University of Nevada, Reno the Role of Hydroperiod and Fluctuating Temperature on Disease Dynamics: a Disease Ecology Approach

University of Nevada, Reno

The role of hydroperiod and fluctuating temperature on disease dynamics: A disease ecology approach to understanding Yosemite toad declines

A thesis submitted in partial fulfillment of the requirements for the degree of Master of Science in Biology

Alexa L. Lindauer

Jamie L. Voyles/Thesis Advisor

August, 2018

Abstract

Environmental factors can alter host-pathogen interactions. Temperature and water availability are critical environmental factors that affect disease susceptibility in amphibian hosts due to amphibian thermal biology. Temperature and water can also alter prevalence and virulence of the amphibian pathogen Batrachochytrium dendrobatidis

(Bd), because Bd requires moisture and temperatures between 2-27°C to replicate.

Understanding disease dynamics under fluctuating temperatures and changes in water availability is important because climate change is likely to affect these environmental factors across local and landscape scales with potential implications for disease.

Investigating the effects of environment on disease outcomes for imperiled species may inform conservation and recovery efforts. To better understand the effects of reduced water availability and fluctuating temperature on disease, I examined (1) the effects of larval development under drought conditions on disease susceptibility post- metamorphosis in Yosemite toads; and (2) the effects of daily fluctuating temperatures on

Bd growth and reproduction rate in vitro.

Acknowledgements

I would like to acknowledge many people for helping to make my research possible. First, I am grateful for the guidance given to me by my advisor, Jamie Voyles.

She encouraged me to pursue my own questions and provided support through the process. In addition, her dedication to scientific communication has helped me become a stronger writer and presenter, invaluable skills for any career. I also thank my committee members, Chris Feldman, Paul Hurtado, and Jenny Ouyang; Voyles Lab members, including Mason Ryan, Goncalo Rosa, Gabi Rios-Sotelo, and Kristin Charles; and the

UNR Evoldoers for feedback on experimental design, statistical analyses and writing, and for general enthusiasm and support. In particular, I thank Voyles Lab undergraduates

Tiffany May and Kristen McCarty for their tireless dedication to amphibian husbandry and Patience Gbafa and Katie Haran for assistance with pathogen culturing. I also extend thanks to Dan Wetzel in the Richards-Zawacki Lab at the University of Pittsburgh for processing skin swabs, and to Paul Maier, PhD candidate at San Diego State University, for lending me temperature data from the field. In general, I thank University of Nevada

Reno Biology and EECB graduate students, faculty, and staff for contributing to my education and providing a sense of community during my time in Reno. I also thank the following funding sources: The Strategic Environmental Research and Development

Program (SERDP); the National Science Foundation (NSF) 1354152; and the Society for

Integrative Biology (SICB) Grant in Aid of Research.

Projects involving threatened species are not possible without the support of state and federal agencies and their non-profit partners. I thank Sarah Markeguard of U.S. Fish and Wildlife Service, Laura Patterson and Justin Garcia of the California Department of iii

Fish and Wildlife, and the Nevada Department of Wildlife for project support and for granting me collection permits in record time. I thank Stephanie Barnes of the U.S. Forest

Service for granting me permission to collect eggs in the Sierra National Forest and for her enthusiasm for Yosemite toad conservation. I thank Rob Grasso of Yosemite National

Park for help with Yosemite toad egg collection and transport and for project advice. I thank Jessie Bushell of the San Francisco Zoo for her guidance on amphibian husbandry.

I also thank the Yosemite Toad Recovery Team for their support and their efforts to preserve imperiled amphibian species and better understand causes of their declines.

I would not have pursued a project studying causes of Yosemite toad decline had it not been for Paul Maier, Steven Lee of the U.S. Geological Survey (USGS) Yosemite

Field Station, and Roland Knapp of the Sierra Nevada Aquatic Research Laboratory,

University of California Santa Barbara. Having never seen or heard of a Yosemite toad,

Paul and Steven hired me as an intern in the summer of 2011 to help identify Yosemite toad occupied meadows and characterize Yosemite toad habitat in Yosemite and Sequoia and Kings Canyon National Parks. I continued to work for the USGS, in collaboration with Roland Knapp and the National Park Service, on yellow-legged frog and Yosemite toad projects in the summers of 2013-2015. During these summers, I traveled extensively on and off trail, having the rare excuse to explore lakes and meadows that few people visit. I developed a sense of place and deep affinity for the high Sierra and its ecological communities, motivating me to study the Yosemite toad during my graduate career. In addition to introducing me to the Yosemite toad, I thank Paul Maier, Steven Lee, and

Roland Knapp for help with project design, writing, navigating the permitting process, and serving as invaluable mentors, each able to provide perspectives associated with their iv unique experiences and personal progression along the research trajectory (Paul as a current graduate student, Steven as a working government ecologist, and Roland as an established career ecologist). I also thank Paul, Steven, and Roland for their dedication to amphibian conservation in the Sierra Nevada.

