SARAH REINTJES-TOLEN UNIVERSITY OF FLORIDA

Transcription

1 GEOGRAPHIC DISTRIBUTION OF CHYTRID FUNGUS (Batrachochytrium dendrobatidis) AND Ranavirus spp. IN AMPHIBIANS IN NORTHERN PENINSULAR AND PANHANDLE FLORIDA: WITH A CASE OF A RANAVIRUS DIE-OFF IN GOLD HEAD BRANCH STATE PARK By SARAH REINTJES-TOLEN A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA

2 2012 Sarah Reintjes-Tolen 2

3 To my parents, for always supporting me in my nature related curiosities and encouraging my intellectual endeavors 3

4 ACKNOWLEDGMENTS I would firstly like to thank the members of my committee, Dr. Raymond R. Carthy, Dr. Kenneth L. Krysko, and Dr. D. Bruce Means for their time, guidance and support. I would like to thank Kevin M. Enge and Paul E. Moler for assisting me in the field with their invaluable knowledge of Florida amphibians. I would like to thank Dr. Jan Landsberg for writing the grant and receiving funding for this project and the FWRI, St. Petersburg, research team for their assistance with this project. I would additionally like to thank Joseph Mansuetti, Lindsay M. Wagner and Stephen Harris for help collecting samples. I am grateful for help from the Florida Fish and Wildlife Conservation Commission, the Central Florida Zoological Gardens, the Wildlife Disease Laboratories at the San Diego Zoo and the Florida Museum of Natural History. I would like to thank Dr. Kent Vliet for giving me the opportunity to receive funding through a Teaching Assistantship with the Biology Department. I would like to thank Natalie C. Williams, Michael A. Gil, Marvin V. Morales, Jason Fidorra, Anthony Lau, Noah L. Mace, Dana J. Ehret, Carly L. Manz, Meghan Godby and Buddy Coleman for their support both academically and emotionally. I am very appreciative of the unconditional love of my dog Syd who was always there for me. Finally I would like to thank my parents, Susan Reintjes and Steve Tolen for their moral support, not only during the past few years but my entire life. Without their continued support, I would have never made it this far. 4

5 TABLE OF CONTENTS page ACKNOWLEDGMENTS... 4 LIST OF TABLES... 7 LIST OF FIGURES... 8 ABSTRACT...10 CHAPTER 1 BACKGROUND INFORMATION...12 Overview...12 Amphibian Anatomy and Life History...14 Batrachochytrium Dendrobatidis Life History...16 Ranavirus spp. Life History...19 Amphibian Declines in the State of Florida...22 Current Accounts of Amphibian Disease in Florida...23 Objectives and Significance of Current Study...23 Overview of Study Area GEOGRAPHIC DISTRIBUTION OF CHYTRID FUNGUS (Batrachochytrium dendrobatidis) AND RANAVIRUS SPP. IN NORTHERN PENINSULAR AND PANHANDLE FLORIDA AMPHIBIANS...36 Introduction...36 Methods...36 Sampling Sites...37 Species Sampled...37 Amphibian Sampling...38 Swabbing for Batrachochytrium Dendrobatidis...39 Swabbing for Ranavirus spp Environmental Factors...40 Disinfection...41 Pathogen Detection...41 Results...42 Discussion RANAVIRUS MORTALITY EVENT AT MIKE ROESS GOLD HEAD BRANCH STATE PARK, FLORIDA, USA...59 Introduction...59 Methods...60 Results

6 Discussion ANALYSIS OF PRESERVED AMPHIBIAN SPECIMENS TO TEST FOR DETECTION OF CHYTRID FUNGUS (Batrachochytrium dendrobatidis)...66 Introduction...66 Methods...66 Results...67 Discussion SUMMARY AND CONSERVATION SIGNIFICANCE...73 APPENDIX A STATE PARK SAMPLING PERMITS...76 SAMPLING PERMIT-A...76 SAMPLING PERMIT-B...78 B ARC APPROVAL FORM...80 LIST OF REFERENCES...81 BIOGRAPHICAL SKETCH

7 LIST OF TABLES Table page 2-1 The locality of each pond where amphibians were sampled for Batrachochytrium dendrobatidis and/or Ranvirus spp Summary table of data collected on chytrid fungus, Batrachochytrium dendrobatidis, in northern peninsular and panhandle Florida Summary table of data collected on Ranavirus spp., in northern peninsular and panhandle Florida Environmental data collected at each pond where amphibians were sampled for Batrachochytrium dendrobatidis and/or Ranvirus spp Desmognathus auriculatus specimens from the Florida Museum of Natural History that were sampled for Batrachochytrium dendrobatidis Specific locality and number of Desmognathus auriculatus specimens from the Florida Museum of Natural History that were sampled for Batrachochytrium dendrobatidis

8 LIST OF FIGURES Figure page 1-1 The life cycle of the chytrid fungus, Batrachochytrium dendrobatidis, illustrating substrate-dependent and substrate-independent stages Photomicrograph of a stained kidney from a green frog (Lithobates clamitans) larva infected with Ranavirus spp Representative pond (Pond 3) within Apalachicola National Forest, Liberty County, Florida, USA. Photo by Kevin M. Enge Representative pond (Pebble Lake) within Gold Head Branch State Park, Clay County, Florida, USA. Photo by Sarah Reintjes-Tolen Representative pond (Pond 3) within Camp Blanding Military Reserve, Clay County, Florida, USA. Photo by Sarah Reintjes-Tolen Representative pond (Pond 1) within Osceola National Forest, Baker County, Florida, USA. Photo by Sarah Reintjes-Tolen Representative pond (Pond 21) within Ocala National Forest-South of Salt Springs, Marion County, Florida, USA. Photo by Sarah Reintjes-Tolen Representative pond (Pond 1) within Ocala National Forest-South of Church Lake, Marion County, Florida, USA. Photo by Sarah Reintjes-Tolen Representative pond (Pond 7) within Etoniah Creek State Forest, Putnam County, Florida, USA. Photo by Sarah Reintjes-Tolen Map illustrating sampling sites where amphibians were swabbed for Batrachochytrium dendrobatidis and/or Ranavirus spp., in Florida, USA Sampling amphibians for Batrachochytrium dendrobatidis and Ranavirus spp. and recording data from individuals caught in Camp Blanding Military Reserve Sampling a Southern Leopard Frog (Lithobates sphenocephalus) tadpole for Batrachochytrium dendrobatidis and Ranavirus spp at Ocala National Forest Sampling an adult Southern Cricket Frog (Acris gryllus) for Batrachochytrium dendrobatidis at Ocala National Forest-South of Church Lake Sampling an adult Southern Cricket Frog (Acris gryllus) for Ranavirus spp. at Ocala National Forest-South of Church Lake

9 2-5 Map of sites sampled for Batrachochytrium dendrobatidis and Ranvirus spp. mapped using Google Earth Number of individual amphibians, sampled for both Batrachochytrium dendrobatidis and Ranavirus spp., by species and life stage Location of Pebble Lake in Mike Roess Gold Head Branch State Park, Clay County, Florida, USA Lateral view of Bullfrog (Lithobates catesbeianus) tadpole with white lesions, indicative of Ranavirus spp. infection, Pebble Lake Ventral view of Bullfrog (Lithobates catesbianus) tadpole with swollen, blotched belly and red swollen vent, both indicative of Ranavirus spp Five southern dusky salamander (Desmognathus aurituculatis) specimens from Deep Springs Canyon, Bay County, FL, which were sampled for Batrachochytrium dendrobatidis Map showing collection location of each Desmognathus auriculatus specimen sampled for Batrachochytrium dendrobatidis

10 Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science GEOGRAPHIC DISTRIBUTION OF CHYTRID FUNGUS (Batrachochytrium dendrobatidis) AND Ranavirus spp. IN AMPHIBIANS IN NORTHERN PENINSULAR AND PANHANDLE FLORIDA WITH A CASE OF A RANAVIRUS DIE-OFF IN GOLD HEAD BRANCH STATE PARK Chair: Raymond R. Carthy Major: Wildlife Ecology and Conservation By Sarah Reintjes-Tolen August 2012 Amphibian species across the globe have been experiencing population declines. There are a number of factors attributed to these declines, including habitat destruction, increased ultraviolet radiation, pollution, and introduced diseases. However, one of the most potentially devastating of these factors has been the emergence of two pathogens, chytrid fungus (Batrachochytrium dendrobatidis) and Ranavirus spp. Determining the distribution of these pathogens is important as it is unknown what kind of impact these pathogens are capable of having on frog populations. In this study, I attempt to document the presence and geographic distribution of two pathogens, B. dendrobatidis and Ranavirus spp., in northern peninsular and panhandle Florida. During March through May of 2011 and February and March of 2012, I surveyed seven sites in northern peninsular and panhandle Florida. After surveying a total of 32 ponds, B. dendrobatidis was detected in two of 32 ponds surveyed, one in Camp Blanding Military Reserve and one in Osceola National Forest. Ranavirus spp. was detected in one pond in Gold Head Branch State Park. Additionally, 10

