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AMphibian Diseases
characteristics such as a septum and discharge tube (Fig. 2), and
hyperplasic epidermis.
It is worth mentioning that the population of P. fernandezae
used in this study has been studied by local herpetologists since
2001, but no moribund or dead specimens have been recorded
to date. Moreover, the individuals on which infection was detected had been engaged in reproductive activities.
Acknowledgments.—We are grateful to Josefina V. Candioti and
Marcelo L. Larramendy for use of their microscope and digital camera. We thank Allan P. Pessier for his critical review of the photos.
Literature Cited
Arellano, M. L., D. P. Ferraro, M. M. Steciow, and E. O. Lavilla. 2009.
Infection by the chytrid fungus Batrachochytrium dendrobatidis
in the yellow belly frog (Elachistocleis bicolor) from Argentina.
Herpetol. J. 19:217–220.
Barrio, A. 1964. Relaciones morfológicas, eto-ecológicas y zoogeográficas entre Physalaemus henseli (Peters) y P. fernandezae (Müller)
(Anura, Leptodactylidae). Acta Zool. Lilloana 20:285–305.
Barrionuevo, S., and S. Mangione. 2006. Chytridiomycosis in two species of Telmatobius (Anura: Leptodactylidae) from Argentina. Dis.
Aquat. Org. 73:171–174.
Berger, L., R. Speare, and A. Kent. 1999. Diagnosis of chytridiomycosis
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edu.au/school/phtm/PHTM/frogs/histo/chhisto.htm. November
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Drury, R. B., and E. A. Wallington. 1980. Carleton’s Histological Technique. Oxford University Press. 520 pp.
Fox, S., J. Yoshioka, M. E. Cuello, and C. Úbeda. 2006. First case of ranavirus-associated morbidity and mortality in natural populations
of the South American frog Atelognathus patagonicus. Dis. Aquat.
Org. 72:87–92.
Ghirardi, R., J. N. Lescano, M. S. Longo, G. Robledo, M. M. Steciow, and
M. G. Perotti. 2009. Batrachochytrium dendrobatidis in Argentina:
first record in Leptodactylus gracilis and another record in Leptodactylus ocellatus. Herpetol. Rev. 40:175–176.
Gutierrez, F. R., M. L. Arellano, L. E. Moreno, and G. S. Natale. 2010.
Batrachochytrium dendrobatidis in Argentina: First record of infection in Hypsiboas cordobae and Odontophrynus occidentalis
tadpoles, in San Luis province, Argentina. Herpetol. Rev. 41:323–
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Herrera, R., M.M. Steciow, and G.S. Natale. 2005. Chytrid fungus parasitizing the wild amphibian Leptodactylus ocellatus (Anura: Leptidactylidae) in Argentina. Dis. Aquat. Org. 64:247–252.
IUCN [International Union for the Conservation of Nature]. 2011.
IUCN Red List of Threatened Species. Available at: www.iucnredlist.org. Accessed 06 August 2011.
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August 2011.
Herpetological Review, 2012, 43(1), 85–87.
© 2012 by Society for the Study of Amphibians and Reptiles
Low Prevalence of Batrachochytrium dendrobatidis in Two
Plethodontid Salamanders from North Carolina, USA
Although the pathogenic fungus Batrachochytrium dendrobatidis (Bd) was first isolated from anuran amphibians, subsequent research has clearly shown that it also infects many species of caudate amphibians. Opportunistic sampling surveys
have shown that at least 56 species of salamanders from the
families Ambystomatidae, Amphiumidae, Salamandridae, Crytobranchidae, and Plethodontidae in the United States harbor
Bd (reviewed in Bryne et al. 2008; Olson 2010). The effect of Bd
on salamander demography, however, is less understood than
its impact on anuran populations. Laboratory challenge experiments have shown that salamanders can be infected by Bd and
show mortality from chytridiomycosis (Chinnadurai et al. 2009;
Vazquez et al. 2009; Weinstein 2009), but more sampling and
monitoring of salamanders will help elucidate potential ecological and/or climatic variables that may influence the susceptibility of salamanders in the wild. Here, we contribute information
on the prevalence and distribution of Bd in salamander populations by testing 166 individuals from two localities in North Carolina, USA. Our results expand upon previous reports of Bd infection in amphibian populations in North Carolina (e.g., Bryne
et al. 2009; Keitzer et al. 2011; Rothermel et al. 2008).
We opportunistically sampled two plethodontid salamander
species at two collection sites on 8–12 August 2010 (Fig. 1). At
Deep Gap, we collected salamanders by turning over rocks and
logs that were located up to 15 m from a stream. The collecting
site at Wayah Bald was a grassy area near the forest edge. We
swabbed each salamander 30 times using sterile swabs (Medical
wire No. 113) in the manner of Boyle et al. (2004) and Kriger et al.