Outside of the scientific community, I thank my husband, Everett Phillips, for love and support, and my family for instilling in me an appreciation for education and lifelong learning. As my grandfather said, “They can take your money, your home, but they can never take what is in your head - education is the one thing they can’t take from you.” I am grateful for the opportunity granted to me to pursue this degree.

Table of Contents

Abstract…………………………………………………………………………………….i

Acknowledgements………………………………………………………………………..ii

Table of Contents………………………………………………………………………….v

List of Tables……………………………………………………………………………viii

List of Figures………………………………………………………………………….....ix

Introduction: A disease ecology approach to understanding drought and disease threats to a threatened alpine endemic, Anaxyrus canorus……………………………1

Chapter 1 overview……………………………………………………………………4

Chapter 2 overview……………………………………………………………………5

Chapter 1: The effects of larval development under drought on Bd susceptibility post-metamorphosis……………………………………………………………………...7

Introduction……………………..……………………………………………………7

Methods…………………………………………...…………………………………11

Egg collection……………………………………………………………………11

Husbandry………………………………………………………………………..12

Larval response to decreased hydroperiod……………………………………….13

Juvenile susceptibility to Bd……………………………………………………..14

Statistical analysis……………………………………………………………….16

Results………………………………………...……………………………………..17

Larval response to decreased hydroperiod……………………………………….17 vi

Juvenile susceptibility to Bd……………………………………………………..19

Effects of axial deformity and larval development under drying conditions on Bd

susceptibility……………………………………………………………………..20

Discussion…………………………………………………………………………...23

Larval response to decreased hydroperiod……………………………………….23

Juvenile susceptibility to Bd……………………………………………………..25

Effects of larval development under drying conditions on Bd susceptibility..…. 27

Conclusions and implications for conservation..………………………………….28

Chapter 2: The effect of daily fluctuating temperatures on Bd growth and reproduction rate in vitro………………………………………………………………31

Introduction…………………………………………………………………………31

Methods…...…………………………………………………………………………36

Incubation temperature selection………………………………………………...36

Isolate selection, culturing, and plate set-up……………………………………..37

Quantification of Bd growth...…………………………………………………...38

Optical density……………………………………………………………….39

Zoospore production…………………………………………………………39

Zoosporangia viability assay…………………………………………………39

Statistical analysis………………………………………………………………..40

Results……………………………………………………………………………….42

Optical density growth measurements….……………………………………….42

Zoospore production…………………………………………………………….44 vii

Culture fecundity………………………………………………………………...44

Zoosporangia viability………………………………………….………………..46

Discussion…………………………………………………………………………...48

Husbandry Supplement………………………………………………………………...54

References……………………………………………………………………………….59

viii

List of Tables

Table 1-1. Cox proportional hazards model selection for survival of Bd-exposed toads using a stepwise approach with AIC……………………………………………………..22

Table 2-1. Parameter estimates for logistic growth models of Bd grown at stable (17.5°C and 27.5°C) and fluctuating temperatures (7.5°C – 27.5°C, “Flux”)……………………43

List of Figures

Figure 1. The disease triangle applied to A. canorus–Bd disease dynamics……………...6

Figure 2. Yosemite toad life history………………………………………………………3

Figure 1-1. Shortened hydroperiod decreases Yosemite toad larval period but does not affect size at metamorphosis……………………………………………………………..18

Figure 1-2. Survival and Bd load in juvenile A. canorus exposed to Bd inoculum or Bd- negative control solution…………………………………………………………………19

Figure 1-3. Differences in survival probability of Bd-exposed juveniles (A) among drought treatments and (B) between toads with and without axial deformities…………21

Figure 1-4. Additive effects of scoliosis and drought on survival time of Bd-exposed toadlets…………………………………………………………………………………...22

Figure 2-1. Observed and experimental diurnal temperature fluctuations………………37

Figure 2-2. Daily fluctuating temperature reduces Bd growth compared to growth at a stable optimal temperature……………………………………………………………….43

Figure 2-3. Zoospore production in Bd cultures grown at stable (7.5°C, 17.5°C, or

27.5°C) and fluctuating temperautres……………………………………………………45

Figure 2-4. Fluctuating and optimal temperature cultures do not differ in fecundity…...45

