thesis may 19 final draft

8/7/2019 Thesis May 19 Final Draft

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EVALUATION OF A NOVEL DETECTION METHOD FOR RANAVIRUS INWATER SAMPLES FROM PINE MOUNTAIN WILDLIFE MANAGEMENT

AREA, LETCHER COUNTY KENTUCKY

By:

Matthew R. Pettus

Thesis Approved:

Chair, Advisory Committee

Member, Advisory Committee

Member, Advisory Committee

Dean, Graduate School


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STATEMENT OF PERMISSION TO USE

In presenting this thesis in partial fulfillment of the requirements for aMaster's degree at Eastern Kentucky University, I agree that the Library shall

make it available to borrowers under rules of the Library. Brief quotations from this

thesis are allowable without special permission, provided that accurateacknowledgment of the source is made.

Permission for extensive quotation from or reproduction of this thesis may begranted by my major professor, or in his absence, by the Head of Interlibrary

Services when, in the opinion of either, the proposed use of the material is forscholarly purposes. Any copying or use of the material in this thesis for financial gain

shall not be allowed without my written permission.

Signature _____________________________________

Date _________________________________________


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EVALUATION OF A NOVEL DETECTION METHOD FOR RANAVIRUS INWATER SAMPLES FROM PINE MOUNTAIN WILDLIFE MANAGEMENT

AREA, LETCHER COUNTY KENTUCKY

By:

Matthew R. Pettus

Bachelors of ScienceElmhurst CollegeElmhurst Illinois

2006

Submitted to the Faculty of the Graduate School ofEastern Kentucky University

in partial fulfillment of the requirementsfor the degree of

MASTER OF SCIENCEApril, 2010


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DEDICATION

This thesis is dedicated to my parents, Robert and Debbie Pettus. Without their

help this thesis would never have been completed. To my father, you have

always been there to support me. I thank you from the bottom of my heart. To

my mother, without the lessons you taught me I would not be here. Little woman,

you will always be loved and missed. You both have made me who I am, thank

you.


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ACKNOWLEGDEMENTS

I would like to thank my advisor and graduate committee chair, Dr. Paul Cupp.

Your knowledge and resources have helped make this project work. I would like

to thank my co-committee chair Dr. Marcia Pierce for all the long hours and

countless times you have stepped in to remedy my research woes. Also, for all

your help in getting this thesis written and ready for review. Dr. Stephen Richter

thank you for the use of your lab, as well as always making me feel welcome

even though I wasnt part of your lab. Finally, Id like to say thank you to all of the

graduate students. This would have been an impossible road without your

camaraderie.


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ABSTRACT

Throughout the world amphibian populations have declined over the last

ten years. Many factors have been implicated, yet emerging diseases are

currently having the largest effects on amphibian populations. This research was

intended to develop a protocol using water samples to detect Ranavirus, a

recently emerging infectious agent, from environmental water samples at Pine

Mountain Wildlife Management Area in Letcher County, Kentucky. Ranaviruses

are dsDNA viruses that have been implicated in localized amphibian declines.

Possible reservoirs for Ranaviruses include adult amphibians, reptiles, and

fishes. Direct transmission has been well documented and indirect transmission

is highly possible. Centrifugal filters were used to concentrate water samples

from a volume of 15 ml down to 200 l. PCR was performed on the concentrated

water samples and PCR products were separated using 1% agarose gel

electrophoresis. Tissue samples from animals living in each pond were also

taken for comparison to the water samples. Total samples obtained included 38

water samples and 98 tissue samples. All of the samples tested negative for

Ranavirus. To determine the lowest concentration of virus detectable by this

novel system, double distilled water (ddH2O) and pond water was seeded with

Ranavirus at a known concentration. This system could detect 13.3 PFU/l in

ddH2O and 106.4 PFU/l in pond water. While there may have been several

factors involved in this result, it is most likely that during the sampling period

Ranaviruswas not present in the Pine Mountain Wildlife Management Area.


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TABLE OF CONTENTS

CHAPTER Page

INTRODUCTION.................................................................................................. 1

Ranavirus Characteristics.................................................................... 3

Viral Reservoirs ................................................................................... 6

Possible Amphibian Reservoirs ................................................. 6

Non-Amphibian Reservoirs........................................................ 7

Ranavirus Transmission ..................................................................... 7

Sampling............................................................................................. 9

II.METHODS....................................................................................................... 11

Field Sampling................................................................................... 11

Laboratory Data Collection ................................................................ 12

Lowest Detectable level of Virus.............................................. 12

DNA Extraction from Water and Tissue Samples .................... 13

Amplification of DNA................................................................ 14

III. RESULTS...................................................................................................... 15

Controls ............................................................................................. 15

Lowest Detectable Level of Virus....................................................... 15

Water Samples .................................................................................. 15

Tissue Samples ................................................................................. 16

IV. DISCUSSION................................................................................................ 17

LITERATURE CITED ......................................................................................... 24


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APPENDIX.. ................................................. 31

VITA ................................................................................................................... 38


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LIST OF TABLES

TABLE

1. Latitude and longitude of ponds with and

without recorded die-offs and parkingplaces close to the ponds.............................................................................. 33

2. The pond number, species collected, andthe collection number of the samples fromJuly 10, 2008 ................................................................................................ 34

3. The pond number, species collected, andthe collection number of the samples fromJuly 25, 2008 ................................................................................................ 35

4. The pond number, species collected, andthe collection number of the samples fromAugust 8, 2008 .............................................................................................. 36

5. The pond number, species collected, andthe collection number of the samples fromAugust 26, 2008 ............................................................................................ 36

6. The pond number, species collected, andthe collection number of the samples fromSeptember 11, 2008 ...................................................................................... 37

7. The pond number, species collected, andthe collection number of the samples fromSeptember 28, 2008 ....................................................................................... 37

8. The pond number, species collected, andthe collection number of the samples fromOctober 9, 2008.............................................................................................. 38

9. The pond number, species collected, andthe collection number of the samples fromOctober 23, 2008.......................................................................................... 38


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LIST OF FIGURES

FIGURE

1. Lowest detectable level of virus in water samples using

20l, 10l, 5l, and 2.5l of Ranavirusstock virus ........................................ 16


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Chapter I

Introduction

Amphibians are the most threatened vertebrate class, more so than

eitherbirds or mammals (Gascon et al. 2007). There are 1856 endangered or

threatened species of amphibians; approximately 32% of amphibians are

globally threatened, as compared to 12% of birds and 23% of mammals

(Gascon et al. 2007). At least 2468 amphibian species (43.2%) are

experiencing some form of population decrease, whereas

28 species (0.5%)

are increasing and only 1552 species (27.2%) are stable. Currently, 1661 or

29.1% of amphibian species have an unknown trend (Gascon et al. 2007).

