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Batrachochytrium dendrobatidis Occurrence in Eurycea cirrigera
Michael W. Byrne, Emily P. Davie, and J. Whitfield Gibbons

Southeastern Naturalist, Volume 7, Number 3 (2008): 551–555

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Batrachochytrium dendrobatidis Occurrence in Eurycea cirrigera Michael W. Byrne1,*, Emily P. Davie1, and J. Whitfield Gibbons2 Abstract - Several pathogens affect amphibians, but a chytridiomycete fungus, Batrachochytrium dendrobatidis, is of particular interest because this pathogen is linked to localized amphibian population declines and extinction of species. Species-specific infections and pathogen distribution are poorly understood, particularly in members of Caudata. We found B. dendrobatidis in adult Eurycea cirrigera (Southern Two-lined Salamander), a species not previously reported as susceptible to this fungal pathogen, and this report is the first record of B. dendrobatidis occurrence in Alabama. Amphibians are host to several pathogens (Eiras 2005, Essbauer and Ahne 2001, Green et al. 2003, Hyatt et al. 2000, Speare et al. 1994), however, the chytridiomycete fungus, Batrachochytrium dendrobatidis (Longcore et al. 1999), has been identified as a causal factor in localized population declines and species’ extinctions (Berger et al. 1998, Bosch et al. 2001, Daszak et al. 1999, Skerratt et al. 2007). The Chydridiomycota (i.e., chytrids) is a phylum of early diverging fungi that form motile asexual spores that are distributed through water. Batrachochytrium dendrobatidis is the only member of this phylum known to be a pathogen of vertebrates (Longcore et al. 1999). Although some species of amphibians die from infection, e.g., Rana muscosa Camp (Mountain Yellow-legged Frog) (Briggs et al. 2005) and Bufo boreas Baird and Girard (Western Toad) (Carey et al. 2005), other species may harbor the pathogen with no signs of distress, e.g., R. catesbeiana Shaw (American Bullfrog) (Daszak et al. 2004). Fundamental understanding of the geographical and host distribution of B. dendrobatidis is limited, particularly for North American caudates, in which the chytrid fungus has been confirmed in 11 species in 7 families (Table 1). Herein we report the detection of B. dendrobatidis in the semi-terrestrial Eurycea cirrigera Green (Southern Two-lined Salamander), a genus not previously reported as susceptible to this fungal pathogen. This report expands the geographical distribution of B. dendrobatidis, extending its distribution to Alabama, a state from which the fungus was not previously reported. This report is part of an ongoing study to identify the distribution of B. dendrobatidis in several national parks in the southeastern US. The ≈825-ha Horseshoe Bend National Military Park (HOBE) is located in east-central Alabama (32.97583ºN, 85.73639ºW) in the Tallapoosa River basin. The Park is bisected by the Tallapoosa River and contains several first- and second-order perennial, ephemeral, and intermittent streams (with predominantly sand/gravel streambeds), both insular and originating outside the Park boundary. Based on the 30-year average during 1971–2000, mean-annual precipitation is 139 cm; March is the wettest month, and October is the driest. Mean high and low temperatures for January are 12 °C and -2 °C, and for July are 33 °C and 19 °C (http://www.ncdc.noaa. gov). HOBE is a mixed-pine hardwood forest primarily composed of Pinus taeda L. (Loblolly Pine) on ridges and Quercus spp. (oak), Carya spp. (hickory), and Fagus grandifolia Ehrh. (Beech) in swales. Elevation ranges from 183 to 217 m. HOBE falls within the predicted fundamental-niche boundary modeled for B. dendrobatidis by Ron (2005). Caudates at HOBE include Ambystoma maculatum (Spotted Salamander), Ambystoma opacum