDOCRINE DISRUPTING CHEMICALS

Endocrine disrupting chemicals are defined differently by various entities; the nuances among definitions can have important implications for legislation and can influence the way in which research on EDCs isconducted. For example, the World Health Organization's (WHO) International Program on Chemical Safety (IPCS) recently organized and conducted a Global Assessment of the State-of-the-Scienceof Endocrine Disruptors (IPCS 2002). The current definition of an endocrine disruptor according to that programme is "an exogenous substance or mixture that alters function(s) of the endocrine system and consequently causes adverse health effects in an intact organism, or its progeny, or (sub)populations"(IPCS 2002; Sanderson 2006). The European Commission's definition is similar to this but also includes a definition of a potential endocrine disruptor which acknowledges that consideration of a chemical's safety can be appropriate if that substance could possibly disrupt the endocrine

3210

AMPHIBIAN BIOLOGY

-

Sensory Input

Steroid Plasrr~a Binding Proteins

'ituitary LIVER nins

f

3

J\ Clean

sterols

~nsformation Excretion

'IPGONAD

\s Steroids

Gene Ekurs331un 1

RECEPTORS-

1,

;;rf,8

1%

Fig. 1. The endocrine system regulates physiological processes by integrating signals from the external environment via the sensory system with internal signals that are conveyed by endogenous (or internal) hormones (e.g., GnRH, FSH, LH, steroids, inhibin, activin). These hormones are secreted by various cells or organs (dark grey boxes), and regulate specific functions (light grey boxes) within target organs. Any physiological, behavioural or developmental process that is controlled or influenced by the endocrine system is susceptible to modification by EDCs. For example, EDCs alter the synthesis of steroid hormones (e.g., within the gonad) by altering key steroidogenic enzymes. Altered steroid production affects feedback mechanisms to the brain that in turn changes endocrine regulation. Altered steroid production also changes gene expression within target cells and organs. EDCs can also directly interfere with gene expression by binding to nuclear steroid receptors. In the liver, EDCs can interfere with hormone degradation and clearance and alter the function of steroid plasma-binding proteins. All of these processes (light grey boxes) are crucial for hormonal homeostasis.

system (European-Commission 1997). However, the US EPA (http://www.epa.gov/scipoly/ oscpendo/edsparchive/2-3attac.htm) defines an endocrine disruptor as "an exogenous agent that interferes with the synthesis, secretion, transport, binding, action, or elimination of natural hormones in the body which are responsible for the maintenance of homeostasis, reproduction, development and/or behaviour". The EPA explicitly states that the agency "does not consider endocrine disruption to be an adverse effect per se, but rather to be a mode or mechanism of action potentially leading to other outcomes, for example carcinogenic, reproductive, or developmental effects . . .". These differences are important because the WHO'S or European Commission's definition suggests that altered endocrine function can induce adverse health effects through any number of mechanisms, while the EPA's industry-friendly definition requires demonstration of a particular mechanism of toxicity that leads to adverse health effects beyond disruption of the endocrine system itself. Although it is clearly extremely informative to understand the exact mechanism of toxicity of a particular environmental pollutant and to identify relevant consequences of exposure, these details can, and do, take many years, and in some cases decades, to work out. One consequence is that many endocrine disrupting chemicals can remain in use for many years. In addition, this definition creates a situation where significant time, and research funding,

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUPTING CHEMICALS

321 1

Membrane bound receptors regulate cell metabolism and rapid responses

EDC

-

protein

lntracellular receptors regulate transcription and slow responses

Fzg. 2. Lipophilic hormones (grey crosses) easily diffuse from a capillary into a target cell and bind to specific intracellular protein receptors (black Vs). This complex (receptors and ligand) binds to the DNA and induces gene expression that modulates a wide range of physiological functions such as cellular growth and proliferation, development and differentiation, and maintenance of homeostasis (slow responses). Most EDCs (white trapezoids) identified to date influence gene expression by interacting with the steroid, thyroid, and retinoid receptors found within the cell (black Vs). Some EDCs can bind to, and activate, a ligand receptor that induces expression of responsive genes, whereas others block gene expression by competing for receptors without mimicking biological function. Recent evidence suggests that EDCs can also bind to membrane receptors and influence rapid cellular responses such as metabolism.

is spent working out mechanisms of action, and less time is spent in the field understanding effects at the population level. Endogenous hormones control a wide variety of living processes and are physiologically active at extremely low concentrations (in the pg/ml or ng/ml range). Consequently, pollutants that behave like endogenous hormones can induce effects at concentrations well below traditionally perceived "toxic" levels. Traditional chemical evaluation and risk-assessment procedures were not designed to manage such an unexpected mode of action. As a result, current regulations on the use of EDCs were made mandatory only after significant environmental damage had already occurred (Matthiessen and Johnson 2007). For the purposes of this chapter, EDCs are defined by paraphrasing Crews and McLachlan (2006) and Tabb and Bloomberg (2006). The resulting definition implies that there are multiple mechanisms through which endocrine disruption can occur and a diversity of effects and consequences to exposure: "Endocrine disrupting chemicals are a class of environmental pollutants that mimic or block transcriptional activation elicited by endogenous hormones such that they resemble (or alter) natural biological signals (hormones) and, thus, can be misinterpreted by the organism leading to abnormal gene expression and phenotype."

3212

AMPHIBIAN BIOLOGY

Fig. 3. Steroid, retinoid, and thyroid hormones are lipophilic substances that can cross the nuclear membrane (dashed circle). These hormones bind specific receptors (grey oval labelled "R") that induce allosteric transformation and enable the hormone-receptor complex to bind to high-affinity sites in the DNA, called response elements (REs). This binding modulates gene transcription. EDCs can interfere with receptor binding, modulate receptor degradation, alter receptor activity, or alter the activity and availability of co-factors (ovals shown above and between the receptor complex and RNA polymerase). All these processes are essential for normal receptor function and gene expression. Altered gene expression can lead to a variety of negative effects, including abnormalities in the structure and function of organs, or induction of cancers.

IV. THE NATURE AND SOURCES OF ENDOCRINE DISRUPTING CHEMICALS

Many endocrine disrupting chemicals are persistent either because they have a long half-life or because they are added to environmental systems at a rate faster than they break down or are metabolized. EDCs are also ubiquitous and many are distributed globally even areas once believed to be pristine are affected by their run off or fallout. Once deposited, EDCs can biomagnify in food chains (Nations and Hallberg 1992; Skaare et al. 2000; Thurman and Cromwell 2000; Bjerregaard et al. 2001; Lie et al. 2003; Goksoyr 2006; Guillette et al. 2006). For example, organochlorine pesticides (e.g., DDT) and industrial chemicals, such as polychlorinated biphenyls (PCBs), are found in high concentrations in arctic ecosystems. Skaare et al. (2000) showed that polar bears (Ursus maritimus) and glaucous gulls (Lam hyperboreus) from Svalbard, in the Arctic Ocean midway between Norway and the North Pole, contain high levels of PCBs. Polar bear PCB levels were greater than eight times higher than those found in ringed seals (Phoca hispzda), an important polar bear prey (Severinsen et al. 2000; Skaare et al. 2000). Similarly, glaucous gull PCBs are up to three orders of magnitude higher than in crustaceans and fish which make up their major food items ( Skaare et al. 2000; Borga et al. 2003).

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUPTING CHEMICALS

3213

Interestingly, there are sex differences in exposure to lipophilic environmental contaminants such as PCBs that are related to differences in excretory pathways. For example, male glaucous gulls and polar bears have higher PCB loads than do females because egg yolk and milk are important, female-specific, excretory pathways for lipophilic compounds (Skaare et al. 2000). In fact, polar bear yearlings typically have higher PCB levels than do their mothers, and are therefore exposed to high levels of PCBs during critical periods of growth and development when they are most sensitive to endocrine disruption (Skaare et al. 2000). The occurrence of high levels of PCBs and DDE (a metabolite of DDT) in the eggs of the green frog (Rana clamitam) and spring peeper (Pseudacris crucijer) in southern Ontario, Canada, suggests that sex-specific exposure and excretory pathways occur in amphibians, and that larvae are exposed to environmental as well as to maternally derived concentrations of contaminants at critical developmental stages (Russell et al. 1995, 1997). These exposures could certainly be negatively affecting amphibian populations in many places, but the ecological implications have not been well studied (Russell et al. 1995, 1997). Some EDCs enter the environment as "natural" sources such as hormones originating from humans, live stock, or plants (phytoestrogens). For example, some estrogenic hormones, such as 17P-estradiol and estrone, are naturally synthesized and excreted by women. Several studies have shown that these chemicals, as well as 17a-ethinylestradiol, the synthetic estrogen in contraceptives, can be found in aquatic environments associated with domestic waste water treatment plants at concentrations up to 5, 12, 47 and 7.5 ng/L respectively (e.g., Belfroid et al. 1999). Although many waste water treatment sites contain concentrations below detection limits, 17a-ethinylestradiol is known to induce the development of intersexed gonads in some gonochoristic fish at levels well below those detected by Belfroid ( E n g e et al. 2001; Jobling et al. 2002; Palace et al. 2002; Pawlowski et al. 2004; Norris and Carr 2006; Woodling et al. 2006). Furthermore, in the Ruhr district of Germany, 17a-ethinylestradiol (used in birth control) has been detected in surface waters in concentrations between 1-4 ng/L, which can induce intersexed gonads. The naturally synthesized estrogens, 17gestradiol and estrone, however, were below the quantification limit of 1 ng/L (Lintelmann et al. 2003). Another source of "naturally derived" EDC pollution involves the hormones released from the waste of animals on feedlots or from waste used as fertilizer (reviewed by Johnson et al. 2006). These hormones are either naturally synthesized, or "prescribed" as growth stimulants, or for other purposes. Intensive livestock rearing or agriculture leads to runoff of androgenic hormones at levels high enough to induce endocrine disruption in wild fathead minnows (Pimephales promelas) (Ankley et al. 2003; Orlando et al. 2004; Soto et al. 2004; Matthiessen et al. 2006). Steroid hormones, such as 17gestradiol and testosterone, have even been found in spring water from mantled karst aquifers in agricultural areas (Peterson et al. 2000). Therefore, hormone contamination from agricultural practices is not limited to surface waters and is likely distributed across long distances via ground water. Phytoestrogens are endogenous compounds found in plants and are structurally and functionally similar to estradiol and other endogenous estrogens. They are released into the environment in a variety of ways, including sewage treatment plant effluent (Spengler et al. 2001; Pawlowski et al. 2003), runoff from agricultural areas treated with manure (Burnison et al. 2003) and effluent from wood pulp mills (Mahmood-Khan and Hall 2003; Clotfelter et al. 2004). Although it is easy to understand how pollution containing natural or synthetic hormones can alter the endocrine systems of wildlife, there is an extraordinarily diverse array of man-made (synthetic) chemicals that also induce endocrine disruption. These chemicals include pesticides, flame retardants, plastics or plasticizers, heavy metals, industrial compounds (e.g., coolants or insulators), pharmaceuticals (e.g., antidepressants) and cosmetics (Propper 2005; Goksoyr 2006). Each of these broad categories contains many different chemicals that are known or suspected to induce endocrine disruption. Importantly, individual chemicals within and among these general categories (e.g., pesticides) can have very different chemical structures or be from different chemical classes, which complicates the ability to identift. or predict chemicals that cause endocrine disruption.

3214

AMPHIBIAN BIOLOGY

Despite the large diversity of chemicals that can disrupt the endocrine system, they often have similar effects, albeit via different mechanisms of toxicity. For example, most known EDCs are estrogenic (induce estrogen-like effects), which can lead to feminization of males or development of intersexed gonads (Crews and McLachlan 2006). However, a growing literature also has shown that aquatic pollutants potentially alter thyroid, androgen, progesterone and retinoic acid signaling (Guillette 2006). In fact, many PCBs and polybrominated biphenyl ethers (PBDEs), used for decades as flame retardants, alter thyroid hormone function (Zoeller et al. 2002). A growing list of fungicides and other pesticides have anti-androgenic activity (Gray et al. 2006). Interestingly, no synthetic environmental compounds have been reported to have androgenic effects (McLachlan 2001) except pharamaceutical agents (i.e., synthetic androgens) prescribed by, and released into, the environment as part of cattle feedlot operations (Durhan et al. 2007). Plant sterols found in paper-mill effluent, however, can be metabolized by native aquatic bacteria to produce androgens, and have been shown to masculinize female fish (e.g., Gambusiu afinis holbrooki) (Denton et al. 1985; Parks et al. 2001; Toft et al. 2004). Thus, it is important to recognize that no single endocrine mechanism is at risk, but rather, EDCs can alter various endocrine signaling pathways via multiple mechanisms.

V. MODES OF ACTION - HOW AND WHY? The most commonly studied mode of action of EDCs is their action as a ligand (e.g., a hormone), thus inducing inappropriate activation or antagonism of nuclear hormone receptors (Fig. 2). However, hormone receptors can be influenced through several other mechanisms. EDCs can modulate receptor degradation, or alter receptor activity, such as by changing the activity or availability of co-factors essential for receptor function (Tabb and Blumberg 2006). In addition, EDCs can modulate the degradation and clearance of endogenous hormones or alter synthesis of hormones, such that inappropriate amounts or types of steroid hormones are formed (Guillette and Gunderson 2001; Sanderson 2006; Tabb and Blumberg 2006). Finally, and possibly most importantly, some EDCs have been shown to alter genome-wide DNA methylation patterns. Methylation controls gene expression and if it occurs during particular developmental windows, gene expression patterns can be permanently altered, and these changes are inherited by offspring (Edwards and Myers 2007). A. Nuclear Receptors

Nuclear receptors are protein receptors that bind to specific hormones (also called ligands) (Figs 2, 3). This binding changes the conformation of the receptor such that it can bind with another receptor-ligand complex or with a coactivator-ligand complex. After this dimerization, the complex binds to specific DNA sequences in the regulatory region of target genes (called responsive elements) (Fig. 3), and triggers gene expression (Janosek et al. 2006). There are 47 known nuclear receptors and, as a group, they constitute the nuclear receptor superfamily. These receptors are involved in modulating a wide range of physiological functions across eukaryotes including cellular growth and proliferation, development and differentiation, and maintenance of homeostasis (Blumberg and Evans 1998; Janosek et al. 2006). EDCs can bind to and activate a receptor and induce expression of responsive genes (McLachlan 2001; Sanderson 2006). Alternatively, other EDCs bind to the receptor and block the endogenous hormone from its receptor, which results in inappropriately low levels of ligand-responsive gene expression. Such influences on sex hormone receptors have been extensively studied over the past few decades (Tabb and Blumberg 2006; for a review see Janosek et al. 2006). Some chemicals, such as ethanol, however, activate transcription of estrogen-controlled genes by influencing up-stream genes, not by binding the estrogen receptor (ER) Uanosek et al. 2006). Many members of the nuclear receptor superfamily are degraded via the ubiquitinproteasome pathway. Ubiquitin labels old or damaged proteins for degradation, which is then carried out by proteosomes (proteins that degrade other proteins). Receptor turnover is one mechanism whereby cells control gene expression and prevent over stimulation by endogenous

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUETING CHEMICALS

3215

hormones (Tabb and Blumberg 2006). EDCs can modulate receptor degradation because they do not induce the same proteosome response that endogenous hormones elicit. For example, Masuyama and Hiramatsu (2004) showed that in the presence of estradiol both estrogen receptor a (ERa) and P (ERP) interacted with active sites within the proteosome and were degraded. However, in the presence of Bisphenol A (a constituent of many plastics) which activates ER transcription, ERP degradation was blocked (Masuyama and Hiramatsu 2004). Reduced degradation of ERP leads to a build up of these receptors in the cell, and thus is expected to lead to increased expression of ERP-responsive genes (Tabb and Blumberg 2006). Other EDCs have been shown to alter ligand receptor activity (Jansen et al. 2004) or the activity of co-activators essential for receptor function (Tabb and Blumberg 2006). B. Hormone Catabolism and Carrier Protein Binding Altering hormone biotransformation (synthesis) and catabolism (break down) is yet another way EDCs can modulate hormone balance (Guillette and Gunderson 2001; Tabb and Blumberg 2006). For example, two nuclear receptors are important regulators of the catabolism of xenobiotics (biologically active pollutant or drug), chemicals, and steroid hormones. These receptors are: (1) human steroid and xenobiotic receptor/rodent pregnane X receptor (SXRJPXR) and (2) constitutive androstane receptor (CAR) (Forman et al. 1998; Xie et al. 2000; Kretschmer and Baldwin 2005; Tabb and Blumberg 2006). Both of these receptors are highly expressed in the liver and intestine, and control induction of several key enzymes that orchestrate the biotransformation, catabolism, and transport of endogenous hormones and xenobiotics. Many EDCs have been found to activate these receptors and increase expression of their target genes (Kretschmer and Baldwin 2005; Tabb and Blumberg 2006). EDC-induced activation of enzymes that degrade hormones and xenobiotics has a two-fold effect. First, endogenous hormones have a shorter half-life and thus are biologically active for less time. Second, in degrading xenobiotics, these enzymes increase levels of EDC metabolites, some of which are more toxic than the parent compounds. Plasma-binding proteins sequester and control the bioavailability of hormones such that they cannot pass through the plasma membrane and bind to nuclear receptors (Nagel et al. 1997). Many xenobiotic chemicals do not bind to plasma-binding proteins and, thus, are readily available to the cell (Arnold et al. 1996; Crain et al. 1998). Thus, they would not be excluded and could induce endocrine disruptive actions. In contrast, Danzo (1997) showed that several xenobiotics induced disassociation of endogenous ligands from their binding proteins and competed for binding proteins just as strongly as did natural ligands. Therefore, xenobiotics can competitively bind to steroid-binding proteins, thereby increasing the bioavailablity of endogenous hormones because they are free to enter the cell (Danzo 1997). In addition, several studies have shown that receptors for these binding proteins can be present on the plasma membranes of some cells. The hormone-binding protein complex binds to these receptors and initiates a signal transduction cascade (for a review see Danzo 1998). Xenobiotics that bind in place of the endogenous hormones can inhibit or stimulate signal transduction (Danzo 1998). C. Steroid Biosynthesis

Steroid hormone synthesis involves a complex network of chemical reactions that convert a cholesterol-based substrate into a product (Fig. 4) through the activity of several substratespecific cytochrome P450 enzymes, steroid dehydrogenases, and reductases (Sanderson 2006; Guillette et al. 2007). The cytochrome P450 enzymes are especially important because each one is responsible for specific chemical conversions that are essential for steroid biosynthesis (Miller 1988). Importantly, the activity of many of these enzymes can be modulated by Parious endocrine-disrupting chemicals, resulting in impaired development, growth, sexual differentiation, reproduction, and the development of particular cancers (for a review see Sanderson 2006). For example, aromatase (CYPl9) is responsible for controlling the ratehniting step in the conversion of androgens to estrogens (e.g., testosterone to 17&estradiol), md its modulation by EDCs has been shown to interfere with the homeostasis and function

AMPHIBIAN BIOLOGY ClIobSteml

20'22-demIaso

I

20a -hyimxylase

20% 22~r- H y ~ c h o l e E f e r o lz 22- IRdrowIase 20. -Hyd-hohstma1

i

Preenemlone 3 w * .

I

Pmprhone

17a -kydmx+

21-mxylase

~ll-Deo~).cmbcorteronemn l=~onicOshone

17a -hpiro&ase

17a-Hgdrca~~re-olo~ 38dompm.

17a-Hydmqpro~enerone

17&~molra.

21 -lra..11-Deowconisol IRdroxy

.

"@ W x y h Conisol

17desmolra.

Dehyhepiandrorterone. ~-3

17B-*tnss

.

Testosterone

'aa 5a-

DihydrotestDnaarone

AfcwnUmgevs upctudvg Aromotas8

Esmne

17~-dohydmgo~l 17gestradial 17~-&~genm-2

Fig. 4. Steroid hormone synthesis involves a complex network of chemical reactions that convert a cholesterol-based substrate into a product (e.g., hormones) through the activity of several substrate-specific cytochrome P450 enzymes, steroid dehydrogenases, and reductases. The activity of many of these enzymes can be modulated by various endocrine-disrupting chemicals, resulting in impaired development, growth, sexual differentiation, reproduction, and the development of particular cancers.

of sex steroid hormones, resulting in feminization and demasculinazation (Crain and Guillette 1997). For example, several triazine herbicides (atrazine, simazine, and propazine) and a number of their metabolites (atrazine desethyl and atrazine desisopropyl) induced aromatase activity and gene expression up to 2.5-fold in human H295R adrenocortical carcinoma cells (Sanderson et al. 2000; Sanderson et al. 2002; Fan et al. 2007). Similar results were shown in alligator neonates (Crain and Guillette 1997). This mechanism could explain why male amphibians exposed to artrazine in laboratory and field studies are feminized and have intersexed gonads (Hayes et al. 2002, 2003). Alternatively, Ankley et al. (2005) demonstrated the fungicides prochloraz and fenarimol inhibit aromatase activity as well as bind to the androgen nuclear receptor in the fathead minnow (Pzmephales promelas). Both fungicides caused significant anti-androgenic alterations in endocrine hnction of the fish, thereby decreasing reproductive success (Ankley et al. 2005). D. Epigenetics

Epigenetics refers to changes in gene function that are heritable (mitotically or meiotically) but do not involve changes in nucleotide sequence. For example, all the cells in the body of a multi-celled organism originate from the same single-celled zygote and thus share the same DNA sequences. Processes that lead to cellular differentiation and maintenance of specific cell functions involve epigenetic changes in genes rather than change in nucleotide sequence (e.g., mutation). In other words, gene function and expression among different cells is altered in a mitotically heritable way, in this case without changing the DNA sequence. One mechanism through which this occurs is by methylation of DNA. This occurs when methyl groups are added to cytosines in the DNA. Methylation is inherited by daughter cells through DNA synthesis and cell division, and typically leads to suppression of gene expression, whereas de-methylation is associated with increased gene expression (Holliday and Pugh 1975; Ballestar and Esteller 2002; Crews and McLachlan 2006; Edwards and Myers 2007). For example, several of the inactive genes on female Barr bodies (redundant sex chromsomes) are methylated and the active and inactive states of these

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUPTING CHEMICALS

3217

chromosomes are inherited from mother cell to daughter cells. Therefore, the pattern of DNA methylation among chromosomes (and alleles) of somatic cells is stably maintained, but germ cells and pre-implantation embryos can experience genome-wide re-programming of the methylation pattern that generates cells with new (meiotically) heritable developmental potential (Reik et al. 2001; Reik and Walter 2001a). Epigenetic processes, via differential methylation, are also involved in gene imprinting which occurs when genes function differently depending on whether they are located on maternal or paternal chromosomes (Reik and Walter 2001b). The DNA of eggs and sperm are methylated differently and inheritance of these differences leads to differential gene expression between the sexes (Zuccotti and Monk 1995). In addition, during gonadal sex determination DNA in primordial germ cells is demethylated and remethylated in a sex-spec&c manner (Reik and Walter 2001a). The methylation patterns that occur in the germ line can be permanent and inherited from parent to offspring. The mechanisms controlling methylation during development are under intense investigation, and estrogens are thought to be involved in some cases (McLachlan 2001). For example, estrogens, through interactions with their receptors, are known to increase the levels of c-fos and c-jun in target cells (Kamiya et al. 1996; McLachlan 2001). C-fos and c-jun are transcription factors that are up-regulated in response to many physiological signals, and in turn up-regulate transcription of other genes involved in a diverse array of functions. Over-expression of c-fos leads to an-upregulation of (for example) cytosine methyltransferase, the enzyme involved in increasing DNA methylation (McLachlan 200 1). Estrogen's influence on methylation, however, is complex and can be tissue and gene specific. For example, 17b-estradiol exposure in male white Leghorn roosters is associated with removal of methyl groups at the estrogen response element (estradiol-receptor binding site) of a vitellogenin (yolk protein) gene, and this demethylation is associated with inappropriate transcription and translation of vitellogenin in males (Saluz et al. 1986; Jost et al. 1990). A similar mechanism of vitellogenin activation is known to occur in frogs (e.g., Xenopus laevis) (Andres et al. 1984; Crews and McLachlan 2006). Hormone exposure early in life can alter methylation, and thus epigenetically alter the set point for later response to the same or different hormones (Saluz et al. 1986). Given the potential role of hormones, such as estrogens, in controlling methylation, and the complexity of the mechanisms involved, one should expect EDCs that modulate plasma concentrations of estrogens, or influence their receptor expression, to induce changes in methylation patterns in ways that could be heritable. Indeed, it has recently been demonstrated that several EDCs can influence epigenetic programming through DNA methylation, and that when these changes occur at particular stages during development they are permanent and inherited by offspring (Crews and McLachlan 2006; Tabb and Blumberg 2006). For example, Anway et al. (2005) showed that rat pups (Rattus rattus) exposed to the fungicide vinclozilin (anti-androgenic) or the insecticide methoxyclor (estrogenic) during sexual differentiation experienced inappropriate genome-wide methylation patterns in the male germ line. The altered methylation patterns were associated with altered sperm development and lower reproductive success (Anway et al. 2005). This methylation pattern and reduction in fertility was transmitted through the male germ line to nearly all males in each generation across the four generations that were examined. The ability for endocrine disrupting chemicals to reprogramme the germ line and induce trans-generational pathologies clearly has important implications for ecology, evolution, and conservation of exposed wildlife, including amphibians.

VI. PHENOTYPIC RESPONSES Examining the effects of EDCs is difficult and complex for a variety of reasons. There are multiple ways to define endocrine disruption, multiple physiological mechanisms through rrhich disruption occurs, diverse abnormalities and pathologies that can be induced by EDCs, and multiple levels at which to study these effects. In addition, most toxicological studies

3218

AMPHIBIAN BIOLOGY

have focused on identifying effects of high concentrations of contaminants (e.g., mgIL), often focusing on measuring end points such as mortality. Fewer studies have examined sublethal effects of chemicals, such as endocrine disruption, at ecologically, or physiologically, relevant concentrations (e.g., yg/L). In particular, it is difficult to understand the effects of EDCs on amphibian populations because relatively little research on the effect of these chemicals has been conducted on this group of vertebrates, and almost all of it has been based on very few species under laboratory conditions. It is known, however, that several chemicals can modulate timing of metamorphosis (e.g., Freeman et al. 2005), as well as drastically alter organ development. For example, Hayes et al. (2003) collected leopard frogs (Rana pipiens) that were intersexes (male and female gonads in the same gonochoristic individual) from sites across the United States that were contaminated with the widely used herbicide atrazine, as well as other agricultural chemicals. In addition, the present authors (in preparation) have found a clear relationship between exposure to agricultural habitats where known EDCs, such as atrazine, are used, and intersexed primary and secondary sexual characteristics in Bufo marinus. They also observed that pollutants such as nitrate, largely from fertilizer use, a ubiquitous global pollutant of fresh water and near marine ecosystems, can alter the rate of development of tadpoles, presumably through an effect on thyroid hormone function, and can alter ovarian steroidogenesis (Barbeau et al. 2007; Edwards and Myers 2007). A. Thyroid

The thyroid system is highly conserved among vertebrate species, so thyroid hormone chemistry, its synthesis and delivery system, its receptors, and its regulation within the hypothalamic-pituitary-thyroid (HIT) axis are all comparable across vertebrates (Zoeller and Tan 2007; Zoeller et al. 2007). The thyroid gland and its hormones regulate growth, development, metabolism, and reproduction, so alterations of the thyroid system caused by EDCs are of concern (Opitz et al. 2006a). The thyroid system, like other hormones, involves many mechanisms that can be altered by EDCs (as discussed above), such as alterations in receptor binding, hormone synthesis, and clearance. These processes and mechanisms are affected in similar ways across diverse taxa, including mammals and amphibians (Opitz et al. 2006a; Fort et al. 2007). Several groups of chemicals are known, or are thought to, induce thyroid disruption (Boas et al. 2006). For example, polychlorinated biphenyls (PCBs) (such as coolant/insulating fluids and flame retardants), dioxins (e.g., agent orange, chemicals found in some insecticides, and industrial byproducts from chlorination) and polybrominated biphenyl ethers (PBDEs) (such as some flame retardants) are known to cause insufficient synthesis of thyroid hormone (hypothyroidism) in exposed animals. Environmentally relevant doses appear to affect human thyroid function as well. Although data on the effects of flame retardants on human populations is limited, there is great concern as the concentrations of these chemicals have dramatically increased in mother's milk over the past decade. These compounds appear to be similar in action to the PCBs, which have been studied in detail for decades and which clearly influence thyroid function. Also of major concern are the phthalates, used as plasticizers and chemical stabilization agents (e.g., in personal care products), as they also affect thyroid function, but appear to be stirnulatory (Boas et al. 2006). 1. Effects of EDCs on the Amphibian Thyroid

Structural and functional changes required for successhl amphibian metamorphosis are dependent on the presence and appropriate concentrations of the thyroid hormones 3,5,3'triiodothyronine (T,), and 3,5,3',5'-tetraiodothyronine(T,) (Sachs et al. 2000; Crump et al. 2002; Fort et al. 2007). Thyroid hormones bind to nuclear thyroid receptors (TR), and this complex can activate, or inhibit, gene transcription in a tissue-specific manner. Particular tissues are proliferated after thyroid hormone exposure, whereas others undergo apoptosis (programmed cell death) (Helbing et al. 1992, 1996; Helbing and Atkinson 1994; Sachs et al. 2000; Crump et al. 2002). Blocking the natural synthesis of thyroid hormone inhibits

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUFTING CHEMICALS

3219

metamorphosis whereas exposing tadpoles to exogenous thyroid hormone induces precocious metamorphosis (Sachs et al. 2000). Therefore, thyroid hormones are essential for stimulating the complex and diverse genetic programmes required for amphibians to successfully complete metamorphosis. The essential nature of thyroid hormones for amphibian metamorphosis, coupled with the fact that thyroid function is well conserved across vertebrates, has lead to the suggestion that tadpoles are an ideal bioassay system to identify thyroid function disruption by environmental contaminants (Crump et al. 2002; Degitz et al. 2005; Opitz et al. 2005; Zhang et al. 2006; Fort et al. 2007). A chemical evaluation protocol referred to as the Xenopus metamorphosis assay ( X E M A ) has been developed and evaluated via a ring test where six laboratories participated to evaluate inter-laboratory variation in results (Opitz et al. 2005). Opitz et al. (2005) demonstrated that the XEMA test can provide a sensitive, robust, and practical approach for the detection and evaluation of chemicals that affect the thyroid system. acetamide] The herbicide acetochlor [2-chloro-N-(ethoxymeth~1)-~\r-(2-ethyl-6-methyphenyl) has been shown to accelerate metamorphosis in several amphibian species (Cheek et al. 1999; Veldhoen and Helbing 2001). Acetocholor, however, only accelerates metamorphosis in animals that are "primed" by previous exposum to thyroid hormone. Therefore, it was suggested that acetochlor did not bind the thyroid receptor directly but enhanced T, action by different receptor-mediated mechanisms (Cheek et al. 1999). Veldhoen and Helbing (2001) showed that an environmentally relevant dose of acetochlor significantly enhanced TRP mRNA levels in T3-primed Rana mtesbeiana within 24 hours. Therefore, acetochlor does not bind to TRP but it does increase -Il43 expression, so that endogenous thyroid hormone can bind (Veldhoen and Helbing 2001). Given that there is a natural spike in thyroid hormones just before metamorphosis, metamorphic animals could be "naturally primed" by thyroid hormone. Under these conditions, acetochlor exposure could induce higher than usual TRP (and other) mRNA levels and cause the animals to proceed through metamorphosis faster than normal. The ecological implications of this acceleration are as of yet unknown, but could lead to higher mortality, smaller size at metamorphosis, and altered morphology. Crump et al. (2002) investigated acetochlor's influence on gene expression in Xenoupus Zuevis by developing a 420-gene cDNA array from known frog genes. They found that 26 genes are modulated by thyroid hormone and acetochlor (after thyroid hormone priming). Twenty-four genes \\-ere up-regulated whereas only two were inhibited. A detailed understanding of the physiological changes induced by altering this number of thyroid-dependent genes required for normal metamorphosis or the long-term effects on juvenile and adult metabolism and health are unknown. Another approach to the investigation of EDCs is to study their effects at the tissue level. Changes in thyroid histology (morphology) are indicative of alterations in the physiology of the pituitary-thyroid axis. k r example, inhibition of thyroid hormone secretion inhibits the negative feed-back on the pituitary which would normally stop secretion of thyroid stimulating hormone (TSH). This induces increased TSH that results in an increase of the size of thyroid gland follicles (hypertrophy of thyrocytes) and this effect can be observed and quantified histologically. For example, perchlorate, an oxidizer used in solidfuel rockets, inhibits thyroid hormone synthesis by inhibiting the sodium-iodide symporter that acts as an ion pump transporting iodide into thyroid epithelial cells so that they can synthesize thyroid hormones (Tietge et al. 2005; Opitz et al. 2006b). The effects of perchlorate on cricket frogs (AcmF crepztanr) from contaminated streams in central Texas were evaluated through histological analyses. Individuals living in streams with the greatest mean water perchlorate concentrations (-25 pg/L) showed significantly greater follicle cell hypertrophy. In addition, a significant positive correlation was found between thyroid follicle cell height and mean water perchlorate concentrations across all sites (Theodorakis et ad. 2006). Similar results were found for Ca?npostomaanomalum, a species of fish, from the same habitats, which suggests that perchlorate affects thyroid function across many taxa through a similar mechanism. Perchlorate reduced thyroid hormone, and the thyroid follicles experienced prolonged stimulation by TSH.

