Geographic Distribution of Chytrid Fungus (Batrachochytrium Dendrobatidis) and Ranavirus Spp. in Amphibians in Northern ...

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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
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1 online resource (88 p.)
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english
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Reintjes-Tolen, Sarah
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University of Florida
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Gainesville, Fla.
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Thesis/Dissertation Information

Degree:
Master's ( M.S.)
Degree Grantor:
University of Florida
Degree Disciplines:
Wildlife Ecology and Conservation
Committee Chair:
Carthy, Raymond R
Committee Members:
Krysko, Kenneth L.
Means, Donald B

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Subjects / Keywords:
amphibians -- chytrid -- florida -- pathogens -- ranavirus
Wildlife Ecology and Conservation -- Dissertations, Academic -- UF
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Wildlife Ecology and Conservation thesis, M.S.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

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Abstract:
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 of32 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, 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.
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In the series University of Florida Digital Collections.
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Includes vita.
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Description based on online resource; title from PDF title page.
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This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility:
by Sarah Reintjes-Tolen.
Thesis:
Thesis (M.S.)--University of Florida, 2012.
Local:
Adviser: Carthy, Raymond R.
Electronic Access:
RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2014-08-31

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lcc - LD1780 2012
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UFE0044510:00001


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1 GEOGRAPHIC DISTRIBUTION OF CHYTRID FUNGUS ( Batrachochytrium dendrobatidis ) AND R anavirus spp IN AMPHIBIANS IN NORTHERN P ENINSULAR 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 2012

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2 2012 Sarah Reintjes Tolen

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

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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. Petersbu rg, research team for their assistance with this project. I would additionally like to thank Jo s e ph Mansuetti, Lindsay M. Wag ner a nd Stephen Harris for help collecting samples. I am grateful for help from the Florida Fish and Wildlife Conservation Commission, the Central Florida Zoo logical Gardens the Wildlife Disease La boratories at the San Diego Zoo and the Florida Museum of Natur al History I would like to thank Dr. Kent Vliet for giving me the opportunity to receive f unding through a Teaching Assis tantship 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. Ehr et, Carly L. Manz, Meghan Godby and Buddy Coleman for their suppor t both academically and emotionally. I am very appreciative of the unconditional love of my dog Syd who was always there for me. Finally I wou ld like to thank my parents, Susan Reintjes and Steve Tolen for their moral support, not only during the pa st few years but my entire life. W ithout their continued support I would have never made it this far.

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5 TABLE OF CONTENTS page ACKNOWLEDGMENTS ................................ ................................ ................................ ........ 4 LIST OF TABLES ................................ ................................ ................................ .................. 7 LIST OF FIGURE S ................................ ................................ ................................ ................ 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 ................................ ................................ ................................ 24 2 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 Sa mpling ................................ ................................ .............................. 38 Swabbing for Batrachochytrium Dendrobatidis ................................ .................... 39 Swabbing for Ranavirus spp. ................................ ................................ ................ 40 Environmental Factors ................................ ................................ ........................... 40 Disinf ection ................................ ................................ ................................ ............. 41 Pathogen Detection ................................ ................................ ............................... 41 Results ................................ ................................ ................................ ........................... 42 Discussion ................................ ................................ ................................ ..................... 43 3 RANAVIRUS MORTALITY EVENT AT MIKE ROESS GOLD HEAD BRANCH STATE PARK, FLORIDA, USA. ................................ ................................ ................... 59 Introduction ................................ ................................ ................................ .................... 59 Methods ................................ ................................ ................................ ......................... 60 Results ................................ ................................ ................................ ........................... 61

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6 Discussion ................................ ................................ ................................ ..................... 61 4 ANALYSIS OF PRESERVED AMPHIBIAN SPECIMENS TO TEST FOR DETECTION OF CHYTRID FUNGUS ( Batrachochytrium dendrobatidis ) ................ 66 Introduction ................................ ................................ ................................ .................... 66 Methods ................................ ................................ ................................ ......................... 66 Results ................................ ................................ ................................ ........................... 67 Discussion ................................ ................................ ................................ ..................... 68 5 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 ................................ ................................ ................................ 88

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7 LIST OF TABLES Table page 2 1 The locality of each pond where amphibians were sampled for Batrachochytrium dendrobatidis and/or Ranvirus spp ................................ ........... 52 2 2 Summary table of data collected on chytrid fungus, Batrachochytrium dendrobatidis, in northern peninsular and panhandle Florida ............................... 53 2 3 Summary table of data collected on Ranavirus spp. in northern peninsular and panhandle Florida ................................ ................................ ............................. 55 2 4 Environmental data collected at each pond where amphibians were sampled for Batrachochytrium dendrobatidis and/or Ranvirus spp ................................ ...... 57 4 1 Desmognathus auriculatus specimens from the Florida Museum of Natural History that were sampled for Batrachochytrium dendrobatidis ............................ 71 4 2 Specific locality and number of Desmognathus auriculatus specimens from the Florida Museum of Natural History that were sa mpled for Batrachochytrium dendrobatidis ................................ ................................ .............. 72

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8 LIST OF FIGURES Figure page 1 1 The life cycle of the chytrid fungus, Batrachochytrium dendrobatidis illustrating substrate dependent and substrate independent stages .................... 26 1 2 Photomicrograph of a stained kidney from a green frog ( Lithobates clamitans ) larva infected with Ranavirus spp ................................ ......................... 27 1 3 Representative pond (Pond 3) within Apalachicola National Forest, Liberty County, Florida, USA. Photo by Kevin M. Enge ................................ ..................... 28 1 4 Representative pond (Pebble Lake) within Gold Head Branch State Park, Clay County, Florida, USA. Photo by Sarah Reintjes Tolen ................................ 29 1 5 Representative pond (Pond 3) within Camp Blanding Military Reserve, Clay County, Florida, USA. Photo by Sarah Reintjes Tolen ................................ .......... 30 1 6 Representative pond (Pond 1) within Osceola National Forest, Baker County, Florida, USA. Photo by Sarah Reintjes Tolen ................................ ........................ 31 1 7 Representative pond (Pond 21) within Ocala National Forest South of Salt Springs, Marion County, Florida, USA. Photo by Sarah Reintjes Tolen ............... 32 1 8 Representative pond (Pond 1) within Ocala National Forest South of Church Lake, Marion County, Florida, USA. Photo by Sarah Reintjes Tolen ................... 33 1 9 Representative pond (Pond 7) within Etoniah Creek State Forest, Putnam County, Florida, USA. Photo by Sarah Reintjes Tolen ................................ .......... 34 1 10 Map illustrating sampling sites where amphibians were swabbed for Batrachochytrium dendrobatidis and/or Ranavirus spp in Florida, USA ............. 35 2 1 Sampling amphibians for Batrachochytrium dendrobatidis and Ranavirus spp and recording data from individuals caught in Camp Blanding Military Reserve ................................ ................................ ................................ ..................... 46 2 2 Sampling a Southern Leopard Frog ( Lithobates sphenocephalus ) tadpole for Batrachochytrium dendrobatidis and Ranavirus spp at O cala National Forest .... 47 2 3 Sampling an adult Southern Cricket Frog ( Acris gryllus ) for Batrachochytrium dendrobatidis at Ocala National Forest South of Church Lake ............................. 48 2 4 Sampling an adult Southern Cricket Frog ( Acris gryllus ) for Ranavirus spp a t Ocala National Fores t South of Church Lake ................................ ......................... 49

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9 2 5 Map of sites sampled for Batrachochytrium dendrobatidis and Ranvirus spp. mapp ................................ ................................ ................ 50 2 6 Number of individual amphibians, sampled for both Batrachochytrium dendrobatidis and Ranavirus spp ., by species and life stage ............................... 51 3 1 Location of Pebble Lake in Mike Roess Gold Head Branch State Park, Clay County, Florida, USA ................................ ................................ ............................... 63 3 2 Lateral view of Bullfrog ( Lithobates catesbeianus ) tadpole with white lesions, indicative of Ranavirus spp infection, Pebble Lake ................................ ............... 64 3 3 Ventral view of Bullfrog (Lithobates catesbianus) tadpole with swollen, blotched belly and red swollen vent, both indicative of Ranavirus spp. ................ 65 4 1 Five southern dusky salamander ( Desmognathus aurituculatis ) specimens from Deep Springs Canyon, Bay County, FL, which were sampled for Batrachochytrium dendrobatidis ................................ ................................ .............. 69 4 2 Map showing collection location of each Desmognathus auriculatus specimen sampled for Batrachochytrium dendrobatidis ................................ ........ 70

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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 sp p IN AMPHIBIANS IN NORTHERN P ENINSULAR AND PANHANDLE FLORIDA WITH A CASE OF A RANAVIRUS DIE OFF IN G OLD HEAD BRANCH STATE PARK By Sarah Reintjes Tolen August 2012 Chair: Raymond R. Carthy Major: Wildlife Ecology and Conservation Amphibian species across the globe have been experiencing population declines. There are a number of facto rs attributed to these declines including habitat destruction, increased ultraviolet radiation, pollution and introduc ed diseases. H owever, one of the most potentially devastating of these factors has been the emergence of two pathogens, chytrid fungus ( Batrachochytrium dendrobatidis ) and R anavirus spp Determining the distribution of these pathogens is important as it is un known 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 pond s surveyed one in Camp Bland ing Military Reserve and one in Osceola National Forest. Ranavirus spp was detected in one pond in Gold Head Branch State Park. Additionally

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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.