Gravenhorst (Marbled Salamander), Ambystoma talpoideum Holbrook (Mole Salamander), Desmognathus fuscus Green (Northern Dusky Salamander), Southern Two-lined Salamander, Eurycea guttolineata Holbrook (Three-lined Notes of the Southeastern Nat u ral ist, Issue 7/3, 2008 551 552 Southeastern Naturalist Notes Vol. 7, No. 3 Salamander), Gyrinophilus porphyriticus Green (Spring Salamander), Notophthalmus viridescens (Eastern Newt), Plethodon glutinosus Green (Slimy Salamander), Pseudotriton montanus Baird (Mud Salamander), and Pseudotriton ruber (Red Salamander) (Tuberville et al. 2005). Batrachochytrium dendrobatidis infections have been detected in Spotted Salamander, Red Salamander, and Eastern Newt in other parts of their range, and in 1 species expected to occur at HOBE, Ambystoma tigrinum (Tiger Salamander) (Table 1). We opportunistically collected samples from Southern Two-lined Salamanders and Three-lined Salamanders during 22–23 May 2006 and from Northern Dusky Salamanders, Southern Two-lined Salamanders, Three-lined Salamanders, Spring Salamanders (larva), Slimy Salamanders, and Red Salamanders during 10–15 October 2006. Swab samples for polymerase chain reaction (PCR) testing were collected from all captured animals during both sampling efforts. Spring 2006 samples were pooled for analysis. The pooled sample was composed of 9 Southern Two-lined Salamanders and 1 Threelined Salamander, and PCR results were positive for presence of B. dendrobatidis. The detection of B. dendrobatidis in the pooled spring 2006 sample prompted us to collect individual samples in fall 2006 to quantify B. dendrobatidis infection by species. We captured animals by hand and dip-net in creek beds, riparian zones, and hillsides at HOBE. Sampling effort was distributed equally across all first- and second-order streams at the park. Captured animals were toe-clipped for individual marking to avoid re-sampling of individuals and were subsequently released at the point of capture. All decontamination protocols set forth in NSW National Parks and Wildlife Service (2001) were followed. Individual swab samples were stored in a 70% EtOH solution in 2-mL vials, and pooled samples were stored in 50-mL vials. All samples were analyzed by Pisces Molecular, Boulder, CO. DNA extracted from swabs and alcohol were assayed for the presence of a 296 bp fragment of the B. dendrobatidis ribosomal RNA Intervening Transcribed Sequence region by 45-cycle Table 1. North American caudate species reported as positive for B. dendrobatidis. WC = wild capture, C = captive. Species Type Source Ambystoma californiense Gray WC Padgett-Flohr and Longcore (2005) (California Tiger Salamander) Ambystoma maculatum Shaw WC Ouellet et al. (2005) (Spotted Salamander) Ambystoma tigrinum Green WC Davidson et al. (2003) (Tiger Salamander) Amphiuma tridactylum Cuvier C Speare and Berger (2000) (Three-toed Amphiuma) Dicamptodon aterrimus Cope WC USGS National Wildlife Health Center (Idaho Giant Salamander) (2001) Necturus maculosus Rafinesque C Speare and Berger (2000) (Common Mudpuppy) Notophthalmus viridescens Rafinesque WC Ouellet et al. (2005) (Eastern Newt) Plethodon neomexicanus Stebbins and Riemer WC Cummer et al. (2005) (Jemez Mountains Salamander) Pseudotriton ruber Sonnini and Latrielle WC Don Nichols, pers. comm. 2002, in (Red Salamander) Speare and Berger (2000) updated April 2004 Siren lacertina Linnaeus (Greater Siren) C Speare and Berger (2000) Taricha torosa Rathke (California Newt) WC Padgett-Flohr and Longcore (2007) 2008 Southeastern Naturalist Notes 553 single-round PCR amplification with an established assay (J. Wood, Pisces Molecular, pers. comm.). Each group of sample PCR reactions included the following control reactions: a) positive DNA, i.e., DNA prepared from a positively identified laboratory