3220

AMPHIBIAN BIOLOGY

Since the mechanism whereby perchlorate effects thyroid function is known, it is an appropriate chemical to use to help understand how effects induced by EDCs at the tissue level can scale up to influence processes, such as metamorphosis, at the individual level. For example, African clawed frogs (Xenopuslaevis) exposed to ecologically relevant concentrations of ammonium perchlorate (59 and 140 ,ug/L) experienced significant hypertrophy of the thyroid follicular epithelium, as well as inhibition of forelimb emergence, tail resorption, and hindlimb development relative to control animals (Goleman et al. 2002). All these findings suggest that perchlorate alters thyroid fbnction (as expected), but they also demonstrate that many thyroid-hormone-dependant developmental programmes required for metamorphosis are negatively affected. Interestingly, only the highest concentration of perchlorate reduced whole-body thyroxine concentrations relative to the control treatment. This finding highlights the importance of measuring multiple endpoints; if thyroid hormone concentrations were measured without consideration of other morphological endpoints, the conclusion would have been drawn that perchlorate does not influence thyroid function in this species. In addition to alterations in metamorphic development, Goleman et al. (2002) found that perchlorate induced a skewed sex ratio such that significantly fewer males were recorded at metamorphosis. They concluded that ammonium perchlorate altered thyroid activity and gonadal differentiation in developing X. laevis, which suggests that animals that proceeded through seemingly normal metamorphosis could suffer lower reproductive success as adults. Importantly, the concentrations of perchlorate used in this study were below concentrations reported in surface waters contaminated with ammonium perchlorate, so perchlorate contamination likely poses a threat to normal development in natural amphibian populations (Goleman et al. 2002). In a similar study, Tietge et al. (2005) exposed X. laevis tadpoles to sodium perchlorate during metamorphosis, and measured thyroid histology as well as metamorphic timing. Histological effects on the thyroid were found at levels as low as 16 pg/L and metamorphosis was retarded significantly by perchlorate concentrations as low as 125 pgl L. Histological effects occurred at concentrations below those required to induce delays in metamorphosis which suggests that thyroid histology is a more sensitive biomarker for detecting thyroid-disruption induced by perchlorate (Tietge et al. 2005). Importantly, it is unclear how such alterations in the thyroid during development influences later life stages (e.g., adult metabolism) independent of whether metamorphosis was delayed. B. Development

- Homeobox Gene Expression

A significant amount of the early work on EDCs focused on developmental endpoints as the developing embryo is sensitive to very small concentrations of hormones required for normal development of many organs and organ systems (Bern 1992). It is recognized today that the developing embryo has sensitive windows of responsiveness when endocrine signals, at appropriate concentrations, induce organizational changes that fundamentally modify cellular differentiation and endocrine responsiveness later in life (Guillette et al. 1995). Many diverse genes, including the Homeobox (Hox) genes, have been shown to be responsive to endocrine signaling, making them targets for EDCs. Hox genes are evolutionarily conserved transcription factors that are essential for orchestrating embryonic development as well as cellular differentiation in adults. The expression of particular collections of Hox genes in specific places within the embryo creates a collinear expression continuum that regulates development in a segment-specific pattern and determines the anterior-posterior axis in the developing embryo (Daftary and Taylor 2006). Hox genes are also essential in the adult where they mediate cellular differentiation and control the developmental plasticity required to specify the function of new cells. Although Hox genes have been studied, and their role in cell fate determination is well known, the mechanisms by which their expression is controlled are not well understood. Recent work has demonstrated that nuclear receptors and their corresponding ligands (hormones) can regulate Hox gene expression in the embryo and adult (reviewed by Daftary and Taylor 2006). For example, in mice (Mus musculus) estrogens are necessary for the normal Hox gene expression required during embryonic development of the female

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUITING CHEMICALS

3221

reproductive tract (Block et al. 2000; Daftary and Taylor 2006). Exposure to diethylstilbestrol (DES), a synthetic non-steroidal estrogen, results in altered endogenous estrogen signaling and a change in the spatial expression of Hoxa9, HoxalO, and Hoxal 1, such that each is expressed more caudally throughout the developing female reproductive tract and leading to alterations in its morphology (Block et al. 2000; Daftary and Taylor 2006). Human females exposed to DES during development exhibit reproductive tract abnormalities with phenotypes that can be explained by a similar shift in Hox gene expression (Block et al. 2000; Daftary and Taylor 2006). For example, glandular tissue normally present in the uterus and cervix is observed in the vagina in women exposed in-utero to DES, which is consistent with caudal displacement of HoxalO and Hoxall expression (Block et al. 2000; Daftary and Taylor 2006). This work demonstrates that endocrine regulation of Hox genes by estrogens during embryogenesis is necessary across a variety of taxa. Estrogens are also thought to control functional differentiation in the adult reproductive tract (Daftary and Taylor 2006). Given the critical role of hormones such as the estrogens in modulating Hox gene expression, any perturbation to these endogenous signals by EDCs is expected to induce developmental abnormalities (Iguchi et al. 2001). Indeed, methoxychlor (a replacement insecticide for DDT) reduces estrogen binding to the estrogen receptor (ER), and disrupts the ability of the l7P-estradiol-ER complex to bind to the estrogen response element (ERE) for the HoxalO gene in endometrial cells, resulting in a decrease in HoxalO gene expression (Fei et al. 2005). Therefore, methoxychlor functions as an endocrine disruptor by affecting 17P-estradiol signaling in endometrial cells. In mice and rats, these changes inhibit modification of the endometrium (to create the dicidua), which blocks implantation of the embryo, and has deleterious effects on fertility. Importantly, in mice, this reduction of HoxalO gene expression is permanent and continues in adults even if they were only exposed as neonates (Fei et al. 2005). The mechanisms inducing continued repression of Hox gene expression in the absence of continued methoxychlor exposure are not known, but likely result from epigenetic modification, possibly via increased methylation (McLachlan 2001; Fei et al. 2005). Indeed, Wu et al. (2005) clearly demonstrated that differences in methylation of HoxalO, and the subsequent decline in HoxalO gene expression, is significantly associated with increased endometiosis in woman, and they argue that endometriosis is an epigenetic disease. Unfortunately, to our knowledge no work has explicitly evaluated the occurrence and importance of EDC exposure on Hox gene expression in amphibians. It is likely that similar effects are operating in amphibians because Hox genes, and their roles in embryonic development and cellular differentiation in adults, are highly conserved. Important aspects of amphibian development and metamorphosis are controlled by Hox genes. For example, the number, position, and type of limbs in amphibians depend on the proper expression of specific Hox genes. Retinoic acid regulates Hox gene expression and altering the quantity of retinoids during development results in serious developmental abnormalities across various taxa (Gardiner et al. 2003; Koussoulakos 2004). Application of retinoic acid to chicken limb buds alters homoeobox gene expression and induces duplication of limbs (Ogura and Evans 1995). Indeed, one of the most compelling explanations for the skeletal dysplasias observed in frogs from Minnesota, and elsewhere, is that the severely malformed limbs are due to exposure to environmental contaminants that function as retinoid (e.g., Vitamin A and retinoic acid) mimics (Gardiner et al. 2003). Gardiner et a1 (1999) analysed skeletal abnormalities in the Minnesota frogs and identified two major classes of abnormalities (Gardiner and Hoppe 1999). First, the initiation of limb development was altered, which induced absent or supernumerary limbs. Second, limb growth and pattern formation were modified such that primary and supernumerary limbs had specific skeletal abnormalities, including truncation and dwarfism. Skeletal elements were also folded back on themselves, forming characteristic "bony triangles" (Gardiner and Hoppe 1999; Gardiner et al. 2003). Retinoid (e.g., retinoic acid) exposure during limb bud development can induce the formation of boney triangles in frogs, chickens, and mice (Gardiner et al. 2003). In addition,

3222

AMPHIBIAN BIOLOGY

retinoic acid can induce supernumerary limbs, incomplete or missing limbs, complete limb duplications, as well as duplicated pelvic girdles across diverse taxa, including several frog species (Gardiner et al. 2003). Most of these abnormalities have been observed in field-collected malformed specimens. Therefore, it is possible that a single pollutant that functions as a retinoid mimic could be responsible for all the malformed phenotypes that have been observed (Gardiner et al. 2003). In addition, Gardiner et al. (2003) extracted hydrophobic substances from water samples collected from sites where malformed frogs were routinely found, and tested them for their ability to activate the retinoic acid receptor (RAR). Biologically active retinoids were found in water samples from severely affected sites. Interestingly, a recent survey of 5 264 hylid and ranid metamorphs in 42 Vermont wetlands demonstrated that proximity to agricultural land was associated with an increase in limb malformations (Taylor et al. 2005). No analysis of the mechanisms underlying these abnormalities have yet been carried out (Gardiner et al. 2003). It seems likely, however, that retinoic acid mimics could modulate Hox gene expression and induce a variety of abnormalities, depending on the developmental stage and length of time of exposure. C. Reproduction

Reproductive organs produce a variety of hormones including steroids such as testosterone, estrogen, and progesterone, as well as peptide hormones such as inhibin, activin, and Miillerian inhibiting hormone, so they function as endocrine glands and they appear to be a major target for EDCs. Several environmental chemicals alter the development and function of the reproductive system, thereby altering endocrine system function. These affects have been documented across all classes of vertebrates including humans (reviewed by Milnes et al. 2006). Although the mechanisms that drive sex determination and gonadal development appear to vary widely among species, the underlying genetic and endocrine control of gonadal development, growth, and function, as well as the roles of hormones on secondary sexual characteristics, are highly conserved among vertebrates. Most EDCs studied to date exhibit estrogenic or anti-androgenic activity and can influence sex ratio, sexual maturation, gonadal morphology and function, as well as spermatogenesis, fertility, hormone levels (steroidogenesis, metabolism), secondary sexual characteristics and reproductive behaviour (Milnes et al. 2006). Clearly, all of these characteristics are important for maintaining healthy and persistent populations, especially for taxa that are known to be declining globally, such as amphibians. 1 . Effects of EDCs on the Amphibian Reproductive System

The effects of EDCs on the development and functioning of the reproductive system of amphibians are not well understood relative to other wild life such as fish, alligators, and mammals. Amphibians, however, could be especially sensitive to EDCs because gonadal differentiation in this taxon is highly sensitive to sex hormones and occurs over an extended period of time, sometimes beginning prior to metamorphosis and lasting for several months (Hayes 1998; Qin et al. 2003). In genetic males, estrogen can induce a complete and permanent sex reversal or induce development of intersexed gonads (ovarian tissue and testicular tissue in the same gonad). Intersexed gonads typically occur in two forms: oocytes can be scattered throughout the testes forming an ovo-testis, or the ovary can be distinct and well delineated from the testicular tissue (Milnes et al. 2006). These different phenotypes likely occur through different molecular mechanisms and/or timing of exposure, but these details have not yet been fully investigated. Although the data on the effects of EDCs on the reproductive system of amphibians are relatively limited, it has been clearly demonstrated that EDCs can induce female-skewed sex ratios and alter gonadal development and concentrations of hormones. Tavera-Mendoza et al. (2002a,b) exposed Xenopus laevis tadpoles to ecologically relevant levels (21 pg/L) of atrazine for 48 hours during sexual differentiation. Histological analysis of the ovaries showed that atrazine-exposed females experienced a 20% reduction in primary germ cells compared to 2% in controls. Atrazine-exposed males had a 5'7% reduction in testicular

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUFTING CHEMICALS

3223

volume, and primary spermatogonial cell nests were reduced by 70%. Furthermore, Sertoli (nurse) cells, which provide nutritive support for the developing germ cells, declined by 74% and testicular resorption occurred in 70% of the males, while 10% failed to filly develop testes. These findings are significant because primary germ cells are thought to comprise the total number of germ cells for the entire reproductive life of the organism, and such a remarkable reduction in only 48 hours suggests that atrazine, even in pulsed exposures, could severely negatively affect reproductive success of both females and males (TaveraMendoza et al. 2002a; Tavera-Mendoza et al. 2002b). As mentioned above, African clawed frogs (&no@ laevk) exposed to environmentally relevant doses of ammonium perchlorate during metamorphosis had female-skewed sex ratios (Goleman et al. 2002). Several other chemicals, however, have been demonstrated to induce female-biased sex ratios. For example, Kloas et al. (1999) exposed X. laevk tadpoles to estradiol (positive control), nonylphenol (used in surfactants, detergents, and pesticides), bisphenol A (plastic or plasticizer), octylphenol (surfactant) and butylhydroxyanisol (pharmaceutical for treating retroviruses). Each chemical induced a significantly higher number of females compared to controls. These results demonstrate that a diverse array of chemicals can, and do, alter sex ratios making them female-biased. There are two ways in which this type of bias can occur. Males could, in general, be less resistant to chemicals and suffer higher mortality than females. Alternatively, the genotypic males in these experiments may undergo sex reversal so that they appear female.

Many chemicals have been shown to induce feminization of male gonadal tissue. Qin et al. (2003) exposed X. laeuis to Aroclor 1242 and Aroclor 1254 (aroclors are mixtures of PCBs) and recorded both gross morphological and histological differences relative to the vehicle control. Control animals had normal ovaries or testes from the standpoint of gross morphology whereas PCB-exposed animals had abnormally sized and asymmetrical testes as well as ovotestes. Ovotestes, in that study, were characterized by ovaries in the cranial andlor caudal portion of the gonad with testes located medially. PCB exposure did not alter the proportion of females across treatments but the portion of males with morphologically normal testes was reduced. Histological examinations revealed that testes that were classified as morphologically normal, as well as those classified as abnormal upon gross evaluation, had oocytes interspersed throughout the tissue. In addition, testes of PCBexposed animals were more loosely organized and had fewer seminiferous tubules, spermatogonia, and spermatozoa than was the case for controls. Qin et al. (2003) argued that Aroclor 1242 and Aroclor 1254 feminize gonadal differentiation in X. laeuis and that X. laeuis could be especially sensitive to endocrine disruption. Thus, this species is an appropriate model for studying endocrine disruption. Hayes et al. (2002) showed that very low ecologically relevant levels of the common herbicide atrazine can induce X. laeuis tadpoles to develop intersexed gonads and exhibit decreased size of the larynx (demasculinized), and they suggested that these changes could reduce fertilization success and alter breeding call characteristics. Indeed, X. laeuis, collected from agricultural areas in South Africa where atrazine and other chemicals are used, suffer decreased testosterone levels (Hecker et al. 2004). Importantly, atrazine is a widespread compound and the concentrations of atrazine used in that study are realistic levels (e.g., "acceptable" for drinking water according to the EPA) to which many amphibian species are exposed in the wild, putting them at risk of impaired sexual development. Indeed, much work has been conducted on X. laeuis and this species has proven to be an important model since it is easy to obtain, breed, and rear in the laboratory. Several studies, however, have documented feminization in amphibian species other than X. laeuis. For example, leopard frogs ( R a m pipiens) exposed to ecologically relevant concentrations of atrazine (>0.1 ppb) had retarded gonadal development (gonadal dysgenesis) and testicular oogenesis (intersexed). Males that developed slowly and were thus exposed to atrazine for longer periods of time, also experienced oocyte growth (oogenesis and vitellogenesis) (Hayes et al. 2003). Hayes et al. (2003) also conducted field surveys of sites

3224

AMPHIBIAN BIOLOGY

that varied in atrazine exposure across the United States and observed gonadal dysgenesis and intersexuality in animals from sites contaminated with atrazine. They concluded that these integrated laboratory and field studies on leopard frogs, coupled with similar findings in X. laevis, demonstrate the effects of atrazine on amphibians in general and suggest that atrazine and other endocrine-disrupting pesticides could be playing a major role in amphibian declines around the world. Another important group of contaminants that is commonly found in the environment due to its importance to humans is plastic that contains bisphenol A or plasticizers (various phthalates) that give plastic its pliability. Bisphenol A has been shown to induce female skewed sex ratios in X. Zuevis (Mosconi et al. 2002). Ohtani et al. (2000) exposed genetically male tadpoles of Rana rugosa to dilute solutions of the plasticizer dibutyl phthalate and three 17P-estradiol-positivecontrols for four days during gonadal sexual differentiation (days 19-23). At day 40, the gonads of the tadpoles were examined histologically. Positive control and dibutylphthalate-treated, genetically male tadpoles had complete ovaries or were intersexed in a dose-dependent manner. Therefore, dibutyl phthalate was able to induce intersexuality, but it was approximately 1000-fold less potent than 17P-estradiol (Ohtani et al. 2000). Even considering the reduced potency, Ohtani et al. (2000) concluded that dibutyl phthalate is an estrogen-like hormone that alters testicular differentiation, and thus is a dangerous environmental contaminant. It is known that gonadal development in many amphibian species can be altered by exposure to estrogens or to estrogenic contaminants (Hayes 1998). Therefore one would expect larvae living in environments containing sewage effluent that is contaminated with human birth-control chemicals to have altered gonadal development. To test this hypothesis, Park and Kidd (2005) added low concentrations of 17a-ethinylestradiol (EE,) to an experimental lake in northwestern Ontario, Canada. A target concentration of 5 ngIL was maintained by adding 17a-ethinylestradiol to the treatment lake three times a week during the open-water seasons (May to October) for three years (2001, 2002, and 2003). Egg masses were reared in cages at the EE, lake and in two reference lakes. In the EE2 lake, hatching success was reduced sigdicantly in green frogs (Ram clamitans) but not in mink frogs (Ram septentrim2uli-s). relative to each species in the reference lakes. Ethinylestradiol had no effect on sex ratios of either species and no intersex gonads were observed in tadpoles of either species at the reference sites. Although no green frog tadpoles were intersexed at the EE2 site, approximately 6% and 13% of the caged mink frog tadpoles were intersexed in 2001 and 2002 respectively. Therefore, these two species responded differently to EE, exposure. Green frogs suffered higher mortality at hatching but survivors did not have intersexed gonads, whereas mink frogs did not have high hatchling mortality but did experience altered gonadal development. This highlights the fact that different species respond differently to the same environmental contaminant. Un-caged mink frog tadpoles were also studied at each of the sites, and EE, had no effect on sex ratios, but by the third year 28.6% of wild EE2-exposedfirst-year tadpoles had intersexed gonads whereas none were intersexed in the reference lakes. These results indicate that concentrations of l7a-ethinylestradiol, comparable to those found in sewage treatment effluents and some surface waters, can affect hatching success as well as gonadal development in native amphibians (Park and Kidd 2005). In addition to intersexed or completely feminized gonads, alterations in circulating steroid concentrations have been reported in amphibians exposed to EDCs. For example, Hayes et al. (2002) exposed adult African clawed frogs to 25 ppb atrazine (level acceptable in drinking water according to the EPA) and found that they exhibited a 10-fold decrease in testosterone concentrations relative to animals housed in 0 ppb. They hypothesized that atrazine increased aromatase, the enzyme that converts testosterone to estrogen, and caused the decrease in testosterone. As discussed above, this mechanism is known to occur in other vertebrate species (Sanderson et al. 2000, 2002; Sanderson 2006; Fan et al. 2007). Altered gonadal development and hormone concentrations associated with EDC exposure induce other abnormal physiological responses. One important example of this

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUFTING CHEMICALS

3223

is the induction of the normally female-specific protein vitellogenin (yolk protein) in males. Typically males do not express the vitellogenin gene but when they are exposed to estrogen or estrogenic chemicals this gene is upregulated. Therefore, plasma vitellogenin in males is a biomarker for estrogenic xenobiotics. For example, adult male red-eared slider turtles (Trachemys scripta) and African clawed frogs given injections of estradiol, the synthetic estrogen diethylstilbestrol (DES), or the insecticide DDT experienced an induction of vitellogenin. In both species, estradiol and DES treatments induced more vitellogenin than did DDT, which demonstrates that DDT is estrogenic, but as expected from previous studies investigating its estrogenicity, it does not elicit as strong a response as do actual estrogens (Palmer and Palmer 1995). Since that study, and many others establishing vitellogenin as a reliable biomarker, researchers have begun to use vitellogenin to detect estrogen exposure in amphibians (Palmer et al. 1998; methodology reviewed by Wheeler et al. 2005). Mosconi et al. (2002) reported a dose-dependent induction of vitellogenin in male European frogs (Rana esculenta) and crested newts (Triturus carnqex) exposed to the surfactant 4-nonylphenol. They also demonstrated that 4-nonylphenol inhibited gonadotropin release from the hypothalamus and prolactin secretion by the pituitary, but increased plasma androgen concentrations. Importantly, the metabolic cost associated with production and elimination of vitellogenin and its impacts on reproductive success of males have not been considered or investigated in detail (Milnes et al. 2006). Other physiological responses that can be altered when EDCs are misinterpreted as hormonal signals involve secondary sexual traits. These include many sexually dimorphic characteristics such as differences in forelimb size, or coloration, or the presence of nuptial pads. For example, female colouration in the reed frog (Hyperolius argus) is estrogen dependent, so Noriega and Hayes (2000) used early induction of female colour pattern as a biomarker for estrogenic activity. H. a r p tadpoles were exposed to the insecticide o,pDDT and six of its congeners and in vivo colour changes were compared among treatments. Estradiol, o,p-DDT, o,pl-DDE and o,p'-DDD prematurely induced adult female colouration in juvenile animals, whilst p,p'-DDT, p,pl-DDE and p,p'-DDD did not (Noriega and Hayes 2000). This concept that estrogen-dependent coloration could be used as a biomarker of estrogenic chemicals was utilized by McCoy et al. (unpubl. data) during field surveys of giant toads (Bufo marinus) living in sugarcane agricultural areas in South Florida. It was found that as many as 40% of the males (defined as having testes and nuptial pads) were intersexed (also had ovaries) and many males, including individuals with morphologically normal testes, exhibited female coloration, whereas those from reference sites were not intersexed, and exhibited the sexually dimorphic male colour pattern. D. Behaviour The endocrine system and the central nervous system (CNS) are integrated such that the proper functioning of one is dependent upon the proper hnctioning of the other. For example, particular regions of the brain are known to be sexually dimorphic and these differences occur in response to different hormonal influences during development. Having the proper hormonal milieu at crucial stages in brain organization is essential for future (adult) sex-specific behavioural responses (Palanza et al. 2002; Moore et al. 2005). Altering this hormonal milieu during fetal development can permanently change adult behaviour. The hypothalamus links the nervous and endocrine systems by releasing stimulating or inhibiting hormones through blood vessels to the anterior pituitary which in turn signals endocrine glands such as the thyroid, adrenal, or gonad via tropic (releasing) hormones (Fig. 1). These endocrine glands can then synthesize and secrete specific hormones that induce various physiological effects and provide feedback that alters pituitary and hypothalamic (CNS) function. Since these systems are integrated, hormonal effects on behaviour (CNS) can be direct, or indirect (e.g., altering thyroid hormones can influence metabolism that in turn influences behaviour) (Zala and Penn 2004). The endocrine and nervous (neuroendocrine) systems are so highly integrated that chemicals which alter the endocrine system also influence the central nervous system and

3226

AMPHIBIAN BIOLOGY

therefore behaviour. There are numerous diverse examples of endocrine disruption leading to alteration of behaviour; thus, Zala and Penn (2004) suggested that endocrine disrupting chemicals be renamed as "neuroendocrine disrupters". Indeed, behaviour is becoming a more common endpoint in standard toxicological assays. Many researchers suggest that behaviour is a useful bioindicator of endocrine disruption because it is an easily measured, integrated physiological response that is related to alterations of specific neural pathways and is dependent on environmental context (Clotfelter et al. 2004; Panzica et al. 2005; Crews et al. 2007). An animal's behaviour is indicative (a surrogate or sentinel) of its health and the tools needed to evaluate behaviour are relatively inexpensive and easy to implement. Many of the physiological mechanisms that control behaviour, however, are not understood in detail, so it can be difficult to determine the mechanism(s) through which specific EDCs modulate behaviour. Thus, there is some debate about the usefulness of behaviour as a measure of endocrine disruption (Clotfelter et al. 2004). If one cannot ascribe an endocrine mechanism, how can one know that disruption of the endocrine system is leading to the altered behaviour? Understanding how EDCs alter the mechanisms controlling and modulating behaviour is an extremely important and open field of exploration. Although the mechanisms of endocrine or neuroendocrine disruption leading to altered behaviour are not always clear, there are numerous examples of known EDCs that induce alterations in behaviour. Recent reviews by Zala and Penn (2005) and Clofelter et al. (2005) demonstrated that locomotion, balance, feeding, antipredator behaviour, communication, aggression, various aspects of learning, and reproductive behavior (including courtship, mate choice, mating, nesting, and parental care) can all be altered by chemicals that are known to disrupt the endocrine system. 1. Effects of EDCs on Amphibian Behaviour

Many aspects of amphibian behaviour are altered by chemical pollutants including locomotion, activity, exploratory behavior, response to cues emanating from predators, communication, learning, and mating. The following discussion is restricted to examples describing how known EDCs alter behaviour in amphibians. PCB 126 (insulating fluid for transformers and capacitors) has been shown to reduce swimming speed in tadpoles of Ram clamitans and R. pipiens, which suggests that both feeding and predator escape could be affected in natural systems; these ecological consequences, however, were not tested (Rosenshield et al. 1999). Bridges (1999) explicitly tested whether responses by tadpoles to predators were altered in the presence of the insecticide carbaryl. Tadpoles of Hyla versicolor were exposed to two ecologically relevant levels of carbaryl (1.25 and 2.50 mg/L), the vehicle control, and a negative control (acetone solvent and water respectively) for only 24 hours. After the exposure, tadpole behaviour was examined in the presence and absence of adult red-spotted newts (Notophthalmus virzdescens) housed in a mesh container. The containers allowed visual, tactile, and chemical cues to be transmitted but did not allow predatory attack. Tadpole activity, measured as percentage of time spent swimming, resting, and feeding and time spent in refugia, was recorded every three minutes for one hour (Bridges 1999). Control tadpoles spent 20% of their time in refugia in the presence of a predator, but did not enter refugia when no predator was present; instead they actively fed. Animals in the lowcarbaryl treatment spent roughly equal amounts of time in refugia whether a predator was present or not, whereas those in high-carbaryl treatment spent less time in refugia when the predator was present than when it was absent. In addition, tadpoles exposed to high levels of carbaryl were, on average, less active than were those in control treatments but they spent more time foraging in the presence of a predator than in its absence. Control tadpoles in Bridges' (1999) experiment were able to effectively detect predators, hide in their presence, and actively forage in their absence. In other words, they behaved adaptively. The tadpoles exposed to carbaryl, however, did not show these adaptive behaviours, especially at the high carbaryl concentration. This study clearly demonstrates that EDCs can alter behaviour in unexpected ways.

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUPTING CHEMICALS

3227

In a similar study, salamander larvae (Ambystoma mcroductylum) exposed to methoxychlor (replacement insecticide for DDT) exhibited a reduced startle response and traveled shorter distances in response to disturbance; they also experienced increased predation by dragonflies (Verrell 2000; Eroschenko et al. 2002). Importantly, this was not the first time that exposure to a known EDC induced behaviour that increased predation. Three decades earlier, Cooke (1970, 1971) showed that common frog tadpoles (Rana temporaria) exposed to DDT were uncoordinated and hyperactive and were preferentially preyed upon by warty newts (Triturus cristatus). In addition to the increased predation that prey experience after chemical exposure, predators could suffer increased bioaccumulation if they preferentially eat exposed individuals (Cooke 1970, 1971). The occurrence of maladaptive behaviour in the presence of EDCs suggests that learning could be altered by exposure to such chemicals. To examine this hypothesis, Steele et al. (1999) exposed tadpoles of Rana clamitans and R. catesbeiana to 0 or 750 microg Pb/L for 5-6 days; then, animals were conditioned to associate a light source (conditioned stimulus) with a mild shock (unconditioned stimulus). Tadpoles exposed to lead had a higher mean response time and showed less avoidance. It was concluded that sublethal exposure to lead adversely affects acquisition learning and retention in tadpoles (Steele et al. 1999). In addition to maladaptive antipredator responses, feeding, and learning behaviour, exposure to EDCs also has been associated with altered communication and reproductive behaviour in amphibians. For example, Park and Propper (2002) exposed male red spotted newts (Notophthalmus viridescens) to ecologically relevant levels of the insecticide endosulfan or to an acetone carrier control for four days. After the exposure period, olfactory response tests were run in a Y-maze in which each arm contained a female used to attract males. Individuals exposed to the low concentration of endosulfan took longer to respond to the presence of females. Morphological measurements of pheromone glands demonstrated that endosulfan decreased the alveolar and luminal area. Park and Popper (2002) also studied the mating success of individuals from endosulfan treatments by placing males with a female and recording the presence of spermatophores after 24 hours. They found that males from endosulfan treatments had dramatically lower mating success and concluded that environmental chemicals may lead to population declines in amphibians by disrupting chemical communication and mating behaviour (Park and Propper 2002). 2. Effects of EDCs on Non-Amphibian Behaviour The effects of EDCs on pheromonal systems are not restricted to amphibians and have been documented in diverse taxa. For example, exposure to low ecologically relevant concentrations of the insecticide cypermethrin reduced or inhibited the olfactory response of mature male Atlantic salmon (Salmo salar) to a priming pheromone released by ovulating females, and resulted in decreased levels of plasma sex steroids and expressible milt (Moore and Waring 2001). Other studies have documented that EDCs also affect chemosensory communication in mammals. For example, vom Saal et al. (1995) fed low doses of the insecticides DDT and methoxychlor to pregnant mice and investigated the scent-marking behaviour of their male progeny as adults. Male mice exposed in utero showed a dosedependent increase in territorial scent-marking as adults (vom Saal et al. 1995). These results indicate that exposure to pesticides such as DDT and methoxychlor can modulate brain development such that social behaviour of adults is altered. Reproductive behaviour is especially important to document since it is directly tied to population health and persistence, and there are many examples of EDCs that alter behaviours such as courtship, mating, and parental care. Howell et al. (1980) compared mosquitofish, (Gambusia afinis holbrooki) from 11 sites that varied in exposure to paper mill diluent. Females from five geographically distinct sites downstream of a paper mill had anal fins that were modified into gonopodia, the intromittent organ of males. Many young males were precociously mature at these sites as well. No masculinized females or precociously mature males were collected upstream of the paper mill, in tributaries, or in

3228

AMPHIBIAN BIOLOGY

nearby ponds (Howell et al. 1980). Howell and colleagues continued their studies in the laboratory, and found that precociously masculinized males displayed male-typical courtship but were more aggressive than were normal control males. The masculinized males were dominant over the normal males, and were able to court normal females without competition. In addition, masculinized females displayed male reproductive behaviour. They chased the control females and attempted to mate by performing gonopodial thrusts. Clearly this type of behaviour could reduce mating success in the wild. In a more recent study in Florida comparing mosquitofish from sites contaminated by papermills and those that were not contaminated, females from contaminated sites were smaller, fewer were gravid, and their estradiol concentration exhibited greater variation (Toft et al. 2004). Indeed, several pesticides, including the fungicides vinclozolin and fenarimol, the insecticides DDT (and its metabolites) and methoxychlor have been shown to alter copulato~y behaviour such as intromission and normal ejaculation, erectile functions, and mounting behaviour (Zala and Penn 2004). For example, exposure to methoxychlor during development inhibits the normal female-induced elevations of testosterone in adult male mice and quail and leads to reduced sexual interest and copulatory behaviour; similar to the effects in salmon mentioned above (Eroschenko et al. 2002). However, when methoxychlor was administered to male rats from weaning age or later, it reduced feeding rate and growth, delayed puberty, testicular testosterone production, epididymal size and sperm numbers, but increased number of attempts to copulate and promoted early ejaculation (Ostby et al. 1999). These results clearly show that the timing of exposure can determine the types of effects that are induced by EDCs, and other contaminants. VII. IMPLICATIONS A. Conservation and Ecology Contaminants are distributed globally; even areas once believed to be pristine are affected by run off, wind blown contaminants, or fallout through precipitation (Thurman and Cromwell 2000; Sparling et al. 2001; Davidson and Knapp 2007). Importantly, many amphibians are very sensitive to contaminants and are exposed throughout their life history. For instance, they have highly permeable skin as adults and larvae. Many reproduce in aquatic environments affected by runoff or precipitation, so adults can be exposed throughout their lives, in transit to breeding sites, and while breeding. Their larvae experience critical hormone-regulated developmental stages while in these potentially polluted environments, making them particularly susceptible to endocrine disruption. Living in, and around, contaminated areas and undergoing critical stages of development while exposed to endocrine-disrupting chemicals is expected to negatively affect individuals as well as populations (Hayes et al. 2006b; but see Glennemeier and Begnoche 2002).

The link between pollutants and amphibian declines has, in some cases, been weak, occurring as a post hoc hypothesis after declines have occurred. For example, the last remaining natural populations of Wyoming toads were known to be exposed to fenthion, an organophosphate insecticide used for mosquito control (Lewis et al. 1985). Importantly, fenthion is known to block androgen receptors (in mammals) and androgen-dependent physiological processes (Tamura et al. 2001). Carey and Bryant (1995) speculated that this exposure could have been involved in the ultimate extinction of wild Wyoming toads (Bufo hemiophrys baxteri). Male toads in the last population showed reduced clasping behaviour during breeding, which is likely an androgen-dependent behaviour, and hatchability of the few fertilized clutches was low (Carey and Bryant 1995). Appropriate studies to evaluate the effects of fenthion on Wyoming toad populations were not conducted because the connection between the contaminant and the population extinction was made too late. Although Wyoming Toads have been reintroduced, they are not currently self sustaining, and research on these "wild" populations is hampered by the scarcity of the toads (Dreitz 2006). Importantly, however, several strong lines of evidence suggest that pollution induces population declines and extinctions of amphibians. For example, population declines of

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUPTING CHEMICALS

3229

several amphibian species are associated with wind-borne agricultural chemicals (Sparling et al. 2001; Davidson and Knapp 2007). In an important and creative study, Sparling et al. (2001) used altered cholinesterase (ChE) activity to demonstrate that exposure to wind-borne pesticide was associated with reduced population status in the Pacific Tree Frog (Hyla regzlla). Cholinesterase (ChE) is an enzyme that rapidly clears the neurotransmitter acetylcholine from neuronal synapses. The clearance of acetylcholine modulates neuron-to-neuron communication and is essential for proper muscle function. Many insecticides and some herbicides that are typically sprayed in agricultural areas contain pyrethoids and organophosphates. These chemicals are known to inhibit ChE activity and reduced ChE activity is a biomarker of exposure to these pesticides. Sparling et al. (2001) found that cholinesterase (ChE) activity in H. regilla tadpoles from mountainous areas downwind of the highly agricultural Central Valley, where the most severe amphibian population declines in California have occurred, was depressed compared to those from sites on the coast or north of the valley where declines are less precipitous (Sparling et al. 2001). In addition, as many as 86% of the populations in areas with reduced ChE had detectable levels of endosulfan, up to 50% had measurable organophosphorus levels, and 40% carried DDT residues. This study clearly establishes an association between pesticide exposure, physiological response to that exposure, and poor population status. Although a cause-and-effect relationship is d&cult to establish with field studies that use such epidemiological approaches, these data are highly suggestive of such a relationship. An alternative hypothesis that could explain population declines in these areas of California is that introduced non-native fish have increased mortality of tadpoles and driven populations to decline. Therefore, Davidson and Knapp (2007) studied the relative effect of introduced fish and pesticides on the mountain yellow-legged frog (Rana muscosa). They conducted an enormous set of field surveys that included 6831 sites studied over a sevenyear period (1995-2002). At each site, they quantified habitat characteristics and presence or absence of R. muscosa and fish. They calculated windborne pesticide exposure for each site from pesticide application records and predominant wind directions. The probability of R. muscosa presence was significantly reduced by the presence of fish and pesticides. However, the effect of pesticides at the landscape scale was much stronger than the effects induced by fish (Davidson and Knapp 2007). Importantly, the degree of protection from windborne pesticides was also a significant predictor of R. muscosa presence. These results demonstrate that windborne pesticides are contributing to amphibian declines in "pristine" locations (Davidson and Knapp 2007). Many agricultural contaminants are known to function as endocrine disrupting chemicals that induce effects at very low concentrations. California's amphibian populations could be suffering declines not because they are exposed to high quantities or lethal concentrations of contaminants, but because they are exposed to very low concentrations of hormonally active agents that alter thyroid function, development, behaviour, and reproduction.