In the last decade, global amphibian population declines have steadily

gained more attention (Blaustein and Wake 1990). Many causes have been

proposed for this decline including increases in UV radiation (Broomhall et al.

2000), introduced species (Adams et al. 1999), and emerging infectious

disease (Daszak et al. 2003; Collins and Storfer 2003; Wake and Vredenburg

2008). While each of these is important, emerging infectious disease (EID)

has been implicated in single and multiple population die offs and is of

increasing importance each year (Jancovich et al. 2003; Daszak et al. 1999;

Carey et al. 2003). Declines have been increasingly seen in pristine areas

with little disturbance (Lips et al. 2003). EIDs have been found in both

pristine and impacted areas, and as the presence of these diseases increase


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we will see far-reaching consequences in our amphibian populations (Lips et

al. 2003; Jancovich et al. 2003).

Emergent infectious diseases are defined as diseases that are

expanding their range, taking on new hosts, or are newly discovered (Daszak

et al. 1999). There are two main emerging diseases implicated in the decline

of amphibian populations: chytrid fungus and Iridovirusinfections.

Chytridiomycosis is a fungal disease caused by Batrachochytrium

dendrobatidis, which was first described in 1998 (Berger et al. 1998). This

fungus attacks keratinized tissue including several skin layers in adult

amphibians and the mouth parts of larval amphibians. Three hypotheses

have been proposed to explain the death of amphibians infected with chytrid

fungus: damage to the skin reduces the ability of amphibians to cutaneously

breathe and/or osmoregulate; there is a release of fungal toxins that are

absorbed; or a combination of these events occur (Berger et al. 1998, Pessier

et al. 1999). Often highly virulent pathogens are dependent on the host

density and as the pathogen suppresses the host population, transmission

stops. This allows the host population to rebound (Anderson and May 1986;

Dobson and May 1986). This does not appear to be the case with chytrid.

The amphibian larval stages have been shown to live on after the death of the

adults, implying that the larvae may be reservoir hosts (Daszak et al. 2000).

This may allow chytrid to remain in reduced amphibian populations. In

addition, chytrid can live as a saprophyte, or survive on decaying matter,


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which may explain the lack of recolonization after amphibian die offs (Lips

1998; Lips 1999).

The second group of pathogens implicated in amphibian declines is the

Iridoviruses. Unlike chytrid fungus, the Iridoviruses are not a single species.

They are a group of viruses that infect fish, salamanders, frogs, and reptiles

(Mao et al. 1999, Johnson et al. 2008, Gray et al. 2009, Ariel et al. 2009).

Those affecting amphibians are in the genus Ranavirus. Several ranaviruses

are common in amphibians and mortalities have been reported at all latitudes,

and in most major families of anurans and urodeles (Carey et al. 2003;

Daszak et al.2003).

RanavirusCharacteristics

Ranaviruses were first isolated by Granoff et al. (1965) from Lithobates

pipiens. Ranaviruses are in the family Iridoviridae (Eaton et al. 2007).

Iridoviridae contains 5 genera. The Iridoviruses and Chloridoviruses infect

invertebrates. The Ranaviruses, Megalocystiviruses, and Lymphocystiviruses

infect vertebrates (Chinchar et al. 2009).

Ranaviruses have a diameter of 160-200 nm and have a linear

double-stranded (ds) DNA genome (Fauquet et al. 2005, Williams et al.

2005). Their nucleoprotein core is made up of a single coiled filament 10 nm

wide. The capsid symmetry is skewed with a T=133 or 147. The unit

genome size is ~105 kbp with a G+C content of ~54% (Fauquet et al. 2005).

Ranaviruses have a distinctive icosahedral shape that is often visible in the


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cytoplasm of infected cells in electron microscope images (Chinchar and

Hyatt 2008). The genome encodes for 100 gene products with several genes

thought to increase virulence and help in evasion of the host immune

response (Chinchar 2002, Chinchar et al. 2009). The viral infection and

immune response can be so overwhelming that cellular death can occur

within hours of infection (Chinchar et al. 2003; Williams et al. 2005).

Ranaviruses are known to infect anurans, urodeles, reptiles, and bony

fish (Mao et al. 1997, Williams et al. 2005). Three species of Ranavirusare

known to infect amphibians. These are Frog Virus 3 (FV3), Bohle iridovirus

(BIV), and Ambystoma tigrinumvirus (ATV) (Chinchar et al. 2005). The major

capsid protein (MCP) of these viruses make up approximately half the virus

weight and is highly conserved (Hyatt et al. 2000).

The major capsid protein (MCP) is an area that has been studied

extensively (Mao et al. 1999). While Hyatt et al. (2000) found that the MCP is

highly conserved there is still variation amongst isolated viruses. The genetic

variation in the MCP of ATV shows geographic differences (Ridenhour and

Storfer 2008), while FV3 seems to be more genetically conserved over its

geographic range (Schock et al. 2008). Despite this variation, Mao e al.

(1997) found that primers created for FV3 were able to adhere to nine wild

strain iridoviruses and that those strains were more closely related to FV3

than other iridoviruses. To this day the MCP remains one of the most

commonly used genes in identifying ranaviruses.


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Viral Reservoirs

Possible Amphibian Reservoirs

Amphibians are a likely reservoir for the ranaviruses. Due to the

biphasic life cycle in amphibians it is possible that reservoirs are both aquatic

and terrestrial. In aquatic environments it is probable that species with larva

that mature in >1 season, have incomplete metamorphosis, and/or are highly

aquatic adults are the most likely reservoirs (Gray et al. 2009). Gray et al.

