Proc. Natl. Acad. Sci. USA
Vol. 95, pp. 9031–9036, July 1998
Population Biology
Chytridiomycosis causes amphibian mortality associated with
population declines in the rain forests of Australia and
Central America
LEE BERGERa,b,c, RICK SPEAREa, PETER DASZAKd, D. EARL GREENe, A NDREW A. CUNNINGHAMf, C. L OUISE GOGGINg,
RON SLOCOMBEh, MARK A. RAGANi, A LEX D. HYATTb, KEITH R. MCDONALDj, HARRY B. HINESk, KAREN R. LIPSl,
GERRY MARANTELLIm, AND HELEN PARKESb
aSchool of Public Health and Tropical Medicine, James Cook University, Townsville, Queensland 4811, Australia; bAustralian Animal Health Laboratory,
Commonwealth Scientific and Industrial Research Organization, Ryrie Street, Geelong, Victoria 3220, Australia; dSchool of Life Sciences, Kingston University,
Kingston-upon-Thames, Surrey KT1 2EE, United Kingdom; eMaryland Animal Health Laboratory, College Park, MD 20740; fInstitute of Zoology, Zoological
Society of London, Regent’s Park, London NW1 4RY, United Kingdom; gCommonwealth Scientific and Industrial Research Organization, Marine Research,
Hobart, Tasmania 7001, Australia; hVeterinary Clinical Centre, University of Melbourne, Werribee, Victoria 3030, Australia; iCanadian Institute for Advanced
Research, Program in Evolutionary Biology, National Research Council of Canada, Halifax, NS Canada B3H 3Z1; jConservation Strategy Branch, Queensland
Department of Environment, Atherton, Queensland 4883, Australia; kConservation Resource Unit, Queensland Department of Environment, Moggill, Queensland
4070, Australia; lDepartment of Zoology, Southern Illinois University, Carbondale, IL 62901-6501; and mAmphibian Research Centre, 15 Suvla Grove, Nth
Coburg, Victoria 3058, Australia
Edited by Robert May, University of Oxford, Oxford, United Kingdom, and approved May 18, 1998 (received for review March 9, 1998)
primary degraders or saprobes, using substrates such as chitin,
plant detritus, and keratin. Some genera are facultative or
obligate anaerobes, and many are obligate parasites of fungi,
algae, vascular plants, rotifers, nematodes, or insects. The
chytrid reported here is the first member of the phylum
Chytridiomycota to be recognized as a parasite of the phylum
Vertebrata (18). A similar discovery of a chytrid fungus in
dying captive dendrobatids in the United States made independently and contemporaneously (19) demonstrates that
chytridiomycosis is widespread in amphibians in the Americas
as well as Australia.
ABSTRACT
Epidermal changes caused by a chytridiomycete fungus (Chytridiomycota; Chytridiales) were found in
sick and dead adult anurans collected from montane rain
forests in Queensland (Australia) and Panama during mass
mortality events associated with significant population declines. We also have found this new disease associated with
morbidity and mortality in wild and captive anurans from
additional locations in Australia and Central America. This is
the first report of parasitism of a vertebrate by a member of
the phylum Chytridiomycota. Experimental data support the
conclusion that cutaneous chytridiomycosis is a fatal disease
of anurans, and we hypothesize that it is the proximate cause
of these recent amphibian declines.
MATERIALS AND METHODS
Collection of Specimens. Large numbers of ill and dead
anurans were found during monitoring programs of amphibian
populations in decline in Big Tableland, Queensland, Australia
(1993–1994) (3 species) (7) and in the Fortuna Forest Reserve,
western Panama (1996–1997) (10 species) (K.R.L., unpublished data). These were the last sightings of many of the
affected species at these locations. Apparently healthy adults
and tadpoles of Taudactylus acutirostris were collected from
Big Tableland in late September 1993 for captive breeding;
however, none survived. Dead or morbid wild and captive
anurans from additional locations in Australia and Central
America also were collected. (Table 1). The captive anurans
included rain forest frogs, Mixophyes fasciolatus, and introduced toads, Bufo marinus, which had been bred in captivity in
Australia and held in large, open collections that experienced
high mortality (.90% of the M. fasciolatus) at 2–3 weeks
postmetamorphosis. Retrospective histological examinations
of the skin of formalin-fixed, ethanol-stored museum specimens were performed on riparian anurans (n 5 28; 7 genera,
13 species) that had been collected during amphibian surveys
in 1987–1995 in three protected sites in the montane regions
of south-central Costa Rica (Las Tablas and Las Cruces,
Puntarenas Province) and western Panama (Fortuna Forest
Reserve), 1–10 years before population declines at those sites.