11 a Ranavirus spp. mortality event was discovered at Pebble Lake in Gold Head Branch State Park. This initial survey of these two amphibian diseases will illustrate the geographic distribution of these pathogens, enabling researchers to track the spread of these diseases and assess which populations of amphibians are most at risk. 11

12 CHAPTER 1 BACKGROUND INFORMATION Overview Currently there are more than 6,300 known species of amphibians, with new species being discovered each year (Cheng et al., 2011; Stuart et al., 2004). Amphibians live in both water and on land and have semipermeable skin that makes them very sensitive to changes in their environment (Shoemaker et al., 1992). Due to their unique physiology, amphibians are considered to be indicator species showing physical or behavioral changes with even the smallest variation in physical, chemical, or ecological make-up of their surroundings (Mawdsley & O Malley, 2009). Some amphibian species, such as salamanders in the Family Plethodontidae, are capable of gas exchange through their skin (Shoemaker et al., 1992). Additionally, amphibians possess a permeable integument, which acts as a conduit between the organism and its environment (Bentley and Main, 1972). This physiological trait makes such amphibians good indicator species, a proverbial canary in the mine as they are often the first species in an ecosystem to show signs of a change in the environment, such as the introduction of a pollutant, a decrease in dissolved oxygen, or the emergence of a pathogen. Amphibians play a vital ecological role as secondary consumers. Adult amphibians consume great quantities of flies and insects, including mosquitos and their larvae (Mohneke and Rodel, 2009). For example, a single adult of certain species of cricket frogs can consume approximately 4,800 small insects per year (Johnson and Christiansen, 1976). Many of the insect species consumed are vectors for human diseases such as malaria, yellow fever and West Nile virus (Mohneke and Rodel, 2009). 12

13 In addition tadpoles provide ecological services in the freshwater environment by consuming algae. A number of tadpole exclusion experiments have been performed to determine the impact that tadpoles have on their aquatic environment. These experiments showed that a loss of tadpoles lead to rapid degradation of aquatic environments, resulting in uncontrolled algal biomass, increased water turbidity, and an increase in the accumulation of organic and inorganic sediments (Mohneke and Rodel, 2009). Additionally, amphibians are an important prey item for many other species. For example; frog eggs are consumed by wasps and spiders; shrimp, fish and dragonfly nymphs eat tadpoles; and birds, snakes and lizards eat frogs (Mohneke and Rodel, 2009). Removing amphibians from an ecosystem in some cases could have detrimental effects (Mohneke and Rodel, 2009). Amphibian populations globally have been experiencing declines over the past few decades. Approximately 40% of all amphibian species are currently experiencing a decline (Cheng et al., 2011). Many different factors have been hypothesized as causing these population declines. Some of these include habitat destruction and modification, climate change, over-exploitation, pollution, introduced species, increase in ultraviolet light, acid rain and introduced species (Kiesecker et al., 2004; Muths et al., 2008; Pounds et al., 2006). Since the 1990s, mass mortality events have been observed in amphibian populations throughout the world (Berger et al., 1998). Berger et al. (1998) attributed a number of these mortality events to chytrid fungus (Batrachochytrium dendrobatidis) because its symptoms were observed in dying amphibians in Australia 13

14 and Central America. Additionally B. dendrobatidis has been attributed to the collapse of amphibian populations in Panama, California, and Peru (Cheng et al., 2011). This recent emergence of B. dendrobatidis could be due to five forms of anthropogenic amphibian movement including; the pet trade, transportation of zoo animals, food trade, laboratory trade (e.g. Xenopus) and inadvertent or deliberate introduction or release of amphibians (Daszak et al., 2003). For example, more than one million bullfrogs are imported into the United States each year for food trade (Daszak et al., 2003). In this study, I attempt to document the distribution of two pathogens, B. dendrobatidis and Ranavirus spp., in Florida, which have been implicated in declining amphibian populations around the world. Amphibian Anatomy and Life History Amphibians possess distinct anatomy and life history unique among vertebrates. The moist skin of amphibians is an interactive interface between the animal and its environment. It serves as a membrane for the exchange of gases, water, as well as any other materials in the environment. The skin is highly permeable and epidermal sculpturing is very important in water conservation (Duellman and Trueb, 1986). Watersaving mechanisms include the curtailment of water loss through the skin, modifications of the excretory products of the kidneys, and storage of water in vesicles and tissues (Duellman and Trueb, 1986). Purely aquatic species have generally smooth skin, whereas terrestrial and arboreal species may have granular, tough, or even dry skin on the ventral surfaces of the body (Duellman and Trueb, 1986). Irregular ventral surfaces provide increased surface area for absorbing water through substrates. The ventral pelvic region of 14

15 anurans is highly specialized as it is the primary source of water uptake for these organisms (Duellman and Trueb, 1986). This region is often referred to as the drink patch (Daszak et al., 2007). The skin of amphibians consists of two layers, the outer epidermis and the inner dermis. The epidermis is composed of four layers, which include the stratum corneum, stratum granulosum, stratum spinosum, and stratum germinativum (Wells, 2007). In most adult amphibians the stratum cornuem is the location of keratinized skin cells (Wells, 2007). However, amphibians have much less keratinization in their skin when compared to other vertebrates. In most anuran larvae the cells around the mouth become highly keratinized to form a beak or jaw sheath as well as rows of denticles or labial teeth, which aid in consumption of food (Wells, 2007). Amphibians periodically undergo molting cycles where the outer layer of the epidermis, the stratum corneum, sheds and is replaced with a new layer of cells. These molting cycles appear to be affected by temperature, metabolic rate, and photoperiod and are under hormonal control (Wells, 2007). The dermis of amphibians is considerably thicker than the epidermis and consists of two layers, the stratum spongiosum and the stratum compactum (Wells, 2007). Amphibians have a complex life cycle beginning as aquatic organisms and then undergoing metamorphosis to become terrestrial organisms. Some species such as the hellbender (Cryptobranchus alleganiensis), the African clawed frog (Xenopus leavis), and the common mudpuppy (Necturus maculosus) remain aquatic for their entire lives (Vondersaar and Stiffler, 1989). However, since most amphibian species occupy both 15

16 aquatic and terrestrial environments at some point in their lives, they depend on both ecosystems. Batrachochytrium Dendrobatidis Life History One introduced pathogen, chytrid fungus (B. dendrobatidis), can cause chytridiomycosis and has been implicated in declines of many montane amphibian populations (Daszak et al., 2003). Batrachochytrium dendrobatidis can affect the skin cells containing keratin with subsequent infections leading to sloughing of the skin, severe weight loss, hyperkeratosis, mild paralysis and delayed reflexes (Cheng et al., 2011). Batrachochytrium dendrobatidis has low survival rates at temperatures above 25 C and its growth stops above 28 C (Piotrowski et al., 2004). Optimal growth occurs between C, but growth may occur at temperature as low as 4 C (Piotrowski et al., 2004). Batrachochytrium dendrobatidis is able to grow and reproduce at ph 4-8, but growth is optimal at ph 6-7 (Piotrowski et al., 2004). Batrachochytrium dendrobatidis has fairly high survival rates in the environment yet it is extremely sensitive to desiccation (Gleason et al., 2007). Batrachochytrium dendrobatidis has three life stages. The first stage is an aquatic, motile infectious stage (zoospore). The second stage is a parasitic stage where the fungus is found in the skin of adult amphibians (thallus). The third stage occurs when infected individuals discharge new zoospores into the environment (zoosporangia) (Figure 1-1). Batrachochytrium dendrobatidis is attracted to keratinized tissue resulting in thickening (i.e., hyperplasia which is an increase in number of cells and hyperkeratosis which is a thickening of the stratum corneum) of the stratum corneum (Daszak et al., 2007; Johnson and Speare, 2003) in both newly metamorphosed 16