(2007) reported that 57% of overwintering tadpoles of Lithobates catesbeiana

were infected with Ranavirus. Many individuals of overwintering tadpoles do

not show physical signs of infection (Miller et al. 2009). This trend has also

been shown in Desmognathus quadramaculatus(Gray et al. 2009). Because

of this, both larval and adult amphibians may help Ranaviruspersist in the

environment (Duffus et al. 2008)

In situations where amphibian larvae do not over winter, adults may

remain as a reservoir for Ranavirus. Sub-lethal infections in tadpoles have

been shown to continue after metamorphosis (Ariel et al. 2009). Brunner et

al. (2004) was able to show that ATV-infected Ambystoma tigrinumcould

survive through metamorphosis and that 7% of returning salamanders the

following year were infected with ATV. In the late 1990s, Jancovich et al.

(1997) hypothesized that tadpoles and asymptomatic adults act as the

reservoir for Ranavirus. As more research has come to light, it appears that

pre- and post-metamorphic amphibians are likely to be the preeminent

reservoirs for Ranavirus.


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2006). Skin contact of one second between an infected animal and an

uninfected animal has been shown to cause disease (Brunner et al. 2007).

Many amphibians are cannibalistic (Alford 1999, Rosen 2007, Pizzatto and

Shine 2008). Cannibalism may be the consumption of any life stage including

eggs, which Ranavirusalso has been found to infect (Tweedell and Granoff

1968, Alford 1999). Studies have shown that predation on infected

individuals can led to infection (Harp and Petranka 2006, Brunner et al. 2007).

Those species that have cannibalistic phenotypes may exhibit a lower

occurrence of this phenotype due to poor survivorship (Pfennig et al. 1991).

Indirect transmission is infection caused by virus circulating through

the environment. This can be through the water or through soil. Salamander

larvae exposed to ATV-infected individuals, but without direct contact, have

become infected (Greer et al. 2008). Harp and Petranka (2006) were able to

induce infection by exposing Lithobates sylvaticusto sediment from an active

natural die-off. These studies suggest that Ranavirusthat is shed into the

surrounding environment can infect new hosts. For indirect transmission to

occur, Ranavirusmust be able to persist in the environment. However, this

aspect of Ranavirusecology is not well understood. Jancovich et al. (1997)

showed that ATV could persist for up to 2 weeks. However, this may be

contingent on some level of moisture being present. Brunner et al. (2007)

found that sediment that was inoculated with ATV and then dried for four days

did not cause infection, while soil inoculated and kept moist caused 87%

mortality in salamanders. The length of viability is unknown for ranaviruses,


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during or after die-offs cannot aid us in understanding transmission or sub-

lethal infections.

Recently, a non-lethal method of sampling has been developed. St.

Amour and Lesbarreres (2007) have been able to detect the Ranavirusin toe

clippings. This has allowed investigators to determine the presence or

absence of the Ranavirusin field samples and determine if lab raised animals

are free of Ranavirus. Since the results are a positive or negative for the

presence of Ranavirusand animals need to be caught, restrained, and toes

removed, a faster method is possible.

The purpose of my research is the detection of Ranavirusin Kentucky

ponds using a water based non-lethal system. I plan to use this method as a

quick and effective means to identify the presence of infection, and hopefully

provide a tool for continued research into indirect transmission of ranaviruses.

My novel detection method using water samples will be compared to tissue

samples processed using the St. Amour and Lesbarreres (2007) protocols.

Water samples will allow quick field sampling and reduce the overall cost of

sampling, while maintaining the speed and reliability of PCR-based detection

methods.


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Chapter II

Materials and Methods

Sampling was done at Pine Mountain Wildlife Management Area

(PMWMA). This management area is one of Kentuckys last contiguous and

least disrupted forests. It is comprised of 4,849 acres located in the far

southeast corner of Kentucky in Letcher and Harlan counties. In 2002, more

than 50 ponds were built by Kentucky Department of Fish and Wildlife as

wildlife watering holes. When these ponds were monitored in 2004 and 2007

for amphibian populations it was found that there were active die-offs at

several ponds.

For this study, ponds were selected from the 20 best amphibian ponds

at PMWMA (J. MacGregor, personal communication, August 30 2008). Of

these 20 ponds, 11 had no previous die-offs and the remaining 9 had

recorded die-offs suspected to be from Ranavirusbut that were never verified

(J. MacGregor, personal communication, May 16 2008). Prior to entering the

field, 5 ponds were randomly selected from each of these two groups. These

ten ponds became the sampled ponds.

Field Sampling

At each pond, water and tissue samples were taken. Water samples

were collected from July 10, 2008 to October 23, 2008 at two week intervals.

Samples were taken at approximately 75 mm under the surface of the pond at


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the edges where amphibian adults and larvae were often seen. Water

samples consisted of 45 ml of pond water in a 50 ml plastic test tube. These

water samples were immediately placed on ice for transport to the laboratory.

Animals were obtained using a dip net. Tissue samples were taken from

adults, larvae, and eggs. Tissue samples were obtained from adults by

removing the third phalanx at the second joint with a pair of suture scissors.

The toe was immediately placed in 1 ml tubes containing 100% ethanol and

placed on ice. Tissue samples from larvae were obtained by tail clipping.

Approximately 2 cm of the distal tail tip were cut using suture scissors. The

samples were placed in 1ml tubes containing 100% ethanol and placed on

ice. While the traditional means of sampling larvae is tissue homogenation,

tail clippings have been used in many amphibian orders to test for Ranavirus

(St. Amour and Lesberreres 2007). Eggs were sampled by excising 3 to 4

eggs from the egg mass. Eggs were placed directly in 1 ml tubes containing

100% ethanol and placed on ice for transport to the laboratory.

Processing Samples in the Laboratory

Minimum Ranavirus Detectable Concentrations

A consistent control volume was needed for each following experiment.

A volume of 20 l was chosen because it was a volume that consistently

produced PCR products that were detected and was easily visible on gel

electrophoresis. These controls were at a concentration of 106.4 plaque

forming units (PFU)/l.


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Double distilled water was used to produce viral dilutions, which were

then used to determine the lowest concentration detectable by the water

testing method. Stock Ranaviruswas obtained from ATCC (Manassas, VA)

at a concentration of 8 x 107 PFU/ml. Double distilled water was seeded with

20 l, 10 l, 5 l, 2.5 l, 1.25 l, 0.625 l, and 0.3125l to create a total

volume was 15 ml. Concentrations of Ranaviruswere 106.4 PFU/l, 53.2

PFU/l, 26.6 PFU/l, 13.3 PFU/l, 6.65 PFU/l, 3.325 PFU/l, and 1.662

PFU/l respectively. The 15 ml sample was then pipetted into an Amicon 15

filter unit (Millipore; Billerica, MA) with a pore size of 3 kDa. The Amicon Ultra

15 centrifugal filter units containing the water samples were centrifuged until

the volume was reduced to 200 l. Using the reduced volume of 200 l, a

DNA extraction was performed using a MiniElute Virus Spin Kit(Qiagen;

Valencia, CA).