culture of B. dendrobatidis previously demonstrated to be positive by PCR for the B. dendrobatidis target fragment; b) negative DNA, i.e., DNA prepared from a laboratory culture of a non-Batrachochytrium, chytrid fungus, previously demonstrated to be negative by PCR for B. dendrobatidis; and c) no DNA, i.e., H2O in place of template DNA. The no-DNA reaction remained uncapped during addition of sample DNAs to the test reactions, and served as a control to detect contaminating DNA in the PCR reagents or carryover of positive DNA during reaction set-up. Pisces Molecular has established a detection sensitivity of <0.1 zoospore equivalent in the PCR reaction (J. Wood, pers. comm.). Individual sample sizes in fall 2006 were small for 5 of the 6 species—Northern Dusky Salamander, n = 12; Three-lined Salamander, n = 7; Spring Salamander larva; n = 1; Slimy Salamander, n = 5; Red Salamander, n = 1—and all were negative for B. dendrobatidis. Sample size for Southern Two-lined Salamanders (n = 50), however, was sufficient for a determination of an infection prevalence of 42% (95% CI 28.2–55.8). The sample of Southern Two-lined Salamanders included three dead specimens, all of which tested positive for B. dendrobatidis. We cannot attribute mortality of the dead individuals sampled to chytridiomycosis, because we did not perform post-mortem examinations to detect other potential causes of death; one dead specimen was preserved for later analysis. All fall 2006 captures, except those for Slimy Salamander, were within 100 m of a Southern Two-lined Salamander individual that tested positive for B. dendrobatidis. Superficial examination of infected individuals from fall 2006 sampling revealed no lesions, skin discoloration, abnormal ecdysis, or exudate. Although several small (<2 mm width) wart-like protuberances were observed on the body and limbs of several individuals, none of those specimens tested positive for B. dendrobatidis. General observations of infected individuals included lethargy and tails that crumbled in response to very light handling (i.e., the tails had little structural integrity). Southern Two-lined Salamander is a relatively common semi-terrestrial species found in rocky streambeds and along stream banks at HOBE. During the spring breeding season, Southern Two-lined Salamanders deposit eggs under rocks and leaves along stream banks (Martof et al. 1980, Weichert 1945), and larvae live in small pools along streams for up to three years before metamorphosis (Mount 1975). The relationship between this prolonged contact with an aquatic environment and B. dendrobatidis infection in Southern Two-lined Salamander is unknown. In the non-breeding season, Southern Two-lined Salamanders can migrate to proximate aquatic habitats (Pauley and Watson 2005), which could explain the relatively equal spatial distribution of infected animals among first- and second-order streams across HOBE. Batrachochytrium dendrobatidis was also detected in several Rana clamitans Latreille (Green Frog) at HOBE; however, the relationship between fungal presence in Southern Two-lined Salamander and Green Frog is unknown. The presence of B. dendrobatidis in a semi-terrestrial salamander is of special concern given the diverse and largely endemic amphibian fauna characteristic of the southeastern US (Conant and Collins 1998, Gibbons and Buhlmann 2001, Kiester 1971, Tuberville et al. 2005) and the increasing levels of habitat stressors concomitant with rapid human-population growth rate in this region (Willson and Dorcas 2003). Despite the high prevalence of B. dendrobatidis detected in Southern Twolined Salamander, our sample sizes were inadequate to determine if B. dendrobatidis is present in other caudates at HOBE. Because B. dendrobatidis can be lethal to 554 Southeastern Naturalist Notes