Officers and Editors for 2013-2014 President

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Publications Secretary BRECK BARTHOLOMEW P.O. Box 58517 Salt Lake City, UT 84158, USA

Immediate Past-President JOSEPH R. MENDELSON III Zoo Atlanta Atlanta, GA 30315, USA

Directors (Class and Category) FRANK BURBRINK (2016 R) College of Staten Island, USA ALISON CREE (2016 Non-US) University of Otago, NEW ZEALAND TIFFANY DOAN (2014 R) Central Connecticut State Univ., USA LISA HAZARD (2016 R) Montclair State University, USA TRAVIS LADUC (2014 Mem. at-Large) University of Texas, USA JENNIFER PRAMUK (2014 Cons) Woodland Park Zoo, USA CAROL SPENCER (2014 R) University of California, Berkeley, USA GREGORY WATKINS-COLWELL

(2016 R) Yale Peabody Mus. of Nat. Hist., USA

Trustee GEORGE PISANI University of Kansas, USA

Journal of Herpetology ERIN MUTHS, Co-Editor U.S. Geological Survey Fort Collins, CO 80526, USA GAD PERRY, Co-Editor Texas Tech University Lubbock, TX 79409, USA

Herpetological Review ROBERT W. HANSEN, Editor 16333 Deer Path Lane Clovis, CA 93619, USA

Contributions to Herpetology KRAIG ADLER, Editor Cornell University Ithaca, NY 14853-2702, USA

Facsimile Reprints in Herpetology AARON BAUER, Editor Villanova University Villanova, PA 19085, USA

Catalogue of American Amphibians and Reptiles

CHRISTOPHER BELL, Co-Editor University of Texas, Austin Austin, TX 78712, USA TRAVIS LADUC, Co-Editor University of Texas, Austin Austin, TX 78758, USA

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Herpetological Review, 2013, 44(3), 455–457.© 2013 by Society for the Study of Amphibians and Reptiles

First Report of Ranavirus in Plethodontid Salamanders from the Mount Rogers National Recreation Area, Virginia, USA Ranaviruses are an emerging pathogen responsible for nu-merous amphibian die-offs throughout Europe and North Amer-ica (Miller et al. 2011). In the southeastern United States, local anuran die-offs due to ranavirus have been observed (Green et al. 2002; Hoverman et al. 2012; Todd-Thompson 2010), and sug-gested to contribute to local species declines (Gray et al. 2009a). The southern Appalachian Mountains have one of the greatest global diversities of plethodontid salamanders (Dodd 2004), and disease emergence could have devastating impacts on commu-nity structure and ecosystem function (Whiles et al. 2006). Rana-virus infections have been reported in 14 species of plethodontid salamanders occurring in the southern Appalachian Mountains (Miller et al. 2011). Initial surveys for ranavirus infections have focused on the Great Smoky Mountains National Park (81% prevalence; Gray et al. 2009b), which is known for high species richness of plethodontid salamanders (Dodd 2004). However, much of the remainder of the southern Appalachian Mountains has not been investigated except for a single study in the Ridge and Valley physiographic province of Wise County, Virginia (33% prevalence; Davidson and Chambers 2011). Many other Appa-lachian peaks have high plethodontid salamander richness and include many species with limited distributions, but the occur-rence of ranavirus in these communities is unknown. We sampled salamanders in the Mount Rogers National Rec-reation Area (MRNA), Virginia from Whitetop and Beech Moun-tains (Grayson, Smyth, and Washington counties). Many species reach the extreme northern limit of their range on these moun-tains with 15 species of plethodontid salamanders found on Whitetop Mountain alone (Organ 1991). Our sampling focused on five species with aquatic larval stages (i.e., complex life cy-cle), which have shown higher prevalence of ranavirus infection (Gray et al. 2009b), and four terrestrial-breeding species with di-rect development. Nine species that we targeted for sampling included: Northern Dusky (Desmognathus fuscus), Seal (D. mon-ticola), Blue Ridge Dusky (D. orestes), Northern Pygmy (D. or-gani), Black-bellied (D. quadramaculatus), Blue Ridge Two-lined (Eurycea wilderae), Northern Gray-cheeked (Plethodon monta-nus), Ravine (P. richmondi), and Weller’s (P. welleri) salamanders. In Virginia, Weller’s, Northern Pygmy, Blue Ridge Dusky, and Blue Ridge Two-lined salamanders are listed as species of great-est conservation need (Virginia Department of Game and Inland Fisheries 2005). Salamanders were captured along transects from June to Au-gust 2008 and May to August 2009 from Byars Creek (1158–1311 m elev.; 36.62755°N, 81.58903°W), Whitetop Creek (1158–1615