(2006). Each animal was handled with a clean pair of latex gloves.
No animals showed any outward signs of disease. After swabbing, animals were retained for additional studies under permits
issued by the North Carolina Wildlife Resources Commission
and the U.S. Forest Service to LDH. Swabs were stored in 100%
ethanol and transported to Cornell University, where molecular
testing was performed by Kiemnec-Tyburczy.
Genomic DNA was extracted from the swabs using Prepman Ultra following the protocol of Boyle et al. (2004). The level
of Bd zoospore load was assessed using the method of Boyle et
al. (2004). Briefly, this method used Taqman quantitative PCR
to determine the total number of Bd zoospore genome equivalents in each unknown sample, based on known standards that
were run simultaneously. To maximize cost efficiency but retain
individual information, PCRs were run in singlicates at 1:10
Karen M. Kiemnec-Tyburczy*
Department of Ecology and Evolutionary Biology, Cornell University
E145 Corson Hall, Ithaca, New York 14853, USA
Sarah L. Eddy
Adam J. Chouinard
Lynne D. Houck
Department of Zoology, Oregon State University, 3029 Cordley Hall
Corvallis, Oregon 97331, USA
*Corresponding author; e-mail: kmk255@cornell.edu
Herpetological Review 43(1), 2012
86
AMphibian Diseases
interval) for each site using Clopper-Pearson binomial confidence intervals (Clopper and Pearson 1934).
We found that 9 of 166 (5.4%) animals tested positive for Bd.
Both species—Desmognathus ocoee and Plethodon shermani—
had Bd-positive individuals (Table 1). Plethodon shermani is a
direct-developing species that is commonly found under leaf
litter, logs, and rocks. In contrast, D. ocoee has an aquatic larval
stage and often is associated with seepages and streams (and
thus may be more likely to come in contact with Bd zoospores).
Despite these differences in life histories, both species had similar Bd prevalence levels. To our knowledge, this is the first published report of Bd infecting these two species, although Keitzer
et al. (2011) tested one P. shermani and 53 D. ocoee at Coweeta
Hydrologic Laboratory (Macon Co., North Carolina) but did not
detect Bd on any individuals. We note, however, that our sampling occurred slightly later in the year and may have occurred at
higher elevations. The elevation of our two localities, Deep Gap
and Wayah Bald—approximately 1300 m and 1630 m, respectively—are in the upper end of the elevation range of the Coweeta
area that was sampled (675–1592 m), and thus our localities may
have been cooler and perhaps more suitable for Bd.
The prevalence of Bd at Deep Gap was 0.050 (95% CI: 0.016–
0.11), whereas the prevalence at Wayah Bald was slightly higher at 0.061 (95% CI: 0.017–0.15). All of the zoospore loads we
quantified were very low. The highest load we calculated was
57 zoospores, and most of the positive samples contained 3–4
zoospores (range 1–57 zoospores). We did not observe any trend
towards particular life stages or sexes being infected within the
two species (Table 1), although the animal with the highest load
was a juvenile P. shermani from Wayah Bald. The low levels of
prevalence, combined the lack of clinical signs of disease, suggests that Bd exists primarily as a subclinical infection in D. ocoee
at Deep Gap and in P. shermani at Wayah Bald, at least during the
summer months.
Our data are consistent with other studies of plethodontid
populations that have shown that Bd prevalence in some populations is relatively low, but that Bd is present in North Carolina
(e.g., Keitzer et al. 2011; Rothermel et al. 2008). Bd also has been
found in other species of Plethodon and Desmognathus in other
areas of the USA. Prevalence in P. glutinosus and P. yonahlossee in Watauga County, North Carolina was 1/40 and 1/41, respectively (Chinnadurai et al. 2009). Desmognathus conanti
Fig 1. Specific location and topography of two sites in North Caroli(Timpe et al. 2008) and D. fuscus (Grant et al. 2008) have also
na, USA (inset; shaded state), where salamanders were tested for the
tested positive for Bd. In fact, Bd has been found on plethodonpresence of Batrachochytrium dendrobatidis (Bd). DG = Deep Gap,
tid species as diverse as Pseudotriton ruber (Montanucci 2009),
WB = Wayah Bald.
