The Role of Emerging Pathogens in Amphibian Population Declines: Experimental Evidence

AN ABSTRACT OF THE THESIS OF

Natalie M. Hambalek for the degree of Master of Science in Zoology presented on May 17, 2016.

Title: The Role of Emerging Pathogens in Amphibian Population Declines: Experimental Evidence

Abstract approved: ______Andrew R. Blaustein

Rapid rates of biodiversity loss have supported the notion that Earth is experiencing a sixth major extinction event. The causes of worldwide biodiversity loss are multifaceted and context dependent. One of the most prominent groups experiencing population declines and extinctions are amphibians. Several pathogens and their associated diseases are especially significant contributors to amphibian population declines. These include the chytrid fungus, Batrachochytrium dendrobatidis, the related but highly divergent fungal pathogen, B. salamandrivorans, and ranaviruses.

In Chapter 1, I summarize the life-history traits of these three amphibian pathogens.

In Chapter 2, I present a synthesis of these three emerging infectious pathogens by assessing their broad effects on amphibian hosts as found in experimental studies. I also examined the interactive effects of these pathogens with other potential and known contributors of amphibian population declines that have been experimentally studied. Well-designed experimental studies are critical for understanding the impacts of disease. However, inconsistencies in experimental methodologies often hinder our ability to form valuable comparisons and conclusions. Chapter 2 highlights the importance of implementing standard experimental protocols and reporting.

Furthermore, we emphasize the significance of investigating the roles of multiple environmental and anthropogenic stressors.

The effects of B. dendrobatidis and ranaviruses on their hosts have been well documented but the effects of coinfection with these pathogens are poorly understood. In Chapter 3, I experimentally examined the effects of simultaneous and independent exposure of B. dendrobatidis and ranavirus on survival, growth, and activity levels in two amphibian host species, the Pacific treefrog (Pseudacris regilla) and the western toad (Anaxyrus boreas). I predicted that coinfection with Bd and Rv would have increased lethal and sublethal effects on hosts compared with hosts that were infected with one of the pathogens independently. The results showed antagonistic effects of concurrent exposure to B. dendrobatidis and ranavirus, with individuals exposed to only one pathogen exhibiting more lethal and sublethal effects compared to individuals exposed to both pathogens. These results support the proposition that the dynamics behind multiple pathogen exposure are complex and that one pathogen may mitigate the effects of another to induce response variation.

The Role of Emerging Pathogens in Amphibian Population Declines: Experimental Evidence

by Natalie M. Hambalek

A THESIS

submitted to

Oregon State University

in partial fulfillment of the requirements for the degree of

Master of Science

Presented on May 17, 2016 Commencement June, 2016

Master of Science thesis of Natalie M. Hambalek presented on May 17th, 2016

APPROVED:

Major Professor, representing Zoology

Chair of the Department of Integrative Biology

Dean of the Graduate School

I understand that my thesis will become part of the permanent collection of Oregon State University libraries. My signature below authorizes release of my thesis to any reader upon request.

Natalie M. Hambalek, Author

ACKNOWLEDGEMENTS

I would first like to thank my advisor Andy Blaustein for his unwavering support, sense of humor, and guidance throughout the last few years. There is no way to fully express my gratitude for giving me this opportunity and challenging me to go above and beyond to become a well-rounded scientist and a balanced human being. I have been lucky to learn from him. I am also grateful for Kathy Blaustein for her support and kindness. Thank you to my amazing committee: Dave Stone, Jeri

Bartholomew, and Lindsay Biga for their thoughtful advice and support. Jim Rivers,

Michelle Hladik, and Kelly Smalling all provided invaluable perspective and support in the beginning of my graduate work. I am grateful for Dede Olson for her helpful advice regarding work and beyond. Jason Hoverman has been an amazing collaborator for my graduate thesis work. Cathy Law of STEM Academy has been an incredible source of encouragement. Working alongside her is something I will take away from my graduate career and I am looking forward to providing impactful service in my future.

I am exceptionally grateful for my fellow Blausteinites, both past and present.

Barbara Han, Steph Gervasi, Julia Buck, Paul Bradley, Tara Chestnut, John

Romansic, Giselle Xie and Linsdsay are all amazing scientists and role models. I could not have made it through these years without the mentorship of Trang Dang and

Jenny Urbina, my lab moms and friends. I will dearly miss working alongside Trang in our tiny Cordley office and sharing laughs with Jenny. They made the lab a welcoming place and I treasure the time we spent together. Cassie Nix, Carmen

Harjoe, and Paul Synder made lab meetings and work fun throughout the ups and downs. I could not have gotten through the last few months of graduate school without jumping over the final hurdles with Delia Negru and I really cherish our friendship. Emily Reynolds, once a student of mine, now a fellow colleague, I will miss our conversations and laughs. Thanks to Elva Manquera for being the trustiest undergraduate lab assistant ever. I know you will go on to accomplish great things!

I am incredibly thankful to have been supported through teaching assistantships throughout the years. I have learned immensely from Lori Kayes’ commitment to education and it has been a pleasure working alongside her, Bob

Mason, and the rest of my fellow TAs. I also wanted to thank the undergraduate students that I have had the pleasure to work with over the years. I have learned as much from them as they have from me. The Zoology department has been a wonderful and constant source of support. Tara, Traci, Trudy, and Jane have been great to work with on logistical matters. I am also extremely grateful to those from my undergraduate career who were instrumental to helping me get to where I am now. Merith Weisman, Karin Jaffe, Derek Girman, Daniel Smith, Gary Cherr,

Cynthia Boaz, and Nick Geist, their perspectives have influenced me to be the best I can be academically, professionally, and personally.

I have the absolute BEST COHORT IN THE WORLD: Brian Tanis, Danielle

Tom, David Taylor, Ian Morelan, Jenna Sullivan, Katie Dziedzic, Kyle Coblentz,

Leah Segui, and Trevor Tivey. I am so grateful for the moments and adventures we have shared both in and outside of work. EBC forever and Gladiators for life. To my friends outside my cohort: Brianna Gil, Adam Karmally, Aurora Tivey, Shannon

Hennessey, Danielle Marias, Cara Keegan, and Alana Rose, thank you for providing countless laughs, escapades, and encouragement during my time here. Katie, Aurora,

Shannon, Emily, and Peregrin, I will miss our Monday nights more than you can imagine!

A HUGE thank you to my family for their constant optimism and keeping me grounded. To my mom, Ani, and my sisters, Meaghan and Sabrina, for always being my #1 cheerleaders and my main source of motivation. My grandma Arsiné for being my best friend and never letting me forget who I am. Tristin, Devin, Dylan, Krysten,

Hampo, Suzy, Patrick, Liza, and Brian, I love you all and your belief in me has kept me going. Thank you to Marabeth and Mohamed Karmally for providing continual guidance and advice throughout the years. My adorable felines, Callie and Beasley, have been the best soundboards and sources of comfort. Finally, thank you to my partner in life, Ali Karmally, for your understanding, compassion, and relentless support through this journey. There are no words that could come close to expressing how lucky I feel to have you by my side.

Behind every able man, there are always.

CONTRIBUTION OF AUTHORS

Emily Reynolds assisted with data collection in Chapter 2 and experiment execution in Chapter 3. Jenny Urbina assisted with data collection and provided helpful editing for Chapter 3. Trang Dang offered perspective on Chapter 2 and assisted with Bd culture and inoculation for Chapter 3. Jason T. Hoverman offered perspective to Chapters 2 and 3 and provided ranavirus culture for the experiment in Chapter 3. Barbara Han provided significant insight to Chapter 2. Dede Olson was involved with the design and writing with Chapters 2 and 3. Andrew Blaustein assisted with design and writing and contributed to data interpretation and conclusions of all Chapters.

TABLE OF CONTENTS

Page

CHAPTER 1: GENERAL INTRODUCTION……………………………………… 1 Batrachochytrium dendrobatidis……………..……………... 4 Batrachochytrium salamandrivorans………….….……….... 5 Ranavirus……………………………………………………. 6

CHAPTER 2: THE IMPACTS OF EMERGING INFECTIOUS DISEASES AND OTHER CO- FACTORS ON AMPHIBIANS: A REVIEW OF EXPERIMENTAL STUDIES Abstract……………………………………………………... 9 Introduction…………………………………...... 10 Methods…………………………………………………….. 13 Results………………………………………...... ……. 13 Discussion………………………………………………….. 34

CHAPTER 3: COMPLEX DYNAMICS OF TWO EMERGING PATHOGENS, BATRACHOCHYTRIUM DENDROBATIDIS AND RANAVIRUS, ON AMPHIBIAN HOSTS Abstract……………………………………………………... 63 Introduction…………………………………...... 64 Methods…………………………………………………….. 65 Results………………………………………...... ……….. 70 Discussion…………………………………………………... 72

CHAPTER 4: UNDERSTANDING THE ROLES OF EMERGING INFECTIOUS DISEASE ON AMPHIBIAN POPULATION DECLINES: SUMMARY AND IMPLICATIONS…………………………………………………...... 80 Bibliography………………………………………………... 83

Figure Page LIST OF FIGURES

2.1 Trends in experimental amphibian disease ecology over time………… 37

2.2 Abiotic and biotic drivers of amphibian host-pathogen dynamics……. 37

2.3 Trends in Batrachochytrium dendrobatidis and ranavirus literature….. 38

2.4 Map depicting experimental studies on Batrachochytrium dendrobatidis and ranavirus with respect to host genus and geographic range………. 39

3.1 Kaplan-Meier survival curves for Pseudacris regilla and Anaxyrus boreas in response to pathogen exposure……………………………………… 77

3.2 Mean Batrachochytrium dendrobatidis infection loads……………….. 78

3.3 Proportion of active tadpoles in response to pathogen exposure……… 79

LIST OF TABLES

Table Page

2.1 Overview of the effects of experimental studies on amphibian hosts …………………………………………………………………… 40 2.1.a Batrachochytrium dendrobatidis………………………………. 40 2.1.b Batrachochytrium salamandrivorans………………………….. 51 2.1.c Ranavirus………………………………………………………. 53

3.1 Final snout-vent-length and mass measurements……………………… 76

3.2 Hazard ratios and associated p-values for within and between-species analyses……………………………………………… 76

CHAPTER 1: GENERAL INTRODUCTION

Rapid rates of biodiversity loss over the last few centuries have supported the notion that

Earth is experiencing a sixth major extinction event (Dirzo et al. 2014; Ceballos et al. 2015).

Current species extinction rates are higher than pre-human background rates suggesting this 21st century crisis is largely attributed to anthropogenic change (Wilson 1992; Pimm et al. 1995;

Wake & Vredenburg 2008; Barnosky et al. 2011; Dirzo et al. 2014; Ceballos et al. 2015). At the forefront of this crisis, amphibians are a rapidly declining vertebrate group, (Stuart et al. 2004;

Alroy 2015), though not one factor alone bears total responsibility for their overall decline

(Blaustein et al. 2011). Although habitat destruction and fragmentation are the most documented factors affecting amphibian populations (Alford & Richards 1999), multiple anthropogenic and natural stressors including pesticides, climate change, over-harvesting for the pet and food trades, invasive species, predation, and infectious diseases have the capacity to work both independently and synergistically to cause additional adverse effects (Alford & Richards 1999;

Green et al. 2002; Blaustein et al. 2003; 2011; Green et al. 2002; Muths et al. 2006). Amphibian population fluctuations may be strong indicators of ecosystem function (Blaustein 1994).

Absorbent skin, unshelled eggs, and a complex life cycle often intimately linked to aquatic habitats make amphibians sensitive to environmental disturbances. They also play significant roles in the food web. Larval amphibians help to control algae and adults consume large quantities of insects that may serve as agricultural pests and disease vectors that can transmit illnesses to humans (Kupferberg 1997; DuRant & Hopkins 2008). Amphibians are also important food sources for a variety of predators (Duellman & Trueb 1986; Wells 2007). As amphibians

have persisted for over 250 million years contributing to ecosystem function, their dramatic declines are of critical conservation concern.

Among the major threats to amphibians are emerging infectious diseases (EIDs) that are diseases that have newly appeared in a population or that have been known for some time but are rapidly increasing in incidence or geographic range (Whittaker et al. 2013). EIDs are globally recognized threats to humans, wildlife, and ecosystems (Tompkins et al. 2015). There are several prominent pathogens and associated EIDs that affect amphibian populations. Batrachochytrium dendrobatidis (hereafter referred to as Bd) is a pathogenic fungus that causes chytridiomycosis

(Daszak et al. 1999a, 2000; Fisher et al. 2009b). A related yet highly divergent fungal pathogen that also causes chytridiomycosis, Batrachochytrium salamandrivorans (hereafter referred to as

Bsal), is a newly discovered pathogen infecting salamanders in northwestern Europe (Martel et al. 2013). Iridioviruses of the genus Ranavirus (hereafter referred to as Rv) have been implicated in declines and mass mortalities of amphibians worldwide (Green et al. 2002; Chinchar et al.

2009; Kik et al. 2011; Miaud et al. 2016). Amphibians are hosts to an assortment of pathogens, including bacteria, viruses, fungi, and helminths (Worthylake & Hovingh 1989; Blaustein 1994;

Cunningham et al. 1996; Daszak et al. 1999a; Johnson et al. 2002). However, we emphasize these three major EIDs given accumulating evidence of their potentially devastating effects on amphibian populations worldwide. In particular, we focus on reviewing the literature reporting the results of controlled experiments conducted with Bd, Bsal and Rv and their amphibian hosts, including laboratory, mesocosm, and field experimental studies. Given the complexity of these host-pathogen systems, experimental approaches have been crucial for disentangeling potential mechanisms driving patterns of transmission and examining variation in lethal and sublethal effects due to species, host life history traits, pathogen strain, and populations.

Since its discovery and first association with mass mortality events in Australian and

Central American frogs (Berger et al. 1998; Lips, 1998, 1999), Bd has been found in multiple host species on every continent where amphibians exist (Fisher et al. 2009b; Olson et al. 2013) and has been associated with numerous population declines and some extinctions (Berger et al.

1998; Lips, 1998; McCallum, 2005). However, the presence of Bd is not always linked to population declines (Daszak et al. 2005; Briggs et al. 2005; Vredenburg et al. 2010). Differences in susceptibility to Bd have been reported across host species (Blaustein et al. 2005; Searle et al.

2011b; Gahl et al. 2011; Gervasi et al. 2013a; Bielby et al. 2015), populations (Tobler & Schmidt

2010; Bradley et al. 2015), life stage (Briggs et al. 2010; Ortiz-Santaliestra et al. 2013), and Bd strain (Berger et al. 2005; Retallick and Miera 2007; Gahl et al. 2011; Doddington et al. 2013;

Gervasi et al. 2013b). Little is known about the newly discovered Bsal pathogen aside from its potential to cause mass mortalities and extinctions in salamander populations, as has been observed in Europe (Martel et al. 2013).

Unlike amphibian-specific Batrachochytrium pathogens, ranaviruses can infect a variety of taxa including reptiles and fishes (Gray et al. 2009a; North et al. 2015). Mass mortality events associated with Rv have had fatality rates as high as 90%, with reoccurring die-offs within the same populations (Green et al. 2002). The effects of Rv vary with host species (Schock et al.

2008; Hoverman et al. 2011; Brenes et al. 2014b), populations (Pearman & Garner 2005), viral isolate (Cunningham et al. 2003; Hoverman et al. 2010), type of exposure (Brunner et al. 2007;

Hoverman et al. 2010), and host life-stage (Schock et al. 2008; Haislip et al. 2011). Due to the global impact of chytridiomycosis and ranaviral disease, the World Organization on Animal

Health (OIE) named Bd and Rv as notifiable pathogens of the OIE Aquatic Animal Health Code

(Schloegel et al. 2009).

Summary of Pathogen Life Histories

Batrachochytrium dendrobatidis

First described by (Longcore et al. 1999), Batrachochytrium dendrobatidis (Bd) is one of three fungal species in the phylum Chytridiomycota that can infect a vertebrate host. Its complex life cycle consists of two critical stages; an infectious free-living aquatic zoospore stage and a non-motile zoosporangium stage. Motile zoospores are chemically attracted to keratinized epidermal cells of amphibian hosts. The zoospores encyst in keratinized larval jaw sheaths or in the keratinized epidermal layers of adult amphibian skin, including the heavily keratinized stratum corneum and stratum granulosum of the anuran pelvic patch and digits (Berger et al.

2005; Greenspan et al. 2012a). In as little as 12 hours, the cellular contents of the zoospore are transferred into deeper cell layers of the epidermis through a germination tube that extends from the zoosporangium (Greenspan et al. 2012a). Bd infection can lead to hyperkeratosis and hyperplasia of the dermal layer, erosions and ulcerations of the skin, disruption of the epidermal cell cycle, and molting (Berger et al. 1998, 2005a; Nichols et al. 2001; Voyles et al. 2009;

Greenspan et al. 2012a). The disruption of intercellular junctions, apoptosis, and necrosis of epidermal cells exposed to proteolytic secreted contents of Bd zoospores have also been observed (Brutyn et al. 2012). The inability to regulate ions through the skin may lead to cardiac arrest (Voyles et al. 2007). Clinical symptoms of chytridiomycosis include lethargy, lack of appetite, abnormal posture, loss of righting reflex, cutaneous erythema, and increased skin sloughing (Voyles et al. 2009). However, not all infected animals show disease symptoms when infected. Amphibian skin is a critical component in maintaining homeostasis, thus the disruption of cutaneous function is one mechanism causing mortality across amphibian species (Voyles et

al. 2009). Once infection occurs within the host, the zoosporangia mature and develop pathogenic zoospores that get released outside the host into the aquatic environment.

Little is known regarding the Bd life cycle outside the host. Bd grows maximally in laboratory conditions at temperatures between 17 and 25°C and a pH of 6-7. Temperatures above

30°C produced a 50% zoospore mortality rate (Piotrowski et al. 2004). Alternative dispersal mechanisms of Bd have yet to be comprehensively explored. Although atmospheric dispersal is thought to be uncommon, quantitative PCR analysis has detected Bd in rainwater (Kolby et al.