m elev.; 36.64016°N, 81.60040°W), Dells Branch (945–1585 m elev.; 36.64774°N, 81.60254°W), Beech Mountain (956–1097 m elev.; 36.63362°N, 81.63030°W), and Daves Ridge (1128–1433 m elev.; 36.6475°N, 81.59185°W). Each transect was positioned perpendicular to the elevational gradient centered on either a stream or ridge line. We searched for salamanders on each side of the transect for one person-hour (~150 m) at every 30.5 m of elevation by turning all cover objects. We placed each captured salamander in individual 1.2-liter plastic bags. We used sterile forceps to remove a small tail section along the natural break point and placed each sample in a sterile microcentrifuge tube with 99% reagent grade isopropyl alcohol. We used a different set of sterilized forceps for each individual tail sample. Between sampling events, forceps were autoclaved to destroy remnant DNA. We extracted genomic DNA from tail samples using a DNeasy Blood and Tissue Kit (Qiagen Inc., Valencia, California), and used a QubitTM fluorometer to quantify the concentration of genomic nuclear DNA from tail samples. Samples from 2008 (N = 99) were extracted and quantified at the Diagnostic and Investigational Laboratory in the College of Veterinary Medicine at the Univer-sity of Georgia (UGA). In 2009, we extracted and quantified the 2009 samples (N = 320) in the Center for Wildlife Health at the University Tennessee (UT). Real-time qPCR was performed us-ing the identical primers and protocol of Gray et al. (2012). The qPCR was conducted using a SmartCycler system (Cepheid, Sunnyvale, California) for 2008 samples and an ABI 7900HT PCR system (Live Technologies Corp., Carlsbad, California) was used for 2009 samples. Controls included two positive controls (cul-tured virus and known positive tadpoles) and two negative con-trols (water and known negative animal). The qPCR was run in

M. KEVIN HAMED* Center for Wildlife Health, Department of Forestry, Wildlife, and Fisheries, Institute of Agriculture, University of Tennessee, 274 Ellington Plant Sciences Building, Knoxville, Tennessee 37966, USA; and Virginia Highlands Community College, 100 VHCC Drive, Abingdon, Virginia 24210, USAMATTHEW J. GRAYDEBRA L. MILLERCenter for Wildlife Health, Department of Forestry, Wildlife, and Fisheries, Institute of Agriculture, University of Tennessee, 274 Ellington Plant Sciences Building, Knoxville, Tennessee 37966, USA

*Corresponding author; e-mail: khamed@vhcc.edu

AMPHIBIAN DISEASESAMPHIBIAN DISEASES

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duplicate and a CT value ≤ 30 was deemed positive, according to

equipment optimization at UGA and UT. Our results document the first known presence of ranavirus infection in amphibians of the MRNA. Salamanders sampled and all other salamanders observed did not display external signs of ranavirus infection such as swelling or ulcerations (Miller et al. 2011). We detected ranavirus in seven salamander species (Table 1)—four of the species were first-time detections: Weller’s, North-ern Pygmy, Blue Ridge Dusky, and Northern Gray-cheeked (Mill-er et al. 2011). Ranavirus prevalence ranged from 3–9% within species, and was 3% among all species that tested positive (Ta-ble 1). Infected salamanders were found between elevations of 945–1554 m, and along all transects except Daves Ridge. Species with a more aquatic life history (D. fuscus and D. monticola) had the highest prevalence of infection (9%), as reported by Gray et al. (2009b). However, there was no association between larval life history (aquatic larvae or terrestrial direct development) and in-fection status (c2 = 0.01, p = 0.920), which may be a consequence of low overall prevalence. Additionally, there was no association between sampling years (c2 = 0.196, p = 0.658). Our low detection of ranavirus infection could be partly in-fluenced by the type of tissue collected. Gray et al. (2012) dem-onstrated that tail clips produced false-negative results for 20% of tail samples in American Bullfrog (Lithobates catebeianus) tadpoles when compared to liver tissue samples and suggested that false negatives could relate to the sensitivity of the amphib-ian host. Even with the possibility of a 20% false detection rate, salamanders from the MRNA appear to have lower ranavirus prevalence than salamanders from the Great Smoky Mountains National Park (81%; Gray et al. 2009b) and Wise County, Virginia (33%; Davidson and Chambers 2011). Future research could in-clude histological analyses to reduce the potential of false nega-tives (Gray et al. 2012) as well as genetic sequencing to identify the strain(s) of ranavirus present in the MRNA. We documented ranavirus infections occurring in secluded areas (often ≥ 300 m from trails, roads, or other areas of public ac-tivities) of the MRNA, indicating that this pathogen is not always associated with human-impacted sites (Gray et al. 2007). We doc-umented infections in four additional species of plethodontid salamanders, including rare species. Weller’s Salamanders are an IUCN (2008) Red List species, and our positive results from this species are a concern suggesting that further monitoring may be