Batrachoseps attenuatus (Weinstein 2009), Bolitoglossa dofleini
(Pasmans et al. 2004) and Plethodon neomexicanus (Cummer
et al. 2005). In other salamander families, population surveys
Table 1. Batrachochytrium dendrobatidis (Bd) prevalence (number of
have reported varying estimates of prevalence. A survey of NoBd-positive animals/total sampled) in plethodontid salamanders in
tophthalmus viridescens found 16 of 63 individuals infected in
North Carolina, USA.
localities in Georgia, North Carolina, and Virginia (Rothermel
Prevalence
et al. 2008), whereas another found prevalence in N. viridescens
LocalitySpecies
♀
♂ JuvenileTotal
varied between 0–100% across multiple ponds in Pennsylvania
(Groner and Relyea 2010). Prevalence in ambystomatid salaDeep Gap
manders also varies widely; Ouellet et al. (2005) found 4/139
(35.0389°N, Desmognathus ocoee 1/23 1/31 2/16 4/70
Ambystoma maculatum infected across Canada while Padgett83.5431°W) Plethodon shermani 1/11 0/6
0/13
1/30
Flohr and Longcore (2005) found 2/11 A. californiense infected
in two ponds. These studies highlight the need for further work
Wayah Bald
investigating how disease-related mortality has impacted natu(35.1803°N, Desmognathus ocoee0/1 0/0 0/0 0/1
ral populations historically. Further testing across multiple sea83.5606°W) Plethodon shermani 1/26 2/14 1/25
4/65
sons and years is necessary to better understand Bd infection
dynamics in salamanders.
dilutions. Studies have shown that there is no statistical difference in sensitivity or specificity between the singlicate and triplicate assays (Kriger et al. 2006). Every swab sample was run with
(1) two negative controls, (2) an internal positive control (ABI
Cat. No. 4308323) to test for potential inhibition and contamination, and (3) a range of Bd standards corresponding to 0.1, 1, 10,
and 100 and 1000 zoospore DNA equivalents in duplicate (Hyatt
et al. 2007). Samples that were positive for Bd in the first round
of qPCR were re-run to confirm loads using the same template
dilution. We categorized an individual as Bd-positive if its swab
sample had ≥ 1 zoospore equivalent in both runs (Boyle et al.
2004). We calculated a mean prevalence (and 95% confidence
Herpetological Review 43(1), 2012
AMphibian Diseases
Acknowledgments.—We thank Corina Wack, Kari Leichty and
Damien Wilburn for assistance with swab collection; Ana Longo for
help with qPCR protocol optimization; and David Rodriguez and
Kelly Christiansen for creating the locality figure. The protocols were
approved by the Cornell University Institutional Animal Care and
Use Committee. This study was funded by a National Science Foundation grant to KMK and LDH (IOS-0808589).
Literature Cited
Boyle, D. G., D. B. Boyle, V. Olsen, J. A. T. Morgan, and A. D. Hyatt. 2004.
Rapid quantitative detection of chytridiomycosis (Batrachochytrium dendrobatidis) in amphibian samples using real-time Taqman
PCR assay. Dis. Aquat. Org. 60:141–148.
Byrne, M. W., P. D. Emily, and J. W. Gibbons. 2008. Batrachochytrium
dendrobatidis occurrence in Eurycea cirrigera. Southeast. Nat.
7:551–555.
Chinnadurai, S. K., D. Cooper, D. S. Dombrowski, M. F. Poore, and M.
G. Levy. 2009. Experimental infection of native North Carolina
salamanders with Batrachochytrium dendrobatidis. J. Wildl. Dis.
45:631–636.
Clopper, C. J., and E. S. Pearson. 1934. The use of confidence or fiducial
limits illustrated in the case of the binomial. Biometrika 26:404–
413.
Cummer, M. R., D. E. Green, and E. M. O’Neil. 2005. Aquatic chytrid
pathogen detected in terrestrial plethodontid salamander. Herpetol. Rev. 36:248–249.
Grant, E. H. C., L. L. Bailey, J. L. Ware, and K. L. Duncan. 2008. Prevalence of the amphibian pathogen Batrachochytrium dendrobatidis in stream and wetland amphibians in Maryland, USA. Applied
Herpetol. 5:233–241.
Groner, M. L., and R. A. Relyea. 2010. Batrachochytrium dendrobatidis
is present in northwest Pennsylvania, USA, with high prevalence
in Notophthalmus viridescens. Herpetol. Rev. 41:462–465.
Hyatt, A. D., D. G. Boyle, V. Olsen, D. B. Boyle, L. Berger, D. Obendorf,
A. Dalton, K. Kriger, M. Hero, H. Hines, R. Phillot, R. Campell, G.
Marantelli, F. Gleason, and A. Colling. 2007. Diagnostic assays and
sampling protocols for the detection of Batrachochytrium dendrobatidis. Dis. Aquat. Org. 73:175–192.