2015). Zoospores can survive for up to 7 weeks in lake water and 3-4 weeks in tap water and deionized water, respectively (Johnson & Speare 2003). Research have detected Bd on bromeliads in the forest canopy (Lindquist et al. 2011) and Bd has been found on tree leaves where infected frogs have recently rested (Kolby et al. 2015). There is the potential of Bd translocation by moist river sand and non-host carriers such as water fowl and crayfish (Johnson

& Speare 2005; Garmyn et al. 2012; Brannelly et al. 2015). Moreover, field studies reveal that despite its thermal optimums as suggested by laboratory studies (Piotrowski et al. 2004), Bd occurs in the environment in certain locations year-round (Chestnut et al. 2014). However, in many of these studies the viability of recovered zoospores has not yet been confirmed.

Batrachochytrium salamandrivorans

The recent isolation and characterization of the fungal pathogen, B. salamandrivorans

(Bsal) may explain some amphibian population declines. Notably, the drastic decline of fire salamanders, Salamandra salamandra, in the Netherlands, Germany, and Belgium, have sparked concern (Spitzen-Van der Sluijs et al. 2014; Sabino-Pinto et al. 2015; Spitzen-van der Sluijs et al.

2016). Martel et al. 2013 proposed that Bsal originated in East Asia and coexisted with

salamander host species for millions of years. The recent introduction of Bsal to Europe is hypothesized to have occurred due to a lack of biosecurity in the international pet trade (Martel et al. 2014). Although both Bd and Bsal infection result in lethal skin erosion, the pathogenic mechanism of Bsal are not yet understood. As with Bd, Bsal produces motile zoospores, contain colonial thalli, and produce germination tubes in vitro (Martel et al. 2013). Bsal possesses a lower thermal optimum compared to Bd, with optimal growth between 10°C and 15°C, and may be able to grow at temperatures as low as 5°C and as high as 25°C (Martel et al. 2013; Blooi et al. 2015a). Infection with Bsal can be alleviated via heat clearance at temperatures above its thermal maximum (Blooi et al. 2015a) and in combination with topical treatments (Blooi et al.

2015b). Due to this recent discovery, it is crucial to incorporate methods to monitor and screen populations for Bsal in worldwide field surveillance efforts. Studies have attempted to assess the presence of Bsal in various amphibian populations in North America (Muletz et al. 2014; Bales et al. 2015) and China (Zhu et al. 2014) utilizing several methods (phalanges histology, nested

PCR, qPCR and duplex qPCR), but its presence has yet to be confirmed in those populations.

Taking into account the abundance of salamander biodiversity in North America, the threat of

Bsal to salamanders in North America is of concern (Yap et al. 2015; Gray et al. 2015; Grant et al. 2016; Richgels et al. 2016). Increased field surveillance of these naïve populations will be critical to contain the potential effects of this newly isolated pathogen.

Ranavirus

Ranaviruses are a group of large double-stranded DNA viruses in the family Iridoviridae with fish, reptile, and amphibian hosts (Chinchar 2002). The first ranaviruses were isolated from

Lithobates pipiens in 1965 (Granoff et al. 1965). The genus Ranavirus is composed of 6

identified viral species, three of which infect amphibians (Ambystoma tigrinum virus (ATV),

Bohle iridovirus (BIV), and Frog Virus 3 (FV3) (Chinchar 2002). Although the effects of Rv are widespread, little is known about the genetic basis for virulence across isolates (Lesbarreres et al.

2012). FV3 and ATV infect many amphibian species, but these isolates are most virulent within the anurans and urodelans, respectively, from which they were isolated (Schock et al. 2008).

Laboratory experiments have shown that introduced Rv isolates may be significantly more virulent than endemic strains (Storfer et al. 2007).

Amphibians become infected with Rv by physical contact (e.g. bumping or fighting), dermal exposure to contaminated water, or by direct ingestion of virions, with the latter resulting in faster mortality rates (Harp & Petranka 2006; Brunner et al. 2007). Rv infection can occur in as little as a one second of direct contact with an infected individual of the same species

(Brunner et al. 2007) or 3 hours of contact with contaminated water (Robert et al. 2011). Vertical transmission of the virus is hypothesized as has occured in invertebrates (Bar Joseph et al. 2001) but empirical studies confirming its potential in amphibians are limited (Greer et al. 2005;

Brunner et al. 2007; Duffus et al. 2008; Gray et al. 2009a). Fish susceptibility to Rv is low, though there is potential for fish to transfer Rv to amphibians in habitats where fish and amphibians overlap (Brenes et al. 2014b; North et al. 2015).

Rvs are capable of inducing cell apoptosis and tissue necrosis within a few hours of infection (Chinchar 2002; Williams et al. 2005). Symptoms vary, but common indicators of Rv infection include erratic swimming, lethargy, erythema, skin sloughing, loss of pigmentation, lordosis (excessive inward curvature of the spine), and ulcerations (Tweedell & Granoff 1968;

Bollinger et al. 1999). Lesions and hemorrhages associated with fatal cases of Rv occur in internal organs, particularly the liver, kidney, intestine, spleen, and reproductive organs

(Cunningham et al. 1996; Docherty et al. 2003; Miller & Gray 2010). However, the precise mechanisms of Rv dissemination within the host are relatively unclear, especially at the earliest stages of infection. A recent study demonstrated that FV3 infection is capable of altering the blood brain barrier in Xenopus laevis tadpoles eventually leading to Rv dissemination into the central nervous system (De Jesús Andino et al. 2016). Death can occur even without external signs of infection (Chinchar 2002; Brunner et al. 2005).

The persistence of Rv in the environment is poorly understood but is significant to understanding indirect transmission dynamics. Nazir et al. (2012) observed that four isolates were resistant to desiccation. Persistence was highest in sterile pond water, followed by unsterile pond water and was lowest in soil. This is supported by Brunner et al.’s 2007 experiment, which demonstrated that desiccated then rehydrated soil was unable to cause Rv infection in larval salamanders. Evidence suggests that Rvs could survive up to 2 weeks in the environment and for long durations at low temperatures (Nazir et al. 2012). Although information on the thermal optimums of Rv are limited, for one strain, FV3, viral replication occured between 12°C and

32°C (Chinchar 2002). Rv occurrence is positively associated with increased urbanization (North et al. 2015) and the international trade has been widely implicated in the spread of Rvs

(Cunningham et al. 2003; Schloegel et al. 2009; Kolby 2014). Field studies and monitoring can add to our understanding of both temporal and spatial prevalence.

CHAPTER 2: THE IMPACTS OF EMERGING INFECTIOUS DISEASES AND OTHER CO-FACTORS ON AMPHIBIANS: A REVIEW OF EXPERIMENTAL STUDIES

Natalie M. Hambalek, Jenny Urbina, Emily Reynolds, Trang Dang, Jason T. Hoverman, Barbara Han, Deanna Olson, and Andrew Blaustein

ABSTRACT

The loss of biodiversity at genetic, species, and population levels is affecting ecosystems worldwide. Complex effects of multiple environmental stressors can act alone or together to drive population losses. Amphibians are among the most prominent groups experiencing population declines and extinctions. The causes of global amphibian population decline are multifaceted and context dependent. One major factor affecting amphibian populations is emerging infectious diseases. Several pathogens and their associated diseases are especially significant contributors to amphibian population declines. These include the chytrid fungus,

Batrachochytrium dendrobatidis, the related but highly divergent fungal pathogen, B. salamandrivorans, and ranaviruses. Here, we present a synthesis of these three emerging infectious diseases by assessing their broad effects on amphibian hosts as found in experimental studies. We also examine the interactive effects of these pathogens with other potential and known contributors of amphibian population declines that have been experimentally studied.

Well-designed experimental studies are critical for understanding the impacts of disease.

However, we note that inconsistencies in experimental methodologies often hinder our ability to form valuable comparisons and conclusions. Our review suggests the importance of implementing standard experimental protocols and reporting. Furthermore, we highlight the significance of investigating the roles of multiple environmental and anthropogenic stressors.

INTRODUCTION

Rapid rates of biodiversity loss over the last few centuries have supported the notion that

Earth is experiencing a sixth major extinction event (Dirzo et al. 2014; Ceballos et al. 2015).

Current species extinction rates are higher than pre-human background rates suggesting this 21st century crisis is largely attributed to anthropogenic change (Wilson 1992; Pimm et al. 1995;

Wake & Vredenburg 2008; Barnosky et al. 2011; Dirzo et al. 2014; Ceballos et al. 2015). At the forefront of this crisis, amphibians are a rapidly declining vertebrate group, (Stuart et al. 2004;

Alroy 2015), though not one factor alone bears total responsibility for their overall decline

Green et al. 2002; Blaustein et al. 2003; 2011; Muths et al. 2006). Amphibian population fluctuations may be strong indicators of ecosystem function (Blaustein 1994). Absorbent skin, unshelled eggs, and a complex life cycle often intimately linked to aquatic habitats make amphibians sensitive to environmental disturbances. For example, larval amphibians help to control algae whereas adults consume large quantities of insects that may serve as agricultural pests or disease vectors that can transmit illnesses to humans (Kupferberg 1997; DuRant &

Hopkins 2008). Amphibians are important food sources for a variety of predators (Duellman &

Trueb 1986; Wells 2007). As amphibians have persisted for over 250 million years contributing to ecosystem function, their dramatic declines are of critical conservation concern.

(Daszak et al. 1999a, 2000; Fisher et al. 2009b). A related yet highly divergent fungal pathogen that also causes chytridiomycosis, Batrachochytrium salamandrivorans (hereafter referred to as

2009; Kik et al. 2011; Miaud et al. 2016). Amphibians are hosts to an assortment of pathogens, including bacteria, viruses, fungi, and helminths (e.g. Worthylake & Hovingh 1989; Blaustein

1994; Cunningham et al. 1996; Daszak et al. 1999; Johnson et al. 2002). However, we emphasize these three major EIDs given accumulating evidence of their potentially devastating effects on amphibian populations worldwide. In particular, we focus on reviewing the literature reporting the results of controlled experiments conducted with Bd, Bsal and Rv and their amphibian hosts, including laboratory, mesocosm, and field experimental studies. Given the complexity of these host-pathogen systems, experimental approaches have been crucial for disentangling potential mechanisms driving patterns of transmission and examining variation in lethal and sublethal effects due to species, host life history traits, pathogen strain, and populations.

Since its discovery and first association with mass mortality events in Australian and

Central American frogs (Berger et al. 1998; Lips, 1998, 1999), Bd has been found in multiple

host species on every continent where amphibians exist (Fisher et al. 2009b; Olson et al. 2013) and has been associated with numerous population declines and some extinctions (Berger et al.

2005; Gahl et al. 2011; Searle et al. 2011b; Gervasi et al. 2013a) , populations (Tobler &

Schmidt 2010; Bradley et al. 2015), life-stage (Briggs et al. 2010; Ortiz-Santaliestra et al. 2013), and Bd strain (Berger et al. 2005; Retallick and Miera 2007; Gahl et al. 2011; Doddington et al.

2013; Gervasi et al. 2013b). Little is known about the newly discovered Bsal pathogen aside from its potential to cause mass mortalities and extinctions in salamander populations, as has been observed in Europe (Martel et al. 2013).

2008; Hoverman et al. 2011; Brenes et al. 2014b), populations (Pearman & Garner 2005), viral isolate (Cunningham et al. 2003; Hoverman et al. 2010), type of exposure (Brunner et al. 2007;

Hoverman et al. 2010), and host life-stage (Schock et al. 2008; Haislip et al. 2011). Due to the global impact of chytridiomycosis and ranaviral disease, the World Organization on Animal

Health (OIE) named Bd and Rv as notifiable pathogens of the OIE Aquatic Animal Health Code

(Schloegel et al. 2009).

We examine the impacts of the interactions between disease and other known drivers of amphibian population declines. Prior to 2009, relatively few studies of amphibian diseases employed standard ecological experimental designs (Blaustein et al. 2009) (Figure 2.1). Since

2009 there has been a surge in the use of experiments to determine how diseases affect amphibians. Experimental design, methods, and interpretation vary, thus it is useful to summarize these aspects to assess generality.

METHODS

The effects of Bd, Bsal, and Rv found in experimental studies are summarized in Table

2.1. Our search was conducted via Web of Science and supplemented with a Google Scholar search using the keywords “Batrachochytrium dendrobatidis + amphibians”, “Batrachochytrium salamandrivorans + amphibians”, and “Ranavirus + amphibians”, respectively. Duplicates and non-experimental studies were removed and the remaining studies were documented. Studies that examined interactive effects (ie. pesticide + pathogen) are included, but only the effect of the pathogen independently is reported. The Bd search resulted in 1435 hits, of which 90 were experimental studies, the Bsal search resulted in 17 hits, of which 4 were experimental studies, and the Rv search yielded 355 hits, of which 31 were experimental studies. If one publication examined multiple species or host life stages, each species and life stage was reported separately.

RESULTS

Results from experimental studies are summarized below. We present general trends across studies according to response (eg. physiology, behavior, etc.) and/or source of response variation (eg. life stage, strain used, etc.). We then focus on interactive effects and summarize the

experimental work with each pathogen in combination with natural or anthropogenic environmental stressors. Our objective was to provide a summary of patterns and gaps in the accumulated experimental work on host-pathogen dynamics of Bd, Bsal, and Rv and their amphibian hosts.

Batrachochytrium dendrobatidis

Experimental studies involving Bd suggest host-pathogen dynamics are influenced by many factors (Figure 2.2). Biotic factors include predator-prey interactions, host and pathogen density, competition, and whether more than one pathogen is present within a host (Parris et al.

2006; Rachowicz & Briggs 2007; Romansic et al. 2011; Groner et al. 2014). Lab and field experiments show that abiotic factors influencing Bd-host dynamics include climate, season, altitude, resource availability, and temperature (Woodhams et al. 2003; Berger et al. 2004; Andre et al. 2008). Experimental studies found dose-dependent differences in development, infection load, and mortality, indicating increased infection virulence associated with inoculum dose

(Garner et al. 2009; Bustamante et al. 2010; Romansic et al. 2011; Gervasi et al. 2013a; Bielby et al. 2015). Experiments have also confirmed temperature as a critical mediating factor in Bd dynamics. Frogs housed in warmer temperatures (22°C) exhibited significantly lower mortality than those housed in cooler temperatures (17°C) (Andre et al. 2008). Several experimental studies have shown that infection in post-metamorphic amphibians can be cleared when temperatures are elevated above the Bd thermal optimum range (Woodhams et al. 2003;

Weinstein 2009; Marquez et al. 2010; Garner et al. 2011; McMahon et al. 2014). While amphibians living near hot springs may clear infection naturally (Schlaepfer et al. 2002, 2005,

2007) adopting this treatment more widely for conservation management is logistically difficult,

especially if species do not select for warmer temperatures even when they are available (Han et al. 2008) .

Bd infection dynamics also can vary due to differences among strains. Experimental studies illustrate strain dependent infection outcomes (Berger et al. 2005a; Retallick & Miera

2007; Fisher et al. 2009b; Kilpatrick et al. 2010; Farrer et al. 2011; Gahl et al. 2012; Gervasi et al. 2013; Doddington et al. 2013; Piovia-Scott et al. 2015). Host survival, body condition at metamorphosis, and even lethal effects are strain dependent (Fisher 2009). Farrer et al. (2011) tested the effect of nine Bd strains on common toad (Bufo bufo) larvae and found that infection with global panzootic lineage (Bd-GPL) strains incurred greater mortality than other strains examined. When investigating Australian Bd strains, Berger et al. (2005) found differences in mortality rates in frogs, with the most recently isolated strain causing the highest mortality rates.

Retallick and Miera (2007) reported that although their tested Bd strains came from localities in close proximity to each other, infection resulted in drastically different host mortality rates. Gahl et al. (2013) further demonstrated that the combination of strain and host-species influences how

Bd affects hosts; four species (Lithobates catesbeianus, L. pipiens, Pseudacris crucifer, and

Ambystoma laterale) did not experience significant mortality from any strain, but two species

(Anaxyrus americanus and Lithobates sylvaticus) died from exposure to both strains, and Rana clamitans only died from infection with a Bd Panama strain (JEL 423). However, survival also may have been affected by factors such as host age at capture and exposure regimes. (Padgett-

Flohr & Hayes 2011) did not find survival differences when exposing Rana pretiosa to two Bd strains, but they showed that the rates of pathogen clearance were strain dependent. Furthermore,

Lithobates catesbeianus, an invasive species previously thought to have low susceptibility to Bd, can experience significant mortality to Bd infection. Gervasi et al. (2013) tested two Bd strains

(one isolated from L. catesbeianus in Oregon, JEL 630, and the second from Anaxyrus boreas in

Colorado, JEL 274) and showed mortality effects only with the Colorado Bd strain.

Alternatively, (Eskew et al. 2015) tested three Bd strains on L. catesbeianus, including one isolated from a population observed to be experiencing a Bd-induced decline, and found no impact on mortality. Other studies have revealed no effect associated with strain differences

(Padgett-Flohr & Hayes 2011; Brannelly et al. 2012).

Comparative strain experiments along with observational amphibian surveys are useful in investigating the relationships between host population trends and Bd virulence variation. For example, Piovia-Scott et al. (2015) and Doddington et al. (2013) linked an observed Rana cascadae population decline to a known intensely infectious and lethal Bd strain through multiple lines of analyses. In an experiment, adult R. cascadae exposed to the Bd strain cultured from the site undergoing a host population decline had significantly lower survival rates compared to uninfected controls (Piovia-Scott et al. 2015). This hypervirulent Bd strain also displayed greater immunotoxicity in experimental assays (Piovia-Scott et al. 2015). Exposure to endemic vs. novel strains also can affect host survival. Doddington et al. 2013 found survival differences in captive-bred Alytes muletensis experimentally exposed to two Bd strains, a local

Mallorcan strain (TF5a1) or a putatively hypervirulent Bd-GPL strain (UKTvB). Toads exposed to the Bd-GPL strain had higher mortality than individuals exposed to the Mallorcan strain or control group (Doddington et al. 2013).