Toeclips collected into formalin during survey work on healthy
Litoria spenceri from southeastern Australia (Kosciuszko Na-
Amphibian population declines in protected habitats are a
serious global concern (1–3), and although several etiologies
have been proposed, none has been substantiated (4–6).
Disease has been hypothesized to be a likely cause, precipitated by an introduction of a pathogen into a naı̈ve population
(7, 8) or secondary to stress (6), but this paper presents
evidence of a specific pathogen as a potential cause of amphibian declines. Previous investigations of sudden declines
where adult mortality was suspected retrospectively (9, 10), or
observed (6, 11, 12), have not included rigorous diagnostic
examinations of large numbers of animals.
Montane riparian amphibian populations have declined
markedly in Queensland (Australia) and in Central America
with no evidence of environmental causes (13, 14). In these
locations, documented declines have appeared to progress
temporally and geographically consistent with an epidemic,
and, in monitored areas, mass mortalities of adult anurans
have been detected (refs. 7, 8, 14, and 15; K.R.L., unpublished
data). Tadpoles were seen at these sites after adults had
disappeared (ref. 7; K.R.L., unpublished data). Diagnostic
investigations to detect infectious and noninfectious diseases
were performed on the dead amphibians, and our results
provide strong evidence that cutaneous chytridiomycosis was
the cause of these mortalities.
Members of the phylum Chytridiomycota are heterotrophic
fungi that are ubiquitous and cosmopolitan (16, 17). They are
found primarily in soil and water where they usually act as
This paper was submitted directly (Track II) to the Proceedings office.
Abbreviation: ssu-rDNA, small-subunit rRNA genes.
Data deposition: The sequence reported in this paper has been
deposited in the GenBank database (accession no. AF051932).
cTo whom reprint requests should be addressed at: Commonwealth
Scientific and Industrial Research Organization, Australian Animal
Health Laboratory, P.O. Bag 24, Geelong, VIC 3220, Australia.
e-mail: Lee.Berger@dah.csiro.au.
The publication costs of this article were defrayed in part by page charge
payment. This article must therefore be hereby marked ‘‘advertisement’’ in
accordance with 18 U.S.C. §1734 solely to indicate this fact.
© 1998 by The National Academy of Sciences 0027-8424y98y959031-6$2.00y0
PNAS is available online at http:yywww.pnas.org.
9031
9032
Population Biology: Berger et al.
Table 1.
Amphibian deaths and morbidity associated with chytridiomycosis
Species
Location
Taudactylus acutirostris
Big Tableland, Australia,
15°429S, 145°169E, and
Captive Townsville,
Australia
Big Tableland, Australia,
15°429S, 145°169E
Big Tableland, Australia,
15°429S, 145°169E
Eungella National Park,
21°049S, 148°389E
Mary River, Australia,
26°379S, 152°419E
Goomburra, Australia,
27°589S, 152°209E
Brisbane, Australia,
27°389S, 152°239E,
Casino, Australia,
28°509S, 153°029E; and
Brisbane, Australia,
27°379S, 152°389E
Adelaide, Australia,
34°469S, 138°329E
Cunningham’s Gap,
Australia, 28°039S,
152°229E
Captive Geelong,
Australia
Captive Melbourne,
Australia
Central Highlands,
Australia, 37°229S,
146°029E
Captive, Cerro Pando,
Panama
Fortuna, Panama,
8°439N, 82°149W
Fortuna, Panama,
8°439N, 82°149W
Fortuna, Panama,
8°439N, 82°149W
Fortuna, Panama,
8°439N, 82°149W
Fortuna, Panama,
8°439N, 82°149W
Fortuna, Panama,
8°439N, 82°149W
Litoria rheocola
Litoria nannotis
Taudactylus eungellensis
Litoria lesueuri
Limnodynastes dumerilii
Litoria caerulea
Limno dynastes
tasmaniensis
Mixophyes fleayi
Bufo marinus
Mixophyes fasciolatus
Litoria spenceri
Atelopus chiriquiensis
Atelopus varius
Bufo haematiticus
Cochranella prosoblepon
Cochranella
albomaculata
Eleutherodactylus
emcelae
Eleutherodactylus
cruentus
Proc. Natl. Acad. Sci. USA 95 (1998)
Date of death
No. with chytridiomycosisy
no. examined
November 1993–January 1994
2y2
October 1993
5y7*
October–November 1993
1y2*
October–November 1993
2y4*
October 1995
1y1
May–June 1996
5y5
December 1996
1y1
July–September 1996
7y8
July 1996
3y4
June–July 1997
5y5
May–June 1996
4y4
August–October 1996
4y5
June–August 1996
18y18
December 1996–February 1997
35y35
February 1998
4y4
February 1994
4y5
January 1997
3y3
January 1997
2y2
January 1997
2y2
January 1997
2y2
January 1997
8y8
January 1997
2y2
*Histological examination of the skin was limited to the dorsum.