These contaminant-associated declines are not surprising given what is known about EDCs, but it is important to note that few studies have investigated the population-level effects of chronic exposure to pollution, nor have any, to our knowledge, explicitly modeled the way in which EDCs could be involved in global amphibian decline. Chemical pollutants, including EDCs, are expected to play an important role in modulating population dynamics because they can directly induce mortality and, at sublethal levels, they influence growth and development, decrease the ability of larvae to avoid predators, and alter reproductive success (Carey and Bryant 1995). In addition, because the immune system interacts directly with the endocrine system, EDCs can affect susceptibility to pathogens (Carey and Bryant 1995; Hayes et al. 2006b). Importantly, EDCs could contribute to recent outbreaks of infectious disease in amphibians, especially when they occur in complex mixtures as is typical of contaminated environments. For example, Hayes et al. (2006) studied the effects of nine different agricultural pesticides used on cornfields in the Midwestern United States by exposing

3230

AMPHIBIAN BIOLOGY

tadpoles of Rana pipiens to very low concentrations (0.1 ppb) of each chemical, separately or in various combinations, and measuring differences in growth, development, gonadal differentiation, and immune function. Many of the single chemical treatments led to differences in larval growth and development but the pesticide mixtures had more severe effects. Pesticide mixtures damaged the thymus which was associated with immunosuppression and the contraction of flavobacteria meningitis (gram negative bacteria). This disease left the animals debilitated; they were in poor condition and unable to hold themselves upright. Interestingly, the control animals tested positive for the presence of this bacteria but did not show signs of the disease (Hayes et al. 2006). This type of interaction between contaminants and disease agents, making the disease more pathogenic, has been hypothesized for many years (Carey and Bryant 1995). Another recent study has investigated the influence of EDCs on disease susceptibility by exposing tiger salamanders (Ambystoma tigrinum) to atrazine and measuring peripheral leukocyte levels and susceptibility to Ambystoma tig.rznum virus (ATV), a pathogen implicated in some amphibian die-offs (Forson and Storfer 2006). Atrazine significantly decreased peripheral leukocyte levels and increased susceptibility of the larvae to ATV infection, illustrating that atrazine influences the immune system. Interestingly, there were familial differences in infection rates, making particular genotypes more (or less) sensitive. Over all, this study shows that in addition to its effects on testicular development, ecologically relevant concentrations of atrazine can have immunosuppressive effects that could contribute to ATV epizootics (Forson and Storfer 2006). B. Evolution Another important consequence of EDC exposure is that it can alter the evolutionary trajectory of a species. A classic example of pollutant-induced evolutionary change is the darkening of the peppered moth (Biston betularia) in response to the darkening of trees by soot in England during the industrial revolution. There are also many taxa that have evolved pesticide resistance. Pollutants can induce strong selective forces by increasing mortality and altering genetic variability. Sublethal levels of pollutants can also influence evolutionary processes. For example, intergenerational transfer of contaminants (through egg yolk or mother's milk) can alter development, reproductive physiology, metabolism, and behaviour in maladaptive ways (Fox 1995). In addition, many recent experiments have demonstrated that gene imprinting and induction of epigenetic modifications through methylation is controlled, at least is some cases, by estrogen or other hormones and can be modulated by EDCs such as methoxyclor. Therefore EDCs can induce persistent, heritable phenotypic changes, independent of mutagenesis. These phenotypic variants will be subject to selection and the evolutionary trajectory of affected species is expected to be altered. Evolutionary changes are in many cases irreversible and they impart an environmental legacy that is broader than that of the pollutant itself (Leblanc 1994). VIII. WHAT NEEDS T O BE KNOWN

Hayes et al. (2006) reported that fewer than 30 published laboratory and field studies have addressed low concentration, endocrine-disrupting effects on amphibians. A few studies have examined the effects of contaminant exposure on wild amphibian populations (Ouellet et al. 1997; Sparling et al. 2001; Davidson and Knapp 2007) but establishing cause and effect in such studies is difficult, and describing physiological alterations inducing effects at the population level is daunting. Regardless of these difficulties, understanding the population-level effects of contaminants on amphibians and the role of EDCs in global amphibian decline is extremely important, albeit understudied. For example, although several studies have demonstrated that some chemicals can induce feminization of males and intersexuality, little work has focused on the population-level effects of male feminization; this problem is not restricted to amphibians (Hayes et al. 2006a).

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUFTING CHEMICALS

3231

Many studies on the effects of contaminants such as EDCs have been conducted in controlled laboratory settings. Although these studies are extremely important, and necessary, for determining the mechanisms of toxicity, they do not even attempt to replicate natural systems. For example, many studies investigate the effects of chemicals at concentrations much higher than individuals experience in the wild. Reported effects from these studies do not help explain whether exposed populations will suffer similar effects because the mechanisms of toxicity of EDCs can depend on concentration. A number of studies have documented inverted U-shaped dose responses to EDCs; that is, low to intermediate concentrations can produce larger effects than high doses due to the integrated nature of physiological responses, and the presence of negative feedback mechanisms. For example, survival was significantly lower for four different amphibian species when they were exposed to 3 ppb of atrazine compared with either 30 or 100 ppb and these mortality patterns depended on tadpole stage (Storrs and Kiesecker 2004). Therefore, the deficiency in studies on the effects of lower, ecologically relevant concentrations of EDCs in amphibians has likely resulted in an underestimation of the impacts of these pesticides (Carey and Bryant 1995; Hayes et al. 2002, 2006a). In addition, wild amphibian populations are exposed to complex mixtures of contaminants and other stressors, such as UV radiation, but most studies have investigated the effects of single contaminants without considering other environmental variables (Hayes et al. 2002; Blaustein et al. 2003; Sullivan and Spence 2003). Contaminant concentrations also fluctuate drastically in natural systems, and these fluctuations can induce unexpected and unpredictable effects, but few studies have investigated fluctuating contaminant exposures (Edwards et al. 2006). Furthermore, most studies focus on effects of contaminants during, but not after, exposure. This can lead to underestimations of the effects of contaminant because survival of later life-history stages (e.g., adults) could be negatively affected whereas earlier, exposed, stages are not (Harris et al. 2000; Rohr and Palmer 2005; Rohr et al. 2006b). More realistic exposures that replicate wild systems, and investigate effects across amphibian life history will help describe the effects that contaminants have on amphibian populations. Questions that attempt to link individual-level effects induced by a contaminant in the laboratory to effects in the field are difficult and require integration across broad fields that have traditionally been isolated from one another. Although recent studies have begun to address these issues, it is not clear how the mechanisms of toxicity or the effects of chemical pollutants change relative to competition, predation, alterations in animal density, food quantity, or the presence of pathogens (Relyea and Mills 2001; Rohr et al. 2004; Forson and Storfer 2006; Rohr et al. 2006b; Romansic et al. 2006). Knowing how ecological factors influence an organism's response to chemicals will help predict the effects of pollutants in the wild, and will advance an understanding of how to restore polluted habitats (Rohr et al. 2006a). The phenomenon of endocrine disruption has been focused on documenting estrogenic, androgenic, antiandrogenic, and antithyroidal actions of specific chemicals, and these pathways of toxicity have now been established in numerous vertebrate species (Guillette 2006). Many potential mechanisms of endocrine disruption, however, have not been studied. For example, little work has been conducted on the effects of EDCs on other steroid pathways, such as those including the progestins (e.g., progesterone) and glucocorticoids (e.g., the stress hormone corticosterone) or on the retinoids (Berube et al. 2005; Guillette 2006; Hinson and Raven 2006; Leiva-Presa and Jenssen 2006). Importantly, it is known that progesterone is important for oocyte maturation in amphibians and that EDCs, such as methoxychlor, disrupt progesterone-induced oocyte maturation in Xenopus leavis (Pickford and Morris 1999, 2000, 2003; Guillette 2006). In addition, the adrenal is thought to be the most common toxicological target organ for EDCs (Harvey et al. 2007) but relatively little work has focused on understanding the effects of EDCs on the stress axis (hypothalamus-pituitary-adrenals) (Pottinger 2003). Evaluation of the adrenals are also neglected in EDC screening and testing for regulatory purposes (Harvey et al. 2007). Finally, retinoid imbalances are associated with multiple effects, including changes in secondary

3232

AMPHIBIAN BIOLOGY

sexual characteristics, inhibition of spermatogenesis, and decreased embryo survival. Importantly, retinoid homeostasis is affected by several contaminants (Berube et al. 2005). Future studies on endocrine disruption must broaden current approaches. The mechanisms and endocrine end points examined need to be expanded, and conduct studies that allow evaluation of how organisms are affected in wild habitats need to be conducted (Guillette 2006). IX. REFERENCES Andres, A. C., Muellener, D. B. and Ryffel, G., 1984. Persistence, methylation and expression of vitellogenin gene derivatives after injection into fertilized-eggs of Xenopus laevis. Nuc. Aczds Res. 12: 2283-2302.

Blaustein, A. R., Romansic, J. M., kesecker, J. M. and Hatch, A. C., 2003. Ultraviolet radiation, toxic chemicals and amphibian population declines. Divec Dist. 9: 123-140.

Ankley, G. T., Jensen, K. M., Durhan, E. J., Makynen, E. A., Butterworth, B. C., Kahl, M. D., Villeneuve, D. L., Linnum, A., Gray, L. E., Cardon, M. and Wilson, V. S., 2005. Effects of two fungicides with multiple modes of action on reproductive endocrine function in the fathead minnow (Pimephales promelas). Toxicol. Sci. 86: 300-308.

Block, K., Kardana, A,, Igarashi, P and Taylor, H. S., 2000. In utero diethylstilbestrol (DES) exposure alters Hox gene expression in the developing Mullerian system. FASEB J. 14: 1101-1 108.

Ankley, G. T, Jensen, K. M., Makynen, E. A, Kahl, M. D., Korte, J. J., Hornung, M. W., Henry, T R., Denney, J. S., Leino, R. L., Wilson, V. S., Cardon, M. C., Hartig, F! E. and Gray, L. E., Jr., 2003. Effects of the androgenic growth promoter 17P-trenbolone on fecundity and reproductive endocrinology of the fathead minnow. Environ. Toxicol. Chem. 22: 1350-1360.

Boas, M., Feldt-Rasmussen, U., Skakkebaek, N. E. and Main, K. M., 2006. Environmental chemicals and thyroid function. Euro. J. Endocrin. 154: 599-61 1.

Blumberg, B. and Evans, R. M., 1998. Orphan nuclear receptors - new ligands and new possibilities. Gene Develop. 12: 3149-3 155.

Borga, K., Gabrielsen, G. W. and Skaare, J. U., 2003. Comparison of organochlorine concentrations and patterns between free-ranging zooplankton and zooplankton sampled from seabirds' stomachs. Chemosphere 53: 685-689.

Anway, M. D., Cupp, A. S., Uzumcu, M. and Skinner, M. K., 2005. Epigenetic transgenerational actions of endocrine dismptors and mate fertility. Science 308: 1466-1469.

Bridges, C. M., 1999. Effects of a pesticide on tadpole activity and predator avoidance behavior.J. Herpetol. 33: 303-306.

Arnold, S. F., Collins, B. M., Robinson, M. K., Guillette, L. J., Jr. and Mclachlan, J. A., 1996. Differential interaction of natural and synthetic estrogens with extracellular binding proteins in a yeast estrogen screen. Steroids 61: 642-646.

Burnison, B. K., Hartmann, A., Lister, A., Servos, M. R., Ternes, T and Van Der Kraak, G., 2003. A toxicity identification evaluation approach to studying estrogenic substances in hog manure and agricultural runoff. Environ. Toxicol. Chem. 22: 2243-2250.

Ballestar, E. and Esteller, M., 2002. The impact of chromatin in human cancer: linking DNA methylation to gene silencing. Carcinogenesis 23: 1103-1 109.

Carey, C. and Bryant, C., 1995. Possible interrelations among environmental toxicants, amphibian development and decline of amphibian populations. Environ. Health Perspec. 103: 13-17.

Barbeau, T. R. and Guillette, L. J., Jr., 2007. Altered ovarian steroids in Xenopus laevis exposed to environmentally relevant concentrations of nitrate. J. Herpetol. 41: 590-596. Belfroid, A. C., Van Der Horst, A., Vethaak, A. D., Schafer, A. J., Rijs, G. B. J., Wegener, J. and Cofino, W. F!, 1999. Analysis and occurrence of estrogenic hormones and their glucuronides in surface water and waste water in the Netherlands. Sci. Total Environ. 225: 101-108. Bern, H., 1992. The fragile fetus. Pp. %15 in "ChemicallyInduced Alterations in Sexual and Functional Development: The WildlifeiHuman Connection", ed by T Colborn and C. Clement. Princeton Scientific Publishing Company, Princeton. Bernbe, V. E., Boily, M. H., Deblois, C., Dassylva, N. and Spear, F! A, 2005. Plasma retinoid profile in b u k g s , Rana catesbeiana, in relation to agricultural intensity of sub-watersheds in the Yamaska River drainage basin, Quebec, Canada. Aquatic Toxicol. 71: 10%120. Bjerregaard, F!, Dewailly, E., Ayotte, F!, Pars, T., Ferron, L. and Mulvad, G., 2001. Exposure of Inuit in Greenland to organochlorines through the marine diet. J. Toxicol. Environ. Health Part A 62: 69-81.

Cheek, A. O., Ide, C. F., Bollinger, J. E., Rider, C. V. and Mclachlan, J. A,, 1999. Alteration of leopard frog (Rana pipiens) metamorphosis by the herbicide acetochlor. Arch. Environ. Contam. Toxicol. 37: 70-77. Clotfelter, E. D., Bell, A. M. and Levering, K. R., 2004. The role of animal behaviour in the study of endocrine-disrupting chemicals. Anim. Behav. 68: 665-676. Colborn, T and Smolen, M. J., 1996. Epidemiological analysis of persistent organochlorine contaminants in Cetaceans. Rev. Envimn. Contam. Toxicol. 146: 91-172. Cooke, A. S., 1970. The effect of p,p'-DDT on tadpoles of the common frog (Rana temporaria). Environ. Pollut. 1: 57-71. Cooke, A. S., 1971. Selective predation by newts on frog tadpoles treated with DDT Nature 229: 275-276. Crain, D. A. and Guillette, L. J., Jr., 1997. Endocrinedisrupting contaminants and reproduction in vertebrate wildlife. Rev. Toxicol. 1: 47-70. Crain, D. A,, Noriega, N., Vonier, F! M., Arnold, S. F., Mclachlan, J. A. and Guillette, L. Jr., 1998. Cellular bioavailability of natural hormones and environmental contaminants as a function of serum and cytosolic binding factors. Toxicol. Indust. Health 14: 261-273.

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUITING CHEMICALS Crews, D. and Mclachlan, J., 2006. Epigenetics, evolution, endocrine disruption, health, and disease. Endocrinology 147: S4-S 10. Crews, D., Gore, A. C., Hsu, T. S., Dangleben, N. L., Spinetta, M., Schallert, T., Anway, M. D. and Skinner, M. K., 2007. Transgenerational epigenetic imprints on mate preference. Proc. Nut. Acad. Sci. USA. 104: 5942-5946. Crump, D., Lean, D. and Trudeau, V. L., 2002. Octylphenol and UV-B radiation alter larval development and hypothalamic gene expression in the leopard frog (Rana pipiens). Enuiron. Health Perspec. 110: 277-284. Daftary, G. S. and Taylor, H. S., 2006. Endocrine regulation of Hox genes. Endo. Rev. 27: 331-355. Danzo, B. J., 1997. Environmental xenobiotics may disrupt normal endocrine function by interfering with the binding of physiological ligands to steroid receptors and binding proteins. Enuiron. Health Perspec. 105: 294-301.

3233

Fan, W. Q., Yanase, T., Morinaga, H., Gondo, S., Okabe, T., Nomura, M., Komatsu, T., Morohashi, K. I., Hayes, T B., Takayanagi, R. and Nawata, H., 2007. Atrazine-induced aromatase expression is SF-1 dependent: implications for endocrine disruption in wildlife and reproductive cancers in Humans. Environ. Health Perspec. 115: 720-722. Fei, X. L., Chung, H. J. and Taylor, H. S., 2005. Methoxychlor disrupts uterine HoxalO gene expression. Endocrinology 146: 3445-345 1. Forman, B. M., Tzameli, I., Choi, H. S., Chen, L., Simha, D., Seol, FV., Evans, R. M. and Moore, D. D., 1998. Androstane metabolites bind to and deactivate the nuclear receptor CAR-P. Nature 395: 612-615. Forson, D. and Storfer, A,, 2006. Effects of atrazine and iridovirus infection on survival and life-history traits of the long-toed salamander (Ambystoma macrodoctylum). Environ. Toxicol. Chem. 25: 168-173.

Danzo, B. J., 1998. The effects of environmental hormones on reproduction. Celluhr Molec. Lqe Sci. 54: 1249-1264.

Fort, D. J., Degitz, S., Tietge, J. and Touart, L. W., 2007. The hypothalamic-pituitary-thyroid (HPT) axis in frogs and its role in frog development and reproduction. Critical Rev. Toxicol. 37: 117-161.

Davidson, C. and Knapp, R. A,, 2007. Multiple stressors and amphibian declines: Dual impacts of pesticides and fish on yellow-legged frogs. Ecol. Appl. 17: 587-597.

Fox, G. A,, 1995. Tinkering with the tinkerer: Pollution verses evolution. Environ. Health Perspec. 103 (Supplement 4): 93-100.

Degitz, S. J., Holcombe, G. W., Flynn, K. M., Kosian, P. A., Korte, J. J . and Tietge, J. E., 2005. Progress towards development of an amphibian-based thyroid screening assay using Xenopus laeuis. Organismal and thyroidal responses to the model compounds 6-propylthiouracil, methimazole, and thyroxine. Toxicol. Sci. 87: 353-364. Denton, T E., Howell, W. M., Allison, J. J., Mccollum, J. and Marks, B., 1985. Masculinization of female mosquitofish by exposure to plant sterols and Mycobacterium smegmatis. Bull. Enuiron. Contam. Toxicol. 35: 627-632. Dreitz, V. J., 2006. Issues in species recovery: An example based on the Wyoming toad. BioScience 56: 765-771. Durhan, E. J., Lambright, C. S., Makynen, E. A,, Lazorchak, J., Hartig, l? C., Wilson, V. S., Gray, L. E. and Ankley, G. T, 2007. Identification of metabolites of trenbolone acetate in androgenic runoff from a beef feedlot. Enuiron. Health Perspec. 114 (Supplement 1): 65-68. W a r d s , T. M. and Myers, J. l?, 2007. Environmental exposures and gene regulation in disease etiology. Enuiron. Health Perspec. 115: 1264-1270.

Fox, G. A., 2001. Effects of endocrine disrupting chemicals on wildlife in Canada: Past, present and future. Water Qual. Res. J. Canada 36: 233-251. Freeman, J. L., Beccue, N. and Rayburn, A. L., 2005. Differential metamorphosis alters the endocrine response in anuran larvae exposed to T, and atrazine. Aquat. Toxicol. 75: 263-276. Gardiner, D. M. and Hoppe, D. M., 1999. Environmentally induced limb malformations in mink frogs (Rana septentrionalis).J. Experi. 2001.284: 207-216. Gardiner, D., Ndayibagira, A,, Grun, F. and Blumberg, B., 2003. Deformed frogs and environmental retinoids. Pure Appl. Chem. 75: 2263-2273. Goksoyr, A., 2006. Endocrine disruptors in the marine environment: mechanisms of toxicity and their influence on reproductive processes in fish. J. Toxicol. Enuiron. Health Part A 69: 175-184. Goleman, W. L., Carr, J. A. and Anderson, T A,, 2002. Environmentally relevant concentrations of ammonium perchlorate inhibit thyroid function and alter sex ratios in developing Xenopw laevis. Environ. Toxicol. Chem. 21: 590-597.

Idrrards, T M., Mccoy, K. A., Barbeau, T, Mccoy, M. \V, Thro, J. M. and Guillette, L. J., Jr., 2006. Environmental context determines nitrate toxicity in southern toad (Bufo terrestris) tadpoles. Aquatic Toxicol. 78: 50-58.

Gray, L. E., Jr., Wilson, V. S., Stoker, T, Lambright, C., Furr, J., Noriega, N., Howdeshell, K., Ankley, G. T and Guillette, L. J., Jr., 2006. Adverse effects of environmental antiandrogens and androgens on reproductive development in mammals. Internat. J. Androl. 29: 96-104.

M e n k o , V. l?, Amstislavsky, S. Y., Schwabel, H. and Ingermann, R. L., 2002. Altered behaviors in male mice, male quail, and salamander larvae following early exposures to the estrogenic pesticide rnethoxychlor. Neurotoxicol. Teratol. 24: 29-36.

Guillette, L. J., Jr. and Iguchi, T, 2003. Contaminantinduced endocrine and reproductive alterations in reptiles. Pure Appl. Chem.. 75: 2275-2286.

T-Commission, 1997. European Workshop on the Impact of Endocrine Disrupters on Human Health and the Environment. In "Environment and Climate k a r c h Programme", Vol. DG XII, European Commission. http://ec.europa.eu/environ menu emlocrineldocuments/reports~conclusions~ en.htm*

Guillette, L. J., Jr., 2004. Contaminants, reproductive endocrinology and wildlife: The evolving field of signal disruption. Harmon. Behau. 46: 130-13 1. Guillette, L. J., Jr., 2006. Endocrine disrupting contaminants - Beyond the dogma. Enuiron. Health Perspec. 114: 9-12.

3234

AMPHIBIAN BIOLOGY

Guillette, L. J., Jr. and Gunderson, M. F!, 2001. Alterations in development of reproductive and endocrine systems of wildlife populations exposed to endocrine-disrupting contaminants. Reproduction 122: 857-864.

Helbing, C. C. and Atkinson, B. G., 1994. 3,5,3'triiodothyronine-induced carbamyl-phosphate synthetase gene-expression is stabilized in the liver of rana catesbeiana tadpoles during heat-shock. J. Biol. Chem. 269: 11743-1 1750.

Guillette, L. J., Jr., Edwards, T. M. and Moore, B. C., 2007. Alligators, environmental contaminants and steroid hormones. Enuiron. Sci., in press.

Helbing, C., Gallimore, C. and Atkinson, B. G., 1996. Characterization of a Rana catesbeiana HSP3O gene and its expression in the liver of this amphibian during both spontaneous and thyroid hormoneinduced metamorphosis. Develop. Genet. 18: 223-233.

Guillette, L. J., Jr., Crain, D. A., Rooney, A. A. and Pickford, D. B., 1995. Organization Versus Activation - the Role of Endocrine-Disrupting Contaminants (Edcs) During Embryonic-Development in Wildlife. Enuiron. Health Perspec. 103: 157-164. Guillette, L. J., Jr., Kools, S., Gunderson, M. F! and Bermudez, D. S., 2006. DDT and its analogues: new insights into their endocrine disruptive effects on wildlife. Pp. 332-355 in "Endocrine Disruption: Biological Basis for Health Effects in Wildlife and Humans", ed by D. 0.Norris and J. A. Carr. Oxford University Press, New York. Harris, M. L., Chora, L., Bishop, C. A. and Bogart, J. P, 2000. Species- and age-related differences in susceptibility to pesticide exposure for two amphibians, Ram p i e . and Bufo americanus. Bull. Enuiron. Conturnin. Toxicol. 64: 263-270. Harvey, P W., Everett, D. J. and Springall, C. J., 2007. Adrenal toxicology: a strategy for assessment of functional toxicity to the adrenal cortex and steroidogenesis. J. Appl. Toxicol. 27: 103-1 15. Hayes, T B., 1998. Sex determination and primary sex differentiation in amphibians: Genetic and developmental mechanisms. J. Experim. 2001. 281: 373-399. Hayes, T. B., Haston, K., Tsui, M., Hoang, A., Haeffele, C. and Vonk, A., 2003. Atazine-induced hermaphroditism at 0.1 ppb in American leopard frogs (Rana pipiens): Laboratory and field evidence. Environ. Health Perspec. 111: 568-575. Hayes, T. B., Collins, A., Lee, M., Mendoza, M., Noriega, N., Stuart, A. A. and Vonk, A., 2002. Hermaphroditic, demasculinized frogs after exposure to the herbicide atrazine at low ecologically relevant doses. Proc. Nut. Acad. Sci. USA. 99: 5476-5480. Hayes, T B., Stuart, A. A,, Mendoza, M., Collins, A., Noriega, N., Vonk, A,, Johnston, G., Liu, R. and Kpodzo, D., 2006b. Characterization of atrazineinduced gonadal malformations in African clawed frogs (Xenqpw laeuis) and comparisons with effects of an androgen antagonist (cyproterone acetate) and exogenous estrogen (l7P-estradiol): Support for the demasculinization/feminization hypothesis. Enuiron. Health Perspec. 114: 134-141.

Helbing, C., Gergely, G. and Atkinson, B. G., 1992. Sequential up-regulation of thyroid-hormone betareceptor, onithine transcarbamylase, and carbamylphosphate synthetase messenger-RNAs in the liver of Rana catesbeiana tadpoles during spontaneous and thyroid hormone-induced metamorphosis. Develop. Genet. 13: 289-30 1. Hinson, J. F! and Raven, F! W., 2006. Effects of endocrine-disrupting chemicals on adrenal function. Best Practice Res. Clinic. Endocrin. Metab. 20: 111-120. Holliday, R. and Pugh, J. E., 1975. DNA modification mechanisms and gene activity during development. Science 187: 226-232. Iguchi, T, Watanabe, H. and Katsu, Y, 2001. Developmental effects of estrogenic agents on mice, fish and frogs: a mini-review. Hormon. Behau. 40: 248-251. IPCS, 2002. Global Assessment or the state of the science of endocrine disruptors (eds T Damstra, S. Barlow, A. Bergman, R. Kavlock and G. Van der Kraak), Vol. WHO/PCS/EDC/O2/2: 1-180. World Health Organization. Janosek, J., Hilscherova, K., Blaha, L. and Holoubek, I., 2006. Environmental xenobiotics and nuclear receptors - Interactions, effects and in vitro assessment. Toxicol. in Ktro 20: 18-37. Jansen, M. S., Nagel, S. C., Miranda, F! J., Lobenhofer, E. K., Afshari, C. A. and Mcdonnell, D. P, 2004. Short-chain fatty acids enhance nuclear receptor activity through mitogen-activated protein kinase activation and histone deacetylase inhibition. Proc. Nut. Acad. Sci. USA. 101: 7199-7204. Jobling, S., Beresford, N., Nolan, M., Rodgers-Gray, T., Brighty, G. C., Sumpter, J. F! and Tyler, C. R., 2002. Altered sexual maturation and gamete production in wild roach (Rutilus rutilus) living in rivers that receive treated sewage effluents. Biol. Repro. 66: 272-281. Johnson, A. C., Williams, R. J. and Matthiessen, P, 2006. The potential steroid hormone contribution of farm animals to freshwaters, the United Kingdom as a case study. Sci. Total Environ. 362: 166-178.

Hayes, T. B., Case, F!, Chui, S., Chung, D., Haeffele, C., Haston, K., Lee, M., Mai, V F!, Ma juoa, Y., Parker, J. and Tsui, M., 2006a. Pesticide mixtures, endocrine disruption, and amphibian declines: Are we underestimating the impact? Enuiron. Health Perspec. 114: 40-50.

Jost, J. P, Saluz, H. P., Mcewan, I., Feavers, I. M., Hughes, M., Reiber, S., Liang, H. M. and Vaccaro, M., 1990. Tissue specific expression of avian vitellogenin gene Is correlated with DNA hypomethylation and invivo specific protein-DNA interactions. Philosoph. Trans. Roy. Soc. Lond. Series B-Biolog. Sci. 326: 231-240.

Hecker, M., Giesy, J. P,Jones, P D., Jooste, A. M., Carr, J. A., Solomon, K. R., Smith, E. E., Van Der Kraak, G., Kendall, R. J. and Du Preez, L., 2004. Plasma sex steroid concentrations and gonadal aromatase activities in African clawed frogs (Xenopus laeuis) from South Africa. Enuiron. Toxicol. Chem. 23: 1996-2007.

Kamiya, K., Sato, T, Nishimura, N., Goto, Y , Kano, K. and Iguchi, T., 1996. Expression of estrogen receptor and proto-oncogene messenger ribonucleic acids in reproductive tracts of neonatally diethylstilbestrol-exposed female mice with or without postpuberal estrogen administration. Experi. Clinic. Endocrin. Diabetes 104: 111-122.

McCOY and GUILLETTE Jr.: END(X R I N E DISRUFTING CHEMICALS

3235

Koussoulakos, S., 2004. Vertebrate limb development: from Harrison's limb disk transplantations to targeted disruption of Hox genes. Anat. Emhyol. 209: 93-105.

Moore, F. L., Boyd, S. K and Kelley, D. B., 2005. Historical perspective: hormonal regulation of behaviors in amphibians. H m m m Behau. 48: 373-383.

Kretschmer, X. C. and Baldwin, W. S., 2005. CAR and PXR: Xenosensors of endocrine disrupters? ChemuoBiolog. Inter: 155: 1 11-128.

Mosconi, G., Carnevali, O., Franzoni, M. F., Cottone, E., Lutz, I., Kloas, W., bmamoto, K., Kikuyama, S. and Polzonetti-Magni, A. M., 2002. Environmental estrogens and reproductive biology in amphibians. Gen. Campar. Enduoin. 146: 125-129.

Lange, R., Hutchinson, T. H., Croudace, C. l?, Siegmund, F., Schweinfurth, H., Hampe, l?, Panter, G. l? and Sumpter, J. l?, 2001. Effects of the synthetic estrogen 17a-ethinylestradiol on the lifecycle of the fathead minnow (Pimephales promelas) Environ. Toxicol. Chem. 20: 1216-1227. Leblanc, G. A,, 1994. Assessing deleterious ecosystemlevel effects of environmental pollutants as a means of avoiding evolutionary consequences. Environ. Health Perspec. 102: 266-267. Leiva-Presa, A. and Jenssen, B. M., 2006. Effects of p,p '-DDE on retinoid homeostasis and sex hormones of adult male European common frogs (Rana temporaria).J. Toxicol. Environ. Health Part A Current Issues 69: 205 1-2062. Lewis, D. L., Baxter, G. T, Johnson, K. M. and Stone, M. D., 1985. Possible extinction of the Wyoming toad, Bufo hemiophrys baxteri.J. Herpetol. 19: 166-168. Lie, E., Bernhoft, A., Riget, F., Belikov, S. E., Boltunov, A. N., Derocher, A. E., Garner, G. W., Wiig, 0. and Skaare, J. U., 2003. Geographical distribution of organochlorine pesticides (OCPs) in polar bears (Ursw maritimus) in the Norwegian and Russian Arctic. Sci. Total Environ. 306: 159-170. Lintelmann, J., Katayama, A,, Kurihara, N., Shore, L. and Wenzel, A., 2003. Endocrine disruptors in the environment. Pure Appl. Chem. 75: 63 1-68 1. Mahmood-Khan, 2. and Hall, E. R., 2003. Occurrence and removal of plant sterols in pulp and paper mill effluents. J. Environ. Engineer. Sci. 2: 17-26. Mangelsdorf, D. J., Thummel, C., Beato, M., Herrlich, l?, Schutz, G., Umesono, K., Blumberg, B., Kastner, l?, Manuel, M., Chambon, l? and Evans, R. M., 1995. The nuclear receptor superfamily: the second decade. Ell 83: 835-839. Masuyama, H. and Hiramatsu, Y, 2004. Involvement of suppressor for Gal 1 in the ubiquitin/proteasome mediated degradation of estrogen receptors. J. Biolog. Chem. 279: 12020-12026. Matthiessen, l? and Johnson, I., 2007. Implications of research on endocrine disruption for the environmental risk assessment, regulation and monitoring of chemicals in the European Union. Environ. Poll. 146: 9-18. Matthiessen, l?, Arnold, D., Johnson, A. C., Pepper, T J., Pottinger, T. G. and Pulman, K. G. T, 2006. Contamination of headwater streams in the United l n g d o m by oestrogenic hormones from livestock farms. Sci. Total Environ. 367: 616-630. Mclachlan, J. A., 200 1. Environmental signaling: what embryos and evolution teach us about endocrine disrupting chemicals. Endocrine Rev. 22: 3 19-34 1.

Mukejee, D., 2006. Special section. Endocrine disruptors - Introduction. Envimn. Res. 100: 1-2. Nagel, S. C., Vom Saal, F. S., Thayer, K. A, Dhar, M. G., Boechler, M. and Welshons, W. V, 1997. Relative binding affinity-serum modified access (RBA-SMA) assay predicts the relative in viva bioactivity of the xenoestrogens Bisphenol A and octylphenol. Envirun. Health Perspec. 105: 70-76. Nations, B. K. and Hallberg, G. R., 1992. Pesticides in Iowa precipitation. J. Environ. Qual. 21: 486492. Noriega, N. C. and Hayes, T B., 2000. DDT congener effects on secondary sex coloration in the reed frog Hyperolius argus: a partial evaluation of the Hyperoliw argw endocrine screen. Compar. Biochem. Physiol. B-Biochem. Molec. Biol. 126: 231-237. Norris, D. 0. and Carr, J. A., eds., 2006. "Endocrine Disruption: Biological Basis for Health Effects in Wildlife and Humans". Oxford University Press, New York. Ogura, T and Evans, R. M., 1995. A retinoic acid triggered cascade of Hoxbl gene activation. Proc. Nut. Acad. Sci. USA. 92: 387-391. Ohtani, H., Miura, I. and Ichikawa, Y , 2000. Effects of dibutyl phthalate as an environmental endocrine disruptor on gonadal sex differentiation of genetic males of the frog Rana rugosa. Environ. Health Perspec. 108: 1189-1 193. Opitz, R., Hartmann, S., Blank, T, Braunbeck, T, Lutz, I. and Kloas, W., 2006a. Evaluation of histological and molecular endpoints for enhanced detection of thyroid system disruption in Xenopus laevis tadpoles. Toxicol. Sci. 90: 337-348. Opitz, R., Trubiroha, A., Lorenz, C., Lutz, I., Hartmann, S., Blank, T, Braunbeck, T and Kloas, W., 2006b. Expression of sodium-iodide symporter mRNA in the thyroid gland of Xenopus laevis tadpoles: developmental expression, effects of antithyroidal compounds, and regulation by TSH. J. Endocrin. 190: 157-170. Opitz, R., Braunbeck, T., Bogi, C., Pickford, D. B., Nentwig, G., Oehlmann, J., Tooi, O., Lutz, I. and Kloas, W., 2005. Description and initial evaluation of a Xenopus metamorphosis assay for detection of thyroid system-disrupting activities of environmental compounds. Environ. Toxicol. Chem. 24: 653-664. Orlando, E. F., Kolok, A. S., Binzcik, G. A., Gates, J. L., Horton, M. K., Lambright, C. S., Gray, L. E., Soto, A. M. and Guillette, L. Jr., 2004. Endocrinedisrupting effects of cattle feedlot effluent on an aquatic sentinel species, the fathead minnow. Environ. Health Perspec. 112: 353-358.