Studies examining Bd strains independent from their host are important for understanding the differences in host-response. Differences in molecular profiles among Bd strains has been linked to in vivo functional virulence (Fisher et al. 2009b; Piovia-Scott et al. 2015). Several

studies have linked genomic and proteomic features to phenotypic traits thought to be involved in pathogenicity such as zoospore production and protease activity (Fisher et al. 2009; Piovia-

Scott et al. 2015). A hypervirulent Bd strain was found to contain genomic regions with higher copy numbers and increased loss of heterozygosity that correspond to putative virulence factors

(i.e. protease genes) (Piovia-Scott et al. 2015). One study found that Bd contains a large expansion of these protease genes compared to other closely related chytrids (Joneson et al.

2011). Other gene expansions of interest include a class of chitin-binding genes which are found in other pathogenic fungi and may serve as a critical component in the Bd cell wall (Abramyan

& Stajich 2012; Liu & Stajich 2015). Understanding the significance of results from different comparative strain experiments are difficult due to variation in methodology. Bd dosage, site of strain isolation, strain passaging history, and several more factors can influence strain experiment outcomes (Retallick & Miera 2007; Fisher et al. 2009b; Kilpatrick et al. 2010; Gahl et al. 2012;

Rosenblum et al. 2013; Langhammer et al. 2014; Voyles et al. 2014). Biases in choice of pathogen strain and host also influence our understanding of this complex disease system.

Accumulating evidence suggests that some host species can vary in their inherent susceptibility to Bd. Some species can persist with infection (Davidson et al. 2003; Daszak et al.

2003; Woodhams et al. 2007; Padgett-Flohr & Hayes 2011; Brannelly et al. 2012; Gervasi et al.

2013; Bielby et al. 2015; Eskew et al. 2015) and others experience mortality rapidly after Bd exposure (Blaustein et al. 2005; Carey et al. 2006; Searle et al. 2011b; Gahl et al. 2012).

Variation in skin composition, including keratin abundance, distribution, and thickness may affect the depth of which the zoospore-produced germination tube can affect the severity of infection among amphibian hosts (Reeder et al. 2012; Greenspan et al. 2012a). Differences in amphibian ability to mount sufficient endocrinological responses, particularly stress responses,

also may play a role (Warne et al. 2011; Gabor et al. 2013, 2015; Peterson et al. 2013; Searle et al. 2014). Furthermore, life history traits such as habitat preference may influence host susceptibility to infection (Rowley & Alford 2007; Bancroft et al. 2011). Habitat preference can influence pathogen exposure, as field studies have demonstrated higher prevalence of Bd infection in boreal toads relative to aquatic toads (Hossack et al. 2013)(Hossack et al. 2013).

Research should consider amphibian life history traits, particularly of species that do not seem to be susceptible to Bd infection, to better understand innate differences in host susceptibility and will be useful to target species which may act as reservoirs for the pathogen.

An important driver of host-pathogen interactions is host behavior (Parris et al. 2006;

Han et al. 2008; Venesky et al. 2011a). Basking, for example, may be an indication of disease infection in amphibians (Lefcort & Eiger 1993; Lefcort & Blaustein 1995; Richards-Zawacki

2010). Altered thermoregulatory behavior (i.e. behavioral fever), may aid in clearing Bd infection, however fever behavior depends on species and life stage (Schlaepfer et al. 2007; Han et al. 2008). Additionally, it has been suggested that aggregation behaviors can increase Bd prevalence, thus species displaying these behaviors may be more at risk thus schooling species may be at greater risk than solitary species (Venesky et al. 2011a). This prediction depends strongly on the assumption that infected hosts shed infectious zoospores, but some recent work shows that spillover infection does not occur in all hosts, suggesting that aspects of life history

(such as body size) and behavioral interactions (such as interspecific competition) between hosts drives infection severity in host communities (Han et al. 2015a). Infected tadpoles have demonstrated altered activity levels which may be an important indicator of anti-predator behavior (Parris et al. 2006; Han et al. 2011). While reduced activity can make tadpoles less visible and thus less at risk for predation, sluggish behavior can hinder an individual’s ability to

escape a predation event. Moreover, increased activity in combination with frequenting available refugia can minimize predation by making capture more difficult. Han et al. (2011) observed Bd- infected toad tadpoles seeking refuge more often than other species tested. Parris et al. (2006) demonstrated that when tadpoles were exposed to only visual predation cues, uninfected individuals positioned themselves farther from the predator than infected animals. Moreover, the ability to utilize avoidance behaviors may also be a reason behind interspecific variation. Carey et al. (2006) observed that post-metamorphic toads exposed to Bd were holding their bodies out of water more than unexposed individuals. In one study, frogs that have never been exposed to

Bd displayed no significant avoidance or attraction to the pathogen, whereas previously infected frogs associated with pathogen-free frogs a majority of the time (McMahon et al. 2014). This indication of potentially learned behavioral avoidance to Bd and other pathogens is an area that warrants further exploration.

Differences in Bd susceptibility are dependent on amphibian life stage, with juveniles and adults being more susceptible than embryos and larvae, most likely due to increased keratin distribution and abundance after the larval stage (Gervasi et al. 2013; Ortiz-Santaliestra et al.

2013). Bd infection in tadpoles rarely results in mortality (but see Blaustein et al. 2005; Fisher et al. 2009a; Gahl et al. 2012), but more often results in reduced foraging efficiency and food intake

(Parris & Beaudoin 2004; Venesky et al. 2009, 2010b; Hanlon et al. 2015). In adult amphibians,

Bd infection is manifested in the keratinized epidermis, thus the effects of foraging efficiency are dependent on the locality of infection. For example in adult salamanders (Plethodon cinereus),

Bd-infected individuals displayed increased feeding behaviors in comparison with uninfected individuals, a behavioral modification that has been suggested as a strategy to offset the costs associated with immune activation (Hess et al. 2015).

Observational field studies have helped build the foundation for experimental examination of Bd infection across and within host-life stages. For example, Smith et al. 2007 found that older tadpoles were more likely to be infected with Bd than younger ones. Both field studies and experimental laboratory studies have shown that newly metamorphosed individuals are at most risk for Bd infection, suffering higher mortality than other life-stage, though this is dependent on species (Kriger & Hero 2006; Ortiz-Santaliestra et al. 2013). Bd related mass mortality events have been documented in recently post-metamorphic frogs while sympatric larvae of the same species survive (Berger et al. 1998; Bosch et al. 2001). This may be attributed to changes that affect the amphibian immune system during developmentally critical window of metamorphosis. In response to these field observations, experimental testing showed that the amphibian immune system is down regulated before, during, and post-metamorphosis, resulting in greater susceptibility in late-stage tadpoles and newly metamorphosed individuals (Rollins-

Smith 1998; Rollins-Smith et al. 2011). Additionally, as tadpoles undergo metamorphosis, mouthparts are lost and the entire skin becomes keratinized, allowing the fungus to manifest over the entire body (Rachowicz & Vredenburg 2004a). Since the surface area of keratin is positively correlated with body size, experiments have shown that individual size also may be an influential factor in Bd susceptibility (Ortiz-Santaliestra et al. 2013). Garner et al. 2009 showed that smaller juvenile toads (Anaxyrus boreas) juveniles were more prone to Bd inflicted mortality compared with larger individuals.

Amphibian physiological responses to Bd have an important role in understanding the interspecific and intraspecific variation in susceptibility to chytridiomycosis. Changes to the immune system have been documented at both the genetic and cellular levels (Ramsey et al.

2010; Rollins-Smith et al. 2011; Gervasi et al. 2014a). Bd alters physiological and

immunological gene expression, including decreased expression of many immune and cytochrome p450 genes (Rosenblum et al. 2009; Ellison et al. 2014). At the cellular level, Bd infection alters lymphocyte proliferation (Fites et al. 2013; Gervasi et al. 2014a; McMahon et al.

2014; Young et al. 2014), though this could be dependent on experimental temperature

(Woodhams et al. 2007).

Experiments on amphibian-Bd interactions have also illuminated physiological stress responses to infection, a somewhat understudied area. In both field and laboratory investigations,

Bd significantly elevated physiological stress hormone (corticosterone) levels in amphibian hosts of multiple species (Gabor et al. 2013, 2015; Peterson et al. 2013; Searle et al. 2014), though exposure to endogenous corticosterone did not always alter amphibian susceptibility to Bd

(Searle et al. 2014). Different strains of Bd elicit significantly distinctive hormonal stress responses from their hosts, with more virulent strains resulting in higher corticosterone levels

(Gabor et al. 2015). New methodologies, such as a non-invasive stress hormone assay (Gabor et al. 2013), enhance the value of field studies coupled with experimental laboratory investigations on physiological stress response. The dynamics between stress response and chronic disease manifestation warrant further exploration.

Multiple experimental studies have found that Bd causes disruption of amphibian skin integrity and ion imbalance (Voyles et al. 2007; Marcum et al. 2010; Rosenblum et al. 2012;

Greenspan et al. 2012a). Additionally, infection with Bd can significantly alter the amphibian microbiome. Studies have confirmed that Bd infected frogs harbor significantly different bacterial communities (Jani & Briggs 2014; Belden et al. 2015). Thus, host microbiome composition may in part, explain interspecific variation in Bd susceptibility. Microbiota possess

antifungal properties that can compete with or eradicate the pathogen (Harris et al. 2006; Brucker et al. 2008; Becker & Harris 2010; Ramsey et al. 2010; Myers et al. 2012). This has been demonstrated in some studies, whereby frogs with experimentally reduced microbiota exposed to

Bd resulted in lower survival and mass gain than frogs in non-immunologically modified treatments (Woodhams et al. 2010; Pask et al. 2013). While amphibian microbiota has been emphasized in the Bd-host system, there are also implications of how the host-microbiota may play a role in other amphibian host-pathogen systems.

Batrachochytrium salamandrivorans

Due to its recent discovery, there are only four experimental studies documenting the effects of Bsal. However, 35 amphibian species have been studied (Table 2.1b). Amphibians differ in their sensitivity to Bsal, with Bsal primarily affecting newts and salamanders rather than anurans. The common midwife toad (Alytes obstetricans), a species highly susceptible to Bd, did not experience any clinical signs of Bsal infection (Martel et al. 2013). Further, Martel et al.

(2014) showed that all ten anuran (frog and toad) species tested were resistant to skin invasion, infection, and disease symptoms when exposed to a dose of 5,000 zoospores of the Bsal type strain, AMFP13/1. Experimental studies conducted with Bsal on potential urodelan (salamander) hosts demonstrated that responses varied across species and within the same genus. Bsal induced lethal effects on Lissotriton italicus, the Italian newt, whereas no infection or disease symptoms were documented in L. helveticus (Martel et al. 2014). The results of Bsal-host experiments also show that Bd and Bsal differ in chytridiomycosis disease symptoms. Experimentally infected fire salamanders, Salamandra salamandra, experienced ataxia, a less reported symptom in experimental studies with Bd. The study also identified three potential reservoir species, the

Japanese fire belly newt (Cynops pyrrhogaster), the Chuxiong fire-bellied newt (Hypselotriton cyanurus), and the Tam Dao salamander (Paramesotriton deloustali), as individuals of these species were able to persist with or clear Bsal infection in some capacity (Martel et al. 2014), but this area warrants further exploration.

Bsal transmission dynamics is not yet well documented. In a study examining transmission between infected and naïve hosts, Martel et al. 2013 found that two days of shared housing in salamanders resulted in infection and mortality of formerly naïve hosts within one month. All of the experimental work done regarding Bsal has used only one pathogen isolate, a small range of doses, and few source populations for each species tested. Experiments conducted on Bd-host dynamics show that responses are heavily dependent on species, population, pathogen isolate, temperature, and exposure dose. Therefore, the effects of Bsal on amphibians must continue to be comprehensively examined.

Ranavirus

Research on ranaviruses (Rv) has dramatically increased in the last decade (Figure 2.3).

Experimental studies have shed light onto the comprehensive effects of Rv on amphibians worldwide (Table 2.1c). Susceptibility to Rv varies greatly depending on species (Jancovich et al. 2001; Duffus et al. 2008; Schock et al. 2008; Hoverman et al. 2010), life-stage (Brunner et al.

2005; Haislip et al. 2011; Echaubard et al. 2014), transmission mode (Cullen & Owens 2002;

Pearman & Garner 2005; Brunner et al. 2007), viral dose (Pearman et al. 2004; Brunner et al.

2005; Warne et al. 2011; Duffus et al. 2014b), and viral isolate (Cunningham et al. 2007; Storfer et al. 2007; Schock et al. 2009; Bayley et al. 2013; Morrison et al. 2014; Echaubard et al. 2014).

Understanding the relative susceptibility of hosts to a pathogen is critical to predicting host- pathogen dynamics. Coevolution between Rvs and their hosts has been hypothesized to be a driving force behind host variation of susceptibility (Miller et al. 2011).

Experimental Rv mortality is influenced by a variety of factors, most notably, viral exposure method. Ingestion of Rv infected carcasses result in infection transmission and reduce survival (Pearman et al. 2004; Harp & Petranka 2006). Likewise, intraperitoneal injection of Rv may increase mortality (Cullen et al. 1995; Cunningham et al. 2007). Exposure to Rv via water induced variable rates of mortality, with most studies showing mortality occurring over slower rates (Brunner et al. 2005; Cunningham et al. 2007). Hoverman et al. 2010 found that infection and mortality rates were greater for tadpoles that were orally inoculated with Rv compared to those exposed via water bath.

It has been hypothesized that aggressive interactions can serve as an efficient transmission route of Rv (Brunner et al. 2007). Cannibalistic behavior may be harmful to the individual exemplifying the behavior because of disease transmission, but an experimental study showed cannibalism can result in decreased contact rates between naive and infected individuals in the population (Brunner et al. 2007). Additionally, experiments have suggested that necrophagy may serve as a common route of Rv transmission, shifting transmission from density-dependent to frequency-dependent (Jancovich et al. 1997, 2001; Pearman et al. 2004;

Harp and Petranka 2006; Brunner et al. 2007).

Experiments that examine disease dynamics by manipulating temperature conditions are useful in helping to explain Rv disease outbreaks in the field. Similarly to Bd, temperature influences Rv infectivity and survival rates in hosts (Rojas et al. 2005; Echaubard et al. 2014).

When exposed to the Rv, ATV, larval Ambystoma tigrinum salamanders experienced higher

survival rates when exposed at 26°C than those exposed at 18°C and 10°C with virus titer being higher in cooler temperatures (Rojas et al. 2005). The study also found viral replication rates were higher at higher temperatures. Thus, it appears that the immune system may have increased functionality at higher temperatures and may minimize Rv pathology. Echaubard et al. (2014) examined effects of temperature along with various Rv isolates and different host species. They also found that the probability of Rv infection increased at lower temperatures (14°C), but further, they showed effects were isolate and species dependent.

It is critical to take a comparative approach to experimentally investigate species variation in susceptibility with regards to Rv. Hoverman et al. 2012 discovered a wide range of lethal effects among 19 larval amphibian species, which resulted in mortality rates spanning from

0 to 100%. Their study showed that anurans in the family Ranidae were typically more susceptible to Rv than the other five families tested.

Previous experimental work has demonstrated infection and virulence variation among isolates and Rv species (Cunningham et al. 2007; Schock et al. 2008; 2009; Hoverman et al.

2012). Phenotypic variation among Rv isolates is not well understood. Schock et al. 2008 determined that FV3 and ATV Rv species vary in their ecology and restriction endonuclease profiles, even though they have identical major capsid protein (MCP) gene sequences. Their results further emphasize the importance of characterizing isolates beyond MCP sequence analysis. Cunningham et al. 2007 detected differences in tissue trophism and pathology between two trains of FV3-like Rvs in common frogs (Rana temporaria). Furthermore, Schock et al. 2009 revealed that ATV strains differed in virulence, but this was dependent upon the origin of the salamander host. This was built upon by Hoverman et al. 2012, who showed that infection by

FV3 and a ranaculture isolate were associated with species with evolutionary trends for breeding

in semi-permanent ponds. These results highlight the importance of controlled experimental studies to elucidate patterns of differential host susceptibility with regards to Rv isolates and species.

Field observations and experimental studies have shown that late-stage larvae that are nearing metamorphosis are the most susceptible to lethal effects of Rv infection (Duffus et al.

2008; Gray et al. 2009a; Warne et al. 2011; Andino et al. 2012; Brenes et al. 2014b). When experimentally exposed to ATV, Ambystoma tigrinum larvae that metamorphosed were five times less likely to be infected than those that remained at the larval stage (Brunner et al. 2005).

Experimental studies suggest that the effects of Rv are more lethal to larvae than any other host life stage. In an experimental study examining seven amphibian species at various developmental stages, Haislip et al. 2011 observed that mortality and infection prevalence were greatest at the hatchling and larval stages in four of the species tested compared with frogs undergoing metamorphosis, and that the embryo was the least susceptible stage, most likely due to the eggs protective membranous properties. Similarly to what has been observed with Bd infections, life stage variation in susceptibility has been attributed to changes that occur in the hypothalamic- pituitary-interrenal axis (the central stress response system) around the time of metamorphosis, which helps to mediate the immune system (Rollins-Smith 1998). Additionally, host gene expression variation may contribute to life-stage differences in susceptibility. Andino et al. 2012 found that larvae experienced greater infection rates and possessed lower and delayed expression of inflammation associated antiviral genes. It has been suggested that impacts of epizoonotic events may be underestimated due to increased difficulty of detecting mass mortality of hatchings and larvae in the field (Haislip et al. 2011).