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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 b e indicator species showing physical or behavioral changes with ev en the smallest variation in physical, chemical, or ecological make up of their surroundin 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 ecologic al role as secon dary consumers. Adult amphibians consume great quantities of flies and insects, including mosquitos and their la rvae (Mohneke and Rodel, 2009). For example a single adult of certain species of cricket f rog s can consume approximately 4 ,800 small in sects pe r year (Johnson and Christia nsen, 1976 ). Many of the insect species consumed are vectors for human diseases such as malar ia, yellow fever and West N ile vir us (Moh neke and Rodel, 2009).

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13 In addition t adpoles provide ecological services in the freshwater envi ronme nt by consuming algae. A number of tadpole exclusion experiments have been performed to determine the impact that tadpoles hav e 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 pr ey item for many other species. For example; frog eggs are consumed by was ps and spiders ; shrimp, fish and dragonfly nymphs eat tadpoles; and birds, snakes and lizards eat frogs (Mohneke and Rodel, 2009). Removing amphibi ans from an ecosystem in some cases could h ave 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 differen t factors have been hypothesized as causing these population declines. Some of these include habitat destruction and modification, climate change, over exploitation, pollution, introduce d 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 pop ulations throughout the world (Berger et al., 1998 ) Berger et al (1998) attributed a number of these mort ality events to chytrid fungus ( Batrachochytrium dendrobatidis ) because it s symptoms were observed in dying amphibians in Aust ralia

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14 and Central America Additionally B dendrobatidis has been attributed to the collapse of amphibian populations in Panama, C alifornia, 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 docume nt the distribution of two pathogens B dendrobatidis and R anavirus spp in Florida, which have been implicated in declining amphibian populations around the world. Amphibian An atomy and Life H istory Amphibians possess distinct anatomy and life history unique among vertebrate s The moist skin of amphibi ans is an interactive interf ace between the animal and its environment. It serves as a membrane for the exchange of gases, water, as well as any other material s in the environment. The skin is highly perm eable and epidermal sculpturing is very im portant in water conservation (D uellman and Trueb, 1986). Water saving 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 tiss ues (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) I rregular ventral surface s provide increased surface area for absorbing water through substrates. The ventral pelvic region of

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15 anurans is highly specialized as it is the primary source of water uptake for the se organism s ( Duellman and Trueb, 1986 ). This reg ion 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. T he epidermis is composed of four layers, which include the stratum corneum, stratum granulosum, str atum 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 aff ected by temperature, metabolic rate, and photoperiod and are under hormonal contro l (Wells, 2007) The dermis of amphi bians is considerably thicker than the epidermis and consists of two layers, the stratum spo n gi osum and the stratum compactum ( Wells, 2007 ) A mphibians have a complex life cycle beginning as aquatic organisms and then undergo ing metamorphosis to become ter restrial 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

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16 aquatic and terrestrial environments at some point in their lives they depend on both ecosystems. Batrachochytrium D endrobatidis Life H istory One introduced pathogen chytrid fungus ( B dendrobatidis ), can cause ch ytridiomycosis and has been impl icated in declines of many montane amphibian popu lations (Daszak et al., 2003 ). B atrachochytrium dendrobatidis can affect the skin cells co ntaining keratin with subsequent infections lead ing 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 17 25 C, but growth may occur at temperature as low as 4 C (Piotrowski et al., 2004). B atrachochytrium dendrobatidis is able to grow and reproduce at pH 4 8, but growth i s opti mal at pH 6 7 (Piotrowski et al., 2004). Batrachochytrium dendrobatidis has fairly high sur vival 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 infectio us st age (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 int o the environment (zoosporangia ) ( Figure 1 1). Batrachochytrium dendrobatidis is attracted to keratinized tissue resulting in thickening (i.e., hyperpl asia which is an increase in number of cells and hyperkeratosi s which is a thickening of the stratum corneum ) of the stratum corneum ( Daszak et al., 200 7; Johnson and Speare, 2003 ) in both newly metamorphosed

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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 gran ulosum of the epidermis. It then matures in the stratum corneum where it enters healthy cells and grows into a zoosporangium within which zoospores devel op (Rollins Smith et al., 2011) Eventually infected skin cells move toward the surface the zoosporangium matures, a discharge papilla (pore) o pens and mature zoospores swim out (Rollins Smith et al., 2011). B atrachochytrium 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 account s show that many larvae are unaffected by infection of B dendrobatidis M ortality 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 u ntil 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 leg s and drink patch are elevated ( Daszak et al. 2007) Other symptoms may include neurologic al signs such as abnormal sitting posture with hind legs adducted lethargy, and s low response to tactile stimuli, Red Leg S yndrome like skin discoloration, and in salamanders b lack spots on the skin and loss of the tail and toe ti ps ( Daszak et al., 2007 )

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18 Laboratory experiments indicate that B. dendrobatidis grows most rapidly, with a 4 5 day generation time at cool temperatures (17 C 25 C) (Piotr owski 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. de ndrobatidis 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 fro g 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 produce s the observed infection pattern where prevalence quickly cha nges from very lo w 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 ) S ome amphibians are known to carry a nd tolerate B dendrobatidis infections. These species may be asymptomatic, live in t he community and act as vectors. Among them are the American Bullfrog ( Lithobates catesbeianus ), Cane Toad ( Rhinella marina ), Xen opus spp., Eastern Tiger Salamander ( Ambystoma tigrinum ) and Cricket Frog ( Acris gryllus ) ( Daszak et al. 2007 ). A ll of these species have been introduced

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19 globally (Collins et al 1988; Kraus, 1999; Mazzoni et al 2003; Hanselmann et al 2004; Weldon et al 2004 ). T reatment of amphibians infected with chytridiomycosis in captivity has been success ful 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 2008). This treatment protocol should not be used on tadpoles or recent metamorphs as death may occur ( Pessier, 2008 ) Multiple treatme nt 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 s p p. Life H istory Another pathogen that has severely imp acted amphibian populations are three viral species grouped togeth er 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., 200 9). Amphibians infected with Ranavirus spp experience hemorrhaging, reddeni ng of the belly, legs, and vent, l ethargy, 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

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20 hemorrhaging within the skeletal muscles and visceral organs (Teacher et al., 2010). I nfection is most easily observed in the kidneys and liver o f 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 viru s (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 var ying 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 fro m 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 sp p 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. M any mortality events therefore, most likely go unnoticed (Gray et al., 2009). Not much atten tion 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 recreation al fish species, cultured and wild frogs, and endanger ed salamanders,

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21 has brought increased attention to the genus (Chinchar, 2002). T hree species of Ranavirus are know n to infect amphibians. They are Ambystoma tigrinum virus (ATV), Bohle iridovirus (BIV) and Frog V irus 3 (FV 3) (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 R anaviruses Each major group most likely contains numerous different R anavirus species that can be difficult t o differentiate without additional analyse s One major R anavirus group commonly found with these PCR primers is most closely related to the type of virus for the genus Ranavirus (Frog V irus 3), but based on PCR and sequencin g products one most often gets results that indicate a 3 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 fast est replication occurring at 30 C and slo wer replication occurring at 1 C (Teacher et al., 2010). F rog V irus 3 replicates at temperatures ranging from 12 32 C making it able to survive an d reproduce at a fairly broad r ange of temperatures (Chinchar, 2002). Transmission of R anavirus is thought to occur by multiple pathways including contaminated soil, direct c ontact, 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 communit ies. It was found that both vertical

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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 l ikely 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 S tate of Florida A number of amphibian species in the state of Florida have been experiencing population declines and in some cases extir pation from historical ranges. Some species that have experienced major decl ines include the striped newt ( Not o p hthalmus perstriatus ) and the southern dusky salamand er ( Desmognathus auriculatus ) ( Means et al ., 2008 and Means & Travis 2007 ) In July of 2008 Means et al. (2008) petition ed the U.S. Fish and Wildlife Service to list the striped newt as a Federally Threatened species under the Endangered Species Act of 1973. This was due to a nu mber 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 purpos es, 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) S outhern dusky salamander ( D auriculatus ) populations have disappeared over the past few decades at several localities in Florida. Some of these area s that historically had D. auriculatus populations include Deep Springs C anyon in Bay County, Lightwood Knot Creek, Garnier C reek, reek o n Egl in Air Force Base in Okaloosa county (D. B. Means, personal

