© 2016 Lauren P. Kane

INVESTIGATING THE EPIDEMIOLOGY OF TERRAPENE HERPESVIRUS 1 IN FREE- RANGING EASTERN BOX TURTLE POPULATIONS

LAUREN P. KANE

THESIS

Submitted in partial fulfillment of the requirements for the degree of VMS - Comparative Biosciences in the Graduate College of the University of Illinois at Urbana-Champaign, 2016

Urbana, Illinois

Master’s Committee:

Assistant Professor Matthew C. Allender, Chair Associate Professor David Bunick Clinical Associate Professor Jennifer Langan Assistant Professor Elizabeth Driskell

ABSTRACT

Herpesviruses are ubiquitous, well described pathogens in captive chelonians worldwide, but their importance on free-ranging populations are less defined. In this thesis, a quantitative PCR was developed that detected a 58 base pair segment of the DNA polymerase gene segment of

Terrapene herpesvirus 1 (Order: Herpesvirales; Family: Herpesviridae; Subfamily:

Alphaherpesvirinae; Genus: Scutavirus). This assay was used to estimate prevalence of herpesvirus infection in 409 free-ranging eastern box turtles (Terrapene carolina carolina) from

Tennessee and Illinois. The overall prevalence in this study population was 31.3% (95% confidence interval: 27-36%), with a significantly higher prevalence in July (52.3%; 95% CI: 41-

59%) compared to May (13.3%; 95% CI: 5-15%) and September (34.4%; 95% CI: 29-47%).

Clinical signs recorded in box turtles were not significantly associated with herpesvirus infection and may be attributed to a latency period. The work presented in this thesis aids in characterizing the epidemiology of herpesvirus in chelonians.

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TABLE OF CONTENTS

CHAPTER 1: DISEASE IMPACT ON TURTLE POPULATIONS……………………………..1

CHAPTER 2: LITERATURE REVIEW………………………………………………………….4

CHAPTER 3: DEVELOPMENT AND VALIDATION OF QUANTITATIVE PCR FOR DETECTION OF TERRAPENE HERPESVIRUS 1 UTILIZING FREE-RANGING EASTERN BOX TURTLES (TERRAPENE CAROLINA CAROLINA)……………………………………...23

CHAPTER 4: PREVALENCE OF TERRAPENE HERPESVIRUS 1 IN FREE-RANGING EASTERN BOX TURTLES (TERRAPENE CAROLINA CAROLINA)…………………………37

CHAPTER 5: CONCLUSIONS AND FUTURE DIRECTIONS……………………………….56

CHAPTER 6: BIBLIOGRAPHY……………………………………………………………..…58

CHAPTER 1

DISEASE IMPACT ON TURTLE POPULATIONS

Disease events can play an important role in wildlife long-term sustainability and several environmental, physiological, and pathogenic factors affect the impact of disease on wildlife

(Cunningham and Daszak, 1998; Pounds et al., 2006; Skerratt et al., 2007). Changes in the environment attributed to anthropogenic factors, such as, fragmentation, habitat loss, pollution and pesticide use, are a contributor to disease epidemiological changes (Daszak et al., 1999;

Holladay et al, 2001; van Dijk, 2013). Ectothermic animals highly dependent on their environment, such as reptiles, may be even more susceptible to disease, when compared to birds or mammals, if their immunologic function becomes impaired due to environmental changes

(Donaldson and Echternacht, 2005; Ultsch, 2006). Lack of information describing the environmental effects in wild turtle populations makes it is imperative to describe disease epidemiology in the face of these various environmental factors.

Across the world, it has been noted that several aquatic and terrestrial chelonians are experiencing population declines (Lederle et al., 1997; Johnson et al., 2008; Willoughby et al.,

2013). According to the International Union for Conservation of Nature (IUCN) Red List of threatened species, 8 different chelonian species are already extinct, 31 chelonian species are listed as critically endangered, 44 species are listed as endangered, 60 species are listed as vulnerable, 36 species are listed as near threatened, and 1 species is listed as conservation dependent in 2015 (van Dijk et al., 2014). Thirty-two chelonian species are noted to have decreasing population trends. Since 2013, 20 chelonian species changed their Red List status; two were downgraded to critically endangered, eight were newly listed as endangered, five were

listed as vulnerable, two were listed as nearly threatened, and only three were upgraded to least concern (van Dijk, 2014). In summation, 143 of the 230 turtle species, or 62% of all currently recognized turtle and tortoise species are either already extinct or threatened with extinction, making it one of the most imperiled vertebrate groups.

One species in particular, the North American box turtle (Terrapene carolina carolina), has been experiencing significant ongoing gradual population declines leading to a disturbing

Red List status downgrade, in 2011, from a near threatened to vulnerable status (van Dijk, 2013).

Specific causes of the population decline are mostly attributed to human-induced factors that include international pet trade, habitat fragmentation, vehicle strike, and vegetation fires (Henry,

2003; Hester et al., 2008; Nazdrowicz et al., 2008; Schrader et al., 2010; van Dijk, 2013). Due to their longevity and low fecundity, populations rely heavily on juvenile survivorship and low adult mortality to maintain stable populations. Therefore, a disease outbreak affecting adult survivorship could have catastrophic long-term population effects (Budischak et al., 2006;

Currylow et al., 2011; Donaldson and Echternact, 2005; Ultsch, 2006). Additionally, life history characteristics, such as sexual maturity and reproduction, may be delayed due to climate changes, which ultimately affect resource availability. These negative impacts have the potential to put box turtles at an increased risk for slower population recovery (Budischak et al., 2006;

Ernst et al., 1994; Klemens, 2000). Therefore, anthropogenic changes, such has habitat fragmentation, climate changes, or international elimination for the pet trade, play a substantial role in box turtle populations survivability. While box turtle population declines are most likely due to a combination of factors, disease outbreaks and enzootics has been increasingly been reported in chelonian species.

Described diseases affecting population stability in free-ranging box turtles are both infectious and non-infectious in nature (Brown and Sleeman, 2002; Gibbons and Steffes, 2013).

Several box turtle health investigations have been described, including the establishment of hematologic and plasma reference values for free ranging box turtles in Maryland and Tennessee

(Adamovicz et al., 2015; Way Rose and Allender, 2011). Other investigations have identified threats to free-ranging box turtle populations including aural abscessation, nutritional disorders, trauma, and infectious disease (Brown and Sleeman, 2002; DeVoe et al., 2004; Feldman et al.,

2006; Sim et al., 2015; Schrader et al., 2010). Captive box turtle populations are likewise vulnerable to infectious diseases. Herpesvirus, in particular, has been emerging across the United

States in eastern box turtles (Kane et al., 2016; Sim et al., 2015; Yonkers et al., 2015). It is vital to further evaluate herpesvirus enzootics seen in box turtles and to ascertain the disease prevalence and the potential role in population declines.

This thesis aims to address key gaps in the epidemiology of herpesvirus disease in the North

American box turtles. The following objectives will be presented:

1. Develop a quantitative (real-time) polymerase chain reaction (PCR) assay for detection of

Terrapene herpesvirus 1 DNA in chelonians.

2. Characterize the epidemiology and determine prevalence of Terrapene herpesvirus 1 in free- ranging eastern box turtles in Tennessee and Illinois.

CHAPTER 2 LITERATURE REVIEW

Importance

Infectious diseases have been an increasing contributor to global population declines in wild animals (Daszak et al., 1999; Friend 2006). Some diseases are well studied, such as ranavirus in amphibians (North et al., 2015) and West Nile virus in birds (Hale, 2015), while others are poorly understood or are underreported. Disease events may be underreported due to lack of recognition or minimal long-term research opportunities, and may be a significant threat to population survival, which increases as population sizes decrease (Daszak, 1999). In reptiles, there are fewer reports of infectious diseases than those of mammals or birds. To understand the role of disease on wild reptiles, it is important to understand the epidemiology of pathogens causing morbidity and mortality. The family Herpesviridae has been widely reported in numerous reptile species and identified as a pathogen that can lead to population decline in wild reptiles (Bicknese et al., 2010; Haines et al., 1977; Jacobson et al., 1986; Johnson et al., 2005;

Teifke et al., 2000), however there is a paucity of information on the epidemiology.

Box Turtle Natural History

Eastern box turtles, T. carolina carolina, are among the most terrestrial North American

Emydid turtles (Donaldson and Echternacht, 2005). Occasionally, box turtles utilize aquatic habitats during high temperature days and are most active during this time (Donaldson and

Echternacht, 2005). Eastern box turtles are active from April to October, followed by an extended state of winter dormancy (Currylow et al., 2011). These chelonians have a small natural home range, 8-40 hectacres, (Hester et al., 2008) and are long-lived with life expectancy of

greater than 50 years (Budischak et al., 2006). Eastern box turtles rely heavily on juvenile survivorship and low adult mortality in order to maintain stable populations (Budischak et al.,

2006; Klemens, 2000). If adult and juvenile survivorship decreases due to disease, populations are highly susceptible to local extirpation, which ultimately can lead to worldwide population declines (Budischak et al., 2006).

Herpesvirus Taxonomy

The order Herpesvirales is divided into three families, Herpesviridae, which incorporates mammal, bird, and reptile viruses, Alloherpesviridae, which contains bony fish and frog viruses, and a third family, Malacoherpesviridae, which contains oyster herpesvirus (Davison, 2010).

They are large, icosahedral enveloped double stranded DNA viruses (Essbauer and Ahne, 2001;

Marschang, 2011; McGeoch and Gatherer, 2005; Sim, et al., 2015; Wellehan, 2004). The family

Herpesviridae is divided further into three subfamilies, Alphaherpesvirinae, Betaherpesvirinae, and Gammaherpesvirinae categorizing approximately 90 different species (Davison, 2010;

ICTV, 2014). Currently, chelonian herpesviruses have DNA sequences most homologous to viruses in the alpha herpesvirus family (Bicknese et al., 2010; Lu et al., 2003; Marschang et al.,

2006; Origgi et al., 2001). There have been several reports of herpesviruses in various chelonians

(Table 1), yet there is a large void of information in box turtle species. However, two molecularly distinct species of herpesviruses have recently been reported in Eastern box turtles:

Terrapene herpesvirus 1 (Sim et al., 2015) and Terrapene herpesvirus 2 (Yonkers et al., 2015).

Terrapene herpesvirus 1 was identified in two separate captive collections of eastern box turtles using histopathology and PCR with DNA sequencing (Sim et al., 2015). A three-month- old box turtle emerged early from brumation with lethargy, dehydration, and dyspnea (Sim et al.,

2015). Therapy was initiated but the animal died a week later with histopathologic lesions of necrotizing and ulcerative rhinitis, stomatitis, and esophagitis (Sim et al., 2015). Eosinophilic intranuclear inclusion bodies were visualized on histopathology and PCR amplified a virus, which was sequenced, revealing a novel herpesvirus in the genus Scutavirus (Sim et al., 2015). A separate captive group of adult eastern box turtles with lethargy, stomatitis, cloacitis, and conjunctivitis due to ranavirus infection were opportunistically tested for herpesvirus and had a total prevalence of 58% by PCR (Sim et al., 2015). Sim et al.’s report presented two manifestations of herpesvirus in box turtles. The hatchling exhibited a primary severe herpesvirus infection as seen in very young animals (Marschang, 2011; Sim et al., 2015), whereas the adults displayed recrudescence of latency attributed to concurrent infection and immunosuppression (Sim et al., 2015).