Vol. 7, No. 3 amphibians and is now known to be in Alabama as well as other states in the Southeast (Green and Dodd 2007, Peterson et al. 2007), we believe that determining its presence and effects on amphibian populations should be a priority. Continued monitoring for B. dendrobatidis at HOBE and other sites in the Southeast will improve our understanding of the relationship between the presence of B. dendrobatidis and short- and long-term changes in amphibian community composition and structure. Acknowledgments. We thank D. Lagana, J. Maxfield, B. Blankley, B. Huston, K. Ivins, and K. Funk for assistance with fieldwork; J. DeVivo, E. DiDonato, J.E. Longcore, and two anonymous reviewers for manuscript review; J. Wood at Pisces Molecular for technical assistance; and the staff at Horseshoe Bend National Military Park for assistance with logistical issues. The authors and all field personnel complied with all applicable Institutional Animal Care guidelines and the State of Alabama scientificcollection permit requirements. This project was funded as part of the USDI National Park Service Inventory and Monitoring Program and the Southeast Coast Network. Literature Cited Berger, L., R. Speare, P. Daszak, D.E. Green, A.A. Cunningham, C.L. Goggin, R. Slocombe, M.R. Ragani, A.D. Hyatt, K.R. McDonald, H.B. Hines, K.R. Lips, G. Marantelli, and H. Parkes. 1998. Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proceedings of the National Academy of Science, USA 95:9031–9036. Bosch, J., I. Martinez-Solano, and M. Garcia-Paris. 2001. Evidence of a chytrid fungus infection involved in the decline of the Common Midwife Toad (Alytes obstetricans) in protected areas of central Spain. Biological Conservation 97:331–337. Briggs, C.J., V.T. Vredenburg, R.A. Knapp, and L.J. Rachowicz. 2005. Investigating the population-level effects of chytridiomycosis: An emerging infectious disease of amphibians. Ecology 86:3149–3159. Carey, C., P.S. Corn, M.S. Jones, L.J. Livo, E. Muths, and C.W. Loeffl er. 2005. Factors limiting the recovery of Boreal Toads (Bufo b. boreas). Pp. 222–236, In M. Lannoo (Ed.), Amphibian Declines: The Conservation Status of United States Species. University of California Press, Los Angeles and Berkeley, CA. 1094 pp. Conant, R., and J.T. Collins. 1998. Reptiles and Amphibians: Eastern/Central North America. Houghton Miffl in Co., Boston, MA. 616 pp. Cummer, M.R., D.E. Green and E.M. O’Neill, 2005. Aquatic chytrid pathogen detected in terrestrial plethodontid salamander. Herpetological Review 36:248–249. Daszak, P., L. Berger, A.A. Cunningham, A.D. Hyatt, D.E. Green, and R. Speare. 1999. Emerging infectious diseases and amphibian population declines. Emerging Infectious Diseases 5:735–748. Daszak, P., A. Strieby, A.A. Cunningham, J.E. Longcore, C.C. Brown, and D. Porter. 2004. Experimental evidence that the Bullfrog (Rana catesbeiana) is a potential carrier of chytridiomycosis, an emerging fungal disease of amphibians. Herpetological Journal 14:201–207. Davidson, E.W., M. Parris, J.P. Collins, J.E. Longcore, A.P. Pessier, and J. Brunner. 2003. Pathogenicity and transmission of chytridiomycosis in Tiger Salamanders (Ambystoma tigrinum). Copeia 2003:601–607. Eiras, J.C. 2005. An overview of the myxosporean parasites in amphibians and reptiles. Acta Parasitologica 50:267–275. Essbauer, S. and W. Ahne. 2001. Viruses of lower vertebrates. Journal of Veterinary Medicine Series B 48:403–475. Gibbons, J.W., and K.A. Buhlmann. 2001. Reptiles and amphibians. Pp. 372-390, In J.G. Dickson (Ed.). Wildlife of Southern Forests: Habitat and Management. Hancock House Publishers, Blaine, WA. 552 pp. Green, D.E., and C.K.J. Dodd. 2007. Presence of amphibian chytrid fungus Batrachochytrium dendrobatidis and other amphibian pathogens at warmwater fish hatcheries in southeastern North America. Herpetological Conservation and Biology 2:43–47. 