Figure 2-5. Differences in zoosporangia viability among Bd cultures grown at fluctuating and stable temperatures (7.5°C, 17.5°C, or 27.5°C)……………………………………...47 1

Introduction: A disease ecology approach to understanding drought and disease threats to a threatened alpine endemic, Anaxyrus canorus

Disease ecology incorporates the effect of the environment on host-pathogen interactions and disease dynamics (Demas and Nelson 2012). Disease ecologists address this complexity and the interaction between host physiology, pathogen prevalence and virulence, and environmental factors through the disease triangle (Fig. 1; McNew 1960;

Demas and Nelson 2012). The disease triangle framework is particularly applicable to amphibian hosts infected with Batrachochytrium dendrobatidis (Bd), the fungus responsible for the amphibian disease chytridiomycosis (Berger et al. 1998; Longcore et al. 1999). Chytridiomycosis has caused amphibian declines throughout the world and is considered one of the greatest disease threats to biodiversity (Skerratt et al. 2007; Wake and Vredenburg 2008). While the fungus is responsible for the declines of over 200 amphibian species (Wake and Vredenburg 2008), not all amphibians are susceptible

(Daszak et al. 2004; Kriger and Hero 2007). Intra- and interspecific differences in mortality from Bd have been associated with differences in host behavior and immune defenses, pathogen virulence, and environmental factors (Richards-Zawacki 2009;

Murphy et al. 2011; Savage and Zamudio 2011; Voyles et al. 2011).

Temperature and water availability are important environmental factors governing this host-pathogen system, as both amphibians and Bd are moisture-reliant and temperature-sensitive (Carey et al. 2003; Piotrowski et al. 2004). The disease ecology of

Bd is inextricably linked to its thermal environment, with a growth-limiting thermal maximum near 27°C (Voyles et al. 2012). Temperature can cause differences in Bd 2 growth patterns in culture and on amphibian hosts by changing the infectivity and virulence of Bd (Woodhams et al. 2008; Voyles et al. 2012, Stevenson et al. 2013; Daskin et al. 2011; Raffel et al. 2013; Greenspan et al. 2017). Decreases in water availability may impose physiological costs on amphibian hosts that increase risk of disease for certain life stages. Due to plasticity in development time, tadpoles inhabiting rapidly drying pools can metamorphose early to escape death by desiccation (Alford and Harris

1988; Newman 1992). However, individuals that avoid desiccation through accelerated development can incur the costs of smaller body sizes and decreased immune function, potentially leading to increases in susceptibility to disease post-metamorphosis (Denver et al. 1998; Rollins-Smith 1998; Gervasi et al. 2008; Garner et al. 2009; Blaustein et al.

2010). In these ways, temperature and water availability can change disease risk for amphibian hosts.

The Yosemite toad (Anaxyrus [Bufo] canorus) is a good model for examining the effects of both temperature and water availability on chytridiomycosis. The species utilizes habitats with large daily and seasonal fluctuations in both temperature and water availability, and their susceptibility to Bd is currently unknown (Mulllally 1953;

Karlstrom 1962; Brown et al. 2015). A. canorus lives exclusively in high elevation meadows (1,950-3,444 m) of the Sierra Nevada of central California where snowmelt runoff and spring recharge create important habitat for breeding and tadpole development

(Karlstrom 1962; Sherman and Morton 1993; Wang 2012). After overwintering in rodent burrows or other hibernacula, adult toads emerge in the summer after snowmelt to breed in flooded meadows, ephemeral pools, or shallow ponds which make up less than 3% of the landscape (Mullally 1953; Karlstrom 1962; Ratliff 1985; Fig. 2). Females lay strands 3 of 1,500-2000 eggs which take 40-60 days to reach metamorphosis in these shallow water bodies (Mullally 1953; Karlstrom 1962; Fig. 2). The shallow pools where A. canorus tadpoles develop undergo large daily temperature fluctuations. Some breeding pools reach daily maxima that exceed the thermal optimum for Bd (28-37°C daily maximum), while other pools fluctuate within the bounds of the Bd thermal range (5-27°C daily range; Maier 2018). In addition, climate change is expected to decrease seasonal water availability, or hydroperiod, in these habitats (Hamlet et al. 2007; PRBO 2011; Viers et al. 2013; Godsey et al. 2014; Ryan et al. 2014; Lee et al. 2015), with potential implications for disease dynamics (Parmesan 2006; Rohr and Raffel 2010; Rohr et al. 2011). As such, the temperature fluctuations and decreases in water availability in this system may shift disease outcomes by altering pathogen growth patterns or host susceptibility.