tional Park) also were examined histologically for chytrids.
Toeclips from 42 frogs had been collected during the 2 years
before any observations of unusual mortality.
Pathology and Microbiology. Frogs were fixed whole in
neutral buffered 10% formalin after opening the body cavity,
or necropsied in a sterile manner when fresh or after being
stored frozen. Skin scrapings were collected from fresh specimens by using sterile, disposable scalpel blades and examined
unstained with a light microscope or preserved in 100%
ethanol for DNA analysis. Giemsa-stained blood smears were
prepared and examined from 24 frogs received live. Formalinfixed tissues were paraffin-embedded, sectioned at 6-mm thick,
and stained with hematoxylin and eosin by using routine
methods for histology. Samples were collected for bacteriology
and mycology by using alginate swabs and immediately plated
onto horse blood agar, chocolate agar, MacConkey agar,
thiosulfate-citrate-bile-sucrose (TCBS) agar, and Sabauraud’s
agar. Bacterial isolates were identified by Gram stain, Mi-
crobact 24 NE (Medvet Science, Adelaide, Australia), and API
20E (BioMerieux, Charbonnier les Bains, France) systems. For
virological testing, frozen samples of liver, lung, kidney, muscle, gonad, and brain were macerated, passed through a
0.45-mm filter, and adsorbed to a monolayer of fat head
minnow [FHM; American Type Culture Collection (ATCC),
Manassas, VA; CCL 442), African clawed toad kidney (A6;
ATCC CCL 102), bullfrog tongue (FT; ATCC CCL 41), or
bluegill fry (BF-2; ATCC CCL 91) cells for Australian samples,
and rainbow trout gonad (RTG-2) (20), chinook salmon
embryo (CHSE) (21), Rana pipiens liver (RPL; KS Tweedell,
University of Notre Dame, IN), and Terrapene carolina heart
(TH; ATCC CCL 50) cells for Panamanian samples. Cells were
observed for two passages of at least 7 days each.
Electron Microscopy. For scanning electron microscopy,
skin was fixed in 2.5% glutaraldehyde, postfixed in 1% osmium
tetroxide, dehydrated, critical point-dried, sputter-coated with
gold, and examined by using a JEOL JSM 840 scanning
Population Biology: Berger et al.
FIG. 1. Histological section of severely infected digital skin of a wild
frog, Litoria caerulea, from Queensland, Australia. The stratum corneum
is markedly thickened because of a massive infection by a chytrid parasite.
Thickness of normal stratum corneum is 2–5 mm, but here it is about 60
mm. This section contains a mass of intracellular sporangia (S) and
developing sporangia. The mature sporangia are 12–20 mm (n 5 25) in
diameter, have refractile walls (0.5- to 2.0-mm thick), and contain
zoospores. Many sporangia have discharged all zoospores. Zoospores are
released through discharge tubes (D). Note the absence of an inflammatory cell reaction in the dermis and epidermis. (Bar 5 30 mm.)
electron microscope at 5 kV. For transmission electron microscopy (TEM), skin was fixed in neutral-buffered 10%
formalin, postfixed in 2.5% glutaraldehyde then 1% osmium
tetroxide, stained en-bloc with 2% uranyl acetate, dehydrated,
embedded in Araldite epoxy resin, sectioned at 70 nm (serial
sectioning was not performed), stained with Reynold’s lead
citrate, and examined on either a Phillips 301 or a Hitachi 600
TEM. In addition, internal organs were retrieved from paraffin
blocks, dewaxed, and prepared for TEM.