Though few studies have examined host physiological responses to Rv exposure, these studies may be important in assessing species-specific impacts of Rv infection. Warne et al. 2011 demonstrated tadpoles infected with an FV3-like isolate had significantly higher whole body corticosterone relative to controls. In a study examining immune function, Maniero et al. 2006 demonstrated that Xenopus laevis frogs are able to develop an effective and persistent humoral immunity after exposure to FV3. Studies that examine Rv and host physiological endpoints such as stress and immune function are sparse and warrant further investigation.

Interactive Effects of Disease, Anthropogenic, and Natural Stressors

Anthropogenic and natural environmental stressors can exacerbate the effects of disease.

It has been hypothesized that an increase in the number and intensity of anthropogenic stressors in the environment are partially accountable for the emergence of wildlife diseases (Carey et al.

1999; Daszak et al. 2000). Though the impact of one factor may be particularly devastating to amphibians in certain regions, considering simultaneous effects of several factors may be more realistic because amphibians, like other organisms are exposed to many abiotic and biotic factors at the same time (Kiesecker et al. 2001; Blaustein et al. 2011). Host-pathogen relationships in amphibians are mediated by, for example, climate, contaminants, disease, predation, and competition (Fisher et al. 2009b; Blaustein et al. 2011; Rollins-Smith et al. 2011) (Figure 2.2).

These factors display a high degree of spatial and temporal variation and can result in complex local interactions that are poorly understood (Blaustein et al. 2011). Realistic insight can be gained by taking a population-specific approach in assessing the variables involved and overall status of a population using long-term field data (Blaustein 1994). Furthermore, experimental approaches can be particularly helpful in disentangling the mechanisms of interacting variables.

Gaining a comprehensive understanding of how environmental factors may influence infection and pathology is critical to amphibian conservation.

Pathogens and Climate Change

Climate change has the ability to alter disease dynamics by fostering conditions more or less hospitable for pathogens and their hosts. Many wildlife disease outbreaks are associated with warming climates. For example, climate-induced fluctuations in water depth are associated with increased ultraviolet-B (UV-B) radiation which results in greater sensitivity to the pathogenic water mold, Saprolegnia (Kiesecker et al. 2001). Information regarding the effects of UV-B radiation on Rv viability is absent from the literature. However, given that many viruses are UV- resistant (Jacquet & Bratbak 2003; Liltved et al. 2006; Eischeid 2009), Rv resistance to UVs could explain its documented environmental persistence. For example, decreased pond depth has been associated with increased Rv prevalence (North et al. 2015). No interaction has been found with increased UV-B radiation and Bd (Garcia et al. 2006; Searle et al. 2010), global climate change appears to increase temperature variability, which can mediate disease dynamics. In an observational field study Bosch et al. 2007 documented that rising temperatures are linked to the occurrence of chytridiomycosis. Since pathogen and its hosts are sensitive to temperature changes, fluctuating temperature regimes have had negative effects on survival and development in the presence of Bd (Hamilton et al. 2012; Rumschlag et al. 2014; Raffel et al. 2015), while higher temperatures often resulted in higher survival rates (Bustamante et al. 2010; Murphy et al.

2011). Raffel et al. 2015 examined varying climatic factors, including temperature variation and soil moisture on host thermal acclimation. They demonstrated that Bd growth and infection- induced mortality on newts, Notophthalmus viridescens, was greater following a shift to a new

cooler temperature, but this was dependent on increased soil moisture. Host thermal acclimation is context dependent and can serve as a key mediator of climate-disease dynamics.

Pathogens and Contaminants

Experiments are particularly important for understanding the impacts of contaminants on host-pathogen systems. Many contaminants break down quickly in the environment, yet exposure can have major carry over effects, and the effects of interactions between multiple contaminants cannot be well understood without experimentation (Relyea & Jones 2009; Relyea

& Edwards 2010). Contaminant exposure may contribute to amphibian population declines.

However, much research on the interactive effects of contaminants and pathogens remains inconclusive. Most studies examining this interaction investigate the hypothesis that pesticides and contaminants play a role in decreasing amphibian immune response, rendering amphibians more susceptible to infectious disease (Taylor et al. 1999; Christin et al. 2003; Gilbertson et al.

2003). However, few experimental studies support this hypothesis (Parris & Baud 2004;

Davidson et al. 2007; Buck et al. 2012, 2015; Paetow et al. 2012; Kleinhenz et al. 2012; Edge et al. 2013; McMahon et al. 2013b; Brown et al. 2013; Hanlon & Parris 2014; Wise et al. 2014).

Rohr et al. 2013 found that early-life exposure to atrazine decreased survival post- metamorphosis when combined with Bd in Osteopilus septentrionalis. Likewise, Buck et al.

2015 demonstrated that exposure to pesticides in tadpoles resulted in higher Bd loads and increased mortality in post-metamorphic Pseudacris regilla, P. crucifer, and Anaxyrus boreas individuals but not in others (Rana cascadae or Lithobates pipiens). A possible reason for these findings with little or no interactive effects may be that certain compounds can inhibit or

diminish the growth or integrity of Bd, as was demonstrated outside of the host species (Hanlon

& Parris 2012; McMahon et al. 2013b; Brown et al. 2013).

For Rv, the use of pesticides has been associated with increased Rv prevalence in the field (North et al. 2015) and may interact with the pathogen or ecosystem to pose additive risks to amphibians by negatively impacting the amphibian immune system. For example, Forson and

Storfer 2006a revealed that ecologically relevant levels of the pesticide atrazine and the fertilizer sodium nitrate significantly decreased Ambystoma tigrinum larvae peripheral leukocyte levels and that larvae exposed to atrazine significantly increased susceptibility to ATV. Furthermore,

Kerby and Storfer 2009 showed that atrazine and Rv exposure marginally decreased survival in larvae of the same species. Conversely, Forson and Storfer 2006b revealed that Ambystoma macrodactylum larvae exposed to both atrazine and ATV had lower levels of mortality and ATV infectivity compared to larvae exposed to ATV alone, suggesting atrazine may compromise virus integrity. Additional research is needed to assess the impacts of pesticides and fertilizers and their metabolites on Rv viability and amphibian physiology. Contaminants are becoming increasingly widespread with over 50% of detected insecticide concentrations exceeding regulatory thresholds (Stehle & Schulz 2015), thus, the importance of researching the interrelationships between contaminants and disease in amphibian disease should not be overlooked. Experiments designed to identify mechanisms that are generalizable across classes of pesticides will also enable better management and conservation planning as known contaminants are phased out and new ones are introduced to market.

Pathogens and Community Composition

Higher biodiversity may influence disease risk through a variety of mechanisms. The dilution effect hypothesizes that greater biodiversity in an assemblage decreases disease risk but this is somewhat controversial (Ostfeld & Keesing 2000; Lafferty 2012; Civitello et al. 2015)).

Thus, an increase in biodiversity (including predators) in an assemblage can increase the availability of less or non-competent hosts in a system leading to disease dilution. Olson et al.

2013 reported association of decreased Bd occurrence at a site with increased species richness.

Experimental evidence supports the dilution effect in the Bd-host system showing increases in tadpole diversity have decreased Bd zoospore abundance (Searle et al. 2011a; Johnson et al.

2013; Venesky et al. 2014; Han et al. 2015b). Searle et al. 2011a demonstrated that the addition of Rana cascadae tadpoles decreases infection risk for Anaxyrus boreas tadpoles. This in large part can be due to differing feeding strategies and life-history traits. Venesky et al. 2013 showed that some tadpoles can filter feed Bd zoospores and that the degree of this feeding was positively associated with pathogen dilution. Moreover, experiments have shown that zooplankton, such as

Daphnia, can consume Bd zoospores, significantly reducing infection probabilities in tadpoles

(Buck et al. 2011; Hamilton et al. 2012; Searle et al. 2013; Schmeller et al. 2014). Additionally, species “reservoirs” may be important for community level Bd dynamics. For example, evidence suggests the Pacific treefrog, Pseudacris regilla, may act as a Bd reservoir; P. regilla thrive and occupy 100% of study sites where a sympatric species has been extirpated by Bd (Reeder et al.

2012).

Predation in combination with disease infection can result in varying effects. The healthy herd hypothesis states that predators may decrease infection prevalence by decreasing overall population size of potential hosts (Lafferty 2004; Duffy et al. 2005). However, salamanders,

important predators of amphibian larvae are also susceptible to Bd. Several hypotheses regarding predator/prey dynamics and disease remain untested. For example, is selective predation occurring, or alternatively, are predators capable of avoiding infected prey? Han et al. 2011 experimentally demonstrated the potential of non-selective predation occurring in the predator/prey interactions in the Bd system. Salamander predators consumed Bd-infected and uninfected tadpoles at the same frequency, and predation risk among prey was not altered by Bd infection. This area warrants further exploration as predation behavior may have significant impacts on outcomes in amphibian disease systems.

Long-term environmental stressors can be detrimental to the immune system (Martin

2009), whereas short-term stress-induction, such as a predation event, can actually lead to immune-enhancement (Groner et al. 2014). For example, the presence of a predator resulted in decreased infection loads in wood frog (Lithobates sylvaticus) larvae (Groner & Relyea 2015) and have resulted in increased developmental rates (Groner et al. 2013; Brown et al. 2013).

Effects of predation in combination with Rv remain inconclusive. Dragonfly predator cues have resulted in decreased survival in combination with Rv exposure (Kerby et al. 2011). However,

Haislip et al. 2012 found no evidence that Rv exposure in combination with predator cues increased mortality in any of four larval anurans tested.

In addition to predator presence, other aspects of community composition can play an influential role in disease dynamics. When reared in higher densities amphibians metamorphose at smaller body masses than when reared individually (Wilbur 1977; Parris & Cornelius 2004).

Furthermore, when these higher densities were combined with the presence of Bd, larvae also experienced a delayed time to metamorphosis (Wilbur 1977; Parris & Cornelius 2004). Increased densities have also been associated with the increased likelihood of Bd infection (Bielby et al.

2015), but other experimental studies have not observed this association (Searle et al. 2011b).

These results are in direct contrast with the effects of density with regards to Rv. In higher densities of larvae and in the presence of Rv, the rate of metamorphosis was documented to be

3x faster and the probability of mortality was 5x lower than in the controls (Reeve et al. 2013).

Coinfection Dynamics

Infection by multiple pathogens is common for most wild animals (Ezenwa & Jolles

2011), though experimental evidence of coinfection patterns in amphibians remain sparse.

Several studies have investigated coinfection dynamics between pathogens and their amphibian hosts in the field. Whitfield et al. 2013 revealed widespread infection by both Bd and ranavirus in wild frogs in Costa Rican lowland wet forests and verified that complex patterns of pathogen occurrence may play a significant role in interspecific infection dynamics. They identified a significant positive association between Bd and Rv infection in one species, Craugastor fitzingeri. Hoverman et al. 2012 documented that Bd, Rv, and trematode infections are widespread in the California San Francisco Bay Area, with over 68% of sampled wetlands harboring at least two of the three pathogens. Shock et al. 2010 investigated over thirty Canadian wetlands for Bd and Rv and revealed only a single site where the pathogens cooccurred.

Evidence of coinfection between Bd and Rv have been shown to exist in the field and in captivity (Kik et al. 2012; Whitfield et al. 2013; Warne et al. 2016) but the results from experimental investigations are limited. Romansic et al. 2011 experimentally investigated the effects of three pathogens: Bd, the trematode Ribeiroia sp., and the water mold, Achyla flagellata, which resulted in little evidence for interactive effects. Thus the interrelationships of coinfection must be further explored in experimental studies and international field monitoring.

DISCUSSION

Host, isolate, and geographic biases

Uneven sampling of hosts is considered to be a source of bias when interpreting the dynamics of host-parasite systems (Kuris and Blaustein 1977). There are 7,479 amphibian species described (ASA 2016), yet our analysis of experimental studies documenting the effects of these pathogens have only reported effects for <1% of species across these pathogens (008% of species with regard to Bd, .004% of species for Bsal, and 0.005% of species with regard to

Rv). Of the species studied in these disease systems, there is a high degree of interspecific variation in disease susceptibility (Blaustein et al. 2005; Searle et al. 2011b; Gahl et al. 2012;

Hoverman et al. 2012a; Gervasi et al. 2013). Furthermore, responses can vary based on strain, population, and host life-stage (Pearman et al. 2004; Rachowicz & Vredenburg 2004a; Pearman

& Garner 2005; Brunner et al. 2005; Blaustein et al. 2005; Retallick & Miera 2007; Schock et al.

2008, 2009; Bradley et al. 2015). Additionally, a distinct disparity exists in species studied and geographic regions (Figure 2.4). The majority of research has focused primarily on host species located in Europe, North America, and Australia. However, we know Bd and Rv have global distribution and effects, yet far less is known about infection in hosts from Africa, Asia, and

South America. Bsal has only documented effects based on one type strain isolated from Europe, and most experimental studies have used a dose of 5,000 zoospores, a low dose in comparison to studies on Bd (Gervasi et al. 2013). Similarly, the bulk of the studies examining Rv pathogen- host dynamics are largely biased in North America, with a minority of studies coming out of

Europe, Africa, and Australia. No doubt, these biases are inherently based on number of researchers in these regions, institution locality, access to collaborators, species, isolates,

feasibility and cost. We must attempt to close the gaps behind variation in susceptibility of populations most vulnerable to the effects of these pathogens can be targeted so effects are mitigated. In order to accomplish this, we must have standardized methods and reporting in our studies.

Non-standard methods and reporting

Experimentation is advantageous because it is repeatable and well-designed studies can examine problems and portray convincing results (Hairston 1989; Underwood 1997). However, there are numerous limitations to experimental work, as is illustrated in amphibian disease ecology. One problem with experimental work on amphibian diseases has been the lack of standardization in experimental methods. Kilpatrick et al. 2010 highlighted the importance of standardizing and reporting all relevant infection protocols within and between species when conducting laboratory studies regarding Bd and its host species. This includes how individuals are collected for experiments, how they are reared, the developmental stage in which they are tested, the population origin, inoculation and exposure protocols, and strains of pathogen being used. For instance, reporting and standardizing the zoospore exposure concentration (total number of zoospores per mL of water in total volume of water) in experimental procedures would make relative species comparisons among experiments more useful. Developmental stage

(i.e. (Gosner 1960) stage) should always be reported as this can also confound the interpretation of results. Additionally, whether hosts are reared from eggs or caught as larvae, juveniles, or adults, or even bought from supply houses can dramatically alter the results of experiments and their interpretation. Our analysis shows that 31%, 13%, and 22% of experiments examining Bd,

Bsal, and Rv, respectively were using animals not reared from eggs. Even when tested for

current infection prior to the experiment, wild-caught individuals have different ecological histories and may have a more or less robust immune system depending upon whether they were previously exposed to a particular pathogen (Gahl et al. 2012). Field surveillance shows that amphibian parasites such as echinostomes are widespread (Johnson & Sutherland 2003;

Hoverman et al. 2012b) and essentially many, if not all individuals, collected from the wild will inevitably possess trematodes. The potential influence of these parasites on amphibian immunological response pose a serious problem for experiments that choose to use individuals not reared as eggs. We emphasize the importance of utilizing subjects raised from the embryo stage in experimental investigations. Because of lack of standardization, each experiment must be taken at face value and applied to those specific individuals at the reported experimental conditions, so interpretation should be reported cautiously. When protocols are standardized we can more easily generalize effects of Bd and Rv on hosts as has been accomplished in several studies (Blaustein et al. 2005; Hoverman et al. 2011; Searle et al. 2011b; Gervasi et al. 2013).

Under controlled environmental conditions, observed effects after pathogen exposure can be attributed to intrinsic biological factors of the host, rather than environmental differences

(Bradley et al. 2015).

Figure 2.1 Trends in amphibian disease ecology experimental studies over time. Bd = Batrachochytrium dendrobatidis, Bsal = Batrachochytrium salamandrivorans, Rv = ranavirus.

Figure 2.2 Abiotic and biotic factors that contribute to host-pathogen dynamics in amphibian disease systems.

Figure 2.3 Trends in all articles published on Batrachochytrium dendrobatidis (A) and ranavirus (B) in the primary literature over time. Publications were compiled using the search strings Batrachochytrium dendrobatidis and amphibians (A) and ranavirus and amphibians (B) in the Web of Science database from which duplicates and articles that were unrelated and from non-primary literature were removed. The Bd search yielded a total of 1,435 hits and the Rv search yielded 355 hits.

Figure 2.4 Experimental studies published on Batrachochytrium dendrobatidis and ranavirus with respect to amphibian host genus and geographic range. Methods to generate number of studies were produced in the same fashion as Table 2.1. N indicates the number of studies for that particular region.

Table 2.1 An overview of the effects of Bd (a), Bsal (b), and Rv (c) on amphibian species based on studies that experimentally infected or observed effects of the pathogen. If one publication examined multiple species or host life stages, each species and life stage was reported separately. We have included each species International Union for Conservation of Nature (IUCN) Red List Status (http://www.iucnredlist.org), a widely recognized mechanism for assessing conservation status. Categories reported in this table include species of Least Concern (LC), Near Threatened (NT), Vulnerable (VU), Endangered (EN), and Critically Endangered (CR).