Figure 2. Yosemite toad life history. A) Eggs in an ephemeral pool. B) Tadpoles in an ephemeral pool. C) Recently metamorphosed juvenile (young of year). D) Subadult, 2-3 years old. E) Adult male, olive coloration. F) Adult female, mottled coloration (sexually dimorphic adults). G) Example of an ephemeral pool in Yosemite National Park, CA, USA. H) Young tadpoles in a rapidly drying habitat with water depth of 1cm. The remaining alive tadpoles (black) are too young to accelerate development and will perish without water influx from a storm. 4

Examining environmental drivers of disease in this host may aid in species recovery initiatives. A. canorus is declining in distribution and abundance (Drost and

Fellers 1994; Jennings and Hayes 1994; Brown and Olsen 2013) and is listed as

"Threatened" by the United States Fish and Wildlife Service (USFWS), "Endangered" by the International Union for the Conservation of Nature (IUCN), and a "Priority 1 Species of Special Concern" by the State of California (Thompson et al. 2016). Although major drivers of decline remain unknown, Bd and climate change, with drought in particular, are major threats to this species (USFWS 2014). A recent range-wide study showed that

Bd is widely distributed across the A. canorus range and is found on metamorphs, subadults and adults (Dodge et al. in prep). However, whether infection leads to development of lethal disease remains unknown for this species. To better understand disease dynamics in this system, it is important to understand if Bd causes direct mortality events in A. canorus populations, and if so, to what extent and under what environmental conditions.

Chapter 1: The effects of larval development under drought on Bd susceptibility post- metamorphosis. In chapter one, I tested the effects of environmental variability (i.e. water level variation) on disease across life stages. Specifically, I examined (1) the responses of A. canorus tadpoles to drought conditions, (2) the susceptibility of post- metamorphic A. canorus to Bd, and (3) the synergistic effects of drought and disease on post-metamorphic toad survival. I predicted that (1) tadpoles reared under experimentally-simulated drought conditions would metamorphose more rapidly and at smaller body sizes as compared to animals reared in tanks with stable water levels; (2) 5 juvenile toads infected with Bd would have higher mortality rates than control toads not exposed to the pathogen, and mortality would be associated with an increase in Bd infection intensity; and (3) tadpoles that developed under drought conditions would have higher Bd infection intensities and mortality rates post-metamorphosis than individuals that developed in a stable water level environment.

Chapter 2: The effect of daily fluctuating temperatures on Bd growth and reproduction rate in vitro. In chapter two, I tested Bd growth dynamics under a fluctuating temperature profile that simulates the natural diurnal temperature fluctuations that A. canorus are exposed to in wild ephemeral habitats. This thermal profile was used to investigate how daily fluctuating temperature affects Bd growth, fecundity, zoospore production, and time to zoospore production. I predicted that Bd grown at a daily fluctuating temperature profile with a daily maximum at the Bd thermal maximum would have a reduced growth rate and reduced zoospore production.

Figure 1. The disease triangle applied to A. canorus–Bd disease dynamics. A. canorus susceptibility to Bd is unknown in both controlled settings and in the wild. Decreases in hydroperiod may impose physiological costs on A. canorus, increasing risk of disease. Daily fluctuating temperatures may alter Bd growth, altering pathogen prevalence and/or virulence. Bd photo credit: Berger et al. 2005. 7

Chapter 1: The effects of larval development under drought on Bd susceptibility post-metamorphosis

Introduction

Disease ecology examines the independent and interactive effects of host, pathogen, and environmental factors on disease outcomes (Demas and Nelson 2012). A common framework for understanding these interactions is the disease triangle, a conceptual model that incorporates environmental factors into host-pathogen interactions

(McNew 1960; Scholthof 2007; Demas and Nelson, 2012). Prior to the disease triangle model, much disease research focused on simplified host-pathogen interactions without incorporating the effects of environmental factors on disease (Scholthof 2007). However, both host susceptibility and pathogen virulence can be influenced by the environment through temporal, spatial, and climatic variables (Demas and Nelson 2012). Changes in environment can alter host immune function, physiology, behavior, and life-history traits as well as pathogen reproduction rate, transmission rate, and evasion of host defenses, such that disease outcome can vary considerably depending on the environment (Demas and Nelson 2012).