Proc. Natl. Acad. Sci. USA 95 (1998)
9033
DNA Sequencing of Chytrid. DNA was extracted from
ethanol-fixed skin scrapings from two wild frogs, Litoria
caerulea, from Queensland that were naturally diseased with
cutaneous chytridiomycosis. Ethanol was removed and tissues
were crushed in extraction buffer (50 mM Tris, pH 8.0y0.7 M
NaCly10 mM EDTAy1% hexadecyltrimethylammonium bromidey0.1% 2-mercaptoethanol) before incubation at 65°C for
1 hr with 100 mgyml proteinase-k added. DNA was extracted
by using phenolychloroform and precipitated in ethanol. The
nuclear gene encoding small-subunit ribosomal RNA (ssurDNA) ('1,800 bp) was amplified by using universal primers
A (59-CCAACCTGGTTGATCCTGCCAGT-39) and B (59GATCCTTCTGCAGGTTCACCTAC-39) modified from
Medlin et al (22). DNA was purified by using QiaQuick spin
columns (Qiagen, Chatsworth, CA) following the protocol
recommended by the manufacturer. Products were sequenced
by using the dye terminator sequencing reaction (Perkin–
Elmer) and run on an acrylamide gel on an automated ABI
Prism 377 DNA sequencer (Applied Biosystems). A total of
1,726 nt (GenBank accession no. AF051932) were collected
and aligned with the ssu-rDNA of 44 other eukaryotes (23).
Ambiguously aligned regions were removed to yield a 45 3
1,607 matrix that corresponds closely to matrix B (23). Neighbor-joining, parsimony, and maximum likelihood trees were
inferred by using PHYLIP 3.53 (24). All inferences used randomaddition order; parsimony and likelihood analyses used global
(43-level) optimization, and neighbor-joining and parsimony
analyses were bootstrapped (n 5 1,000).
Transmission Experiment. Experimental transmission of
cutaneous chytridiomycosis was conducted by using captivebred sibling frogs, M. fasciolatus, which were housed individually at 24°C in plastic tubs with gravel and 800 ml aged tap
water. Before experimental infection, 10 frogs were tested for
the presence of chytrids by histological examination of clipped
toes and all were negative for the fungus. For the experiment
FIG. 2. Scanning electron micrographs of infected digital skin of a wild frog, Litoria lesueuri, from Queensland, Australia, which died with
cutaneous chytridiomycosis. (A) A cluster of mature sporangia are evident within the cells of the epidermis. The discharge tubes of the sporangia
are visible projecting outwards from the cell surface. The plug of material within the discharge tube has been released in one of the sporangia
(arrowhead). (Bar 5 10 mm.) (B) Most of the cells in this field of the superficial layer of the epidermis contain sporangia, as evident by the bulging
surfaces and the protrusion of unopened discharge tubes through infected cells (arrowheads). (Bar 5 10 mm.)
9034
Population Biology: Berger et al.
Proc. Natl. Acad. Sci. USA 95 (1998)
FIG. 3. Transmission electron micrographs of the zoospores found within fungal sporangia in the epidermis of naturally infected amphibians from
Panama (Eleutherodactylus emcelae, A and C) and from Australia (Litoria caerulea, B and D). (A) Longitudinal section through a zoospore. The ribosomal
area (Rb) is surrounded by a single cisterna of endoplasmic reticulum and bounded by a nucleus (N), mitochondria (Mi), and a microbody–lipid globule
complex (asterisk). A single flagellum (Fl) connects to the posterior of the zoospore. (Bar 5 1 mm.) (B) A transverse–oblique section through the anterior
aspect of a zoospore demonstrates the unusual microbody–lipid globule complex. The microbody (asterisk) lies adjacent to four lipid globules (L) in this
section. There is no evidence of a cisterna bounding these lipid globules. Rb, ribosomal area. (Bar 5 1 mm.) (C) Transverse section through the anterior
aspect of a zoospore. Note the discoidal cristae within mitochondria (Mi). The ribosomal area is bounded by the microbody–lipid globule complex (asterisk)
and the mitochondria. No cisterna bound the lipid globules in this section. (Bar 5 0.5 mm.) (D) A longitudinal–oblique section through a zoospore in
the epidermis of an Australian amphibian for comparison with A. (Bar 5 0.75 mm.)
14 frogs (6 exposed, 8 controls) from the previously sampled
cohort were used. Each of six frogs was exposed to 800 ml of
an aqueous suspension of approximately 3 3 103 sporangia in
fresh skin scrapings taken from a dead frog of the same species
that had naturally acquired cutaneous chytridiomycosis. Four
frogs were exposed to water that had contained infected skin
scrapings and had been passed through a 0.45-mm filter, and
four were kept as untreated controls.