2008 Southeastern Naturalist Notes 555 Green D.E., S.H. Feldman, and J. Wimsatt. 2003. Emergence of a Perkinsus-like agent in anuran liver during die-offs of local populations: PCR detection and phylogenetic characterization. Proceedings of the American Association of Zoo Veterinarians 2003:120–121. Hyatt A.D., A.R. Gould, Z. Zupanovic, A.A. Cunningham, S. Hengstberger, R.J. Whittington, J. Kattenbelt, and B.E.H. Coupar. 2000. Comparative studies of piscine and amphibian iridoviruses. Archives of Virology 145:301–331. Kiester, A.R. 1971. Species density of North American amphibians and reptiles. Systematic Zoology 20:127–137. Longcore, J.E., A.P. Pessier, and D.K. Nichols. 1999. Batrachochytrium dendrobatidis gen. et sp. nov., a chytrid pathogenic to amphibians. Mycologia 91:219–227. Martof, B.S., W.M. Palmer, J.R. Bailey, and J.R. Harrison III. 1980. Amphibians and Reptiles of the Carolinas and Virginia. University of North Carolina Press, Chapel Hill, NC. 264 pp. Mount, R.H. 1975. The Reptiles and Amphibians of Alabama. Agricultural Experimental Station, Auburn University Press, Auburn, AL. 347 pp. NSW National Parks and Wildlife Service. 2001. Hygiene protocol for the control of disease in frogs. Information Circular Number 6. NSW NPWS, Hurstville NSW, Australia. 20 pp. 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. Conservation Biology 19:1431–1440. Padgett-Flohr, G.E., and J.E. Longcore. 2005. Ambystoma californiense (California Tiger Salamander). Fungal infection. Herpetological Review 36:50–51. Padgett-Flohr, G.E., and J.E. Longcore. 2007. Taricha torosa. Fungal infection. Herpetological Review 38:176–177. Pauley, T.K., and M.B. Watson. 2005. Eurycea cirrigera (Green, 1830); Southern Two-lined Salamander. Pp. 740–743, In M. Lannoo (Ed.). Amphibian Declines: The Conservation Status of United States Species. University of California Press, Los Angeles and Berkeley, CA. 1094 pp. Peterson, J.D., M.B. Wood, W.A. Hopkins, J.M. Unrine and M.T. Mendonca. 2007. Prevalence of Batrachochytrium dendrobatidis in American Bullfrog and Southern Leopard Frog larvae from wetlands on the Savannah River Site, South Carolina. Journal of Wildlife Diseases 43:450–460 Ron, S.R. 2005. Predicting the distribution of the amphibian pathogen Batrachochytrium dendrobatidis in the New World. Biotropica 37:209–221. Skerratt, L.F., L. Berger, R. Speare, S. Cashins, K.R. McDonald, A.D. Phillott, H.B. Hines, and N. Kenyon. 2007. Spread of chytridiomycosis has caused the rapid global decline and extinction of frogs. EcoHealth 4:125–134. Speare R., and L. Berger. 2000. Global distribution of chytridiomycosis in amphibians. Available online at Accessed 4 June 2007. Speare R., A.D. Thomas, P. O’Shea, and W.A. Shipton. 1994. Mucor amphibiorum in the Cane Toad, Bufo marinus, in Australia. Journal of Wildlife Diseases 30:399–407. Tuberville, T.D., J.D. Willson, M.E. Dorcas, J.W. Gibbons. 2005. Herpetofaunal species richness of southeastern National Parks. Southeastern Naturalist 4:537–569. USGS National Wildlife Health Center. 2001. USGS National Wildlife Heath Center quarterly mortality report. Available online at 2001_qtr_4.jsp. Accessed 4 June 2007. Weichert, C.K. 1945. Seasonal variation in the mental gland and reproductive organs of the male Eurycea bislineata. Copeia 1945:78–84. Willson, J.D., and M.E. Dorcas. 2003. Effects of habitat disturbance on stream salamanders: Implications for buffer zones and watershed management. Conservation Biology 17:763–771. 1National Park Service, Southeast Coast Inventory and Monitoring Network, Cumberland Island National Seashore, PO Box 806, Saint Marys, GA 31558, USA. 2 University of Georgia, Savannah River Ecology Laboratory, Aiken, SC 29802. *Corresponding author - Michael_W_