DNA Extraction from Water and Tissue Samples

Pond water samples were vortexed to homogenize the contents. The

maximum volume of water an Amicon 15 can process is 15 ml. Therefore,

sub-samples of 15 ml were taken from each 45 ml sample. These were pre-

filtered using a 0.45 micron syringe filter (Corning; Corning NY) to remove

bacteria, detritus, and other debris. The filtrate was then placed in an Amicon

15 filtration unit. Each Amicon 15 was centrifuged until the volume was

reduced to 200 l. DNA extraction was done using the MiniElute Virus Spin

Kit(Qiagen; Valencia, CA).


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DNA extractions from tissue samples were done using a DNeasy

Blood and Tissuekit (Qiagen; Valencia, CA).

Amplification of DNA

Forward and reverse primers for the MCP developed by Mao et al.

(1996) were used to detect the presence of Ranavirusthrough PCR. PCR

conditions during amplification followed protocols described in Mao et al.

(1996). Reactions were incubated for 5 minutes at 94C followed by 28

cycles consisting of 94C for 1 minute, 45C for 2 minutes, 55C for 3 minutes,

and a single cycle of 55C for 5 minutes. Gel electrophoresis was performed

using 1% agarose gels (50 ml volume). Each gel was produced using 45ml

distilled water, 5 ml of 10X TAE buffer, and 0.5g agarose. Samples were

prepared for electrophoresis by adding a 10 l aliquot of each PCR product

sample to 6.7 l distilled water and 3.3 l orange buffer (Invitrogen; Carlsbad,

CA). Samples were vortexed and 10 l of each were pipetted in a single lane

on the gel. 5 l of a 100 bp ladder was placed in the first and last wells of

each gel. Gels were electrophoresed at 100 volts and stained with ethidium

bromide (0.02%) before examination using a UV light box.


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Chapter III

Results

Controls

Initial testing determined Ranaviruscould be recovered from ddH2O

samples at 53.2 PFU/l, 106.4PFU/l, 159.6PFU/l, and 266.0PFU/l. A

consistent control volume was needed for each following experiment. These

controls each contained 106.4 PFU/l.

Lowest Concentration of RanavirusDetected

The sample of 2.5l was the lowest volume visible on gel

electrophoresis (Figure 1). The 2.5l of Ranavirusin 15ml distilled water was

a concentration of 13.3 PFU/l.

To determine the lowest detectable concentration in pond water, pond

water samples were seeded to concentrations of 266 PFU/l, 106.4 PFU/l,

53.2 PFU/l, 26.6 PFU/l, and 13.3 PFU/l. The lowest concentration that

had PCR products detectable by electrophoresis was 106.4 PFU/l.

Water Samples

A total of 38 water samples were taken from 10 ponds at PMWMA.

Ranaviruswas not detected in any environmental sample at PMWMA.

Six randomly selected environment samples from PMWMA were seeded with

virus stock solution at 106.4 PFU/l to determine if there were PCR inhibitors


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present in the water samples. Of these samples, four were visible after gel

electrophoresis. PCR and gel electrophoresis were attempted twice more on

the samples that tested negative. These samples in subsequent tests

continued to test negative.

Figure 1. Lowest detectable level of virus in water samples using 20l, 10l,5l, 2.5l of Ranavirusstock virus (8 x 107 PFU/ml). L-DNA Ladder,E-Empty Lane

Tissue Samples

A total of 90 tissue samples were taken from the experimental

ponds at PMWMA. No evidence of Ranavirusinfection were found in any

tissue samples collected at PMWMA.

L E E E E 2.5l 5l 10l20l L


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Chapter IV

Discussion

I was able to create a novel protocol for the detection of Ranavirusin

both seeded ddH20 and seeded pond water that is non-destructive. This

method differs from other water sampling protocols for emerging infectious

diseases (EID) in that it uses lower water sample volumes (see Krishtein et al.

2007). The goal for this novel detection method was to be used to detect

Ranavirusin small ephemeral ponds where the volume of water can be very

low, amphibian larval densities can be high, and Ranavirusinfection is

common. However, we were unable to detect Ranavirusin any

environmental water samples, so more research needs to be conducted to

further develop this method for use in the field.

This is the first attempt to create a water based sampling method for

the detection of Ranavirus. However, water sampling has been used to

detect other EIDs. Kirshtein et al. (2007) proposed a sampling protocol for

the detection of Batrachochytrium dendrobatidisin water and sediment.

Using 0.2 m filters and sampling between 64 l and 50 ml, they were able to

detect as little as 0.06 zoospores, or 10 copies of DNA. However, the lowest

concentration detectable in 50 ml of water was 30 zoospores per liter. The

protocol proposed for Ranavirusdetection in this paper uses lower volumes of

water, samples were filtered after collection from the field, and isolates a

smaller causative agent. The use of smaller water volumes may have caused


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an increase in false negative results. Samples were collected in the field and

put on ice until they could be returned to the lab for processing. This reduced

the amount of equipment and time needed to sample. Ranaviruses are

between 160-200 nm in size; because of their small size they are difficult to

filter out of water. The Amicon 15 (Millipor; Billerica, MA) was selected

because it had the ability to retain Ranavirusabove the filter and was easily

used in the lab, however it is limited to a 15 ml sample size. In future studies

using similar protocols for water sampling, it may be necessary to filter

multiple sub-samples of 15 ml through a single filter to reduce the chance of

false negative results. My goal was to keep the field sampling quick and the

laboratory work simple and thereby keep the cost, both in time and money

low, however using this system I was unable to detect Ranavirusin any

environmental water samples.

For water sampling to work an EID must be able to persist in the

environment. While this is not well established for Ranavirus, Brunner et al.

(2007) and Jancovich et al. (1997) showed Ranavirusinfections could be

initiated through inoculated sediment. Harp and Petranka (2006) also

showed that Ranaviruscould be transmitted through water between

Lithobates sylvaticatadpoles without contact. The novel detection method

presented here also anecdotally supports that Ranaviruscan be detected in

water, though only in seeded samples.