RESULTS
At necropsy, no consistent gross lesions were found in the sick
and dying anurans. However, areas of abnormal epidermal
sloughing were seen in a number of amphibians from both
Australia and Central America. Examinations of fresh, unstained wet mounts of this dysecdic skin consistently revealed
large numbers of spherical to subspherical sporangia of a
nonhyphal, previously unreported fungus. Histologically, significant abnormalities were restricted to the skin, in which
sporangia were found in the stratum corneum and stratum
granulosum (Fig. 1) principally of the digits and ventral body,
especially the hypervascularized pelvic patch (‘‘drink patch’’).
Epidermal changes associated with these parasites consisted of
irregular cell loss, erosions, and segments of marked thickening of the stratum corneum (parakeratotic hyperkeratosis).
Deeper epidermal changes consisted of moderate hyperplasia
of the stratum intermedium (acanthosis), but there were negligible inflammatory cell infiltrates. Sporangia were also seen
on histological examination of the keratinized mouth parts of
wild Panamian tadpoles (three of seven) (three genera) and in
killed, otherwise-healthy, captive M. fasciolatus (three of four)
and B. marinus (three of eight) tadpoles from the spawning
groups that had high mortality rates after metamorphosis. In
these tadpoles, the fungus was found only in the mouth parts
and not in the unkeratinized skin of the body and tail.
There was no evidence of epidermal viral infection in adult
anurans based on histological, ultrastructural, and culture
findings, including culture attempts from 23 wild and captive
frogs originating from Big Tableland (three species) and 26
wild anurans from Fortuna (8 species). A range of bacteria was
isolated from these specimens, including Aeromonas hydrophila from the Australian frogs and Flavobacterium indologenes from the Panamanian amphibians. No species of bacteria
was isolated from more than 44% of sick and dead amphibians
in any wild group, and the histological findings were not
consistent with primary bacterial disease. Aeromonas spp.
were not isolated from any of 26 frozen anurans collected from
the mass mortality incident in Panama. Infectious organisms
were not detected on light microscopic examination of Giemsastained blood smears from 24 diseased Australian frogs received alive. No epidermal myxozoa or protozoa were detected
histologically or ultrastructurally in tissues from either sick or
dead amphibians. Mycological cultures of frozen tissue failed
to detect chytrids.
Chytrid fungi were not found on histologic examination of
the 28 archived Panamanian and Costa Rican specimens
collected from the wild before population declines, or the 42
archived toeclips from healthy Australian frogs.
Scanning electron microscopical examination of the surface
of digital skin from infected Australian frogs showed marked
roughening and sloughing of the skin surface compared with
the smooth and intact appearance of uninfected epidermis
from the control frogs. Infection of the superficial keratinized
cells was evident by the presence of prominent discharge tubes
of the fungus protruding through the bulging cells (Fig. 2).
Transmission electron microscopical examination of wild and
captive Australian and wild Panamanian anurans identified
this parasite as a chytrid fungus (Chytridiomycota; Chytridiales) primarily based on zoospore ultrastructure (17). The
Population Biology: Berger et al.
FIG. 4. A composite line drawing of a longitudinal section through the
zoospore of the chytridialean parasite found in the epidermis of Australian and Panamanian amphibians. The drawing is based on examination
of thin sections of zoospores within the sporangium, not of serially
sectioned, cultured, glutaraldehyde-fixed specimens. For these reasons,
some details of the kinetosomal region were poorly preserved or may not
have been visible. The unusual microbody–lipid globule complex and
some details of the flagellar attachment are shown. In some sections, it
appears that two microbody–lipid globule complexes are present, but it is
unknown whether this represents a single complex wrapped around the
ribosomal region. The flagellum in longitudinal and transverse section is
drawn slightly larger than scale to adequately demonstrate the aspects of
the flagellar attachment. Rb, ribosomal area; N, nucleus; L, lipid globule;
m, microbody; mi, mitochondrion; k, kinetosome; p, prop attaching
kinetosome to plasmalemma; tp, terminal plate; nfc, nonfunctioning
centriole; fl, flagellum. (Bar 5 1 mm.)