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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 1997 2007 did not detect B dendrobatidis (K. M. Enge personal communication). After 2007, h owever, 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 pe rsonal communication). Rizkalla (2009) detected B dendrobatidis in both L. catesbeianus and Acris gryllus at both the Wa lt 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. D etermining the geographic distribu tion of these pathogens in northern Florida is a critical first step in assessing it the impact that these pathogens are having on the amphibian populations. Objective s and Significance of Current Study The objective s 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 distri bution of these pathogens, and enable the tracking of the spread of these diseases. This is important for the protection and

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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 group s 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 area s 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 causin g amphibian population declines recently observed in Florida and other sites worldwide This documentation of the distribution of B dendrobatidis and Ranavirus spp can assist st ate wildlife agencies in implement ing the appropriate conservation m easures 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, Blackwa ter, East Bay, Yellow, Choctawhatchee, Choctawhatchee Bay, St. Andrews Bay, Apalachicola, Whiskey George Creek, Crooked, Ochlockonee, St. Marks, A veilla, Upper Suwannee, Lower Suwannee, St. Marys, Ecofina, Fenholloway, Spring Warrior Creek, Blue Creek, Ste inhatchee, Amason Creek, California Creek, Waccasassa, Withcacoochee, Nassau, and St. Johns (Robert H. Robins, personal

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25 communication) Many of the ponds in my study area were ephemeral and are therefore not connected to any of these drainages. Samp les we re 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 S outh of C hurch L ake Etoniah Creek State Forest and Osceola National Forest (Figures 1 3 to 1 9) All sites were located on land owned by the sta te or federal government and included h abitats of upland sandhill s, grassy pond s, cypress gum pond s in flatwoods borrow pit s, sandhill/scrub flatwoods scrub ecotone reclaimed mine land ( formerly scrub) sinkholes, sandpine scrub/flatwoods and long leaf/slash pine flatwoods ( Figure 1 10)

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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)

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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)

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28 Fig ure 1 3 Representative pond (P ond 3) within Apalachicola National Forest Liberty County, Florida, USA. Photo by Kevin M. Enge

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

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

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31 Figure 1 6. Representative pond (P ond 1) within Osce ola National Forest, Baker County, Florida, USA Photo by Sarah Reintjes Tolen

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32 Figure 1 7 Representative pond (P ond 21) within Ocala National Forest S outh of S alt S prings Marion County, Florida, USA Photo by Sarah Reintjes Tolen

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33 Figure 1 8 Representative pond (P ond 1) within O cala National Forest S outh of C hurch L ake Marion County, Florida, USA Photo by Sarah Reintjes Tolen

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34 Figure 1 9 R epresentative pond (P ond 7) within Etoniah Creek State Forest Putnam C ounty, Florida, USA Photo by Sarah Reintjes Tolen

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35 Figure 1 10 Map illustrating sampling sites where amphibians were swabbed for Batrachochytrium dendrobatidis and/or Ranavirus spp in Florida, USA

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36 CHAPTER 2 GEOGRAPHIC DISTRIBUT ION OF CHYTRID FUNGU S ( BATRACHOCHYTRIUM DENDROBATIDIS ) AND RANAVIRUS SP P IN NORTH ERN PENINSULAR AND PANHANDLE FLORIDA AMPHIBIANS Introduction In order to determine the geographic distribution of these two amphibian pathogens, a mphibians from seven sites in northern peninsular and panhandle Florida were sampled for Batrachochytrium dendrobatidis and Ranavirus spp This survey was the first of it s 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 an d Conservation Area, Orange County (Rizka lla, 2009) and in Ocala National Forest, Putnam County (K. M. Enge, personal commu n ication) 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 pon ds and 296 individuals, across nine species, were sampled. Field surveys were conducted between t he months of November and April, in which ambient air temperat ures allowed for optimal growth and survival of B dendrobatidis This time

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37 period also coincided with the winter amphibian breeding season in Florida Skerratt et al. (2008) noted that there a re a number of key aspects to consider when surveying amphibian populations across large areas such as ar ea, species, life stage, sample size, planning a surveying strategy, season, and diagnostic tests. Sampling S ites I attempted to sample a minimum of 3 0 individual amphibians of a variety of species at each of the seven sites Within each site amphibians were sampled fro m 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 diff erent species and increase the likelihood of detecting B dendrobatidis and/or Ranavirus spp D ue to low water levels I was only able to obtain samples from one pond in Gold Head Branch State Park. At each site, excep t for those within A palachicola N ational F orest 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 as sessed for either of these pathogens. Species S ampled 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 in cluded the Tiger Salamander ( Ambystoma tigrinum ), Reticulated Flatwoods Salamander and Frosted Flatwoods Salamander ( A bishopi and A cingulatum ), Striped Newt ( Notophthalmus perstriatus ), Ornate C h orus Frog ( Pseudacris ornata ) and Gopher Frog ( Lithobates capito ). T hese species are considered species of greatest conservation need because of their declining or low

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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 So u thern Leopard Frog ( L sphenocephalus ), Bullfrog ( L catesbeianus ), and Southern Cricket Frog ( Acris gryllus ) A dult A. gryllus and L. catesbeianus are kn own to be carriers of B. dendrobatidis which made these species a sampling priority ( Daszak et al., 2007 ) 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); oad, Anaxyur a 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 ; Ouelle t 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 ) T hese species were given priority as part of the 30 miscellaneous individuals that were sampled at each site. Amphibian S ampling Amphibians were collected us ing 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. E ach 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

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39 320 g) ; and any atypica l physical charact eristics such as lesions, deformities, swollen limbs or a b d omen, 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 amphib i ans: 70% ethanol, co tton 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 D endrobatidis T o 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 tota l sweeps per animal). These areas commonly have a high concentration of B. dendrobatidis sporangia (B rem et al., 2007) To properly swab each area of adult amphibian s it is best to grip the animal mid dorsally anterior to the hind legs wit h the ventral side up. For e xtremel y small individuals or salamanders the position or technique may need to be altered slightly in order obtain a n adequate swab ( Figure 2 3) (Brem et al., 2007) T o test for B dendrobatidis in larvae the pharyngeal mucosa was swabbed. T hen, t he 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 c ross contamination between individuals and/or samples

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40 Swabbing for Ranavirus s pp. T o 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 n ecropsy (A. P. Pessier, personal communication ). The kidneys and liver are removed and placed in a lab eled 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 environmen tal factors were recorded. A biotic factors included ambient air, soil, and water temperature s pH, diss olved oxygen, and specific conductivity ( Table s 2 1 and 2 2) A Garmin GPS 76Cx was used to obt ain the latitude and longitude at each site. A Hannah Instruments HI 9142 dissolved oxygen meter was used to obtain dissolved oxygen measur e ments 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

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41 were taken to the nearest .01 C. A Hannah Instruments 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 pond s 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 suscepti ble to infection from pathogens. Pathogen D etection The most effective method of detection involves euthanizing the animal and then performing a necropsy to remove the liver and kidneys. T issue samples from t hese organs are then analyzed using either a histological or PCR analysis to determine the presence or a bsence of the pathogen (A. P. Pe ssier, 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 i s 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, maki ng 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

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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 s ampled for B dendrobatidis including nine species at 32 different localities. Some individuals were s ampled for both B dendrobatidis and Ranavirus spp while others were only s ampl ed for B. dendrobatidis S amples of Ranavirus spp were limited by the fact that Ranavirus spp i s difficult to detect from pharyngeal mucosa swabs. This resulted in slightly fewer Ranavirus spp. samples in the study. Two swab s from Southern Cricket F rogs 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 F rogs that were asymptomatic and behaved normally at the time of collection ( Figure 2 5) Two swabs one from six Bullfrog s the other from four Leopard Frog tadpoles gave positive results for Ranavirus spp Both of these swabs were from individuals from Pebble Lake in Go ld 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 woul d give an infection rate of 0. 68 % which is the lowest possible infection rate The next possible scenario simply takes into account the two positi ve swabs out of a total of 103 swabs. This gives an infection rate of 1 8 % The highest possible infection rate assumes that all

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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 posit ive 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 infec tion rate of 4.0% All of the environmental factors measured came back with in normal ranges (T able 2.1). This suggests that none of the infected individuals were directly compromised by existing environmental conditions. Discussion My results sh ow 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 pa thogens are present in Florida. This means that there is the possibility of infection and extirpat ion of vulnerable populations of amphibians. Based on the infection rates found in this study i t appears that t here is a low background infection rate for B dendrobatidis with individuals that are simply carriers

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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 infectio n 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 positi ve for Ranavirus spp. were obtained from dead and moribund tadpoles. Despite the fact t hat neither B dendrobatidis nor Ranavirus spp had been dete cted in Florida until 2008 and 2011 does not mean that these pathogens were not prese nt in Florida before this time. These pathogens could have been present for an u nknown amoun t of time and not s ampled, or if they were sampled, not detected. Additionally, due to the short time period during whic h 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 s cale 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 o ccurrence of two amphibian diseases in the state of Florida. The small scale of this study warrants caution in drawi ng conclusions using these data: because the diseases were found in a few localities does mean they are not mor e 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 R anavirus spp. infections/ die off s can be us eful to park and forest officials Florida Fish and Wildlife Conservation Commission biologists T hese diseases could certainly

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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 Nia years in 2010 2012 (NOAA) that resulted in severe droughts throughout the state of Florida many species of greatest conservation need were not known to breed d uring 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 a n unusually large percent age of ephemeral ponds were dry (K. M. Enge, personal c ommunication). 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). Bli hovde (2 006) 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 (Bl i hovde, 2006). Rainfall has considerable effects on the behavior of amphibian species. L ack of moisture may therefore result in very low levels of activity and low reproduction. This site fidelity in combination with drought condi tions could lead to densit y dependent infection rates as amphibians are forced to congregate in remaining water sources.