Ranaviruswas detected in concentrations as low as 13.3 PFU per l in

ddH2O and 106.4PFU per l in pond water, or13.3 x 103 PFU ml-1 and 10.6 x


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10 4 PFU ml-1respectively. The concentration of Ranavirusthat is biologically

significant is not known. However, Rojas et al. (2005) used a laboratory study

to suggest that an environmentally relevant concentration of Ranavirusvirions

in water inhabited with infected salamanders may be between 103 to 104 PFU

ml-1. The concentrations detected by my novel detection method in pond

water falls just above the projected biologically significant concentrations of

Ranavirus. It may be possible to reduce the lowest detectable volume by

adjusting PCR parameters, and reducing the chance of inhibitors in the pond

water. Inhibitors in the water are a possible problem with this system.

When six randomly selected samples of pond water were seeded with

20 l ofRanavirusat 8 x 107 PFU/ml, then filtered and processed normally,

only four tested positive for Ranavirus. This could lead to a high level of false

negatives. If Ranaviruswas present in these samples it is possible that the

primers do not adhere well to the wild strain of Ranaviruspresent at PMWMA.

However, this is a highly conserved region. Mao et al. 1997 tested nine

Iridovirus wild strains and was able to detect each of them using these

primers. In subsequent tests, positive results were not consistent and the

lowest detectable level of Ranavirusfluctuated. This may indicate that there

are PCR inhibitors in the water samples. However, more field research is

necessary to determine if these projected concentrations are indeed

biologically significant and whether or not inhibitors in the water are affecting

the lowest concentration detectable in pond water samples.


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The prevalence of Ranavirusin known infected areas has been

studied. In China, Ranaviruswas found to infect 5.7% of adults and 42.5% of

larva of Rana dybowskii(Xu et al. 2010). Gray et al. (2007) found that

between 15% and 40% of Rana clamitanstadpoles were infected with

Ranavirusdepending on time of the year and whether there was cattle access

to the wetland. These high percentages of infected animals were not found in

sampling sites at PMWMA. There may be several explanations for these

results. First, it is possible that sublethally infected animals tested negative

due to reduced levels of Ranaviruswithin their tissues. Brunner et al. (2007)

found that Ambystoma tigrinumlarvae that survived with sub-lethal infections

of ATV did not always test positive. Seven of ten sublethally infected animals

tested positive only once out of three tests. However, only a small portion of

individuals were sub-lethally infected. While this could have reduced the total

number of animals that tested positive it is not likely to cause all animals

sampled to test negative. Ninety-eight animals were sampled in this study

with most being tadpoles. With others reporting high percentages of tadpoles

being infected (Gray et al. 2007, Xu et al. 2010) it is not probable that sub-

lethal infections were a factor in PMWMA negative persistence of Ranavirus.

A second possibility is that through multiple exposures to Ranavirus,

the population at PMWMA is gaining immunity to Ranavirus. Gantress (2006)

found that Ranaviruswas cleared from Xenopus laevismore quickly after a

second exposure, and Majji et al. (2006) found that exposure to one

Ranavirusgave partial immunity to other Ranaviruses. Die-offs have


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occurred over the last six years at PMWMA (J. MacGregor, personal

communication, August 30 2008) and may be reducing a portion of the

susceptible adults and increasing the number of metamorphs that have

cleared a Ranavirusinfection. It is also possible that breeding has shifted to

reduce the presence of genes that lead to high-susceptibility to Ranavirus.

Teacher et al. (2009) found that Ranavirushad imposed selection for

particular haplotypes and that some amphibians were adapting to the

presence of Ranavirus. While more research would be necessary to

determine if this was occurring at PMWMA, it is possible that susceptible

individuals are dying and being replaced by animals that are less susceptible.

It is also possible that Ranaviruswas not detected at PMWMA

because it was not present at the time of sampling. Gray et al. (2007) found

that there was a higher percentage of infected tadpoles in the fall and winter.

Sampling in this study was conducted from July to October. This time frame

was chosen based on observations made at PMWMA (J. MacGregor,

personal communication, May 16 2008). Die-offs were observed historically

in May at PMWMA. Since sampling was done after May there is a possibility

that the concentration of Ranavirusin the water was lower than it would have

been if sampling was done earlier in the year. At PMWMA die-offs in wood

frogs and green frogs were most common. Due to my later sampling it is

possible that wood frog die-offs had already occurred and the level of

Ranaviruspresent was reduced. In this study a large amount of green frogs

were captured and tested without any evidence of Ranavirusbeing present.


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In future work at PMWMA it will be necessary to sample as early as possible

to determine if wood frog die-offs occur and whether or not there is an

increase in Ranavirusin water samples during this time. However, it is

possible that winter and fall infections and die-offs occur also at this location.

The lack of Ranavirusat PMWMA may have been due to the

misdiagnosis of the causative agent behind the die-offs that have occurred

there. The diagnosis of the causative agent was based on a few physical

symptoms and the fact that Ranavirushad been found in elsewhere in

Kentucky. While only one pond in Kentucky has had confirmed amphibian

Ranavirusdie-offs, Ranavirushas been confirmed in Terrapene carolinain

Ohio County, Kentucky via oral swabs (J. MacGregor, personal

communication, May 12 2009).

A recently emerging disease exhibits similar symptoms to Ranavirus.

Perkinsus-like disease, or Dermomycoidessp. causes lethargy, bloat, and

hemorrhaging of the skin (Cook 2009). While these previous symptoms are

similar to both fungal and viral EIDs, Perkinsus-like disease also causes

erratic swimming, and Davis et al. (2007) found that they were able to hand

catch infected animals. These symptoms have not been reported in the die-

offs at PMWMA; however, it will be necessary to continue to sample at

PMWMA during an active outbreak to determine definitively the causative

agent of the die-offs.

Another possibility is that after several drought years Ranavirusno

longer persisted in PMWMA. Die-offs may be affected by rainfall (J.


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23

MacGregor, personal communication, August 23 2007). Kentucky had lower

than average rainfall in Letcher County in both 2006 and 2007 (Robert Watts,

unpublished data). When sampling was done in 2008, Letcher County had

average rainfall for the year (National Weather Service Forecast Office).