fungus in the epidermis of the anurans had a thallus bearing
a network of rhizoids and smooth-walled, spherical to subspherical, inoperculate sporangia. Each sporangium produced
a single discharge tube. Zoospores (Fig. 3) had an elongate–
ovoidal body and a single, posterior flagellum and possessed a
core area of ribosomes often with membrane-bound spheres of
ribosomes within the main ribosomal mass. A small spur was
located at the posterior of the cell body, adjacent to the
flagellum. It is unknown whether this represents an artifact in
these formalin-fixed specimens. The core area of ribosomes
was surrounded by a single cisterna of endoplasmic reticulum,
two to three mitochondria, and an extensive microbody–lipid
globule complex. The microbody closely apposed and almost
surrounded four to six lipid globules (three anterior and one to
three laterally), some of which appeared bound by a cisterna.
Some zoospores appeared to contain more lipid globules;
however, this may have been a result of a plane-of-sectioning
effect, because the globules were often lobed in the zoospores
examined (Fig. 4). A rumposome was not observed. A nonfunctioning centriole lay adjacent to the kinetosome. Nine
interconnected props attached the kinetosome to the plasmalemma, and a terminal plate was present in the transitional
zone (Fig. 4). An inner ring-like structure attached to the
tubules of the flagellar doublets within the transitional zone
was observed in transverse section. No roots associated with
the kinetosome were observed. In many zoospores, the nucleus
lay partially within the aggregation of ribosomes and was
invariably situated laterally. Small vacuoles and a Golgi body
with stacked cisternae occurred within the cytoplasm outside
the ribosomal area. Mitochondria, which often contained a
Proc. Natl. Acad. Sci. USA 95 (1998)
9035
FIG. 5. Maximum-likelihood tree inferred from ssu-rDNA sequences
of the amphibian parasite and 44 other eukaryotes, based on 1,607 stably
aligned positions at a transitionytransversion ratio of 2.0. GenBank
accession numbers of ssu-rDNAs other than the amphibian parasite in
this tree have been published (23). Bootstrap values (percent of 1,000
replicates) are taken from the topologically identical parsimony tree.
Distance matrices (corrected for superimposed substitutions by a generalized Kimura two-parameter model) were calculated by using DNADIST
from the PHYLIP package, and neighbor-joining trees were inferred by
using NEIGHBOR (24). Unweighted parsimony trees were inferred by using
DNAPARS with 1,000 iterations (jumbles). Neighbor-joining and parsimony
analyses were bootstrapped (n 5 1000) by sequential use of SEQBOOT, the
inference program(s) above, and CONSENSE (24, 25). A maximumlikelihood tree was inferred by using DNAML. All inferences used randomaddition order and, for parsimony and likelihood, global (43-level)
optimization. The position of Apusomonas proboscidea ssu-rDNA was
unstable, and in likelihood inference depended on the transitiony
transversion ratio.
small number of ribosomes, were densely staining with discoidal cristae.
DNA analyses showed that the amphibian parasite ssurDNA groups were among those of the chytrid fungi, with the
closest match to Chytridium confervae (Fig. 5).
Between 10 and 18 days postexperimental exposure, all six
frogs exposed to unfiltered skin scrapings became moribund;
one died and the rest were euthanized with 0.2% tricaine
methanesulfonate (Ruth Consolidated Industries, Annandale,
Australia). Cutaneous chytridiomycosis was diagnosed cytologically, histologically, and ultrastructurally. The eight control
frogs remained healthy and were free of chytrids when killed
after 22 days and examined histologically.
DISCUSSION
The extensive pathological investigations reported in this
paper provide evidence for a pathogen causing amphibian
mortality in areas where declines are marked and well documented. Furthermore, the similarity between pathological
findings from Australian and Central American amphibians is
remarkable, and it appears that a similar fungal pathogen is
present in these two widely separated regions.
9036
Population Biology: Berger et al.
The ultrastructural studies and DNA analyses firmly place
the epidermal fungus within the phylum Chytridiomycota,
class Chytridiomycetes, order Chytridiales (18, 26). It was not
possible to further classify the organism by using ultrastructural characters because cultured, glutaraldehyde-fixed specimens were not available and serial sectioning was not performed. However, the occurrence of several lipid globules in
the microbody–lipid globule complex with some bounded by a
cisterna, is unusual within the Chytridiales (27, 28). These and
other ultrastructural characters, together with parasitism of
the amphibian epidermis, suggest that this parasite represents
a new chytridialean genus. Despite extensive ultrastructural
examination of zoospores and other developmental stages, no
significant differences could be found between chytrids from
wild and captive Australian amphibians and chytrids from wild
Panamanian amphibians, suggesting that animals from both
continents were infected by the same fungus.