17 individuals and adults as well as mouthparts of larvae (Daszak et al., 2007). Infection of an individual animal occurs when mobile zoospores land on the skin or mouthparts and encyst (Rollins-Smith et al., 2011). The pathogen then moves from the surface of the skin to the stratum granulosum of the epidermis. It then matures in the stratum corneum where it enters healthy cells and grows into a zoosporangium within which zoospores develop (Rollins-Smith et al., 2011). Eventually infected skin cells move toward the surface, the zoosporangium matures, a discharge papilla (pore) opens, and mature zoospores swim out (Rollins-Smith et al., 2011). Batrachochytrium dendrobatidis is difficult to diagnose in larvae, because it is an aclinical infection. However, it can affect mouthparts, growth rates and other developmental features of larvae (Daszak et al., 2003; Parris and Baud, 2004; Daszak et al., 2007). Current accounts show that many larvae are unaffected by infection of B. dendrobatidis. Mortality however typically occurs following the metamorphosis of infected individuals (Mitchell et al., 2008). Signs of infection may or may not be present, and many times B. dendrobatidis is not obvious until an animal is close to death (Daszak et al., 2007). Symptoms of infection may include the animal becoming lethargic, increased shedding of skin (especially on feet and ventral surface in adults) and sitting in a posture where the hind legs and drink patch are elevated (Daszak et al., 2007). Other symptoms may include neurological signs such as abnormal sitting posture with hind legs adducted, lethargy, and slow response to tactile stimuli, Red Leg Syndrome-like skin discoloration, and in salamanders black spots on the skin and loss of the tail and toe tips (Daszak et al., 2007). 17

18 Laboratory experiments indicate that B. dendrobatidis grows most rapidly, with a 4 5 day generation time, at cool temperatures (17 C 25 C) (Piotrowski et al., 2004). This suggests that its physiology relating to environmental conditions (i.e., ambient air temperature changes) may alter how it interacts with its amphibian host (Woodhams et al., 2003). Because of the lack of many obvious clinical signs, the presence of B. dendrobatidis must often be confirmed with microscopy (e.g., skin histopathology of the feet or groin) or polymerase-chain reaction (PCR) (Berger et al., 1999; Daszak et al., 2007). Chytridiomycosis outbreaks typically spread by a combination of frog-to-frog and environment-to-frog transmission (Lips et al., 2006). As prevalence increases within the community, infected amphibians shed zoospores into the environment and/or directly pass them to other amphibians by contact (Lips et al., 2006). Daszak et al. (2007) hypothesized that saturation of the environment with zoospores by the long-term persistence of zoospores in the environment coupled with infectivity of amphibians produces the observed infection pattern where prevalence quickly changes from very low to very high. This is followed by widespread mortality and also yields a mechanism for the most significant impact of chytridiomycosis: the extirpation of frog populations from an entire region (Daszak et al., 2007). Some amphibians are known to carry and tolerate B. dendrobatidis infections. These species may be asymptomatic, live in the community, and act as vectors. Among them are the American Bullfrog (Lithobates catesbeianus), Cane Toad (Rhinella marina), Xenopus spp., Eastern Tiger Salamander (Ambystoma tigrinum) and Cricket Frog (Acris gryllus) (Daszak et al., 2007). All of these species have been introduced 18

19 globally (Collins et al., 1988; Kraus, 1999; Mazzoni et al., 2003; Hanselmann et al., 2004; Weldon et al., 2004). Treatment of amphibians infected with chytridiomycosis in captivity has been successful. The most common treatment used to eradicate chytridiomycosis in a single individual is a bath of 0.01% itraconazole applied for five minutes daily for 11 days (Pessier, 2008). The bath can be prepared by diluting 1% itraconazole oral solution with 0.6% saline or Amphibian Ringer s solution (Pessier, 2008). This treatment protocol should not be used on tadpoles or recent metamorphs as death may occur (Pessier, 2008). Multiple treatment cycles 1 to 2 weeks apart might be necessary for some species (Pessier, 2008). After treatment the individual can be released, however, infection may reoccur from frog-to-frog or frog-to-environment exposure. Ranavirus spp. Life History Another pathogen that has severely impacted amphibian populations are three viral species grouped together as Ranavirus spp. Ranavirus is one of five genera of viruses within the family Iridoviridae, which is one of the five families of nucleocytoplasmic large family DNA viruses. Ranavirus is the only genus within this family that includes viruses that are infectious to amphibians and reptiles (Teacher et al., 2010). Ranaviruses were first isolated from Northern Leopard Frogs (L. pipiens) in the mid-1960s (Gray et al., 2009). Amphibians infected with Ranavirus spp. experience hemorrhaging, reddening of the belly, legs, and vent, lethargy, emaciation, edema, bloat, skin ulcers, and in many cases death (Teacher et al., 2010). There are two disease syndromes of Ranavirus spp. in amphibians. The first is ulcerative skin syndrome, characterized by dermal ulceration. The second is hemorrhaging syndrome, which is characterized by systematic 19

20 hemorrhaging within the skeletal muscles and visceral organs (Teacher et al., 2010). Infection is most easily observed in the kidneys and liver of amphibians as seen in Figure 1-2. Green et al. (2002) observed over 90% mortality in 25 Ranavirus spp. amphibian mortality events, with larvae being the most susceptible to mortality by this virus (Gray et al., 2007). It is thought that some individuals are able to recover from Ranavirus spp. infections because scars characteristic of healed skin ulcers have been found on individual amphibians with current infections (Teacher et al., 2010). Ranavirus spp. associated mortalities have been reported on five continents, at varying latitudes and elevations. These mortalities have occurred in most of the major families within Anura and Caudata (Gray et al., 2009). Mass mortalities of amphibians from Ranaviruses have been reported in the Americas, Europe, and Asia (Gray et al., 2009). A number of amphibian die-offs in cultured frogs in China and Thailand have been linked to a Ranavirus which appears to be related to, if not identical to Frog Virus 3 (Chinchar, 2002). Ranavirus spp. is also thought to be the cause of a Sonora Tiger Salamander (A. mavoritium stebbinsi) die-off that occurred in western North America in 1985 (Chinchar, 2002). Approximately one to three U.S. new states report Ranavirus spp. die-offs each year (Gray et al., 2009). It is difficult to quantify the extent of Ranavirus spp. mortality events, especially if they occur in common species. Many mortality events, therefore, most likely go unnoticed (Gray et al., 2009). Not much attention had been paid to this genus of viruses as it does not infect mammals or birds. However, recent outbreaks (since ca. the mid 1980s) in commercial and recreational fish species, cultured and wild frogs, and endangered salamanders, 20

21 has brought increased attention to the genus (Chinchar, 2002). Three species of Ranavirus are known to infect amphibians. They are Ambystoma tigrinum virus (ATV), Bohle iridovirus (BIV) and Frog Virus 3 (FV3) (Chinchar et al., 2005). Detection of Ranavirus spp. in infection is done using Polymerase-Chain Reaction (PCR). The PCR primers used for the samples in this study have the ability to detect all three major groups of Ranaviruses. Each major group most likely contains numerous different Ranavirus species that can be difficult to differentiate without additional analyses. One major Ranavirus group commonly found with these PCR primers is most closely related to the type of virus for the genus Ranavirus (Frog Virus 3), but based on PCR and sequencing products, one most often gets results that indicate a Frog Virus 3-like viral infection (A. Pessier, personal communication). Ranavirus spp. isolates from the United Kingdom have been shown to grow in vitro between 8 C and 30 C, with the fastest replication occurring at 30 C and slower replication occurring at 1 C (Teacher et al., 2010). Frog Virus 3 replicates at temperatures ranging from C, making it able to survive and reproduce at a fairly broad range of temperatures (Chinchar, 2002). Transmission of Ranavirus is thought to occur by multiple pathways, including contaminated soil, direct contact, waterborne exposure, as well as ingestion of infected tissue during predation, necrophagy or cannibalism (Gray et al., 2009). Ranaviruses are relatively stable in aquatic environments, persisting for several weeks or longer outside a host organism (Brunner et al., 2004). Duffus et al. (2008) conducted a study on the transmission of Frog Virus 3-like infections in aquatic amphibian communities. It was found that both vertical 21