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46 Figure 2 1. S ampling amphibians for Batrachochytrium dendrobatidis and Ranavirus spp and recording data from individuals caught in Ca mp Blanding Military Rese rve, (P ond 26), Clay County, Florida, USA Photo by Kevin M. Enge

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

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

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

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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

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51 Figure 2 6. Number of individual amphibians, s ampled for both Batrachochytrium dendrobatidis and Ranavirus spp ., by species and life stage 0 20 40 60 80 100 120 140 160 Number of Individuals Species larvae adults

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52 Table 2 1 The locality of each pond where amphibians were s ampled for Batrachochytrium dendrobatidis and/or Ranvirus spp Location Pond name/number Lat/Long Blackwater SF 002 BSF 002 30.889167, 86.851389 Apalachicola NF 71.02 ANF 71.02 30.1048, 85.0583 Apalachicola NF 001 ANF 001 30.073056, 85.03 Apalachicola NF 100.01 ANF 100.01 30.0570, 85.0184 Apalachicola NF 002 ANF 002 30.033611, 84.996389 Apalachicola NF 003 ANF 003 30.040278, 84.983611 Apalachicola NF 004 ANF 004 30.296667, 85.019167 Camp Blanding MR 026 CBMR 026 29.8663, 82.0392 Camp Blanding MR 004 CBMR 004 29.88, 82.034722 Camp Blanding MR 001 CBMR 001 29.899722, 82.04 Camp Blanding MR 065 CBMR 065 29.896944, 82.041944 Camp Blanding MR 063 CMBR 063 30.028611, 82.0346 Gold Head Branch SP 001 GHBSP 001 29.825, 81.953611 Gold Head Branch SP 001 GHBSP 001 29.825, 81.953611 Ocala NF 003 ONF SSS 003 29.314444, 81.734167 Ocala NF 022 ONF SSS 022 29.3125, 81.735556 Ocala NF 006 ONF SSS 006 29.319167, 81.730556 Ocala NF 020 ONF SSS 020 29.318889, 81.725 Ocala NF 005 ONF SSS 005 29.328333, 81.724444 Ocala NF 021 ONF SSS 021 29.318889, 81.727222 Etoniah Creek SF 011 ECSF 011 29.770833, 81.867222 Etoniah Creek SF 007 ECSF 007 29.786667, 81.860556 Etoniah Creek SF 005 ECSF 005 29.753056, 81.859444 Etoniah Creek SF 006 ECSF 006 29.748889, 81.837778 Etoniah Creek SF 022 ECSF 022 29.748889, 81.834167 Ocala NF 001 ONF SCL 001 29.180278, 81.931111 Ocala NF 014 ONF SCL 014 29.178056, 81.913889 Ocala NF 017 ONF SCL 017 29.169444, 81.907222 Osceola NF 001 OsNF 001 30.304722, 82.373611 Osceloa NF 002 OsNF 002 30.18381, 82.45189 Osceloa NF 003 OsNF 003 30.22112, 82.57258 Osceola NF 004 OsNF 004 30.25071, 82.39259 Big Bend WMA 002 BBSCU 002 29.984444, 83.691111

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53 Table 2 2. Summary table of data collected on chytrid fungus, Batrachochytrium dendrobatidis i n northern peninsular and panhandle Florida Site Latitude/Longitude Date Species sampled Chytrid (positive/negative) Blackwater State Forest 30.889167, 86.851389 3/8/11 Lithobates sphenocephalus 0/1 Apalachicola National Forest 30.073056, 85.03 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.0570, 85.0184 3/9/11 Ambystoma cingulatum 0/10 Apalachicola National Forest 30.0570, 85.0184 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.033611, 84.99638 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.033611, 84.99638 3/9/11 Lithobates sphenocephalus 0/5 Apalachicola National Forest 30.296667, 85.01916 3/9/11 Lithobates sphenocephalus 0/5 Gold Head Branch State Park 29.825, 81.95361 4/7/11 Acris gryllus 0/5 Gold Head Branch State Park 29.825, 81.95361 4/7/11 Lithobates sphenocephalus 0/9 Gold Head Branch State Park 29.825, 81.95361 4/7/11 Lithobates catesbeianus 0/9 Camp Blanding Military Reserve 29.8663, 82.0392 4/7/11 Acris gryllus 0/5 Camp Blanding Military Reserve 29.8663, 82.0392 4/7/11 Lithobates catesbeianus 0/1 Camp Blanding Military Reserve 29.8663, 82.0392 4/7/11 Notophthalmus perstriatus 0/2 Camp Blanding Military Reserve 29.8663, 82.0392 4/7/11 Lithobates capito 0/6 Camp Blanding Military Reserve 29.88, 82.03472 4/7/11 Lithobates sphenocephalus 0/5 Camp Blanding Military Reserve 29.88, 82.03472 4/7/11 Acris gryllus 0/5 Camp Blanding Military Reserve 29.88, 82.03472 4/7/11 Lithobates capito 0/3 Camp Blanding Military Reserve 29.899722, 82.04 4/7/11 Lithobates sphenocephalus 0/5 Camp Blanding Military Reserve 29.899722, 82.04 4/7/11 Acris gryllus (5/5) Gold Head Branch State Park 29.899722, 82.04 4/25/11 Lithobates capito 0/3 Camp Blanding Military Reserve 29.89694, 82.04194 4/25/11 Lithobates sphenocephalus 0/5 Camp Blanding Military Reserve 29.896944, 82.04194 4/25/11 Acris gryllus 0/5 Camp Blanding Military Reserve 30.028611, 82.0346 4/25/11 Acris gryllus 0/5 Camp Blanding Military Reserve 30.02861, 82.0346 4/25/11 Lithobates sphenocephalus 0/5 Big Bend WMA 29.98444, 83.69111 5/2/11 Lithobates sphenocephalus 0/1 Osceola National Forest 30.30472, 82.37361 5/4/11 Acris gryllus 0/2 Ocala National Forest 29.31444, 81.73416 5/11/11 Acris gryllus 0/10 Ocala National Forest 29.31444, 81.73416 5/11/11 Lithobates catesbeianus 0/3 Ocala National Forest 29.3125, 81.73555 5/11/11 Acris gryllus 0/10 Ocala National Forest 29.3125, 81.73555 5/11/11 Notophthalmus perstriatus 0/2 Ocala National Forest 29.318889, 81.725 5/11/11 Lithobates catesbeianus 0/1 Ocala National Forest 29.31888, 81.725 5/11/11 Lithobates sphenocephalus 0/4 Ocala National Forest 29.32833, 81.72444 5/11/11 Lithobates catesbeianus 0/4 Ocala National Forest 29.31888, 81.72722 5/11/11 Acris gryllus 0/5 Etoniah Creek State Forest 29.77083, 81.86722 5/18/11 Acris gryllus 0/10 Etoniah Creek State Forest 29.78666, 81.86055 5/18/11 Acris gryllus 0/10 Etoniah Creek State Forest 29.75305, 81.85944 5/18/11 Acris gryllus 0/10

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54 Table 2 2. Continued Site Latitude/Longitude Date Species sampled Chytrid (positive/negative) Etoniah Creek State Forest 29.74888, 81.83777 5/18/11 Acris gryllus 0/10 Etoniah Creek State Forest 29.74888, 81.83777 5/18/11 Lithobates sphenocephalus 0/1 Etoniah Creek State Forest 29.74888, 81.83777 5/18/11 Lithobates capito 0/3 Etoniah Creek State Forest 29.74888, 81.83416 5/18/11 Acris gryllus 0/10 Ocala National Forest 29.18027, 81.93111 2/23/12 Acris gryllus 0/6 Ocala National Forest 29.18027, 81.93111 2/23/12 Lithobates sphenocephalus 0/5 Ocala National Forest 29.18027, 81.93111 2/23/12 Lithobates capito 0/4 Ocala National Forest 29.17805, 81.91388 2/23/12 Acris gryllus 0/10 Ocala National Forest 29.16944, 81.90722 2/23/12 Lithobates sphenocephalus 0/1 Ocala National Forest 29.16944, 81.90722 2/23/12 Acris gryllus 0/8 Osceola National Forest 30.18381, 82.45189 4/5/12 Lithobates sphenocephalus 0/2 Osceola National Forest 30.18381, 82.45189 4/5/12 Acris gryllus 0/4 Osceola National Forest 30.22112, 82.57258 4/5/12 Acris gryllus 0/11 Osceola National Forest 30.22112, 82.57258 4/5/12 Lithobates heckscheri 0/1 Osceola National Forest 30.25071, 82.39259 4/5/12 Acris gryllus (7/7)