warranted. These results provide additional evidence that rana-virus could be distributed throughout the Southern Appalachian Mountains. In the MRNA, ranavirus was found to infect both rare and more abundant salamanders as well as those with aquatic or terrestrial embryonic development, suggesting that ranavirus is not selective and any southern Appalachian salamander could be at risk. Additionally, six other species of plethodontid sala-manders, Eastern Newts (Notophthalmus viridescens), and five anuran species are found in the MRNA, but were not part of this study (Hamed, unpubl. data). Given the tendency for anurans to be more susceptible to ranavirus than salamanders (Hover-man et al. 2011), frogs may be important reservoirs and amplify-ing hosts for ranavirus (Paull et al. 2012). It would be prudent to include all amphibian species in future monitoring designs to provide a more comprehensive understanding of ranavirus prev-alence in the MRNA, and to increase the likelihood of identifying amplifying species and disease hotspots (Paull et al. 2012).

Acknowledgments.—This research was completed with funds provided by the Virginia Department of Game and Inland Fisheries through a State Wildlife Grant from the U.S. Fish and Wildlife Service and additional funding from the University of Tennessee Institute of Agriculture. We extend special thanks to James and Della Organ for providing sampling locations and guidance. We thank Gary Poe and Brian Parks for assistance in collecting and Becky Hardman, Kate Carpenter, and Lisa Whittington for assistance with molecular test-ing and extraction. Additionally, we are grateful to the MRNA staff for project assistance and to many private land owners for access to our study sites. This manuscript was improved with comments from Trang Nguyen, Dede Olson, and two anonymous reviewers. All sampling was approved by the Virginia Department of Game and In-land Fisheries (Scientific Collection Permit #41396) and followed UT Institutional Animal Care and Use Committee protocol #2084-0412.

literature cited

daVidson, s. r. a., and d. l. chamBers. 2011. Ranavirus prevalence in amphibian populations of Wise County, Virginia, USA. Herpetol. Rev. 42:540–542.

dodd, c. k., Jr. 2004. The amphibians of Great Smoky Mountains Na-tional Park. University of Tennessee Press, Knoxville, Tennessee. 283 pp.

Gray, m. J., d. l. miller, and J. t. hoVerman. 2012. Reliability of non-lethal surveillance methods for detecting ranavirus infection. Dis. Aquat. Org. 99:1–6.

———, ———, and ———. 2009a. Ecology and pathology of amphib-ian ranaviruses. Dis. Aquat. Org. 87:243–266.

———, ———, and ———. 2009b. First report of ranavirus infecting lungless salamanders. Herpetol. Rev 40:316–319.

———, ———, a. c. schmutzer, and c. a. Baldwin. 2007. Frog virus 3 prevalence in tadpole populations inhabiting cattle-access and non-access wetlands in Tennessee, USA. Dis. Aquat. Org. 77:97–103.

Green, d. e., k. a. conVerse, and a. k. schrader. 2002. Epizootiology of sixty-four amphibian morbidity and mortality events in the USA, 1996–2001. Ann. New York Acad. Sci. 969:323–339.

hoVerman, J. t., m. J. Gray, n. a. haisliP, and d. l. miller. 2011. Phylog-eny, life history, and ecology contribute to differences in amphib-ian susceptibility to ranaviruses. EcoHealth 8:301–319.

———, ———, d. l. miller, and n. a. haisliP. 2012. Widespread oc-currence of ranavirus in pond-breeding amphibian populations. EcoHealth 9:36–48.

iucn, conserVation international, and natureserVe. 2008. An analysis of amphibians on the 2008 IUCN Red List. <www.iucnredlist.org/amphibians>. Accessed 17 October 2012.

taBle 1. Ranavirus prevalence in salamander communities inhabit-ing Whitetop Mountain (Grayson, Smyth, Washington counties), Vir-ginia, USA.