A second herpesvirus report describes herpesvirus infection in Terrapene sp. was in an eastern box turtle with a papillomatous growth behind both rear legs presented to a wildlife rehabilitation center in Florida (Yonkers et al., 2015). Surgical excision resulted in an eight- month remission, however lesions reoccurred, requiring an additional surgery and acyclovir therapy, and the animal was later euthanized due to poor quality of life (Yonkers et al., 2015).

Histopathology of the mass revealed subacute superficial inflammation and stroma consistent with cutaneous papillomas (Yonkers et al., 2015). Nested PCR amplification of the DNA polymerase gene was performed on the extracted lesion, which identified the second novel alphaherpesvirus in eastern box turtles (Yonkers et al., 2015).

While these investigations were integral to the discovery of these viruses in this species, they failed to determine the impact or extent of this pathogen in free-ranging populations. To better characterize the epidemiology of herpesvirus in Terrapene sp., we must gain knowledge

about the effects on wild chelonian populations from captive chelonian reports and other well studied chelonian species.

Molecular Characteristics

Herpesviruses are enveloped, approximately 150 nm in diameter, and contain an icosahedral nucelocapsid measuring 100 nm in diameter (Essbauer and Ahne, 2001; McGeoch and Gatherer, 2005; Sim et al, 2015). The genome consists of a single linear molecule of double stranded DNA, 125-235 kbp in size (Tidona and Darai, 2002). There is a remarkable degree of variation in the composition, size, and organization of the genomes of the herpesviruses (Murphy et al., 1999b). The virion is comprised of core, capsid, tegument, and an envelope (van

Regenmortel et al., 2000). Herpesvirus virions contain over thirty structural proteins and six are present in the nucleocapsid. (Murphy et al.,1999b). These virus-coded proteins play a significant role in the pathogenesis of herpesvirus infections by membrane fusion (Glauser et al., 2012). The viral DNA is transcribed and replicated in the host cell nucleus and the viral mRNA is translated in the cytoplasm (Ahne, 1993).

All known DNA viruses encode their own DNA polymerases, which is essential for viral replication (Coen, 1996). All herpesviruses encode a DNA polymerase in which the catalytic activity remains in the large subunit and the smaller subunit binds DNA and stimulates polymerase activity (Coen, 1996). The last step of herpesvirus replication requires the recruitment of another subunit polymerase, UL30 (Ward and Weller, 2011).

Clinical Signs and Pathology

Clinical and histopathological descriptions of herpesviruses have been best characterized

and described in chelonians (Table 2) as compared to other reptiles and all tortoises share several common clinical and pathologic characteristics (Marschang, 2011). This subfamily of viruses has a predilection for epithelial cells at mucosal surfaces and leads to lesions of stomatitis, rhinitis, glossitis, and conjunctivitis (Drury et al., 1999; Essbauer and Ahne, 2001; Johnson et al., 2005;

Marschang et al., 2001; Marschang 2011; Origgi et al., 2001; Soares et al., 2004; Teifke 2000;).

While some reported infections are fatal before any premonitory signs (Harper et al., 1982;

Jungwirth et al., 2014), there are examples of antemortem clinical signs described in chelonian species.

In captive Mediterranean tortoises, there is an association between herpesvirus presence and upper respiratory tract disease (Origgi et al., 2001). In captive Greek tortoises inoculated with a tortoise herpesvirus, necrotizing stomatitis and rhinitis developed (Origgi et al., 2004).

There are several herpesvirus reports in captive desert tortoises (Gopherus agassizii) (Harper et al., 1982; Jacobson et al., 2012; Johnson et al., 2005; Pettan-Brewer et al., 1996). Necropsy of a captive six-year-old male tortoise revealed caseonecrotic debris over the pharyngeal mucosa, which was attributed to a herpesvirus as seen by enveloped icosahedral viral particles measuring

143 nm in size (Harper et al., 1982). Another report observed herpesvirus particles via electron microscopy in a spur-thighed tortoise (Testudo graeca) displaying necrotic stomatitis (Drury et al., 1999). In an epidemic of chronic rhinitis in fifty captive spur-thighed tortoises, electron microscopy revealed the inciting agent to be a herpesvirus infection (Muro et al., 1998).

A subsequent report described herpesvirus in a captive desert tortoise with caseous necrosis of the trachea, lungs, and oral cavity (Pettan-Brewer et al., 1996). Diagnosis was via visualization of intranuclear inclusion bodies on electron microscopy (Pettan-Brewer et al.,

1996). In a more recent report, a captive California desert tortoise (Gopherus agassizii)

exhibiting clinical signs of anorexia, lethargy, and caseous glossal plaques was diagnosed with a herpesvirus infection (Johnson et al., 2005). Intranuclear inclusion bodies were detected in mucosal epithelial cells by light microscopy followed by observation of herpeslike-particles via electron microscopy (Johnson et al., 2005). Additionally, nested PCR was utilized to amplify the

DNA-dependent DNA polymerase gene of herpesvirus and nucleic acid (Johnson et al., 2005).

Field observation of wild desert tortoises with oral lesions revealed antibody prevalence to

Testudinid herpesvirus of 30.9% (Jacobson et al., 2012). A novel tortoise herpesvirus was identified via PCR in a clinically healthy wild Bowspirit tortoise (Chersina angulata) (Bicknese et al., 2010). In a Great Britain breeding colony, herpesvirus-like particles identified by electron microscopy were described in Hermann’s (Testudo hermanni) and leopard tortoises (Geochelone pardalis) exhibiting ocular and nasal discharge and intermittent paralysis (Drury et al., 1998). An adult Russian tortoise (Testudo horsfeldii) was diagnosed with a herpesvirus infection on necropsy with multifocal necrotic hepatitis and displaying no clinical signs or lesion in the respiratory system or oral cavity (Hervas et al., 2002).

In marine turtles, herpesvirus infections display different clinical signs compared to terrestrial chelonians and has been linked to neoplastic disease, respiratory disease, and skin disease (Alfaro-Nunez and Gilbert, 2014; Ariel, 2011; Essbauer and Ahne, 2001; Greenblatt et al., 2005, Marschang, 2011; Teifke et al., 2000;). One of the first herpesviruses described in chelonians was gray patch disease, associated with Chelonid herpesvirus 1 (Rebell et al., 1975).

This disease affects green sea turtle (Chelonia mydas) hatchlings with circular papular skin lesions, which coalesce into diffuse gray skin lesions (Rebell et al., 1975). Five cases of herpesvirus infections have been described in water turtles; two Pacific pond turtles (Clemmys marmorata) (Frye et al., 1977), a captive painted turtle (Chrysemys picta) (Cox et al., 1980), map

turtles (Jacobson et al., 1982), West African mud turtles (Marschang et al.,2015), and freshwater turtles (Pseudemys concinna concinna) (Jungwirth et al., 2014). Reported necropsy findings include pulmonary edema, hepatomegaly, and hepatic lipidosis with intranuclear inclusion bodies seen in hepatocytic nuclei. Chelonid herpesvirus 6 has been associated with lung-eye- trachea disease in marine turtles, and the eyes and respiratory tract are affected in ill turtles with signs lasting 2-3 weeks (Ariel, 2011; Coberely et al., 2002; Curry et al., 2000; Jacobson et al.,

1986; Marschang, 2011). Chelonid herpesvirus 5 is associated with a neoplastic disease seen in many different species of marine turtles; including green, loggerhead, Hawksbill, and Olive

Ridley sea turtles (Jacobson et al., 1991; Quackenbush et al., 2001). Infected turtles develop fibropapillomas externally all over the body. Severe infections may lead to tumors of internal organs (Alfaro-Nunez and Gilbert, 2014; Ariel, 2011; Chaloupka et al., 2009; Stacy et al., 2008).

Two herpesvirus-associated diseases from wild-caught loggerhead sea turtles (Caretta caretta) have been described: loggerhead genital-respiratory herpesvirus and loggerhead orocuteanous herpesvirus (Ariel, 2011; Stacy et al., 2008). Loggerhead genital-respiratory herpesvirus was associated with tracheal and cloacal ulcers and loggerhead orocuteanous herpesvirus was associated with oral ulcers and exudative plaques (Marschang, 2011; Stacy et al., 2008).

Few herpesviruses have been documented in snakes while several have been discussed in lizards (Hauser et al., 1983; Literak et al., 2010; Simpson et al., 1979; Watson, 1993; Wellehan et al., 2004; Wellehan et al., 2005) A decrease in venom production in cobras (Simpson et al.,

1979) as well as anemia and acute hepatitis in boas (Hauser et al., 1983) has been noted in snakes infected with herpesvirus. Oral lesions have been associated with herpesvirus in lizards

(Marschang, 2011). In green tree monitor lizards (Varanus prasinus), herpesvirus, in the subfamily alphaherpesvirinae, was characterized as proliferative stomatitis (Wellehan et al.,

2005). Another alphaherpesvirinae stomatitis infection was identified in Sudan plated lizards

(Gerrhosaurus major) and a black-lined plated lizard (Gerrhosaurus nigrolineatus) (Wellehan et al., 2004). Herpesvirus associated liver lesions were detected by electron microscopy in red- headed agamas (Agama agama) (Watson, 1993). A San Esteban chuckwalla (Sauromalus varius) was also identified to have diffuse hepatic necrosis attributed to an alphaherpesvirinae infection.

(Wellehan et al., 2003) Hepatocellular necrosis due to a herpesvirus infection, related to Iguanid herpesvirus 1, was identified in a green iguana (Iguana iguana) (Wilkinson et al., 2005). More recently, herpesvirus DNA was detected by PCR from papillomas in green lizards (Literak et al.,

2010) and was found to have sequence similarity to fibropapilloma-associated turtle herpesvirus from sea turtles. Hepatitis and enteritis caused by a novel herpesvirus in the family Varanidae was identified in two monitor lizards (Varanus sp.) (Hughes-Hanks et al., 2010).

Herpesvirus infections have been documented in two crocodilian species. Juvenile alligators (Alligator mississippiensis) exhibiting multifocal hyperemic cloacal nodules were diagnosed with a herpesvirus infection via PCR (Govett et al., 2005) and shared 85% of sequence identify to tortoise herpesvirus 1. A herpesvirus infection was documented in an

Australian saltwater crocodile (Crocodylus porosus) that presented with grey-colored abdominal skin lesions (McCowen et al., 2005).

Pathogenesis

Transmission of mammalian herpesviruses is by contact of infected cells in saliva, urogenital excretions, or aerosols (Fields et al., 1996). In mammalian species, herpesviruses can survive via persistent, often latent, infections, from which virus is periodically reactivated and shed (Alfaro-Nunez and Gilbert, 2014; Ariel, 2011; Hardman et al., 2012; Marschang et al,

2011). These shedding events may reactive the virus and are associated with stress caused by other infections, shipping, cold, or crowding (Govett et al., 2005; Hoff et al., 1984).

The mechanism of transmission of chelonian herpesvirus is largely unknown and differs species to species (Hoff et al., 1984). A transmission study with Greek tortoises (Testudo graeca) demonstrated that ELISA detected seroconversion of herpesvirus within 4-7 weeks post experimental inoculation which is earlier than serum neutralization (Origgi et al., 2001). Another herpesvirus transmission study in Greek tortoises (Testudo graeca) was successful and all infected tortoises, either intranasally or intramuscularly, exhibited conjunctivitis and diphtheritic oral plaques, consistent with a herpesvirus infection after a secondary challenge (Origgi, et al.,

2004). Chelonian herpesvirus is frequently detected in pharyngeal swabs, suggestive of a route of natural transmission through nasal and oral discharge (Soares et al., 2004).