22 (reproductive-dependent) and horizontal (non-reproductive-dependent) transmission could be possible, although the latter is the most likely mode of transmission (Duffus et al., 2008). It was found that salamanders are likely both hosts and reservoirs of this pathogen. The movement of salamanders from one pond to another could potentially spread the Frog Virus 3-like pathogen (Duffus et al., 2008). Amphibian Declines in the State of Florida A number of amphibian species in the state of Florida have been experiencing population declines and in some cases, extirpation from historical ranges. Some species that have experienced major declines include the striped newt (Notophthalmus perstriatus) and the southern dusky salamander (Desmognathus auriculatus) (Means et al., 2008 and Means & Travis, 2007). In July of 2008 Means et al. (2008) petitioned the U.S. Fish and Wildlife Service to list the striped newt as a Federally Threatened species under the Endangered Species Act of This was due to a number of factors, which have been, and currently are impacting this species. These factors include present and future modification or destruction of habitat/range, over-exploitation for commercial, scientific, or educational purposes, disease or predation, lack of existing regulatory mechanisms, and any other natural or unnatural forces affecting the continued existence of this species (Means et al., 2008). Southern dusky salamander (D. auriculatus) populations have disappeared over the past few decades at several localities in Florida. Some of these areas that historically had D. auriculatus populations include Deep Springs Canyon in Bay County, Lightwood Knot Creek, Garnier Creek, Tom s Creek on Eglin Air Force Base in Okaloosa county, and Devil s Millhopper in Alachua County (D. B. Means, personal 22

23 communication). No cause for these extirpations has been discovered (Means and Travis, 2007). Current Accounts of Amphibian Disease in Florida Surveys of amphibian populations in Florida from did not detect B. dendrobatidis (K. M. Enge, personal communication). After 2007, however, amphibians infected with B. dendrobatidis have been confirmed in four localities in central Florida. Batrachochytrium dendrobatidis was first detected in 2008 in a bullfrog (L. catesbeianus) at the Central Florida Zoo and Botanical Gardens, Seminole County (J. L. Stabile, personal communication). Rizkalla (2009) detected B. dendrobatidis in both L. catesbeianus and Acris gryllus at both the Walt Disney Animal Kingdom and the Disney Wildlife Management and Conservation Area, Orange County, respectively. In May 2009, Kevin M. Enge (personal communication) collected three Gopher Frog (L. capito) tadpoles from Ocala National Forest, Putnam County, which tested positive for B. dendrobatidis (St-Amour et al., 2010). Nevertheless, a comprehensive survey of the occurrence of B. dendrobatidis and Ranavirus spp. in Florida has yet to be performed. Determining the geographic distribution of these pathogens in northern Florida is a critical first step in assessing it the impact that these pathogens are having on the state s amphibian populations. Objectives and Significance of Current Study The objectives of this project is to ascertain the current infection status, by determining the geographic, taxonomic, and temporal (seasonality and life stage) distribution of B. dendrobatidis and Ranavirus spp. in sampled Florida amphibians. The results will illustrate the geographic distribution of these pathogens, and enable the tracking of the spread of these diseases. This is important for the protection and 23

24 conservation of threatened species. The results of this study will provide guidance to managers and conservationists evaluating the threat that B. dendrobatidis and Ranavirus spp. pose to the amphibian populations of northern peninsular and panhandle Florida. This study will benefit a number of groups by providing initial data for researchers, land managers, state and federal conservancy programs, and the public. The methods and results can inform future research and help other amphibian conservation groups, not only in Florida but in other areas where B. dendrobatidis or Ranavirus spp. has been detected in amphibian populations. Once the distribution of B. dendrobatidis and Ranavirus spp. in northern peninsular and panhandle Florida has been determined, then further studies can be conducted in order to determine if B. dendrobatidis, Ranavirus spp., or a combination of the two are causing amphibian population declines recently observed in Florida and other sites worldwide. This documentation of the distribution of B. dendrobatidis and Ranavirus spp. can assist state wildlife agencies in implementing the appropriate conservation measures. Overview of Study Area The study area included the northern peninsula and panhandle of Florida. There are 28 major drainages in this region. These primary drainages include: Escambia, Perdido, Blackwater, East Bay, Yellow, Choctawhatchee, Choctawhatchee Bay, St. Andrews Bay, Apalachicola, Whiskey George Creek, Crooked, Ochlockonee, St. Marks, Aveilla, Upper Suwannee, Lower Suwannee, St. Marys, Ecofina, Fenholloway, Spring Warrior Creek, Blue Creek, Steinhatchee, Amason Creek, California Creek, Waccasassa, Withcacoochee, Nassau, and St. Johns (Robert H. Robins, personal 24

25 communication). Many of the ponds in my study area were ephemeral and are therefore not connected to any of these drainages. Samples were collected from 7 sites, including, Apalachicola National Forest, Gold Head Branch State Park, Camp Blanding Military Reserve, Ocala National Forest-South of Salt Springs, Ocala National Forest-South of Church Lake, Etoniah Creek State Forest and Osceola National Forest (Figures 1-3 to 1-9). All sites were located on land owned by the state or federal government and included habitats of upland sandhills, grassy ponds, cypress gum ponds in flatwoods, borrow pits, sandhill/scrub, flatwoods scrub ecotone, reclaimed mine land (formerly scrub), sinkholes, sandpine scrub/flatwoods and long leaf/slash pine flatwoods (Figure 1-10). 25

26 Figure 1-1. The life cycle of the chytrid fungus, Batrachochytrium dendrobatidis, illustrating substrate-dependent and substrate-independent stages. (with permission from Rosenblum et al., 2008) 26

27 Figure 1-2. Photomicrograph of a stained kidney from a green frog (Lithobates clamitans) larva infected with Ranavirus spp. (with permission from Miller et al., 2009) 27

28 Figure 1-3. Representative pond (Pond 3) within Apalachicola National Forest, Liberty County, Florida, USA. Photo by Kevin M. Enge 28

29 Figure 1-4. Representative pond (Pebble Lake) within Gold Head Branch State Park, Clay County, Florida, USA. Photo by Sarah Reintjes-Tolen 29

30 Figure 1-5. Representative pond (Pond 3) within Camp Blanding Military Reserve, Clay County, Florida, USA. Photo by Sarah Reintjes-Tolen 30

31 Figure 1-6. Representative pond (Pond 1) within Osceola National Forest, Baker County, Florida, USA. Photo by Sarah Reintjes-Tolen 31

32 Figure 1-7. Representative pond (Pond 21) within Ocala National Forest-South of Salt Springs, Marion County, Florida, USA. Photo by Sarah Reintjes-Tolen 32

33 Figure 1-8. Representative pond (Pond 1) within Ocala National Forest-South of Church Lake, Marion County, Florida, USA. Photo by Sarah Reintjes-Tolen 33

34 Figure 1-9. Representative pond (Pond 7) within Etoniah Creek State Forest, Putnam County, Florida, USA. Photo by Sarah Reintjes-Tolen 34

35 Figure Map illustrating sampling sites where amphibians were swabbed for Batrachochytrium dendrobatidis and/or Ranavirus spp., in Florida, USA 35

36 CHAPTER 2 GEOGRAPHIC DISTRIBUTION OF CHYTRID FUNGUS (BATRACHOCHYTRIUM DENDROBATIDIS) AND RANAVIRUS SPP. IN NORTHERN PENINSULAR AND PANHANDLE FLORIDA AMPHIBIANS Introduction In order to determine the geographic distribution of these two amphibian pathogens, amphibians from seven sites in northern peninsular and panhandle Florida were sampled for Batrachochytrium dendrobatidis and Ranavirus spp. This survey was the first of its kind in the state of Florida. An amphibian being infected with B. dendrobatidis was not recorded in the state until 2008 (J. L. Stabile, personal communication). Since then amphibians infected with this fungus have been recorded at four localities, including the Central Florida Zoo and Botanical Gardens, Seminole County (J. L. Stabile, personal communication), both the Walt Disney Animal Kingdom and its Wildlife Management and Conservation Area, Orange County (Rizkalla, 2009) and in Ocala National Forest, Putnam County (K. M. Enge, personal communication). Due to this apparent recent emergence of infected amphibians, surveys were conducted with the help of partners including the University of Florida, Florida Fish and Wildlife Conservation Commission and Central Florida Zoological Gardens. These organizations decided to fund a survey of both B. dendrobatidis and Ranavirus spp. in order to determine geographic distribution in Florida and to determine the level of threat to local populations. Methods A total of 32 ponds and 296 individuals, across nine species, were sampled. Field surveys were conducted between the months of November and April, in which ambient air temperatures allowed for optimal growth and survival of B. dendrobatidis. This time 36