The disease triangle is frequently applied to understand infectious disease in amphibians. As ectotherms that depend on water for breeding and larval development, amphibian physiology is inextricably tied to changes in the environment, particularly shifts in temperature and water availability (Carey and Alexander 2003). These environmental changes may alter disease outcomes for amphibian hosts (Parmesan 2006;

Alford et al. 2007; Rohr and Raffel 2010; Rohr et al. 2011). One disease that has 8 garnered a considerable amount of attention is chytridiomycosis, which is caused by the pathogenic fungus, Batrachochytrium dendrobatidis (Bd; Berger et al. 1998; Longcore et al. 1999). Chytridiomycosis has been associated with amphibian declines throughout the world and is considered one of the greatest disease threats to biodiversity (Skerratt et al.

2007; Wake and Vredenburg 2008).

The pathophysiology of chytridiomycosis differs across amphibian life stages. In adults, Bd infects keratinized tissues in the skin (Berger et al. 1998; Longcore et al. 1999;

Pessier et al. 1999) which damages the amphibian epidermis, leading to electrolyte imbalance and cardiac arrest (Voyles et al. 2009). In tadpoles, Bd infects the keratinized cells found in larval mouthparts (Fellers et al. 2001; Rachowicz and Vredenburg 2004;

Marantelli et al. 2004; Knapp and Morgan 2006). While damage of mouthparts from Bd infection can impair feeding and therefore growth in tadpoles, mortality from Bd infection is less likely during the larval stage as compared to highly susceptible post- metamorphic life stages (Bosch et al. 2001; Rachowicz and Vredenburg 2004; Garner et al. 2009).

Aquatic amphibian larvae are unlikely to develop lethal chytridiomycosis, but this life stage is prone to physiological costs and mortality associated with decreases in water availability that may result in downstream consequences for disease susceptibility (Alford et al. 2007; Gervasi and Foufopolous 2008; Crespi and Warne 2013). Tadpoles are vulnerable to desiccation and can die before reaching metamorphosis as water habitats evaporate (Newman 1992; Sherman and Morton 1993; Denver 2009). With predicted increases in temperature and frequency and intensity of drought associated with climate change, risk of death by desiccation may be amplified for amphibians in warming and 9 drying regions (McMenamin et al. 2008; Ryan et al. 2014; Brown et al. 2015). However, tadpoles of some species can metamorphose early and escape their drying habitat due to phenotypic plasticity in development time (Alford and Harris 1988; Newman 1992;

Denver 2009). While this shortened larval period increases survival probability, it may come with costs of smaller body sizes and decreased immune function that reduce fitness later in life relative to individuals that develop at a normal rate (Denver et al. 1998;

Rollins-Smith 1998; Altwegg and Reyer 2003; Gervasi et al. 2008; Garner et al. 2009;

Blaustein et al. 2010). Premature metamorphosis may increase susceptibility to pathogens, such as Bd, due to incomplete immune development, increased relative surface area for infection, altered sloughing rates, and differences in skin microbiome diversity (Rollins-Smith et al. 1988; Rollins-Smith 1998; Chammas et al. 2015; Bakar et al. 2016; Bates et al. 2018; Wu et al. 2018). The amphibian immune system also undergoes a substantial reconstruction and a brief period of immunosuppression during metamorphosis, leaving juveniles more vulnerable to disease (Rollins-Smith et al. 2011).

The potential fitness costs of drought-induced early metamorphosis paired with immune system reorganization and immunosuppression may increase disease risk for recently metamorphosed amphibians.

The Yosemite toad (Anaxyrus [Bufo] canorus) is an ideal species for examining the synergistic effects of reduced hydroperiod (i.e. seasonally available water level) during the larval stage and susceptibility to disease for multiple reasons. First, A. canorus tadpoles develop in shallow water bodies subject to seasonal drying (Cunningham 1963;

Sherman and Morton 1993; Brown et al. 2013). Species, such as ephemeral pool breeders, that utilize highly variable environments are more likely to express high levels 10 of plasticity (Bradshaw 1965; Van Buskirk 2002). Therefore, it is possible that A. canorus will exhibit some phenotypic plasticity in response to reduced hydroperiod.

Second, A. canorus is thought to be susceptible to Bd infection (Dodge et al. in prep). A recent study showed that Bd is widely distributed across the Yosemite toad’s range and is found on metamorphs, subadults, and adults (Dodge et al. in prep). However, Bd infection has not been connected to development of chytridiomycosis and subsequent mortality in this species. Third, A. canorus is currently listed as federally threatened, and understanding the roles of reduced hydroperiod and chytridiomycosis may improve understanding of species decline. Causes of declines in distribution and abundance are currently unknown for this species, (Drost and Fellers 1994; Jennings and Hayes 1994;

Brown and Olsen 2013), but chytridiomycosis and drought are considered serious threats to A. canorus (USFWS 2014; Brown et al. 2015). Investigating the potential additive effects of reduced hydroperiod on Bd susceptibility in A. canorus may help to inform recovery efforts for this species.