In marine turtles, herpesviruses from infected turtles may be transmitted to uninfected turtles via vectors, direct contact with the virus, or in seawater (Curry et al., 2000). In aquaculture reared C. mydas, gray-patch disease sheds from skin lesions into the water, allowing subsequent transmission to susceptible turtles (Rebell et al., 1975). In green sea turtles, shedding of gray patch disease and severity of disease is affected by water temperature and other stress factors (Haines and Kleese, 1977). Marine leeches (Ozobranchus spp.) are a discussed mode of transmission of herpesvirus for green turtles, in which they feed in the cloacal region as well as mouth, eyelids, and integument (Greenblatt et al., 2004; Stacy et al., 2008). Experimental transmission of fibropapilloma to green turtles has been successful using filtered, cell-free tumor extracts (Herbst et al., 1995). A transmission study of lung-eye-trachea-disease in marine turtles remained persistent, as measured by PCR, in seawater for as long as 120 hours (Curry et al.,

2000). Caution should be taken with handling and proper sanitation of equipment in contact with

marine turtles should be utilized to minimize the risk of transmission (Curry et al., 2000). A more recent paper discusses viral presence of herpesvirus in genital lesions of loggerhead sea turtles (Caretta caretta); one presented with several linear ulcers on the phallus and a second presented with a distinct circumferential ulcer around the cloacal mucocutaneous junction (Stacy et al., 2008). This is the first paper eluding to sexual transmission of herpesvirus (Stacy et al.,

2008), which can play a significant role in conservation efforts, especially if this mode of transmission occurs in other chelonid species.

In crocodilian species, transmission is less commonly believed to occur through aerosolization of infectious droplets and more through mucosal contact (Govett et al., 2005).

Once infected, herpesvirus persists in lymph tissue, epithelial tissue, or nervous tissue and is spread through continuous or intermittent shedding (Govett et al., 2005; Hunt, 2006; Ritchie,

2006; Thiry et al., 1986)

Diagnosis

There are several methods that have been used to diagnose herpesvirus infection in chelonians (Table 2), including, polymerase chain reaction (PCR), in situ hybridization, histopathology, electron microscopy, serum neutralization, and serology such as enzyme-linked immunosorbent assay (ELISA) and immunohistochemistry. The most documented method for diagnosis of herpesvirus in many species is to utilize a consensus PCR that amplifies a region of the herpesviral DNA polymerase gene. This method can detect herpesvirus presence even when no prior DNA sequence information is available (VanDevanter et al., 1996). The assay successfully amplifies a 215- to 315-bp of a moderately variable region (VanDevanter et al.,

1996). However, it is labor intensive and time consuming and requires sequencing of each

positive sample, making it less practical for large-scale epidemiologic studies. Other conventional PCR primers were developed targeting two specific primers of the DNA polymerase gene, U-73 and L-588, for a 535 bp sequence specific for tortoise herpesvirus

(Murakami et al., 2001). This technique allowed for the diagnosis of tortoise herpesvirus infection as well as the detection of possible latent infections and carriers (Murakami et al,

2001). A 307 bp DNA probe was developed targeting the UL5 homologue from tortoises using in situ hybridization and PCR (Teifke et al., 2000). This technique utilized formalin-fixed, paraffin embedded tissues and concluded to be highly specific for tortoise herpesvirus infections.

Greek tortoises were experimentally inoculated with tortoise herpesvirus isolates and PCR targeting the UL39 homologue was utilized to fulfill Koch’s postulates for a herpesvirus associated stomatitis-rhinitis in Mediterranean tortoises (Origgi et al., 2004). PCRs targeting the

DNA polymerase gene, the UL5 homologue and the UL39 homologue were evaluated in 11 different known-positive tortoise isolates (Marschang et al., 2006). The nested PCR targeting the

DNA polymerase gene, by VanDevanter, was the most sensitive and the only PCR capable of detecting all of the 11 tortoise herpesvirus isolates used in that study (Marschang et al., 2006;

VanDevanter et al., 1996). PCR has also been utilized in marine turtles to amplify a herpesvirus

DNA polymerase gene segment from fibropapillomas (Quackenbush et al., 1998). While extremely useful and widespread, PCR only indicates viral DNA presence and does not indicate active disease. Furthermore, another limitation of this method is that it does not allow quantification.

An inverse polymerase chain reaction based genomic walking technique was utilized for the amplification of a 483 base pair segment of a green turtle herpesvirus fibropapilloma and reduction of non-specific herpesvirus amplification (Yu et al., 2001). Reverse transcription

polymerase chain reaction (RT-PCR) and quantitative PCR have been used to support that herpesvirus in green turtles cause fibropapillomas for green, olive ridley, and loggerhead turtles from a variety of locations (Lu et al., 2003 and Quackenbush et al., 2001).

The diagnosis of herpesvirus has also been made by the presence of eosinophilic or amphophilic intranuclear inclusion bodies by light microscopy followed by visualization of virus particles by electron microscopy (Drury et al., 1998; Muro et al., 1998; Harper et al., 1982;

Heldstab and Bestetti, 1989; Johnson et al., 2005; Murakami et al., 2001; Sim et al., 2015; Une et al., 1999). Virus particles are most frequently identified in epithelial cells of the tongue, oral mucosa, and upper respiratory tract (Marschang et al., 2011). This technique requires biopsy of infected tissue and this method may not detect subclinical carriers and will not detect latent infections (Ariel 2011; Essbauer and Ahne 2001; Farkas and Gal, 2009; Jacobson et al., 1982;

Johnson et al., 2005; Origgi et al., 2004).

A serum neutralization technique, which detects and quantifies antiviral antibodies, has been developed and is considered the gold standard for determining exposure, but has limited application due to geographic restriction and availability in a few research laboratories in

Europe, as well as being a laborious technique, and containing a long result retrieval time of 10-

14 days (Frost and Schmidt, 1997; Murphy et al., 1999a; Origgi et al., 2001).

In tortoises, plasma samples have been evaluated for the presence of anti-herpesvirus antibodies by ELISA, which is a more efficient and quicker, yet equally reliable method to serum neutralization (Johnson et al., 2010; Origgi et al., 2001). This technique is simpler than serum neutralization and has a shorter procedure time, with results available in one day. ELISA was able to detect the presence of antiherpesvirus antibodies 2 to 5 weeks earlier than serum neutralization when performed in Mediterranean tortoises (Origgi et al., 2001). ELISA can detect

antibodies that bind to multiple viral antigenic determinants, where as serum neutralization detects antibodies against surface glycoproteins (Origgi et al., 2001). ELISA can detect recent infections as well as infections that occurred before plasma was collected (Johnson et al., 2010).

One of the main disadvantages of an ELISA test is that the antigen immobilization method utilized is not specific (Gan and Patel, 2013). An ELISA was developed for the detection of anti- herpesvirus antibodies associated with lung-eye-trachea disease seen in culture-reared green turtles (Coberley et al., 2001). This assay distinguished herpesvirus infections due to lung-eye- trachea disease from antibodies due to fibropapillomatosis with confirmation from immunohistochemistry (Coberley et al., 2001).

Immunohistochemistry is useful in quickly confirming the presence of a pathogen in formalin fixed tissue generally through biopsy or necropsy (Gupta et al., 2012). Two sensitive immunohistochemical investigative tools, direct and indirect immunoperoxidase assays, were developed as a more rapid technique for detecting the presence of herpesvirus antigen in infected tissue and anti-herpesvirus antibodies in exposed tortoises, respectively (Mesa-Tejada et al.,

1977; Origgi et al., 2003). The direct assay detects the presence of herpesvirus antigen in cells being used to screen for the presence of herpesvirus, whereas the indirect detects the presence of specific antibody by direct observation of positive staining (Origgi et al., 2003). This technique may be utilized independently or complementary to the existing serum neutralization or ELISA techniques.

Treatment

Herpesvirus treatment is primarily supportive therapy with parenteral nutrition, maintenance fluids, minimizing stress, proper environmental hygiene, and antibiotics to prevent

secondary bacterial infections (Jacobson et al., 1986; Jacobson et al., 1991; Kirchgessner and

Mitchell, 2008; Ritchie 2006). Infected animals with upper respiratory tract disease may benefit therapeutically with nebulization of responsive antifungals and antimicrobials as well as an increase in ambient temperature and humidity (Schumacher, 1996; Schumacher, 2003). It is also recommended to treat herpesvirus associated oral lesions, topically, with dilute chlorhexidine every twelve to twenty-four hours (Hunt, 2006; Kirchgessner and Mitchell, 2008). If a reptile survives a herpesvirus infection, they should be viewed as latently infected or a carrier, and should be separated from susceptible individuals (Ritchie, 2006).

Herpesvirus infections in humans treated with acyclovir has been shown to have shorter viral shedding time and reduced re-infection (Ritchie, 2006). There are few reports of acyclovir use in reptiles and the role acyclovir plays in minimizing recrudescence of disease has not yet been reported (Frye et al., 1977; Jacobson et al., 1986; Ritchie, 2006; Schumacher, 1996).

Therapy is most effective before reptiles are symptomatic (Ritchie, 2006). The only anti- herpesviral drug study in tortoises showed gancyclovir and acyclovir to inhibit herpesvirus replication in vitro (Marschang et al., 1997; Origgi, 2006). There is a necessity for anti- herpesvirus drug studies in vivo to formulate the most effective drug treatment and route of administration (Origgi, 2006). Acyclovir at 80 mg/kg every three days has been used in

California desert tortoises to treat herpesvirus-associated ulcerative stomatitis (Schumacher,

1996). The same dose was successfully used in Mediterranean tortoises once daily, for three months, with nasal discharge attributed to a herpesvirus infection (Hunt, 2006). The application of 5% acyclovir ointment in spur-thighed tortoises improved oral lesions (Schumacher,1996). A pharmacokinetics study of acyclovir and valacyclovir in twelve North American box turtles

concluded that 40 mg/kg of oral valacyclovir every 24 hours can be utilized as proposed treatment for herpesvirus infections (Allender et al., 2013).

If herpesvirus infection is uncertain, quarantine should be instilled while specific diagnostic tests are performed (Ritchie 2006; Schumacher, 2003).

Epidemiology

Chelonid herpesvirus was first isolated in green sea turtles in the 1970’s (Rebell et al.,

1975). Since that time, reports of many reptilian species affected by herpesvirus have been described including other species of chelonians (Table 1), snakes (Hauser et al., 1983; Lunger and Clark, 1978; Monroe et al., 1968), and lizards (Clark et al., 1972; Watson, 1993; Literak et al., 2010; McCowan et al., 2004; Wellehan et al., 2004). More recently, herpesvirus has been reported in the family Emydidae, as demonstrated in eastern box turtles (Terrapene carolina carolina ) (Sim et al., 2015; Yonkers et al., 2015). Other species in the family Emydidae, comprising of marsh turtles found in the western hemisphere, have recently been reported as well.

A herpesvirus infection was diagnosed in a captive, juvenile, female northern map turtle

(Graptemys geographica) displaying weakness and frothy nasal discharge (Ossiboff et al.,

2015a). Despite supportive care with a broad-spectrum antibiotic for suspected bacterial pneumonia, the animal was found shortly after (Ossiboff et al., 2015a). Histologic examination revealed significant lesions in the lung, liver, and spleen, consistent with pulmonary epithelial necrosis with intranuclear viral inclusions present in all three organs (Ossiboff et al., 2015a).

PCR sequence analysis of the liver revealed resemblance to Emydid herpesvirus 1, Terrapene herpesvirus 1, and tortoise herpesvirus (Ossiboff et al., 2015a). Twenty-seven additional turtles

housed in an enclosure with shared circulation were tested by PCR using a combined choanal and cloacal swab (Ossiboff et al., 2015a). Four asymptomatic painted turtles (Chrysemys picta) and one asymptomatic northern map turtle were PCR positive (Ossiboff et al., 2015a).