Miller, W. L., 1988. Molecular biology of steroid hormone synthesis. Endocrine Rev. 9: 295-318.

Ouellet, M., Bonin, J., Rodrigue, J., Desgranges, J. L. and Lair, S., 1997. Hindlimb deformities (ectromelia, ectrodactyly) in free-living anurans from agricultural habitats. J. Wildl. Dis. 33: 95-104.

Milnes, M. R., Bermudez, D. S., Bryan, T A,, Edwards, T M., Gunderson, M. E, Larkin, I. L. V., Moore, B. C. and Guillette, L. Jr., 2006. Contaminant-induced feminization and demasculinization of nonmammalian vertebrate males in aquatic environments. Environ. Res. 100: 3-17.

Palace, V. l?, Evans, R. E., Wautier, K., Baron, C., Vandenbyllardt, L., Vandersteen, W. and Kidd, K., 2002. Induction of vitellogenin and histological effects in wild fathead minnows from a lake experimentally treated with the synthetic estrogen, ethynylestradiol. Water Qual. Res. J. Canada 37.

3236

AMPHIBIAN BIOLOGY

Palanza, P., Morellini, F., Parmigiani, S. and Vom Saal, F. S., 2002. Ethological methods to study the effects of maternal exposure to estrogenic endocrine disrupters - A study with methoxychlor. Neurotoxicol. Teratol. 24: 55-69. Palmer, B. D. and Palmer, S. K., 1995. Vitellogenin Induction by xenobiotic estrogens in the red-eared turtle and African clawed frog. Environ. Health Perspec. 103: 19-25. Palmer, B. D., Huth, L. K., Pieto, D. L. and Selcer, K. W., 1998. Vitellogenin as a biomarker for xenobiotic estrogens in an amphibian model system. Enuiron. Toxicol. Chem. 17: 30-36. Panzica, G. C., Viglietti-Panzica, C. and Ottinger, M. A,, 2005. Introduction: neurobiological impact of environmental estmgens. Brain Res. Bull. 65: 187-191. Park, B. J. and Kidd, K., 2005. Effects of the synthetic estrogen ethinylestradiol on early life stages of mink frogs and green frogs in the wild and in situ. Enuiron. Toxicol. Chem. 24: 2027-2036. Park, D. and Propper, C. R., 2002. Endosulfan affects pheromonal detection and glands in the male redspotted newt, Notophthalmw uindescens. Bull. Enuiron. Contam. Toxicol. 69: 609-616. Parks, L. G., Lambright, C. S., Orlando, E. F., Guillette, L. J., Jr., Ankley, G. T and Gray, L. E., Jr., 2001. Masculinization of female mosquitofish in kraft mill effluent-contaminated Fenholloway river water is associated with androgen receptor agonist activity. Toxicol. Sci. 62: 257-267. Pawlowski, S., Van Aerle, R., Tyler, C. R. and T. B., 2004. Effects of 17a-ethinylestradiol in a fathead minnow (A'mephales promelas) gonadal recrudescence assay. Ecotoxicol. Environ. Safety 57: 330-349. Pawlowski, S., Temes, I,Bonerz, M., Kluczka, T., \'an Der Burg, B., Nau, H., Errlinger, L. and Braunbeck, T, 2003. Combined in situ and in vitro assessment of the estmgenicactivity of sewage and surface water samples. Toxicol. Sci. 75: 57-65. Peterson, F. W., Davis, R. K. and Omdorff, H. A,, 2000. 17P-estradiol as an indicator of animal waste contamination in mantled karst aquifers. J. Enuiron. Qual. 29: 826-834. Pickford, D. B. and Morris, I. D., 1999. Effects of endocrine-disrupting contaminants on amphibian oogenesis: methoxychlor inhibits progesteroneinduced maturation of Xmopus laevis oocytes in vitro. Enuiron. Health Perspec. 107: 285-292. Pickford, D. B. and Morris, I. D., 2003. Inhibition of gonadotropin-induced oviposition and ovarian steroidogenesis in the African clawed frog (Xenopus laeuis) by the pesticide methoxychlor. Aquat. Toxicol. 62: 179-194. Pottinger, T G., 2003. Interactions of endocrine disrupting chemicals with stress responses in wildlife. Pure Appl. Chem. 75: 2321-2333. Propper, C. R., 2005. The study of endocrine disrupting compounds: past approaches and new directions. Zntegrat. Compa?: Biol. 45: 194-200. Qin, Z. F., Zhou, J. M., Chu, S. G. and Xu, X. B., 2003. Effects of Chinese domestic polychlorinated biphenyls (PCBs) on gonadal differentiation in Xenopw laenis. Enuiron. Health Perspec. 111: 553-556. Reik, W. and Walter, J., 2001a. Evolution of imprinting mechanisms: the battle of the sexes begins in the zygote. Nut. Genet. 27: 255-256.

Reik, W. and Walter, J., 2001b. Genomic imprinting: Parental influence on the genome. Nut. Rev. G m d . 2: 21-32. Reik, W., Dean, W. and Walter, J., 2001. Epigenetic reprogramming in mammalian development. Science 293: 1089-1093. Relyea, R. A. and Mills, N., 2001. Predator-induced stress makes the pesticide carbaryl more deadly to gray treefrog tadpoles (Hyla uersicolor). Proc. Not. Acad. Sci. USA. 98: 2491-2496. Rohr, J. R. and Palmer, B. D., 2005. Aquatic herbicide exposure increases salamander desiccation risk eight months later in a terrestrial environment. Enuiron. Toxicol. Chem. 24: 1253-1258. Rohr, J. R., Kerby, J. L. and Sih, A., 2006a. Community ecology as a framework for predicting contaminant effects. Trend. Ecol. Evol. 21: 606-613. Rohr, J. R., Sager, T, Sesterhenn, T. M. and Palmer, B. D., 2006b. Exposure, postexposure, and densitymediated effects of atrazine on amphibians: Breaking down net effects into their parts. Enuiron. Health Perspec. 114: 46-50. Rohr, J. R., Elskus, A. A,, Shepherd, B. S., Crowley, P. H., Mccarthy, T. M., Niedmiecki, J. H., Sager, T., Sih, A. and Palmer, B. D., 2004. Multiple stressors and salamanders: Effects of an herbicide, food limitation, and hydroperiod. Ecolog. Appl. 14: 1028-1040. Romansic, J. M., Diez, K. A., Higashi, E. M. and Blaustein, A. R., 2006. Effects of nitrate and the pathogenic water mold Saprolegnia on survival of amphibian larvae. Dis. Aquat. Organ. 68: 235-243. Rosenshield, M. L., Jofre, M. B. and Karasov, W. H., 1999. Effects of polychlorinated biphenyl 126 on green frog (Rana clamitans) and leopard frog (Rana pipas) hatching success, development, and metamorphosis. Environ. Toxicol. Chem. 18: 2478-2486. Russell, R. W., Gillan, K. A. and Haffner, G. D., 1997. Polychlorinated biphenyls and chlorinated pesticides in southern Ontario, Canada, green frogs. Enuiron. Toxicol. Chem. 16: 2258-2263. Russell, R. W., Hecnar, S. J. and Haffner, G. D., 1995. Organochlorine pesticide residues in southern Ontario spring peepers. Enuiron. Toxicol. Chem. 14: 815-817. Sachs, L. M., Damjanovski, S., Jones, P L., Li, Q., Amano, T, Ueda, S., Shi, Y. B. and Ishizuya-Oka, A., 2000. Dual functions of thyroid hormone receptors during Xenopus development. Compar: Biochem. Physiol. B-Biochem. Molec. Biol. 126: 199-2 11. Saluz, H. P., Jiricny, J. and Jost, J. E, 1986. Genomic sequencing reveals a positive correlation between the kinetics of strand-specific DNA demethylation of the overlapping estradiol glucocorticoid receptorbinding sites and the rate of avian vitellogenin messenger RNA synthesis. Proc. Nut. Acad. Sci. USA. 83: 7167-7171. Sanderson, J. T, 2006. The steroid hormone biosynthesis pathway as a target for endocrinedisrupting chemicals. Toxicol. Sci. 94: 3-21. Sanderson, J. T, Boerma, J., Lansbergen, G. W. A. and Van Den Berg, M., 2002. Induction and inhibition of aromatase (CYP19) activity by various classes of pesticides in H295R human adrenocortical carcinoma cells. Toxicol. Appl. Pharmacol. 182: 44-54.

McCOY and GUILLETTE Jr.: ENDOCRINE DISRUF'TING CHEMICALS Sanderson, J. T, Seinen, W., Giesy, J. F! and Van Den Berg, M., 2000. 2-chloro-s-triazine herbicides induce aromatase (CYP19) activity in H295R human adrenocortical carcinoma cells: A novel mechanism for estrogenicity? Toxicol. Sci. 54: 121-127. Severinsen, T., Skaare, J. U. and Lydersen, C., 2000. Spatial distribution of persistent organochlorines in ringed seal (Phoca hispiah) blubber. Marine Environ. Res. 49: 291-302. Skaare, J. U., Bernhoft, A., Derocher, A,, Gabrielsen, G. W., Goks~yr,A., Henriksen, E., Larsen, H. J., Lie, 2000. Organochlorines in top E. and Wiig, 0., predators at Svalbard-occurrence, levels and effects. Toxicol. letters 112-113: 103-109. Soto, A. M., Calabro, J. M., Prechtl, N. V., Yau, A. Y., Orlando, E. F., Daxenberger, A,, Kolok, A. S., Guillette, L. J. J., Le Bizec, B., Lange, I. G. and Sonnenschein, C., 2004. Androgenic and estrogenic activity in water bodies receiving cattle feedlot effluent in eastern Nebraska, USA. Enuiron. Health Perspec. 112: 34C352. Sparling, D. W., Fellers, G. M. and Mcconnell, L. L., 2001. Pesticides and amphibian population declines in California, USA. Enuiron. Toxicol. Chem. 20: 1591-1595. Spengler, P, Korner, W. and Metzger, J. W., 2001. Substances with estrogenic activity in effluents of sewage treatment plants in southwestern Germany. 1. Chemical analysis. Enuiron. Toxicol. Chem. 20: 2133-2141. Steele, C. W., Strickler-Shaw, S. and Taylor, D. H., 1999. Effects of sublethal lead exposure on the behaviours of green frog (Rana clamitans), bullfrog (Rana catesbiana) and American toad (Bufo americanus) tadpoles. Mar: Freshw. Behau. Physiol. 32: 1-16. Storrs, S. I. and Kiesecker, J. M., 2004. Survivorship patterns of larval amphibians exposed to low concentrations of atrazine. Environ. Health Perspec. 112: 1054-1057. Sullivan, K. B. and Spence, K. M., 2003. Effects of sublethal concentrations of atrazine and nitrate on metamorphosis of the African clawed frog. Enuiron. Toxicol. Chem. 22: 627-635. Tabb, M. M. and Blumberg, B., 2006. New modes of action for endocrine-disrupting chemicals. Molec. Endocrin. 20: 475-482. Tamura, H., Maness, S. C., Reischmann, K., Dorman, D. C., Gray, L. E. and Gaido, K. W., 2001. Androgen receptor antagonism by the organophosphate insecticide fenitrothion. Toxicol. Sci. 60: 5G62. Tavera-Mendoza, L., Ruby, S., Brousseau, F!, Fournier, M., Cyr, D. and Marcogliese, D., 2002a. Response of the amphibian tadpole Xenopus laeuis to atrazine during sexual differentiation of the ovary. Enuirm. Toxicol. Chem. 21: 1264-1 267. Tavera-Mendoza, L., Ruby, S., Brousseau, P, Fournier, M., Cyr, D. and Marcogliese, D., 2002b. Response of the amphibian tadpole Xenopus laeuis to atrazine during sexual differentiation of the testis. Enuiron. Toxicol. Chem. 21: 527-53 1. Taylor, B., Skelly, D., Demarchis, L. K., Slade, M. D., Galusha, D. and Rabinowitz, F! M., 2005. Proximity to pollution sources and risk of amphibian limb malformation. Enuirm. Health Perspec. 113: 1497-150 l .

3237

Terasah, S., Ata, Y, Inoue, A., Hayashi, S., Nishigaki, M., Aoyagi, K, Sasaki, H., Wda-Kiyama, Y., Sakuma, Y., Akaba, S., Tanaka, J., Sone, H., Yonemoto, J., Tanji, M. and Kiyarna, R., 2004. Using a customized DNA microarray for expression profiling of the estrogen-responsive genes to evaluate estrogen activity among natural estrogens and industrial chemicals. Enuimn. HeaWI PPrpec. 112: 773-781. Theodorakis, C. W.,Rinchard, J., Carr, J. A, Park, J. W., Mcdaniel, L., Liu, F. J. and Wages, M., 2006. Thyroid endocrine disruption in stonerollers and cricket frogs from perchloratecontaminated streams in east-central Texas. E c o t o x i c o b ~15: 31-50. Thurman, E. M. and Cromwell, A, 2000. Armospheric transport, deposition, and fate of &ine herbicides and their metabolites in pristine areas at Isle Royale National Park. Enuiron. Sci. Technol. 34: 3079-3085. Tietge, J. E., Holcombe, G. W., Flynn, K. M., Kosian, F! A., Korte, J. J., Anderson, L. E., Wolf, D. C. and Degitz, S. J., 2005. Metamorphic inhibition of Xenopus laeuis by sodium perchlorate: Effects on development and thyroid histology. Enuiron. Toxuol. Chem. 24: 926-933. Toft, G., Baatrup, E. and Guillette, L. J., Jr., 2004. Altered social behavior and sexual characteristics in mosquitofish (Gambwia holbrooki) living downstream of a paper mill. Aquatic Toxicol. 70: 2 13-222. Veldhoen, N. and Helbing, C. C., 2001. Detection of environmental endocrine-disruptor effects on gene expression in live Rana catesbeiana tadpoles using a tail fin biopsy technique. Enuiron. Toxicol. Chem. 20: 2704-2708. Verrell, P, 2000. Methoxychlor increases susceptibility to predation in the salamander Ambystoma macrodactylum. Bull. Enuiron. Contam. Toxicol. 64: 85-92. Watanabe, H., Suzuki, A,, Kobayashi, M., Takahashi, E., Itamoto, M., Lubahn, D. B., Handa, H. and Iguchi, T., 2003. Analysis of temporal changes in the expression of estrogen-regulated genes in the uterus. J. MoLc. Endocrinol. 30: 347-358. Wheeler, J. R., Gimeno, S., Crane, M., Lopez-Juez, E. and Morritt, D., 2005. Vitellogenin: A review of analytical methods to detect (anti) estrogenic activity in fish. Toxicol. M e c h n . Methods 15: 293-306. Woodlmg,J. D., Lopez, E. M., Maldonado, 7: A., Norris, D. 0 . and Vajda, A. M., 2006. Intersex and other reproductive disruption of fish in wastewater emuent dominated Colorado streams. Compar. Biochem. Physiol., Part C 144: 10-15. Xie, W., Ba~wick,J. L., Simon, C. M., Pierce, A. M., Safe, S., Blumberg, B., Guzelian, P S. and Evans, R. M., 2000. Reciprocal activation of xenobiotic response genes by nuclear receptors SXWPXR and CAR. Genes and Develop. 14: 3014-3023. Yamamoto, K. R., 1985. Steroid receptor regulated transcription of specific genes and gene networks. Ann. Rev. Genet. 19: 209-252. Zala, S. M. and Penn, D. J., 2004. Abnormal behaviours induced by chemical pollution: a review of the evidence and new challenges. Anim. Behau. 68: 649-664. Zhang, F., Degitz, S. J., Holcombe, G. W., Kosian, P A., Tietge, J., Veldhoen, N. and Helbing, C. C., 2006. Evaluation of gene expression endpoints in the context of a Xenopus laeuis metamorphosis-based bioassay to detect thyroid hormone disruptors. Aquatic Toxicol. 76: 24-36.

3238

AMPHIBIAN BIOLOGY

Zoeller, R. T and Tan, S. W., 2007. Implications of research on assays to characterize thyroid toxicants. Crit. Rev. Toxicol. 37: 195-210. Zoeller, R. T, Tan, S. W. and Tyl, R. W., 2007. General background on the hypothalamic-pituitary-thyroid (HET) axis. Crit. Rev. Toxicol. 37: 11-53. Zoeller, R. T, Dowling, A L. S., Herzig, C. T A, Iannacone, E. A., Gauger, K. J. and Bansal, R., 2002. Thyroid hormone, brain development, and the environment. Environ. Health Perspec. 110 (Supplement 3): 355-361.

Zuccotti, M. and Monk, M., 1995. Methylation of the mouse Xist gene in sperm and eggs correlates with imprinted Xist expression and paternal X-inactivation. Nut. Genet. 9: 316-320.

CHAPTER 10

Role of Petrochemicals and Heavy Metals in Amphibian Declines Luca Luiselli and Jerry Lea

I. Introduction II. General Effects of Petrochemical Activities on Terrestrial Fauna

V. Changes in the Composition of Species Assemblages VI. References

Ill. Toxicity of Petrochemicals and Heavy Metals IV. o i l Pollution A. Petrochemical Pollution of Air and its Effect on Terrestrial Fauna Acronyms used in the text: AGlP = Anonima Gas ltaliana Petroli; EN1 = Ente Nazionale Idrocarburi.

I. INTRODUCTION

P

ETROCHEMICAL activity produces pollution that is extremely hazardous to wildlife because of its persistence in the environment, its bioaccumulation in the food chain and its toxicity (Fu et al. 2003). Pollutants may be carried long distances by air, rivers and ocean currents to contaminate regions remote from the source. Most studies of petrochemical pollution have concentrated on the effects on marine life of oil spills at sea, road-traffic pollution, or human health (e.g., the carcinogenic properties of hydrocarbons). Despite the fact that a great deal of petrochemical industrial activity occurs onshore, however, very little is known about the effects of petrochemical pollution on terrestrial organisms, and hardly anything is known about its consequences for amphibians. Because amphibian larvae feed off both the substrate and attached algae, and continuously process water for respiration, they are exposed to a wide variety of pollutants including dissolved toxins, airborne pollution, and sediment contamination (Hall and Mulhern 1984). Below, the focus is on the possible effects of terrestrial petrochemical industrial activity on amphibians. 11. GENERAL EFFECTS OF PETROCHEMICAL ACTIVITIES ON TERRESTRIAL FAUNA

Studies on the direct impact of petrochemical activity on amphibians are very rare in the literature, although there is a generic mention of many frogs being killed by a huge oil spill on the St. Lawrence River (Alexander et al. 1981). This lack of any serious assessments of the consequences of petrochemical activity is very worrying because many tropical areas that are important for petrochemical exploitation are also important "hotspots"

3240

AMPHIBIAN BIOLOGY

of amphibian biodiversity, and also because petrochemical pollution would probably have long-term consequences at the community level for amphibian assemblages. Such long-term consequences on community functioning have been demonstrated in tropical turtle assemblages, where oil spills not only had an immediate and devastating impact on the survival of the various species, but also resulted in major eco-ethological changes in the different populations (Luiselli and Akani 2003). One important region where these threats are current is the Niger Delta in southern Nigeria. With a crude oil production of 2.04 million barrels per day, the Federal Republic of Nigeria is the main oil producer of sub-Saharan Africa and most of Nigeria's oil-industry is found in the Niger Delta region (Luiselli and Akani 2003). The Niger Delta contains several ecologically significant areas, such as swamp-rainforests and mangroves, as well as many endemic species of divergent faunal and floral groups (Kingdon 1990). Unfortunately, the area has been devastated by both deforestation and overpopulation (both related to the economic growth generated by oil companies) and by thousands of oil spills, some of them of huge proportions (Carbone 2003). Akani et al. (1999) studied the herpetofauna of this area as part of a long-term ecological study sponsored in part by Ente Nazionale Idrocarburi (ENI), the parent of Anonima Gas Italiana Petroli (AGIP), the Italian petroleum company. They demonstrated clear negative effects of industrial gas and oil transmission and extraction activities on the biodiversity and abundance of reptiles. Some of the most damaging consequences of the petrochemical activities that they highlighted include: (1) the contamination of wastewater by industrial effluents, which is especially noticeable in areas where local people crack pipelines to extract oil to sell on the black market, (2) destruction of forests and modification of river characteristics (e.g., the destruction of bank plants and aquatic vegetation and the alteration of river courses) to facilitate the placement of pipelines, which results in habitat loss and fragmentation, (3) direct mortality of terrestrial fauna as a result of heavy machinery and insensitive working practices, (4) acoustic disturbance (by men and machinery) and local habitat-alteration, which possibly affects the territorial or breeding behaviour of individual animals, and ( 5 ) the creation of access roads into forest and mangrove sites, which encourages people to utilize these new areas to establish cultivation-plots, markets and settlements. Akani et al. (1999) pointed out that patches of primary and secondary rainforest in the Niger Delta contain the highest diversity and density of reptiles in West-Central Africa. They therefore suggested that petrochemical companies should avoid laying their pipelines through these areas and should instead locate them on land that is already developed or cultivated. Unfortunately, current studies show that the oil companies have generally ignored these suggestions, so it is very likely that the negative effects on the general environment of the Niger Delta (and on its amphibian fauna) will continue in years to come. In addition to the problems listed above, community-development programmes (which petrochemical companies establish in areas where they work) foster immigration of people. This brings further threats to faunal communities in the form of habitat destruction for urban expansion, and also increases hunting pressure for the bush-meat market. West African amphibians (e.g., Ptychadena, Hoplobatrachus spp.) form a major component of the bushmeat diet of many indigenous people (see for example Akani et al. 1998; Eniang et al. 2002). 111. TOXICITY OF PETROCHEMICALS AND HEAVY METALS

Effluents from petrochemical industries contain different groups of non-related organic substances that can be highly toxic in isolation (Bridik et al. 1979). These chemicals are additive, however, and in combination have toxic effects on Xenopus laevis at extremely low levels that individually are non-toxic. This occurs regardless of differing modes of action of the various chemicals (Zwart and Slooff 1987). Other toxins produced from petrochemical industries are non-essential heavy metals that commonly accumulate in soils and bodies of water around installations. These become mobilized when soils become acidic (Freda 1991) and are toxic when they compete with essential metals and each other in transportation

LUISELLI and LEA: PETROCHEMICALS AND HEAVY METALS IN AMPHIBIAN DECLINES

3141

and metabolic processes (Anderson et al. 1997). Zwart and Slooff (1987) specifically examined mixtures of petrochemicals and heavy metals and found such combinations to be particularly dangerous to Xenopus laevis . Petroleum-laden effluents are a real problem, not just because of their toxicity, but also because of their persistence in the environment. For instance, petroleum residues that have an impact on the quality of life of native fauna were still present in the sediment of a Louisiana waterway more than 25 years after the discharge of petroleum wastes had been discontinued (Anderson et al. 1997). In that waterway, high levels of both hydrocarbons and non-essential heavy metals had accumulated in the tissues of crayfish that had been in contact with sediments. The accumulation of these metals (lead, cadmium, aluminium and arsenic) reflected environmental contamination levels and was especially concentrated in the gills. This study is particularly worrying with regard to the effects of this kind of pollution on amphibians because of the high contact amphibians have with the substrate at all stages of their lifecycle (Hall and Mulhern 1984). If tadpoles accumulate these substances in their gills, then gaseous exchange and the regulation of ion fluxes could be affected, while in metamorphs or adults these poisons could impinge upon cutaneous respiration and osmoregulation as well as other vital processes. The toxic effects may or may not be lethal, but even if they only have a slight impact on resource acquisition or energy allocation, then growth, reproduction and ultimately population structure could suffer. Such non-lethal effects of pollution have been demonstrated in tadpoles of R a m c a t e s b e k (Rowe et al. 1996). One of the most persistent petrochemical pollutants is lead, with a half-life in soils of about 20 years (Nriagu 1978). Berzins and Bundy (2002) found that lead in water and sediment at a former petrochemical disposal-area was actively taken up by, and caused suppressed development in, Xenopus laevis tadpoles. Suppressed larval development at worst can lead to larval death and at best to reduced adult fecundity (for example, see Smith 1987), and so again it would affect population structure. Lead also causes a wide range of amphibian deformities including underdeveloped gills, sigmoidal bodies and stunted tails in Bufo arenarum embryos (Perez-Coll and Herkovits 1990) and neural tube defects, tail curvatures and lordoscoliosis in Xenopus laevis tadpoles (Sobotka and Rahwan 1995). Additionally, lead inhibits learning and memory in Rana catesbiuna (Strickler-Shaw and Taylor 1991) and inhibits avoidance of predators by Rana luteiventris (Lefcort et al. 1998). Moreover, some amphibians show no avoidance of lead-enriched waters (Steele et al. 1991), which could lead to chronic exposure. Such long-term exposure to lead pollution has been demonstrated in Bufo arenarum near a petrochemical complex in Argentina, where the average content of lead in the blood of the toads remained constant over three years, indicating a cycle of sustained air-water-soil pollution (Arrieta et al. 2001). In that study, environmental lead concentrations were 20-50 times higher than the maximum level permitted under Argentine legislation for the protection of freshwater life. Lead pollution from the petrochemical industry could therefore be a major factor in the health of many amphibian populations. IV. OIL POLLUTION There is little evidence of the direct effects of oil pollution on amphibians except for the studies by Alexander et al. (1981) and Lefcort et ul. (1997) on tiger salamanders. In the latter paper, the authors found that larvae living in silty and oil-polluted ponds metamorphosed earlier and at a smaller size than did salamanders living in pristine ponds. As mentioned above, these stresses on larval growth and development can profoundly alter adult population structure and, given the regularity of pipeline oil-leaks or sabotage in many developing countries, one must assume that oil pollution probably directly affects many aniphibian populations. A, Petrochemical Pollution of Air and its Effect on Terrestrial Fauna

Examples were given above of how direct petrochemical pollution of soil and water affects amphibian populations. Not previously mentioned, however, are the consequences

3242

AMPHIBIAN BIOLOGY

of air pollution by petrochemicals; these may indeed be a serious threat to amphibians. For instance, the volume of hazardous emissions into the atmosphere (including phenol, hydrogen sulphide, aromatic hydrocarbons and other toxins) exceeds 30 000 metric tons per year from just one Russian petrochemical complex (Naidenko and Grechkanev 2002). This situation is likely to be reflected in many ecologically sensitive areas in the world where petrochemical production occurs. These emissions are taken up by vegetation and eventually find their way into the soil to be accumulated by organisms that participate in the destruction and mineralization of proteins and carbohydrates (Khot'ko and Vetrova 1982). Petrochemical air pollutants have been shown to stunt the growth of ground beetles either through direct soil contact, through consumption of pollutants with their food or via inhaling vapours (Naidenko and Grechkanev 2002). As amphibians are predators of soil entomofauna they are likely to build up high levels of airborne toxins through their diet, as well as through the active uptake of toxins directly from the soil.

M CHANGES I N THE COMPOSITION OF SPECIES-ASSEMBLAGES So far, mention has been made only of the negative effects of petrochemical activity on arnphibian populations. It should be pointed out, however, that habitat change due to petrochemical expansion does not necessarily lead to a reduction in the size of anuran species-assemblages. On the contrary, Lea et al. (2002) found that, although forest specialists disappeared, the destruction of more than 90% of tropical riparian forest as a result of petrochemical expansion (over a twenty-year period) actually led to a greater diversity of amphibians. Destruction of rainforest is not advocated; it is merely pointed out that marly amphibians adapt to changing environments (in this case the increase in species-richness was due to a flood of ranids occupying the newly formed mosaic landscape). VI. REFERENCES Akani, G. C., Luiselli, L., Angelici, F. M. and Politano, E., 1998. Bushmen and herpetofauna: notes on amphibians traded in bush-meat markets of local people in the Niger Delta (Port Harcourt, Rivers State, Nigeria). Anthropozoologica 27: 21-26.

Eniang, E. A,, King, R., Lea, J., Capizzi, D. and Luiselli, L., 2002. Trophic Niches of four sympatric rainforest anurans from southern Nigeria: Does resource partitioning play a role in structuring the community? Revue d'Ecologie (Terre et Vw)57: 19-28.

Akani, G. C., Luiselli, L. and Politano, E., 1999. Ecological and conservation considerations on the reptile fauna of the eastern Niger Delta (Nigeria). Herpetozoa 11: 141-153.

Freda, J., 1991. The effects of Aluminium and other metals on amphibians. Environ. Pollut. 71: 305-328.

Alexander, M. M., Longabucco, P. and Phillips, D. M., 1981. The impact of oil on marsh communities in the St. Lawrence River. Pp. 333-340 in "Oil-Spill Conference". American Petroleum Institute, Washington D.C. Anderson, M. B., Reddy, I!, Preslan, J . E., Fingerman, M., Bollinger, J., Jolibois, L., Maheshwarudu, G. and G e o ~ eW , . -I., 1997. Metal accumulation in crayfish, Procambarus clarkii, exposed to a petroleumcontaminated bayou in Louisiana. Ecotoxicol. Environ. Safety 37: 267-272.

-

Arrieta, M. A., Apartin, C., Rosenberg, C. E., Fink, N. E. and Salibian, A,, 2001. Blood lead content in a peri-urban population of the South American toad Bufo arenarum. Sci. Total Environ. 271: 99-105. Berzins, D. W. and Bundy, K. J., 2002. Bioaccumulation of lead in Xenopus laevis tadpoles from water and sediment. Environ. Internat. 28: 69-77. Bridie, A. L., Wolfe, C. J. M. and Winter, M., 1979. The acute toxicity of some petrochemicals to gold fish. Water Res. 13: 623-626. Carbone, M., 2003. Morte di un delta. Airone 260: 52-60.

Fu, J., Mai, B., Sheng, G., Zhang, G., Wang, X., Peng, E, Xiao, X., Ran, R., Cheng, F., Peng, X., Wang, Z. and Tang, U. W., 2003. Persistent oragnic pollutants in environment of the Pear; River Delta, China: an overview. Chemosphere 52: 141 1-1422. Hall, R. J. and Mulhern, B. M., 1984. Are anuran amphibians heavy metal accumulators? Pp. 445466 in "Vertebrate Ecology and Systematics, a Tribute to Henry S. Fitch, ed by R. A. Seigel, L. E. Hunt, J. L. Knight, L. Malaret and N. L. Zuschlag. Universitv of Kansas Museum of Natural Histow. Lawrence. Khot'ko, E. I. and Vetrova, A. O., 1982. On the influence of industrial emissions on the forest entomofauna. Ekol. Zashch. Lesa 3: 19-22. Kingdon, J., 1990. "Island Africa". Academic Press, New York. Lea, J. M., Politano, E. and Luiselli, L. 2002. Changes in the herpetofauna of a fresh water river in Southern Nigeria, after 20 years of development. Rus. J. Herp. 10: 191-198. Lefcort, H., Hancock, K., Maur, K. and Rostal, D., 1997. The effects of used motor oil and silt on the growth, survival, and the ability to detect predators by tiger salamanders, Ambystoma tigrinum. Arch. Envron. Contam. Toxicol. 32: 383-388.

LUISELLI and LEA: PETROCHEMICALS AND HEAVY METALS IN AMPHIBIAN DECLINES Lefcort, H., Mepire, R. A., Wilson, L. H. and Ettinger, W. F., 1998. Heavy metals alter the survival, growth, metamorphosis, and antipredatory behavior of Columbia Spotted Frog (Rana luteiuentris) tadpoles. Arch. Environ. Contam. Toxicol. 35: 447-456. Luiselli, L. and Akani, G. C., 2003. An indirect assessment of the effects of oil pollution on the diversity and functioning of turtle communities in the Niger Delta, Nigeria. Anim. Biodiv. Cons. 26: 57-65. Naidenko, V V and Grechkanev, 0 . M., 2002. The state of components of the biota as an indicator of disturbances in natural ecosystems exposed to petrochemical pollution. R w . J. Ecol. 33: 62-64.

3243

Rowe, C. L., knney, 0. M., Fiori, A. F? and Congdon, J. D., 1996. Oral deformities in tadpoles (Rana catesbeiana)associated with coal ash deposition: effects on grazing ability and growth. Ereshw. Bwl. 36: 723-730. Smith, D. C., 1987. Adult recruitment in chorus frogs: effects of size and date of metamorphosis. Ecology 68: 344-350. Sobotka, J. M. and Rahwan, R. G., 1995. Teratogenesis induced by short- and long-term exposure of Xenopm lueuis progeny to lead. j. Toxicol. Environ. Health 44: 469-484. Steele, C. W., Strickler-Shaw, S. and Taylor, D. H., 1991. Failure of Bufo americanw tadpoles to avoid leadenriched water. J. Herpetol. 25: 241-243.

Nriagu, J. O., 1978. Lead in soils, sediment and major rock types. Pp. 15-72 in "The Biogeochemistry of Lead in the Environment: Part A. Ecological Cycles", ed by J. 0 . Nriagu. Elsevier/North-Holland Biomedical Press, New York.

Strickler-Shaw, S. and Taylor, D. H., 1991. Lead inhibits aquisition and retention learning in bullfrog tadpoles. Neurotoxicol. Teratol. 13: 167-173.

Perez-Coll, C. S. and Herkov~ts,J., 1990. Stage dependent suscepib~lityto lead in Bufo arenarum embryos. Environ. Pollut. 62: 239-245.

Zwart, D. D. and Slooff, W., 1987. Toxicity of mixtures of heavy metals and petrochemicals to Xenopw laevis. Bull. Enuiron. Contam. Toxicol. 38: 345-35 1.