37 period also coincided with the winter amphibian breeding season in Florida. Skerratt et al. (2008) noted that there are a number of key aspects to consider when surveying amphibian populations across large areas, such as area, species, life stage, sample size, planning a surveying strategy, season, and diagnostic tests. Sampling Sites I attempted to sample a minimum of 30 individual amphibians of a variety of species at each of the seven sites. Within each site amphibians were sampled from a minimum of three separate bodies of water. This was done in order to obtain samples from a variety of habitats with a number of different species and increase the likelihood of detecting B. dendrobatidis and/or Ranavirus spp. Due to low water levels I was only able to obtain samples from one pond in Gold Head Branch State Park. At each site, except for those within Apalachicola National Forest, one to ten individuals of the same species were pooled (i.e., sampled with the same swab) in order to increase the likelihood of detecting either B. dendrobatidis and/or Ranavirus spp. This was done because these populations had not been assessed for either of these pathogens. Species Sampled At each site, targeted species included common species as well as species of greatest conservation need (K. M. Enge-FWC, personal communication). Species of greatest conservation need included the Tiger Salamander (Ambystoma tigrinum), Reticulated Flatwoods Salamander and Frosted Flatwoods Salamander (A. bishopi and A. cingulatum), Striped Newt (Notophthalmus perstriatus), Ornate Chorus Frog (Pseudacris ornata) and Gopher Frog (Lithobates capito). These species are considered species of greatest conservation need because of their declining or low 37

38 population levels in Florida (K. M. Enge, personal communication, Means and Travis, 2007, Means et al., 2008). Common species that were sampled included the Southern Leopard Frog (L. sphenocephalus), Bullfrog (L. catesbeianus), and Southern Cricket Frog (Acris gryllus). Adult A. gryllus and L. catesbeianus are known to be carriers of B. dendrobatidis which made these species a sampling priority (Daszak et al., 2007). Some of Florida s amphibian species that have been found to be infected with B. dendrobatidis include the Northern Cricket Frog, A. crepitans (Pessier et al., 1999; Steiner and Lehtinen, 2008); Fowler s Toad, Anaxyura fowleri tadpoles (Parris and Cornelius, 2004); Gray Treefrog, Hyla chrysoscelis/versicolor and its tadpoles (Ouellet et al., 2005; Parris and Baud, 2004; Parris and Cornelius, 2004); Bullfrog, L. catesbeianus (Mitchell and Green, 2002; Daszak et al., 2004; Ouellet et al., 2005; Pearl and Green, 2005; Longcore et al., 2007); Green Frog, L. clamitans (Longcore et al., 2007; Ouellet et al., 2005); and Southern Leopard Frog, L. sphenocephalus ( Mitchell and Green, 2002; Ouellet et al., 2005). These species were given priority as part of the 30 miscellaneous individuals that were sampled at each site. Amphibian Sampling Amphibians were collected using 0.3mm mesh size dip nets. Additionally other specimens were caught by hand. Once captured, each amphibian was placed in a separate plastic bag to reduce the chance of transmission of B. dendrobatidis or Ranavirus spp. among individuals. Each individual amphibian was sampled for B. dendrobatidis and/or Ranavirus spp. and the following measurements were taken; snout to vent length (SVL) to the nearest mm was recorded using Mitutoyo brand calipers; mass to the nearest 0.1 g was recorded using an Ohaus HH 320 hand-held scale (0.1-38

39 320 g); and any atypical physical characteristics such as lesions, deformities, swollen limbs or abdomen, missing digits, and jaw deformation were recorded (Figure 2-1) (Daszak et al., 2007; Teacher et al., 2010). Different protocols were followed in order to test for B. dendrobatidis and Ranavirus spp., following the procedures of Hyatt et al. (2006). The following materials were needed to swab amphibians: 70% ethanol, cotton tipped applicators (swabs), 2ml vials with self-sealing screw caps, nitrile gloves, alcohol sanitizer for hands, water- and alcohol-proof pens, waterproof notebook, waste bag, collection bags, closeable bags (Zip-Loc recommended), vial storage containers, and bleach solution (Figure 2-2) (Brem et al., 2007). Swabbing for Batrachochytrium Dendrobatidis To test for B. dendrobatidis on an adult amphibian, a sterile cotton swab was swept five times on each of the ventral surface of the thighs, abdomen and feet (25 total sweeps per animal). These areas commonly have a high concentration of B. dendrobatidis sporangia (Brem et al., 2007). To properly swab each area of adult amphibians it is best to grip the animal mid-dorsally, anterior to the hind legs with the ventral side up. For extremely small individuals or salamanders the position or technique may need to be altered slightly in order obtain an adequate swab (Figure 2-3) (Brem et al., 2007). To test for B. dendrobatidis in larvae, the pharyngeal mucosa was swabbed. Then, the bare end of the cotton swab was stuck into the ground in a sunny area to dry for five minutes. After drying, the cotton tip was broken off and placed in an empty vial and labeled. Between working up individual animals gloves were carefully changed to prevent cross contamination between individuals and/or samples. 39

40 Swabbing for Ranavirus spp. To test for Ranavirus spp. in adult as well as tadpole or larval amphibians, the pharyngeal mucosa was swabbed with a cotton tip (Figure 2-4). The cotton tip was then broken off and placed in a labeled vial filled with 70% ethanol. Gloves were worn at all times and were switched out between each individual to prevent the spread of any pathogens and to prevent cross-contamination of samples. The optimal way to test for the presence of Ranavirus spp. in adult amphibians is to euthanize the animal and perform a necropsy (A. P. Pessier, personal communication). The kidneys and liver are removed and placed in a labeled vial with 70% ethanol, then PCR analysis is used for a more accurate detection of Ranavirus spp. (A. P. Pessier, personal communication). Environmental Factors After the proper number of individuals were collected and processed at each sampling location; the habitat type, latitude and longitude, and a number of abiotic environmental factors were recorded. Abiotic factors included ambient air, soil, and water temperatures, ph, dissolved oxygen, and specific conductivity (Tables 2-1 and 2-2). A Garmin GPS 76Cx was used to obtain the latitude and longitude at each site. A Hannah Instruments HI 9142 dissolved oxygen meter was used to obtain dissolved oxygen measurements in units of milligram per liter. An EcoTestr ph2 brand meter was used to measure ph. A RadioShack digital indoor/outdoor thermometer was used to measure air and soil temperature in degrees Celsius. Water temperatures in degrees Celsius were obtained by a Traceable brand lollipop thermometer. All temperatures 40

41 were taken to the nearest.01 C. A Hannah Instruments Watercheck ph and TDS meter was used to measure specific conductivity in siemens per meter. Disinfection All equipment (waders, boot, nets, etc.) was soaked in a 10% bleach solution for one to two minutes between successive ponds in order to prevent the spread of B. dendrobatidis and Ranavirus spp. (Phillott et al., 2010). We attempted to minimize the amount of stress each individual amphibian experienced during the capture and swabbing process as stressed animals are at a greater risk of infection. Additionally great care was used when handling larvae as they are vulnerable to skin damage from traumatic handling (Phillott et al., 2010). Wounds could make larvae more susceptible to infection from pathogens. Pathogen Detection The most effective method of detection involves euthanizing the animal and then performing a necropsy to remove the liver and kidneys. Tissue samples from these organs are then analyzed using either a histological or PCR analysis to determine the presence or absence of the pathogen (A. P. Pessier, personal communication). The PCR assay used to detect the presence of B. dendrobatidis is extremely sensitive as it is able to detect a single zoospore (Brem et al., 2007). As long as proper decontamination protocol is followed to eliminate the chance of cross contamination, the PCR assay is unlikely to produce a false-positive. A false-negative (where the animal is infected but the result comes back as negative) can occur if the animal has a very light infection, making it difficult to detect the presence of spores. Sampling multiple areas of the body that are the most common sites of B. dendrobatidis infection, such as ventral 41

42 surface and toes of adults and the pharyngeal mucosa of tadpoles and metamorphs, reduced the chances of a false-negative (Brem et al., 2007). Results A total of 296 individual amphibians were sampled for B. dendrobatidis, including nine species at 32 different localities. Some individuals were sampled for both B. dendrobatidis and Ranavirus spp., while others were only sampled for B. dendrobatidis. Samples of Ranavirus spp. were limited by the fact that Ranavirus spp. is difficult to detect from pharyngeal mucosa swabs. This resulted in slightly fewer Ranavirus spp. samples in the study. Two swabs from Southern Cricket Frogs came back positive for B. dendrobatidis. These individuals were caught in Camp Blanding Military Reserve (five individuals) and Osceola National Forest (seven individuals). These were all adult Southern Cricket Frogs that were asymptomatic and behaved normally at the time of collection (Figure 2-5). Two swabs, one from six Bullfrogs the other from four Leopard Frog tadpoles gave positive results for Ranavirus spp. Both of these swabs were from individuals from Pebble Lake in Gold Head Branch State Park (Figure 2-5). When trying to determine the possible infection rate of B. dendrobatidis there are three scenarios, due to the fact that swabs were pooled (one swab used to swab multiple individuals). The first scenario assumes that there were only two positive individuals out of the total number of individuals sampled, 296. This would give an infection rate of 0.68%, which is the lowest possible infection rate. The next possible scenario simply takes into account the two positive swabs out of a total of 103 swabs. This gives an infection rate of 1.8%. The highest possible infection rate assumes that all 42