An epidemiological study was performed in northeastern United States investigating herpesvirus infection by PCR in endangered or critically endangered Emydids (Ossiboff et al.,

2015b). Prevalence in bog turtles (Glyptemys muhlenbergii) was 51.5%, wood turtles had a prevalence of 55.6% (Glyptemys insculpta), and 5.8% of spotted turtles (Clemmys guttata) were positive for herpesvirus (Ossiboff et al., 2015b).

Herpesvirus infections have been more commonly reported in adult chelonians compared to juveniles (Cowan et al., 2015; Cox et al., 1980; Jacobson et al., 2012; Johnson et al., 2005;

Pettan-Brewer et al., 1996). This could be due to adult chelonians being more commonly sampled than juveniles or due to adult chelonians having recrudescence of latency. There has been no reported increase in herpesvirus detection attributed to sex predilection or geographical location; however, there has been an increase in association with underlying illness, which may cause reactivation of latent infection (Stacy et al., 2008). There is also a suspected association between seasonality and occurrence of herpesvirus infections in tortoises (Origgi, 2006).

Herpesvirus infections have been more frequently observed in the beginning of the spring and fall seasons, which correlates to the end and beginning of brumation, which may be attributed to decreased immunocompetence during periods when animals are outside of their preferred optimal temperature zone (Campbell et al., 2014; Origgi, 2006; Zapata et al., 1992).

The global distribution of herpesviruses in reptilian populations has been previously documented in the United States (Jacobson et al., 1985; Harper et al., 1982; Pettan-Brewar et al.,

1996; Bickenese et al., 2010; Sim et al., 2015, Watson, 1993), Africa (Oettle et al., 1990; Duarte

et al., 2012; Marschang et al., 2009), and Europe (Cooper et al., 1988; Drury et al., 1998;

Heldstab and Bestetti, 1989; Salinas et al., 2011; Literak et al., 2010; Drury et al., 1999;

Jungwirth et al., 2014; Hauser et al., 1983; Soares et al., 2004; Hunt, 2006).

Summary

In summary, herpesvirus is a well-studied DNA virus in humans, tortoises, and marine turtles; however, little is known of herpesvirus epidemiology on the genus Terrapene, but it is increasingly being reported. The remainder of this thesis will provide insight in characterizing the epidemiology of herpesvirus in eastern box turtles.

Table 1. Chelonian species herpesviruses have been isolated from.

Family Species Reference Testudinidae California desert tortoises (Gopherus agassizii) Harper et al., 1982 Pettan-Brewer et al., 1996 Soares et al., 2004 Johnson et al., 2005 Salinas et al., 2011 Jacobson et al., 2012 Bowspirit tortoises (Chersina angulata) Bicknese et al., 2010 Hermann's tortoises (Testudo hermanni) Heldstab and Bestetti, 1989 Lange et al., 1989 Drury et al., 1998 Teifke et al., 2000 Herbst et al., 2004 Soares et al., 2004 Hunt 2006 Leopard tortoises (Geochelone pardalis) Drury et al., 1998 Soares et al., 2004 Marginated tortoise (Testudo marginata) Soares et al., 2004 Hunt 2006 Russian tortoises (Testudo horsfeldii) Lange et al., 1989 Une et al., 1999 Teifke et al., 2000 Hervas et al., 2002 Soares et al., 2004 Hunt 2006 Argentine tortoises (Geochelone chilensis) Jacobson et al., 1985 Soares et al., 2004 Egyptian tortoises (Testudo kleinmanni) Hunt 2006 Marschang et al., 2009 Pancake tortoises (Malacochersus tornieri) Une et al., 2000 Une et al., 1999 Greek tortoises (Testudo graeca) Muro et al., 1998 Drury et al., 1999 Une et al., 2000 Soares et al., 2004 Hunt 2006 Fresh water turtles (Pseudemys concinna Emydidae concinna) Jungwirth et al., 2014 Northern map turtles (Graptemys geographica) Jacobson et al., 1982 Ossiboff et al., 2015a Eastern box turtles (Terrapene carolina carolina) Sim et al., 2015 Yonkers et al., 2015 Bog turtles (Glyptemys muhlenbergii) Ossiboff et al., 2015b Wood turtles (Glyptemys insculpta) Ossiboff et al., 2015b Spotted turtles (Clemmys guttata) Ossiboff et al., 2015b Painted turtles (Chrysemys picta) Cox 1980 Ossiboff et al., 2015a Pacific pond turtles (Actinemys marmorata) Frye et al., 1977 Pelomedusidae West African mud turtles (Pelusios castaneuis) Marschang et al., 2015 Chelidae Krefft's river turtle (Emydura macquarii krefftii) Cowan et al., 2015 Cheloniidae Green turtles (Chelonia mydas) Jacobson et al., 1986 Rebell et al., 1975 Jacobson et al., 1991 Cobereley et al., 2001 Quakenbush et al., 2001 Duarte et al., 2012 Loggerhead turtles (Caretta caretta) Herbst et al., 1994 Olive ridley turtles (Lepidochely olivacea) Quackenbush et al., 2001 Stacy et al., 2008

Table 2. Herpesvirus clinical signs and diagnostic methods reported in chelonians.

Species Clinical Signs Diagnosis Reference Bowspirit tortoises (Chersina angulata) None PCR Bicknese et al., 2010 Green turtles (Chelonia mydas) Fibropapillomatosis ELISA Cobereley et al., 2001 Skin lesions, lethargy, chronic decreased Krefft's river turtle (Emydura macquarii krefftii) appetite H, PCR Cowan et al., 2015 Painted turtles (Chrysemys picta) Cranial abscess H, EM Cox 1980 Hermann's tortoises (Testudo hermanni) Ocular/nasal discharge, paralysis EM Drury et al., 1998 Leopard tortoises (Geochelone pardalis) Greek tortoises (Testudo graeca) Stomatitis, nasal discharge, facial edema EM Drury et al., 1999 Green turtles (Chelonia mydas) Fibropapillomatosis H, PCR Duarte et al., 2012 California desert tortoises (Gopherus agassizii) None H, EM Harper et al., 1982 Russian tortoises (Testudo horsfeldii) Multifocal necrotic hepatitis H, EM Hervas et al., 2002 Hermann's tortoises (Testudo hermanni) Russian tortoises (Testudo horsfeldii) Egyptian tortoises (Testudo kleinmanni) None, anorexia, stomatitis, mucopurulent PCR, H Hunt 2006 Greek tortoises (Testudo graeca) nasal/oral discharge, neurologic Marginated tortoise (Testudo marginata) Northern map turtles (Graptemys geographica) Lethargy, anorexia, subcutaneous edema H, EM Jacobson et al., 1982 Green turtles (Chelonia mydas) Conjunctivitis, tracheitis, pneumonia H, EM, VI Jacobson et al., 1986 Green turtles (Chelonia mydas) Fibropapillomatosis H, EM Jacobson et al., 1991 California desert tortoises (Gopherus agassizii) None PCR, ELISA Jacobson et al., 2012 H, EM, PCR, VI, California desert tortoises (Gopherus agassizii) Early brumation exit, anorexia, lethargy ELISA Johnson et al., 2005 Fresh water turtles (Pseudemys concinna concinna) None H, EM, PCR Jungwirth et al., 2014 Egyptian tortoises (Testudo kleinmanni) Stomatitis PCR, VI Marschang et al., 2009 Greek tortoises (Testudo graeca) West African mud turtles (Pelusios castaneuis) None PCR Marschang et al., 2015 Greek tortoises (Testudo graeca) Rhinitis, stomatitis, glossitis EM Muro et al., 1998 Northern map turtles (Graptemys geographica) None, weakness, nasal discharge H, PCR Ossiboff et al., 2015a Painted turtles (Chrysemys picta) Wood turtles (Glyptemys insculpta) None Ossiboff et al., 2015b Spotted turtles (Clemmys guttata) PCR Bog turtles (Glyptemys muhlenbergii) Pettan-Brewer et al., California desert tortoises (Gopherus agassizii) Weight loss, anorexia H, EM 1996 Green turtles (Chelonia mydas) Fibropapillomatosis PCR Quakenbush et al., 2001 Olive ridley turtles (Lepidochely olivacea) Green turtles (Chelonia mydas) Skin lesions H, EM Rebell et al., 1975 Nasal discharge, oral/respiratory lesions, California desert tortoises (Gopherus agassizii) conjunctivitis, esophagitis, glossitis PCR Salinas et al., 2011 Eastern box turtles (Terrapene carolina Lethargy, dehydration, dyspnea, stomatitis, carolina) cloacitis, conjunctivitis H, PCR Sim et al., 2015 California desert tortoises (Gopherus agassizii) Hermann's tortoises (Testudo hermanni) Leopard tortoises (Geochelone pardalis) Nasal discharge, oral lesions, PCR Soares et al., 2004 Greek tortoises (Testudo graeca) rhinitis, conjunctivitis, stomatitis, glossitis Marginated tortoise (Testudo marginata) Argentine tortoises (Geochelone chilensis) Ulcerative/necrotic oral, respiratory, Olive ridley turtles (Lepidochely olivacea) cutaneous, & genital lesions PCR Stacy et al., 2008 Hermann's tortoises (Testudo hermanni) Stomatitis–rhinitis complex EM, VI, PCR, ISH Teifke et al., 2000 Russian tortoises (Testudo horsfeldii) Russian tortoises (Testudo horsfeldii) N/A PCR Une et al., 2000 Pancake tortoises (Malacochersus tornieri) Greek tortoises (Testudo graeca) Pancake tortoises (Malacochersus tornieri) None H, EM, PCR Une et al., 1999 Russian tortoises (Testudo horsfeldii) Eastern box turtles (Terrapene carolina carolina) Papilloma H, PCR Yonkers et al., 2015

Key: EM - Electron microscopy, H - Histology, PCR - polymerase chain reaction, ISH - In situ hybridization, ELISA - enzyme linked immunosorbent assay, VI - virus isolation

CHAPTER 3

DEVELOPMENT AND VALIDATION OF QUANTITATIVE PCR FOR DETECTION OF TERRAPENE HERPESVIRUS 1 UTILIZING FREE-RANGING EASTERN BOX TURTLES (TERRAPENE CAROLINA CAROLINA)1

Abstract

Diseases that affect the upper respiratory tract (URT) in chelonians have been well described as a significant contributor of morbidity and mortality. Specifically, herpesviruses are common pathogens in captive chelonians worldwide, but their importance on free-ranging populations is less well known. Historical methods for the diagnosis of herpesvirus infections include virus isolation and conventional PCR. Real-time PCR has become an essential tool for detection and quantitation of many pathogens, but has not yet been developed for herpesviruses in box turtles. Two quantitative real-time TaqMan PCR assays, TerHV58 and TerHV64, were developed targeting the DNA polymerase gene of Terrapene herpesvirus 1 (TerHV1). The assay detected a viral DNA segment cloned within a plasmid with 10-fold serial dilutions from

1.04×107 to 1.04×101 viral copies per reaction. Even though both primers had acceptable levels of efficiency and variation, TerHV58 was utilized to test clinical samples based on less variation and increased efficiency. This assay detected as few as 10 viral copies per reaction and should be utilized in free-ranging and captive box turtles to aid in the characterization of the epidemiology of this disease.

Keywords

Herpesvirus, Eastern box turtle, Epidemiology, qPCR, Terrapene carolina

1 Content of this chapter has been partially or fully submitted for publication as cited in bibliography as Kane et al., 2016.