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Keitzer, S. C., R. Goforth, A. P. Pessier, and A. J. Johnson. 2011. Survey
for the pathogenic chytrid fungus Batrachochytrium dendrobatidis in southwestern North Carolina salamander populations. J.
Wildl. Dis. 47: 455–458.
Kriger, K. M., J. M. Hero, and K. J. Ashton. 2006. Cost efficiency in the
detection of chytridiomycosis using PCR assay. Dis. Aquat. Org.
71:149–154.
Montanucci, R. R. 2009. The chytrid fungus in the red salamander,
Pseudotriton ruber, in South Carolina, USA. Herpetol. Rev. 40:188.
Olson, D. H. 2010. Global Batrachochytrium dendrobatidis Mapping
Project.
http://www.parcplace.org/news-a-events/207-globalbd-mapping-project-update-7-july-2010.html
Ouellet, M., I. Mikaelian, B.D. Pauli, J. Rodrigue, and D.M. Green. 2005.
Historical evidence of widespread chytrid infection in North
American amphibian populations. Cons. Biol. 19:1431–1440.
Padgett-Flohr, G.E., and J.E. Longcore. 2005. Ambystoma californiense (California Tiger Salamander). Fungal infection. Herpet. Rev.
36:50–51.
Pasmans, R., R. Zwart, and A. D. Hyatt. 2004. Chytridiomycosis in the
Central American Bolitoglossine salamander (Bolitoglossa dofleini). Vet. Rec. 154:153.
Rothermel, B. B., S. C. Walls, J. C. Mitchell, C. K. Dodd, L. K. Irwin,
D. E. Green, V. M. Vazquez, J. W. Petranka, and D. J. Stevenson. 2008.
Widespread occurrence of the amphibian chytrid fungus Batrachochytrium dendrobatidis in the southeastern USA. Dis. Aquat.
Org. 82:3–18.
Timpe, E. K., S. P. Graham, R. W. Gagliardo, R. L. Hill, and M. G. Levy.
2008. Occurrence of the fungal pathogen Batrachochytrium dendrobatidis in Georgia’s amphibian populations. Herpetol. Rev.
39:447–449.
Vazquez, V. M., B. B. Rothermel, and A. P. Pessier. 2009. Experimental
infection of North American plethodontid salamanders with the
fungus Batrachochytrium dendrobatidis. Dis. Aquat. Org 84:1–7.
Weinstein, S. B. 2009. An aquatic disease on a terrestrial salamander:
individual and population level effects of the amphibian chytrid
fungus, Batrachochytrium dendrobatidis, on Batrachoseps attenuatus (Plethodontidae). Copeia 2009:653–660.
Herpetological Review, 2012, 43(1), 87–89.
© 2012 by Society for the Study of Amphibians and Reptiles
Detection of Batrachochytrium dendrobatidis in Amphibian
Populations of Northeast Ohio
The first North American occurrence of Batrachochytrium
dendrobatidis (Bd) was documented in Quebec in 1961 (Ouellet
et al. 2005). As of 2009, Bd had been detected in five Canadian
provinces and at least 38 US states (for example, Hossack et al.
2010; Sadinski et al. 2010; Fisher et al. 2009; Slough 2009; Muths
et al. 2008; Rothermel et al. 2008; Adams et al. 2007; Longcore et
al. 2007; Steiner and Lehtinen 2007; Ouellet et al. 2005, and many
others, see www.Bd-maps.net). In Ohio, Bd has been detected via
PCR analysis in wild Northern Leopard Frogs (Lithobates pipiens), American Toads (Anaxyrus americanus) (Scott and Sheafor, unpubl.), and Blanchard’s Cricket Frogs (Acris blanchardi)
(Steiner and Lehtinen 2008). In addition, Fowler’s Toads (Anaxyrus fowleri) and cricket frogs captured in Ohio and held at the
Toledo Zoo were later found to be infected with Bd (G. Lipps,
pers. comm.). It is uncertain, however, whether these animals
were infected prior to arriving at the zoo, or if they contracted it
from other infected species already present at the zoo. Histological surveys of Fowler’s Toad museum specimens originally collected from Ashtabula County, Ohio in 1977 determined that the
samples were Bd-positive (Scott and Sheafor, unpubl.), indicating that Bd has been in the northeastern Ohio region for at least
three decades. Because Bd has been detected in field-collected
amphibians from northeast Ohio, the possibility exists that the
Timothy J. Krynak *
Terry L. Robison
Cleveland Metroparks, 4101 Fulton Parkway, Cleveland, Ohio 44144, USA
Jonathan J. Scott
University of Mount Union, Academic Affairs, Biology Department, Alliance,
Ohio 44601, USA
*Corresponding author; e-mail: tjk@clevelandmetroparks.com
Herpetological Review 43(1), 2012