43 of the 12 individuals sampled with the two swabs that came back positive for B. dendrobatidis were infected with the pathogen. This scenario gives an infection rate of 4.0%. When trying to determine the possible infection rate of Ranavirus spp there are three scenarios, due to the fact that swabs were pooled (one swab swabbed multiple individuals). The first scenario is the lowest possible infection rate, assuming that there were only two positive individuals out of the total number of individuals sampled, 250. This would give an infection rate of 0.8%. The next possible scenario simply takes into account the two positive swabs out of a total of 96 swabs. This gives an infection rate of 2.1%. The third scenario assumes that all of the 10 individuals sampled with the two swabs that came back positive for Ranavirus spp. were infected with the pathogen. This gives an infection rate of 4.0% All of the environmental factors measured came back within normal ranges (Table 2.1). This suggests that none of the infected individuals were directly compromised by existing environmental conditions. Discussion My results show a very low frequency of the pathogens examined however, based on the scale of my study and considering all of the available waterways in northern peninsular and panhandle Florida, these results are most likely an underestimate of the frequency of these pathogens. Nonetheless, my study does show that these pathogens are present in Florida. This means that there is the possibility of infection and extirpation of vulnerable populations of amphibians. Based on the infection rates found in this study it appears that there is a low background infection rate for B. dendrobatidis with individuals that are simply carriers 43

44 and appear healthy. For Ranavirus spp. it appears that the pathogen is either present or not present. If it is present then there is a very high infection rate and therefore a high mortality rate. This was observed at Pebble Lake in Gold Head Branch State Park, where a large-scale mortality event was observed and samples testing positive for Ranavirus spp. were obtained from dead and moribund tadpoles. Despite the fact that neither B. dendrobatidis nor Ranavirus spp. had been detected in Florida until 2008 and 2011 does not mean that these pathogens were not present in Florida before this time. These pathogens could have been present for an unknown amount of time and not sampled, or if they were sampled, not detected. Additionally, due to the short time period during which amphibians were impacted by Ranavirus spp. at Pebble Lake in Gold Head Branch State Park it is likely that many more events are taking place and going undetected. The small scale of my study along with the small window of time available to detect a mortality event means that the threats these pathogens pose to Florida amphibians remains unknown. My project is a preliminary study of the occurrence of two amphibian diseases in the state of Florida. The small scale of this study warrants caution in drawing conclusions using these data: because the diseases were found in a few localities does mean they are not more widespread in the state. The data show the importance of doing a large-scale study designed to understand the distribution of these pathogens more thoroughly. Knowledge of these initial findings and of indicators of B. dendrobatidis and Ranavirus spp. infections/die-offs can be useful to park and forest officials, Florida Fish and Wildlife Conservation Commission biologists These diseases could certainly 44

45 become more of a problem in the future as amphibians become more vulnerable due to a combination of climate change, habitat loss/fragmentation and increased exposure to ultraviolet light. (Daszak et al., 2003) Due to La Niña years in (NOAA) that resulted in severe droughts throughout the state of Florida, many species of greatest conservation need were not known to breed during the winter reproductive season. This lack of breeding was evident when surveys were conducted across many parts of northern and peninsular Florida. During these surveys an unusually large percentage of ephemeral ponds were dry (K. M. Enge, personal communication). Most species of greatest conservation need rely on ephemeral ponds to breed. Because of these weather conditions the majority of species sampled in my study were common species. Frogs and salamanders are known to migrate considerable distances into upland habitats surrounding breeding sites. However, during times of little rainfall this migration may not occur as the energy expenditure is not worth it (Blihovde, 2006). Blihovde (2006) noted that drought contributes to a sedentary behavior in the Gopher Frog. After successive years of drought, ponds in central Florida held very little water, which may have led to an increase in the level of site fidelity that the frogs exhibited (Blihovde, 2006). Rainfall has considerable effects on the behavior of amphibian species. Lack of moisture may therefore result in very low levels of activity and low reproduction. This site fidelity in combination with drought conditions could lead to density-dependent infection rates as amphibians are forced to congregate in remaining water sources. 45

46 Figure 2-1. Sampling amphibians for Batrachochytrium dendrobatidis and Ranavirus spp. and recording data from individuals caught in Camp Blanding Military Reserve, (Pond 26), Clay County, Florida, USA. Photo by Kevin M. Enge 46

47 Figure 2-2. Sampling a Southern Leopard Frog (Lithobates sphenocephalus) tadpole for Batrachochytrium dendrobatidis and Ranavirus spp at Ocala National Forest- South of Church Lake, (Pond 1), Marion County, Florida, USA. Photo by Kevin M. Enge 47

48 Figure 2-3. Sampling an adult Southern Cricket Frog (Acris gryllus) for Batrachochytrium dendrobatidis at Ocala National Forest-South of Church Lake, (Pond 14), Marion County, Florida, USA. Photo by Kevin M. Enge 48

49 Figure 2-4. Sampling an adult Southern Cricket Frog (Acris gryllus) for Ranavirus spp. at Ocala National Forest-South of Church Lake, (Pond 14), Marion County, Florida, USA. Photo by Kevin M. Enge 49

50 Figure 2-5. Map of sites sampled with negative for Batrachochytrium dendrobatidis and Ranvirus spp. sampling localities in blue, positive for B. dendrobatidis localities in yellow, and positive for Ranvirus spp. in red, mapped using Google Earth 50

51 Number of Individuals larvae adults Species Figure 2-6. Number of individual amphibians, sampled for both Batrachochytrium dendrobatidis and Ranavirus spp., by species and life stage 51

52 Table 2-1. The locality of each pond where amphibians were sampled for Batrachochytrium dendrobatidis and/or Ranvirus spp Location Pond name/number Lat/Long Blackwater SF-002 BSF , Apalachicola NF ANF , Apalachicola NF-001 ANF , Apalachicola NF ANF , Apalachicola NF-002 ANF , Apalachicola NF-003 ANF , Apalachicola NF-004 ANF , Camp Blanding MR-026 CBMR , Camp Blanding MR-004 CBMR , Camp Blanding MR-001 CBMR , Camp Blanding MR-065 CBMR , Camp Blanding MR-063 CMBR , Gold Head Branch SP-001 GHBSP , Gold Head Branch SP-001 GHBSP , Ocala NF-003 ONF SSS , Ocala NF-022 ONF SSS , Ocala NF-006 ONF SSS , Ocala NF-020 ONF SSS , Ocala NF-005 ONF SSS , Ocala NF-021 ONF SSS , Etoniah Creek SF-011 ECSF , Etoniah Creek SF-007 ECSF , Etoniah Creek SF-005 ECSF , Etoniah Creek SF-006 ECSF , Etoniah Creek SF-022 ECSF , Ocala NF-001 ONF SCL , Ocala NF-014 ONF SCL , Ocala NF-017 ONF SCL , Osceola NF-001 OsNF , Osceloa NF-002 OsNF , Osceloa NF-003 OsNF , Osceola NF-004 OsNF , Big Bend WMA-002 BBSCU ,