1. Introduction

Viruses from the order Herpesvirales are large, icosahedral, enveloped, double stranded,

DNA viruses (McGeoch and Gatherer, 2005; Wellehan et al., 2004; Sim et al., 2015). This order is divided into three families, Herpesviridae, Alloherpesviridae, and Malacoherpesviridae

(Davison, 2010). The family Herpesviridae is further divided into three subfamilies,

Alphaherpesvirinae, Betaherpesvirinae, and Gammaherpesvirinae encompassing approximately

90 different species (Davison, 2010; ICTV, 2014). Currently, chelonian herpesviruses, in the genus Scutavirus, have DNA sequences most homologous to viruses in the alphaherpesvirus family (Bicknese et al., 2010; Greenblatt et al., 2005, Ossiboff et al., 2015a; Ossiboff et al.,

2015b; Herbst et al., 2004; Origgi et al., 2015).

As has been seen in mammals and birds, several chelonian herpesviruses have been associated with clinical signs (Salinas et al., 2011; Stacy et al., 2008; Pettan-Brewer et al., 1996;

Johnson et al., 2005). In tortoises, this virus has a predilection for mucoepithelial cells, commonly associated with rhinitis, conjunctivitis, stomatitis and glossitis (Salina et al., 2011;

Pettan-Brewer et al., 1996; Johnson et al., 2005; Ossiboff et al., 2015a). Captive Greek tortoises inoculated with herpesvirus developed necrotizing stomatitis and rhinitis (Origgi et al., 2004). A captive desert tortoise diagnosed with a herpesvirus infection exhibited anorexia, lethargy, and caseous glossal plaques (Johnson et al., 2005). In aquatic turtles, herpesvirus infections have been linked to skin disease (Rebell et al., 1975; Cowan et al., 2015; Yonkers et al., 2015). Marine turtles affected with gray patch disease, associated with Chelonid herpesvirus 1, develop circular gray skin lesions (Rebell et al., 1975). The most common clinical sign seen in marine turtles, with a herpesvirus infection due to Chelonid herpesvirus 5, is development of neoplastic fibropapillomas (Jacobson et al., 1991; Quackenbush et al., 2001; Chaloupka et al., 2009).

Chelonid herpesvirus 6 has been associated with lung-eye-trachea disease in marine turtles

(Curry et al., 2000; Coberely et al., 2002; Jacobson et al., 1986). However, some reported herpesvirus infections are fatal before any premonitory signs are noted (Harper et al., 1982;

Jungwirth et al., 2014).

Reports of box turtle herpesvirus are scarce. The presence of a unique virus has been demonstrated to occur in three different collections (Sim et al., 2015; Yonkers et al., 2015). An eastern box turtle (Terrapene carolina carolina) which emerged early from brumation with lethargy, dehydration, and dyspnea died a week later and was diagnosed with herpesvirus on histopathology and PCR (Sim et al., 2015). A separate collection of turtles infected with ranavirus was opportunistically tested for herpesvirus via PCR and was found to have a 58% prevalence (Sim et al., 2015). A free ranging eastern box turtle with a papillomatous growth was diagnosed with a novel herpesvirus by nested PCR (Yonkers et al., 2015). Herpesvirus was diagnosed by PCR in a captive Australian Krefft’s river turtle (Emydura macquarii kreffti), which presented with severe proliferative and ulcerative skin and shell lesions (Cowan et al.,

2015). A captive, juvenile, female northern map turtle (Graptemys geographica) was diagnosed with herpesviral pneumonia, which was confirmed by PCR, after exhibiting weakness and nasal discharge (Ossiboff et al., 2015a). Three novel herpesviruses were identified in free-ranging bog turtles (Glyptemys muhlenbergii), wood turtles (Glyptemys muhlenbergii), and spotted turtles

(Clemmys guttata) by PCR (Ossiboff et al., 2015b). Additionally, two clinically healthy West

African mud turtles (Pelusios castaneus) were diagnosed with a novel herpesvirus infection by

PCR (Marschang et al., 2015). Nonetheless, the epidemiology in free-ranging eastern box turtle populations is still unknown.

Several diagnostic methods have been used to diagnose herpesvirus infections in

chelonians, including virus isolation and conventional PCR. Quantitative PCR (qPCR) has not been previously developed for the detection of Terrapene herpesvirus 1 (TerHV1) in chelonians and would provide increased sensitivity as compared to conventional PCR. This method would allow quantitation of virus copy numbers. Furthermore, it may allow detection of carriers with low levels of virus particles. Animals with subclinical infections may serve as important carriers or reservoirs for infectious disease; therefore, it is crucial for assay development to be capable of detecting pathogens in these animals. The quantification of viral copy numbers may also allow for therapeutic monitoring of infections. To date, no such assay has been reported for herpesvirus quantification in free-ranging or captive box turtles.

The purpose of this study was to develop and evaluate diagnostic methods for detecting

TerHV1 in eastern box turtles. The hypothesis tested in this study was that a Taqman qPCR assay would be sensitive for detecting Terrapene herpesvirus 1 (Order Herpesvirales, Family

Herpesviridae, Subfamily Alphaherpesvirinae, Genus Scutavirus) in eastern box turtles. This quantitative method will be useful for early detection and disease monitoring in both captive and free-ranging chelonian populations.

2. Materials and Methods

2.1. Conventional PCR, Sequencing, and Cloning

An oral swab was collected from a suspect positive free-ranging eastern box turtle in

Tennessee determined by associated clinical signs, such as rhinitis, glossitis, or stomatitis. DNA was extracted according to the manufacturer’s instructions (QIAamp DNA blood mini kit,

Qiagen, Valencia, California, USA). Nested PCR was performed as previously described for herpesvirus detection (VanDevanter et al., 1996) and later documented for detection of

herpesviruses in chelonians (Marschang et al., 2006). Primary PCR contained two sense (DFA,

5’-GAYTTYGCNAGYYTNTAYCC-3’; and ILK, 5’-

TCCTGGACAAGCAGCARNYSGCNMTNAA-3’) and one anti-sense (KG1, 5’-

GTCTTGCTCACCAGNTCNACNCCYTT-3’) primers. Secondary PCR assays were performed with sense (TGV, 5’-TGTAACTCGGTGTAYGGNTTYACNGGNGT-3’) and anti-sense (IYG,

5’-CACAGAGTCCGTRTCNCCRTADAT-3’) primers targeting a herpesvirus-consensus region of the DNA polymerase gene. Reactions were performed in 25 µL total volume using Premix taq

(Takara Taq version) containing hot start Taq (TaKaRa Bio Inc.). Five µl of template DNA from the first step was used in the second reaction. Feline herpesvirus 1 and canine herpesvirus 1 were used as positive controls, while DNase/RNase-free water was used as a negative control. PCR products were resolved by electrophoresis in 1.4 % agarose gels. Products were sequenced in both directions using the Molecular Biology Resource Facility at the University of Tennessee and compared to known sequences in GenBank using TBLASTX. Phylogenetic analysis of herpesvirus sequences obtained from clinical samples and the literature (Sim et al., 2015;

KJ004665) were used to design a consensus 480 base pair gene segment (Integrated DNA

Technologies, Coralville, Iowa) that was then cloned in E. coli (TOPO TA Cloning® kit,

Invitrogen, Carlsbad, CA). The cloning products were verified through sequencing in both directions (Keck Biotechnology Center at the University of Illinois). Plasmids were linearized with BamH1 (Clontech Laboratories, Mountain View, CA), purified (QIAfilter plasmid Maxi kit,

Qiagen, Valencia, CA), and quantified using a spectrophotometer (Nanodrop spectrophotometer,

Thermo Scientific, Wilmington, DE). Ten-fold serial dilutions of linearized plasmids were made from 1.04×109 ng/µl to 1.04×100 ng/µl. Viral genome (DNA) copy number was calculated using the following formula:

# copies/µL = (ng DNA of plasmid + clone/ µL) (6.022 ×1023 copies/mol)

(base pair length)(1×109 ng/g) (650 g/mol of base pair)

The final copy number for ten-fold serial dilutions ranged from 1.04×107 to 1.04×101 viral copies per reaction.

2.2. Real Time qPCR Assay

Two TaqMan-MGB (Taqman® primers, FAM dye labeled, Applied Biosystems,

Carslbad, CA) qPCR assays, TerHV58 and TerHV64, for TaqMan-MGB (Taqman® primers,

FAM dye labeled, Applied Biosystems, Carslbad, CA) qPCR assays were designed using a commercial software program (Primer Express®, Applied Biosystems, Carlsbad, CA) based on the sequence of the TerHV1 DNA polymerase gene segment in our clone. The TaqMan with assay TerHV58 was performed using forward (5’-TCTATTGGGCGAGCTGTTGAC-3’), reverse (5’-GAAAAGCCATACGCCTACGC-3’), and probe (5’-ACTGGCTGGCCTTG-3’), targeting a 58 base pair segment of the herpesvirus DNA polymerase gene. The TaqMan with assay TerHV64 was performed using forward (5’-GGCCACCCGAGATTATATCCA-3’) reverse (5’-TTCTGGCCGACTTCCCG-3’), and probe (5’-

CCCATTGGAGCGAACGGGAAAAG-3’), targeting a 64 base pair segment of the herpesvirus

DNA polymerase gene. Real-time qPCR assays were performed using a real-time PCR thermocycler (7500 ABI real-time PCR System, Applied Biosystems, Carlsbad, CA) and data was analyzed using associated software (Sequence Detection Software v2.05, Applied

Biosystems, Carlsbad, CA). Each reaction contained 12.5 µl of 20x TaqMan Platinum PCR

Supermix-UDG with ROX (TaqMan Platinum PCR Supermix-UDG with ROX, Invitrogen,

Carlsbad, CA), 1.25 µl TaqMan primer-probe, 2.5 µl plasmid dilution, and water to a final

volume of 25 µl. Cycling parameters were as follows: 1 cycle at 50°C for 2 minutes followed by

95°C for 10 minutes, then 40 cycles at 95°C for 15 seconds and 60°C for 60 seconds, and a final cycle of 72°C for 10 minutes.

2.3. Standard Curve and Sensitivity

To determine the sensitivity, assays were performed in four technical repeats on dilutions of turtle-derived positive control plasmid of TerHV1 DNA (1.04×107 – 1.04×101 copies/rxn) within a single run. Standard curves were generated using the cycle threshold (Ct) values of the positive control plasmid dilutions. Intra-assay variation was determined for both assays by calculating the mean Ct values, standard deviations (SD), and coefficient of variations (CV) separately for each control plasmid DNA dilution (107 – 101). Efficiency curves of the dilutions were performed in uninfected cell culture lysates (spiked with plasmid dilutions) and turtle oral swab extracts from negative samples, with high (67.56 ng/µl) and low (8.61 ng/µl) total DNA concentrations, as measured by spectrophotometry.

2.4. Box Turtle Samples

DNA extracts from twenty-seven eastern box turtles from Tennessee were assayed using conventional PCR in 2007. These samples were assayed by qPCR and repeated for conventional

PCR seven years later. The level of agreement was compared between the results of the conventional PCR and qPCR. The quantity of TerHV1 target DNA in oral swabs was determined using a standard curve method. The copy number of the target DNA was determined from the standard curve generated with ten-fold dilutions of the positive control plasmid for each assay.

2.5. Statistical Analysis

Copy numbers were tabulated and evaluated for normality using the Shapiro-Wilk test.

Mean, median, standard deviation, 95% confidence interval, and 10-90% percentiles were determined for positive cases (copy number) for each assay. The prevalence of TerHV1 was determined (categorical variable assigned; 1=positive, 0=negative) and 95% binomial confidence intervals were determined. Level of agreement (kappa) was determined between both the real- time PCR assays and the conventional PCR previously reported based on prevalence. All statistical analysis was performed using statistical software (SPSS Statistics 22, IBM, Chicago,

IL).

