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Proportion of Hosts Carrying Batrachochytrium dendrobatidis, Causal Agent of Amphibian Chytridiomycosis, in Oswego County, NY in 2012
Sofia T. Windstam and Jennifer C. Olori

Northeastern Naturalist, Volume 21, Issue 1 (2014): NENHC-25—NENHC-34

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Northeastern Naturalist Vol. 21, No. 1 S.T. Windstam and J.C. Olori 2014 NENHC-25 2014 NORTHEASTERN NATURALIST 21(1):NENHC-25–NENHC-34 Proportion of Hosts Carrying Batrachochytrium dendrobatidis, Causal Agent of Amphibian Chytridiomycosis, in Oswego County, NY in 2012 Sofia T. Windstam1 and Jennifer C. Olori1,* Abstract - Although the fungus Batrachochytrium dendrobatidis is a causal agent behind precipitous amphibian declines globally, little is known about its regional distribution in New York State (NYS). With an aim toward increased understanding of B. dendrobatidis prevalence locally, we collected amphibians between April through November 2012 at the Rice Creek Field Station in Oswego County, NY, and took swabs of the ventral surfaces of all individuals caught. Polymerase chain reaction on DNA extracted from swabs and comparison with B. dendrobatidis control DNA showed that 30% of amphibians sampled carried the fungus, with prevalence ranging between 20–50% for Lithobates catesbeianus (Bull Frogs), Lithobates clamitans (Green Frogs), Pseudacris crucifer (Spring Peepers), and Eurycea bislineata (Two-lined Salamanders). We detected Batrachochytrium dendrobatidis only during the months of April, May, June, August, and September of the sampling period. June and September had the highest percentage of amphibians infected with B. dendrobatidis at 32 and 48%, respectively. This study represents the first time that B. dendrobatidis has been documented in Oswego County and only the second time that the fungus has been documented in NYS. The documented prevalence levels in combination with lack of observed mass amphibian declines suggest that the fungus may be endemic in local amphibian populations, but additional research is needed to establish the relative importance of these data for the health of amphibian populations in Oswego County a nd NYS. Introduction Amphibian chytridiomycosis is an emerging infectious disease of concern caused by the chytrid fungus Batrachochytrium dendrobatidis Longcore, Pessier, & D.K. Nichols (Longcore et al. 1999). Chytridiomycosis was first detected in mass die-offs of amphibian populations in Central America and Australia in the late 1990s. Since then, infection by B. dendrobatidis has been linked to rapid amphibian declines and extinctions around the world (Berger et al. 1998, Bosch et al. 2001, Briggs et al. 2005, Wake and Vredenberg 2008). Batrachochytrium dendrobatidis has been documented to infect 508 different amphibian species and more recently also Crayfish (Procambrus spp. and Orconectes virilis (Hagen) [Virile or Northern Crayfish]), a non-amphibian host, in Louisiana (Fisher et al. 2009a, 2009b; McMahon et al. 2013; Olson et al. 2013). Batrachochytrium dendrobatidis forms sporangia in the skin of infected amphibian hosts which shed the flagellated, aquatic, motile zoospores responsible for re-infecting the original host as well as infecting new hosts (Longcore et al. 1999, 1Department of Biological Sciences, State University of New York at Oswego, Oswego, NY. *Corresponding author - jennifer.olori@oswego.edu. Manuscript Editor: Todd Rimkus Northeastern Naturalist NENHC-26 S.T. Windstam and J.C. Olori 2014 Vol. 21, No. 1 Rosenblum et al. 2010, Wake and Vredenberg 2008). Zoospore locomotion relies on the presence of free-water films. The fungal infection is restricted to keratin present in the epidermis of adult amphibians and larval salamanders and also in the mouthparts of tadpoles. Although in tadpoles the specific effect of B. dendrobatidis infection varies by species (Blaustein et al. 2005), tadpoles often do not die as a direct result of infection, and the infection is lost at metamorphosis. However, newly metamorphosed individuals can become re-infected and then die (Carey et al. 2003). Adults of different host species also vary in their susceptibility to B. dendrobatidis; Lithobates catesbeianus (Shaw) (Bullfrog) and Ambystoma tigrinum Green (Tiger Salamander), both native to NYS, sometimes are considered to be reservoirs or vectors because they may carry fungal infections without suffering from chytridiomycosis (Carey et al. 2003, Gahl et al. 2011, Searle et al. 2011; but see Gervasi et al. 2013). However, in many species, the mortality rate following infection is 55–99% of the population (Carey et al. 2003). The pathogen causes a characteristic thickening of the stratum corneum (upper-most layer of epidermis) that disrupts the ability of infected animals to regulate ion transport across the skin, ultimately causing decreased blood-plasma potassium and sodium levels that leads to cardiac arrest (Voyles et al. 2009). However, there are also indications that B. dendrobatidis may release some sort of a chemical that is partially responsible for the observed host pathology (Carey et al. 2003, McMahon et al. 