53 Table 2-2. Summary table of data collected on chytrid fungus, Batrachochytrium dendrobatidis, in northern peninsular and panhandle Florida Site Latitude/Longitude Date Species sampled Chytrid (positive/negative) Blackwater State Forest , /8/11 Lithobates sphenocephalus 0/1 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Ambystoma cingulatum 0/10 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Lithobates sphenocephalus 0/5 Apalachicola National Forest , /9/11 Lithobates sphenocephalus 0/5 Gold Head Branch State Park , /7/11 Acris gryllus 0/5 Gold Head Branch State Park , /7/11 Lithobates sphenocephalus 0/9 Gold Head Branch State Park , /7/11 Lithobates catesbeianus 0/9 Camp Blanding Military Reserve , /7/11 Acris gryllus 0/5 Camp Blanding Military Reserve , /7/11 Lithobates catesbeianus 0/1 Camp Blanding Military Reserve , /7/11 Notophthalmus perstriatus 0/2 Camp Blanding Military Reserve , /7/11 Lithobates capito 0/6 Camp Blanding Military Reserve 29.88, /7/11 Lithobates sphenocephalus 0/5 Camp Blanding Military Reserve 29.88, /7/11 Acris gryllus 0/5 Camp Blanding Military Reserve 29.88, /7/11 Lithobates capito 0/3 Camp Blanding Military Reserve , /7/11 Lithobates sphenocephalus 0/5 Camp Blanding Military Reserve , /7/11 Acris gryllus (5/5) Gold Head Branch State Park , /25/11 Lithobates capito 0/3 Camp Blanding Military Reserve , /25/11 Lithobates sphenocephalus 0/5 Camp Blanding Military Reserve , /25/11 Acris gryllus 0/5 Camp Blanding Military Reserve , /25/11 Acris gryllus 0/5 Camp Blanding Military Reserve , /25/11 Lithobates sphenocephalus 0/5 Big Bend WMA , /2/11 Lithobates sphenocephalus 0/1 Osceola National Forest , /4/11 Acris gryllus 0/2 Ocala National Forest , /11/11 Acris gryllus 0/10 Ocala National Forest , /11/11 Lithobates catesbeianus 0/3 Ocala National Forest , /11/11 Acris gryllus 0/10 Ocala National Forest , /11/11 Notophthalmus perstriatus 0/2 Ocala National Forest , /11/11 Lithobates catesbeianus 0/1 Ocala National Forest , /11/11 Lithobates sphenocephalus 0/4 Ocala National Forest , /11/11 Lithobates catesbeianus 0/4 Ocala National Forest , /11/11 Acris gryllus 0/5 Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 53

54 Table 2-2. Continued Site Latitude/Longitude Date Species sampled Chytrid (positive/negative) Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 Etoniah Creek State Forest , /18/11 Lithobates sphenocephalus 0/1 Etoniah Creek State Forest , /18/11 Lithobates capito 0/3 Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 Ocala National Forest , /23/12 Acris gryllus 0/6 Ocala National Forest , /23/12 Lithobates sphenocephalus 0/5 Ocala National Forest , /23/12 Lithobates capito 0/4 Ocala National Forest , /23/12 Acris gryllus 0/10 Ocala National Forest , /23/12 Lithobates sphenocephalus 0/1 Ocala National Forest , /23/12 Acris gryllus 0/8 Osceola National Forest , /5/12 Lithobates sphenocephalus 0/2 Osceola National Forest , /5/12 Acris gryllus 0/4 Osceola National Forest , /5/12 Acris gryllus 0/11 Osceola National Forest , /5/12 Lithobates heckscheri 0/1 Osceola National Forest , /5/12 Acris gryllus (7/7) 54

55 Table 2-3. Summary table of data collected on Ranavirus spp., in northern peninsular and panhandle Florida Site Latitude/Longitude Date Species sampled Ranavirus spp. (positive/negative) Blackwater State Forest , /8/11 Lithobates sphenocephalus 0/6 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Ambystoma cingulatum 0/10 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Pseudacris ornata 0/5 Apalachicola National Forest , /9/11 Lithobates sphenocephalus 0/5 Gold Head Branch State Park , /7/11 Lithobates sphenocephalus (4/4) Gold Head Branch State Park , /7/11 Lithobates catesbeianus (6/6) Gold Head Branch State Park , /25/11 Lithobates capito 0/3 Ocala National Forest , /11/11 Acris gryllus 0/10 Ocala National Forest , /11/11 Lithobates catesbeianus 0/3 Ocala National Forest , /11/11 Acris gryllus 0/10 Ocala National Forest , /11/11 Notophthalmus perstriatus 0/2 Ocala National Forest , /11/11 Lithobates catesbeianus 0/1 Ocala National Forest , /11/11 Lithobates sphenocephalus 0/4 Ocala National Forest , /11/11 Lithobates catesbeianus 0/4 Ocala National Forest , /11/11 Acris gryllus 0/5 Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 Etoniah Creek State Forest , /18/11 Lithobates sphenocephalus 0/1 Etoniah Creek State Forest , /18/11 Lithobates capito 0/3 Etoniah Creek State Forest , /18/11 Acris gryllus 0/10 Ocala National Forest , /23/12 Acris gryllus 0/6 Ocala National Forest , /23/12 Lithobates sphenocephalus 0/5 Ocala National Forest , /23/12 Lithobates capito 0/4 Ocala National Forest , /23/12 Acris gryllus 0/10 Ocala National Forest , /23/12 Lithobates sphenocephalus 0/1 Ocala National Forest , /23/12 Acris gryllus 0/8 Osceola National Forest , /5/12 Lithobates sphenocephalus 0/2 Osceola National Forest , /5/12 Acris gryllus 0/4 55

56 Table 2-3. Continued Site Latitude/Longitude Date Species sampled Ranavirus spp. (positive/negative) Osceola National Forest , /5/12 Acris gryllus 0/11 Osceola National Forest , /5/12 Lithobates heckscheri 0/1 Osceola National Forest , /5/12 Acris gryllus 0/7 56

57 Table 2-4. Environmental data collected at each pond where amphibians were sampled for Batrachochytrium dendrobatidis and/or Ranvirus spp Location Habitat type Water temp Air temp Soil temp ph DO Conductivity collection date Blackwater SF-002 grassy pond / / / / / / 3/8/11 Apalachicola NF cypress gum pond/flatwoods / / / / / / 3/9/11 Apalachicola NF-001 cypress gum pond/flatwoods / / / / / / 3/9/11 Apalachicola NF cypress gum pond/flatwoods / / / / / / 3/9/11 Apalachicola NF-002 borrow pit / / / / / / 3/9/11 Apalachicola NF-003 ditch / / / / / / 3/9/11 Apalachicola NF-004 borrow pit / / / / / / 3/9/11 Camp Blanding MR-026 borrow pit /7/11 Camp Blanding MR-004 scrub /7/11 Camp Blanding MR-001 scrub /7/11 Camp Blanding MR-065 scrub /25/11 Camp Blanding MR-063 scrub /25/11 Gold Head Branch SP-001 sinkhole pond/upland sandhill /7/11 Gold Head Branch SP-001 sinkhole pond/upland sandhill /25/11 Ocala NF-003 ring pond/sandpine scrub /11/11 Ocala NF-022 sandpine scrub /11/11 Ocala NF-006 sandpine scrub /11/11 Ocala NF-020 sandpine scrub /11/11 Ocala NF-005 scrub /11/11 Ocala NF-021 scrub /11/11 Etoniah Creek SF-011 sandhill scrub /18/11 Etoniah Creek SF-007 sandhill scrub /18/11 Etoniah Creek SF-005 sandhill scrub /18/11 Etoniah Creek SF-006 flatwoods scrub ecotone /18/11 Etoniah Creek SF-022 scrub /18/11 Ocala NF-001 sandhill /23/12 57

58 Table 2-4. Continued Location Habitat type Water temp Air temp Soil temp ph DO Conductivity collection date Ocala NF-014 sandhill /23/12 Ocala NF-017 sandhill /23/12 Osceola NF-001 flatwoods /4/11 Osceloa NF-002 flatwoods /5/12 Osceloa NF-003 flatwoods /5/12 Osceola NF-004 flatwoods /5/12 Big Bend WMA-002 sandpine scrub/flatwoods /2/11 58

59 CHAPTER 3 RANAVIRUS MORTALITY EVENT AT MIKE ROESS GOLD HEAD BRANCH STATE PARK, FLORIDA, USA. Introduction Amphibian populations have been experiencing declines globally over the past few decades. These declines have been associated with a number of factors, including habitat alteration and introduced pathogens (Cheng et al., 2011; Stuart et al., 2004). Outbreaks caused by pathogens of the genus Ranavirus (Family Iridoviridae) were believed to be the largest single cause of reported amphibian mass mortality events in the United States from (Daszak et al., 1999; Green et al., 2002) and have been associated with a number of amphibian mortality events in the Americas, Europe, and Asia (Cunningham et al., 1996). Despite these widespread die-offs, Ranavirus spp. have yet to be reported in any amphibian populations in the state of Florida. Ranavirus spp. can impact a number of different vertebrates, including bony fishes, reptiles, and amphibians (Chinchar, 2002). Despite the widespread infections of Ranavirus spp., the actual threat that this virus poses to herpetofaunal species is still unknown (Gray et al., 2009; Schock et al., 2008). Herein, we document the first known amphibian die-off related to Ranavirus in Florida. On 7 April 2011 during an amphibian disease survey, we observed an amphibian mortality event at Pebble Lake in Mike Roess Gold Head Branch State Park, Clay County, ( N, W; Figure 3-1). Pebble Lake is a sinkhole pond in upland sandhill habitat. We observed hundreds of dead or dying Bullfrog (Lithobates catesbeianus) and Southern Leopard Frog (L. sphenocephalus) tadpoles. After collecting 10 tadpoles (six L. catesbeianus and four L. sphenocephalus), we noticed 59