I hypothesized that larval development under a reduced hydroperiod would increase susceptibility to Bd post-metamorphosis. To examine how amphibians respond to drought and disease threats across life stages, I collected A. canorus eggs, reared tadpoles under simulated drought conditions, and infected metamorphosed toads with a standard dose of Bd known to cause mortality in previous inoculation experiments with other species (Carey et al. 2006; Voyles et al. 2009; Murphy et al. 2011). I predicted that

(1) tadpoles reared under experimentally-simulated drought conditions would metamorphose more rapidly and have smaller body sizes as compared to animals reared in tanks with stable water levels; (2) toads infected with Bd would have higher mortality 11 rates than control toads not exposed to the pathogen, and mortality would be associated with an increase in Bd infection intensity and clinical signs and symptoms of chytridiomycosis including a decrease in mass; and (3) toads that developed under drought conditions would have higher mortality rates than toads that developed in an environment with stable water levels. My results provide a first look at the interplay between disease and drought for A. canorus that improve our understanding of threats to this species.

Methods

Egg collection. I collected 200 live A. canorus eggs on 21 June 2017 from Kaiser

Meadow in the Sierra National Forest, Fresno County, California, USA (37.2967,

-119.1047). Kaiser Meadow has a large, stable, yearly breeding population with a history of long-term monitoring (pers. comm. Stephanie Barnes), and the removal of 200 eggs was unlikely to affect population persistence at this site. I collected 40 eggs from 5 distinct egg clutches of unknown parentage. Eggs are an ideal stage to collect because they are the least likely to carry Bd, and their removal is least likely to impact population dynamics given the high mortality of eggs and larvae in the wild (Sherman 1980;

Sherman and Morton 1993; Biek et al. 2002; Scherff-Norris et al. 2002; Sadinski 2004).

To ensure that all collected eggs were fertilized, I examined all eggs for signs of development and cleavage (Gosner 1960).

I transported eggs to the University of Nevada Reno following protocols outlined in Grasso et al. (2017). In brief, I transferred eggs to five disinfected containers filled with stream water from Kaiser Meadow (40 eggs per container), placed the egg 12 containers in a roto-molded cooler packed with bagged snow, and supplied aeration to each of the five egg containers. I kept the air temperatures within the cooler between 8°C and 11°C during. I allowed eggs to warm to 22°C upon arrival before transferring them to aquaria of the same water temperature.

Husbandry. During the egg and tadpole stages preceding initiation of drought treatments,

I housed animals at low densities in disinfected polycarbonate tanks (Cambro, 53cm L x

33cm W x 20cm H) filled to a depth of 14 cm with aged tap water on constant aeration.

This water depth falls within the range of pool depths used by A. canorus tadpoles in the wild and eliminated cannibalism by decreasing tadpole density. I placed tanks under

UVB lights on a 12:12 L/D cycle and maintained room temperature at 21-22°C. I measured water depth, temperature, and quality daily (see Husbandry Supplement) and recorded tadpole health and approximate tadpole development stage daily. I cleaned tanks and replaced tank water daily, treating all tanks equally. I fed tadpoles a diverse tadpole diet ad libitum (see Husbandry Supplement). Once tadpoles reached metamorphosis, I placed perches in the tanks to allow tadpoles nearing metamorphic climax to climb out of the water.

After completing metamorphosis, I housed juvenile toads separately in individual enclosures (17.3cm L x 12.2cm W x 5.6cm H) to track individual identity and to help control Bd zoospore concentration after exposure (Carey et al. 2006). I provided terrestrial and aquatic habitats in each enclosure to mimic A. canorus habitat needs in the wild by placing a damp paper towel along the enclosure floor, resting the enclosure on an incline, and allowing water to pool at one end. As Bd cannot survive without moisture, 13 paper towels were always kept damp. Before beginning the susceptibility trial, I allowed juveniles to acclimate to their new containers for a minimum of nine days. After the initiation of the susceptibility trial, I replaced aged tap water with Holtfretter's artificial pond water (20% Holtfretter’s solution: 250 mL, in mMol: 6.0 NaCl, 0.06 KCL, 0.09

CaCl2, 0.24 NaCO3; pH 7.0) in toadlet enclosures to control for potential effects of tap water on Bd infection (Voyles et al. 2009). I fed juveniles fruit flies coated in vitamin D3 and calcium (RepCal) and changed paper towels and Holtfretter’s solution every other day. I maintained all toads in individual containers under UVB light on a 12:12 L/D cycle in a room with air temperatures between 21-22°C, which falls within the Bd thermal optimum (Piotrowski et al. 2004; Voyles et al. 2012; Stevenson et al. 2013).