2013). Amphibian chytridiomycosis is unusual not only because fungi of the phylum Chytridiomycota rarely are severe animal pathogens, but because B. dendrobatidis is the first chytrid fungus known to infect vertebrates (Longcore et al. 1999). Furthermore, it is currently not known if chytridiomycosis in amphibians has become widespread because of a recent change in virulence of B. dendrobatidis or susceptibility of its amphibian hosts (perhaps caused by environmental changes or anthropogenic stressors). Another possibility is that the fungus was recently introduced and has since spread to multiple new regions (Carey et al. 2003, Kilpatrick et al. 2009, Lips et al. 2008). Despite the presence of B. dendrobatidis in Quebec, Canada (Ouelett et al. 2005), and the Northeast US (Longcore et al. 2007), specimens from NYS were not reported to test positive for the presence of B. dendrobatidis until 2012 (Becker et al. 2012), and amphibian declines connected to chytridiomycosis have not been described in the Northeast. Current hypotheses suggest that either declines are ongoing but have not been detected because of a lack of long-term population data in the Northeast, or that the absence of declines are real and may result from lessvirulent local strains of B. dendrobatidis or a natural resistance to the pathogen in at least some local species (Gahl et al. 2011). Moreover, results from recent studies in the Southeast suggested strong seasonality in the prevalence of B. dendrobatidis infections, with peak prevalence (45% of individuals) occurring in the spring, whereas levels are minimal in the fall (2%) (Pullen et al. 2010 ). The apparent lack of widespread, pathogenic outbreaks of chytridiomycosis and mass-die offs of amphibians in the Northeast also could result from a combination of those factors proposed above, such that a local strain of B. dendrobatidis Northeastern Naturalist Vol. 21, No. 1 S.T. Windstam and J.C. Olori 2014 NENHC-27 may be controlled by climactic and seasonal variables specific to the Northeast in conjunction with the resistance of at least some native species (Gahl et al. 2011). However, because local amphibian population trends are not actively monitored in many areas, and local species have not been tested regularly for the presence and prevalence of B. dendrobatidis, it is unknown which or how many strains could be present in the area, and little information exists on the annual pathogenicity and transmission patterns of B. dendrobatidis in the Northeast. Our goal was to investigate the prevalence of B. dendrobatidis infection in local populations of amphibians in order to address the following questions: 1) Is B. dendrobatidis present in local populations? 2) Do patterns of prevalence in the Northeast reflect similar seasonal patterns detected in the Sout heast? The first step in this long-term project was to address whether or not B. dendrobatidis presently is infecting local amphibians. Materials and Methods Field-site description and sampling of amphibians We sampled a total of 82 amphibians representing five different species (6 Bull Frogs, 49 Lithobates clamitans Latreille [Green Frogs], 1 Lithobates pipiens (Schreber) [Northern Leopard Frog], 5 Pseudacris crucifer (Wied-Neuwied) [Spring Peepers], and 16 Eurycea bislineata (Green) [Two-lined Salamanders]) over 11 visits to the SUNY Oswego Rice Creek Field Station (RCFS) between the months of April through November 2012 (Fig. 1). The majority of RCFS is abandoned agricultural land that is being allowed to undergo natural succession, with Figure 1. Number of individuals of each species of amphibian sa mpled by month. Northeastern Naturalist NENHC-28 S.T. Windstam and J.C. Olori 2014 Vol. 21, No. 1 different areas at various stages of succession represented (Weeks and Cox 1988). The site contains multiple wetlands (e.g., large pond, creek, marshes, and vernal pools), meadows, old woodlots, and hardwood forest. All sampled individuals were adults or sub-adults, and the vast majority of animals sampled were collected from or adjacent to the pond, creek, or vernal pools. We captured animals either using nets or by hand with sterile nitrile gloves. We then identified individuals to species and swabbed (Medical Wire and Equipment Co., UK) each one five times along each of the ventral surfaces of the hands, legs, feet, abdomen, thighs, and pelvic patch. Swab tips were broken off into sterile 1.5-ml microcentrifuge tubes containing one ml of 70% ethanol, which were then stored at -4 ºC until processed. We also collected standard data such as snout–vent length, weight, sex, and condition (external appearance and overall health), along with environmental information (temperature, humidity, weather, etc.) for each swabbed individual and entered the information into a database for long-term population monitoring. During sampling, we placed each amphibian into a new plastic bag to prevent cross-contamination of equipment, and all gloves were changed and all equipment was sterilized using either 10% bleach solution or 70% ethanol after each individual was processed. DNA extraction We transferred swabs to sterile 2-ml polypropylene tubes (Biospec, OK) containing 30–40 mg of sterile 0.5-mm zirconium/silica beads (Biospec). We added fifty μl of PrepMan Ultra DNA extraction solution (Applied Biosystems, CA) to each tube and homogenized the samples for 50 s using a Mini-Beadbeater-1 (Biospec). We then centrifuged the samples for 30 s at 13,000 × g, after which we repeated homogenization and spinning of the samples. Thereafter, samples were boiled for 10 min, cooled for 2 min on ice, and centrifuged for 3 min at 13,000 × g. We collected 20 μl of the resulting supernatant to be stored long term at -80 ºC (Hyatt et al. 2007). PCR amplification and gel electrophoresis Although quantitative PCR (qPCR) has been the standard when diagnosing B. dendrobatidis infections of amphibians, a recent publication demonstrated that conventional end-point PCR is equally sensitive in detecting B. dendrobatidis and therefore is appropriate for analyses in which prevalence data are collected (Boyle et al. 2004, Garland et al. 2011). We diluted all DNA extracts tenfold in molecular grade water prior to subjecting them to PCR. PCR reactions contained 0.8 u Fast- Start Taq polymerase (Roche Applied Science, IN), 0.25 mM each of dATP, dCTP, dGTP, and dTTP (Invitrogen, Life Technologies, NY), 0.9 μM forward primer ITS1-3 Chytr (5’-CCTTGATATAATACAGTGTGCCATATGTC-3’, Invitrogen), 0.9 μM reverse primer 5.8S Chytr (5’-AGCCAAGAGATCCGTTGTCAAA-3’, Invitrogen), 3 mM MgCl2, 4 μl diluted DNA template, and sterile molecular-grade water to achieve a final reaction volume of 20 μl (Boyle et al. 2004, Garland et al. 2011). We performed PCR using the following temperature protocol: initial denaturation at 95 ºC for 4 min followed by 50 cycles of denaturing at 95 ºC for 30 s, annealing at 55 ºC for 30 s, and extension at 72 ºC for 45 s. We carried out all PCR Northeastern Naturalist Vol. 21, No. 1 S.T. Windstam and J.C. Olori 2014 NENHC-29 reactions in individual 0.2-ml PCR tubes, and each round of PCR included a negative and a positive control using sterile molecular-grade water and DNA isolated from B. dendrobatidis cultures, respectively. We mixed 10-μl aliquots of PCR products with loading dye and analyzed them using 2% agarose gels in 1X TAE buffer. The positive control DNA from B. dendrobatidis generated an anticipated band size of 146 bp, and we deemed samples to be positive for the presence of B. dendrobatidis if: 1) controls yielded the expected results (no amplification in negative control and amplification in positive control) and 2) the sample produced a band of the anticipated size. Band sizes were ascertained by comparing migrations of DNA fragments in a 100-bp DNA ladder (Fisher BioReagents, PA) as well as comparing to the relative band location of the positive control. Results and Discussion Batrachochytrium dendrobatidis was detected in 30% of the sampled amphibians (data not shown). The fungus was detected on Green Frogs, Bull Frogs, Spring Peepers, and Two-lined Salamanders, with a prevalence of 33, 50, 20, and 20%, respectively (Fig. 2). The prevalence levels for Bull Frogs and Spring Peepers in particular have to be viewed as tentative due to the low number of animals sampled (6 Bull Frogs and 5 Spring Peepers). Likewise, the absence of B. dendobatidis on Northern Leopard Frogs has to be viewed with similarly guarded skepticism due to only one sampled individual. Survey data of B. dendrobatidis prevalence levels for Two-lined Salamanders at other Northeast sites is lacking, and our study presents a preliminary glance at prevalence levels of the chytrid fungus on this amphibian. A recently published survey in the Adirondacks, NY, found B. dendrobatidis to be Figure 2. Overall B. dendrobatidis prevalence rates on all amphibian species found to test positive for the fungus. Northeastern Naturalist NENHC-30 S.T. Windstam and J.C. Olori 2014 Vol. 21, No. 1 present on approximately 25% of Green Frogs sampled in the latter part of June at two pond sites (Becker et al. 2012). Thirty-three percent of Green Frogs at RCFS tested positive for Bd, demonstrating that 25–33% of Green Frogs carry B. dendrobatidis at two geographically distinct locations in NYS. How indicative these B. dendrobatidis prevalence levels are among Green Frogs in NYS remains to be established, as this study along with Becker et al. (2012) are the only studies to date that have examined B. dendrobatidis prevalence. The significance of B. dendrobatidis prevalence levels is not yet clear because of the lack of published reports of amphibian population declines connected to chytridiomycosis in the Northeast. It is possible that population declines are ongoing, declines have already taken place, or the lack of decline is actually real. Lack of current observable population declines could result from one or a combination of factors, including the introductory wave of infection having already swept through the amphibian population locally, the presence of less virulent local isolates of B. dendrobatidis in NYS, some level of protection afforded by local environmental or climatic conditions, or a higher natural resistance against the pathogen in our local populations (Gahl et al. 2011). Using Green