3. Results

3.1. PCR, Standard Curve, Reproducibility

Serial ten-fold dilutions of positive control plasmids assayed with our TaqMan assay generated standard curves based on Ct values (Figure 1). The linear range for TerHV58 was between 1.04×107 to 1.04×101 with an R2 of 1 (slope = -3.389) (Figure 1). Low and high DNA dilution efficiency curves carried out on spiked-uninfected cell lysates and turtle oral swab extracts resulted in an equal performance between the low (slope = -3.327; R2 = 0.998) and the high DNA samples (slope = -3.354; R2 = 1). The amplification plots of TaqMan qPCR TerHV58 are presented in figure 2.

The intra- and inter-assay reproducibility was evaluated for the serial dilutions of the control plasmids. The intra-assay CVs for TaqMan TerHV58 was between 0.04 – 0.27% and the inter-assay CVs were 0.28 – 1.70%, respectively (Table 3). The intra-assay CVs for TaqMan

TerHV64 was between 0.15 – 0.33% and the inter-assay CVs were 0.34 – 1.95%, respectively.

The results indicate high reproducibility between assays at all dilutions. The dynamic range for

7 1 qPCR assays at which point the Ct value of the technical repeats was from 1.04×10 to 1.04×10 .

The TaqMan primer sets, TerHV58 and TerHV64, were successful in detecting either a

58 or 64 base pair length gene segment, respectively, of a conserved portion of the DNA polymerase gene of TerHV1. Both primers had acceptable level of efficiency and variation but

TerHV58 was more efficient, based on the standard curve slope, and had less variation, R2 value, as compared to TerHV64, and was the primer utilized in testing clinical samples.

3.2. Box Turtle samples

Twenty-seven oral swabs were collected from eastern box turtles that presented to the

University of Tennessee from March through October 2007. Previous conventional PCR assay determined an 11% (n=3) prevalence rate for these samples (data not shown). The same samples were re-tested by conventional PCR for this study and a 3.4% (n=1) prevalence rate was determined. Quantitative PCR utilizing primer TerHV58 determined an 11% (n=3) prevalence of

TerHV1, in the same three individuals as conventional PCR in 2007. Observed kappa statistic was 1.0 when comparing 2007 results with qPCR, demonstrating perfect agreement, however agreement (kappa=0.36) was low when comparing 2015 conventional PCR results with qPCR.

The DNA viral copy numbers of the three positive samples determined by qPCR was 890.17,

25.80, 112.83 with a mean purity (A260/A280) of 1.74, as measured by spectrophotometry.

4. Discussion

Herpesviruses have been well described in various reptilian species, especially chelonians, and may pose a threat to affected populations (Johnson et al., 2005; Stacy et al.,

2008; Sim et al., 2015, Ossiboff et al., 2015b). Overall, there is limited published information regarding herpesviruses in box turtle species, particularly in free-ranging populations (Sims et al., Yonkers et al., Ossiboff et al., 2015a; Ossiboff et al., 2015b; Cowan et al., 2015; Marschang et al., 2015). Epidemiologic surveys that characterize the species range and disease extent rely on a sensitive diagnostic assay. A quantitative PCR assay was developed that allows detection of a

DNA polymerase gene segment of TerHV1 with as few as 10 viral copies per reaction. The detection and quantification of viral copies may allow for earlier therapeutic intervention to reduce morbidity and mortality rates in box turtles as has been observed with other species

(Staton et al., 2010).

Taqman qPCR assays utilize a third hybridization or probe in the selection step, which allows for increased specificity of qPCR when testing clinical samples. The assay had highly reproducible intra- and inter-assay variability coefficient of variation of less than 5%. Assay

TerHV58 was more sensitive than assay TerHV64, as seen with a lower coefficient of variation at all dilutions. TerHV58 was designed to be specific for a segment of DNA polymerase gene which previous studies have targeted (Murakami et al., 2001; Marschang et al., 2006). One of the biggest limitations of this assay was lack of specificity determination. While this assay was developed specifically for the detection of a unique sequence of TerHV1, future studies should evaluate the specificity of this assay and its application in diagnosing presence of viral DNA in turtle samples that may posses additional herpesviruses.

DNA from twenty-seven turtle oral swabs collected in 2007 was screened for TerHV1 using conventional at the time of collection and again in 2015, as well as with the developed qPCR. Conventional PCR detection decreased in the 8 subsequent years since collection.

Differences between prevalence detection by conventional PCR may be attributed to the DNA

samples being stored at -20oC with multiple freeze-thaw cycles (Shao et al. 2012). In contrast, use of qPCR revealed equal prevalence detection as the original conventional PCR screening.

Despite the high agreement, as stated by kappa, between the conventional PCR eight years prior and the qPCR assay, qPCR allowed quantification of the viral copies where conventional PCR could not. The storage method may also have degraded DNA of weak positive samples overtime and prevalence may have been higher if qPCR was available at the time of collection. Dual testing (conventional and qPCR methods) on the same samples suggests that indeed the prevalence of infection was low in this population of box turtles.

The current study demonstrates that a TaqMan assay is reliable and sensitive for the detection of a DNA polymerase gene segment of TerHV1. The TaqMan assay has low variability and is a recommended assay for free-ranging eastern box turtle herpesvirus surveys. This assay is capable of detecting a low number of viral copies, which may allow for early or subclinical identification. This assay should be used as a conservation tool in eastern box turtles and can help characterize the epidemiology of herpesvirus.

Figure 1. Standard curve for a Taqman probe-based primer TerHV58 obtained with 10-fold serial dilutions from 1.04×107 to 1.04×101 viral copies per reaction. The graph is plotted against a logarithmic concentration of the serial dilutions.

Figure 2. Amplification plot for the standard curve for TaqMan probe-based primer TerHV58 obtained with 10-fold serial dilutions from 1.04×107 to 1.04×101 viral copies per reaction.

Table 3. Intra- and inter-assay variability for a Taqman probe-based primer TerHV58 obtained with 10-fold serial dilutions from 1.04×107 to 1.04×101 viral copies per reaction for detection of

Terrapene herpesvirus 1 DNA polymerase gene.

Viral Copy Intra-assay Inter-assay Taqman CT mean CT SD CV CT mean CT SD CV 10,400,000 15.93 0.03 0.19% 15.81 0.1 0.63% 1,040,000 19.32 0.03 0.16% 19.15 0.2 1.04% 104,000 22.89 0.01 0.04% 22.8 0.1 0.44% 10,400 26.32 0.03 0.11% 26.12 0.2 0.77% 1040 29.79 0.08 0.27% 29.39 0.5 1.70% 104 32.88 0.08 0.24% 32.67 0.3 0.92% 10 36.13 0.06 0.17% 35.81 0.1 0.28% NTC 38.81

CHAPTER 4

PREVALENCE OF TERRAPENE HERPESVIRUS 1 IN FREE-RANGING EASTERN BOX TURTLES (TERRAPENE CAROLINA CAROLINA)

Abstract: Upper respiratory tract diseases in chelonians, including herpesviruses, have been well described in captive chelonians worldwide but their importance in free-ranging populations is less well known. To help characterize the disease epidemiology of Terrapene herpesvirus 1

(TerHV1), 409 free-ranging eastern box turtles (Terrapene carolina carolina) in Tennessee and

Illinois were sampled in 2013 and 2014. Whole blood and swabs of the oral mucosa were collected from 365 adults (154 female, 195 male, 16 unknown sex) and 44 juveniles. A Taqman qPCR assay targeting a portion of the DNA polymerase gene was performed on DNA from all oral swabs. The prevalence of detection was 31.3% (n=128). TerHV1 prevalence was significantly higher in July (50%; n=67) compared to September (38.3%; n=44) and May

(10.6%; n=17) (p<0.0001). The prevalence was not variable across state (p=0.492), sex

(p=0.506), age class (p=0.235), or habitat (p=0.328). Turtles sampled in 2014 had a significantly higher prevalence (50%; n=98) than in 2013 (14.1%; n=30) (p<0.001). Packed cell volume

(25.5%) and total solids (4.9 mg/dl) were significantly higher in positive turtles than negative turtles (23.0%, p=0.001; 4.3 mg/dl, p<0.0001), respectively. Positive turtles had higher eosinophils (p=0.035), lower lymphocytes (p<0.001), and lower monocytes (p<0.001). No clinical sign was associated with detection of herpesvirus (p=0.313 - 0.677). Eastern box turtles were detected with widespread DNA evidence of Terrapene herpesvirus 1 infection, and the epidemiology of this virus may aid in management of free-ranging and captive individuals.

Key Words: Disease, epidemiology, herpesvirus, Terrapene carolina, qPCR.

INTRODUCTION

Eastern box turtles (Terrapene carolina carolina) are small, terrestrial turtles that are widespread throughout eastern United States forests (Donaldson and Echternacht, 2005). Their status was recently downgraded to vulnerable by the International Union on the Conservation of

Nature (IUCN) due to population declines in several areas throughout their range (van Dijk,

2013). Specific causes for the decline are not entirely known, but are attributed to anthropogenic factors including road and mowing mortalities, collection, nest depredation, disturbances of nest sites by off-road vehicles, and habitat loss (Henry, 2003; Nazdrowicz et al., 2008; Schrader et al.,

2010). While a combination of factors is likely playing a role in the population declines of the box turtle, disease outbreaks due to upper respiratory pathogens have been emerging across the eastern United States in chelonians (Sim et al., 2015; Yonkers et al., 2015). Several upper respiratory pathogens have been observed during these outbreaks in captive and free-ranging box turtles, including ranavirus, herpesvirus, and Mycoplasma (Brown and Sleeman, 2002; De Voe et al., 2004; Pettan-Brewer et al., 1996; Soares et al., 2004).

Herpesviruses are large, enveloped viruses with a double-stranded DNA genome

(McGeoch and Gatherer, 2005). Reptilian herpesviruses fall into the family Herpesviridae, which is divided into three subfamilies, Alpha-, Beta-, and Gammaherpesvirinae (Davison, 2010;

ICTV, 2014). Currently, all reptilian herpesviruses appear to be most homologous to sequences from the subfamily Alphaherpesvirinae (Bicknese et al., 2010; Marschang et al., 2006). Reports of clinical and pathologic findings of herpesvirus in marine turtles and tortoises are frequent

(Chaloupka et al., 2009; Johnson et al., 2005; Pettan-Brewer et al., 1996; Salinas et al., 2011;

Stacy et al., 2008). However, there are few investigations in Emydid turtles, including box turtles, and it is unknown if free-ranging populations experience similar disease events (Sim et al., 2015;

Yonkers et al., 2015).

Historically, herpesvirus in emydid turtles was restricted to a captive painted turtle

(Chrysemys picta) that died with marked necrosis and intranuclear inclusion bodies in the liver and lungs consistent with herpesvirus like-particles (Cox et al., 1980) and two captive map turtles (Graptemys spp.) exhibiting lethargy, anorexia, and subcutaneous edema for a week

(Jacobson et al., 2012). Recent evidence, suggests that herpesvirus affecting the family Emydidae might be more common than previously believed (Ossiboff et al., 2015a; Ossiboff et al., 2015b;

Sim et al., 2015; Yonkers et al., 2015). A case report described a molecularly distinct novel herpesvirus, Terrapene herpesvirus 1, that caused upper respiratory signs in two separate collections of eastern box turtles in Georgia and Maryland (Sim et al., 2015). A second herpesvirus (Terrapene herpesvirus 2) was discovered in Florida in an eastern box turtle associated with a fibropapilloma lesion (Yonkers et al., 2015). Herpesvirus infection was diagnosed via PCR in two asymptomatic turtle species, painted turtles (Chrysemys picta) and a northern map turtle (Graptemys geographica) (Ossiboff et al., 2015a). A northern map turtle displaying lethargy and nasal discharge was also diagnosed with herpesvirus by PCR (Ossiboff et al., 2015a). Additionally, three novel herpesvirus’ were identified in bog turtles (Glyptemys muhlenbergii), wood turtles (Glyptemys insculpta), and spotted turtles (Clemmys guttata)

(Ossiboff et al., 2015b). Prevalence of herpesvirus in bog and woods turtles was over 50% whereas the prevalence in spotted turtles was 6% (Ossiboff et al., 2015b). Herpesvirus infection in Emydid’s has yet to be reported in Tennessee and Illinois.