Chytrids were observed to infect tadpoles, but the localized
distribution in the only keratinized body region, the mouth
parts, may explain the lack of mortality. We assume that only
after metamorphosis, when the skin becomes keratinized (29),
does the chytrid cause a widespread, fatal epidermal infection.
The absence of infection in nonkeratinized epithelial surfaces
of tadpoles (body, limbs, tail, mouth, gills) and postmetamorphic anurans (conjunctiva, nasal cavity, mouth, tongue, intestines) emphasizes the strictly keratinophilic affinity of this
newly recognized pathogen.
Although the transmission experiment was a preliminary
trial that used unpurified material, it demonstrates that
chytrids are associated with a transmissible fatal disease of
anurans and supports our diagnostic findings that cutaneous
chytridiomycosis was the cause of the mortality events in wild
amphibians. Furthermore, the failure to identify alternative
causes of death after examination of the wild amphibians and
their environment (refs. 13 and 30; K.R.L., unpublished data)
supports our theory that cutaneous chytridiomycosis was the
cause of the riparian amphibian population declines in the
montane rain forests of Queensland and Panama.
The mechanism(s) by which cutaneous chytridiomycosis
becomes a fatal infection in postmetamorphic anurans is not
clear. The epidermal hyperplasia may seriously impair cutaneous respiration and osmoregulation, particularly as chytrids
consistently infect the pelvic patch, a major site of water
absorption in some anurans (31). Alternatively, death may be
a result of absorption of a toxic product released by the fungus.
An analysis of declining amphibian species at high altitudes
in eastern Australia (J.-M. Hero, unpublished data) found that
affected species have low clutch sizes and occupy restricted
geographic ranges. This may be consistent with the impact of
a disease affecting a range of amphibian species, but with
recovery of more robust populations. There are many examples of infectious disease affecting wildlife populations (32),
including examples where introduction of pathogens has decimated populations (33–35). There may be several explanations as to why chytridiomycosis has emerged as a disease of
frogs in Australia and Panama. The chytrid may be an introduced pathogen spreading through naı̈ve populations (7, 8), or
it may be a widespread organism that has emerged as a
pathogen to frogs because of either an increase in virulence or
an increased host susceptibility caused by other factors, such as
environmental changes or as yet undetected coinfections.
For collection of frogs and technical and other assistance we thank, in
Australia, G. Russell, M. Braun, T. Wise, F. Fillipi, T. Stephens, J.
Humphrey, P. Hooper, M. Williamson, G. Rowe, J. Muschialli, D.
Carlson, T. Chamberlain, S. Larkin, and S. Daglas (Australian Animal
Health Laboratory); K. Field and R. Ritallick (James Cook University);
J.-M. Hero (Griffith University); G. Gillespie (Arthur Rylah Institute); N.
Murphy (Commonwealth Scientific and Industrial Research Organization, Marine Research); J. Koehler (Townsville General Hospital); H.
Proc. Natl. Acad. Sci. USA 95 (1998)
McCracken (Royal Melbourne Zoo); M. Tyler, R. Short, and C. Williams
(University of Adelaide); L. Skerratt (University of Melbourne); D.
Charley (NSW, National Parks and Wildlife Service); R Natrass and B.
Dadds (Queensland Department of Environment); P. O’Donoghue (University of Queensland); M. Mahony (University of Newcastle); and
members of the public who found sick frogs. In the United States we
gratefully acknowledge many contributions of R. Hess, V. Beasley, B.
Jakstys, L. Miller, J. Cochran, J. Cheney, and B. Ujhelyi (University of
Illinois at Urbana–Champaign); G. Rabb and T. Meehan (Brookfield
Zoo, Chicago), R. Mast (Conservation International); and J. Savage
(CRE Collections). This work was supported by the Australian Nature
Conservation Agency, Biodiversity Australia, Wet Tropics Authority,
IUCNySSCyDeclining Amphibian Populations Task Force, the University of Illinois at Urbana–Champaign, and American Airlines.
1.
2.
3
4.
5.
6.
7.
8.
9.
10.
11.
12.
13.
14.
15.
16.
17.
18.
19.
20.
21.
22.
23.
24.
25.
26.
27.
28.
29.
30.
31.
32.
33.
34.
35.