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55 Table 2 3. Summary table of data collected on Ranavirus spp. i n northern p eninsular and panhandle Florida Site Latitude/Longitude Date Species sampled Ranavirus spp. (positive/negative) Blackwater State Forest 30.88916, 86.85138 3/8/11 Lithobates sphenocephalus 0/6 Apalachicola National Forest 30.1048, 85.0583 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.07305, 85.03 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.0570, 85.0184 3/9/11 Ambystoma cingulatum 0/10 Apalachicola National Forest 30.0570, 85.0184 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.03361, 84.99638 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.03361, 84.99638 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.04027, 84.98361 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.29666, 85.01916 3/9/11 Pseudacris ornata 0/5 Apalachicola National Forest 30.29666, 85.01916 3/9/11 Lithobates sphenocephalus 0/5 Gold Head Branch State Park 29.825, 81.95361 4/7/11 Lithobates sphenocephalus (4/4) Gold Head Branch State Park 29.825, 81.95361 4/7/11 Lithobates catesbeianus (6/6) Gold Head Branch State Park 29.825, 81.95361 4/25/11 Lithobates capito 0/3 Ocala National Forest 29.31444, 81.73416 5/11/11 Acris gryllus 0/10 Ocala National Forest 29.31444, 81.73416 5/11/11 Lithobates catesbeianus 0/3 Ocala National Forest 29.3125, 81.73555 5/11/11 Acris gryllus 0/10 Ocala National Forest 29.3125, 81.73555 5/11/11 Notophthalmus perstriatus 0/2 Ocala National Forest 29.318889, 81.725 5/11/11 Lithobates catesbeianus 0/1 Ocala National Forest 29.31888, 81.725 5/11/11 Lithobates sphenocephalus 0/4 Ocala National Forest 29.32833, 81.72444 5/11/11 Lithobates catesbeianus 0/4 Ocala National Forest 29.31888, 81.72722 5/11/11 Acris gryllus 0/5 Etoniah Creek State Forest 29.77083, 81.86722 5/18/11 Acris gryllus 0/10 Etoniah Creek State Forest 29.78666, 81.86055 5/18/11 Acris gryllus 0/10 Etoniah Creek State Forest 29.75305, 81.85944 5/18/11 Acris gryllus 0/10 Etoniah Creek State Forest 29.74888, 81.83777 5/18/11 Acris gryllus 0/10 Etoniah Creek State Forest 29.74888, 81.83777 5/18/11 Lithobates sphenocephalus 0/1 Etoniah Creek State Forest 29.74888, 81.83777 5/18/11 Lithobates capito 0/3 Etoniah Creek State Forest 29.74888, 81.83416 5/18/11 Acris gryllus 0/10 Ocala National Forest 29.18027, 81.93111 2/23/12 Acris gryllus 0/6 Ocala National Forest 29.18027, 81.93111 2/23/12 Lithobates sphenocephalus 0/5 Ocala National Forest 29.18027, 81.93111 2/23/12 Lithobates capito 0/4 Ocala National Forest 29.17805, 81.91388 2/23/12 Acris gryllus 0/10 Ocala National Forest 29.16944, 81.90722 2/23/12 Lithobates sphenocephalus 0/1 Ocala National Forest 29.16944, 81.90722 2/23/12 Acris gryllus 0/8 Osceola National Forest 30.18381, 82.45189 4/5/12 Lithobates sphenocephalus 0/2 Osceola National Forest 30.18381, 82.45189 4/5/12 Acris gryllus 0/4

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56 Table 2 3. Continued Site Latitude/Longitude Date Species sampled Ranavirus spp. (positive/negative) Osceola National Forest 30.22112, 82.57258 4/5/12 Acris gryllus 0/11 Osceola National Forest 30.22112, 82.57258 4/5/12 Lithobates heckscheri 0/1 Osceola National Forest 30.25071, 82.39259 4/5/12 Acris gryllus 0/7

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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 71.02 cypress gum pond/flatwoods / / / / / / 3/9/11 Apalachicola NF 001 cypress gum pond/flatwoods / / / / / / 3/9/11 Apalachicola NF 100.01 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 27 28.7 26.7 7.1 8.2 20 4/7/11 Camp Blanding MR 004 scrub 26.8 32.1 27.2 6.8 8.1 30 4/7/11 Camp Blanding MR 001 scrub 28.1 32.5 26.8 7.6 8.8 30 4/7/11 Camp Blanding MR 065 scrub 31.5 45.1 38.9 5.8 4.8 20 4/25/11 Camp Blanding MR 063 scrub 25.8 35.3 24 5.9 7.8 10 4/25/11 Gold Head Branch SP 001 sinkhole pond/upland sandhill 26.3 31 25.8 6.7 7.9 10 4/7/11 Gold Head Branch SP 001 sinkhole pond/upland sandhill 25.6 28.1 25.8 6.8 7.3 10 4/25/11 Ocala NF 003 ring pond/sandpine scrub 26.5 37.6 39.6 4.8 7.6 20 5/11/11 Ocala NF 022 sandpine scrub 28.8 41.7 39.6 4.7 8 20 5/11/11 Ocala NF 006 sandpine scrub 27.2 40.9 29.1 5 7.8 20 5/11/11 Ocala NF 020 sandpine scrub 32 42.4 30.6 5.6 8.1 20 5/11/11 Ocala NF 005 scrub 32.5 44.1 30.7 3.9 8.7 20 5/11/11 Ocala NF 021 scrub 34 35.3 33.3 6.5 8.2 20 5/11/11 Etoniah Creek SF 011 sandhill scrub 19.3 21.4 27.1 6.8 7.8 20 5/18/11 Etoniah Creek SF 007 sandhill scrub 23.8 24.6 37.6 6 7.7 20 5/18/11 Etoniah Creek SF 005 sandhill scrub 24.8 36.6 27.9 5.4 7 20 5/18/11 Etoniah Creek SF 006 flatwoods scrub ecotone 31.4 37.7 30.6 6.6 8.2 30 5/18/11 Etoniah Creek SF 022 scrub 26.4 37.9 26.7 5.5 7.8 20 5/18/11 Ocala NF 001 sandhill 22.4 26.1 21.3 6.9 8.1 10 2/23/12

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58 Table 2 4 Continued Location Habitat type Water temp Air temp Soil temp pH DO Conductivity collection date Ocala NF 014 sandhill 26.2 26.4 22.5 7.2 7.9 20 2/23/12 Ocala NF 017 sandhill 25.3 26.4 21.5 7 8.2 20 2/23/12 Osceola NF 001 flatwoods 26 24.7 25.2 4.8 8.1 10 5/4/11 Osceloa NF 002 flatwoods 26.2 27.8 23.3 6.6 8.3 20 4/5/12 Osceloa NF 003 flatwoods 26.8 31.6 23.3 6.9 8.6 10 4/5/12 Osceola NF 004 flatwoods 27.7 33.3 29.5 7.7 8.1 10 4/5/12 Big Bend WMA 002 sandpine scrub/flatwoods 22.1 33.1 22.3 7.4 8 20 5/2/11

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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). Outbrea ks 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 1996 2001 (Daszak et al. 1999; Green et al. 2002) and have been associa ted 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, (29.8254N, 81.9535 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

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60 whitish lesions on the dorsal surface of the body, hemorrhaging around the vent and ventral surface, and swollen edematous a bdomens on some specimens (Figures 3 2 and 3 3). Methods Freshly dead or dying tadpoles were placed on ice and later necropsied. Ea ch 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 a nd 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 fo ur 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 a ppeared 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.