Table 2.1a: Effects of Batrachochytrium dendrobatidis on amphibian hosts

Species IUCN Bd Strain Bd Dose Life Stage Effect on host Reference Status (total zoospores in mL water, if specified)

Alytes muletensis VU UKTvB, 2,3000 Through Strain differences in infection (Doddington et al. 2013) (Majorcan midwife toad) TF5al metamorphosis

Alytes obstetricans LC n/a n/a Through Population differences in survival (Tobler & Schmidt (Common midwife toad) metamorphosis 2010)** Ambystoma VU JEL 270 1,000, 100,000 Juveniles No significant differences in survival or (Padgett-Flohr 2008)** californiense mass (California tiger salamander)

Ambystoma laterale LC JEL 423, 10^6-10^7 and Juveniles No significant differences in survival (Gahl et al. 2012) (Marbled salamander) JEL 404 10^5-10^6 zoosporangia Ambystoma opacum LC 277 250,000 Larvae No infection detected, no significant (Venesky et al. 2010a) (Blue-spotted differences in survival salamander) Ambystoma tigrinum LC A-277, R- 9,000,000, Juveniles No significant differences in survival (Davidson et al. 2003) (Tiger salamander) 230 6,000,000

Bd-GPL 10,000, Juveniles No differences in zoospore outputs (Peterson & McKenzie isolate 200,000 2014)** Anaxyrus americanus LC JEL 197 500,000 Juveniles Age dependent effect of Bd (Ortiz-Santaliestra et al. (American toad) susceptibility 2013) JEL 423, 10^6-10^7 Larvae Reduced survival (Gahl et al. 2012) JEL 404 zoospores and 10^5-10^6 zoosporangia

JEL 213 2.10x10^6 Juveniles Reduced survival (Wise et al. 2014) Anaxyrus boreas NT JEL 215 12,600 Larvae Reduced survival (Blaustein et al. 2005) (Western toad) JEL 274 2.4 x 10^4 Larvae Higher stress hormones, increased (Searle et al. 2014) length JEL 274 2 culture dishes Larvae Did not avoid infected conspecifics, (Han et al. 2008) inoculated in increased activity, no differences in batches with 20 temperature selection larvae JEL 215 6.18 x 10^6/mL Larvae Decreased activity, no difference in (Han et al. 2011) refuge use JEL 274 100,000, 50,000, Larvae No significant differences in survival (Gervasi et al. 2013a 10,00 JEL 274 100,000, 50,000, Juveniles No significant differences in survival (Gervasi et al. 2013a) 10,00 JEL 215 2.08 × 10^7 / plate Juveniles Reduced survival (Garcia et al. 2006)* JEL 275 10^6 daily Juveniles Mass dependent survival time, exposed (Carey et al. 2006) individuals held bodies out of water as much as possible JEL 275 5.8x10^5/mL Adults Reduced survival (Murphy et al. 2011) Anaxyrus boreas boreas JEL 275 100,000 Adults Electrolyte alterations, lymphocytic (Marcum et al. 2010)*** infiltration Anaxyrus fowleri LC n/a n/a Larvae Reduced foraging efficiency (Venesky et al. 2009) (Fowler’s toad) FMB 001 6,000,000 Larvae Negatively impacts growth (Venesky et al. 2011) USA isolate 6,000,000 Larvae Reduced foraging efficiency (Venesky et al. 2010b) 284 Anaxyrus terrestris LC JEL 274 2.6 × 10^5 Juveniles Reduced survival, decreased feeding (Searle et al. 2011b) (Southern toad) Anaxyrus woodhousii LC Bd-GPL 10,000, 200,000 Juveniles No significant differences in zoospore (Peterson & McKenzie (Woodhouse’s toad) isolate outputs 2014)** Atelopus zeteki CR JEL 423 30,000 Adults Infection intensity and zoospore output (DiRenzo et al. 2014) (Panamanian golden were positively correlated frog) JEL 423 30,000 Adults Significant differences in expression of (Ellison et al. 2014)** numerous genes involved in innate and inflammatory responses JEL 408 100, 10^4, 10^6 Adults Dose and temperature dependent (Bustamante et al. 2010) effects

Batrachoseps attenuates LC n/a 3x10^9 Adults Cleared infection, wild caught infected (Weinstein 2009)** (California slender individuals experienced 100% mortality salamander) in the laboratory Bufo bufo LC IA042, 3,000-17,000 Larvae Strain differences in mortality and (Farrer et al. 2011) (Common toad) IA043, 0711, infection dynamics VAo2, VAo4, VAo5, CCB1, TF5a, TF1.1 UKTvB, 190, 190,00 Larvae Reduced survival, differences in mass, (Fisher et al. 2009a) TF5a1, strain differences in virulence and IA042 infection IA-42 160, 16,000 Juveniles Reduced survival, mass dependent (Bielby et al. 2015) effects IA2004 043 30 - 70, 3000 - Through Dose, size, and age dependent effects (Garner et al. 2009) 15,000 metamorphosis (Garner et al. 2009) IA2004 043 30 - 70, 3000 - Juveniles Increased mortality rates, dose, and 15,000 size dependent effects

n/a 120-300, 12,000 - Juveniles Warmer overwintering regime increases (Garner et al. 2011) 30,000 the probability of infection. Proliferation of Bd in the host was better in toadlets that experienced a colder winter Bufo quercicus LC SRS 812 60,000 Adults Learned behavioral avoidance to Bd (McMahon et al. 2014) (Oak toad) Dendropsophus EN BdLEcat10C 9x10^6 Juveniles Reduced survival (Villarroel et al. 2013)** meridensis G-1 (Merida tree frog) Dendrobates auratus LC n/a n/a Juveniles Reduced survival (Nichols et al. 2001) (Green and black poison dart frog) Dendrobates tinctorius LC n/a n/a Juveniles Reduced survival, skin lesions (Nichols et al. 2001) (Dyeing dart frog) Desmognathus LC JEL 197 1.068x10^7 Adults Reduced survival (Vazquez et al. 2009)** monticola (Seal salamander) Desmognathus orestes LC BD 197 1,000,000 Adults No clinical signs of infection (Chinnadurai et al. (Blue Ridge dusky 2009)** salamander) Eleutherodactylus coqui LC JEL 427 50,000, 100,000 Juveniles Reduced survival, population (Langhammer et al. (Common coquí) differences 2014)** 42

JEL 427 10^6,10^5/mL in Adults No significant differences in survival, (Langhammer et al. 10 mL cleared or reduced infection 2014)** Hyla chrysoscelis LC n/a 7000/mL Through No significant differences in survival, (Parris & Baud 2004) (Cope's gray tree frog) metamorphosis reduced metamorphic body mass, delayed time to metamorphosis JEL 646, 8x10^3 Through No significant differences in survival, (Gaietto et al. 2014) JEL 423, metamorphosis growth, or time to metamorphosis JEL 213, JEL 660, FMB 003, JEL 404 n/a 125,000 Larvae Reduced foraging efficiency (Venesky et al. 2009) n/a 6,000,000 Larvae Reduced foraging efficiency (Venesky et al. 2010b) Hyla cinerea LC JEL 423, 76.7 x 10^6, 4.7 Juveniles and No clinical signs of infection. Infection (Brannelly et al. 2012)** (American green tree SRS810 x10^6 Adults did not negatively affect body condition frog) or growth rate for either strain or life- stage Hyla versicolor LC JEL 274 2.6 × 10^5 Juveniles Reduced survival (Searle et al. 2011b) (Gray tree frog) FMB 003 75,000 Larvae Reduced survival, age dependent (Hanlon & Parris 2012) effects FMB 001 6,000,000 Larvae Negatively impacts growth (Venesky et al. 2012) Hypsiboas crepitans LC Bd1006 9,000,000 Juveniles Cleared infection (Marquez et al. 2010)** (Emerald-eyed tree frog) Leiopelma archeyi CR JEL 197 250,000 Adults Cleared infection (Shaw et al. 2010)** (Archey's frog) Limnodynastes peronei LC Gibbo River- 20x10^6 Larvae and Reduced survival, infection loads (Stockwell et al. 2010) (Striped marsh frog) Llesueuri- Juveniles increased over time 00-LB-1 Limnodynastes LC GibboRiver- 5,000 + 2mL Juveniles No significant differences in survival (Woodhams et al. 2007) tasmaniensis Llesueuri- (Spotted grass frog) 00-LB-1

Lithobates blairi/ n/a n/a 7,000/mL Larvae No significant differences in survival, (Parris 2004) Lithobates reduced metamorphic body mass sphenocephala (Plains leopard frog/Southern leopard frog)

Lissotriton helveticus LC n/a ~2,000 Adults Decreased mass, no evidence of (Cheatsazan et al. (Palmate newt) hastened secondary sexual trait 2013)** regression, exposure associated with a 50% earlier initiation of the terrestrial phase Lithobates catesbeianus LC JEL 274 48,000 Larvae Higher stress hormones and increased (Searle et al. 2014) (American bullfrog) length JEL 215 8400 Larvae No significant differences in survival (Blaustein et al. 2005) JEL 274, 1.7 x 10^4/ml in 15 Juveniles Strain differences in infection (Gervasi et al. 2013b) JEL 630 mL JEL 423 8x10^7 to 2x10^8 Juveniles Disruption of the epidermal cell (Greenspan et al. maturation cycle 2012a)** JEL 423, 10^6-10^7 Juveniles No significant differences in survival (Gahl et al. 2012) JEL 404 zoospores and 10^5-10^6 zoosporangia Bd-GPL 10,000 or 200,000 Juveniles Produces more infective zoospore (Peterson & McKenzie isolate stage than other species tested 2014)** Crater 10^6 and 2x10^6 Juveniles No significant differences in survival, (Eskew et al. 2015)** Meadow low infection prevalence, strain isolate, differences Finley Lake isolate Lithobates clamitans LC JEL 423, 10^6-10^7 Juveniles Strain differences in infection (Gahl et al. 2012) (Green frog) JEL 404 zoospores and 10^5-10^6 zoosporangia

Lithobates pipiens LC n/a 2,800,000 Larvae Reduced activity (Parris et al. 2006) (Northern leopard frog) JEL 275 10^4 Juveniles Reduced survival (Pask et al. 2013)**

JEL 274 2.6 x 10^5 Juveniles Reduced survival (Searle et al. 2011b)

JEL 423, 10^6-10^7 Juveniles No significant differences in survival (Gahl et al. 2012) JEL 404 zoospores and 10^5-10^6 zoosporangia

JEL 423 3.98x10^6 Juveniles Increased skin shedding, no significant (Paetow et al. 2012)** differences in survival or immunological endpoints tested JEL 197 500,000 Juveniles No significant differences in survival (Ortiz-Santaliestra et al. regardless of age 2011) JEL 423 1.69x10^7 - 7.43 x Adults Lower peak jumping velocity in infected (Chatfield et al. 2013)** 10^8 subjects, testes width significantly greater in infected individuals

Lithobates LC n/a 2.88 x 10^6 Larvae No significant differences in survival, (Hanlon et al. 2015) sphenocephalus reduced foraging efficiency (Southern leopard frog) n/a 400,000 Larvae Low protein diets resulted in smaller (Venesky et al. 2012) and less developed Larvae and reduced immune responses, high dietary protein significantly increased resistance to Bd JEL 197 10^6 Juveniles Increased pathogen skin burden within (Holden et al. 2015) two weeks of exposure, higher pathogen burden in deceased frogs, decrease in pathogen loads over time Lithobates sylvaticus LC JEL 404, 10^6-10^7 Larvae Reduced survival, no differences in (Gahl et al. 2012) (Wood frog) JEL 423 zoospores and growth or time to metamorphosis 10^5-10^6 zoosporangia JEL 404, 10^6-10^7 Larvae Reduced survival (Gahl et al. 2012) JEL 423 zoospores and 10^5-10^6 zoosporangia JEL 197 10^4 Juveniles No significant differences in survival (Ortiz-Santaliestra et al. regardless of age 2013) JEL 274 2.6 × 10^5 Juveniles Reduced survival (Searle et al. 2011b) JEL 274 1.55 x 10^5 Juveniles Population differences in survival (Bradley et al. 2015) JEL 423 1 x 10^7 to 2 x Juveniles Disruption of the epidermal cell (Greenspan et al. 10^7 maturation cycle 2012a)**

Lithobates yavapaiensis LC A-277, R- 8.5 x 10^3/mL Juveniles No significant differences in survival (Davidson et al. 2003) (Lowland leopard frog) 230

Litoria aurea VU Gibbo River- 20x10^6 Larvae No significant differences in survival, (Stockwell et al. 2010) (Green and golden bell Llesueuri- Juveniles decrease in pathogen loads over time frog) 00-LB-1

Litoria booroolongensis CR Abercrombie 750,000 in 5mL Juveniles No evidence that prior Bd infection (Cashins et al. 2013) (Booroolong frog) NP- increases protective immunity L.booroolon gensis-09- LB-P7

LC GibboRiver- 5,000 in 2mL Juveniles Reduced survival (Woodhams et al. 2007) Litoria caerulea Llesueuri- (Australian green tree 00-LB-1 frog)

98 1469/10, 50,000 Juveniles Differences in survival rates among (Berger et al. 2005b) 99 1385/1, infected groups 00 545 n/a n/a Adults Decreased blood pH, low plasma (Voyles et al. 2007) osmolality and reduced concentrations of sodium, potassium, chloride and magnesium EPS4 250,000 Adults Increased skin sloughing rate with (Ohmer et al. 2014)** increased infection intensity Gibboriver- 93x10^4/mL-1 Adults No significant differences in survival or (Voyles et al. 2014)** L.lesueuri- mass 00-LB-1P50 and P10 n/a 250,000 Adults Impaired immune response (Young et al. 2014)** n/a n/a Adults Impaired stress and immune response, (Peterson et al. 2013)* increased skin shedding Litoria chloris LC GibboRiver- 5,000 in 2mL Juveniles Reduced survival (Woodhams et al. 2007) (Red-eyed green Llesueuri- treefrog) 00-LB-1 GibboRiver- 15,000 in 2mL Juveniles Temperature did not influence leukocyte (Woodhams et al. 2007) Llesueuri- populations 00-LB-1 n/a 15,000 Juveniles Temperature dependent effects on (Woodhams et al. survival 2003)*** Litoria infrafrenata LC n/a 250,000 Adults Reduction in white blood cells and (Young et al. 2014)** (White-lipped tree frog) serum globulin concentrations

Litoria raniformis EN n/a 100,000 Adults Compromised ability to osmoregulate (Carver et al. 2010)** (Growling grass frog) and rehydrate, no significant difference in metabolic or breathing rates Mixophyes fasciolatus LC GibboRiver- 5,000 in 2mL Juveniles Reduced survival (Woodhams et al. 2007) (Great barred frog) Llesueuri- 00-LB-1 No. 00/545 1,000 Adults Lower temperatures enhanced (Berger et al. 2004)* pathogenicity Osteopilus LC SRS 812 3 x 10 ^4/mL in Larvae The loss of keratin in the mouthparts (McMahon & Rohr 2015) septentrionalis 2mL associated with a loss of Bd (Cuban tree frog) SRS 812 3mL of 6x10^4 Larvae Reduced survival (McMahon et al. 2013a) (after each water change) SRS 812 3 x 10^6/mL Juveniles Pathogen loads decreased over time, (McMahon et al. 2014) increased lymphocyte proliferation with increased exposures, previous exposure increased chances of survival Pelophylax esculentus LC TG 739 1.5-2 x 10^5 Adults Reduction in skin peptide and (Woodhams et al. 2012)** (Edible frog) microbiota immune defenses caused less weight gain and increased infection rates Pelophylax lessonae LC TG 739 1.5-2 x 10^5 Adults Reduction in skin peptide and (Woodhams et al. 2012)** (Pool frog) microbiota immune defenses caused less weight gain and increased infection rates Plethodon cinereus LC JEL 660/JS 7 x 10^5 in 5mL Adults Increased feeding activity (Hess et al. 2015)* (Red-backed OH-1 salamander) Plethodon glutinosus LC BD 197 10,000, Adults Clinical symptoms of infection (Chinnadurai et al. (Northern slimy 100,000, 2009)** salamander) 1,000,000 Adults No significant differences in survival (Chinnadurai et al. 2009)** Plethodon metcalfi LC JEL 197 1.068 x 10^7 Adults Reduced survival (Vazquez et al. 2009)** (Southern gray-cheeked salamander) Pseudacris crucifer LC JEL 423, 10^6-10^7 Adults No significant differences in survival (Gahl et al. 2012) (Spring peeper) JEL 404 zoospores and 10^5-10^6 zoosporangia Pseudacris feriarum LC JEL 274 2.6 × 10^5 Juveniles Reduced survival (Searle et al. 2011b) 47

(Upland chorus frog) Pseudacris regilla LC JEL 215 12,600 Larvae No significant differences in survival (Blaustein et al. 2005) (Pacific tree frog) JEL 626 27,800 Larvae Reduced survival and activity, delayed (Kleinhenz et al. 2012) time to metamorphosis JEL 215 2 culture dishes Larvae No differences in temperature selection (Han et al. 2008) inoculated in batches with 20 larvae JEL 215 6.18 x 10^6/mL Larvae No significant differences in activity or (Han et al. 2011) refuge use

JEL 274 10,00, 50,000, Larvae No significant differences in survival, (Gervasi et al. 2013a) 100,000 dose-dependent infection loads JEL 274 10,00, 50,000, Juveniles Reduced survival, dose-dependent (Gervasi et al. 2013a) 100,000 infection loads JEL 215 2.08 × 10^7 Juveniles No significant differences in survival (Garcia et al. 2006)* JEL 274 50,000 Juveniles Reduced survival, Infection load (Gervasi et al. 2014) increased over time, lower lymphocyte levels JEL 274 2.6x10^7 and Through Dose-dependent effects, accelerated (Romansic et al. 2011) 1.1x10^6/L metamorphosis development Pseudacris triseriata LC Bd-GPL 10,000 and Juveniles No differences in zoospore outputs (Peterson & McKenzie (Western chorus frog) 200,000 2014)

27-mile lake 8 x 10^4 Adults Strain differences in infection (Retallick & Miera 2007) isolate, Lost lake isolate

Rana aurora LC JEL 215 2 culture dishes Larvae No differences in temperature selection (Han et al. 2008) (Northern red-legged inoculated in frog) batches with 20 larvae n/a 2x10^5 added Larvae High temperature variability in the (Hamilton et al. 2012) every other day for presence of Bd had decreased growth 8 days JEL 215 6.18 x 10^6/mL Larvae No significant differences in activity or (Han et al. 2011) refuge use