,

CHAPTER 11

Acidification and its Effects on Amphibian Populations Katja Rasanen and David M. Green

I. Introduction II. Acidification A Temporal and Spatial Trends B. Correlated Abiotic and Biotic Changes Ill. Effects of Acidification on Amphibians A. Effects of Acidity at the Level of the Individual 1. Direct Effects 2. Carry-Over Effects 3. Interactive Effects 8. Are Laboratory Results Observable in the Wild? C. Effects at the Level of the Population

V. Spatial and Temporal Relationships in the Wild A. Amphibian Distribution 6. Local Long-Term Patterns C. Global Trends VI. Amphibians as Indicators of Environmental Acidification VII. Conclusions: Acidification as a Cause of Amphibian Decline VIII. Acknowledgements IX. References

IV. Mechanisms for Counteracting the Negative Effects of Acidity A. Migration and Acclimation 6. Evolutionary Adaptation 1. Genetic Variation 2. Maternal Effects 3. Life-History Variation Abbreviations and acronyms used in the text and references: AMAP = Arctic Monitoring and Assessment Programme, Oslo; DOC = dissolved organic carbon; UV-B = ultraviolet-B.

I. INTRODUCTION

N

ATURAL environments vary greatly in pH (hydrogen ion [H+]concentration) ranging from alkaline sites on calcareous soils to acidic soils and black waters. Naturally acidic environments are common and result mainly from high concentration of organic acids. They are formed gradually, over hundreds or thousands of years (Renberg et al. 1993; Battarbee and Charles 1994) thereby allowing organisms to adapt to them (Collier et al. 1990; Dangles et al. 2004). Over the past hundred years or so, however, large parts of the world have become acidified as a result of industrialization (Brodin 1993; Mannion 1999). Consequently acidification emerged as an environmental issue of great concern in the 1960s, particularly in Scandinavia and in the northeastern parts of North America (Brodin 1993; Mannion 1999). Anthropogenic acidification occurs rapidly, over time frames of years or decades (AMAP 1998). It affects terrestrial ecosystems to some degree but the effects are most severe in freshwater ecosystems (Schindler 1988; Pleijel et al. 1999). Anthropogenic acidification

&ANEN

and GREEN: ACIDIFICATION AND AMPHIBIANS

3245

allows little time for most organisms to adapt and hence its negative effects can be far more acute than are those of natural acidification (Collier et al. 1990; Dangles et al. 2004). Furthermore, although measures taken to reduce emissions of acidifying compounds have had some effect (Stoddard et al. 1999), there is still little evidence for biological recovery, and acidification will probably remain an environmental problem for years to come (Alewell et al. 2000; Pleijel et al. 1999). Human induced acidification has detrimental effects on organisms at all taxonomic levels, and in both terrestrial and aquatic environments (Haines 1981; Schindler 1988; Huckabee et al. 1989; Rusek and Marshall 2000). Amphibians are one of the affected groups (Pierce 1985; Freda 1986). Gosner and Black (1957), who compared tolerance to acidity among a large number of North American amphibian species, were among the first to realize that environmental acidity may adversely affect amphibian populations. In the midst of increasing concern about acidification during the 1980s, many studies were conducted on amphibian tolerance to acidity and, when in the early 1990s an alert was raised about global amphibian declines, environmental acidification was suggested as one of the major causes (Blaustein and Wake 1990; Wake 1991). This impetus for research has so far resulted in over 100 published case studies and several reviews (Pierce 1985, 1993; Freda 1986, 1991; Bohmer and Rahmann 1990; Rowe and Freda 2000). Most case studies of amphibians have dealt with direct toxicity of acidity to aquatic embryos and larvae. Overall, however, the studies range from examining the physiological and genetic basis to behavioural responses, as well as to attempts to explore effects at the population level (Freda 1986; Rowe and Freda 2000). To shed light on whether anthropogenic acidification is a major agent of amphibian decline, the currently available evidence is reviewed. The main effects of acidification acting on amphibians at the individual level are considered first and then the evidence for effects of acidification on wild populations is examined. On the basis of existing evidence, it is concluded that acidification has a multitude of negative effects on amphibians but that it is probably not a major factor contributing to global amphibian decline. It is clear, however, that acidification may contribute to local declines in amphibian populations, especially in combination with other threats. 11. ACIDIFICATION A. Temporal and Spatial Trends Anthropogenic acidification is mainly due to emissions of sulphur dioxide (SO2)and nitrogen oxides (NO,) to the atmosphere via the combustion of fossil fuels. This results in acidic precipitation, i.e., "acid rain". The term "acid rain" was first coined by Robert Angus Smith in 1852 after surveying the polluted air of Manchester, England, and refers to precipitation with a pH below that of the 5.6 of uncontaminated rain water (Mannion 1999). Although natural sources, such as volcanic eruptions and algae in oceanic surface water, also release acidifying compounds into the atmosphere, the levels are very low compared to those produced by industrial activities (AMAP 1998; Mannion 1999). In the long term, acidification is a gradual phenomenon that results in reduced pH over several years or decades (Renberg et al. 1993; Battarbee and Charles 1994). In the short term, however, acidification is often episodic. In areas where acid rain is a concern, severe declines in pH can occur over a few hours during periods of rainfall and snow melt, even in habitats that at other times are nearly neutral. The net effect of acid rain depends on both the buffering capacity of the local environment and on the amount of acid deposited. Alkaline soils with naturally high pH and high levels of calcium carbonate or magnesium carbonate have a high acid-neutralizing capacity. On the other hand, acidic bedrock composed of granites or naturally acidic soils have little or no capacity to buffer additional deposition of acid. These soils thus become progressively poorer and more acidic (Mannion 1999). The distribution of affected habitats

3246

AMPHIBIAN BIOLOGY

depends also to a large degree on the locations of sources of acidifying emissions. These coincide with areas of industrial activity in developed and developing countries (Mannion 1999) and are largely in the Northern Hemisphere, particularly eastern North America, Europe, India and eastern Asia. The distribution of acidic deposition, coupled with the geographic variation in buffering capacity of soil and water, results in a patchwork of regions affected to greater or lesser degrees by acidifying precipitation. The occurrence of naturally acid-sensitive soils and lakes is more widespread and includes eastern and northern North America, northern Europe, and the tropical regions of South America, Africa and Indonesia. Thus, although acidification occurs locally in many places throughout the world, the phenomenon is of greatest importance where there is a combination of acidic precipitation and a lack of buffering capacity, such as in eastern North America and northern Europe. B. Correlated Abiotic and Biotic Changes A range of abiotic and biotic environmental changes are directly related to acidification, foremost of these being elevated levels of metals. Under acidic conditions, metal ions are more soluble and leach from the soil into surface waters (Brodin 1993; Mannion 1999). Solubility, especially of aluminium (Al), is strongly increased as pH of the water declines below 5. In addition, dissolved organic carbon (DOC) levels are often reduced in acidified waters (Schindler et al. 1996), which results in low levels of complexation of metals. Hence, A1 is often present in its most toxic form, inorganic monomeric aluminium. Furthermore, as DOC levels decrease, acidified waters become clearer and permit transmission of increased levels of detrimental W-B radiation in the water column (Schindler et al. 1996). As DOC levels can also be reduced over the course of global warming, in areas where acidification is a problem these major environmental changes may be tightly correlated and expose populations to a combined threat of acidification, metal toxicity, increased W-B and changing temperature. Acidification has strong negative effects on plants and animals both in terrestrial and aquatic environments (Brodin 1993; Steinberg and Wright 1994). The many changes that occur range from altered algal composition and disappearance of small crustaceans, such as Daphnza, to disappearance of fish and to an increase in abundance of acid-tolerant insect predators. As a result of the differential sensitivity of species, acidification results in large shifts within ecosystems (Brakke et al. 1994). Together with the many abiotic changes, the potential threats of acidification on organisms, hence, consist of complex interactions among a multitude of abiotic and biotic changes. 111. EFFECTS OF ACIDIFICATION ON AMPHIBIANS Amphibians occupy a large range of terrestrial and aquatic environments (temporary ponds to lakes and small streams), which also encompass a large variation in pH (Freda 1991; Rowe and Freda 2000). Both the terrestrial and aquatic habitats used by amphibians can suffer locally and globally from acidification. Even natural levels of acidity limit amphibian distribution and abundance (e.g., Gosner and Black 1957; Saber and Dunson 1978; Freda and Dunson 1986a; Bradford et al. 1998). However, the increased spatial and temporal variation in pH due to anthropogenic acidification result in pH levels that are both much lower and more rapidly changing than at naturally acidic sites. Furthermore, temporary ponds and small streams, which are important breeding habitats of amphibians, rely on rainwater or melted snow. At such sites, rapid, periodic declines in pH are especially common (e.g., Freda et al. 1991; Sadinski and Dunson 1992). A. Effects of Acidity at the Level of the Individual The potential sensitivity of amphibians to low pH and other environmental perturbations stems to a large extent from their complex life histories and their physiological characteristics, especially the respiratory function of their permeable skin. Due to their biphasic life cycle, many amphibians are exposed to environmental acidity during at least one significant lifehistory stage. For instance, in areas with cold, snowy winters, acidity of surface water may

&ANEN

and GREEN: ACIDIFICATION AND AMPHIBIANS

3247

peak during the spring snow melt (Corn 2003). In such areas, embryos of species that breed early in the spring, such as R a m sylvatica and R. temporark, may have a high risk of exposure to acidic conditions (but see Vertucci and Corn 1996). Likewise, heavy rainfalls resulting in rapid declines in pH can occur at any time and hence expose any life stage to stressful conditions. The effect of such strong temporal decline in pH is most severe in species breeding in seasonal ponds or in small streams (Freda et al. 1991) but is pertinent also for species that breed at permanently acidified sites. Through acidification of soils even terrestrial amphibians, such as the juveniles of Notophthalmus virzdescens and adults of Plethodon cinereus (Wyman and Hawksley-Lescault 198'7) may be exposed to low pH (Rowe and Freda 2000). 1 . Direct Efects A. EMBRYONIC STAGES

The negative effects of low pH may begin at fertilization, although the evidence is currently inconclusive. Schlichter (1981) found that acidic conditions inhibited sperm motility in Rana pipiens, potentially impairing fertilization success. No reduced fertilization under acidic conditions, however, was observed in R . arvalis (AndrCn et al. 1988) or in R. temporaria (Beattie et al. 1991). In addition, Dale et al. (1985a) found the acetate buffer used by Schlichter (1981) to be toxic to Xenopus laevis, which may have biased Schlichter's results. The most severe direct effect of acidity is its impact on embryonic mortality, which carries with it potentially devastating immediate consequences for reproductive success. In accordance with early observations by Gosner and Black (195'7), most studies since have aimed at finding the critical (50% mortality) and lethal (100% mortality) pH levels for embryos (Rowe and Freda 2000). In most amphibians, these range from pH 3.8 to pH 5.5, and pH 3.5 to 4.3, respectively (Table l), denoting reasonable tolerance to acid compared to that of many other organisms. The lethality of acidity to embryos is a sum of various effects. Firstly, extreme acidity may disrupt development at early embryonic stages (Leuven et al. 1986; Haidacher and Fachbach 1991) and result in abnormalities, such as failure of the yolk plug to retract (Tome and Pough 1982; Haidacher and Fachbach 1991). The most common cause of embryonic mortality under acidic conditions, however, seems to be the "curling defect", whereby embryos may develop normally but curl and fail to hatch (Gosner and Black 1957; Salthe 1963; Pough and Wilson 1977; Dunson and Connell 1982). The tight coiling during development also results in abnormalities in spine and tail in those embryos that do manage to hatch (Tyler-Jones et al. 1989; Haidacher and Fachbach 1991). These abnormalities severely affect survival (Beattie et al. 1992). The curling defect of embryos apparently results from shrinkage of the egg capsules together with reduced embryonic movements and malfunctioning of the hatching enzyme. In acidic water, parts of the gelatinous egg capsule often become opaque and have reduced pliability, indicating changes in the mechanical or chemical properties of the jelly coat (Pierce 1985; Picker et al. 1993; Riisanen et al. 2003b). Hatching-enzymes may fail to function properly below an optimal pH, which at least in Xenopus laevis (Urch and Hedrick 1981) and Rana pirica (Kitamura and Katagiri 1998) is about pH 7. Removal of jelly capsules strongly increases embryonic survival at low pH (Dunson and Connell 1992; Picker et al. 1993; E s a n e n et al. 2003b), suggesting that very low pH may affect the egg capsules more than it does the embryos themselves. B. LARVAL STAGES

Slow growth and development is a general response to sub-lethal acidity in both embryos and larvae. In embryos, this generally results in delayed hatching at a smaller size (Rowe and Freda 2000; Rasanen et al. 2003a). In Triturus vulgaris, however, hatching occurred earlier under acidic conditions, and at an earlier developmental stage (Griffiths et al. 1993). Larvae or tadpoles are usually more tolerant than are embryos. At levels causing significant embryonic mortality, effects on larvae are mainly sub-lethal and are manifest as reduced

3248

AMPHIBIAN BIOLOGY

Table 1. Minimum and maximum values reported for embryonic mortality in laboratory studies. "Lethal pH" refers to the pH at which 85-100% mortality occurs, and "Critical pH" refers to the pH at which 50% mortality occurs. - = not reported.

Species PELOBATIDAE Scaphiopus holbrookii Scaphbpus intermontanus MICROHYLIDAE Gastrophryne carolinensis Microhyla ornata HYLIDAE Acris gryllus Hyla andersonii Hyla arborea Hyla crucqer Hyla chrysoscelis Hyla femoralis Hyla gratiosa Hyla squirella Hyla versicolor Pseudacris crucqer Pseudacris ornata Pseudacrk nigrita Pseudacris triseriata PIPIDAE Xenopus gzlli Xenopus laevis

Lethal pH

Critical pH

Reference

4.2 4.3

Warner and Dunson (1998) Pierce (1991) in Pierce (1993)

4.2 3.7

Warner and Dunson (1998) Padhye and Ghate (1988) Gosner and Black (1957) Gosner and Black (1957); Freda and Dunson (1986a) Warner et al. (199 1); Haidacher and Fachbach (1991) Gosner and Black (1957); Karns (1983); Dale et al. (1985a) Warner and Dunson (1998) Warner and Dunson (1998) Grant and Licht (1993); Warner and Dunson (1998) Warner and Dunson (1998) Gosner and Black (1957) Dale et al. (1985a) Warner and Dunson (1998) Gosner and Black (1957) Corn et al. (1989); Karns (1992)

3.8 3.5-5

RANIDAE Rana arvalis

Picker et al. (1993) Saber and Dunson (1978); Tome and Pough (1982); Dale st al. (1985a); Dunson and Connell (1992); Picker et a1. (1993) Leuven et al. (1986); Andrkn et al. (1988, 1989); Esanen et al. (2003a) Gosner and Black (1957); Saber and Dunson (1978); Grant and Licht (1993) Saber and Dunson (1978); Freda and Dunson (1986a) Andrkn et al. (1988) Leuven et al. (1986) Bradford et al. (1992) Gosner and Black 1957, Dale et al. (1985a) Karns (1983); Freda and Dunson (1985b); Corn et al. (1989); Freda and McDonald (1990); Smith et al. (1990); Freda et al. (1990a); Long et al. (1995) Haidacher and Fachbach (1991) Gosner and Black (1957); Warner and Dunson (1998) Gosner and Black (1957), Tome and Pough (1982), Karns (1983); Pierce et al. (1984); Clark and LaZerte (1985); Clark and Hall (1985); Freda and Dunson (1985b); Dale et al. (1985a); Pierce and Sikand (1985); Pierce and Harvey (1987); Corn et al. (1989); Sadinski and Dunson (1992*); Karns (1992); Grant and Licht (1993) Beebee and Gran (1977); Leuven et al. (1986); Gebhardt et al. (1987); Andren et al. (1988); Haidacher and Fachbach (1991) Gosner and Black (1957)

Rana catesbeiana Runa clamitans Rana dalmatina Rana esculenta Runa muscosa Rana palustrk Rana pipiens Rana ridibunda Rana sphenocephala Runa sylvatica

Rana temporaria Rana virgatipes BUFONIDAE Bufo americanus

4.04.3

Bufo boreas

44.5

Bufo bufo Bufo calamita Bufo canorus Bufo punctatus Bufo woodhousii BOMBINATORIDAE Bombina variegata AMBYSTOMATIDAE Ambystoma jeffmsoninnum

4.44.6 4.8 4.6 4.0 4.0-4.2

Freda and Dunson (1985b); Dale et al. (1985a); Clark and LaZerte (1985, 1987); Karns (1983); ~ r e d aand McDonald (1990) porter and Hakanson (1976, in Freda et al. [1991]); Corn et al. (1989) Leuven et (11. (1986); Haidacher and Fachbach (1991) Beebee and Griffin (1977); Beebee (1986) Bradford et al. (1992) Pierce (1991) in Pierce (1993) Freda and Dunson (198513, 1986); Karns (1983)

4.6

Haidacher and Fachbach (1991)

4.04.6

Pough and Wilson (1977); Cook (1978, in Leuven et al. [1987]); %me and Pough (1982); Freda and Dunson (198513); Sadinski and Dunson (1992)*

RASANEN and GREEN: ACIDIFICATION AND AMPHIBIANS

3249

Table 1 - continued

Species

Lethal DH

Critical pH

AMBYSTOMATIDAE - continued Ambystoma laterale 4.2 Ambystoma maculatum 4.04.5

Ambystoma texanum Ambystoma t i p i n u m SALAMANDRIDAE Triturus alpestris Triturus cristatw Triturw helveticw Triturus vulparis

Reference Karns (1992) Pough (1976); Pough and Wilson (1977); Cook (1978); Dale et al. (1985a); Freda and Dunson (198513); Clark and LaZerte (1985); Blem and Blem (1989); Smith (1990); Sadinski and Dunson (1992)* Punzo (1983); Pierce and Wooten (1992b) Whiteman et al. (1995) Bohmer 1988; Bohmer et a1 (1988) Haidacher and Fachbach (1991); Griffiths et al. (1993) Bohmer (1988) Bohmer (1988); Haidacher and Fachbach (1991)

*De-jellied eggs used, which may affect tolerance (Dunson and Connell 1992; Picker et al. 1993; Esanen et al. 2003b).

rates of development and growth as well as delayed metamorphosis at a smaller size (Freda and Dunson 1985a; Rowe and Freda 2000). Impaired rates of larval growth and development have been reported for many amphibian species, including Ambystoma maculatum (Rowe et al. 1992; Sadinski and Dunson 1992), Bufo canorus (Bradford et al. 1992), Bufo fowleri and Hyla andersonii (Freda and Dunson 1986b), Hyla gratiosa (Warner et al. 1991), Rana temporaria (Cummins 1986, 1989; Beattie and Tyler-Jones 1992; R%sanenet al. 2002), Rana arvalis (Rasanen et al. 2003a) and Rana sylvatica (Rowe et al. 1992; Horne and Dunson 1995b). Larval tolerance tends to increase with larval age and size (Pierce et al. 1984; Freda and Dunson 1985a; Bohmer and Rahmann 1990; Verma and Pierce 1994). Low pH can also alter behaviour of larval amphibians. Commonly, embryos and tadpoles are less active under acidic conditions (Freda and Taylor 1992; Kutka 1994). These effects can differ among species. In Triturus helveticus and ?: vulgaris (Griffiths 1993), Ambystoma maculatum (Preest 1993) and Rana temporaria (Rasanen et al. 2002) reduced activity was manifest as reduced feeding rates, whereas such response was not seen in ?: cristatus (Griffiths 1993). Due to reduced activity and possibly impaired co-ordination, acidic conditions can also affect the ability of tadpoles to avoid predators. For instance, the ability of Hyla cinerea tadpoles to avoid predation by dragonfly larvae was impaired under acidic conditions (Jung and Jagoe 1995). On the other hand, acidic conditions can lower the effectiveness of predators; Ambystoma maculatum displayed reduced ability to capture prey (Daphnia) under acidic conditions (Preest 1993). C. ADULTS AND TERRESTRIAL STAGES

Relatively little is known about how exposure to acidic conditions affects terrestrial stages of amphibians. In some situations at least, even adult survival may be reduced at low pH. For instance, when exposed to moderate acidity (pH 5.5), Rana pipiens adults experienced 100% and 58% mortality, respectively, when recently emerged from hibernation and just after breeding (Vatnick et al. 1999). Plethodon cinereus adults died within a month at pH 2.5-3.0 and within eight months at pHs between 3.0 and 4.0 (Wyman and Hawksley-Lescault 1987). Not all adult amphibians are equally strongly affected, however. Rana temporaria adults did not suffer increased mortality when exposed to pH 4.0 during over-wintering (Pasanen et al. 1998). In stresshl environments, individuals may need to allocate more energy to maintenance, growth and tolerance, and hence have fewer resources available for reproduction (Maltby 1999). Evidence from fish suggests that reproduction and age structure can be adversely affected by acidity (Haines 1981). For instance, white sucker, Catostomus commersoni, from acid lakes matured at older age and had shorter reproductive life-spans than did those from circumneutral lakes (Trippel and Harvey 1987). Low pH can affect the reproductive performance of amphibians also. For example, females make a major physiological investment in egg production and there may be a trade-off between maintenance and

3250

AMPHIBIAN BIOLOGY

reproduction under acidic conditions. Accordingly, females of Rana arvalis from acidic habitats have reduced fecundity, and both sexes tend to have delayed age, slower growth and smaller size at first reproduction compared to individuals from circumneutral habitats (Esanen 2002; Soderman 2006; Soderman et al., 2007). Very little attention has been paid to such effects so far, however, and more studies are needed before one can understand the effects of acidification on the whole life cycle. The physiological basis of the effects of acidity on larval and terrestrial stages primarily seems to result from the disruption of ionic balance (Freda and Dunson 1984; Ireland 199l), in particular sodium balance. In tadpoles and salamander larvae, sodium efnux is increased and sodium uptake reduced under acidic conditions, resulting in death when about half of the body sodium has been lost (Freda and Dunson 1984). Sodium balance is disrupted at pH 3.54.5 in recently metamorphosed Ambystoma jeffersonkznum (Horne and Dunson 199413) and at pH 3.0, compared to pH 5.0, in adult Plethodon cinereus (Frisbie and Wyman 1995). In Bufo americanus embryos and larvae, moderate acidity (pH 6.0) disrupted nitrogen balance (Tattersall and Wright 1996). Other physiological effects of acidity on terrestrial amphibians include disruption of immune function and water balance. For instance, adult Rana pipiens exposed to pH 5.5 had bacterial infections in the spleen and had lowered splenic blood counts (Simon et al. 2002; Brodkin et al. 2003), suggesting that immune functions are suppressed under acid stress (Carey 1993). In addition, terrestrial amphibians and tadpoles lose body water under acidic conditions, possibly affecting their performance at acidic sites (Frisbie and Wyman 1991; Horne and Dunson 1994b). 2. Carry-Over Effects

The negative effects of stressful environmental conditions during one stage of life can carry-over to later life, possibly with far-reaching consequences for fitness (Pechenik et al. 1998; Lindstrom 1999). Similarly, exposure to acidity during one life stage can affect future performance in amphibians (Bradford et al. 1992; Rasanen et al. 2002). Such effects have been found in amphibian responses to other stressors, such as UV-B radiation (Pahkala et al. 2001a), but few studies have explored potential carry-over from exposure to acidity. In Ambystoma maculatum, a short-term increase in acidity during the embryonic stage reduced survival three weeks later (Clark and Hall 1985). However, a three-day exposure of Xenopw laevis, Bufo valliceps, and B. woodhousii tadpoles to pH 4 reduced growth rates of tadpoles, but the effects were no longer observable a week after the exposure to acidity was terminated (Pierce and Montgomery 1989). Exposure of embryos of Rana temporaria to moderately acidic conditions (pH 4.5) likewise had no negative effects on the size or age at which metamorphosis occurred (Esanen et al. 2002). These results suggest that, at least when resources are unlimited and when conditions are otherwise benign, amphibians may be able to compensate for the negative effects of short-term exposure to acidity. More information, however, is needed on potential carry-over effects. For example, because amphibians often are exposed to acidic conditions during their aquatic stages, studies investigating how this exposure affects their subsequent terrestrial life would be valuable. 3. Interactive Effects A. ABIOTIC INTERACTIONS

Numerous abiotic factors interact with low pH and can either accentuate or ameliorate its effects. The main factors identified to date are water chemistry, UV radiation and temperature. Due to their correlated nature (Schindler et al. 1996) these interactive effects could potentially have major effects on amphibians during ongoing climatic changes. Water Chemistry: Water chemistry has a major influence on the toxicity of H+ ions. As mentioned above, various metals increase in toxicity at low pH (Spry and Wiener 1991; Linder and Grillitsch 2000), foremost of these being inorganic monomeric aluminium. Increased levels of Al+++ions increase toxicity of H+ to both fish (Spry and Wiener 1991) and amphibians (Freda 1991). In amphibians, high aluminium ion concentration in

-ANEN

and GREEN: ACIDIFICATION AND AMPHIBLANS

3251

combination with low pH can increase mortality (Dale et al. 1985; Tyler-Jones et al. 1989; Freda and McDonald 1990) and alter behaviour. Exposure of Hyla cinerea tadpoles to high A1 levels impaired swimming behaviour and, in combination with low pH, increased risk of predation by dragonfly larvae (Jung and Jagoe 1995). The effects of aluminium are ions have been found to ameliorate the toxicity complex, however, and in some studies, A+++ of acid (reviewed by Freda 1991). Moreover, although very high levels of DOC in themselves may be directly harmful (Dunson and Connell 1982; Freda and Dunson 1986a; reviewed by Steinberg et al. 2006), aluminium toxicity in acidic waters with high DOC levels (such as many bog habitats) is often lower due to aluminium complexation with various organic compounds (Freda 1991; Spry and Wiener 1991). Amphibian habitats are increasingly exposed to toxic compounds due to abundant use of fertilizers and pesticides (Hecnar 1995; Cowman and Manzati 2000), and potential interactive effects of pH and such toxins may be important in determining the fate of populations simultaneously exposed to these different stressors. The interactive effects between pH and toxins have not been extensively studied. High nitrate levels in combination with low pH increased mortality of Rana cascadae larvae (Hatch and Blaustein 2000). The effects of the fungicide, triphenyltin, on survival, growth and development of Runa lessonae and R. esculenta tadpoles depended on pH but appeared to be less severe at lower pH (pH 6.4 versus pH 8.1) (Fioramonti et al. 1997). Likewise, the negative effect of the herbicide uson@ on survival of R. pipiens larvae was reduced at pH 5.5 compared to pH 7.5 (Chen et al. 2004). Calcium is the most important of the ameliorating water chemistry variables. Increased calcium levels can increase survival at low pH (Dale et al. 1985; Freda and Dunson 1985b; Leuven et al. 1986) and moderate the effect of low pH on sodium balance in larvae (Horne and Dunson 1995b). However, the concentrations of dissolved calcium, or other ameliorating ions, such as magnesium or sodium, that are required to produce a recognizable effect are higher than those normally found in breeding ponds (Rowe and Freda 2000) and hence may only rarely counteract the negative effects of acidity in natural ponds. Liming of lakes and forests is commonly used to treat areas that suffer heavily from acidification (Andersson et al. 2002) and, due to the ameliorating effects of Ca++, this practice may benefit amphibians living in acidified habitats. For instance, liming of acidic ponds increased hatching success of Rana arvalis from 5% to 95% (Bellemakers and VanDam 1992) and of R . ternporaria from below 25% to over 60% (Beattie et al. 1993). The effects of increasing concentrations of calcium, however, depend on the species and may sometimes even be harmful (Dale et al. 1985a). For instance, hatching success of R. sylvatica at pH 4.5 decreased with increasing levels of calcium (from 100% at 10 ppm to 20% at 90 ppm). Furthermore, although liming increases pH, it may result in other unwanted changes in the ecosystem (Andersson et al. 2002), with potentially unpredictable consequences for amphibians. Ultraviolet radiation: Increased ultraviolet radiation has been implicated as a major factor contributing to global amphibian declines (Blaustein and Wake 1990; Alford and Richards 1999). In acidified areas, UV-B effects may become even more pronounced. Because DOC levels decline in the course of acidification, this may result in increased UV-B penetration through the water column (Schindler et al. 1996) and expose amphibians simultaneously to stresses from both low pH and elevated UV-B radiation. So far, few studies have investigated the interactive effects of Iow pH and increased UV-B but strong synergistic effects of these two stressors on embryonic mortality have been implicated. Such effects were found in Rana pipiens (Long et al. 1995), R. temporaria (Pahkala et al. 2002) and R. cascadae (Hatch and Blaustein 2000). In contrast, Runa arvalis exposed to natural UV-B levels at pH 5.0 demonstrated no synergistic effects (Pahkala et al. 2001b), probably because of the relatively moderate exposure levels. Temperature: Toxicity of low pH can increase at physiologically suboptimal cold or warm conditions (Rowe and Freda 2000). In addition, low temperatures slow down development of embryos and larvae and result in longer exposure times to low pH and, hence, potentially have stronger negative effects. As extreme temperatures are commonplace in most natural

3252

AMPHIBIAN BIOLOGY

environments, temperature-dependence of pH effects could have major consequences on organisms. For example, in temperate zone species, breeding begins early in the spring soon after snowmelt when occasional cold periods are common. This can expose embryos simultaneously to both low pH and low temperatures. Embryonic tolerance of Triturus cristatus, 7: helveticus and T. vulgaris to acid, however, was not reduced at 12°C compared to 17°C. On the other hand, exposure of adult Rana pipiens to cold (mimicking emergence from hibernation) followed by exposure to low pH (but not to cold alone) induced bacterial infection (Brodkin et al. 2003). At the other end of the temperature scale, the combination of warm water (36-39°C) and low pH (4.64.7) increased sodium loss by Rana clamitans and Acris gryllus tadpoles (Moore and Klerks 1998), suggesting that during hot summer months the toxic effects of acidity could increase. B. BIOTIC INTERACTIONS

At the community level, acidification can influence amphibian populations by altering interactions among individuals within a species, interactions among amphibian species, or interactions between amphibians and other taxa. All else being equal, increased mortality at acidified sites will result in fewer individuals exploiting the same limited resources. As a result of increased food availability, or due to reductions in competition, reduced population densities at acidified sites may ameliorate some of the negative effects for those individuals that are able to tolerate acidity. Because food availability may also be reduced at acidified sites (Horne and Dunson 1995a), however, possible beneficial effects of reduced density may not be great. Moreover, growth of tadpoles of Rana temporaria was depressed to a lesser extent by acidity when tadpoles were raised at higher densities (Cummins 1989). This was possibly because water at pH 4.0 tended to weed out intolerant individuals leaving those with resistance to acidic conditions able to grow and develop with little or no retardation. Likewise, the very acid-tolerant Hyla fimoralis performed better under competition from H. gratiosa at pH 4.5 than at pH 6.0 (Warner et al. 1993). The interactions between acidity and densitylcompetition have not been studied in any detail, but they appear to be complex. Because species differ in their tolerance of acidic conditions, relationships between species will change as a result of acidification. In this vein, predator-prey interactions between different amphibian species and between amphibians and other taxa can be affected by acidification (Warner et al. 1993). Acidity may also have negative effects on the prey or algae upon which amphibians feed (Horne and Dunson 1995b), resulting in indirect negative effects of acidification on amphibians. When acidification eliminates species that prey upon amphibians, this may enhance survival of some amphibian species. For instance, the predatory salamanders Ambystoma maculatum and A. jeffersonianum are more sensitive to acidity than are their prey species Pseudacris triserata and Rana sylvatica. Consequently, acidity results in less predation and higher survival of the more acid-tolerant El triseriata and R. sylvatica (Sadinski and Dunson 1992; Kiesecker 1996). Likewise, fish are common predators of amphibians and have negative impacts on amphibian populations (Bradford et al. 1998; Hecnar and M'Closkey 1998; Zimmer et al. 2002). As fish are sensitive to acidity and high aluminium concentration, many fish species are unable to persist in acidified waters (Driscoll et al. 1980; Haines 1981). The disappearance of fish may benefit those amphibian species that are sensitive to predation by fish but are tolerant of acidity. On the other hand, as fish disappear from acidified lakes, acid-tolerant insect predators, such as water boatmen and dragonfly larvae, increase in abundance. These can be ferocious predators on amphibian embryos and larvae, resulting in elevated levels of insect predation in acidified breeding sites (Henrikson 1989). Because acidity can reduce activity and swimming speed of amphibian larvae it can lessen their ability to elude capture by predatory dragonfly larvae (Jung and Jagoe 1995). Moreover, as some predators, such as caddis fly larvae, are more tolerant of acidity than many amphibian species, they can have greater negative impact upon amphibian egg and larval survival under acidic than under neutral conditions (Rowe et al. 1994).