Despite the occurrence of herpesviruses in the box turtles, the impact on in free-ranging populations is still unknown. Therefore, the specific objectives of this study were to: 1) estimate the prevalence of Terrapene herpesvirus 1 from free-ranging populations of eastern box turtles in

two states in the US using qPCR, and 2) Determine if qPCR prevalence will differ based on sex, age class, location, hematologic findings, or clinical signs.

MATERIALS AND METHODS

Sample populations

Free-ranging turtles, as part of multi-year long-term health assessments, were sampled using human and canine search (Rose and Allender et al., 2011). Turtles were sampled in 2013 and 2014 in Tennessee (36.00°, -84.22°) and Illinois (Kennekuk County Park [40.19°, -87.72],

Forest Glen Nature Preserve [40.01°, -87.57°], Kickapoo State Park [40.14°, -87.74], and

Middlefork Woods [40.23°, -87.789°]). GPS locations were collected on all animals encountered.

Sampling period for the turtles occurred during May, July, and September, for both years.

Habitat, categorized as forest, field, or edge, was recorded for all sampled turtles.

Field methods

Turtles were weighed to the nearest gram, measured to the nearest centimeter, and categorized by sex and age class before venipuncture was performed. Age class classification

(adult, juvenile, unknown) was assigned based on carapace length and annuli count. Turtles with a carapace length less than 9 cm (3.5 in) and an annuli count less than or equal to seven were characterized as juveniles (Dodd, 2001). Sex was classified as male, female, or unknown based on plastron concavity and tail length. Blood samples (less than 0.8% body weight) were collected from the subcarapacial sinus using 3 mL syringe and a 22 gauge needle under manual restraint.

The samples were immediately placed in lithium heparin coated microtainers (Becton Dickinson

Co., Franklin Lakes, NJ 07417) and gently inverted several times. Oral epithelium and mucus

was collected from within the oral cavity using cotton-tipped plastic handled applicators (Fisher

Scientific, Pittsburgh, PA 15275). The swab tips were placed in a 2.0 mL polypropylene microcentrifuge tube (Fisher Scientific, Pittsburgh, PA 15275) for storage at -20C. The blood and swabs were labeled with a unique identification number and remained on wet ice until transport to the lab within 4 hours.

A single veterinarian evaluated clinical signs (MCA). The presence or absence of clinical signs compatible with those previously described for herpesvirus infections was recorded as absent (0) or present (1). Specific clinical signs included nasal discharge/rhinitis, ocular discharge, oral discharge/stomatitis, oral plaques, and respiratory distress/open mouth breathing.

All procedures were approved by the Institutional Animal Care and Use Committee approval through the University of Illinois (Protocol: 13061).

Complete blood counts

Twenty-five microliters of lithium heparinized whole blood were aliquoted for total white blood cell count determination according to the manufacturer's instructions (Avian Leukopet,

Vetlab Supply, Palmetto Bay, FL 33157). Blood smears were then made by conventional methods (Campbell and Ellis, 2007), air dried, and stained with Diff Quick (Jorgensen

Laboratories, Inc., Loveland, CO 80538). Differential blood cell counts were performed by a clinical pathologist (ARM) or certified veterinary technician (JP). Packed cell volume was manually performed from two microhematocrit tubes and averaged. Total solids were determined using a refractometer of plasma recovered from the microhematocrit tubes and averaged.

Polymerase chain reaction

Quantitative PCR was performed as previously described (Kane et al., 2016). Briefly,

DNA was extracted from oral swabs using manufacturer’s instructions (QIAmp Blood Mini Kit,

QIAGEN Inc., Valencia, CA, 91355). TaqMan assay was performed using forward (5’-

TCTATTGGGCGAGCTGTTGAC-3’), reverse (5’-GAAAAGCCATACGCCTACGC-3’), and probe (5’-ACTGGCTGGCCTTG-3’), targeting a 58 base pair segment of the herpesvirus DNA polymerase gene. All samples were assayed in three technical repeats using a real-time PCR thermocycler (7500 ABI real-time PCR System, Applied Biosystems, Carlsbad, CA, 92008), analyzed using commercial software (Sequence Detection Software v2.05, Applied Biosystems), and results averaged.

Geospatial analysis

Mapping and analysis was performed using ArcGIS software (ArcGIS 10.1, ESRI,

Redlands, CA 92373). Basemaps were acquired from ArcGIS. Ortho images were used to demonstrate urban features more accurately. Original data points (that corresponded to capture location of each turtle) were entered into an excel spreadsheet in degree minutes and imported into ArcCatalog. Known locations of TerHV1-positive turtles were mapped using satellite images and point data. The hot spot analysis tool identified statistically significant hot spots and cold spots using Getis-Ord Gi* statistic, which was then performed. It created a new output feature class with a z-value and p-value for each. Z-values are an illustration of the standard deviations and p-value is the probability the observed pattern was created by a random process. This analysis graphically illustrated prevalence of the disease and health parameters that could be influencing infection across the region.

Statistical analyses

Descriptive statistics were tabulated for all continuous variables (weight, viral copy number, pack cell volume, total solids, white blood cell count). Normality was assessed using the

Shapiro-Wilk test, Q-Q plots, skewness, and kurtosis. Mean, standard deviation, and 95% confidence interval are reported for normally distributed data. Median and 10-90% are reported for non-normally distributed data. An independent samples t-test and a one-way analysis of variance (ANOVA) were used to determine between and within group differences respectively.

Whereas a Mann-Whitney U and Kruskal-Wallis one-way analysis of variance were used when data was non-normally distributed. Overall prevalence proportions (based on qPCR) and their

95% confidence intervals were computed. Pearson’s Chi-square and Fisher’s exact tests were used to evaluate age class, sex, season, state, location, habitat, and clinical signs based on qPCR test results. Statistical significance was considered when p<0.05. Odds ratios were calculated for each season based on Terrapene herpesvirus 1 status. All analysis was performed using statistical software (SPSS 23, IBM statistics, Chicago, IL 60606).

RESULTS

There were 409 turtles sampled from 2013 and 2014, with 212 (51.8%) turtles sampled in

2013 and 197 (48.1%) in 2014. There were 200 (48.9%) individuals sampled from Illinois and

209 (51.1%) from Tennessee. All turtles had sampling season recorded; 160 (39.1%) from May,

134 (32.8%) from July, and 115 (28.1%) in September. Of the 409 animals with age and sex recorded, 195 (47.7%) were male, 154 (37.7%) were female, 365 (89.2%) were adult, and 44

(10.8%) were juvenile. The majority of turtles were found in the forest (n=273, 67%) as compared to edge habitats (n=46, 11%), or fields (n=27, 7%).

Median weight for females (403 g, Min/Max: 142 – 710, 10-90 percentiles: 286 – 608) and males (410g, Min/Max: 178 – 761, 10-90 percentiles: 316 – 584) was not significantly different (p=0.835). Median weight for adults 410.5 g (Max/Min: 126 –761, 10-90 percentiles:

312 – 600) was significantly more than juveniles (median: 159.5 g; Max/Min: 30 – 347, 10-90 percentiles: 66.5 – 286) (p<0.001). Mean weight of turtles sampled in Illinois (466.4 g) were significantly heavier than those sampled from Tennessee (356.6g; p<0.001).

One hundred and twenty-eight turtles tested positive by qPCR for TerHV1 resulting in an overall prevalence of 31.3% (95% CI: 27-36%). The majority of positive turtles were male

(n=69; 53.9%; 95% CI: 27-40%) and adults (n=116; 90.6%; 95% CI: 27-37%). However, there was no significant difference in prevalence based on age class (p=0.235) or sex (p=0.506). There were significantly fewer (n=30; 23.4%; 95% CI: 10-18%) positive turtles in 2013 than in 2014

(n=98; 76.6%; 95% CI: 43-57%) (p<0.001). Sixty-six of the 128 positive turtles were sampled in

Tennessee (51.6%; 95% CI: 25-38%) and 62 in Illinois (48.4%; 95% CI: 24-38%); however, there was no significant difference in prevalence when evaluating state (p=0.492). The total number of positive turtles was significantly higher in July (n=67; 52.3%; p<0.001; 95% CI: 41-

59%) (Figure 3 (TN), IL data not shown), than both May (n=17; 13.3%; p<0.001; 95% CI: 5-

15%) (Figure 4 (TN), IL data not shown) and September (n=44; 34.4%; p=0.042; 95% CI: 29-

47%) (Figure 5 (TN), IL data not shown) and turtles sampled in July were four times more likely to be positive than turtles in May and September (95% CI: 2.25-5.46). There was no statistically significant difference in prevalence based on habitat (p=0.328). Prevalence was not significantly

different between the four locations within Illinois (p=0.097). Detection of positive turtles was non-uniformly distributed across the landscape in TN (Figure 6).

There were 409 oral swab samples collected. DNA viral copy was significantly higher in

2014 (p=0.001) as compared to 2013. The median TerHV1 virus copy number in positive turtles from both years was 106 (Min/Max: 10.83 – 730993, 10-90 percentiles: 19.7 – 542.6).

Packed cell volume (PCV) was significantly higher in positive turtles than negative turtles (p=0.039), where as total solids (TS) was not significantly higher (Table 4). Adults had a significantly higher (p<0.001) PCV than juveniles (Table 4). Eosinophils were higher in positive turtles where as lymphocytes and monocytes were lower (Table 5).

All 409 animals had full physical examinations performed and clinical signs were not significantly associated with TerHV1 prevalence; nasal discharge (n=3; p=0.677), ocular discharge (n=2; p=0471), ocular swelling (n=2; p=0.529), oral plaque (n=2; p=0.471), or respiratory distress (n=1; p=0.313). Herpesvirus status in association with clinical signs was only seen in two turtles; one had nasal discharge and ocular swelling, and the other was open mouth breathing. The remainder of clinical signs were seen in TerHV1 negative box turtles.

DISCUSSION

Herpesvirus infections in eastern box turtles have previously been seen in Georgia,

Maryland (Sim et al., 2015), and Florida (Yonkers et al., 2015), but this is the first report from

Tennessee and Illinois. The prevalence in this study was lower than previously reported in captive box turtles (Sim et al., 2015) and free-ranging bog turtles (Ossiboff et al., 2015b), similar to that reported in free-ranging desert tortoises using an ELISA (Jacobson et al., 2012), and higher than that reported for Mediterranean and Hermann’s tortoises (Origgi et al., 2001). Box

turtle populations are gradually decreasing over time and disease may be an attribution, as has been seen in other free-ranging emydids (Ossiboff et al., 2015b; Todd et al., 2010). The prevalence seen in this study and the minimal disease associated clinical signs warrants further study to determine if herpesvirus is a contributor to population declines.