60 whitish lesions on the dorsal surface of the body, hemorrhaging around the vent and ventral surface, and swollen, edematous abdomens on some specimens (Figures 3-2 and 3-3). Methods Freshly dead or dying tadpoles were placed on ice and later necropsied. Each specimen was placed in an individual Ziploc bag, and a new pair of nitrile gloves was used when handling each successive specimen to prevent cross contamination. A sagittal (from inferior to superior along the mid-ventral surface) incision was made on the ventral surface of each tadpole. A large amount of blood was observed in the abdominal cavity, likely the result of massive hemorrhaging; additionally, the livers and kidneys contained a large amount of blood that was also most likely due to internal hemorrhaging. Livers were removed and placed into a container with 70% ethanol. The six livers from the L. catesbeianus tadpoles were combined into one vial while the four livers from the L. sphenocephalus tadpoles were combined into a separate vial. Samples were sent to the Wildlife Disease Laboratories, San Diego Zoo, to test for Ranavirus spp. presence by PCR analysis. Although water and vegetation at the site appeared normal, we tested the following water quality parameters: dissolved oxygen, ph, and conductivity. On 25 April 2011, we returned to Pebble Lake and observed thousands of Southern Toad (Anaxyrus terrestris) and a few Gopher Frog (L. capito) tadpoles. All appeared healthy with normal behavior and no gross abnormalities. The A. terrestris tadpoles were too small to sample for Ranavirus spp., but three L. capito tadpoles were sampled for Ranavirus spp. 60

61 Results Combined samples for both L. catesbeianus and L. sphenocephalus tested positive for Ranavirus spp. However, L. capito samples tested negative for Ranavirus spp. Water-quality data measurements collected on 7 April 2011 were dissolved oxygen = 7.9, ph = 6.7, and conductivity = 10. The water quality data measurements collected on 25 April 2011 were dissolved oxygen = 8.1, ph = 6.8, and conductivity = 10. Discussion The results of this study illustrate that Ranavirus spp. is present in amphibian populations in Florida. This mortality event suggests that Ranavirus spp. is a threat to amphibian populations, at least on a local scale. Although mortality of amphibians in a single event can be high (Green et al., 2002), mortalities are often restricted to a small geographical area, often a single pond (Teacher et al., 2010). Despite the high mortality associated with such events, it is not yet known whether Ranavirus spp. is capable of causing long-term population declines (Teacher et al., 2010). Kevin M. Enge (personal observation) has surveyed thousands of ponds across Florida while working on amphibians. During this time, he has observed only two mass mortality events where dead and moribund tadpoles exhibited similar physical signs as we documented herein for Pebble Lake. However, this is the first documented case of Ranavirus spp. infections in Florida amphibians, and possible linkage to a sizeable mortality event. The sizes of the infected L. catesbeianus and L. sphenocephalus tadpoles indicate that the former species overwintered in Pebble Lake and the latter species were born that winter. The smaller and younger L. capito tadpoles appeared healthy and uninfected. Gray et al. (2007) found 57% of overwintering L. catesbeianus tadpoles 61

62 exhibited similar pathological signs of infection and tested positive for Ranavirus spp. Ranavirus mortality events are most likely under-reported as like this one, they may occur across short time spans and in remote areas. 62

63 Figure Fig Location of Pebble of Lake in in Mike Roess Gold Head Branch State Park, Park, Clay County, Florida, USA, where Ranavirus spp. was detected during a mortality event 63

64 Figure 3-2. Lateral view of Bullfrog (Lithobates catesbeianus) tadpole with white lesions, indicative of Ranavirus spp. infection, Pebble Lake, Clay County, Florida, USA. Photo by Kevin M. Enge 64

65 Figure 3-3. Ventral view of Bullfrog (Lithobates catesbianus) tadpole with swollen, blotched belly and red swollen vent, both indicative of Ranavirus spp. infection. Pebble Lake, Clay County, Florida, USA. Photo by Kevin M. Enge 65

66 CHAPTER 4 ANALYSIS OF PRESERVED AMPHIBIAN SPECIMENS TO TEST FOR DETECTION OF CHYTRID FUNGUS (Batrachochytrium dendrobatidis) Introduction Historically the southern dusky salamander (Desmognathus auriculatus) was abundant in steephead ravines throughout the panhandle and the northern peninsula of Florida (Means and Travis, 2007). This species has now disappeared from most of its historical range. Means and Travis (2007) determined that as of the 1990s, D. auriculatus is effectively extinct from the steephead ravines on Eglin Air Force Base, covering Santa Rosa, Okaloosa, and Walton Counties, Florida. This area once sustained thriving populations of D. auriculatus (Means and Travis, 2007). In order to determine whether B. dendrobatidis could have been the cause of this extirpation, preserved specimens of this species of concern were tested for the presence of B. dendrobatidis (Figure 4-1). Batrachochytrium dendrobatidis has been detected in specimens preserved for up to six decades (Weldon et al., 2004). This process is an excellent tool for determining whether or not B. dendrobatidis was the cause of the extirpation of a population when individuals of these populations have been properly preserved and stored. Methods A total of 20 southern dusky salamander (D. aurituculatis) specimens from the Florida Museum of Natural History (FLMNH) Herpetology Collection were used in this study. The specimens were originally fixed with formalin and then put in 70% ethanol for preservation. Since preservation, all specimens have been stored in glass jars in the Herpetology Collection at the FLMNH. 66

67 A list of catalogued specimens from the appropriate localities was compiled using the herpetology database at the FLMNH. This list of specimens was narrowed down by looking at the time of year each specimen was collected. Since B. dendrobatidis thrives in cooler temperatures, only specimens collected between the months of November and April were considered for swabbing. A total of twenty individual specimens were selected to be sampled. Four individuals from Devil s Millhopper in Alachua County, five individuals from Tom s Creek in Eglin Air Force Base in Okaloosa County, five individuals from Deep Springs Canyon in Bay County, and six individuals from Silver Glen Springs in Marion County were selected (Tables 4-1 and 4-2) Procedures from Soto-Azat et al. (2010) were followed in order to ensure that no samples became contaminated from other preserved specimens. Specimens which were in the same jar as one another were washed with 70% ethanol prior to swabbing to wash off any free-floating B. dendrobatidis zoospores. After being rinsed with ethanol each specimen was thoroughly swabbed using a small cotton tipped swab. The ventral abdomen and pelvis, fore and hind limbs, as well as fore and hind feet were firmly swabbed. A new pair of nitrile gloves were used for each specimen in order to prevent cross-contamination. Results Of the 20 samples submitted for analysis one swab came back positive for B. dendrobatidis. The positive specimen came from Deep Spring Canyon in Bay County, Florida. Four other individuals collected at the same locality on the same day were also sampled but came back negative for B. dendrobatidis (Figure 4-2). 67

68 Discussion The results of this study are inconclusive as they do show that B. dendrobatidis was present on one specimen, however, 19 of the specimens came back negative for the pathogen. The pathogen was present in at least one individual in one population of D. aurituculatis in Deep Spring Canyon, Bay County. However, this single positive sample is not enough to draw conclusions about the impact that B. dendrobatidis had on this population. This species has declined throughout most of its historic range throughout Florida and more sampling needs to be conducted in order to determine what caused the extirpation of these populations. Other studies, such as those by Weldon et al., 2004, Gleason et al., 2007, and Ouellet et al., 2005 have used the same type of methods described above and have been able to show B. dendrobatidis as at least a main factor in the extirpation of an amphibian population. Therefore it is possible to show that this pathogen could be the cause of this species decline. However, in order to determine whether B. dendrobatidis can be implicated in these D. auriculatus population declines, a larger number of specimens from the historical range will need to be tested for the pathogen. 68

69 Figure 4-1. Five southern dusky salamander (Desmognathus aurituculatis) specimens from Deep Springs Canyon, Bay County, FL, which were sampled for Batrachochytrium dendrobatidis. Photo by Sarah Reintjes-Tolen 69