Blaustein, A. R. & Wake, D. B. (1990) Trends Ecol. Evol. 5,
203–204.
Wake, D. B. (1991) Science 253, 860.
Drost, C. A. & Fellers, G. M. (1996) Conserv. Biol. 10, 414–425.
Blaustein, A. R., Hoffman, P. D., Hokit, D.G., Kiesecker, J. M.,
Walls, S. C. & Hays, J. B. (1994) Proc. Natl. Acad. Sci. USA 91,
1791–1795.
Kiesecker, J. M. & Blaustein, A. R. (1995) Proc. Natl. Acad. Sci.
USA 92, 11049- 11052.
Carey, C. (1993) Conserv. Biol. 7, 355–362.
Laurance, W. F., McDonald, K. R. & Speare, R. (1996) Conserv.
Biol. 10, 406–413.
Laurance, W. F., McDonald, K. R. & Speare, R. (1997) Conserv.
Biol. 11, 1030–1034.
Pounds, J. A. & Crump, M. L. (1994) Conserv. Biol. 8, 72–85.
Heyer, W. R., Rand, A. S., Goncalvez da Cruz, C. A. & Peixoto,
O. L. (1988) Biotropica 20, 230–235.
Bradford, D. F. (1991) J. Herpetol. 25, 174–177.
Sherman, C. K. & Morton, M. L. (1993) J. Herpetol. 27, 186–198.
Richards, S. R., McDonald, K. R. & Alford, R. A. (1993) Pac.
Conserv. Biol. 1, 66–77.
Lips, K. R. (1998) Conserv. Biol. 12, 106–117.
Trenerry, M. P., Laurance, W. F. & McDonald, K. R. (1994) Pac.
Conserv. Biol. 1, 150–153.
Sparrow, F. K. (1960) Aquatic Phycomycetes (The Univ. of
Michigan Press, Ann Arbor, MI), pp. 16–18.
Karling, J. S. (1977) Chytridiomycetarum Iconographia: An Illustrated and Brief Descriptive Guide to the Chytridomycetous Genera
with a Supplement of the Hyphochytriomycetes (Lubrecht and
Cramer, Monticello, NY).
Barr, D. J. S. (1990) in Handbook of Protoctista, eds. Margulis, L.,
Corliss, J. O., Melkonian, M. & Chapman, D. J. (Jones and
Bartlett, Boston), pp. 454–466.
Pessier, A. P., Nichols, D. K., Longcore, J. E. & Fuller, M. S.
(1998) J. Vet. Diag. Invest., in press.
Wolf, K. & Quimby, M. L. (1962) Science 135, 1065–1066.
Lannan, C. N., Winton, J. R. & Fryer, J. L. (1984) In Vitro 20,
671–676.
Medlin, L., Elwood, H. J., Stickel, S. & Sogin, M. L. (1988) Gene
71, 491–499.
Ragan, M. A., Goggin, C. L., Cawthorn, R. J., Cerenius, L.,
Jamieson, A. V. C., Plourde, S. M., Rand, T. G., Soderhall, K. &
Gutell, R. R. (1996) Proc. Natl. Acad. Sci. USA 93, 11907–11912.
Felsenstein, J. (1989) Cladistics 5, 164–166.
Felsenstein, J. (1985) Evolution 39, 783–791.
Barr, D. J. S. (1980) Can. J. Bot. 58, 2380–2394.
Powell, M. J. & Roychoudhury, S. (1992) Can. J. Bot. 70, 750–761.
Longcore, J. E. (1993) Can. J. Bot. 71, 414–425.
Warburg, M. R., Lewinson, D. & Rosenberg, M. (1994) in
Amphibian Biology, The Integument, eds. Heatwole, H. & Barthalmus, G. T. (Surrey Beatty & Sons, Chipping Norton, Australia), Vol. 1, pp. 33–63.
Laurance, W. F. (1996) Biol. Conserv. 77, 203–212.
Parsons, R. H. (1994) in Amphibian Biology, The Integument, eds.
Heatwole, H. & Barthalmus, G. T. (Surrey Beatty & Sons,
Chipping Norton, Australia), Vol. 1, pp. 132–146.
May, R. M. (1988) Conserv. Biol. 2, 28–30.
Hess, G. (1996) Ecology 77, 1617–1632.
Warner, R. E. (1968) The Condor 70, 101–120.
Leberg, P. L. & Vrijenhoek, R. C. (1994) Conserv. Biol. 8,
419–424.