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61 Results Combined samples for both L catesbeianus and L. s phenocephalus 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 Ran avirus 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 si milar physical signs as we documented herein for Pebble Lake. However, this is the first documented c ase 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 latte r 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

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62 exhibited similar pathological signs of infection and tested positive for R anavirus spp. Ranavirus mortali ty events are most likely under reported as like this one they may occur across short time span s and in remote areas

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63 Figure 3 1. Location of Pebble Lake in Mike Roess Gold Head Branch State Park, Clay County, Florida, USA, where Ranavirus spp. was detected during a mortality event

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

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

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66 CHAPTER 4 ANALYSIS OF PRESERVE D AMPHIBIAN SPECIMEN S TO TEST FOR DETECT ION 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 historica l range. Means and Tr avis (2007) determined that as of the 1990s D auriculatus is effectively extinct from the steephead r avines o n Eglin Air Force Base, covering Santa Rosa, Okaloos a, and Walton C ounties, Florida This area once sustained thriving populations of D auriculatus (Means and Travis, 2007) I n order to determine whether B dendrobatidis could have been the cause of this extirpation preserved specim ens of this species of concern were tested for the presence of B d endrobatidis ( Figure 4 1) Batrachochytrium dendrobatidis has been detec ted 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 ind ividuals of these populations have been properly preserved and stored. Methods A total of 20 southern dusky salamander ( D aurituculatis ) s pecimens from the Fl orida Museum of Natural History (FLMNH) H erpetology C ollection were used in this study. The speci mens were originally fixed with formalin and then put in 70% ethanol for preservation Since preservation all specimens have be en stored in glass jars in the H erpetology C ollection at the FLMNH

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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 yea r 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 Millhopper in Alachua County, five indivi individuals from Deep Springs Canyon in Bay County, and six individuals from Silver Glen Springs in Marion County were selected (T able s 4 1 and 4 2 ) Procedures from Soto Azat et al. (2010 ) we re 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 zo ospores 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 f or 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 o ther individuals collected at the same locality on the same day were also sampled but came back negative for B. dendrobatidis ( Figure 4 2)

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68 Discussio n The results of this study are inconclusive as they do show that B dendrobatidis was present on one specimen, however, 19 of the spec i mens 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 sampl e is not enough to draw conclusions about the impact that B dendrobatidis had on this population. T his species has declined throughout most of it s 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 a n amphibian population. T herefore it is possible to show that this pathogen could be the However, i n 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

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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

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70 Figure 4 2. Map showing collection location of each Desmognathus auriculatus specimen sampled for Batrachochytrium dendrobatidis Blue dots indicate sites where specimens that were neg a tive for B. dendrobatidis were collected. Red dots indicate the collection site of a single individual that was found to be positive for B. dendrobatidis

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71 Table 4 1. Desmognathus auriculatus specimens from the Florida Museum of Natural History that were sampled for Batrachochytrium dendrobatidis specimen # Bd swab # county month year chytrid 94067 610 Alachua March 1960 negative 94068 611 Alachua March 1960 negative 94069 612 Alachua March 1960 negative 97102 613 Alachua March 1965 negative 161436 614 Okaloosa December 1972 negative 161151 615 Okaloosa December 1972 negative 161158 616 Okaloosa December 1972 negative 161161 617 Okaloosa December 1972 negative 161162 619 Okaloosa December 1972 negative 165232 619 Bay February 1973 negative 157974 620 Bay February 1973 negative 157979 621 Bay February 1973 negative 157981 622 Bay February 1973 positive 157982 623 Bay February 1973 negative 166806 624 Marion February 1972 negative 164297 625 Marion February 1972 negative 164299 626 Marion February 1972 negative 164300 627 Marion February 1972 negative 163634 628 Marion February 1972 negative 163635 629 Marion February 1972 negative

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72 Table 4 2. Specific l ocality and number of Desmognathus auriculatus specimens from the Florida Museum of Natural History that were sampled for Batrachochytrium dendrobatidis Site Sample size Chytrid (positive/negative) Devil's Millhopper, Alachua Co. 4 0/4 Tom's Creek, Okaloosa Co. 5 0/5 Deep Springs Canyon, Bay Co. 5 (1/4) Silver Glen Springs, Marion Co. 6 0/6

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73 CHAPTER 5 SUMMARY AND CONSERVA TION SIGNIFICANCE This study has provided information on the occurrence of two amphibian pathogens, Batrachochytrium dendrobatidis and Ranavirus spp. in northern peninsular and panhandle Florida. B atrachochytrium dendrobatidis and Ranavirus spp. are present i n Florida amphibian populations, and therefore this study provides a starti ng point fo r future researchers interested in the ir distribution as well as the ecological effects of these pathogens on amphibian populations. Additionally this study provides insight into possible explanations of Florida amphibian population declines and extirpations. The results obta ined within the small sampling scale of this study suggest that much still remains to be determined regarding the distribution and prevalence of these pathogens Amphibians in colder, montane regions appear to be more vulnerable to B dendrobatidis than a mphibians in the southeastern U.S., as exemplified by the numerous outbreaks in montane tropical rainforests of Australia and Central America (Daszak et al., 1999). Studies performed by Hossack et al. (2010) in the cooler headwater streams of the United St ates show a low prevalence of B. dendrobatidis in amphibians as compared with wetland associated amphibians. This study suggests that B. dendrobatidis may be more likely to be found in slower flowing, warmer wetland habitats as opposed to snow melt cold, quick flowing headwater streams (Hossack et al., 2010). There is concern about the impact of pathogens in these pristine environments with multiple endemic species. It is speculated that the extinction of the golden toad ( Anaxyrus periglenes ) of Costa Rica and the decline of as many as seven Australian species may be the result of emerging pathogens, specifically B. dendrobatidis (Daszak et al., 1999). Unlike B. dendrobatidis there is no specific

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74 environment that has been shown to be more conducive to the p roliferation of Ranavirus spp (Gray et al., 2009). Buck et al. (2011) performed an experiment in which they found a negative correlation between the infective stage of B. dendrobatidis (which is a free living aquatic flagellated zoospore) and the zooplankter Daphnia magna This study supports the hypothesis that Daphnia magna consume B. dendrobatidis zoospores and could potentially be used as biological control of B. dendrobatidis (Buck et al., 2011). Despite this encouraging study by Buck et al. (2011), B dendrobatidis has already been found on every continent inhabited by amphibians. This means that this pathogen is already present in many populations of amphibians, even isolated popu lations. The fact that this pathogen is so widespread means that it must be carefully watched and survey s must be performed frequently. As discussed in Murray et al. (2011) the native range and rate of expansion of B. dendrobatidis remains unknown, leading it to potentially be detrimental to a wide number of host species across a wide geographic distribution. Amphibians are facing greater population declines than they ever have before, due to climate change, habitat destruction and modification, climate cha nge, 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). It is estimated that current extinction rates may be 211 times the bac kground extinction rate, with estimates of over 400 amphibian species facing rapid declines ( Blaustein et al., 2011 ). The future survival of amphibians depends on the proper and immediate implementation of conservation initiatives.

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75 It is important to recog nize that there are many different factors involved in the curre nt decline of amphibian species; many of these factors work in concert to create the massive extinction rate that has been observed (Blaustein et al., 2011). There are multiple stressors that are affecting amphibians on the molecular, physiological, individual, population, and community levels (Blaustein et al., 2011). It is critical for management strategies to recognize the synergistic effect of these stressors and to include conservation pla n s that employ this more holistic view point

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76 APPENDIX A STATE PARK SAMPLING PERMITS SAMPLING PERMIT A

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78 SAMPLING PERMIT B

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80 APPENDIX B ARC APPROVAL FORM IFAS Office of the Dean for Research 1022 McCarty Hall D P.O. Box 110200 Gainesville, FL 32611 0200 Telephone 352 392 1784 Fax 352 392 4965 Approval Number: 006 11WEC Approved: May 12, 2011 Expires: 2014 05 11 Responsible Faculty: Kenneth L. Krysko Address: Division of HerpetologyFlorida Muse um of Natural HistoryP.O. Box 117800Museum Road, Dickinson HallUniversity of FloridaGainesville, FL 32611 Project Title(s): Prevalence of chytrid fungus in amphibian species of Florida A project using animals has been approved for the research activity shown above. This approval has been granted by IFAS ARC at the University of Florida. This approval expires in three years. Any significant change in this approved animal project must be reviewed and approved by the ARC. Visit our web s ite for more details: http://research.ifas.ufl.edu/administration/animal_subjects_arc.html Dr. Charles Staples Research Foundation Professor Department of Animal Scien ces 204C "Red" Larson Building University of Florida Gainesville, FL 32611 352 392 1958 office phone 352 538 2789 cell phone 352 392 1931 FAX