Rana boylii NT LJR 119 9.4x10^6 in 50mL Juveniles No significant differences in survival, (Davidson et al. 2007)* (Foothill yellow-legged reduced growth, increased skin peptide frog) concentrations A-227, R- 127,500, Juveniles No significant differences in survival (Davidson et al. 2003) 230 1,275,000 Rana cascadae LC JEL 215 12,600 Larvae No significant differences in survival, (Blaustein et al. 2005) (Cascades frog) increased incidence of mouthpart abnormalities JEL 274 48,000 Larvae Higher stress hormones and increased (Searle et al. 2014) length and mass JEL 274 4 culture dishes Larvae Non-infected individuals were observed (Han et al. 2008) inoculated in more frequently on Bd+ side of test batches with 90 chamber larvae JEL 274 10,00, 50,000, Larvae No significant differences in survival (Gervasi et al. 2013a) 100,000

JEL 215 6.18 x 10^6/mL Larvae No differences in activity or refuge use (Han et al. 2011)

JEL 274 50,000 Juveniles No significant differences in mortality, (Gervasi et al. 2014) Infection load decreased over time, stronger bacterial killing response over time, elevated neutrophil levels JEL 274 10,00, 50,000, Juveniles Reduced survival (Gervasi et al. 2013a) 100,000 JEL 215 2 culture dishes Juveniles No differences in temperature selection (Han et al. 2008) inoculated in batches with 20 larvae JEL 274 8.5x10^4 Juveniles Lower stress hormone levels (Searle et al. 2014) Section line 2.2 x 10^5 Juveniles Strain differences in mortality and (Piovia-Scott et al. 2015) lake and infection dynamic, no differences in Carter survivorship between populations. Bd Meadow prevalence and infection intensity differed between populations JEL 215 2.08 × 10^7 Juveniles Reduced survival (Garcia et al. 2006)* Rana draytonii VU JEL 270 n/a Juveniles No significant difference in survival or (Padgett-Flohr 2008)** (California red-legged mass frog) (Rachowicz & Rana muscosa EN JEL 217 3.6 x 10^9 Larvae Infected but appear healthy, loss of Vredenburg 2004a)** (Mountain yellow-legged mouth pigmentation 49

frog)

JEL 217 n/a Larvae Transmitted infection to each other and (Rachowicz & to post-metamorphic individuals Vredenburg 2004a)**

LJR089 1x10^7 Larvae Proportion of hosts that became (Rachowicz & Briggs infected increased with the number of 2007) previously infected R. muscosa larvae to which they were exposed

LJR089 1x10^7 Juveniles Temperature dependent effects on (Andre et al. 2008) survival, increased skin shedding

n/a >100,000 in 1mL Adults Disruption of skin integrity, ion (Rosenblum et al. 2012) imbalance

Rana sierra EN TST75,CJB 200,000 Juveniles Altered microbiome (Jani & Briggs 2014)** (Sierra Nevada yellow- 4, CJB5, legged frog) CJB7 Rana temporaria LC BdGPL IA- 160,16,000 Juveniles No significant differences in survival, (Bielby et al. 2015) (Common frog) 42 high dose resulted in less weight gain or weight loss Xenopus tropicalis LC IA042 10^6 Adults Temperature dependent effects on (Ribas et al. 2009)** (Western clawed frog) immune response n/a n/a Adults Altered gene expression to (Rosenblum et al. 2009)** physiological and immunological genes Xenopus laevis LC JEL 197 and n/a Adults Impaired lymphocyte proliferation and (Fites et al. 2013) (African clawed frog) JEL 275 induced splenocyte apoptosis JEL 197 and 10^6 Adult Peptide-depleted frogs became more (Ramsey et al. 2010)** JEL 275 susceptible to Bd infection with higher burdens and weight loss

Table 2.1b: Effects of Batrachochytrium salamandrivorans on amphibian hosts

Species IUCN Bsal Dose Life Stage Effect Reference Status Strain

Alytes obstetricans LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014) (Common midwife toad) AMFP13/1 5,000 in 1mL Adults No significant effect (Martel et al. 2013) Ambystoma maculatum LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014) (Spotted salamander) Ambystoma opacum LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014)** (Marbled salamander) Bombina variegata LC AMFP13/1 5,000 in 1mL Adults No infection or disease detected (Martel et al. 2014) (Yellow-bellied toad) Cynops pyrrhogaster LC AMFP13/1 5,000 in 1mL <1 year Susceptible to infection and (Martel et al. 2014) (Japanese fire-bellied disease newt) Discoglossus scovazzi LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014) (Moroccan painted frog) Epidalea calamita LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014) (Natterjack Toad) Euproctus platycephalus EN AMFP13/1 5,000 in 1mL Adults Reduced survival, confirmed (Martel et al. 2014) (Sardinian brook invasion of the skin salamander) Gyrinophilus LC AMFP13/1 5,000 in 1mL Adults No infection or disease detected (Martel et al. 2014) porphyriticus (Spring salamander) Hyla arborea LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014) (European tree frog) Hynobius retardatus LC AMFP13/1 5,000 in 1mL Adults No infection or disease detected (Martel et al. 2014) (Ezo salamander) Hypselotriton cyanurus LC AMFP13/1 5,000 in 1mL Adults Susceptible to infection and (Martel et al. 2014)** (Chuxiong fire-bellied disease newt) Ichthyosaura alpestris LC AMFP13/1 5,000 in 1mL <1 year Reduced survival, confirmed (Martel et al. 2014) (Alpine newt) invasion of the skin Lissotriton helveticus LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014) (Palmate newt) Lissotriton italicus LC AMFP13/1 5,000 in 1mL <1 year Reduced survival (Martel et al. 2014) (Italian newt) Lithobates catesbeianus LC AMFP13/1 5,000 in 1mL Adults No infection or disease detected (Martel et al. 2014) (American bullfrog) 51

Neurergus crocatus VU AMFP13/1 5,000 in 1mL Adults Reduced survival, confirmed (Martel et al. 2014) (Yellow-spotted newt) invasion of the skin Notophthalmus LC AMFP13/1 5,000 in 1mL Adults Reduced survival, confirmed (Martel et al. 2014)** viridescens invasion of the skin (Eastern newt) Pachyhynobius VU AMFP13/1 5,000 in 1mL Adults No infection or disease detected (Martel et al. 2014) shangchengensis (Shangcheng stout salamander) Paramesotriton deloustali VU AMFP13/1 5,000 in 1mL Adults Susceptible to infection and (Martel et al. 2014) (Tam Dao salamander) disease Pelobates fascus LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014) (Common spadefoot toad) Plethodon glutinosus LC AMFP13/1 5,000 in 1mL Adults Confirmed infection of the skin, no (Martel et al. 2014)** (Northern slimy disease detected salamander) Pleurodeles waltl NT AMFP13/1 5,000 in 1mL <1 year Reduced survival, confirmed (Martel et al. 2014) (Iberian ribbed newt) invasion of the skin Rana temporaria LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014) (European common frog) Salamandra salamandra LC AMFP13/1 5,000 in 1mL Adults Reduced survival, ataxia. (Martel et al. 2013) (Fire salamander) Cohousing effectively transmits infection AMFP13/1 5,000 in 1mL Adults Warmer temperatures can clear (Blooi et al. 2015a) infection AMFP13/1 10^5 in 1mL Adults Topical treatments can reduce (Blooi et al. 2015b) fungal loads and in combination with warmer temperature can clear infection AMFP13/1 5,000 in 1mL <1 year Reduced survival, confirmed (Martel et al. 2014) invasion of the skin Salamandrella LC AMFP13/1 5,000 in 1mL Adults Confirmed infection but no effects (Martel et al. 2014) keyserlingii of disease or on survival (Siberian salamander) Salamandrina LC AMFP13/1 5,000 in 1mL <1 year Reduced survival (Martel et al. 2014) perspicillata (Northern spectacled salamander) Silurana tropicalis LC AMFP13/1 5,000 in 1mL <1 year No infection or disease detected (Martel et al. 2014) (Western clawed frog) Siren intermedia (Lesser LC AMFP13/1 5,000 in 1mL Adults Confirmed infection but no effects (Martel et al. 2014) siren) of disease or on survival Speleomantes NT AMFP13/1 5,000 in 1mL Adults Reduced survival (Martel et al. 2014)** 52

strinatii (French Cave Salamander) Taricha granulosa LC AMFP13/1 5,000 in 1mL <1 year Reduced survival (Martel et al. 2014) (Rough-skinned newt) Triturus cristatus LC AMFP13/1 5,000 in 1mL <1 year Reduced survival, confirmed (Martel et al. 2014) (Northern crested newt) invasion of the skin Tylototriton wenxianensis VU AMFP13/1 5,000 in 1mL <1 year Reduced survival (Martel et al. 2014) (Wenxian knobby newt) Typhlonectes LC AMFP13/1 5,000 in 1mL Adults No infection or disease detected (Martel et al. 2014) compressicauda (Cayenne caecilian)

Table 2.1c: Effects of ranavirus on amphibian hosts

Species IUCN Isolate Dose Type of Life Stage Effect Reference Status Exposure

Ambystoma VU ATV 200uL of inoculum w/ Injection Adults Reduced survival (Picco et al. 2007)** californiense 1,000 virions of ATV in (California tiger APBS solution salamander) Ambystoma LC ATV n/a Contaminated Larvae Reduced survival (Jancovich et al. 2001)* gracile water (Northwestern salamander)

Ambystoma LC FV3 and 10^3 PFU/mL Water bath Larvae Reduced survival, strain (Hoverman et al. 2012a) maculatum FV3-like obtained from 5x10^6 differences in infection (Spotted isolate PFUs in 30uL of salamander) Eagle's MEM Ambystoma n/a ATV 1 x 10^3.3 and 7.1 x Water bath Larvae Population differences in (Schock et al. 2009) mavortium 10^3 TCID50/mL (1.4 infection (Barred tiger million virions per salamander) animal) Ambystoma LC FV3 and 10^3 PFU/mL Water bath Larvae Reduced survival, no strain (Hoverman et al. 2012a) opacum FV3-like obtained from 5x10^6 differences in infection (Marbled isolate PFUs in 30uL of salamander) Eagle's MEM

Ambystoma LC FV3 and 10^3 PFU/ml obtained Water bath Larvae No significant differences in (Hoverman et al. 2012a) talpoideum FV3-like from 5x10^6 PFUs in survival, no strain (Mole isolate 30uL of Eagle's MEM differences in infection salamander) Ambystoma LC ATV 10^2, 10^2.5, 10^3, Water bath Larvae Dose dependent infection (Brunner et al. 2005) tigrinum (ATV- 10^3.5, 10^4, 10^5 and survival rates (Tiger DO211) PFU from original salamander) plaque assay of 4.5x10^7 ATV 2x10^6 from 200mL of Water bath Larvae No significant differences (Brunner et al. 2007) 10^4 PFU/mL in aged between transmission rates tap water ATV 2x10^7 of ATV for a Water bath with Larvae No infection when exposed (Brunner et al. 2007) final concentration of pond sediment to virus in dried substrate, 6.67x10^4 PFU/mL but when substrate was kept moist they became infected and experienced reduced survival ATV 500 PFU in 200uL Injection Larvae 1s ventral surface to ventral (Brunner et al. 2007) surface contact results in infection ATV 4x10^6 PFU from 400 Water bath Larvae Infection rate increases with (Brunner et al. 2007) mL of 10^4 PFU/mL in time and increased SVL aged tap water ATV 10^3 PFU/mL, 10^4 Water bath Larvae Temperature influences (Rojas et al. 2005) PFU/mL infectivity, survival, and time to death. Sub-lethal infections result in carrier status ATV 10^2, 10^2.5, 10^3, Water bath Larvae Dose and developmental (Brunner et al. 2005) 10^3.5, 10^4, 10^5 stage dependent infection PFU from original rates plaque assay of 4.5x10^7 FV3 and 10^3 PFU/ml obtained Water bath Larvae Reduced survival, strain (Hoverman et al. 2012a) FV3-like from 5x10^6 PFUs in differences in infection isolate 30uL of Eagle's MEM ATV 10^3 PFU/mL Water bath Larvae No significant differences in (Brunner et al. 2004) survival rates between larvae and juveniles

ATV 10^3 PFU/mL Water bath Juveniles Reduced survival (Brunner et al. 2004)

ATV n/a Water bath Adults Reduced survival (Jancovich et al. 2001)

Ambystoma n/a ATV 200 uL of inoculum w/ Injection Adults Reduced survival (Picco et al. 2007)* tigrinum 1,000 virions of ATV in mavortium APBS solution (Barred tiger salamander) Ambystoma n/a ATV 200 uL of inoculum w/ Injection Adults Reduced survival (Picco et al. 2007) tigrinum 1,000 virions of ATV in nebulosum APBS solution (Arizona tiger salamander)

Ambystoma n/a ATV 200uL of inoculum w/ Injection Adults Reduced survival (Picco et al. 2007)* tigrinum 1,000 virions of ATV in stebbinsi APBS solution (Sonora tiger salamander)

Anaxyrus LC FV3-like 10^3 PFU/mL Water bath Embryo through Developmental stage (Haislip et al. 2011) americanus isolate metamorphosis dependent infection and (American survival rates toad) FV3 and 10^3 PFU/ml obtained Water bath Larvae No significant differences in (Hoverman et al. 2012a) FV3-like from 10^6 PFUs in survival, no strain isolate 60uL of Eagle's MEM differences in infection Bufo bufo LC RUK 11, 10^6 PFU, 10^4 PFU Water bath Larvae Reduced survival, dose (Duffus et al. 2014a) (Common toad) RUK 13, in 30mL dependent infection and BUK 2, survival, strain differences BUK 3 in infection Cophixalus LC BIV 10^3 TCID50/mL Water bath, Adults Reduced survival (Cullen & Owens 2002)* ornatus injection, contact (Ornate frog) Gastrophryne LC FV3 and 10^6 PFUs in 10uL of Oral dose, water Larvae No significant differences in (Hoverman et al. 2010) carolinensis FV3-like Eagle's MEM bath survival and no strain (Eastern isolate differences in viral load narrow- mouthed toad)

FV3 and 10^3 PFU/mL Water bath Larvae No significant differences in (Hoverman et al. 2012a) FV3-like obtained from 10^6 survival, no strain isolate PFUs in 60uL of differences in infection Eagle's MEM Hyla LC FV3-like 10^3 PFU/mL Water bath Embryo through Reduced survival (Haislip et al. 2011) chrysoscelis isolate metamorphosis (Cope's gray tree frog) FV3 and 10^6 PFUs in 10 uL of Oral dose, water Larvae Reduced survival, exposure (Hoverman et al. 2010) FV3-like Eagle's MEM bath type dependent effects on isolate survival and infection FV3 and 10^3 PFU/ml obtained Water bath Larvae Reduced survival, strain (Hoverman et al. 2012a) FV3-like from 10^6 PFUs in differences in infection isolate 60uL of Eagle's MEM FV-3 10^3 PFU/mL Water bath Larvae Transmission can occur (Brenes et al. 2014a) between vertebrate classes. Amphibian larvae more susceptible to Rv than other vertebrate classes Limnodynastes LC BIV 10^0, 10^1, 10^2.5, Water bath, Larvae Reduced survival, renal, (Cullen et al. 1995)* terraereginae and 10^4 TCID50/mL injection hepatic, splenic, and (Northern banjo (bath); 0.1 mL of 10^3 pulmonary necrosis frog) TCID50/mL (injection) BIV 10^0, 10^1, 10^2.5, Water bath, Juveniles Reduced survival, renal, (Cullen et al. 1995)* 10^4 TCID50/mL injection hepatic, splenic, and (bath); 0.1 mL of 10^3 pulmonary necrosis TCID50/mL (injection) Lithobates LC ATV n/a Feeding on Larvae No signs of infection (Jancovich et al. 2001)* catesbeianus infected (American salamander bullfrog) FV3 and 10^3 PFU/mL Water bath Larvae No differences in survival, (Hoverman et al. 2012a) FV3-like obtained from 10^6 no strain differences in isolate PFUs in 60uL of infection Eagle's MEM ATV 200uL ATV/EPC with Injection Adults No signs of infection (Jancovich et al. 2001)* 4x10^5 PFU/mL Lithobates LC FV3-like 10^3 PFU/mL Water bath Embryo through Reduced survival (Haislip et al. 2011) clamitans isolate metamorphosis (Green frog) FV3 and 10^3 PFU/mL Water bath Larvae Reduced survival, strain (Hoverman et al. 2012a) FV3-like obtained from 10^6 differences in infection isolate PFUs in 60uL of 56

Eagle's MEM

Lithobates LC FV3 and 10^6 PFUs in 10uL of Oral dose, water Larvae Reduced survival, exposure (Hoverman et al. 2010) palustris FV3-like Eagle's MEM bath type dependent effects on (Pickerel frog) isolate survival and infection

FV3 and 10^3 PFU/mL Water bath Larvae Reduced survival, strain (Hoverman et al. 2012a) FV3-like obtained from 10^6 differences in infection isolate PFUs in 60uL of Eagle's MEM Lithobates LC FV3-like 10^3 PFU/mL Water bath Embryo through Reduced survival (Haislip et al. 2011) pipiens isolate metamorphosis (Northern leopard frog) FV3 and 10^3 PFU/mL Water bath Larvae Reduced survival, isolate (Hoverman et al. 2012a) FV3-like obtained from 10^6 differences in infection isolate PFUs in 60uL of Eagle's MEM FV3 50 mL of water with Water bath Larvae Strain dependent effects on (Morrison et al. 2014)* strains 10,000 PFU/mL survival (SSME, wt-FV3, aza-C) FV3 10,000 PFU/mL Water bath Larvae Temperature and strain (Echaubard et al. 2014) isolate dependent effects (wt-FV3), azacR, SsMeV

ATV 100uL of ATV/EPC Injection Adults No signs of infection (Jancovich et al. 2001)* which had 4x10^5 PFU/mL in EPC cells

Lithobates CR FV3-like 10^3 PFU/mL with 400 Water bath, Adults Reduced survival, exposure (Sutton et al. 2014) sevosus isolate mL of water injection, oral type dependent effects on (Mississippi dose survival gopher frog)