R A S ~ E Nand GREEN: ACIDIFICATION AND AMPHIBIANS

3253

B. Are Laboratory Results Observable in the Wild?

Experiments conducted under controlled conditions in the laboratory can disentangle the effects of different factors that influence individual performance with respect to other potentially confounding factors, including temperature and density. Due to the multifarious interactions observable between low pH and different abiotic and biotic factors, however, it may be difficult to predict from simple laboratory experiments how an environmental factor, such as low pH, affects amphibians in the wild (Skelly and Kiesecker 2001; Relyea 2006). Few investigators have conducted the necessary, multilevel studies combining laboratory and field approaches in analysing amphibian responses to acidity (Sadinski and Dunson 1992). Several comparisons of laboratory and field studies, however, have shown that the negative effects of acidic environments on survival, growth and development observed in the laboratory are also observable in the field (Pough 1976; Harte and Hoffman 1989; Karns 1992; Sadinski and Dunson 1992; Surova 2002). In Rana sylvatica, Ambystom mculatum and A. jefersonianum, estimates of laboratory tolerances were good predictors of embryonic mortality in the field (Sadinski and Dunson 1992) and, consistent with laboratory results, body concentrations of sodium in R . sylvatica tadpoles were lower in acidic ponds (pH 4.14.9) than in ponds with higher pH (Freda and Dunson 1985). Furthermore, many studies that investigated species interactions were conducted in a mesocosm setting, which usually reflects more realistic conditions and the data are more readily applicable to the natural situation (Rowe and Dunson 1994; but see Skelly 2002). C. Effects at the Level of the Population Although not directly evaluated, acidification should have major consequences on individual fitness and, consequently, on population viability. Although the net effect is difficult to predict, high embryonic mortality will have a direct impact on reproductive output and the density of populations. The sublethal effects of acidity, though, will be more indirect. Slowed growth rates during the larval stages can affect post-metamorphic performance through their negative effects on metamorphic size. Smaller metamorphic size and delayed metamorphosis both have detrimental consequences for the survival and prosperity of individuals coping with competition and predation and with the changing conditions in the ponds they inhabit (Wilbur 1982; Travis et al. 1985; Rowe and Dunson 1993; Rowe and Freda 2000). Individuals that are larger at metamorphosis tend to have higher survival, as well as better reproductive success, than do smaller ones (Semlitsch et al. 1988; Berven 1990; Scott 1990, 1994) and any reductions in body size could reduce individual fitness. Also, amphibians breeding in temporary ponds or vernal pools must complete metamorphosis before the body of water dries up, whereas amphibians that inhabit cold climates but are unable to over-winter as larvae, must complete metamorphosis before the breeding pond freezes. The reductions in rates of growth or development resulting from acidification could hence be detrimental to natural populations but, as yet, not much is known about the transmission of individual-level effects to the population. IV. MECHANISMS FOR COUNTERACTING THE NEGATIVE EFFECTS OF ACIDITY A. Migration and Acclimation A key component of a population's ability to withstand environmental changes, such as acidification, is the movement of individuals to more benign areas. Many amphibians occur in patchy landscapes where the characteristics of breeding ponds as well as those of the surrounding habitat may differ greatly, thereby permitting a choice of habitat. Accordingly, the abundance of amphibians is often lower in acidic breeding ponds than in less acidic ones, perhaps because of active avoidance of acidic habitats (see below). Some amphibians are relatively poor dispersers, however, and this potentially limits their ability to actively evade acidified areas (but see Smith and Green 2005). Moreover, low numbers

3254

AMPHIBIAN BIOLOGY

at acidic sites in the field could also reflect high fidelity to breeding sites and poor recruitment at the more acidic ones. Successful avoidance of unfavourably acidified sites depends not only on the animals' dispersal capabilities but also on their ability to discriminate between habitats of different pH, and on the availability of less acidic habitats. Several studies have shown that at-least some amphibians are able to physiologically discriminate between acidic and neutral environments. Tadpoles of Bufo anzericanus, Pseudacris crucqer, R a m clamitans, R . sylvatica and R . pipiens (Freda and Taylor 1992), as well as adult Ambystoma tigrinum (Whiteman et al. 1995), R . clamitans and R. pipiens (Vatnick et al. 1999), can discriminate between neutral and acidic water when given the choice. On the other hand, adult R . temporaria showed only weak preference for higher pH when given the choice between pH 4.0 and 6.5 (Pasanen et al. 1998). Adults of A. tigrinum and R . t~mporariaoriginating from acidic ponds were less likely to avoid acidic water than were individuals from more neutral sites (Whiteman et al. 1995; Pasanen et al. 1998). This suggests that the origin of individuals may affect their ability, or willingness, to choose between acidic and neutral sites, but it is not known whether such habitat choice reflects genetic differences in habitat preference or ability to discriminate, or merely the result of experience. One important mechanism for copinq with stressful conditions, aside from avoidance, is physiological acclimation. The ability amphibians to acclimate to acidic environments has been little studied. In Rana clamitans, a 12-day prior exposure of tadpoles to pH 5 did not reduce ionoregulatory disturbance when later exposed to pH 4.0 (McDonald et al. 1984), suggesting limited ability to acclimate. In Rana temporaria, however, growth during the first three weeks was lower in individuals that were grown at pH 7.5 during the embryonic stage but transferred to water of pH 4.5 as larvae, than it was for larvae that where grown at pH 4.5 during both embryonic and larval stages (Rasanen et al. 2002). This constitutes indirect evidence that larvae acclimated somewhat to an initially low pH.

df

B. Evolutionary Adaptation 1.

Genetic Variation

In situations in which neither migration nor acclimation is possible or sufficient, the long-term persistence of populations depends on their ability to genetically adapt to changed conditions (Forbes and Calow 1997). Accordingly, stresshl environmental conditions often cause strong selection (Hoffman and Parsons 1997). Although differences in experimental methods and conditions partly confound the results, it is evident that there are large differences among species in their tolerance of acidity (Table 1). This may reflect their different evolutionary histories as species living in naturally acidic habitats appear to have the highest tolerances. For example, Hyla andersonii breeds in some of the most acidic (pH < 4.5) freshwater ecosystems in North America (Gosner and Black 1957; Pehek 1995) and correspondingly has high embryonic tolerance (critical pH = 3.6-3.8; Table 1). The same is true for embryos of Xenopus gilli (critical pH = 3.8) that inhabit acidic black waters in South Africa where pH may occasionally drop to 3.0 (Picker et al. 1993). Direct investigations of intra-specific local adaptation to acidification have been conducted for very few species: Rana arvalis (AndrCn et al. 1989; Rasanen et al. 2002, 2003a,b), R . temporaria (Tyler-Jones et al. 1989; Glos et al. 2003), R. sylvatica (Pierce and Harvey 1987; Karns 1992) and A. maculatum (Clark and LaZerte 1985). Of these, local adaptation has been found in the ranid frogs R . arualis and R. temporaria in Europe and R. sylvatica in North America. The chances of populations adapting to changing environmental conditions depends critically on the availability of heritable genetic variation. Genetic differences are suggested by the large variation in tolerance of acidic conditions and other aspects of water chemistry among amphibian species, among different populations and among clutches within populations (Pough 1976; Clark and LaZerte 1985; Clark 1986; Andrkn et al. 1989; Beattie

ANE EN and GREEN: ACIDIFICATION AND AMPHIBLANS

3255

et al. 1992; Karns 1992; Semlitsch et al. 2000; Bridges and Semlitsch 2001; Johansson et al. 2001). However, as most studies have been conducted on full-siblings (but see Pierce and Harvey 1987; Esanen et al. 2003b), variation in tolerance of acid among clutches may be due to maternal effects, which may or may not be adaptive (see below) and can be either environmentally or genetically determined (Mousseau and Fox 1998).

Very few studies to date have attempted to determine whether there is indeed a genetic basis for variation in acid tolerances within and among populations of amphibians (Pierce and Wooten 1992a; Pakkasmaa et al. 2003; Usanen et al. 2003a; Merila et al. 2004). The few studies that have done so seem to confirm that maternal effects determine both withinpopulation and between-population variation in embryonic survival under acidic conditions. This was the case in Rana sylvatica (Pierce and Sikand 1985; Pierce and Harvey 1987) and R . aroalis (&sanen et al. 2003b; Merila et al. 2004) which are among the most tolerant species and among the few in which local adaptation has been found. In Rana tempmaria there was no evidence for a genetic basis for acid tolerance in embryos but maternal effects also were weak (Pakkasmaa et al. 2003). Larval survival under acidic conditions seems to be determined by genetic effects in Rana syluatica (Pierce and Sikand 1985; Pierce and Harvey 1987) whereas no genetic basis was found for response of rates of growth or development to acidity in either R . arualis (Esanen et al. 2003a; Merila et al. 2004) or R. temporaria (Pakkasmaa et al. 2003). 2. Maternal Effects Maternal effects, whereby a mother's phenotype influences offspring phenotype, may either be environmental noise or a source of adaptive variation (Mousseau and Fox 1998). Current evidence suggests that variation in maternally derived traits may partly explain variation in the persistence of some amphibian species in acidic environments. The most obvious maternal effects influencing performance of amphibians under acidic conditions possibly arise from variation in the composition of the jelly envelope, the size of eggs and/or the amount or composition of yolk (Picker et al. 1993; Rosenberg and Pierce 1995; Rasanen et al. 2003b). Jelly capsules have a strong effect on embryonic survival in acidic water (Dunson and Connell 1982; Picker et al. 1993; Esanen et al. 2003b). The difference in embryonic survival between Xenobus gilli and X. laevis, species that have and have not adapted to acidity, respectively, presumably depends on variation in the chemical composition of their jelly capsules (Picker et al. 1993). Likewise, the difference in tolerance to acid among populations of R. arvalis embryos seems to depend on the nature of the jelly capsules as evidenced by the fact that removal of capsules greatly increased embryonic survival of a neutral-origin population, but did not influence survival of a embryos originating from a highly acidtolerant population (Rasanen et al. 2003b). Egg size seems to be of relatively little importance for embryonic acid tolerance in amphibians (AndrCn et al. 1989; Rosenberg and Pierce 1995; Rasanen et al. 2003a,b). Egg size, however, often has a strong effect on offspring size and growth in amphibians (e.g., Kaplan 1998). Thus, variation in egg size may determine differences between populations from acidified versus neutral habitats in terms of larval growth and development under acidic conditions. This seems to be the case for Rana arualis where maternal investment in large eggs in a population from an acidic habitat enables tadpoles to reach metamorphosis faster and without reduction in metamorphic size when reared in acidic water (Usanen et al. 2005). 3. Lzfi-History Variation

Due to its strong negative effects on life-history traits, such as size and age at metamorphosis, acidity might be expected to affect the evolution of life histories in amphibians (Esanen 2002; Esanen et al. 2005). Very few studies are available on this topic, however. When tested under identical conditions, Rana arvalis tadpoles from a population originating from an acidic environment had higher growth rates and metamorphosed at larger size than did

3256

AMPHIBIAN BIOLOGY

tadpoles from a population with a neutral origin (AndrCn et al. 1989; Rasanen 2002). Although larger scale studies are needed, these differences suggest adaptive variation in growth rates (Arendt 1997). Optimal maternal investment patterns may also differ between acidic and neutral sites. In Rana arvalis, females from acidified sites invest in fewer, but larger, eggs (Msanen 2002; Soderman 2006), presumably due to the positive effect of these traits on the performance of offspring (Rasanen et al. 2005). Whether these changes in reproduction are adaptive changes or are plastic responses to the environment is currently not known. Rigorous studies on local adaptation of amphibians to acidity have so far been conducted on just three species. This kind of information, however, is critical for understanding the mechanisms that allow some species to persist in acidic environments whereas other species disappear. Further studies on geographic variation in stress tolerance and in variation of key life-history traits in response to acidity, as well as the quantitative genetic basis of such traits, are clearly needed before reliable generalizations can be made.

V. SPATIAL AND TEMPORAL RELATIONSHIPS IN THE WILD A. Amphibian Distribution The species richness of both terrestrial and aquatic amphibians declines with increasing acidity (Wyman and Jancola 1992; Eason and Fauth 2001) and often breeding populations are smaller at more highly acidic sites than at less acidic ones (Strijbosch 1979; Clark 1986; Wyman and Hawksley-Lescault 1987; Blem and Blem 1991; Wyman and Jancola 1992). If known responses to low pH directly predict the effects of acidification on amphibian populations, one would expect a negative relationship between acidification and those fitness factors that are most strongly affected by acidity. For example, since acidic conditions strongly decrease embryonic viability, which is one of the major components of fitness in amphibians, amphibian distribution may be related to environmental pHs at breeding sites. Accordingly, a relatively strong positive relationship (r = 0.2'7, P < 0.001) is found between critical pH and the lowest pHs of sites occupied by a given species for the 27 species for

5.5

-

A

='5

5 -

111

5

.-

A

4.5 -

C

0

0 0

0

I CI

A

0 D O .

4 -

A

M

O

Microhylidae A

A

A

A

m Hylidae Ranidae Bufonidae Ambystomatidae

A

3

5

3,s -

A

A

0

3 3

I

I

3.5

4

I

4.5 Critical pH

Salamandridae

I

I

5

5.5

Fig. 1. The relationship between embryonic acid tolerance (critical p H = the lowest reported pH for ca. 50% mortality) and the lowest p H where the species has been reported from the wild. Each dot represents a species.

R & ~ E Nand GREEN: ACIDIFICATION AND AMPHIBIANS

3257

which data were available on both variables (Fig. 1). This provides indirect evidence that embryonic acid tolerarlce contributes to a species' ability to persist in acidic environments and that for many amphibian species pH in the range of 4.5 to 5.0 can affect distribution andlor the density of populations. Since the work of Gosner and Black (1957), many studies have found a positive relationship between habitat acidity and the distribution of individual species (Table 2). Such positive relationships between pH and a species' distribution and/or abundance, suggest that species' distributions may be altered by acidification. The evidence is not consistent, however, and several studies have found no relationship between amphibian distribution and the pH of breeding ponds (Table 2). For instance, there were no significant differences in pH or acid-neutralizing capacity between ponds that contained Pseudacris regilh larvae and those that did not in the southern Sierra Nevada of California (Soiseth 1992). Likewise, in a more comprehensive study in the same region, there were no significant differences in water chemistry between sites with and without breeding I? regilla, R a m muscosa and Bufo canorus (Bradford et al. 1994). Hence, acidification was deemed unlikely to be the cause Table 2. Relationship between pH and distribution of amphibians in studies that have investigated ponds over a range of acid and neutral sites. N = number of ponds (or quadrats in the case of Phthodon cinereus) studied. Evidence of a trend between pH and abundance (measured as number of egg masses or adults) or occurrence (present,' absent) is given as (positive), - (negative), -C (either positive or negative) or 0 (no trend), n = not available. Lowest pH refers to the lowest pH at which the species was encountered. Differential methods (egg mass counts, adult numbers) are not separated.

+

Continent North America

Europe

Taxon HYLIDAE Hyla cinerea Hyla crucqer

RANIDAE Rana arualis Rana esculenta Rana pipiem Rana syluatua Rana temporaria

North America

Rana catesbeiana Rana clamitam Rana Rana Rana Rana Rana Rana Rana

Europe

grylzo mwcosa palwtrir pipiem septentrzonalis sphenocephala syluatica

BUFONIDAE Bufo bufo

Bufo calamita

North America

Bufo americanw Bufo terrestris

pH range

lowest pH

Trend

Reference Eason and Fauth (2001) Dale et al. (1985b) Glooschenko et al. (1992) Clark (1986) Leuven et al. (1986) Strand (2002) Leuven et al. (1986) Glooschenko et al. (1992) Glooschenko et al. (1992) Clark (1986) Aston et al. 1987 Laurila (1998) Leuven et al. (1986) Bohmer and Rahmann (1990) Strand (2002) Dale et al. (198513) Clark ( 1 986) Glooschenko et al. (1992) Clark (1986) Dale et al. (1985b) Eason and Fauth (2001) Bradford et al. (1998) Dale et al. (1985b) Dale et al. (198513) Dale et al. (1985b) Warner and Dunson (1998) Gascon and Planas (1986) Rowe and Dunson (1993) Dale et al. (1985b) Leuven et al. (1986) Bohmer and Rahmann (1990) Strand (2002) Laurila (1998) Leuven et al. (1986) Bohmer and Rahmann (1990) Dale et al. (198513) Clark (1986) Warner and Dunson (1998)

AMPHIBIAN BIOLOGY

3258 Table 2

-

continued

Continent North America

Europe

North America

North America

Taxon

N

AMBYSTOMATIDAE Ambystoma 48 jeffersonianum 35 Ambystoma laterale 159 Ambystoma maculatum 17 35 159 SALAMANDRI DAE Triturus alpestk 96 18 Triturus cristatus 96 65 Triturw helveticw 96 18 Triturus vulgaris 96 18 225 158 Notophthalmus 21 uirzdescem 159 PLETHODONTIDAE PZethodon 22-167(2)1 cinereus

pH range 4.4-5.8 4.2-6.3 3.6-9 4.5-7.5 4.2-6.2 3.6-9 3.5-9 3.7-8.1 3.5-9 4.0-9 3.5-9 3.7-8.1 3.5-9 3.7-8.1 4.0-9.5 3.5-8.8 3.8-5.8 3.6-9 2.7-5.9

lowest pH 4.5 4.2 6.6 4.5 4.2 3.9 4-5 4.2 4-5 5.0-5.4 <4 4.2 4-5 6.3 4.5-4.9 4.9 4.2 5 3.4

Trend

+ + 0 n

+

0 f

+ ? + f + + n + n + 0

+

Reference Home and Dunson (1994a) Rowe and Dunson (1993) Dale et al. (1985b) Pough (1976) Rowe and Dunson (1993*) Dale et al. (1985b) Leuven et al. (1986) Bohmer and Rahmann (1990) Leuven et al. (1986) Strand (2002) Leuven et al. (1986) Bohmer and Rahmann (1990) Leuven et al. (1986) Bohmer and Rahmann (1990) Strand (2002) Laurila (1998) Eason and Fauth (2001) Dale et al. (1985b) Wyman and Hawskley-Lescault (1987)

t Refers to buffering status, pH influence not separated. * Studies were conducted 1991 and 1992, but only 1991 presented as 35 ponds, as opposed

:: Terrestrial quadrats within two sites.

15 ponds, were studied.

behind the declines of R. muscosa or B. canorus populations in that region. Sometimes there are even more egg masses in acidic ponds than there are in less acidic ones, as was the case for Rana temporaria in Great Britain, perhaps reflecting the influence of areas surrounding the breeding ponds (Aston et al. 1987). B. Local Long-Term Patterns

There is some evidence that amphibian habitats in regions affected by acidifying precipitation have become more acidic (Freda et al. 1991) but the evidence for a related decline in amphibian populations is very limited and mainly indirect. Apparent local declines resulting from response to acidification have been reported for Rana arvalis in Germany (Clausnitzer 1987), for R. temporaria in Sweden (Hagstrom 1980), for Bufo calamita in Great Britain (Beebee et al. 1990) and for Ambystoma tigrinum in the United States (Harte and Hoffman 1989). Of these, R . arvalis and R. temporaria are common species in Northern Europe (Leuven et al. 1986; Aston et al. 1987). R . arvalis is the more acid-tolerant of these two species and, as expected from their differential sensitivity during the embryonic stage, R. temporaria is usually excluded from extremely acidic sites whereas R. arvalis can occur at a pH of about 4.0 (AndrCn et al. 1989; Rasanen et al. 2003a; Soderman 2006). There is no clear evidence, however, for a large-scale decline in either of these species. On the contrary, both of them seem to be able to adapt to the locally prevailing acidity (AndrCn et al. 1989; Tyler-Jones et al. 1989; Glos et al. 2003; Rasanen et al. 2003a). To make rigorous conclusions about the effects of acidification on amphibian populations, longitudinal data on populations inhabiting areas that suffer from acidification are needed, but few such data exist to date. Among the few examples, Harte and Hoffman (1989) interpreted a decline in Ambystoma tzpinum populations during 1982-1988 in central Colorado as likely being the result of acidification, but in a follow-up study of the same population during 1988-1992, Wissinger and Whiteman (1992) found no evidence for that interpretation. Likewise, Vertucci and Corn (1996) found no evidence for anthropogenic acidification of amphibian habitats in the Rocky Mountains despite the decline of some amphibian species in the area.

RASANEN and GREEN: ACIDIFICATION AND AMPHIBIANS

3259

C. Global Trends

The European species Ram temporaria and Bufo bufo are among the few species for which several time-series are available from regions affected by differing levels of acidification over extended periods of time (Green 2003). Hence, these species, if any, can be expected to show the effects of historical anthropogenic acidification. Yet there are no indications that these species are in decline as a result of acidification. There are no detectable differences in trends from year to year or from place to place in either species (Green 2003), irrespective of overall levels of acidification. Part of this may be explained by the fact that R. tempora.l-ia, at least, seems to be able to adapt to acidification to some degree. As both of these species have high variance in population size, however, any effect that might occur would be difficult to detect. If acidification were a major cause of amphibian decline on a global scale, one would expect negative trends in amphibian populations in areas that have suffered heavily from anthropogenic acidification. These include northeastern North America, northern Europe (especially Norway, Sweden, Finland and Scotland) and some parts of Central Europe, such as the Czech Republic and southern Germany. T h e areas that seem most affected by amphibian decline, however, including western North America, eastern Australia and Central America, do not concur with this pattern of acidification. Lack of clear relationships between acidity (pH) of the breeding site and a species' distribution may arise from failure to account for other factors that likely confound the effects of p H on amphibians in the wild. These include very broad tolerances to acid of the species under study, movement of individuals between sites, as well as other environmental features that may hide the impact of acidity. In a landscape that consists of patchily distributed breeding ponds, some more highly acidic than others, large numbers of breeding adults may arrive from elsewhere to congregate at acidic sites even if recruitment from those sites is low (Aston et al. 198'7). Subpopulations occurring at the more benign sites may contribute to subpopulations at the acidified sites and hence accessibility of less acidic sites can be critical for species that are sensitive to acidity but unable to adapt to it. Acidified areas, however, may often be low in buffering capacity over a large scale, resulting in few or no benign habitats available. In such areas, breeding numbers of amphibians could be severely reduced. Also, the landscape surrounding the breeding ponds may be of great importance in determining the distribution and abundance of animals (Skelly 2001; Houlahan and Findley 2003) and confound the effect of pH at the breeding ponds. VI. AMPHIBIANS AS INDICATORS OF ENVIRONMENTAL ACIDIFICATION

At the time the problem of declining amphibian populations was raised, amphibians were touted as sensitive indicators of environmental stress (Barinaga 1990; Blaustein and Wake 1990; Wynlan 1990). Their utility as environmental indicators has been disputed in general terms (Pechmann and Wilbur 1994), as well as defended (Blaustein 1994), but it seems that the suitability of amphibians as bio-indicators of acidification is limited. Apart from a few exceptionally sensitive species, amphibians are relatively acid-tolerant and most effects of acidification become evident only when acidification has reached extreme levels. Furthermore, many of the effects on amphibians are sublethal, with possibly far-reaching consequences at the population level, but difficult to detect in the wild. First, suitable measures for bioassays are limited. The main attributes of amphibians that have been suggested as parameters for bioassays in acidified aquatic and terrestrial habitats are the embryonic curling defect, net loss of body sodium, and developmental instability (Dunson et al. 1992; McCoy and Harris 2003). Estimating the proportion of embryos with a curling defect should be indicative and an easy measure of acidification. Unless embryonic survival as a result of the curling defect is extremely low, however, there is no easy way to predict how this embryonic mortality will influence population viability. Measurements of net body sodium may indicate to what degree aquatic or terrestrial stages suffer from acidity but this method will inevitably require specific equipment and the

3260

AMPHIBIAN BIOLOGY

development of species-specific baseline measures for comparison. Fluctuating asymmetry may indicate stress, although its usability is disputed in general (Dongen 2006). R. arvalis adults from acidified areas did show higher levels of fluctuating asymmetry than did adults from neutral areas, suggesting a negative effect of acidity on developmental stability (Soderman et al., 2007'). By contrast, in Ambystoma maculatum correlates of traditional fitness, such as size at metamorphosis, were correlated with pH stress whereas fluctuating asymmetry was not, suggesting that it may not be suitable as an indicator of acidification (McCoy and Harris 2003). Hence, although amphibians are susceptible to many environmental stressors, including excessive acidification, they also possess many attributes by which they can counteract negative effects. They may acclimate physiologically, adapt genetically, or avoid acidic habitats by moving to less acidic ones. Very few species are able to withstand extremely acidic conditions but mild acidification may have no easily measured effects on long-term presence of amphibians. Thus, the value of amphibian populations as bio-indicators of acidification appears to be rather low. Second, shifting abundances of individuals are not necessarily indicative of any particular form of environmental degradation. For instance, populations of pond-breeding amphibians naturally fluctuate to a great extent owing to variance in juvenile recruitment and adult survivorship (Green 2003). Amphibian breeding congregations may also move from year to year and thus estimates of decline and distributional change can be extremely sensitive to the duration of resurvey effort and the type of historical data used (Skelly et al. 2003). For example, numbers off egg masses of spotted salamanders, Ambystoma maculatum, fluctuated five-fold both among years and among sites in New Brunswick in acidified ponds with a pH of about 5.5 (Clay 1997'). In Massachusetts, the same species did not lay eggs in ponds below pH 4.5 whereas there was no relationship between numbers of egg masses and pH in ponds less acidic than that (Portnoy 1990). Brodman (2002) monitored breeding populations of A. maculatum and A. jeffersonianum in two wetlands with pH 5.8-6.5 in northern Ohio during 1990-200 1. Survival of embryos of A. maculatum correlated positively with pH but there was no significant trend over time in the rlurnber of embryos surviving to hatch each year for either species. Therefore, despite assumptions that amphibians are particularly susceptible to environmental stressors, there is little evidence that they are either good predictors or indicators of environmental acidification. Although at least some species are able to discriminate between acidic and neutral sites, adults of many species will breed at pH levels that are harmful to their offspring. Furthermore, because of the large fluctuations in amphibian population size at any given locality (Green 2003), very long-term datasets are required to detect negative effects of acidification at the population level. VII. CONCLUSIONS: ACIDIFICATION AS A CAUSE OF AMPHIBIAN DECLINE

There is ample empirical evidence for many direct and indirect negative effects of acidity on amphibians. There is little evidence, however, that acidifying precipitation is a primary cause of amphibian declines. If acidification were a major cause of amphibian decline on a global scale, one would expect negative trends in amphibian populations in areas that have suffered heavily from anthropogenic acidification but current evidence does not suggest this to be the case. Well over a decade ago, there were no data at hand to demonstrate whether or not anthropogenic acidification causes amphibian populations to decline (Dunson et al. 1992). This remains true even today although more long-term studies have now been conducted on amphibian populations (Houlahan et al. 2000; Green 2003). Studies that can infer causes behind detected declines are few and those attempting to detect the effects of acidification are practically non-existent (Alford and Richards 1999). It may be that acidification does not have a major influence on amphibian populations in nature or that local weather or hydroperiod (Wissinger and Whiteman 1992; Muths et al. 2003) or other hurnan-induced

RASANEN and GREEN: ACIDIFICATION AND AMPHIBIANS

3261

environmental changes, such as habitat fragmentation, have a greater impact on persistence of populations than does acidity. Or it may simply be that there is a lack of adequate study and information. It is clear that many areas require more work before one can understand the full suite of effects that acidification may have on amphibian populations. The effects of acidification are complex when other factors, such as substrate buffering capacity, predatorlprey relationships, competition with other species, metal ion concentrations, temperature, UV-B radiation and the physical properties of habitats are taken into account. With this in mind, the suitability of amphibians as bio-indicators of acidification may be limited by the relatively high tolerance for acidity of many species and by their abilities to acclimate and adapt to acidic conditions. Environmental acidification exposes amphibians to stressful conditions and may have far reaching consequences on amphibian populations. Whereas these effects can contribute to the observable global decline, they are likely to exert their influence at a local scale rather than on a global level. VIII. ACKNOWLEDGEMENTS

This work was supported by a Post Doctoral Grant from the Swedish Research Council to KR and a Discovery Grant from NSERC Canada to DMG. IX. REFERENCES Alewell, C., Maderscheid, H., Meesenburg, H. and Bittersohl, J., 2000. Is acidification still an ecological threat? Nature 407: 856-857. Alford, R. A. and Richards, S. J., 1999. Global amphibian declines: a problem in applied ecology. Ann. Rev. Ecol. System. 30: 109-132. AMAP, 1998. Acidifying Pollutants, arctic haze, and acidification in the Arctic. Arctic Monitoring and Assessment Programme (AMAP), Oslo. AndrCn, C . , Henrikson, L., Olsson, M. and Nilsson, G., 1988. Effects of pH and aluminium on embryonic and early larval stages of Swedish brown frogs Rana arualis, R. temporaria and R. dalmatim. Holarctic Ecol. 11: 127-135. AndrCn, C., M%rdCn, M. and Nilsson, G., 1989. Tolerance to low pH in a population of moor frogs, Rana arualis, from an acid and a neutral environment: a possible case of rapid evolutionary response to acidification. Oikos 56: 215-223. B0rg7 H'* Edbergl F' and Hu1tberg2 H', Andersson, av sjbar. Observerade och 2002. Aterf~jrsurnin~ forvantade biologiska och kemiska effekter. NaturvArdsverket, Stockholm. Arendt, J . D., 1997. Adaptive intrinsic growth rates: an integration across taxa. Quart. Rev. Biol. 72: 149-1 - - - 77.

Beattie, R. C., Aston, R. J. and Milner, A. G. P, 1991. A field study of fertilization and embryonic development in the common frog, Ram temporaria, with particular reference to acidity and temperature. J. Appl. Ecol. 28: 346-357. Beattie, R. C., Aston, R. J. and Milner, A. G. P., 1993. Embryonic and larval survival of the common frog (Rana temporaria L) in acidic and limed ponds. Herpetol. J. 3: 43-48. Beattie, R. C. and Tyler-Jones, R., 1992. The effects of low pH and aluminium on breeding success in the frog Rana temporaria.J. Herpetol. 26: 353-360. Beattie, R. C., Tyler-Jones, R. and Baxter, M. J., 1992. The effects of pH, aluminium concentration and temperature on the embryonic development of the European common frog, Rana temporaria. J. Zool. (London) 228: 557-570. Beebee, T J. C. and Griffin, J. R., 1977. A preliminary investigation into natterjack toad (Bufo calamita) breeding site characteristics in Britain. J. Zool. 181: 341-350. Beebee, T. J. C., Flower, R. J., Stevenson, A. C., Patrick, S. T, Appleby, P G., Fletcher, C., Marsh, C., Natkanski, J., Rippey, B. and Battarbee, R. W., 1990. Decline of natteriack toad Bufo calamita in and Britain: paleoecological, documentary experimental evidence for breeding site acidification. Biol. Cons. 53: 1-20.

Barinaga, M., 1990. Where have all the froggies gone? Science 247: 1033-1034.

Brakke, D. F., Baker, J. P., Bohmer, J., Hartmann, A., Havas, M., Jenkins, A., Kelly, A. C., Ormerod, S. J., Paces, T, Putz, R., Rosseland, B. O., Schindler, D. W. and Segner, H., 1994. Physiological and ecolopical effects of acidification on aquatic biota. Pp. 575-312 in "Acidification of ^~reshwater Ecosystems: Implications for the Future", ed by C. E. W. Steinberg and R. F. Wright. Wiley, New York.

Battarbee, R. W. and Charles, D. F., 1994. Lake acidification and the role of paleolimnology. Pp. 51-65 in "Acidfication of Freshwater Ecosystems: Implications for the Future", ed by C. E. W. Steinberg and R. F. Wright. Wiley, New York.

Bellemakers, M. J. S. and Van Dam, H., 1992. Improvement of breeding success of the moor frog (Rana arualis) by liming of acid moorland pools and the consequences of liming for water chemistry and diatoms. Environ. Pollu. 78: 165-17 1.

Aston, R. J., Beattie, R. C. and Milner, A. G. P., 1987. Characteristics of spawning sites of the common frog (Rana temporaria) with particular reference to acidity. . "I. Zool. (London) 213: 233-242.

3262

AMPHIBIAN BIOLOGY

Berven, K. A,, 1990. Factors affecting population fluctuations in larval and adult stages of the wood frog (Rana syluatica). Ecology 71: 1599-1608. Blaustein, A. R., 1994. Chicken Little or Nero's fiddle? A perspective on declining amphibian populations. Herpetologica 50: 85-97. Blaustein, A. R. and Wake, D. B., 1990. Declining amphibian populations: a global phenomenon? Trendr i n Ecol. Euol. 5: 203-204. Blem, C. R. and Blem, L. B., 1991. Cation concentration and acidity in breeding ponds of the spotted salamander, Ambystoma maculatum (Shaw), in Virginia. Brimleyana 17: 67-76. Bradford, D. F., Swanson, C. and Gordon, M. S., 1992. Effects of low p H and aluminum on 2 declining species of amphibians in the Sierra Nevada, California. J. Herpetol. 26: 369-377. Bradford, D. F., Gordon, M. S., Johnson, D. F., Andrews, R. D. and Jennings, W. B., 1994. Acidic deposition as an unlikely cause for amphibian population declines in the Sierra Nevada, California. Biol.Cons. 69: 155-161. Bradford, D. F., Cooper, S. D., Jenkins, T M., Kratz, K., Sarnelle, 0. and Brown, A. D., 1998. Influences of natural acidity and introduced fish on faunal assemblages in California alpine lakes. Can. J. Fish. Aquat. Sci. 55: 2478-2491.

Clark, K. L. and Hall, R. J., 1985. Effects of elevated hydrogen ion and aluminium concentrations on the survival of amphibian embryos and larvae. Can. J. Zool. 63: 116-123. Clark, K. L. and LaZerte, B. D., 1985. A laboratory study on effects of aluminium and pH on amphibian eggs and tadpoles. Can. J. Fish. Aquat. Sci. 42: 1544-1551. Clark, K. L. and LaZerte, B. D., 1987. Intraspecific variation in hydrogen ion and aluminium toxicity in Bufo americanus and Ambystoma maculatum. Can. J. Fish. Aquat. Sci. 44: 1622-1628. Clausnitzer, H.-J., 1987. Gefahrdung des Moorfrosches (Rana arualis Nilsson) durch Versauemng der Laichgewasser. Beiheft Schnftenreihe Naturschutz und Landrchaftspj&ge i n Niedersachen 19: 131-1 37. Clay, D., 1997. The effects of temperature and acidity on spawning of the spotted salamander, Ambystoma maculatum, in Fundy National Park. Pp. 226-232 in "Amphibians in Decline. Canadian Studies of a Global Problem", ed by D. M. Green. Herpetological Conservation, No. 1, Society for the Study of Amphibians and Reptiles, Saint Louis. Collier, K. J., Ball, 0.J., Graesser, A. K., Main, M. R. and Winterbourn, M. J., 1990. Do organic and anthropogenic acidity have similar effects on aquatic fauna? Oikos 59: 3-38.

Bridges, C. M. and Semlitsch, R. D., 2001. Genetic variation in insecticide tolerance in a population of southern leopard frogs (Rana sphenocephala): Implications for amphibian conservation. Copeia 2001: 7-13.

Cook, R. P., 1978. Effects of acid precipitation on embryonic mortality of spotted salamanders (Ambystoma maculatum) and Jefferson salamanders (Ambystoma jefersonianum) in the Connecticut Valley of Massachussets. Thesis, University of Massachussets, Amherst.

Brodin, Y.-W., 1993. "Acidification. An everlasting problem, or is there hope? The environment in Sweden." Swedish Environmental Protection Agency, Varnamo.

Corn, P. S., 2003. Amphibian breeding and climate change: the importance of snow in the mountains. Cons. Biol. 17: 621-624.

Brodkin, M., Vatnick, I., Simon, M., Hopey, H., ButlerHolston, K. and Leonard, M., 2003. Effects of acid smss on adult R a m p*i. J. Exp. Zool. 298A: 1622. Brodman, R., 2002. Twelve-year study monitoring two species of pond-breeding salamanders in northeast Ohio. Ohio J. Sci. 102: 123-127. Bohmer, J., 1988. Labor- und Feldstudien zum EinfluR niedriger pH-Werte auf die friihe Entwicklungs von Molchen im Schwarzwald. Diploma thesis, University of Hohenheim, Federal Republic of Germany.