Brunner el at. (2007) found that sediment that had dried for four days could

not cause infection. During sampling in 2008, 7 of 10 ponds dried for some

period of time over the sampling period, despite an average amount of

rainfall. It is a possibility that due to the occurrence of drought over a two

year span that Ranavirusdid not persist at PMWMA.

While many questions about the findings of this study persist, I am

confident that I have laid the ground work for water sampling to detect

Ranavirus. In the future it will be necessary to further refine this sampling

method by optimizing the PCR reaction and eliminating the possibility of

inhibitors in the water samples. Consistent long term sampling of PMWMA

will help to determine a base line of Ranavirusactivity in the area and

increase the sample size of both water and tissue collected. A positive

Ranavirusdie-off needs to be sampled to determine if Ranaviruscan be

detected in actual environmental samples and what concentrations of

Ranavirusare actually biologically relevant. Despite the future work needed, I

believe that this study has begun to establish the ground work necessary to

found a quick, inexpensive, and non-destructive testing method using water

samples that will elucidate the ecology of Ranaviruses.


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Literature Cited

Adams MJ. (1999). Correlated factors in amphibian decline: Exotic species

and habitat change in western Washington. J Wildl Man63: 1162-1171

Alford RA (1999). Ecology: resource use, competition and predation. In:McDiarmid RW, Altig R (eds) Tadpoles: the biology of anuran larvae.University of Chicago Press, Chicago, IL, p 240278

Allender MC, Fry MM, Irizarry AR, Craig L, Johnson AJ, Jones M (2006).Intracytoplasmic inclusions in circulating leukocytes from an easternbox turtle (Terrapene Carolina carolina) with iridoviral infection. J WildlDis42:677684

Anderson RM, May RM. (1998). The invasion, persistence and spread ofinfectious diseases within animal and plant communities. Philos TransR Soc Lond B Biol Sci314: 533-70

Ariel E (1997). Pathology and serological aspects of Bohle iridovirusinfections in six selected water-associated reptiles in NorthQueensland. PhD thesis, James Cook University,Townsville

Ariel E, Kielgast J, Svart HE, Larsen K, Tapiovaara H, Jensen B, HolopainenR (2009). Ranavirus in wild edible frogs Pelophylaxkl. esculentusinDenmark. Dis Aquat Org85:714

Berger L, Speare R, Daszak P, Green DE, Cunningham AA, Goggin CL.(1998). Chytridiomycosis causes amphibian mortality associated withpopulation declines in the rainforests of Australia and Central America.Proc Natl Acad Sci USA 95: 9031-6

Blaustein AR, Wake DB. (1990). Declining amphibian populations: a globalphenomenon? Trends Ecol Evol5:203204

Broomhall SD, Osborne WS, Cunningham R. (2000). Comparative Effects ofAmbient Ultraviolet-B Radiation on Two Sympatric Species ofAustralian Frogs. Conser Bio14: 420-427

Brunner JL, Schock DM, Davidson EW, Collins JP. (2004). IntraspecificReservoirs: Complex Life History and the Persistence of a LethalRanavirus. Ecology85(2): 560566


35/48

25

Brunner JL, Richards K, Collins JP (2005). Dose and host characteristicsinfluence virulence of ranavirus infections. Oecologia144:399406

Brunner JL, Schock DM, Collins JP (2007). Transmission dynamics of theamphibian ranavirus Ambystoma tigrinumvirus. Dis Aquat Org77:87

95

Carey C (1993). Hypothesis concerning the causes of the disappearance ofboreal toads from the mountains of Colorado. Conser Bio7:355-62

Carey C, Pessier AP, Peace AD (2003). Pathogens, infectious disease, andimmune defenses. In: Semlitsch RD(ed) Amphibian conservation.Smithsonian Institute, Washington, DC, p 127136

Chinchar VG (2002). Ranaviruses (family Iridoviridae): emerging cold-bloodedkillers. Arch Virol147:447470

Chinchar VG, Bryan L, Wang J, Long S, Chinchar GD (2003). Induction ofapoptosis in frog virus 3-infected cells. Dis Aquat Org87: 243266

Chinchar VG, Essbauer S, He JG, Hyatt A, Miyazaki T, Seligy V, Williams T(2005). Iridoviridae. In: Fauguet CM, Mayo MA, Maniloff J,Desselberger U, Ball LA (eds) Virus taxonomy: 8th report of theinternational committee on the taxonomy of viruses. Elsevier, London,p 163175

Chinchar VG, Hyatt AD, Miyazaki T, Williams T (2009). Family Iridoviridae:poor viral relations no longer. In: Van Etten JL (ed) Current topics inmicrobiology and immunology, Vol 328. Lesser known large dsDNAviruses. Springer-Verlag, Berlin, p 123170

Collins JP, Storfer A (2003). Global amphibian declines: sorting thehypotheses. Divers Distrib9:8998

Cook JO (2009). An alveolate agent in southeastern amphibians.Southeastern Partners in Amphibian and Reptile Conservation,Disease, Pathogens and Parasites Task Team, Information Sheet #5.

Daszak P, Berger L, Cunningham AA, Hyatt AD, Green DE, Spear R (1999).Emerging infectious diseases and amphibian population declines.Emerg Infect Dis5:735748

Daszak P, Cunningham AA, Hyatt AD. (2000). Emerging infectious diseasesof wildlife threats to biodiversity and human health. Science287: 443449


36/48

26

Daszak P, Cunningham AA, Hyatt AD (2003). Infectious disease andamphibian population declines. Divers Distrib9: 141150

Davis AK, Yabsley M, Keel K, Maerz JC (2007). Discovery of a novelpathogen affecting southern leopard frogs in Georgia: description of

the disease and host effects. EcoHealth4:310-317

Dobson AP, May RM. (1986). Disease and conservation. In: Soul M, editor.Conservation biology: The science of scarcity and diversity.Sunderland (MA):Sinauer Assoc Inc.: 345-65

Duffus ALJ, Pauli BD, Wozney K, Brunetti CR, Berrill M (2008). Frog virus 3like infections in aquatic amphibian communities. J Wildl Dis44:109120

Eaton HE, Metcalf J, Penny E, Tcherepanov V, Upton C, Brunetti CR (2007).

Comparative genomic analysis of the family Iridoviridae: re-annotatingand defining the core set of iridovirus genes. Virol J4:117

Fauquet CM, Mayo MA, Maniloff J, Desselberger U, Ball LA. (2005). VirusTaxonomy: VIIIth Report of the International Committee on Taxonomyof Viruses. New York: Academic Press.