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81 LIST OF REFERENCES Bentley, P. J. and A. R. Main. Zonal differences in permeability of the skin of some anu ran Amphibia. 1972. America n Journal of Physiology 223 : 361 363. Berger, L., R. Speare, P. Daszak, D. E. Green, A. A. Cuningham, C. L. Goggin, R. Slocombe, M. A. Ragan, A. D. Hyatt, K. R. McDonald, H. B. Hines, K. R. Lips, G. Marantelli, and H. Parkes. 1998 Chytridiomycosis causes amphibian mortality associated with population declines in the rainforests of Australia and Central America. Proceedings of the N ational Academy of Sciences 95: 9031 9036. Berger, L, R. Speare, and A. Kent. 1999. Diagnosis of chytr idiomycosis of amphibians by histological examination. Zoos Print Journal 1999:184 190. Bli hovde, W. B. 2006. Terrestrial movements and upland habitat use of go pher frogs in central Florida. Southeastern Naturalist 5 : 265 276 Blaustein, A. R., B. A. Han, R. A. Relyea, P. T. J. Johnson, J. C. Buck, S. S. Gervasi, and L. B. Kats. 2001. The complexity of amphibian population declines: understanding the role of cofactors in driving amphibian losses. Annals of the New York Academy of Sciences 1223: 108 119. Boyle, D. G., D. B. Boyle, V. Olsen, J. A. T. Morgan, and A. D. Hyatt. 2004. Rapid quantitative detection of chytridiomycosis ( Batrachochytrium dendrobatidis ) in amphibian samples using real time Taqman PCR assay. Diseases of Aquatic Organisms 60:141 148. Brem, F., J. R. Mendelson III, and K. R. Lips. 2007. Field sampling p rotocol for Batra chochytrium dendrobatidis from living a mphibians, usin g alcohol preserved s wabs. Version 1.0 (18 July 2007). Electronic document accessible at http://www.amphibians.org Conser vation International, Virginia, USA. Brunner, J. L., D. M. Schock, E. W. Davidson and J. P. Collins. 2004. Intraspecific Reservoirs: Complex Life History and the Persistence of a L ethal Ranavirus. Ecology 85 : 560 566. Buck, J. C., L. Truong, A. R. Blaustein. 2011. Predation by zooplankton on Batrachochytrium dendrobatidis : biological control of the deadly amphibian chytrid fun gus? Bi odiversity Conservation 20 : 3549 3553 Cheng, T. L., Rovito, S. M., Wake, D. B. Vredenburg, V. T. 201 1. Coincident mass extinction of neotropical amphibians with the emergence of the infectious fungal pathogen Batrachochytrium dendrobatidis Proceedings of the National Academy of Sciences 108 : 9502 9507. Chin char, V. G. 2002. Ranaviru ses (F amily Iridoviridae ): emerging cold blooded killers. Archives of Virology 147: 447 470.

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82 Chinchar VG, Essbauer S, He JG, Hyatt A, Miyazaki T, Seligy V, Williams T (2005). "Family Iridoviridae 145 162. In Fauquet CM, Mayo MA, Maniloff J, Desselburger U, Ball LA (eds). Virus Taxonomy, Eighth report of the International Committee on Taxonomy of Viruses. Academic Press, USA. Collins, J. P., Brunner, J. L., Jancovich, J. and D. M. Schock. 2004. A model host pathogen system for studying infectious disease dyn amics in amphibians: tiger salamaders ( Ambysotma tigrinum ) and Ambystoma tigrinum virus. Herpetological Journal 14:195 200. Cunningham, A. A., T. E. S. Langton, P. M. Bennett, J. F. Lewin, S. E. N. Drury, R. E. Gough and S. K. Macgregor. 1996. Pathological and microbiological findings from incidents of unusual mortality of the common frog ( Rana temporaria ). Philosophical Transactions: Biological Sciences 351 : 1539 1557. Daszak, P., L. Berger, A. A. Cunningham, A. D. Hyatt, D. E. Green, and R. Speare. 1999. E merging infectious diseases and amphibian population declines. Emerging Infectious Diseases 5: 735 748. Daszak, P., A. A. Cunningham, and A. D. Hyatt. 2003. Infectious disease and amphibian population declines. Diversity and Distributions 9: 141 150. Daszak, P., K. Lips, R. Alford, C. Carey, J.P. Collins, A. Cunningham, R. Harris, and S. Ron. Infectious Diseases. p p. 21 25 In: Gascon, C., J. P. Collins R. D. Moore D. R. Church J. E. McKay, and J. R. Mendelson III (eds). 2007. Amphibian Conservation Action Plan. IUCN/SSC Amphibian Specialist Group. Gland, Switzerland and Cambridge, UK. 64pp. Daszak, P., A. Strieby, A. A. Cunningham, J. E. Longcore, C. C. Brown, and D. Porter. 2004. Experimental evidence that the Bullfrog ( Rana catesbeiana ) is a potential car rier of chytridiomycosis, an emerging fungal disease of amphibi ans. Herpetological Journal 14: 201 207. Davis, A. K., M. J. Yabsley, M. K. Keel, and J. C. Maerz. 2007. Discovery of a novel alveolate pathogen affecting southern Leopard Frogs in Georgia: desc ription of the disease and host effects. EcoHealth 4: 310 317. DiGiacomo, R. F., and T. D. Koepsell. 1986. Sampling for detection of infection or disease in animal populations. Journal of American Veterinary Medical Association 189:22 23. Duellman, W. E. an d L. Trueb. 1986. Biology of Amphibians. McGraw Hill Book Company, USA. Duffus, A. L. J., B. D. Paul, K. Wozney, C. R. Brunetti, and M. Berrill. 2008. Frog virus 3 like infections in aquatic amphibian communities. Jour nal of Wildlife diseases 44 : 109 120.

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83 Gleason, F. H., S. E. Mozley Standridge, D. Porter D. G. Boyle and A. D. Hyatt 2007. Preservation of Chytridio mycota in culture collections. Mycological Research 3: 129 136. Gleason, F. H., S. K. Schmid t and A. V. Marano. 2010. Can zoosporic true fungi grow or survive in extr eme or stressful environments? Extremophiles 14: 417 425. Gray, M. J., D. L. Miller, A. C. Schumutzer, C. A. Baldwin. 2007. Frog virus 3 prevalence in tadpole populations inhabiting cattle access and non access wetlands in Tennessee, USA. Diseases of Aquatic Organisms 77:97 103. Gray, M. J., D. L. Miller and J. T. Hoverman. 2009. Ecology and pathology of amphibian ranaviruses. Di seases of Aquatic Organisms 87: 243 266. Green, D.E., K. A. C onverse, and A. K. Schrader. 2002 Epizootiolo gy of sixty four amphibian morbidity and mortality events in the USA. Ann uals: The New York Academy of Sciences 969: 323 339. Green, D. E. and C. K. Dodd, Jr. 2007. Presence of amphibian chytrid fungus Batrachochytrium dendrobatidis and other amphibian pathogens at warm water fish hatcheries in Southeastern North America. Herpetological Conservation and Biology 2 : 43 47. Hanselmann, R., Rodriguez, A., Lampo, M., Fajardo Ramos, L., Aguirre, A. A., Kilpatrick, A. M., Rodriguez, J. P. an d P. Daszak. 2004. Presence of an emerging pathogen of amphibians in introduced bullfrogs ( Rana catesbeiana ) in Venezuela. Biological Conservation 120: 115 119. Hossack, B. R., M. J. Adams E. H. Campbell Grant C. A. Pearl J. B. Bettaso W. J. Barichivich W. H. Lowe K. True J. L. Ware, and P. S. Corn. 2010. Low prevalence of chytrid fungus ( Batrachochytrium dendrobatidis ) in amphibians of U.S. headwater streams Journal of Herpetology 44 : 253 260. Hyatt A., D. Boyle, V. Olsen, D. Boyle, L. Berger, D. Obe ndorf, A. Dalton, K. Kriger, M. Hero, H. Hines, R. Phillott, R. Campbell, G. Marantelli, F. Gleason, A. Colling. 2006. Diagnostic assays and sampling protocols for the detection of Batrachochytrium dendrobatidis Diseases of Aquatic Organisms 73:175 192. Johnson, M.L., and R. Speare. 2003. Survival of Batrachochytrium dendrobatidis in water: quarantine and disease control implications. Emerging Infectious Diseases 9: 922 925. Johnson, B. K., and J. L. Christiansen. 1976. The food and food habits of Blanchar cricket frog, Acris crepitans blanchardi (Amphibia, Anura, Hylidae), in Iowa Journal of Herpetology 10 : 63 74. Kiesecker, J.M., L.K. Belden, K. Shea, and M.J. Rubbo. 2004. Amphibian decline and emerging disease. Scientific American 92:138 147.

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84 Kraus, F. 1999. Alien Reptiles and Amphibians: a scientific compendium and analysis. Springer Science, Japan. Lips, K. R., F. Brem, R. Brenes, J. D. Reeve, R. A. Alford, J. Voyles, C. Carey, L. Livo, A. P. Pessier, and J. P. Collins. 2006. Emerging infectious dis ease and the loss of biodiversity in a Neotropical amphibian community. Proceedings of the Na tional Academy Science USA 103: 3165 3170. Longcore, J. R., J. E. Longcore, A. P. Pessier, and W. A. Halteman. 2007. Chytridiomycosis widespread in anurans of North eastern United States. Journal of Wildlife Management 71:435 444. 2009. Development of multi species indicators for th e Nevada wildlife action p lan. Ecological Indicators 9 : 1030 1036. Mazzoni, R., A. A. Cunningham, P. Daszak, A. Apolo, E. Perdomo, and G. Speranza. 2003. Emerging pathogen of wild amphibians in frogs ( Rana catesbeiana ) farmed for international trade. Emerging Infectious Diseases 9: 995 998. Means, D. B., R. C. Means, and R. P. M. M eans. 2008. Petition to list the striped newt, Notophthalmus perstriatus as a federally threatened species under the Endangered Species Act of 1973. Petition to the U.S. Fish and Wildlife Service. Means, D. B. and J. Travis. 2007. Declines in ravine inhabiting dusky salamanders of the Southeastern US coastal plain Southeastern Naturalist 6 : 83 96. Mitchell, J. C., and D. E. Green. 2002. Chytridiomycosis in two species of ranid frogs in the southeastern United States. Joint Meeting of the American Society of Ichthyologists and Herpetolo Study of Amphibians and Repti les, 4 8 July 2002, USA. Mitchell, K. M., T. S. Churcher T. W. J. G arner and M. C. Fisher. 2008. Persiste nce of the emerging pathogen Batrachochytrium dendrobatidis outside t he amphibian host greatly increases the probability of host extinction. Procee dings of The Royal Society 275: 329 334. Mohneke, M. and M. O. Rodel. 2009. Declining amphibian populations and possible ecological consequen ces a review. Salamandra 45 : 203 210. M urray, K. A., R. W. R. Retallick, R. Puschendorf, L. F. Skerratt, D. Rosauer, H. I. McCallum, L. Berger, R. Speare, and J. VanDerWal. 2011. Assessing spatial patterns of disease risk to biodiversity: implications for the management of the amphibian pathoge n Batrachochytrium dendrobatidis Journal of Applied Ecology 48: 163 173. Mutschmann, F., L. Berger, P. Zwart, and C. Gaedicke. 2000. Chytridiomycosis on amphibians first report from Europe. Berliner und Munchener T ierarztliche Wochenschrift 113: 380 383.