Several species in the order Testudinidae have been fatally infected with herpesvirus infections (Cox et al., 1980; Hervas et al., 2002; Jacobson et al., 1982; Jungwirth et al., 2014;

Muro et al., 1998). Historically, herpesvirus associated mortality rates in tortoises have ranged between 50-100% (Drury et al., 1998; Lange et al., 1989). More recently, herpesvirus via PCR was diagnosed in a northern map turtle exhibiting weakness and nasal discharge as well as four asymptomatic turtles from the same enclosure (Ossiboff et al., 2015a). PCR was also used to diagnose herpesvirus in a captive juvenile eastern box turtle that died after emerging early from brumation with lethargy, dehydration, and dyspnea (Sim et al., 2015). Mortality rates in the box turtles found in this study were not assessed due to loss of follow up.

Several clinical signs have historically been associated with herpesvirus infection in chelonians, including nasal discharge, ocular discharge, ocular swelling, oral plaques, and respiratory distress (Johnson et al., 2005; Origgi et al., 2004; Soares et al., 2004). Few clinical signs were seen in positive turtles in this study, and there was no statistically significant relationship between clinical signs and TerHV1 status. The clinical signs seen may be associated with other disease processes, which were not examined during this study. A transmission study evaluating herpesvirus associated clinical signs in which other factors are controlled is warranted.

In this study, we found that adult turtles were not more likely to be detected with herpesvirus than juveniles and this is opposite to what has been reported in the literature for desert tortoises and painted turtles (Cox et al., 1980; Johnson et al., 2005; Pettan-Brewer et al.,

1996). The higher prevalence seen in adults, similar to that seen in wood mice, may be due to an unknown physiological process, such as breeding or basking, or increased virus exposure overtime compared to juveniles (Knowles et al., 2012). The higher prevalence seen in 2014 as compared to 2013 may be due to environmental factors affecting disease expression

(precipitation, temperature, microhabitat) or yet unknown factors of herpesvirus pathogenesis.

Seasonal differences and environmental temperatures may play a role in prevalence and may be attributed to increased activity of turtles, increased interactions with conspecifics, or increased viral persistence at warmer temperatures. Increased infectivity and severity of herpesvirus due to higher temperatures has been demonstrated in green sea turtles, and may explain the seasonality associated with the results of this study (Haines and Kleese, 1977). Future studies should focus on characterizing the epidemiology in relation to environmental temperatures by conducting temperature transmission trials or an ongoing investigation of prevalence in relation to date and location.

Turtles found in the forest had higher herpesvirus detection than either turtles found in the field or edge habitats. However, herpesvirus was ubiquitous, and overall, habitat had minimal effect on TerHV1 prevalence. Additional research should focus on microhabitats, such as logs or substrates, to further differentiate factors that may be contributing to increased detection. Forests and fields are natural habitats of eastern box turtles, while edge habitats are created during habitat destruction or fragmentation (Budischak et al., 2006; Dodd, 2001). The higher prevalence found in the forest may be attributed to a type 2 error in which fewer turtles were found in field habitats. Future studies should encourage a larger sample size in deteriorating habitats.

Positive turtles had a higher PCV, which may be attributed to dehydration associated with disease, however, these parameters, in this population have previously been shown to increase

with season (Kimble and Williams, 2012). Therefore, changes observed here may reflect seasonal differences rather than the presence of herpesvirus. The perceived lymphopenia and monocytopenia in positive turtles is not unusual as this is commonly seen in reptiles with viral infections (Campbell, 2014). The eosinophilia in positive turtles may be a seasonal influence, attributed to herpesvirus prevalence, or immune stimulation due to incidental parasitic infections

(Campbell, 2014).

The mode of transmission of herpesviruses in chelonians is largely unknown but suspected to spread via nasal and oral discharge (Hoff et al., 1984; Origgi et al., 2004; Soares et al., 2004). Viral detection is demonstrated in several different tissues post mortem, (Hervas et al.,

2002; Marschang, 2011) however; clinical antemortem samples typically involve swabs of the oral mucosa due to the viral tissue predilection of mucoepithelial cells (Hervas et al., 2002;

Origgi et al., 2001; Sim et al., 2015; Teifke et al., 2000) and the infrequent nature of viremia in the literature. However, future investigations should include whole blood to further characterize the pathogenesis of this virus in box turtles.

This study utilized qPCR to survey populations of eastern box turtles for herpesvirus.

This method is considered more sensitive than conventional PCR, allowing the possible detection of subclinical or latent carriers (Alfaro-Nunez and Gilbert, 2014; Greenblatt et al.,

2004; Murakami et al., 2001). However, there is no evidence whether to support subclinical or latency of herpesvirus in chelonians as in mammals (Hardman et al., 2012; Landolfi et al., 2005).

It is possible that positive turtles may experience a latency period; however, due to loss of follow up, it is unknown if the positive turtles experienced disease-related mortality events. To better aid in characterizing the epidemiology of chelonian herpesvirus, future studies should perform long-term surveillance tracking programs.

This was the largest cross-sectional study to estimate the prevalence of TerHV1 in free ranging eastern box turtles performed to date. It demonstrated that TerHV1 prevalence in free ranging chelonians is ubiquitous and age, sex, and geographical influences were minor whereas seasonal influences were major. Further investigation involving environmental factors is needed to understand the relationship between disease expression and season. Clinical signs were not associated with herpesvirus detection and further research is needed to evaluate the association of herpesvirus susceptibility and severity with co-infections. Detection in oral swabs was reliable, but future studies should utilize blood to aid in understanding this virus’ pathogenesis. Findings were consistent with mild disease involving minimally associated clinical signs and no significant sex or age-specific prevalence. Based on this study and other published reports, free ranging eastern box turtles are sensitive to TerHV1 infections. Further work is needed to determine the threat of Terrapene herpesvirus 1 in free-ranging eastern box turtle sustainability.

Based on the results of this study, qPCR is recommended as the diagnostic modality for testing clinical samples against TerHV1 in box turtles. Little is known of this virus in eastern box turtles; however, this report aids in characterizing the presence of TerHV1 and minimizes a deficiency that previously existed.

Table 4. Packed cell volume (PCV) and total solids (TS) of Eastern box turtles (Terrapene carolina carolina) in Tennessee and Illinois.

10%-90% Test Status Median Number Min/Max p value percentiles PCV Positive 25.5 132 21.5-35.4 6.0-43.0 0.039 Negative 23 273 19.3-30.0 4.0-45.5 2013 23.5 213 15.0-30.5 6.0-45.5 0.327 2014 25 192 15.2-34.0 4.0-45.5 Males 25.5 201 17.1-33.9 6.0-45.5 0.803 Females 22 162 13.2-30.4 4.0-45.5 Adult 24.4 362 15.2-33.0 4.0-45.5 <0.001 Juvenile 21.8 42 14.6-27.0 9.0-29.0 TS Positive 4.8 131 3.4-6.2 0.7-20.0 0.569 Negative 4.2 273 2.5-6.1 1.7-9.7 2013 4.2 213 2.5-6.1 1.5-20.0 0.192 2014 4.6 191 3.1-6.3 0.7-8.7 Males 4.3 200 2.9-6.0 1.1-20.0 0.218 Females 4.8 162 3.0-6.7 0.7-9.7 Adult 4.5 361 2.9-6.2 0.7-20.0 0.214 Juvenile 4 42 2.4-5.6 1.8-6.9

Table 5. White blood cell differential counts of eastern box turtles (Terrapene carolina carolina) with and without presence of Terrapene herpesvirus 1.

10%-90% Test Status Median Number Min/Max p value percentiles WBC Positive 7499 128 4348-15325 2406-24440 0.383 Negative 7948 281 4353-13636 156-21641 Eosinophils Positive 2212 128 864-4721 0-11103 0.041 Negative 1869 281 564-4472 0-12408 Heterophils Positive 2558 128 1315-5670 0-20529 0.725 Negative 2502 281 862-5310 0-10727 Lymphocytes Positive 775 128 223-1660 0-4941 <0.001 Negative 1031 281 322-2267 0-6497 Monocytes Positive 175 128 0-2178 75-661 <0.001 Negative 306 281 0-1597 58-954 Basophils Positive 1131 128 176-3255 0-8360 0.166 Negative 1237 281 251-3419 0-10697

Figure 3. Hotspot analysis of individual eastern box turtles (Terrapene carolina carolina) in

Tennessee based on detection of Terrapene herpesvirus 1 in May using quantitative PCR.

Figure 4. Hotspot analysis of individual eastern box turtles (Terrapene carolina carolina) in

Tennessee based on detection of Terrapene herpesvirus 1 in July using quantitative PCR.

Figure 5. Hotspot analysis of individual eastern box turtles (Terrapene carolina carolina) in

Tennessee based on detection of Terrapene herpesvirus 1 in September using quantitative PCR.

Figure 6. Overall hotspot analysis of individual eastern box turtles (Terrapene carolina carolina) in Tennessee based on detection of Terrapene herpesvirus 1 using quantitative PCR.

CHAPTER 5 CONCLUSIONS AND FUTURE DIRECTIONS

Chelonians are threatened with significant population declines and disease impacts in these species have been minimally investigated. The epidemiology of both common and rare diseases is needed to determine effects on ecosystem health. The research in this thesis contributes to further characterization of the epidemiology of herpesvirus disease in chelonians and can be used to develop a better understanding of the diagnosis, transmission, and management of this virus in captive and free-ranging populations.

A quantitative PCR assay was developed that can detect the DNA polymerase gene of

Terrapene herpesvirus 1. The TaqMan qPCR assay was found to be highly sensitive for use in chelonians and detected low quantities of viral DNA. qPCR may be used for detecting lower viral quantities in turtles during periods of latency or in subclinical carriers or reservoirs, which have yet to be identified and needs to be investigated. While PCR has historically been utilized as the diagnostic modality for herpesvirus infections, and still remains useful for identifying novel herpesviruses, the evidence in this thesis provides a superior diagnostic technique for

TerHV1, specifically. Future studies should investigate this assay in multiple species to assess and determine specificity.

While this thesis provides new insight into the epidemiology of herpesvirus disease in chelonians, there is much more that needs to be investigated. To date, there have been numerous reports of herpesvirus epizootics in tortoises and marine turtles; however, minimal reports have been documented in Emydids. Epizootics may be occurring more frequently than suspected and may be underreported or unnoticed due to the natural history characteristics of these chelonians.

This project was a short term investigation describing and monitoring the health of these animals

and future long-term studies, involving radiotelemetry, should be performed to evaluate and further strengthen our understanding of the epidemiology of herpesvirus infections in eastern box turtles to be able to protect this species.

One aspect of the epidemiology of herpesvirus, which warrants further investigation, is this virus’ transmission in box turtles. Transmission studies, utilizing pathogen free turtles, should be performed in these species, to help understand the role this virus plays during co- infections. This study confirms the necessity of a transmission study to elucidate the connection between the clinical signs seen in herpesvirus positive turtles with other disease processes since presence of disease was minimally associated clinical signs. A transmission study should additionally focus on the unknown latency period with disease recrudescence, as seen in mammals.

Ecothermic animals rely heavily on their environment and this thesis concludes an association between viral presence and seasonal differences. Future studies should investigate and provide additional insight into the effect of the environment on the pathogenesis of this virus as well as the role that preferred optimal temperature zones play on virus replication.

Temperature may also allow persistence of viral particles in the environment and may increase transmission between conspecifics.

In conclusion, the research in this thesis revealed and contributed valuable information on herpesvirus previously unknown and aided in characterizing this disease in box turtles. This project can be used as a template for further investigations of the heath and disease in other chelonian populations. Future studies should continue to monitor this disease as well as other diseases effecting box turtles, as these animals are continual sentinels of environmental health.

CHAPTER 6

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