Species No. infected / Prevalence of total no. sampled infection (95% confidence interval)

Desmognathus fuscus 1 / 11 0.09 (0.02–0.37)

Desmognathus monticola 2 / 22 0.09 (0.03–0.28)

Desmognathus orestes 3 / 145 0.02 (0.01–0.05)

Desmognathus organi 3 / 70 0.04 (0.01–0.12)

Desmognathus quadramaculatus 1 / 32 0.03 (0.01–0.16)

Eurycea wilderae 0 / 5 0 (0–0.44)

Plethodon montanus 3 / 101 0.03 (0.01–0.08)

Plethodon richmondi 0 / 3 0 (0–0.56)

Plethodon welleri 1 / 30 0.03 (0.01–0.17)

Total 14 / 419 0.03 (0.02–0.06)

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AMPHIBIAN DISEASES 457

miller, d., m. Gray, and a. storFer. 2011. Ecopathology of ranaviruses infecting amphibians. Viruses 3:2351–2373.

orGan, J. a. 1991. Salamander survey of the Mount Rogers Nation Recreation Area section two. U.S. Department of Agriculture, Marion, Virigina. 210 pp. Available from Mount Rogers National Recreation Area, Marion, Virginia.

Paull, s. h., s. sonG, k. m. mcclure, l. c. sackett, a. m. kilPatrick, and P. t. J. Johnson. 2012. From superspreaders to disease hotspots: linking transmission across hosts and space. Front. Ecol. Environ. 10:75–82.

semlitsch, r. d. (ed.). 2003. Amphibian Conservation. Smithsonian Books, Washington, DC. 320 pp.

stuart, s. n., J. s. chanson, n. a. cox, B. e. younG, a. s. l. rodriGues, d. l. Fischman, and r. w. waller. 2004. Status and trends of amphibian declines and extinctions worldwide. Science 306:1783–1786.

todd-thomPson, m. 2010. Seasonality, variation in species prevalence, and localized disease for Ranavirus in Cades Cove (Great Smoky Mountains National Park) amphibians. Unpubl. Master’s thesis, Uni-versity of Tennessee. http://trace.tennessee.edu/utk_gradthes/665/

VirGinia dePartment oF Game and inland Fisheries. 2005. Virginia’s Com-prehensive Wildlife Strategy. Virginia Department of Game and Inland Fisheries, Richmond, Virginia.

wake, d. B., and V. t. VredenBurG. 2008. Are we in the midst of the sixth mass extinction? A view from the world of amphibians. Proc. Natl. Acad. Sci. USA 105:11466–11473.

whiles, m. r., k. r. liPs, c. m. PrinGle, s. s. kilham, r. J. BixBy, r. Brenes, s. connelly, J. c. colon-Gaud, m. hunte-Brown, a. d. hury-na, c. montGomery, and s. Peterson. 2006. The effects of amphibian population declines on the structure and function of Neotropical stream ecosystems. Front. Ecol. Environ. 4:27–34.

Herpetological Review, 2013, 44(3), 457–458.© 2013 by Society for the Study of Amphibians and Reptiles

Batrachochytrium dendrobatidis in South Dakota, USA Amphibians

Although the amphibian chytrid fungus, Batrachochytri-um dendrobatidis (Bd), has been well-studied across the United States, there are still several re-gions where little is known re-garding its presence or preva-lence (Oullet et al. 2005; www.Bd-maps.net). Currently, there is no information regarding Bd occurrence in South Dakota, yet of the 15 amphibians species known to reside in South Dako-ta, many have been documented as hosts to Bd in other states (Oullet et al. 2005; Woodhams et al. 2008). Additionally, neighbor-ing states have documented the presence of Bd in these and oth-er species (Hossack et al. 2010; Rodriguez et al. 2009). To help fill the knowledge gap for South Da-kota, we sampled for the occur-rence of Bd at several locations across the state.

We opportunistically sampled amphibians for Bd in seven regions of South Dakota (Fig. 1) from April 2010–August 2011. Using nylon swabs, amphibians were rubbed along the ventral side of their hands, feet, legs and abdomen five times each for a total of 25 swipes per amphibian. To prevent cross contamina-tion, animals were handled and swabbed separately, and swabs were stored individually in tubes. Samples were kept on ice until brought to a molecular biology facility at the University of South Dakota where they were kept at 0°C until processed.

DNA was extracted from swabs using the included protocol for DNeasy® spin column kits (Qiagen, Inc., Germantown, Maryland). Each sample was run using an Applied Biosystems StepOne Plus quantitative PCR machine (Applied Biosystems, Inc., California)

according to the fast qPCR protocol as described by Kerby et al. (2013). Each sample was run in triplicate and was considered posi-tive if at least two of the three replicates were positive. A negative control was placed on each plate and the Bd standards utilized were from CSIRO laboratories in Australia. The pathogen load esti-mates are provided in zoospore genome equivalents per swab.

JENNIFER BROWNJACOB KERBY*University of South Dakota, Biology Department, 414 E. Clark St, Vermillion, South Dakota 57069, USA

*Corresponding author; e-mail: Jacob.Kerby@usd.edu

FiG. 1. Locations sampled for Batrachochytrium dendrobatidis (Bd) within South Dakota, USA. Blue dots represent sites with Bd detections.