(Hoverman et al. 2012a) Lithobates LC FV3 and 10^3 PFU/mL Water bath Larvae Reduced survival, strain Sphenocephala FV3-like obtained from 10^6 differences in infection (Southern isolate PFUs in 60uL of 57

leopard frog) Eagle's MEM (Haislip et al. 2011) Lithobates LC FV3-like 10^3 PFU/mL Water bath Embryo through Reduced survival sylvatica isolate metamorphosis (Wood frog) FV3 10,000 PFU/mL with Water bath Larvae Infection dependent on (Echaubard et al. 2014) isolate 50mL water temperature and strain (wt-FV3), azacR, SsMeV n/a n/a Contact and Larvae Reduced survival (Harp & Petranka 2006)* feeding on infected individuals n/a n/a Exposure to Larvae No significant differences in (Harp & Petranka 2006)* contaminated survival or growth, but sediment and tested positive for Rv water FV3 and 10^3 PFU/ml obtained Water bath Larvae Reduced survival, no strain (Hoverman et al. 2012a) FV3-like from 10^6 PFUs in differences in infection isolate 60uL of Eagle's MEM

FV3-like 10 fold dilutions from Water bath Larvae Dose dependent survival (Warne et al. 2011) isolates 2.36x10^1 through rates, no strain differences 2.36 x 10^5 PFU/mL in infection for wood frog isolate and 2.51x10^1 through 2.51 x 10^5 PFU/mL for spotted salamander isolate)

FV3-like 2.36 x 10^3 PFU/mL Water bath Larvae Higher stress hormone (Warne et al. 2011) isolate levels

FV3 67, 670, 6,700 Water bath Larvae Horizontal transmission the (Duffus et al. 2008) PFU/mL most likely means of FV3 transmission Litoria caerulea LC BIV 10^3 TCID50/mL; Water bath, Juveniles Reduced survival, exposure (Cullen & Owens 2002)* (Australian 10^4.5 TCID50/mL injection type dependent effects on green tree frog) survival

BIV 10^3 TCID50/mL Water bath, Adults No significant differences in (Cullen and Owens 2002)* injection, contact survival Litoria inermis LC BIV 10^3 TCID50/mL Injection Adults Tested negative for (Cullen and Owens 2002)* (Bumpy rocket infection frog) Litoria LC BIV 10^3 TCID50/mL Injection Larvae Reduced survival, renal, (Cullen et al. 1995)* latopalmata hepatic, splenic, and (Broad-palmed pulmonary necrosis frog) LC BIV 10^3 TCID50/mL Injection Juveniles Reduced survival, renal, (Cullen et al. 1995)* hepatic, splenic, and pulmonary necrosis Litoria rubella LC BIV 10^4.5 TCID50/mL Injection Adults No significant differences in (Cullen and Owens 2002)* (Desert tree survival frog) Notophthalmus LC ATV n/a Contaminated Adults Reduced survival (Jancovich et al. 2001)* viridescens water (Eastern newt) FV3 and 10^3 PFU/mL Water bath Larvae No significant differences in (Hoverman et al. 2012a) FV3-like obtained from 10^6 survival, no strain isolate PFUs in 60uL of differences in infection Eagle's MEM

Pseudacris LC FV3 and 10^3 PFU/ml obtained Water bath Larvae Reduced survival, strain (Hoverman et al. 2012a) brachyphona FV3-like from 10^6 PFUs in differences in infection (Mountain isolate 60uL of Eagle's MEM chorus frog) Pseudacris LC FV3-like 10^3 PFU/mL Water bath Embryo through Reduced survival (Haislip et al. 2011) feriarum isolate metamorphosis (Upland chorus frog) FV3 and 10^3 PFU/mL Water bath Larvae No significant differences in (Hoverman et al. 2012a) FV3-like obtained from 10^6 survival, no strain isolate PFUs in 60uL of differences in infection Eagle's MEM Pseudacris LC FV3 and 10^3 PFU/mL Water bath Larvae Reduced survival, no strain (Hoverman et al. 2012a) triseriata FV3-like obtained from 10^6 differences in infection (Western isolate PFUs in 60uL of chorus frog) Eagle's MEM Rana capito NT FV3 and 10^3 PFU/ml obtained Water bath Larvae Reduced survival, no strain (Hoverman et al. 2012a) (Gopher frog) FV3-like from 10^6 PFUs in differences in infection isolate 60uL of Eagle's MEM

Rana latastei VU FV3 2.25 x 10^6 PFU/mL Water bath Larvae Reduced survival (Pearman et al. 2004) (Italian agile (aliquots of 10 mL) frog) from 70mL of stock solution with 5.5x10^8 PFU/mL added to aged tap water FV3 4.5 x 10^6 Water bath Larvae Dose dependent survival (Pearman et al. 2004) PFU/mL(aliquots of 10 and survival rates mL), 4.5 x 10^5, 4.5 x 10^4, 4.5 x 10^3, 4.5 x 10^2 FV3 Feeder tadpoles Consuming Larvae Exposure type dependent (Pearman et al. 2004) infected with 4.5x10^6 infected survival rate PFU/mL carcasses FV3 4.5 x 10^4 PFU/mL, Water bath Larvae Dose dependent survival, (Pearman & Garner 2005) 4.5 x 10^6 PFU/mL effect of genetic diversity on (this was achieved by survival adding 2.796x10^8 PFU of FV3 to 615mL of aged water, low exposure was a 1:100 dilution of this)

Rana LC RUK 11, 10^6 PFU, 10^4 PFU Water bath Larvae Dose and strain dependent (Duffus et al. 2014) temporaria RUK 13, in 30mL effects on survival (Common frog) BUK 2, BUK 3 BIV, 10^4 TCID50/mL Water bath Larvae Strain and temperature (Bayley et al. 2013) DFV, dependent effects on ECV, survival EHNV, FV3, GV6, PPIV, REV, and SERV BIV, 10^4 TCID50/mL Water bath Juveniles Strain dependent effects on (Bayley et al. 2013) DFV, survival ECV, EHNV, FV3, GV6, 60

PPIV, REV, and SERV

No 67 0.25mL Injection Adults Reduced survival (Cunningham et al. 2007)** intraperitoneally, 0.25 subcutaneously both from 10^5-7 TCID50/mL stock

Scaphiopus LC FV3-like 10^3 PFU/mL Water bath Embryo through Reduced survival (Haislip et al. 2011) holbrookii isolate metamorphosis (Eastern spadefoot toad) FV3 and 10^3 PFU/mL Water bath Larvae Reduced survival, isolate (Hoverman et al. 2012a) FV3-like obtained from 10^6 differences in infection isolate PFUs in 60uL of Eagle's MEM Taudactylus CR BIV 10^3 TCID50/mL Water bath Adults Reduced survival (Cullen and Owens 2002)* acutirostris (Sharp snouted day frog) Xenopus laevis LC FV3 10^4 PFU in 10uL Injection Larvae Developmental stage (Grayfer et al. 2015) (African clawed differences in immune frog) response

FV3 10^4 PFU in 10uL for Water bath, Larvae Developmental stage (Andino et al. 2012) injection; 10uL of 10^5 injection, oral dependent immune function PFU for oral ingestion; ingestion and infection rates and 2mL of 5 x 10^6 PFU for water bath

FV3 0.1mL volume of Injection Juveniles Developmental stage (Andino et al. 2012) 1x10^6 PFU dependent immune function and infection rates FV3 7.2 x 10^7 TCID50 in Injection Adults Virus transcription was (Robert et al. 2007) 300uL PBS detected in macrophages up to 12 days after infection FV3 1x10^6 to 5x10^6 PFU Injection Adults Host cell differences in viral (Morales et al. 2010) in 300uL clearance

FV3 1 x 10^6 PFU n/a Adults Immunocompromised (Robert et al. 2011) adults can transmit infection within 3 hours FV3 5x10^6 PFU in 100uL Injection Adults Developmental stage (Grayfer et al. 2015) differences in immune response

FV3 5 x 10^7 PFU in 300 Injection Adults Develop long-lasting (Maniero et al. 2006) uL PBS protective immunity post initial exposure to FV3

* Indicates animals were not reared from eggs ** Indicates animals were not reared from eggs but were verified as Bd or Rv negative before the start of the experiment *** Indicates collection information unavailable

CHAPTER 3: COMPLEX DYNAMICS OF TWO EMERGING PATHOGENS, BATRACHOCHYTRIUM DENDROBATIDIS AND RANAVIRUS, IN AMPHIBIAN HOSTS

Natalie M. Hambalek, Emily Reynolds, Trang Dang, Jason Hoverman, Deanna Olson and Andrew Blaustein

ABSTRACT

Emerging infectious diseases are a contributing factor to global biodiversity loss. Amphibian populations are notable examples of the current biodiversity crisis, undergoing unprecedented declines and extinctions. The amphibian chytrid fungus, Batrachochytrium dendrobatidis, and ranaviruses are two emerging pathogens that are contributing to the global loss of amphibian biodiversity. The effects of B. dendrobatidis and ranaviruses on their hosts have been well documented but the effects of co-infection with these pathogens are poorly understood. We experimentally examined the effects of simultaneous and independent exposure of ranavirus and

B. dendrobatidis on survival, growth, and activity levels in two amphibian host species, the

Pacific treefrog (Pseudacris regilla) and the western toad (Anaxyrus boreas). We predicted that coinfection with Bd and Rv would have different effects on hosts compared with hosts that were infected with one of the pathogens alone. Our results showed antagonistic effects of concurrent exposure to B. dendrobatidis and ranavirus. P. regilla exhibited significant mortality in response to ranavirus independently, but not in combination with B. dendrobatidis. Pathogen treatment had no influence on mass in either species, but in both species, independent exposure to ranavirus resulted in decreased snout-vent length (SVL). Furthermore, there was a trend of increased activity levels for tadpoles exposed to both pathogens in comparison to each independently. Pseudacris regilla tadpoles exposed to B. dendrobatidis were significantly less active than those in the control and B. dendrobatidis x ranavirus treatments. We also present

evidence for interspecific variation in host sensitivity to these pathogens. Anaxyrus boreas did not experience any significant effects of pathogen exposure. However, compared to P. regilla, A. boreas tadpoles experienced decreased survival when exposed to B. dendrobatidis independently and in combination with ranavirus. We conclude that the dynamics behind multiple pathogen exposure are complex and that one pathogen may mitigate the effects of another to induce response variation.

INTRODUCTION

Emerging infectious diseases (EIDs) are one of the primary threats to global biodiversity

(Daszak et al. 2000; Harvell et al. 2002; Fisher et al. 2012; McCallum 2012; Tompkins et al.

2015). However, there is a gap in our knowledge concerning how both host and pathogen diversity affect disease dynamics (Holt & Dobson 2006; Pedersen & Fenton 2007; Pongsiri et al.

2009; Keesing et al. 2010). Hosts often encounter multiple pathogens simultaneously, with pathogens combining to produce complex effects within individual hosts (Petney & Andrews

1998; Bentwich et al. 1999; Jolles et al. 2008). For example, synergistic effects of multiple pathogens on individual hosts can increase infection prevalence and severity and have the potential to reduce host population size (Joly & Messier 2005; Munson et al. 2008). This may be especially significant for amphibians, whose populations are declining worldwide (Alford &

Richards 1999; Stuart et al. 2004). However, potential interactive effects of multiple pathogens contributing to amphibian population declines has received little attention (but see for examples

(Cunningham et al. 1996; Johnson & Buller 2011; Romansic et al. 2011), even though amphibians serve as hosts to a multitude of pathogens and parasites including bacteria, viruses, fungi, and helminths (Worthylake & Hovingh 1989; Blaustein 1994; Cunningham et al. 1996; Green et al.

2002; Johnson et al. 2002).

Two pathogens have been especially important contributors to amphibian population declines: Batrachochytrium dendrobatidis (Bd), and ranavirus (Rv). Both pathogens have been associated with amphibian mortality events and population declines worldwide (Lips 1998,

1999; Daszak et al. 1999b; Rachowicz et al. 2006; Mazzoni et al. 2009; Kik et al. 2011) and have been listed as notifiable pathogens by the World Organization for Animal Health

(Schloegel et al. 2010). Bd, the cause of amphibian chytridiomycosis, includes the thickening and sloughing of the skin which may cause death via disruption of vital functions such as cutaneous respiration, osmoregulation, and ion exchange (Voyles et al. 2009; Kilpatrick et al.

2010). Ranaviral disease, ranavirosis, causes systemic hemorrhage and tissue necrosis, ultimately leading to organ failure (Gray et al. 2009b). Ranaviruses are especially lethal in larval amphibians (Gantress et al. 2003; Converse & Green 2005; Haislip et al. 2011), whereas post- metamorphic amphibians are most vulnerable to fatal effects of Bd (Rachowicz & Vredenburg

2004b; Gervasi et al. 2013a). Moreover, there is evidence that these two pathogens cooccur in amphibian hosts (e.g., Shock et al. 2010; Hoverman et al. 2012; Whitfield et al. 2013; Warne et al. 2016). Although they appear ubiquitous and co-occur in the wild and in captivity, the effects of simultaneous exposure to both Rv and Bd have not been investigated in controlled experiments. We experimentally investigated effects of single exposure and potential complex effects of simultaneous exposure of Bd and Rv within two species of amphibian hosts.

METHODS

Animal Collection and Husbandry

We collected recently oviposited (within 24 hours) egg masses of western toads

(Anaxyrus boreas) from Deschutes County, Oregon, USA on 20 May 2015. We collected recently oviposited (within 24 hours) Pacific treefrog (Pseudacris regilla) egg masses from 65

Lane County Oregon, USA on 25 January 2015. Egg masses were placed in 37L aquaria

containing dechlorinated water supplied with aeration within four hours of collection. Upon

hatching, tadpoles separated by species were placed in aquaria in densities of approximately 100

individuals per 37L tank. Tadpoles were fed a 3:1 ratio of ground alfalfa pellets and fish flakes

ad libitum. All animals were kept between 14 and 16°C on a 14:10 hour light:dark photoperiod

for the duration of the experiment. Since tadpoles were collected at different times, to ensure

similar developmental stages were used for each species at the start of the experiment, the

experiments were staggered by six weeks. Mass and snout-vent length (SVL) were measured for

each individual prior to initiation of the experiment. SVL was measured utilizing imaging

software, CascadeSMT (mean ± 1SE, P. regilla = 7.68mm ± 0.17 and A. boreas = 8.5mm ±

0.14).

For each species tested, we used a 2 x 2 randomized factorial design consisting of four treatment groups: Control, Bd only, Rv only, and Bd x Rv, with 25 replicates per treatment.

Treatments were applied on the same day within 30 minutes. When each species reached Gosner stages 27-29 (Gosner 1960), tadpoles were individually and randomly assigned to beakers filled with 800mL of dechlorinated water for treatment inoculation. Beakers with individual treatment replicates were randomly blocked and placed along the laboratory bench. Tadpoles were allowed to acclimate to their individual containers for 24 hours. Tadpoles were fed approximately 0.05g every-other day and full water changes occurred every seven days. Animals were checked daily for activity and survival for the duration of the 60-day experiment. Behavioral activity levels were quantified utilizing scan sampling whereby each animal was observed five times, consecutively per scan, per day, and marked either active (moving vertically or horizontally in the water column with notable tail swiping), or non-active (remaining in place).

Pathogen contaminated water, aquaria, and materials were treated with a 10% bleach solution, a conservative concentration that ensures deactivation of Bd and Rv (Bryan et al. 2009;

Gold et al. 2013). Dead individuals were swabbed for the presence of Bd and preserved in 90% ethanol. All larvae remaining at the end of the experiment were removed, swabbed, measured, euthanized using MS-222, and preserved in 90% ethanol in accordance with Institutional Animal

Care and Use Committee (IACUC) policies (ACUP # 4441, Oregon State University).

Batrachochytrium dendrobatidis

Culture methods for Bd followed studies conducted previously (e.g., Gervasi et al. 2013a,

2014; Searle 2013). Culture plates (100 mm×15 mm, 1% Tryptone and agar) containing Bd (JEL

646, obtained from the Point Reyes, CA, from larval P. regilla from J. Longcore, University of

Main) were flooded with 10mL of dechlorinated water for 15 minutes to allow the discharge of motile zoospores from zoosporangia. Zoospores were gently scraped from culture plates and the liquid contents from each plate were combined with dechlorinated water until it reached an average concentration of 10,000 zoospores/mL, calculated using a haemocytometer. Tadpoles in all Bd exposure treatments were inoculated with 5mL of the 10,000 zoospore/mL solution resulting in 50,000 zoospores total with a final concentration of 62.5 zoospores/mL. This dose was chosen because it is considered to be an intermediate dose in a previous study, which resulted in dose-dependent survival in P. regilla (Gervasi et al. 2013a). We aimed for a low-to- moderate pathogen dose for both pathogens as we were interested in measuring sublethal effects.

Control treatments and treatments with no Bd (i.e., ranavirus only treatment) were given 5mL of sham inoculum consisting of liquid contents from 1% Tryptone and agar-only plates.

We collected DNA swabs by forcefully rotating sterile, fine-tipped, dry synthetic cotton swabs across the mouthparts of each tadpole for ten full rotations ensuring the swab stroked the keratinized jaw sheath, a modified method from Retallick et al. 2006. Swabs were placed into sterile 1.5mL micro-centrifuge tubes and kept in -20°C until analysis. DNA was extracted from swabs by adding 60L of Prepman Ultra® (Life Technologies), heating the vial for 10 min at

100°C, cooling the vials for two min, and extracting the supernatant. We diluted the supernatant to a 10% solution and then analyzed this sample utilizing a quantitative polymerase chain reaction (qPCR) assay using methods of Boyle et al. 2004. We conducted qPCR using the ABI

PRISM® 7500 Sequence Detection System (Applied Biosystems) to quantify the amount of Bd genomic material on each individual exposed to Bd (n=25/species) and Bd x Rv (n=25/species) treatments for each species. We used genetic markers specific for Bd consisting of primers from the internal transcribed spacer (ITS) region and compared each sample to standards (four serial dilutions from concentrations 0.1 to 100 zoospore genomic equivalents). We also quantified Bd- infection status in five randomly sampled unexposed individuals from the control treatments and the Rv only treatments to test for the presence of Bd. All samples including negatives and standards were run in triplicate and were considered positive if two of three samples indicated the presence of Bd genomic material.