Gahl MK, Calhoun AJK (2010). The role of multiple stressors in Ranavirus-caused mortalities in Arcadia National Park wetlands. Can J Zoo88:108-121

Gantress J, Maniero GD, Cohen N, Robert J (2003). Development andcharacterization of a model system to study amphibian immuneresponses to iridoviruses. Virology311: 254262

Gascon C, Collins JP, Moore RD, Church DR, McKay JE, Mendelson, JR III(eds). (2007). Amphibian Conservation Action Plan. IUCN/SSCAmphibian Specialist Group. Gland, Switzerland and Cambridge, UK:64

Granoff A, Came PE, Rafferty KA (1965). The isolation andproperties ofviruses from Rana pipiens: their possible relationship to the renaladenocarcinoma of the leopard frog. Ann NY Acad Sci126:237255

Gray MJ, Miller DL, Schmutzer AC, Baldwin CA (2007). Frog virus 3prevalence in tadpole populations inhabiting cattle-access and non-access wetlands in Tennessee, USA. Dis Aquat Org77: 97103

Gray MJ, Miller DL, Hoverman JT (2009). First report of Ranavirusinfectinglungless salamanders. Herpetol Rev40:316319


37/48

27

Greer AL, Berrill M, Wilson PJ. (2005). Five amphibian mortality eventsassociated with ranavirus infection in south central Ontario, Canada.Dis Aquat Org67:914

Greer AL, Briggs CI, Collins JP (2008). Testing a key assumption of hostpathogen theory: density and disease transmission. Oikos117:16671673

Greer AL, Brunner JL, Collins JP (2009). Spatial and temporal patterns ofAmbystoma tigrinumvirus (ATV) prevalence in tiger salamandersAmbystoma tigrinum nebulosum. Dis Aquat Org85:16

Harp EM, Petranka JW (2006). Ranavirus in wood frogs (Rana sylvatica):potential sources of transmission within and between ponds. J WildlDis42:307318

Hedrick RP, McDowell TS, Ahne W, Torhy C, Dekinkelin P (1992). Propertiesof three iridovirus-like agents associated with systemic infections offish. Dis Aquat Org13:203209

Hyatt AD, Gould AR, Zupanovic Z, Cunningham AA and others (2000).Comparative studies of piscine and amphibian iridoviruses. Arch Virol145:301331

Hyatt AD, Williamson M, Coupar BEH, Middleton D and others (2002). Firstidentification of a ranavirus from green pythons (Chondropythonviridis). J Wildl Dis38:239252

Jancovich JK, Davidson EW, Morado JF, Jacobs BL, Collins JP. (1997).Isolation of a lethal virus from the endangered tiger salamanderAmbystoma tigrinum stebbinsi. Dis Aquat Org31:161-7

Jancovich JK, Davidson E, Seiler A, Jacobs BL, Collins JP. (2001).Transmission of the Ambystoma tigrinumvirus to alternative hosts. DisAquat Org46:159163

Jancovich JK, Mao J, Chinchar VG, Wyatt C, Case ST, Kumar S,Valente G,Subramanian S, Davidson EW, Collins JP, Jacobs BL. (2003).Genomic sequence of a ranavirus (family Iridoviridae) associated withsalamander mortalities in North America. Virology316:90103


38/48

28

Johnson AJ, Pessier AP, Wellehan JF, Childress A, Norton TM, Stedman NL,Bloom DC, Belzer W, Titus VR, Wagner R, Brooks JW, Spratt J,Jacobson ER. (2008). Ranavirus Infection Of Free-Ranging AndCaptive Box Turtles And Tortoises In The United States. J. Wildl. Dis44(4): 851-63

Kristten JD, Anderson DW, Wood JS, Longcore JE, Voytek MA (2007).Quantitative PCR detection of Batrachochytrium dendrobatidisDNAfrom sediments and water. Dis aquat Org77: 11-15

Langdon JS, Humphrey JD, Williams LM, Hyatt AD, Westbury HA (1986).First virus isolation from Australian fish: an iridovirus-like pathogenfrom redfin perch, Perca fluviatilisL. J Fish Dis9:263268

Lips KR. (1998). Decline of a tropical montane fauna. Conser Bio12:106-17

Lips KR. (1999). Mass mortality and population declines of anurans at anupland site in Western Panama. Conser Bio13:117-25

Lips K, Green ED, Papendick R (2003). Chytridiomycosis in wild frogs fromSouthern Costa Rica. J Herpetol37:215218

Majji S, LaPatra S, Long SM, Sample R, Bryan L, Sinning A, Chinchar VG(2006). Rana catesbeianavirus Z (RCV-Z): a novel pathogenicranavirus. Dis Aquat Org73:111

Mao J, Hedrick RP, Chinchar VG. (1997). Molecular characterization,sequence analysis, and taxonomic position of newly isolated fishiridoviruses. Virology229:212220

Mao J, Green DE, Fellers G, Chinchar VG (1999). Molecular characterizationof iridoviruses isolated from sympatric amphibians and fish. Virus Res63:4552

Marschang RE, Braun S, Becher P (2005). Isolation of a ranavirus from agecko (Uroplatus fimbriatus). J Zoo Wildl Med36:295300

Miller DL, Gray MJ, Rajeev S, Schmutzer AC, Burton EC, Merrill A, BaldwinCA (2009). Pathological findings in larval and juvenile anuransinhabiting farm ponds in Tennessee,USA. J Wildl Dis45:314324

Pearman PB, Garner TWJ (2005). Susceptibility of Italian agile frogpopulations to an emerging strain of Ranavirusparallels populationgenetic diversity. Ecol Lett8:401408


39/48


40/48

30

Westhouse RA, Jacobson ER, Harris RK, Winter KR, Homer BL (1996).Respiratory and pharyngo-esophageal iridovirus infection in a gophertortoise (Gopherus polyphemus). J Wildl Dis32:682686

Williams T, Barbosa-Solomieu V, Chinchar GD (2005). A decade of advancesin iridovirus research. In: Maramorosch K, Shatkin A (eds) Advances invirus research, Vol 65. Academic Press, New York, p 173248

Xu K, Zhu D, Wei Z, Schloegel LM, Chen X, Wang X (2010). BroadDistribution of Ranavirus in Free-Ranging Rana dybowskiiinHeilongjiang, China. EcohealthDOI: 10.1007/s10393-010-0289-y


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APPENDIX


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Table 1. Latitude and longitude of ponds with and without previous die-offs

and parking places close to the ponds.