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85 M uths, E., D. S. Pilliod and L. J. Livo. 2008. Distribution and environmental limitations of an amphibian pathoge n in the Rocky Mountains, USA. Biological Conservation 141 : 1484 1492. Olson, D., and K. Ronnenberg. 2007. Batrachochytrium dendrobatidis mapping project. PARC: Partners in Amphibian and Reptile Conservation. . 7 Feb 2009. Ouellett, M., I. Mikaelian, B. D. Paul, J. Rodrigue, and D. M. Green. 2005. Historical evidence of widespread chytrid infec tion in North American amphibian popula tions. Conservation Biology 19: 1431 1440. Parris, M. J., and D. R. Baud. 2004. Interactive effect of a heavy metal and chytridiomycosis on gray treefrog larvae ( Hyla chrysoscelis ). Copeia 2004:344 350. Parris, M. J., and T. O. Cornelius. 2004. Fungal pathogen causes competitive and developmental stress in larval amphibian communities. Ecology 85:3385 3395. Pearl, C. A., and D. E. Green. 2005. Rana catesbeiana (American Bullfrog), Chytridiomycosis. Herpetological Revie w 36:305 306. Pessier, A. P. 2008. Management of disease as a threat to amphibian conservation. International Zoo Yearbook 42:30 39. Pessier, A. P., D. K. Nichols, J. E. Longcore, and M. S. Fuller. 1999. Cutaneous chytridiomycosis in poison dart frogs ( Dendrobates spp.) and White's tree frogs ( Litoria caerulea ). Journal of Veterinary Diagnostic Investigation 11:194 199. Phillott, A. D., R. Speare, H. B. Hines, L. F. Skerratt, E. Meyer, K. R. McDonald, S. D. Cashins, D. Mendez, and L. Berger. 2010. Minimi sing exposure of amphibians to pathogens during field studies. Diseases of Aquatic Organisms. Piotrowski, J. S., S. L. Annis, and J. E. Longcore. 2004. Physiology of Batrachochytrium dendrobatidis a chytrid pathog en of amphibians. Mycologia 96: 9 15. Pound s, J.A., M.R. Bustamante, L.A. Coloma, J.A. Consuegra, M.P.L. Fogden, P.N. Foster, E. La Marca, K.L. Masters, A. Merino Viteri, R. Puschendorff, S.R. Ron, G.A. Snchez Azofeifa, C.J. Still, and B.E. Young. 2006. Widespread amphibian extinctions from epide mic disease driven by global warming. Nature 439:161 167. Rizkalla, C. E. 2009. First reported detection of Batrachochytrium dendrobatidis in Florida, USA. Herpetological Review 40:189 190. Rollins Smith, L. A., J. P. Ramsey J. D. Pask L. K. Reinert and D. C. Woodhams. 2011. Amphibian immune defenses against chytridiomycosis: impacts of changing environments. Integrative and Comparative Biology 51 : 552 562.

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86 Rothermel, B. B., S. C. Walls, J. C. Mitchell, C. K. Dodd, Jr., L. K. Irwin, D. E. Green, V. M. Vaz quez, J. W. Petranka, and D. J. Stevenson. 2009. Widespread occurrence of the amphibian chytrid fungus Batrachochytrium dendrobatidis in the southeastern USA. Di seases of Aquatic Organisms 82: 3 18. Rosenblum, E. B., J. E. Stajich N. Maddox and M. B. Eisen 2008. Global gene expression profiles for life stages of the deadly amphibian pathogen Batrachochytrium dendrobatidis Proceedings of the National Academy of Sciences of the Un ited States of America 105 : 17034 17039. Shoemaker, V. H., S. S. Hillman, S. D. Hillyard, D. C. Jackson, L. L. McClanahan, P. C. Withers, and M. L. Wygoda. 1992. Exchange of water, ions, and respiratory gases in terrestrial amphibians. In M. E. Feder and W. W. Burggren (Eds.), Environmental Physiology of the Amphibians (pp. 125 150 ). The University of Chicago Press, USA. Skerratt, L. F., L. Berger, H. B. Hines, K. R. McDonald, D. Mendez, and R. Speare. 2008. Survey for detecting chytridiomycosis in all Australian frog populations. Di seases of Aquatic Organisms 80: 85 94. Soto Azat C., B. T. Clarke, J. C. Poynton, and A. A. Cunningham. 2010. Widespread historical presence of Batrachochytrium dendrobatidis in African pipid frogs. Diversity and Distributions 16: 126 131. St Amo u r, V., T. W. J. Garner A. I. Schulte Hostedde and D. Lesbarreres 2010. Effects of two amphibian pathogens on the developmental stability of green frog s. Conservation Biology 24 : 788 794. Steiner, S. L. and R. M. Lehtinen. 2008. Occurrence of the amphibian pathogen Batrachochytrium dendrobatidis in Blanchard Acris crepitans blanchardi ) in the U.S. Midwest. Herpetological Review 39:193 196. Stuart, S.N., J.S. Chanson, N.A. Cox, B.E. Young, A.S.L. Rodrigues, D.L. Fischman, and R.W. Waller. 2004. Status and trends of amphibian declines and extinc tions worldwide. Science 306:1783 1786. Teacher, A. G. F., A. A. Cunningham and T. W. J. Garner. 2010. Assessing the long term impact of Ranavirus infection in wild common frog popul ations. Animal Conservation 13: 514 522. Vondersaar, M. E. and D. F. Stiff ler. 1989. Renal function in amphibians: a comparison of strictly aquatic and amphibious species with observations on the effects of anesthesia. Journal of Comparitive Bio chemistry and Physiology 94 : 243 248. Weldon, C., L. H. du Preez, A. D. Hyatt, R. Mull er, and R. Speare. 2004. Origin of the amphibian chytrid fungus. Emerging Infectious Diseases 10:2100 2105.

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87 Wells, K. D. 2007. The Ecology and Behavior of Amphibians. The University of Chicago Press, USA. Woodhams, D. C., R. A. Alford, and G. Marantelli. 2 003. Emerging disease of amphibians cured by elevated body temperature. Di seases of Aquatic Organisms 55: 65 67. Young, B. E., S. N. Stuart, J. S. Chanson, N. A. Cox, and T. M. Boucher. 2004. Disappearing Jewels: The Status of New World Amphibians. NatureServe, Virginia.

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88 BIOGRAPHICAL SKETCH Sarah Reintjes Tolen was born in Siler City, North Carolina and lived with her parents in Chapel Hill, North Carolina until entering college. She attended Grady Brown Elementary, A. L. Stanback Middle Scho ol and Chapel Hill High School. Upon graduating high school she studied Biology and Environmental Science at the University of North Carolina at Chapel Hill. She received her Bachelor of Arts degree in Biology and Environmental Science in 2009 under the gu idance of Dr. Kenneth J. Lohmann. She graduated with honors from UNC CH for her work on the navigational abilities of loggerhead sea turtles. While at UNC CH she worked with Dr. Lohmann on loggerhead sea turtle behavior and navigational abilities from 2008 2009. During the summer of 2008 she studied abroad with The School for Field Studies in Baja Mexico, working on sea turtle conservation. In 2010 she entered the University of Florida Department of Wildlife Ecology and Conservation and began to work on he amphibian disease ecology in the state of Florida. Upon completion of her masters degree she will spend a year working in the Biology Department at the University of Florida, researching the regenerative abilities of amphibian s pecies with application to biomedical engineering She will then con tinue her studies with a focus on behavioral ecology at the Biology Department at the University of Florida obtaining her PhD She hopes to dedicate her life to th e conservation of reptil e a nd amphibian species in the tropics