Ranavirus

We used an FV3-like isolate obtained from a wood frog die-off event in Michigan.

Culture methods for Rv follow the methods of Hoverman et al. 2012. We passaged the FV3 isolate through fathead minnow cells fed with Eagle’s minimum essential medium (MEM) with

Hank’s salts, containing 5% fetal bovine serum. The isolate was on the second passage since

original isolation. It was stored at -80°C until the start of the experiments. Each Rv treatment subject received 222L of the virus solution resulting in a final concentration of 8x104 plaque forming units (PFUs), or 100 PFUs/mL. The no-Rv treatments were given 222L of sham inoculum, MEM.

DNA extractions were conducted following the methods of Hoverman et al. 2012. DNA was extracted utilizing pooled liver and kidney samples from each Rv treated individual using

DNeasy Blood and Tissue Kits (Qiagen). To determine Rv infection status and viral load of each individual we used qPCR following the methods of Forson & Storfer 2006. qPCR was performed using a Bio-Rad real-time qPCR system (Bio-Rad). Each qPCR run included a standard curve and a negative sample. A synthetic double-stranded DNA standard was used by synthesizing a

250 bp fragment of the major capsid protein (MCP) gene (gBlocks Gene Fragments; Integrated

DNA Technologies). We prepared a log-based dilution series (4.014 x 109 to 4.014 x 106 viral copies L-1) for the standard curve. The standard curve and unknown samples were run in duplicate. Five individuals from the control treatments and the Bd-only treatments were tested for the presence of Rv. All duplicated unknowns that peaked before cycle 40 were considered positive. After completion of qPCR, we quantified the concentration of genomic DNA in the samples (ng of DNAL-1) using a NanoDrop 2000c (Thermo Scientific). Then we used the genomic DNA concentration along with the viral concentration data to calculate the viral load

(i.e., viral copies ng-1 of DNA) of the positive samples.

Analyses

All statistical analyses were conducted using the R statistical software (Version 3.1.2, R

Foundation for Statistical Computing, 2014). Statistical analyses used to determine effects on 69

survival and infection load were conducted within species (among-treatment levels) and between species (within-treatment levels). We used a Cox Proportional Hazards (CPH) model to compare the probability of mortality through differences in survival curves with the “coxph” function and in the Survival package for survival analyses. As this was a randomized block design, we incorporated individual placement into the model. A CPH model determines a hazard ratio (HR), a comparative indicator that represents the association of a given factor with the probability of mortality at any given time point. A HR > 1 demonstrates an increase in the probability of mortality while a HR < 1 shows a decrease in the probability of mortality. We transformed Bd loads from qPCR (log-average genome equivalents per individual +1) to normalize data and analyzed the effects of treatment and mass on log-transformed infection loads with an analysis of covariance. To test for differences in growth (SVL) and mass, we performed a one-way analysis of variance followed by a Tukey’s honest significant difference test when significance was detected. To examine pathogen treatment effects on tadpole activity, we fit a generalized linear model with a binomial error distribution and the logit link function followed by a general linear hypotheses and multiple comparisons test using the “glht” function in the Multcomp package.

RESULTS

Survival and Pathogen Loads

There were no effects of placement (block) on survival for P. regilla (p = 0.178) or A. boreas (p = 0.358). The effects of pathogen treatment on survival differed with host-species and pathogen identity. In the treatment comparisons within species, there was no effect of Bd alone on P. regilla survival compared to the control group (HR = 0.306, p > 0.05). However, P. regilla in the Rv-only treatment group experienced a 10.48 fold increase in mortality (p < 0.001), but not in the treatment with Bd and Rv together (HR = 0.312, p > 0.05) (Figure 3.1). Although 70

mortality was observed across pathogen treatments, there were no significant effects of pathogen exposure on survival in A. boreas when controls were compared to pathogen-exposed animals due to unexplained mortality in the control group.

Pathogen loads

In P. regilla, Bd infection was observed in 48% and 64% in the Bd-only and Bd x Rv treatments, respectively but load differences were not significant. Rv infection was detected in

88% of Rv-only treated individuals. No Rv infection was detected in the Bd x Rv exposed individuals, although we observed 16% of these individuals with hemorrhaging, a common symptom of Rv infection. In A. boreas, Bd infection was observed in 76% and 52% of the Bd- only and Bd x Rv treatments, respectively. Bd loads did not differ between species or across Bd-

only and Bd x Rv treatments within a species (ANOVA, F1, 25 = 2.89, p > 0.05) (Figure 3.2).

There was no association between pathogen loads and mass for either pathogen or species (p >

0.05). Average Bd infection levels (in genome equivalents Bd ± 1SE) for P. regilla were 3.8 ±

1.44 and 4.16 ± 1.27 for Bd-only and Bd x Rv treatments and 8.06 ± 3.2 and 5.46 ± 2.34 for A. boreas. Average Rv viral load (copies/ng) for P. regilla in the Rv only treatment was 9.6x105 ±

1.6x104. No Rv infections were detected in any A. boreas individuals. We cannot rule out the possibility that some individuals died from minor infections that were not detected by qPCR as the assay does not rule out the possibility that Rv could have been present in the brain or digestive tract of the tested individual.

Growth and Activity

An analysis of variance on larval SVL and mass yielded no significant differences between treatment groups at the initiation of the experiment. At the end of the 60-day experiment, there were no significant differences of pathogen treatment on mass in either species

(ANOVA, df= 3, p > 0.05). However, tadpoles in the Rv-only treatment had shorter SVL in both

P. regilla (ANOVA, df =3, p = 0.029) and A. boreas (ANOVA, df =3, p = 0.0069) relative to controls (Table 3.1).

A generalized linear model revealed that exposure to pathogen treatment had an effect on activity in P. regilla (2!=!9.5, df!=!3, p = 0.023). Bd treatment alone resulted in a 28% decrease in activity compared with the control group (p = 0.03) and a 25% decrease in activity compared with the Bd x Rv treatment group (p = 0.04). There were no significant effects of pathogen treatment on A. boreas activity levels. However, there was a trend of reduced activity levels in the independent pathogen treatments relative to the control and the combination pathogen treatments for both species (Figure 3.3).

DISCUSSION

Our results suggest complex dynamics regarding coinfection and that different effects are manifested in hosts depending upon whether a host was infected with a single pathogen or two pathogens. Moreover, the host-pathogen dynamics differed among host species.

Batrachochytrium dendrobatidis

Our within-species comparisons showed no effect of any pathogen treatment on A. boreas when compared with controls. In previous studies, the Bd strain we used (JEL 646) caused mortality in A. boreas from the same population we studied and at the same 72

developmental stage (Gervasi et al. 2013a) and induced sublethal effects in other species (Gaietto et al. 2014; Caseltine et al. 2016). Since A. boreas is generally sensitive to Bd, lack of differences in mortality between Bd-exposed A. boreas and controls was a result of unexplained mortality in the control group. However, in the between-species comparisons, A. boreas exhibited a 12.8 fold increase in the probability of mortality in response to Bd than P. regilla, which is consistent with previous studies (Blaustein et al. 2005; Searle et al. 2011; Gervasi et al.

2013a). There was no effect of Bd alone on P. regilla survival in comparison with the control group. Pseudacris regilla seems to be more tolerant to low-level Bd infection compared with some other species, including A. boreas, especially as larvae (Blaustein et al. 2005; Searle et al.

2011b; Gervasi et al. 2013a).

Bd exposure alone resulted in decreased levels of tadpole activity in P. regilla and a trend of decreased activity levels in A. boreas, which is consistent with other experimental studies

(Parris et al. 2006; Venesky et al. 2010b; Han et al. 2011; Kleinhenz et al. 2012). Whereas reduction in activity levels may reduce predation risk (Lambert et al. 2004), decreased activity levels also can be associated with reduced foraging and prolonged time to metamorphosis

(Venesky et al. 2009; Kleinhenz et al. 2012).

Ranavirus

Pseudacris regilla exposed to Rv independently experienced over a 10-fold increase in mortality when compared with controls. We did not detect an effect of Rv on A. boreas.

Exposure to the Rv-only treatment also resulted in decreased SVL for both species. Smaller SVL may have deleterious effects for disease susceptibility. For example, smaller animals at metamorphosis resulted in increased vulnerability to Bd infection as juveniles (Garner et al.

2009). This is the first study documenting the effects of Rv in P. regilla and A. boreas. 73

Coinfection

Although Bd and Rv coinfection has been reported in the wild (Whitfield et al. 2013;

Warne et al. 2016) and in captivity (Miller et al. 2008; Kik et al. 2012), we did not detect Rv after experimental coinfection with Bd. However, Bd infections were confirmed in all combined pathogen treatments. We found no strong evidence that combined exposure to Bd and Rv caused additive effects in P. regilla or A. boreas. Rather, our results demonstrated antagonistic effects whereby independent pathogen exposure (i.e. exposure to Bd or Rv alone) resulted in higher mortality, reduced growth (Rv), and reduced activity (Bd). Pseudacris regilla was significantly affected by Rv alone, but the negative effect on mortality was negated with the addition of Bd.

This could be due to Bd and Rv targeting different localities in amphibian hosts.

Motile Bd zoospores target the keratinized mouthparts of tadpoles and the keratinized integument of post-metamorphic amphibians (Greenspan et al. 2012b). In contrast, non-motile

Rv virions are transmitted by ingestion or direct contact with their host (Brunner et al. 2007) but the mechanisms of Rv dissemination within the host are unclear, especially at the earliest stages of infection. Rv infection proceeds through the digestive tract and leads to infection in the kidney

(Robert et al. 2011), yet a recent study demonstrated that FV3 infection is capable of altering the blood brain barrier in larvae Xenopus laevis, eventually leading to Rv dissemination the central nervous and excretory systems (De Jesús Andino et al. 2016).

Interactions between pathogens and parasites may be occurring directly or indirectly.

Direct interaction, for example, can be illustrated by acanthocephalan worms displacing tapeworms from prime attachment sites within their hosts (Holmes 1959). Indirect interactions could occur from cross-reacting immune cells, whereby T-cells produced in response to one

parasite can cross-react with antigens from another (Lafferty 2010). This cross-reactive immune response can provide a protective effect (Lafferty 2010). For instance, being infected with one species of human schistosome may protect against new infections by other pathogens (Taylor et al. 1973). This dynamic has been observed in amphibians where prior infection with one trematode reduced later infection success of another type of trematode (Hoverman et al. 2013).

Such an interaction may have occurred between Bd and Rv. Bd can cause changes in amphibian skin microbiota (Walke et al. 2014; Jani & Briggs 2014; Federici et al. 2015) and in host physiology in ways which may render the host uninhabitable for Rv. There is also the possibility that through its pathology, Rv could be altering the immunology or some aspect of host making hosts less susceptible to Bd. However, the precise mechanism of interaction for these two pathogens is unknown and warrants further study.

Acknowledgements

We thank the Center for Genome Research and Biocomputing at Oregon State University for providing laboratory space for qPCR. Thank you to A. Hehrer for her assistance with behavioral data. Thanks to A. Karmally, C. Beasley, J. Urbina, and D. Negru for their support and thoughtful insight throughout this project. This research was conducted under Oregon State

University IACUC animal care and use permit #4661. Support was provided by the U.S. Forest

Service Pacific Northwest Research Station, Corvallis, Oregon and NSF/NIH EID grant

R01GM109499.

Table 3.1 Final larvae snout-vent length and mass measurements (mean ± SE) from Pseudacris regilla and Anaxyrus boreas (Oregon, USA) individuals that survived the 60-day experiment. Bd = Batrachochytrium dendrobatidis, Rv = ranavirus.

Treatment Control Bd Rv Bd x Rv

P. regilla 10.1 ± 0.67 11.9 ± 0.42 9.2 ± 0.07 11.8 ± 0.57 SVL 0.339 ± 0.018 0.345 ± 0.018 0.299 ± 0.067 0.354 ± 0.016 Mass

A. boreas 11.2 ± 0.40 11.8 ± 0.66 8.6 ± 1.17 11.3 ± 0.47 SVL Mass 0.204 ± 0.02 0.184 ± 0.003 0.164 ± 0.017 0.177 ± 0.012

Table 3.2 Hazard ratios and associated p-values for within and between-species analyses. Abbreviations are used to identify species and treatments. PR = Pseudacris regilla, AB = Anaxyrus boreas (Oregon, USA). Bd = Batrachochytrium dendrobatidis, Rv = ranavirus, C= control. For all hazard ratios the more extreme treatment or severely effected species is compared to the less extreme or severely effected species (i.e. the more extreme treatment or severely effected species is listed first).

Comparison Treatment Hazard p-value Ratio P. regilla Bd vs. C 0.306 0.30 Rv vs. C 10.48 < 0.001 *** Bd x Rv vs. C 0.312 0.31 Rv vs. Bd 37.37 < 0.001 *** Rv vs. Bd x Rv 35.7 < 0.001 *** Bd x Rv vs. Bd 1.02 0.98

A. boreas Bd vs. C 1.41 0.46 Rv vs. C 1.31 0.56 Bd x Rv vs. C 0.75 0.57 Bd vs. Rv 0.93 0.86 Rv vs. Bd x Rv 1.86 0.20 Bd x Rv vs. Bd 1.83 0.22

Species Bd - AB vs. PR 12.8 0.015 *** Rv - PR vs. AB 2.49 0.01 *** Bd x Rv - AB vs. PR 7.6 0.057 **

** Denotes marginal statistical significance *** Denotes statistical significance 76

Figure 3.1 Kaplan-Meier survival curves indicating mortality rate according to species (A = Pseudacris regilla, B= Anaxyrus boreas) and pathogen exposure group. Bd = Batrachochytrium dendrobatidis, Rv = ranavirus.

Figure 3.2 Mean Bd ± SE infection loads (log genome equivalents +1) according to species (Pseudacris regilla and Anaxyrus boreas) and pathogen treatment. Bd = Batrachochytrium dendrobatidis, Rv = ranavirus.

Figure 3.3 Proportion of active tadpoles for Pseudacris regilla (A) and Anaxyrus boreas (B) individuals reared in the presence of different pathogen treatments. Bd = Batrachochytrium dendrobatidis, Rv = ranavirus. Error bars represent ± 1SE.

CHAPTER 4: UNDERSTANDING THE ROLES OF EMERGING INFECTIOUS DISEASE ON AMPHIBIAN POPULATION DECLINES: SUMMARY AND IMPLICATIONS

The initial sounding of the alarm for amphibian population declines in the

1990s (Blaustein & Wake 1990) prompted a multitude of interdisciplinary investigations regarding various aspects of amphibian disease ecology. Well-designed experiments coupled with natural field observations have been a powerful approach

(Jenkins 2004) and may help us more fully understand the dynamics of amphibian population declines. While field experiments are useful in mimicking natural conditions, they are not always feasible or possible to have control over most environmental variables. Laboratory and mesocosm studies can augment field studies and have been used successfully to examine a variety of ecological processes

(Hairston 1989; Underwood 2007) including various aspects of amphibian population declines (Jenkins 2004).

Bullfrogs (Lithobates catesbeianus) are one example of how experiments may help elucidate the importance of Bd to a specific host. Because of their ubiquity, bullfrogs are widely reported to be resistant hosts and carriers of Bd (Daszak et al.

2004; Schloegel et al. 2010). However, experiments, controlling for all variables except Bd strain found bullfrogs were more or less vulnerable to Bd depending upon the strain used in the experiment (Gervasi et al. 2013b).

The experimental results using different methods for the same host species illustrates the difficulties in making generalizations of how specific pathogens affect a host. For example, western toads (Anaxyrus boreas) have been investigated in a

number of experimental studies (Table 2.1a). These studies used different Bd strains, different Bd doses and different life stages and the results of how the host was affected differed among the studies. For example, some studies showed reduced survival after exposure to Bd, others did not. Even experiments by the same author

(Han 2008, 2011) on western toads showed certain differences in how toads responded to Bd. In these studies, western toads were examined at the same life stage but were exposed to different Bd strains and Bd doses.

Our experimental study investigated the dynamics of a multi-pathogen assemblage in amphibian hosts. Most studies of amphibian disease concentrate on single-pathogen/single-stressor approaches. Our study suggests that the dynamics of at least two pathogens simultaneously occupying a host are complex. Our two different host species were affected by single and multiple pathogens in different ways relative to survival, body size, and behavior. This seems especially significant because the two pathogens we studied are important contributors to amphibian population declines and extinction events (Cunningham et al. 1996; Berger et al.

1998; Lips 1999; Green et al. 2002). We suggest that further studies be initiated with more than one pathogen because amphibians host multiple pathogens in captivity and in natural habitats. Knowledge of the interactions between multiple amphibian pathogens would greatly enhance our ability for amphibian conservation efforts. The role of diseases in amphibian population declines provides an added urgency to understanding the pathogen interactions in amphibian assemblages, thus management efforts will need to consider coinfection if they are to be effective.

We suggest more studies examine differences in susceptibility at the species and population levels as well as those that investigate strain variability, using controlled experiments. Controlled experimental studies examining differences in susceptibility to pathogens can aid in our understanding of the dynamics of epizoonotic outbreaks. Standardizing experimental methods is an essential component of investigating the role of pathogens in amphibian population declines. Moreover, studies that focus on a single cause contributing to amphibian population declines may underestimate the roles of multiple factors working synergistically to cause both direct and indirect effects. Developing a mechanistic understanding of how biotic and abiotic factors can drive disease dynamics will allow us to better predict outbreaks and better manage and alleviate consequences associated with emerging infectious diseases (Keesing et al. 2006).

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