Larval response to decreased hydroperiod. To examine the effects of reduced hydroperiod on body size and time to metamorphosis, I removed water at fast or slow rates from designated treatment tanks to simulate pool drying and measured tadpole size and development on a weekly basis. After tadpoles reached a median Gosner stage of 36, I haphazardly assigned tadpoles (n=108) to one of three hydroperiod treatments, with six tadpoles per tank and six tanks per treatment. I included three hydroperiod treatments: severe drought (water depth lowered by 2 cm every day); mild drought (water depth lowered by 1cm every day); and a no drought control (water maintained at a constant depth of 14 cm). To avoid death by desiccation, I maintained a constant water depth of 2 cm in each tank if tadpoles had not started to metamorphose by the time a tank should be completely dry. 14

Before assignment to experimental tank and at one-week intervals after initiation of the drought treatment, I weighed individuals to the nearest 0.1g, measured snout-vent- length (SVL) to the nearest 0.1mm using calipers, and visually determined each tadpole’s development stage using the system of Gosner (1960). In addition, I measured individual mass and SVL at metamorphosis completion and recorded date of metamorphosis.

During development between Gosner stages 25-30, some tadpoles developed axial malformations including scoliosis (abnormal lateral curvature of the spine), kyphosis

(abnormal backward curvature of the spine), or kinked or S-shaped tails (Fig. 1-3, inset).

I noted individuals with severe axial deformities and incorporated this metric into my analyses. Of the 93 juveniles used in the susceptibility trial, 17 exhibited severe axial deformities. I used Pearson’s chi-squared to test if incidence of axial deformities was correlated with drought treatment.

Juvenile susceptibility to Bd. I used a Bd isolate collected from the Sierra Nevada,

MYLF-16343, that is likely genetically similar to Bd isolates found at the egg collection site (Morgan et al. 2007; Rosenblum et al. 2010). I revived an aliquot of the cryoarchived

MYLF 16343 culture on agar plates following a standard protocol (Boyle et al. 2004).

Once the cultures revived, I passaged Bd in TGhL liquid growth media (16 g tryptone,

4 g gelatin hydrolysate, 2 g lactose, in 1000 mL distilled water, autoclaved) in 75 cm2 tissue culture flasks (20mL TGhL, 4 mL Bd culture) at 18°C for 7-9 days until peak zoospore release (determined via inspection using an inverted microscope; Voyles 2011).

At peak zoospore release, I harvested Bd zoospores for the exposure inoculum by filtering liquid Bd cultures through sterile filter paper to remove sporangia (Voyles 2011). 15

Using a hemocytometer, I determined an inoculum zoospore concentration of 278 ± 21 x

104 zoospores per mL. I generated a negative control solution by using an equal volume of sterilized TGhL that did not contain Bd zoospores.

I randomly assigned toadlets within each drought treatment group to either exposed (Bd+) or unexposed (Bd-) groups. Before inoculation, I swabbed all toadlets for

Bd using a standardized swabbing procedure to determine if toadlets were free of Bd before the exposure (Boyle et al. 2004; Hyatt et al. 2007). I exposed 46 toads to Bd inoculum and 45 toads to a control solution of TGhL via immersion bath in small exposure containers for 20 hours (4.4cm diameter x 3cm H, 60mL capacity). I added 1 mL of exposure solution to 4 mL of Holtfretter's artificial pond water for a final zoospore concentration of approximately 5.6 x 105 zoospores per mL. After the 20-hour exposure period, I moved juveniles to cleaned and disinfected enclosures with 15 mL 20%

Holtfretter’s solution and terrestrial and aquatic conditions (as described above).

Over the course of the experiment, I collected diagnostic skin swabs, measured toadlet body size, and noted toadlet health every two weeks, and again when each individual toadlet died. I used the diagnostic skin swabs to estimate infection intensity using quantitative polymerase chain reaction (qPCR; Boyle et al. 2004). Swabs that returned a Bd-positive PCR result with a cycle threshold greater than 38 were assumed to be Bd-negative because positive PCR results at this cycle threshold represent presence of

0.1 zoospore equivalents and are likely due to contamination (Hyatt et al. 2007). I measured toadlet health using a qualitative scale between 0 and 3, where 0 represented a healthy individual and 3 represented an individual with severe signs and symptoms of chytridiomycosis (Voyles et al. 2009). When I found terminally ill individuals with a 16 health score of 3, I humanely euthanized animals with the chemical anesthetic MS-222

(Scherff-Norris et al. 2002; HACC ASIH, 2004). At time of death, I preserved specimens in 10% Neutral buffered formalin for deposit in educational collections.