Pond #

Latitude andLongitude for

Parking

Latitude andLongitude of Pond

LocationsPrevious Die-Offs

Documented

52 N 57 04' 14.8'' N 57 04' 11.1'' N

W 82 49' 10.0'' W 82 49' 08.0''

26 N 37 02' 36.3'' N 37 02' 41.6'' N

W 82 52' 48.0'' W 82 52' 45.8''

23 N 37 02' 23.1'' N 37 02' 19.5'' N

W 82 53' 04.5'' W 82 53' 04.5''

18 N 37 01' 47.8'' N 37 01' 44.8'' N

W 82 54' 00.4'' W 82 53' 57.6''

13 N 37 01' 32.5'' N 37 01' 31.9'' N

W 82 54' 49.9'' W 82 53' 49.7''

47 N 37 03' 47.2'' N 37 03' 43.1'' Y

W 82 50' 21.7'' W 82 50' 20.4''

44 N 37 03' 42.6'' N 37 03' 39.6'' Y

W 82 50' 38.3'' W 82 50' 37.6''

40 N 37 03' 11.3'' N 37 03' 10.9'' Y

W 82 51' 43.3'' W 82 51' 45.0''

36 N 37 03' 04.6'' N 37 03' 02.8'' Y

W 82 51' 57.6'' W 82 51' 01.0''

27 N 37 02' 42.6'' N 37 02' 38.3'' Y

W 82 52' 34.3 W 82 52' 34.3


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Table 2. The pond number, species collected, and the collection number of

the samples from July 10, 2008.

Pond Number Species Sampled Collection Number

26 N. viridescens Tail (adult) 26-1R. clamitans tail clip (Tadpole) 26-2

R. clamitans tail clip (Tadpole) 26-3

N. viridescens Tail (adult) 26-4



13 Rana clamitans toe (Adult) 13-1



Rana clamitans toe (Adult) 13-4

36 R. clamitans tail clip (Tadpole) 36-1


R. clamitans tail clip (Tadpole) 36-3R. clamitans tail clip (Tadpole) 36-4

40 Rana clamitans toe (Sm Adult) 40-1




44 Rana clamitans toe (Adult) 44-2

Rana clamitans (Adult) 44-2

47 Dry


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the samples from July 25, 2008.


52 Ambystoma sp. (larva) 52b-1Ambystoma sp. (larva) 52b-2

Rana clamitans tail clip (Tadpole) 52b-3


26 Notophthalmus viridescens Tail 26b-1

Rana clamitans toe (Sm. Meta) 26b-2


Rana eggs 26b-4

23 Dry

18 Dry


Rana clamitans toe (Adult) 13b-2

Hyla sp. (Tadpole) 13b-327 Notophthalmus viridescens Tail 27b-1

Rana clamitans toe (Adult) 27b-2


Rana clamitans toe (Adult)*** 36b-2Rana clamitans tail clip(Tadpole)*** 36b-3

Rana clamitans tail clip (Meta)*** 36b-4Rana clamitans tail clip(Tadpole)*** 36b-5




Rana clamitans tail clip (Tadpole) 40b-444 Rana clamitans toe (Adult) 36-1

Notophthalmus viridescens Tail 36-2


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the samples from August 8, 2008.


52 Ambystoma sp. 52C-1Notophthalmus viridescens 52C-2

Rana clamitans 52C-3

26Emerging Tads- Ranaclamitans 26C-1

Notophthalmus viridescens 26C-2

Rana clamitans 26C-3

23 Rana clamitans 23C-1

13 Rana clamitans tadpoles 13C-1


27 Rana clamitans 27C-1

Rana sylvatica 27C-2

36 Notophthalmus viridescens 36C-1

40 Rana clamitans tadpole 40C-1


Rana clamitans tadpole 40C-3

Rana clamitans adult 40C-4

44 Notophthalmus viridescens 44C-1

47 Dry


the samples from August 26, 2008.

Pond Number Species SampledCollectionNumber

52 Bufo sp. Tadpoles 52D-1

23 Dry

13 Notophthalmus viridescens 13D-1

Rana clamitans Tadpoles 13D-2

Rana clamitans Tadpoles 13D-3

36 Rana clamitans 36D-1

47 Dry


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the samples from September 11, 2008.


13 Rana clamitans Tadpole 13E -1

Rana clamitans Tadpole 13E -2

27 Rana clamitans Adult 27E -1

36 Rana clamitans Tadpole 36E -1

Rana clamitans Tadpole 36E -2

47 Dry


the samples from September 28, 2008.


52 Ambystoma sp. Larva 52F-1

Ambystoma sp. Larva 52F-2

26 Rana clamitans Tadpoles 26F-1

Rana clamitans Tadpoles 26F-2

Rana clamitans Tadpoles 26F-3

18 Dry

13 Rana clamitans Tadpoles 13F -1

Rana clamitans Tadpoles 13F -2Rana clamitans Tadpoles 13F -3

Rana clamitans Tadpoles 13F -4

27 Rana clamitans Adult 27F -1

44 Rana clamitans Adult 44F-1

47 Dry


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the samples from October 9, 2008.


52 Ambystoma sp. 52F2-1

Ambystoma sp. 52F2-2

26 Dry

23 Dry

18 Dry

13 Rana clamitans Tadpoles 13F2 -1

Rana clamitans Tadpoles 13F2 -2

Rana clamitans Tadpoles 13F2 -3

27 Rana clamitans Adult 27F2 -1

47 Dry


the samples from October 23, 2008.


13 Rana clamitans Tadpole 13G-1

47 Dry


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Vita

Mathew Robert Pettus was born on January 7, 1979 to Robert and

Debbie Pettus in St. Louis, Missouri. At age 3, he and his family moved to the

northwest suburbs of Chicago. He went to elementary school at Central

School, junior high school at Westfield Junior High School, and high school at

Lake Park High School. In 1997, Matthew graduated high school and began

a long college career. In 2006, Matthew graduated from Elmhurst College

with a B.S. in biology. Matthew began work on a masters degree at Eastern

Kentucky University later that year. In 2010, Matthew received his M.S.

degree in biology from Eastern Kentucky University.

thesis may 19 final draft

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