Corn, I? S., Stolzenburg, W. and Bury, R. B., 1989. "Acid Precipitation Studies in Colorado and Wyoming: Interim Report of Surveys of Montane Amphibians and Water Chemistry". Biological Report 80 (40.26). U.S. Fish and Wildlife Service, Washington DC. Corn, P S. and Vertucci, E A, 1992. Descriptive risk assessment of the effects of acidic deposition on Rocky Mountain amphibians. J. Herpetol. 26: 361-369. Cowman, D. F. and Mazanti, L. E., 2000. Ecotoxicology of "new generation" pesticides to amphibians. Pp. 233-268 in "Ecotoxicology of Amphibians and Reptiles", ed by D. W. Sparling, G. Linder and C. A. Bishop. Society of Environmental Toxicology and Chemistry, Pensacola.

Bohmer, J., Bauser-Eckstein, A. and Rahmann, H., 1988. Labor- und Feldstudien zum EinfluR niedriger pHWerte auf Molche. Pp. 225-228 in "Gefahrdung und Schutz von Gewassern", ed by A. Kohler and H. Rahmann. Hohenheimer Arbeiten, Ulmer Verlag.

Cummins, C. P, 1986. Effects of aluminum and low pH on growth and development in Rana temporaria tadpoles. Oecologta 69: 248-252.

Bohmer, J. and Rahmann, H., 1990. Influence of surface water acidification on amphibians. Fortschritte der Zoologie 38: 287-309.

Cummins, C. I?, 1989. Interaction between the effects of pH and density on growth and development in R a m temporaria L. tadpoles. Funct. Ecol. 3: 45-52.

Carey, C., 1993. Hypothesis concerning the causes of the disappearance of Boreal toads from the mountains of Colorado. Cons. Biol. 7: 355-362.

Dale, J., Freedman, B. and Kerekes, J., 1985a. Experimental studies of the effects of acidity and associated water chemistry on amphibians. Proc. Nova Scotia Znst. Sci. 35: 35-54.

Chen, C. Y., Hathaway, K. M. and Folt, C. L., 2004. Multiple stress effects of Vision (R) herbicide, pH, and food on zooplankton and larval amphibian species from forest wetlands. Enuiron. Toxicol. Chem. 23: 823-831. Clark, K. L., 1986. Distributions of anuran populations in central Ontario relative to habitat acidity. Wate~; Air and Soil Pollut. 30: 727-734.

Dale, J., Freedman, B. and Kerekes, J., 1985b. Acidity and associated water chemistry of amphibian habitats in Nova Scotia. Can. J. Zool. 63: 97-105. Dangles, O., Malmqvist, B. and Hjalmar Laudon, H., 2004. Naturally acid freshwater ecosystems are diverse and functional: evidence from boreal streams. Oikos 104: 149-155.

&ANEN

and GREEN: ACIDIFICATION AND AMPHIBIANS

Dongen, S. V., 2006. Fluctuating asymmetry and developmental instability in evolutionary biology: past, present and hture. J. Evol. Biol. 19: 1727-1743. Driscoll, C. T., Baker, J. P. Jr, Bisogni, J. J. Jr and Schofield, C. L., 1980. Effect of aluminium speciation on fish in dilute acidified waters. Nature 284: 161-164. Dunson, W. A. and Connell, J., 1982. Specific inhibition of hatching in amphibian embryos by low pH. J. Herpetol. 16: 314-3 16. Dunson, W. A,, Wyman, R. L. and Corbett, E. S., 1992. A symposium on amphibian declines and habitat acidification. J. Herpetol. 26: 349-352. Eason, G. W. and Fauth, J. E., 2001. Ecological correlates of anuran species richness in temporary pools: A field study in South Carolina, USA. Israel J. Zool. 47: 347-365. Fioramonti, E., Semlitsch, R. D., Reyer, H.-U. and Fent, K., 1997. Effects of triphenyltin and pH o n the growth and development of Rana lessonae and Rana esculenta tadpoles. Environ. Toxicol. Chem. 16: 1940-1947. Forbes, V. E. and Calow, P., 1997. Responses of aquatic organisms to pollutant stress: theoretical and practical implications. Pp. 25-41 in "Environmental Stress, Adaptation and Evolution", ed by R. Biljsma and V. Loeschcke. Birkhauser Verlag, Basel. Freda, J., 1986. The influence of acidic pond water on amphibians: A review. Wate?; Air and Soil Pollut. 30: 439-450. Freda, J., 1991. The effects of aluminum and other metals on amphibians. Environ. Pollut. 71: 305-328. Freda, J. and Dunson, W. A., 1984. Sodium balance of amphibian larvae exposed to low environmental pH. Physiolog. 2001. 57: 435-443. Freda, J. and Dunson, W. A., 1985a. Field and laboratory studies of ion balance and growth rates of ranid tadpoles exposed to low pH. Copeia 1985: 415423. Freda, J. and Dunson, W. A., 1985b. The influence of external cation concentration on the hatching of amphibian embryos in water of low pH. Can. J. Zool. 63: 2649-2656.

3263

Frisbie, M. P. and Wyman, R. L., 1995. A field simulation of the effect of acidic rain on ion balance in a woodland salamander. Arch. Environ. Contnm. Toxicol. 28: 327-333. Gascon, C. and Planas, D., 1986. Spring pond water chemistry and the reproduction of the wood frog, Rana sylvatica. Can. J. 2001. 64: 543-550. Gebhardt, H., Kreimes, K. and Linnenbach, M., 1987. Untersuchungen zur Beeintrachtigung der Ei- und Larvenstadien von Amphibien in sauren Gewassern. Natur und Landtchaj? 62: 20-23. Glooschenko, V, Weller, \V F., Smith, El G. R., Alvo, R. and Archbold, J. H. G., 1992. Amphibian dismbution with respect to pond water chemistry near Sudbury, Ontario. Can. J. Fish. Aqwzt. Sci. 49 (Sl): 11 4 1 21. Glos, J., Grafe, T U., Rodel, M. 0. and Linsenmair, K. E., 2003. Geographic variation in pH tolerance of two populations of the common frog, Rana temporaria. Copeia 2003: 650-656. Gosner, K. L. and Black, I. H., 1957. The effects of acidity on the development and hatching of New Jersey frogs. Ecology 38: 256-262. Grant, K. P. and Licht, L. E., 1993. Acid tolerance of anuran embryos and larvae from Central Ontario. J. Herpetol. 27: 1-6. Green, D. M., 2003. The ecology of extinction: population fluctuation and decline in amphibians. Biol. Cons. 111: 331-343. Griffiths, R. A,, 1993. The effect of pH on feeding behavior in newt larvae (Triurus, Amphibia).J. Zool. 231: 285-290. Griffiths, R. A., de Wijer, P. and Brady, L., 1993. The effect of pH on embryonic and larval development in smooth and palmate newts, Triturw vulgaris and ?: helveticw. J. 2001. (London) 230: 401-409. Hagstrom, T., 1980. Reproductive strategy and success of amphibians in waters acidified by atmospheric pollution. Pp. 55-57 in Proceedings of the European Herpetological Symposium, 1980. Cotswold Wildlife Park, Oxford. Haidacher, S. and Fachbach, G., 1991. Experimental acid tolerance analyses of the embryos and larvae of European amphibians. Salamandra 27: 108-1 18.

Freda, J. and Dunson, W. A,, 1986a. Effects of low pH and other chemical variables on the local distribution of amphibians. Copeia 1986: 454-466.

Haines, T. A,, 1981. Acidic precipitation and its consequences for aquatic ecosystems: a review. Trans. Ame7: Fish. Sac. 110: 669-707.

Freda, J. and Dunson, W. A., 1986b. The effect of prior exposure on sodium uptake in tadpoles exposed to low pH water. J. Compal: Physiol. 156: 649-654.

Harte, J. and Hoffman, E., 1989. Possible effects of acidic deposition on a Rocky Mountain population of the tiger salamander Ambystoma tigrinum. Cons. Biol. 3: 149-158.

Freda, J. and McDonald, D. G., 1990. Effects of aluminium on the Leopard frog, Rana pipiens: life stage comparisons and aluminium uptake. Can. J. Fish. Aquat. Sci. 47: 210-216. Freda, J . and Taylor, D. H., 1992. Behavioral response of amphibian larvae to acidic water. J. Herpetol. 26: 429-433. Freda, J., Sadinski, W. J. and Dunson, W. A., 1991. Long term monitoring of amphibian populations with respect to the effects of acidic deposition. Wate?;Air and Soil Pollut. 55: 445462. Frisbie, M. P. and Wyman, R. L., 1991. The effects of soil pH on sodium balance in the red-backed salamander, Plethodon cinereus, and three other terrestrial salamanders. Physwlog. Zool. 64: 1050-1068.

Hatch, A. C. and Blaustein, A. R., 2000. Combined effects of W - B , nitrate, and low pH reduce the survival and activity level of larval cascades frogs (Rana cascadae). Arch. Environ. Contam. Toxicol. 39: 494-499. Hecnar, S. J., 1995. Acute and chronic toxicity of ammonium-nitrate fertilizer to amphibians from Southern Ontario. Environ. Toxicol. Chem. 14: 2131-2137. Hecnar, S. J . and M'Closkey, R. T , 1998. Species richness patterns of amphibians in southwestern Ontario ponds. J. Biogeo. 25: 763-772. Henrikson, B.-I., 1989. Predation on amphibian eggs and tadpoles by common predators in acidified lakes. Holarctic Ecology 13: 201-206.

3264

AMPHIBIAN BIOLOGY

Hoffman, A. A. and Parsons, P. A., 1997. "Extreme Environmental Change and Evolution". Cambridge University Press, Cambridge.

Kutka, F. J., 1994. Low pH effects on swimming activity of Ambystoma salamander larvae. Environ. Toxicol. Chem. 13: 1821-1824.

Horne, M. T. and Dunson, W A,, 1994a. Exclusion of the Jefferson salamander, Ambystoma jeffersonianum, fmm some potential breeding ponds in Pennsylvania - effects of pH, temperature, and metals on embryonic development. Arch. Environ. Contam. Toxicol. 27: 323-330.

Laurila, A., 1998. Breeding habitat selection and larval performance of two anurans in freshwater rockpools. Ecography 21: 484494.

Home, M. T and Dunson, W A,, 1994b. Behavioural and physiological responses of the terrestrial life stages of the Jefferson salamander, Ambystoma jefferssonianum, to low soil pH. Arch. Environ. Contam. Toxicol. 27: 232-238. Horne, M. T. and Dunson, W. A,, 1995a. Effects of low pH, metals, and water hardness on larval amphibians. Arch. Environ. Contam. Toxicol. 29: 500-505. Horne, M. T and Dunson, W. A,, 199513. The interactive effects of low pH, toxic metals, and DOC on a simulated temporary pond community. Environ. Pollut. 89: 155-161. Houlahan, J. E., Findlay, C. S., Schmidt, B. R., Meyer, A H. and Kuzmin, S. L., 2000. Quantitative evidence for global amphibian population declines. Nature 404: 752-755. Houlahan, J. E. and Findlay, C. S., 2003. The effects of adjacent land use on wetland amphibian species richness and community composition. Can. J. Fish. Aquat. Sci. 60: 1078-1094. Huckabee, J. W., Mattice, J. S., Pitelka, L. F., Porcella, D. B. and Goldstein, R. A,, 1989. An assessment of the ecological effects of acidic deposition. Arch. Environ. Contam. Toxicol. 18: 3-27.

Leuven, R. S. E. W., den Hartog, C . , Christiaans, M. M. C. and Hejligers, W. H. C., 1986. Effects of surface water acidification on the distribution pattern and the reproductive success of amphibians. Experientia 42: 495-503. Linder, G. and Grillitsch. B., 2000. Ecotoxicology of metals. Pp. 325-459 in "Ecotoxicology of Amphibians and Reptiles", ed by D. W. Sparling, G. Linder and C. A. Bishop. Society of Environmental Toxicology and Chemistry (SETAC) Press, Pensacola. Lindstrom, J., 1999. Early development and fitness in birds and mammals. Trends Ecol. Evol. 14: 343-348. Long, L. E., Saylor, L. S. and Soul.6, M. E., 1995. A pH/UV-B synergism in amphibians. Cons. Biol. 9: 1301-1303. Mannion, A. M., 1999. Acid precipitation: a review. University of Reading, Geographical Paper 127: 1-27. Maltby, L., 1999. Studying stress: the importance of organism-level responses. Ecol. Applic. 9: 43 1-440. McCoy, K. A. and Harris, R. N., 2003. Integrating developmental stability analysis and current amphibian monitoring techniques: An experimental evaluation with the salamander Ambystoma maculatum. Her$etologica 59: 22-36.

Ireland, E, 1991. Separate effects of acid-derived anions and cations on growth of larval salamanders of Ambystoma maculatum. Copeia 1991: 132-137.

McDonald, D. G., Ozog, J. L. and Simons, B. E, 1984. The influence of low pH environments on ion regulation in the larval stages of the anuran amphibians, Rana clamitans. Can. J. 2001. 62: 2171-2177.

Johansson, M., Risanen, K. and Merila, J., 2001. Comparison of nitrate tolerance between different populations of the common frog, Rana temporaria. Aquat. Toxicol. 54: 1-14.

Merila, J., Soderman, F., O'Hara, R., R;isanen, K. and Laurila, A,, 2004. Local adaptation and genetics of acid-stress tolerance in the moor frog, Rana arualis. Cons. Genet. 5: 513-527.

Jung, R. E. and Jagoe, C. H., 1995. Effects of low pH and aluminum on body size, swimming performance, and susceptibility to predation of green frog (Hyla cinerea) tadpoles. Can. J. Zool. 73: 2171-2183.

Moore, M. K. and Klerks, E L., 1998. Interactive effect of high temperature and low pH on sodium flux in tadpoles. J. Herpetol. 32: 588-592.

Kaplan, R. H., 1998. Maternal effects, developmental plasticity, and life history evolution: an amphibian model. Pp. 244-260 in "Maternal Effects as Adaptations", ed by T A. Mousseau and C. W. Fox. Oxford University Press, New York. Karns, D. R., 1983. Toxic bog water in northern Minnesota peatlands: ecological and evolutionary consequences for breeding amphibians. PhD dissertation, University of Minnesota. Karns, D. R., 1992. Effects of acidic bog habitats on amphibian reproduction in a northern Minnesota peatland. J. Herpetol. 26: 401412. Kiesecker, J., 1996. pH-mediated predator-prey interactions between Ambystoma tigrinum and Pseudacris triseriata. Ecol. Applic. 6: 1325-1 33 1. Kitamura, Y. and Katagiri, C., 1998. Characterization of the hatching enzyme from embryos of a n anuran amphibian, Rana pirica. Biochimica et Biophysica Acta Protein structure and Molecular Enzymology 1387: 153-164.

Mousseau, T A. and Fox, C. W., 1998. (Editors). "Maternal Effects as Adaptations". Oxford University Press, Oxford. Muths, E., Campbell, D. H. and Corn, P. S., 2003. Hatching success in salamanders and chorus frogs at two sites in Colorado, USA: effects of acidic deposition and climate. Amphibia-Reptilia 24: 27-36. Padhye, A. D. and Ghate, H. V., 1988. Effect of altered pH on embryos and tadpoles of the frog Microhyla ornata. Herpetol. J. 1: 276-279. Pahkala, M., Laurila, A. and Merila, J., 2001a. Carryover effects of ultraviolet-B radiation on larval fitness in R a m temporama. Proc. Roy. Soc. Lond., B. 268: 1699-1706. Pahkala, M., Laurila, A. Bjorn, L. 0 . and Merila. J., 2001b. Effects of ultraviolet-B radiation and pH on early development of the moor frog Rana arva1i.s. J. Appl. Ecol. 38: 628-636. Pahkala, M., RBsanen, K., Laurila, A., Johansson, O., Bjorn, L. 0. and Merila, J., 2002. Lethal and sublethal effects of UV-B/pH synergism on common frog embryos. Cons. Biol. 16: 1063-1073.

&ANEN

and GREEN: ACIDIFICATION AND AMPHIBIANS

Pakkasmaa, S., Merila, J . and O'Hara, R. B., 2003. Genetic and maternal effect influences on viability of common frog tadpoles under different environmental conditions. Heredity 91: 1 17-1 24. Pasanen, S., Laitinen, M. and Alhonen, T, 1998. Effects of pH on the wintering of the common frog (Rana temporaria L.). Ann. 2001. Fennici 35: 29-36. Pechenik, J. A., Wendt, D. E. and Jarrett, J. N., 1998. Metamorphosis is not a new beginning. Bioscience 48: 901-910. Pechmann, J. H. K. and Wilbur, H. M., 1994. Putting declining amphibian populations in perspective: natural fluctuations and human impacts. Herpetologica 50: 65-94. Pehek, E. L., 1995. Competition, pH, and the ecology of larval Hyle andersonii. Ecology 76: 1786-1793. Picker, M. D., McKenzie, C. J. and Fielding, I?, 1993. Embryonic tolerance of Xenopus (Anura) to acidic blackwater. Copeia 1993: 1072-1081. Pierce, B. A,, 1985. Acid tolerance in amphibians. BioScience 35: 239-243. Pierce, B. A., 1993. Effects of acid precipitation on amphibians. Ecotoxicology 2: 65-77. Pierce, B. A. and Harvey, J. M., 1987. Geographic variation in acid tolerance of Connecticut Wood Frogs. Copeia 1987: 94-103. Pierce, B. A. and Sikand, N., 1985. Variation in acid tolerance of Connecticut wood frogs: genetic and maternal effects. Can. J. Zool. 63: 1647-165 1. Pierce, B. A. and Montgomery, J., 1989. Effects of shortterm acidification on growth rates of tadpoles. J. Herpetol. 23: 97-102. Pierce, B. A. and Wooten, D. K., 1992a. Genetic variation in tolerance of amphibians to low pH. J. Herpetol. 26: 422429.

3265

Preest, M., 1993. Mechanisms of growth rate in acidexposed larval salamanders, Ambystoma maculatum. Physiol. Zool. 66: 686-707. Punzo, F., 1983. Effects of environmental pH and temperature on embryonic survival capacity and metabolic rates in small mouth salamander, Ambystoma texanum. Bull. Envirm. Cmtum. Toxicol. 31: 467-473. Relyea, R. A,, 2006. The effects of pesticides, pH, and predatory stress on amphibians under mesocosm conditions. Ecotoxicology 15: 503-5 11. Renberg, I., Korsman, T. and Anderson, N. J., 1993. A temporal perspective of lake acidification in Sweden. Ambio 22: 264-271. Rosenberg, E. A. and Pierce, B. A., 1995. Effect of initial mass on growth and mortality at low pH in tadpoles of Pseudacris clarkii and Bufo valliceps. J. Herpetol. 29: 181-185. Rowe, C. L. and Dunson, W. A., 1993. Relationships among abiotic parameters and breeding effort by three amphibians in temporary wetlands of Central Pennsylvania. Wetlands 13: 237-246. Rowe, C. L. and Dunson, W. A,, 1994. The value of simulated pond communities in mesocosms for studies of amphibian ecology and ecotoxicology.J. Herpetol. 28: 346-356. Rowe, C. L., Sadinski, W. J. and Dunson, W. A., 1992. Effects of acute and chronic acidification on three larval amphibians that breed in temporary ponds. Arch. Environ. Cmtam. Toxicol. 23: 339-350. Rowe, C. L., Sadinski, W. J. and Dunson, W. A,, 1994. Predation on larval and embryonic amphibians by acid-tolerant caddisly larvae (Ptilostomis postica). J. Herbetol. 28: 357-364.

Pierce, B. A. and Wooten, D. K., 1992b. Acid tolerance of Ambystoma texanum from central Texas. J. Herpetol. 26: 230-232.

Rowe, C. L. and Freda, J., 2000. Effects of acidification on amphibians at multiple levels of biological organization. Pp. 545-571 in "Ecotoxicology of Amphibians and Reptiles", ed by D. W. Sparling, G. Linder and C. A. Bishop. Society of Environmental Toxicology and Chemistry (SETAC) Press, Pensacola.

Pierce, B. A., Hoskins, J. B. and Epstein, J. B., 1984. Acid tolerance in Connecticut wood frogs (Rana sylvatica). J. Herpetol. 18: 159-167.

Rusek, J. and Marshall, V G., 2000. Impacts of airborne pollutants on soil fauna. Ann. Rev. Ecol. System. 31: 95423.

Pleijel, H., Andersson, I. and Lovbla, G., 1999. Acidification in 2010: An assessment of the situation at the end of the next decade. Air Pollutants and Climate, Series 10: 1-30. The Swedish NGO Secretariat on Acid Rain, Gothenburg.

Rasanen, K., 2002. Evolutionary implications of acidification: a frog's eye-view. Ph.D. Dissertation, Uppsala University, Uppsala, Sweden.

Porter, K. R. and Hakanson, D. E., 1976. Toxicity of mine drainage to embryonic and larval boreal toads (Bufonidae - Bufo boreas). Copeia 1976: 327-33 1. Portnoy, J. W., 1990. Breeding biology of the spotted salamander Ambystoma maculatum (Shaw) in acidic temporary ponds at Cape Cod, USA. Biol. Cons. 53: 61-76. Pough, F. H., 1976. Acid precipitation and embryonic mortality of Spotted Salamanders Ambystoma maculatum. Science 192: 68-72. Pough, F. H. and Wilson, R. E., 1977. Acid precipitation and reproductive success of Ambystoma salamanders. Watel; Air and Soil Pollut. 7: 531-544.

Riisanen, R,Laurila, A. and Merila, J., 2002. Carry-over effects of embryonic acid conditions on development and growth of Rana temporaria tadpoles. Freshw. Biol. 47: 19-30. Rasaneu, K., Laurila, A. and Merila, J., 2003a. Geographic variation in acid stress tolerance of the moor frog, Rana arvalis. I. Local adaptation. Evolution 57: 352-362. Riisanen, K., Laurila, A. and Meril, J., 2003b. Geographic variation in acid stress tolerance of the moor frog, Rana arvalis. 11. Adaptive maternal effects. Evolution 57: 363-371. Riisanen, K., Laurila, A. and Merila, J., 2005. Maternal investment in egg size: environment- and populationspecific effects on offspring performance. Oecologia 142: 546-553.

3266

AMPHIBIAN BIOLOGY

Saber, P A. and Dunson, W. A., 1978. Toxicity of bog water to embryonic and larval anuran amphibians. J. Exp. Zool. 204: 3 3 4 2 . Sadinski, W. J. and Dunson, W. A., 1992. A multilevel study of effects of low pH on amphibians of temporary ponds. J. Herpetol. 26: 413422. Salthe, S. N., 1963. Egg capsules in amphibia. J . Morphol. 113: 161-171. Schindler, D. W., 1988. Effects of acid rain on freshwater ecosystems. Science 239: 149-157. Schindler, D. W., Curtic, P J., Parker, B. R. and Stainton, M. P, 1996. Consequences of climate warming and lake acidification for W - B penetration in North American boreal lakes. Nature 379: 705-708. Schlichter, L. C., 1981. Low pH affects the fertilization and development of Rana pipiens eggs. Can. J. Zool. 59: 1693-1699. Scott, D. E., 1990. Effects of larval density in Ambystoma opacum: an experiment in large-scale field enclosures. Ecology 71: 296-306. Scott, D. E., 1994. Effects of larval density on life history traits in Ambystoma opacum. Ecology 75: 1383-1396. Semlitsch, R. D., Scott, D. E. and Pechmann, J. H. K., 1988. Time and size at metamorphosis related to adult fitness in Ambystoma talpoideum. Ecology 69: 184-192. Semlitsch, R. D., Bridges, C . M. and Welch, A. M., 2000. Genetic variation and a fitness tradeoff in the tolerance of gray treefrog (Hyla uersicolor) tadpoles to the insecticide carbaryl. Oecologia 125: 179-185. Simon, M. P, Vatnick, I., Hopey, H. A,, Butler, K., K o N ~ C., ~ , Hilton, C., Weimann, R. S. and Brodkin, M. A,, 2002. Effects of acid exposure on natural resistance and mortality of adult Rana pipiens. J. Herpetol. 36: 697-699. Skelly, D. K., 2001. Distributions of pond-breeding anurans: An overview of mechanisms. Israel J. Zool. 47: 313-332. Skelly, D. K., 2002. Experimental venue and estimation of interaction strength. Ecology 83: 2097-2 101. Skelly, D. K. and Kiesecker, J. M., 2001. Venue and outcome in ecological experiments: manipulations of larval anurans. Oikos 94: 198-208. Skelly, D. K., Yurewicz, K. L., Werner, E E. and Relyea, R. A,, 2003. Estimating decline and distributional change in amphibians. Cons. Biol. 17: 744-75 1. Smith, M. A. and Green, D. M., 2005. Dispersal and the metapopulation paradigm in amphibian ecology and conservation: are all amphibian populations metapopulations? Ecography 28: 110-1 28. Soiseth, C. R., 1992. The p H and acid neutralizing capacity of ponds containing Pseudacris regzlla larvae in an alpine basin of the Sierra Nevada. Cal$ornia Fish and Game 78: 11-19.

Steinberg, C. E. W., Kamara, S., Prokhotskaya, V. Yu., Manusadzianas, L., Karasyova, T A., Timofeyev, M. A,, Jie, Z., Paul, A., Meinelt, T., Farjalla, V. F., Matsuo, A. Y O., Burnison, B. K. and Menzel, R., 2006. Dissolved humic substances - ecological driving forces from the individual to the ecosystem level? Freshw. Biol. 51: 1189-1210. Stoddard, J. L., Jeffries, D. S., Liikewille, A., Clair, T A, Dillon, P J., Driscoll, C. T, Forsius, M., Johannessen, M., Kahl, J. S., Kellogg, J. H., Kemp, A., Mannio, J., Monteith, D. T., Murdoch, P S., Patrick, S., Rebsdorf, A,, Skjelkvsle, B. L., Stainton, M. P, Traaen, T, Van Dam, H., Webster, K. E., Wieting J. and Wilander, A. J., 1999. Regional trends in aquatic recovery from acidification in North America and Europe. Nature 401: 575-578. Strand, L. A., 2002. Reproduksjon hos amfibier i vann med ekstreme pH-verdier. Fauna 55: 108-1 14. Strijbosch, H., 1979. Habitat selection of amphibians during their aquatic phase. Oikos 33: 363-372. Surova, G. S., 2002. The influence of high water acidity on the viability of large and small eggs of Rana temporaria. Zoologzchesky Zhurnal 81: 608-6 16. Soderman, F., 2006. Comparative Population Ecology in Moor Frogs with Particular Reference to Acidity. Ph.D. Dissertation, Uppsala University, Uppsala, Sweden. Soderman, F., van Dongen, S., Pakkasmaa, S. and Merila, J. Environmental stress increases skeletal fluctuating asymmetry in the moor frog, Rana arualis. Oecologia 151: 593-604. Tattersall, G. J. and Wright, P A,, 1996. The effects of ambient pH on nitrogen excretion in early life stages of the American toad (Bufo americanus). Compar: Biochem. Physiol. A 113: 369-374. Tome, M. A. and Pough, F. H., 1982. Responses of amphibians to acid precipitation. Pp. 245-254 i n "Acid RainIFisheries", ed by R. E. Johnson. American Fisheries Society, Bethesda, Maryland. Travis, J., Keen, W. H. and Julianna, J., 1985. The role of relative body size in a predator-prey relationship between dragonfly naiads and larval anurans. Oikos 45: 59-65. Trippel, E. A. and Harvey, H. H., 1987. Reproductive responses of 5 white sucker (Catostomus commersoni) populations in relation to lake acidity. Can. J. Fish. Aquat. Sci. 44: 1018-1023. Tyler-Jones, R., Beattie, R. C. and Aston, R. J., 1989. The effects of acid water and aluminum on the embryonic development of the common frog, Rana temporaria. J. Zool. 219: 355-372. Urch, U. A. and Hedrick, J. L., 1981. Isolation and characterization of the hatching enzyme from the amphibian, Xenopus laevis. Arch. Bzochem. Biophys. 206: 424-43 1.

Spry, D. J. and Wiener, J. G., 1991. Metal bioavailability and toxicity to fish in low-alkalinity lakes: a critical review. Environ. Pollut. 71: 243-304.

Vamick, I., Brodkin, M. A., Simon, M. P, Grant, B. W., Conte, C. R., Gleave, M., Myers, R. and Sadoff, M. M., 1999. The effects of exposure to mild acidic conditions on adult fmgs (Rana pipiem and Rana clamitans): Mortality rates and pH preferences. J. Herpetol. 33: 370-374.

Steinberg, C . E. W. and Wright, R. W. (Editors), 1994. "Acidification of Freshwater Ecosystems: Implications for the Future". Wiley, New York.

Verma, N. and Pierce, B. A,, 1994. Body masss, developmental stage, and interspecific differences in acid tolerance of larval anurans. TexasJ. Sci. 46: 319-327.

and GREEN: ACIDIFICATION AND AMPHIBIANS

-ANEN

3267

Vertucci, F. A. and Corn, P. S., 1996. Evaluation of episodic acidification and amphibian declines in the Rocky Mountains. Ecol. Applic. 6: 449457.

Wilbur, H. M., 1982. Competition between tadpoles of Hyla femoralis and H y h gratiosa in laboratory experiments. Ecology 63: 278-282.

Wake, D. B., 1991. Declining amphibian populations. Science 253: 860.

Wissinger, S. A and \ a t e m a n , H. H., 1992. Fluctuation in a Rocky Mountain population of salamanders: anthropogenic acidification of natural variation? J. Herpetol. 26: 377-391.

Warner, S. C. and Dunson, W. A., 1998. The effect of low DH on a m ~ h i b i a n sbreeding in t e m ~ o r a r v ponds in North lorida. Fla ~ame"and~ r e s d Fish Comm., Final Report, Tallahassee. 88 pp + vi.

water

Warner, S. C., Dunson, W. A,, Travis, J., 1991. Interaction o f p ~density, , and priority on the suIvivors~p and growth of 2 species of hylid tadpoles. Oecologta 88: 331-339. Warner, S. C., Travis, J. and Dunson, W. A,, 1993. Effect of pH variation on interspecific competition between two species of Hylid tadpoles. Ecology 74: 183-194. Whiteman. H. H.. Howard. R. D. and Whitten. K. A,. 1995.'~ffects'ofp~ on'embryo tolerance and adul; behaviour in the tiger salamander, Ambystoma tigrinum t i p n u m . Can. J . Zool. 73: 1529-1537. --

L,lw,tat.r hpFoiog Cons. Bwl. 4: 390-352.

amphbms?

Wyman, R. L. and Hawksley-Lescault, D. S., 1987. Soil acidity affects distribution, behavior, and physiology of the salamander Plethodon cinereus. Ecology 68: 1819-1827. W ~ m a n ,R. L. and Jancola, J., 1992. DeFFe and scale of terrestrial acidification and amphibian community structure. J. Herpetol. 26: 392401.

- - --

Zimmer, K. D., Hanson, M. A, Butler, M. G., 2002. meets of fathead minnows and restoration on prairie wetland ecosystems. Freshw. Biol. 47: 2071-2086.