Frogs as an example, Gahl et al. (2011) demonstrated that these particular amphibians are susceptible to chytrid infection and die at appreciable rates once infected, but that there exists an isolatespecific effect. Green Frogs were more susceptible to a novel isolate originating from Panama, and not at all sensitive to a strain isolated from the Northeast (Gahl et al. 2011). Consequently, in order to tease apart whether observed B. dendrobatidis prevalence levels may be linked to any past or future declines it will be critical to explore which isolates of B. dendrobatidis are present locally in order to ascertain potential effects of the fungus on local amphibian populations. Lack of disease may simply be a matter of local isolates being less virulent, but until this has been established experimentally, it remains mere speculation. Obtaining local B. dendrobatidis isolates would also allow for further controlled studies on potential modulating effects of climatic conditions on infection outcomes. Furthermore, resistance in some amphibians against B. dendrobatidis may be conferred by resident skin-microbe populations (Harris et al. 2006, 2009a, 2009b), but it is unclear whether or not such resistance mechanisms are in place in our local populations, and this possibility remains to be explored. Until recently, NYS had no published records of amphibians testing positive for the presence of B. dendrobatidis (Becker et al. 2012). Our study represents the first time that B. dendrobatidis has been documented in amphibians of Oswego County. In Virginia, a strong seasonality has been described for B. dendrobatidis prevalence at six different sites in both rural and urban areas, where rates of amphibians carrying B. dendrobatidis peaked in spring between March–June and declined to 2% in the fall (Pullen et al. 2010). In NYS, it is unclear whether such seasonality is expected to take place, but our data suggest that the prevalence of B. dendrobatidis fluctuates, which may be a random pattern or the result of seasonality. Potentially seasonal effects could simply be a function of physiology because B. dendrobatidis displays optimal growth at 17–25 °C, with temperatures below 5 °C and at or above Northeastern Naturalist Vol. 21, No. 1 S.T. Windstam and J.C. Olori 2014 NENHC-31 28 °C significantly restricting growth (Piotrowski et al. 2004). We detected B. dendrobatidis on amphibians only during April, May, June, August, and September of the sampling period, during which June and September had the highest proportions of amphibians infected with B. dendrobatidis at 32 and 48%, respectively (Fig. 3). Data from the Adirondacks in NYS provides plausible support for the effect of temperature on infection because a site with increased canopy cover experiencing lower temperatures had higher B. dendrobatidis infection loads on Green Frogs than individuals from a site with less cover and increased temperature (Becker et al. 2012). Considering the effect temperature has on B. dendrobatidis physiology it stands to reason that NYS, with a climate that experiences significant seasonal variation in temperature, would exhibit some form of seasonal variation (Becker et al. 2012, Piotrowski et al. 2004, Pullen et al. 2010). However, additional data from multiple years and sites locally are necessary prior to inferring with certainty the nature of these fluctuations. This study is the first documentation of B. dendrobatidis in Oswego County in NYS and represents a first step in addressing the effect this pathogen may have on our local amphibian populations. In order to interpret the significance of our observations, additional research on many aspects of infection is required, including continued monitoring and surveying of amphibian populations for B. dendrobatidis across different seasons. For future work, isolation and description of the fungal isolates present locally is necessary in order to determine pathogen population variation and structure, which are vital for explaining variation in pathogenicity and virulence should this be an underlying factor that explains lack of observed population declines (Fisher et al. 2009a, Gahl et al. 2011). The genetic population structure of B. dendrobatidis may also provide information on whether or not the Figure 3. Prevalence rate of B. dendrobatidis on all sampled amphibians per month sampled. Northeastern Naturalist NENHC-32 S.T. Windstam and J.C. Olori 2014 Vol. 21, No. 1 fungus was recently introduced regionally as opposed to displaying some endemism (Morgan et al. 2007). Acknowledgments Positive control DNA extracted from B. dendrobatidis cultures was very generously donated by Dr. Kelly Zamudio at the Department of Ecology and Evolutionary Biology at Cornell University. This research was supported by a grant from the Rice Creek Associates to J.C. Olori and S.T. Windstam. We are indebted to the numerous undergraduate students, staff, and faculty members who have participated in the field sampling of amphibians and molecular diagnostics. Literature Cited Becker, C.G., D. Rodriguez, A.V. Longo, A.L. Talaba, and K.R. Zamudio. 2012. 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