CHAPTER 12

Climatic Change and Amphibian Declines Patricia A. Burrowes

I. Introduction 11. Climatic Change and Amphibian Biology Ill. Studies Linking Amphibian Declines to Climatic Change

B. Distributional Shifts and Inability to Adapt to Rapid Changes C. Synergies Leading to Epidemic Disease 1. Chytrids and Climatic Change 2. Saprolegnia ferax and Climatic Change

V. Conclusions IV. Consequences to Amphibians of Climatic Change A. Change in Breeding Phenology

VI. References

I. INTRODUCTION

A

NTHROPOGENIC-DRIVEN climatic change in the 20th century has triggered scientist's concern for its consequences on the earth's biodiversity (Parmesan and Yohe 2003; Root et al. 2003, 2005; Thomas et al. 2004; Parmesan 2006). These scientists have been termed "alarmists" by the media on the basis that climatic change has been a constant in the history of the Earth. Studies of long-term palaeoecological records, however, have shown that although Earth has a history of intense temperature changes, these did not occur at the rapid rates that have been documented for the past few decades, and that are predicted for the 21st century (Bush et al. 2004). The potential for climatic change to decrease the quality of humans' lives has given rise to scientific-based, socio-economic debates at many international fora such as the Intergovernmental Panel on Climate Change (IPCC) and the World Summit on Sustainable Development, Johannesburg, 2002. Major concerns centre around some frightening facts that global mean surface temperatures have increased 0.74"C over the past 100 years and that, in the period between 1906 and 2006, 11 of the 12 warmest years occurred recently, between 1995 and 2006 (Meehl et al. 2007). Analysis of maximum and minimum surface daily temperatures show that minimum daily temperatures are rising twice as fast as daily maxima: 0.2"C versus O.l°C per decade (Houghton et al. 2001). In addition, studies have shown that surface sea temperatures (SST) in the tropics increased significantly in the mid-1970s leading to a rise in elevation of about 100 m for freezing conditions at the surface in tropical mountains (Beniston et al. 1997; Keisecker et al. 2001b). The correlation between tropical SST and surface temperatures in the mountains suggests that high-elevation environments in the tropics may be particularly sensitive to changes in SST, and that prolonged El Nifio-like episodes could have a marked

BURROWES: CLIMATIC CHANGE AND AMPHIBIAN DECLINES

3269

impact on their biota (Beniston et al. 1997; Pounds et al. 1999, 2006). Although many uncertainties have been acknowledged to influence prediction of species' extinction risks under scenarios of climatic change, it is recognized that man-driven global warming is likely to be the greatest present threat to the biodiversity of the planet (Thomas et al. 2004). The present chapter reviews studies within the past 15 years that link climatic change to amphibian population declines, discusses the consequences for amphibian biology, and describes hypothesized climate-driven synergistic interactions that may be putting amphibians under a serious threat of extinction. 11. CLIMATIC CHANGE AND AMPHIBIAN BIOLOGY

When it became apparent that amphibian populations were declining worldwide (Wake and Morowitz 1991) and an appeal went out from the Declining Amphibian Populations Task Force (DAPTF) to assemble evidence of these declines and search for potential causes, many herpetologists considered climatic change as a potential culprit. This is understandable because amphibians are terrestrial ectotherms with thin skins and their biology is highly determined by the temperature and humidity of their environments (Duellman and Trueb 1994; Corn 2005). Heat exchange with their surroundings determines their body temperatures (Hutchinson and Dupre 1992) and hence affectstheir metabolic processes (Rome et al. 1992). Gametogenesis and the growth of larvae and post-metamorphic individuals are temperaturedependent (Beebee 1995; Carey et al. 2003). These temperature-driven physiological responses are also tied to specific behaviours, such as emergence from hibernation, and to onset of reproduction (Blaustein et al. 2001; Carey et al. 2003; Corn 2005) in temperate species, as well as to activity patterns in tropical species (Pough et al. 1983). Water availability has been considered even more important than temperature in determining amphibian distributions around the globe (Duellman 1999; Araujo et al. 2006). Carey and Alexander (2003) discussed the importance of water availability for the -sss-d\sk~hie?e-s -4iLa~pkikiassas&<s t. kestakiky afpapxkatizsizes.&ter;at_ian-ix the rainfall pattern resulting in drier years can (1) increase vulnerability to desiccation of terrestrial eggs, larvae and metamorphosing young; (2) reduce sizes of ponds, thereby altering density-dependent mechanisms such as size at metamorphosis and predator-prey interactions; (3) increase the concentration of contaminants in bodies of water and (4) affect the ability of adults to maintain water balance (Carey and Alexander 2003). Extension of the dry season in tropical environments is another aspect of climatic change that has been linked to the decline of amphibians (Donnelly and Crump 1998). Some examples are the extinction of Bufo periglenes in Costa Rica (Crump et al. 1992) and the decline of highmountain, direct-developing anurans in Puerto Rico (Stewart 1995; Burrowes et al. 2004). Terrestrial amphibians experience a greater risk of desiccation because of their higher rates of water loss through the skin and respiratory system, and because of their increased levels of urine concentration (Shoemaker et al. 1992). For example, drought affects the behaviour and activity pattern of Eleutheroductylus coqui. Three days without rain can kill juveniles, and adult males stop calling and remain at their retreat sites after five days without water (Pough et al. 1983; Stewart 1995). This behaviour can lead to population declines through reduced reproductive activity and limited recruitment of juveniles. Thus, at times of extensive dry periods, amphibians could suffer drastic population declines should they be unable to find moist sites for rehydration. Their ability to respond rapidly to changes in water availability, therefore, represents a selective pressure that may drive different patterns of amphibian declines across geographical areas and taxonomic groups. Later in this chapter there is a discussion of how factors related to local climatic change can act synergistically to decrease amphibian immunity, and thus vulnerability, to disease. 111. STUDIES LINKING AMPHIBIAN DECLINES T O CLIMATIC CHANGE

The majority of studies that consider the response of organisms to climatic change are based on correlations or on anecdotal associations (McCarty 2001; Corn 2005). This is the

3270

AMPHIBIAN BIOLOGY

case for most amphibians. Carey and Alexander (2003) pointed out that cause and effect must be determined by experiments where variables can be controlled and suggest several approaches from the field of epidemiology to help scientists determine if there is a direct, or indirect, link between climatic change and amphibian declines. Testing the hypotheses that increasing temperatures or droughts are driving amphibian declines is complicated because climatic variables are so indistinctly associated with other factors that it is difficult to design experiments that will demonstrate a direct causation (Araujo et al. 2006). The challenge for investigators interested in this issue is to collect data that will differentiate real long-term climatic patterns from yearly fluctuations - "detection", and that can distinguish climatic change as the cause of amphibian declines within the midst of confounding factors - "attribution" (Parmesan 2006). Nonetheless, it is important to pay attention and study these associations because they may explain the role of other factors interacting with climate and resulting in the loss of biodiversity at the unprecedented rate reported for amphibians (Stuart et al. 2005). The first studies to associate climatic change to amphibian population declines, extinctions or community changes were those by Heyer et al. (1988) and Weygoldt (1989) in southern Brazil. Heyer et al. (1988) hypothesized that severe frost was the cause for the disappearance of five species in Brazil from 1979 to 1982 and Weygoldt (1989) linked drastic population declines of other amphibians to an unusually dry winter. Although their work was mostly anecdotal, it raised concern among herpetologists. Soon afterwards, several efforts to correlate amphibian abundance with climatic variables became available in the literature. Years of drought were correlated with local extinctions of the northern leopard frog, Rana pipiens, in the United States (Corn and Fogleman 1984) and with massive declines of streamdwelling species in Australia (Ingram 1990). In the cloud forests of Costa Rica, the decline of several species of frogs, including the disappearance of the golden toad, Bufo perzglenes, and the harlequin frog, Atelopus varius, in the late 1980s was correlated with extension of the dry season and with decrease in the frequency of precipitation of mist in the dry season (Crump et al. 1992; Pounds and Crump 1994; Pounds et al. 1999). Declines in lowland species of anurans in Costa Rica were also correlated with the extension of the dry season during the 1980s (Donnelly and Cmmp 1998). In South America, amphibian declines became evident in Ecuador beginning in the mid-1980s when biologists noticed species absent from localities where they had been abundant in the past (Vial and Saylor 1993; Coloma 1995, 2002; Ron and Merino-Viteri 2000). Quantitative evidence of these declines was presented by Ron et al. (2003) and Bustamente et al. (2005). Analysis of climatic data from 1891 to 1999 showed abnormalities in the Andes of Ecuador for the last two decades of the 20th century, consisting of an unusual combination of high temperature, dry days and low precipitation (Ron et al. 2003; Merino Viteri et al. 2005). Twenty-four amphibian species are considered threatened in Ecuador, of which 79% were last seen between 1988 and 1996, 96% occur at mid to high elevations and 87% have aquatic larvae (calculated from Ron et al. 2001-2006). It is evident from the studies cited above that climatic changes unfavourable for amphibians occurred in the Andes during the time in which amphibian declines were more pronounced. The facts that most affected species in Ecuador live in the Andes and are dependent on water for reproduction suggest a potential causative effect between climatic anomalies and declines (Merino-Viteri 2001; Ron et al. 2003; Bustamante et al. 2005; Merino-Viteri et al. 2005). In the Caribbean, declines of the Puerto Rican coqui, Eleutherodactylus coqui, from the Luquillo Mountain range between 1983 and 1989 were correlated with an increase in number of extended dry periods consisting of more than five days with less than 3 mm of rain (Stewart 1995). Later, Burrowes et al. (2004) analysed weather data for the last 30 years of the 20th century and showed that populations of eight different species of Eleutherodactylus, and the disappearance of three species (E. karlschmidti, E. jasperz and E. enezdae) from forested montane habitat in Puerto Rico were correlated with significantly drier years in the early 1970s and 1990s. That study also showed a marked extension of the dry season and a significantly greater number of dry periods for the years in which declines

BURROWES: CLIMATIC CHANGE AND AMPHIBIAN DECLINES

3271

were observed in Puerto Rico (Burrowes et al. 2004). Thus, that study corroborates the observations by Stewart (1995) and Donelly and Crump (1998). Some studies have suggested that climatic change cannot be the sole cause of amphibian declines because no major climatic anomalies were found concurrent with declines (Laurance 1996; Lips et al. 2006). Furthermore, studies that used comprehensive time-series of climatic datasets from remote and ground stations concluded that although amphibian declines in Colorado, Puerto Rico, Costa Rica, Panama and Australia coincide with times when temperature and drought increased, variations of these climatic factors were not extreme enough to be considered the direct cause (Alexander and Eischeid 2001; Stallard 2001). Davidson et al. (2001) found that the patterns of declines of the California red-legged frog (Rana aurora) were not consistent with those predicted by climatechange hypotheses because they did not follow the anticipated north to south gradient in declines, decrease in declines with elevation, or the association of declines with mean precipitation. Caution is advised, however, when ruling out the effect of climatic change on amphibian declines. Recent data and model simulations show that the effects of climatic change are not uniform around the globe, but rather display different patterns in different regions (Christensen et al. 2007). Separating the effect of multiple climatic variables on amphibian populations is very difficult and will require experimental manipulation of temperature, moisture, wind, and their combinations, on such attributes as amphibian tolerance, adaptability, and vulnerability to potential diseases. It is thus very difficult to predict when or where climate-driven amphibian declines are really observed. Discussed below are several efforts to describe the effect that global climatic changes and their influence in local climate variability may have on amphibian populations. IV. CONSEQUENCES TO AMPHIBIANS OF CLIMATIC CHANGE A. Change in Breeding Phenology The timing of amphibian breeding is driven by various environmental signals (Duellman 1999; Carey and Alexander 2003) and thus global warming and related changes in water availability may directly affect breeding phenology. In temperate regions, amphibians spend a large portion of the year aestivating or hibernating, thereby escaping the extreme temperatures of both summer and winter. Subtle increases in temperature or moisture may trigger emergence from hibernation and immediate migration to ponds to breed. One of the predicted effects of global warming is therefore an early onset of reproduction (Beebee 1995; Blaustein et al. 2001; Gibbs and Breisch 2001). This has been documented in 53% of the amphibians for which breeding patterns were studied in North America and England (calculated from Blaustein et al. 2003). It has been hypothesized that early breeding may result in population declines due to increased risk of mortality of embryos from snowmeltinduced floods, and from episodic freezes in early spring (reviewed by Corn 2005). Variation among temperate-zone amphibians in the tendency to breed earlier in response to warmer temperatures suggests that, as with other environmental factors, there are taxonomic differences in their physiological responses (Blaustein et al. 2003). Therefore, one would expect some species to be more vulnerable than others and efforts should be made to identify species that may need intervention of management programmes. B. Distributional Shifts and Inability to Adapt to Rapid Changes Upward elevational shifts in the distribution of amphibians, anoline lizards and some bird species in the cloud forest of Monteverde, Costa Rica, have been related to global warming (Pounds et al. 1999). In a comprehensive study of the response of 1 700 species (trees, herbs, shrubs, reptiles, amphibians, fish, marine zooplankton and invertebrates, mammals, birds and butterflies) to climatic change in the twentieth century, Parmesan and Yohe (2003) found that 87% of the species showed phenological changes in the direction predicted by climatic change. The trends observed included anticipation of breeding time

3272

AMPHIBIAN BIOLOGY

by anurans, nesting by birds, flowering of plans, budbursts of trees, and arrival of migrant populations of birds and butterflies (Parmesan and Yohe 2003). These authors also found that approximately 50% of the 893 species considered showed significant distributional changes over a mean period of 66 years. Of these, 80% shifted latitudinally or elevationally toward cooler regions, as predicted by scenarios based on change in climate (Parmesan and Yohe 2003). Distributional shifts do not always imply range expansions. Many times it involves range contractions and a risk of population decline due to inability to adapt to new biotic and/or abiotic factors (Parra et al. 2005; Araujo et al. 2006). This is already evident in Puerto Rico for two montane species, Eleutherodactulus g ~ l l u sand E. unicolor (Joglar 1998). Using four different methods to model species distributions as a response to scenarios of climatic change predicted for 2050, Araujo et al. (2006) examined potential distributional changes for European amphibians and reptiles. These authors considered models that assumed unlimited versus limited dispersal abilities for the species. The results of their work revealed that climate-space would expand for the majority of amphibian and reptile species considered. This is predicted because climatic change will bring higher temperatures. The physiological tolerances of amphibians are greater for warmer than for cooler conditions (Snyder and Weathers 1975), thereby promoting range expansions into previously cooler areas. Considering the low viability of amphibians, however, the authors acknowledged unlimited dispersal models to be unrealistic for this group of animals (Araujo et al. 2006). Assuming limited dispersal abilities, their models anticipated major losses and/or range contractions among amphibian species in the Iberian Peninsula, southern France, Italy and Eastern European countries. Geographic areas with warmer, and particularly drier, climates were predicted to become unsuitable for many species of amphibians by the year 2050, suggesting that water shortage might be a significant climatic factor driving amphibian declines (Araujo et al. 2006). In the Andes of Ecuador, where temperature increased an average of 1.9"C per month in the period 1990-1999 (four times greater than the average increase reported for the world), six species of anurans (three Eleutherodactylus and three Hyla) have shifted upward by an average of 430 m to cooler conditions (Bustamante et al. 2005). Although it is evident that some species can adapt to climatic change by making phenological or distributional shifts, organisms that lack the ability for rapid change may face extinction. This is the case for many montane, cool-adapted amphibians that have low tolerance for thermal change. Bernardo and Spotila (2005) studied two montane salamanders, Desmognathus carolinensis and D. ocoee, along an elevational cline in the Appalachian Mountains of eastern North America. Populations at different elevations were exposed to an array of temperatures between 5°C and 20°C. The magnitude of metabolic depression with increasing temperature was inversely related to native elevation suggesting that salamanders from low to mid elevations were already living near the limit of their physiological tolerance. If other montane amphibian species behave similarly, the projection for loss of biodiversity in montane ecosystems is frightening (Bernardo and Spotil 2005). Parra et al. (2005) used Ecological Niche Modelling to predict the distributional shifts of two montane species of plethodontid salamanders in central Mexico under the predicted scenario of climatic change for the year 2050. Results showed drastic reduction and fragmentation of potential distributional areas characterized by an elevational shift to volcanic peaks. At present, these volcanic areas do not exhibit suitable habitat for terrestrial amphibians. Because habitat is unlikely to change so rapidly (50 years) and salamanders are known to have low dispersal abilities, the risk of extinction of these amphibians is very high (Parra et al. 2005). In Europe, the trend toward range contraction under the predicted climatic scenario for 2050 was significantly higher for the order Caudata compared to Anurans and to three orders of reptiles (Araujo et al. 2006). Elevational shifts in distribution have also been observed in typical lowland-adapted species. For example, the cane toad (Bufo mminus) has been found up to 2 000 m in Guatemala (Campbell 1999) and 2 500 m in Ecuador (Bustamante et al. 2005), and Eleutherodactylus antillensis, a typically lowland species, can now be heard up to 850 m in Puerto Rico (Joglar

BURROWES: CLIMATIC CHANGE AND AMPHIBIAN DECLINES

3273

1998). Species' distributional changes result in alteration of ecological communities that can lead to the decline of native species due to predation andlor competition (Knapp and Mathews 2000; Kiesecker et al. 2001b; Kiesecker 2003). When the invader is a highly competitive exotic, however, there is also the potential for spread of pathogens to more vulnerable native species (Kiesecker 2003). Known potential carriers of amphibian diseases are hatchery-reared fish introduced into freshwater lakes for recreational fishing in the Pacific Northwest of the United States (Kiesecker et al. 2001c, 2003), Ram catesbeiuna in South America (Mazzoni et al. 2003; Daszak et al. 2004) and B u . marinus in Australia (Speare et al. 1990) and Puerto Rico (Burrowes et al. 2004). C. Synergies Leading to Epidemic Disease

Possible consequences of an increase in temperature on host-pathogen relationships include change in the pathogen's developmental rate, dispersal, transmission or lifespan and/or change in the host's vulnerability to infection (Harvell et al. 2002). Climatic change may create an environmental scenario that facilitates outbreaks of disease or triggers physiological and behavioural responses that lead to amphibian declines. The influence of climatic change on an amphibian's response to pathogens will depend, however, on its natural history, geographical location, evolutionary history and other interacting factors (Pounds 2001). The three most common diseases implicated in amphibian mass mortalities or population declines are (1) cutaneous infection by a pathogenic chytrid fungus, Batrachochytrium dendrobatzdis (Longcore et al. 1999), (2) infestation of eggs by a pathogenic oomycete fungus, Saprolegniu firax (Kiesecker and Blaustein 1995; Kiesecker et al. 2001b) and (3) infection by a group of iridoviruses of the genus Ranavirus (Daszak et al. 1999). Of these, the first two have been suggested to interact synergistically with climatic factors that increase the severity of the disease on amphibian populations or on entire communities. These potential interactions are reviewed in the following paragraphs. 1 . Chytrids and Climatic Change

Pounds and Crump (1994) proposed the "climate-linked epidemic" hypothesis as a potential explanation for the declines of amphibians at Monte Verde, Costa Rica. This hypothesis suggests that, when climatic change creates suboptimal regimens of temperature and humidity, amphibians suffer a negative impact on their behaviour and on their energy budget. As a consequence, populations tend to move from a dispersed to a clumped distribution, which makes them more vulnerable to disease. This was also suggested as an explanation for chytrid-related declines of several species of terrestrial frogs in the genus Eleutherodactylzls in Puerto Rico (Burrowes et al. 2004). It was proposed that drought-stressed E. coqui infected by B. dendrobatidis would tend to clump in humid patches of the forest and share the few wet retreats still available. In addition, it was proposed that terrestrial fi-ogs would be more likely to die from B. dendrobatzdzs infections during the dry season because cutaneous infection may interfere with ability of the frogs to absorb dew, the only source of water during droughts. The presence of this pathogen in Puerto Rico at approximately the same time that anuran populations declined and extirpations began, and when the climate was significantly drier than average, suggests a potential synergy between dimate and disease (Burrowes et al. 2004). In recent years the present author has performed controlled laboratory experiments to test the hypothesis linking climate with epidemics of B. &robatidis in Puerto Rico. It was found that, indeed, frogs stressed by dehydration tend to clump on humid substrates in terraria by sharing daily retreats, whereas waterindulged frogs exhibit a uniform distribution (Longo et al. 2006). This may be a general amphibian response as Heatwole (1960) found that red-backed salamanders, Plethodon auretls, also clumped in response to drying conditions in Michigan. When inoculated with A t dendrobatzdis zoospores, Puerto Rican frogs exposed to drought treatments (Fig. 1) died signilicantly faster than did frogs with unlimited access to water (Burrowes and Longo, apublished data). Field studies also corroborated these results, showing that prevalence dlevels of infections by B. dendrobatidis in adult frogs are higher during the dry months

3274

AMPHIBIAN BIOLOGY

Fzg. 1. A frog (Eleutherodactylus coqui) dying from infection by Batmchochytrium dendrobatidis, under an experimental drought regime. Note the areas of skin necrosis (white arrow), the glossy skin, and the stretched-out hind legs, typical of very sick frogs. Photo by C. Andres Rodriguez.

of the year. These data provide evidence in favour of the climate-linked epidemic hypothesis as an explanation for B. dendrobatzdis persistence in a seasonal tropical environment. In addition, it suggests that under increasing drought, as has been predicted for the Caribbean under the scenario of climatic change for the next 50 years (Christensen et al. 2007), there is a potential for a chytrid epizootic that may result in drastic adult mortality. The association between climate and prevalence of B. dendrobatidis in amphibians was also noticed in Australia. Retallick et al. (2004) conducted a longitudinal mark-recapture study of a community of stream-dwelling frogs in northern Queensland where several species were declining, or had disappeared, due to infection by B. dendrobatidis. They found that over a four-year period (1994-1998) the prevalence of chytridiomycosis was consistently higher during the cooler and drier months of the year (Retallick et al. 2004). This pattern was confirmed by another study on distribution, incidence and pathogenicity of B. dendrobatidis in eastern Australia held between the years 1993 and 2000 (Berger et al. 2004). Those authors reported that the number of wild frogs that died from chytridiomycosis in Queensland and New South Wales was higher in the winter, and that the pathogenicity of B. dendrobatzdis in laboratory experiments increased at lower temperatures (17-23°C). A more recent study sampled a population of Litoria wilcoxii for a distance of 1 km at low elevation in southeastern Queensland (Kriger and Hero 2006). There was a significant effect of seasonality on the prevalence of B. dendrobatzdis among this non-declining species. The peak of chytridiomycosis occurred during the cooler months of the year when temperatures ranged between 12.3"C and 19.4"C. The highest infection levels were also associated with the drier months but the effect of rainfall was confounded by the fact that winters are typically dry in this part of Australia (Kriger and Hero 2006). It is not surprising to find higher prevalence of B. dendrobatidis during cooler periods, because laboratory experiments have shown that the thermal optimum for growth of this

BURROWES: CLIMATIC CHANGE AND AMPHIBIAN DECLINES

3275

fungus is between 17°C and 25°C (Piotrowski et al. 2004), which is lower than summer temperatures in temperate regions, or lowland temperatures in tropical areas. One mysterious part of the puzzle is that this fungus, which has an infective flagellated zoospore stage (Longcore et al. 1999), appears to be more virulent during drier seasons in different parts of the world (Burrowes 2004; Retallik et al. 2004; Kriger and Hero 2006). It is possible that the stress from drought alone may render the frogs more susceptible to the disease, or that the physiology of the disease in the skin may prevent uptake of water horn the substrate at a time when rain is scarce. Another enigma is that the pathogen seems to flourish in very warm years. Data on the status of the brightly coloured harlequin frogs (Bufonidae: Atelopus) from South and Central America show that many species disappeared immediately after a relatively warm year (La Marca et al. 2005; Ron et al. 2003). Analysing patterns from the Atelopus database, Pounds et al. (2006) found that the number of species lost was greatest at middle elevations (1 001-2 399 m) and related this to thermal optimum for growth of B. dendrobatzdis. Lower elevations would be too warm for the pathogen and higher elevations too cold. Those authors analysed climatic data for the tropics and showed that rising mean air temperatures and surface sea temperatures are positively correlated with the number of dry periods and with minimum temperatures at Monteverde, a mountain reserve in Costa Rica. They proposed that global warming in the tropics is accelerating evaporation and causing increased cloud cover in the mountains, particularly at midelevation. The clouds hinder solar radiation during the day, thereby decreasing daily maximum temperatures, but trap heat during the night and increase minimum temperatures (Pounds et al. 2006). The result is a localized thermal envelope that is optimum for growth of B. dendrobatzdis. They proposed the "chytrid-thermal-optimum hypothesis" to explain the declines and disappearances of many species of frogs in Central and South America at middle elevations (Pounds et al. 2006). The effect of global warming in different regions of the world may create diverse environmental conditions that may favour growth of B. dendrobatzdis. Were temperature the only factor affecting B. dendrobatzdis growth, Kriger and Hero (2006) suggested that global warming trends would reduce the susceptibility of lowland tropical and subtropical amphibians to chytridiomycosis because maximum daily temperatures would be too high. On the other hand, tropical upland and temperate amphibians would be more vulnerable, because a favourable thermal envelope for B. dendrobatzdis (sensu Pounds et al. 2006) would prevail in those regions (Kriger and Hero 2006). Precipitation patterns, community ecology, evolutionary history of the amphibians, and the physiology of the disease, however, may act synergistically to create more complex scenarios that trigger an epidemic, or favour reservoir species that can carry the pathogen at low loads. In order to further understand climatedriven synergies, it is necessary to monitor how different climatic signals affect amphibian populations and their susceptibility to disease. At present, it is advisable that conservation and management programmes prioritize efforts to protect montane tropical and lowland temperate species and be ready to implement emergency plans involving captive breeding as recommended by the Amphibian Conservation Action Plan (Mendelson et al. 2006). 2. Saprolegnia ferax and Climatic Change

Mortality of amphibian embryos in the Pacific Northwest of the United States has been associated with outbreaks of an aquatic oomycete fungus, Saprolegnia ferax, in conjunction with increased W - B radiation (Kiesecker and Blaustein 1995). In 2001, these authors and collaborators reported an interesting synergistic effect between global climatic change, decrease in local precipitation, pond depth, larval exposure to W-B radiation and mortality of amphibian embryos due to infection by S. ferax (Kiesecker et al. 2001). Their study took place in Oregon's North Cascade Mountains where they monitored mortality of embryos of Bufo boreas in relation to S. ferax, UV-B exposure and different pond depths in field and laboratory experiments. To understand the effect of global climate, they analysed local precipitation during the winter months (developmental time for B. boreas) in relation to corresponding summer indices of El Nifio Southern Oscillation Events (ENSO), which are

AMPHIBUN BIOLOGY

3276

known to influence precipitation patterns in the Pacific Northwest (Kiesecker et al. 2001b). It was found that (1) mortality of embryos due to S. ferax was significantly correlated with decreased pond depth, (2) pond depth was related to winter precipitation (3) local winter precipitation decreased with the intensity of ENS0 events and (4) exposure to UV-B radiation significantly decreased hatching success of B. boreas in shallow (<20 cm) ponds as compared to those reared in deeper (50-100 cm) water. Thus, these authors showed how a global climatic phenomenon can cause local reductions in precipitation that result in the deterioration of an amphibian oviposition habitat. The effect is much more complicated, however, because shallow ponds increase exposure of eggs to UV-B radiation and hence increase susceptibility of embryos to disease (Kiesecker and Blaustein 1995; Kiesecker et al. 2001b). Although the projected amplitude and frequency of ENS0 events for the 21st century are uncertain, it is evident that strong El Nifio events can affect local precipitation in many places in the world (Meehl et al. 2007). The work of Kiesecker et al. (2001b) is an example of the complex synergies that can occur between global climatic change, local climatic fluctuations and disease.

V CONCLUSIONS The prognostics for global climatic change in the 21st century are not favourable for amphibians. The latest report of the Intergovernmental Panel for Climate Change (IPCC) states that "continued greenhouse gas emissions at or above current rates will cause further warming and induce many changes in the global climate system during the 21st century that would very likely be larger than those observed during the 20th century" (Meehl et al. 2007). Studies highlighted in the present chapter show how the proximal causes of climatic change can affect amphibian biology, ecology and vulnerability to disease, thereby driving population declines and extinctions of some species (Fig. 2). Under the striking evidence of direct and indirect effects of global warming, herpetologists face the challenge of conserving what is left of amphibian biodiversity. This will require rigorous scientific research

-.. bal Clin?ateCh:ange

-

Ultim Amp! rsiologica11 limitatio lationldet~ydration

Earlier Springs /

Disrupition of Breeding

\

Chanc

Fig. 2. Relationship between proximal causes of global climatic change and ultimate causes of amphibian declines around the world.

3278

AMPHIBIAN BIOLOGY

Daszak, E!, Berger, L., Cunningham, A. A,, Hyatt, A. D., Green, D. E. and Speare, R., 1999. Emerging infectious diseases and amphibian population declines. Emerg. Infect. Dis. 5: 73548. Daszak, F,! Strieby, A., Cunningham, A. A., Longcore, J. E., Brown, C. C. and Porter, D., 2004. Experimental evidence that the bullfrog (Rana catesbeiana) is a potential carrier of chytridiomycosis, an emerging fungal disease of amphibians. Hmpetol. J. 14: 201-207. Davidson, C., Shaffer, H. B. and Jennings, M. R., 2001. Declines of the California red-legged frog: climate, UV-B, habitat, and pesticides hypotheses. Ecol. Applic. 11: 464-479. Donnelly, M. and Crump, M., 1998. Potential effects of climate change on two neotropical amphibian assemblages. Climatic Change 39: 54 1-56 1. Duellman, W. E., 1999. "Patterns of Distribution of Amphibians". McGraw-Hill, New York. Duellman, W. and Trueb, L., 1994. "Biology of Amphibians". The Johns Hopkins University Press, Baltimore. Gibbs, J. E! and Breisch, A. R., 2001. Climate warming and calling phenology of frogs near Ithaca, New York, 1900-1999. Cons. Biol. 15: 1175-1 178. Harvell, C. D., Mitchell, C. E., Ward, J. R., Altizer, S., Doobson, A. E, Ostfeld, R. S. and Samuel, M. D., 2002. Climate warming and disease risks for terrestrial and marine biota. Sci 296: 2 158-2162. Heatwole, H., 1960. Burrowing ability ad behavioral responses to desiccation of the salamander, PIethodon cinerew. Ecol. 41: 661-668. Heyer, W. R., Rand, A. S., Goncalves da Cruz, C. A. and Peixoto, 0. L., 1988. Decimations, extinctions, and colonizations of frog populations in southeast Brazil and their e v ~ l u t i o ~ ~ i m ~ l i c a t iBiotropica ons. 20: 230-235. Houghton, J. Z,Ding, Y., Griggs, D. J., Noguer, M., van der Linden, I? J., Dai, X., Maskell, K. and Johnson, C. A., 2001. "Climate Change 2001: The Scientific Basis". Thinl assessment Report of the Intergovenzmental 2001, "Climate Change 2001: The Scientific Basis". Third assessment Report of the Intergovernmental Panel on Climate Change. Cambridge University Press, Cambridge. Hutchinson, V H. and Duprk, JS., 1992. Thermoregulation, Pp. 206-249 in "Environmental Physiology of the Amphibian, ed by M. E. Feder and W. W. Burggren. Univesity of Chicago Press, Chicago. Ingram, G. J., 1990. The mystery of the disappearing frog. Wildl. Aust. 27: 6-7. Joglar, R. L., 1998. Los Coquies de Puerto Rico: Su Historia Natural y Conservaci6n. Editorial de la Universidad de Puerto Rico, San Juan. Kiesecker, J. M., 2003. Invasive species as a global problem. Pp. 113-126 in "Amphibian Conservation", ed by R. D. Semlitsch. Smithsonian Institution, Washington DC. Keisecker, J. M. and Blaustein, A. R., 1995. Synergism between UV-B radiation and a pathogen magnifies amphibian embryo mortality in nature. Procl. Nut. Acad. Sci. (USA) 92: 11049-1 1052. Kiesecker, J. M., Blaustein, A. R. and Miller, C. L., 2001a. Potential mechanisms underlying the displacement of native red-legged frogs by introduced bullfrogs. Ecol. 82: 1964-1970. Kiesecker, J. M., Blaustein, A. R. and Belden, L. K., 2001b. Complex causes of amphibian population declines. Nat. 410: 681-684.

Kiesecker, J. M., Blaustein, A. R. and Miller, C. L., 2 0 0 1 ~ . Transfer of a pathogen from fish to amphibians. Cons. Biol. 15: 1064-1070. Knapp, R. A and Mathews, K. R., 2000. Non-native fish introductions and the decline of the mountain yellow-legged frog from protected areas. Cons.. Biol. 14: 428438. Kriger, K. M. and Hero, J.-M., 2006. Large-scale seasonal variation in the prevalence and severity of chytridiomycosis.J. 2001. 2006: 1-8.

La Marca, E., Lips, K. R., Etters, S., Puschendorf, R., Ibhiiez, R., Rueda-Almonacid, J. V, Rainer Schulte, C. M., Castro, F., Manzanilla-Puppo,J., Garcia-Pkrez, J. E., Bolaiios, F., Chaves, G., Pounds, J. A., Toral, E. and Young, B. E., 2005. Catastrophic population declines and extinctions in Neotropical harlequin frogs (Bufonidae: Atelopw). Biotropica 37: 190-201. Laurance, W. F., McDonald, K. R. and Speare, R., 1996. Epidemic disease and the catastrophic decline of Australian rain forest frogs. Cons. Biol. 10: 408-413. Lips, K. R., Brem, F., Brenes, R., Reeve, J. D., Alford, R. A., Voyles, J., Carey, C., Livo, L., Pessier, A. F! and Collins, J. E!, 2006. Emerging infectious disease and the loss of biodiversity in a Neotropical amphibian community. PNAS 103: 3165-3170. Longcore, J. E., Pessier, A. E! and Nichols, D. K., 1999. Batrachochvtrium dendrobatidis en. Et SD. Nov.. a a chytrid pithogenic to amphibians. ~ y ' c o l o ~ i91: 219-227. Longo, A. V, Burrowes, E! A. and Joglar, R. L., 2006. Study of the effect of drought on the dispersion pattern of rainforest Eleutherodactylus. Focus I v 2 2005: 64. Published Abstract for the 20th AAAS conference UIA, Bayam6n. Mazzoni, R., Cunningham, A. A,, Daszak, E!, Apolo, A,, Perdomo, E. and Speranza, G., 2003. Emerging pathogen of wild amphibians in frogs (Rana catesbeiana) farmed for International trade. Emerg. Infec. DL-. 9: 995-998. McCarty, J. l?, 2001. Ecological consequences of recent climate change. Cons. Biol. 15: 320-33 1. Meehl, G. A., Stocker, T F., Collins, W. D., Friedlingstein, E!, Gaye, A. T., Gregory, J. M., I t o h , A., Knutti, R., Murphy, J. M., Noda, A., Raper, S. C. B., Watterson, I. G., Weaver, A. J. and Zhao, 2.C., 2007. Global climate projections. Pp. 747-845 in "Climate Change 2007: The Physical Science Basis. Contribution of Working Group I to the Fourth Assessment Report of the Intergovernmental Panel on Climate Change", ed by S. Solomon, D. Qin, M. Manning, 2. Chen, M. Marquis, K. B. Averyt, M. Tignor and H. L. Miller. Cambridge University Press, Cambridge. Mendelson, J . R. III., Lips, K. R., Gagliardo, R. W., Rabb, G. B., Collins, J. I?, Diffendorfer, J. E., Daszak, E!, Ibhiiez, R. D., Zippel, K. C., Lawson, D. E!, Wright, K. M., Stuart, S. N., Gascon, C., da Silva, H. R., Burrowes, E! A., Joglar, R. L., La Marca, E., Lotters, S., du Preez, L. H., Weldon, C., Hyatt, A,, Vicente Rodriguez-Mahecha, J. V., Hunt, S., Robertson, H., Lock, B., Raxworthy, C. J., Frost, D. R., Lacy, R. C., Alford, R. A, Campbell, J. A, ParraOlea, G., Bolafios, F., Calvo Domingo, J. J., Halliday, T, Murphy, J. B., Wake, M. H., Coloma, L. A., Kuzmin, S. L., Price, M. S., Howell, K. M., Lau, M., Pethiyagoda, R., Boone, M., Lannoo, M. J., Blaustein, A. R., Dobson, A., Griffiths, R. A., Crump, M. L., Wake, D. B. and Brodie, E. D. Jr., 2006. Confronting amphibian declines and extinctions. Sci. 313: 48.

BURROWES: CLIMATIC CHANGE AND AMPHIBIAN DECLINES Merino-Viteri, A. R., 2001. Anilisis de las posibles causas de las disminuciones de las poblaciones de anfibios en 10s Andes de Ecuador. Thesis, Pontificia Universidad Catdlica del Ecuador, Quito, Ecuador. Merino-Viteri, A. R., Coloma, L. A. and Almendariz, A,, 2005. Los Telmatobius de 10s Andes de Ecuador y su disminucidn poblacional. Monogrufuzs de Herpetologia 7: 9-37. Parmesan, C., 2006. Ecological and evolutionary responses to recent climate change. Annu. Rm. Ecol. Evol. Syst. 37: 637-669. Parmesan, C. and Yohe, G., 2003. A globally coherent fingerprint of climate change impacts across natural systems. Nut. 421: 37-42. Parra-Olea, G., Martinez-Meyer, E. and Perez-Ponce de Leon, G., 2005. Forecasting climate change effects on salamander distribution in the highlands of central Mexico. Biotropica 37: 202-208. Plotrowski, J. S., Annis, S. L. and Longcore, J. E., 2004. Physiology of Batrachochytrium dendrobatzdis, a chytrid pathogen of amphibians. Mycologia 96: 9-15. Pough, F. H., Taigen, T., Stewart, M. and Brussard, l?, 1983. Behavioral modification of evaporative water loss by a Puerto &can fmg. Ecol. 64: 244-252. Pounds, J. A., 2001. Climate and amphibian declines. Nut. 410: 639-640.

3279

Ron, S. R., Coloma, L. A., Merino, A., Guayasamin, J. M. and Bustamante, M., 2001-2006. Declinaciones de anfibios en el Ecuador. [on line] Museo de Zoologia, Pontificia Universidad Cat6lica del Ecuador, Quito, Ecuador. <www.puce.edu.ed Zoologia/declinac.html> 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

--

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