Statistical analysis. I conducted all analyses using R v3.4.3 (R Core Team 2017). To test the difference in larval period, mass at metamorphosis, SVL at metamorphosis, and mass at Gosner stage 41 among different drought treatments, I used mixed model ANOVA with tank as a random variable using packages “lme4” v1.1.17 (Bates et al. 2015) and

“lmerTest” (Kuznetsova et al. 2017). Tank accounted for 0 variance in all four models. If mixed model ANOVAs suggested significant differences among drought treatments, I ran

Tukey’s post-hoc tests on mixed model ANOVA using the “multcomp” package v1.4.8

(Hothorn et al. 2008).

To test for differences in survival among Bd-exposed and Bd-negative control toadlets, I used a Kaplan-Meier estimator for survival curves and log-rank tests with a

Weibull distribution using the “survival” package v2.41.3 (Therneau 2015). I removed four toadlets that died during the 20-hour exposure period from all survival analyses, as their death was likely caused by drowning and was not attributed to chytridiomycosis. In addition to survival, I looked at changes in mass two weeks post exposure to Bd both within and among Bd treatment groups using paired t-tests and Welch’s t-tests respectively.

For Bd-exposed toadlets only, I tested differences in Bd load at time of death among drought treatments using ANOVA, and differences in Bd load at time of death among toadlets with and without axial deformities using a Wilcoxon signed-rank test. I 17

log10 transformed raw Bd zoospore equivalents to reduce skewness. I tested for singular effects of drought treatment and axial deformities on survival using Cox proportional hazards models with the “survival” package. In addition, I tested the potential role of drought, scoliosis, the interaction between drought and scoliosis, and age, mass, and SVL at initiation of the susceptibility trial on survival and risk of death. I used stepwise model selection with an AIC criterion to select the most parsimonious model with the best fit, followed by analysis of deviance to compare models using a three-leveled drought variable and a simplified drought variable that combined mild and severe drought treatment groups. I did not detect a difference between models using a three-level or binary drought treatment (Analysis of Deviance, χ2=1.457, df=1, p=0.2274), so I chose to use the simplified drought variable in multivariate analyses to improve sample size. All models met assumptions of proportional hazards.

Unless otherwise noted, summary statistics in figures and text represent mean + standard error (SE).

Results

Larval response to decreased hydroperiod. Tadpoles exposed to drought treatments metamorphosed earlier than control tadpoles in tanks where the water levels were held constant. Compared to tadpoles from control tanks, tadpoles from mild and severe drought tanks completed metamorphosis 2.55 + 0.61 and 2.88 + 0.60 days earlier, respectively (mild, t(91)= -4.15, p<0.001; severe, t(91)= -4.789, p<0.001; Fig. 1-1a). I did not detect a significant difference in time to metamorphosis between severe and mild drought treatment groups (Tukey’s, severe-control p<0.001, mild-control p<0.001, 18 severe-mild p=0.592; Fig. 1-1a). While a reduced hydroperiod accelerated the time to metamorphosis in A. canorus tadpoles, tadpole and metamorph body size did not differ at metamorphosis. There were no significant differences in snout-vent length (severe: t(91)= -

0.549, p=0.584; mild: t(91)= -0.477, p=0.635) or body mass (severe: t(91)= -0.709, p=0.480; mild: t(91)= -0.016, p=0.988) at time of metamorphosis between different drought treatment groups (Fig. 1-1), nor was there a difference in snout-vent length

(severe: t(79)=0.107, p=0.915; mild: t(91)= -0.968, p=0.336) or mass (severe: t(79)=0.367 , p=0.715; mild: t(91)=1.589 , p=0.116) between drought treatments at Gosner stage 41, the development stage at which tadpoles have invested most in growth.

Figure 1-1. Shortened hydroperiod decreases Yosemite toad larval period but does not affect size at metamorphosis. Blue bars represent tadpoles from control tanks (n=31) that experienced a constant water depth throughout development. Green bars represent tadpoles from mild drought tanks (n=30) that experienced a 1 cm reduction in daily water depth. Yellow bars represent tadpoles from severe drought treatments (n=33) that experienced a 2 cm reduction in daily water depth. Different letters represent statistically different groups, with asterisks represent p<0.001. Midline represents mean, boxes represent one standard error, and whiskers represent maximum and minimum values. 19

Juvenile susceptibility to Bd. Survival results were strikingly different between Bd- exposed and Bd-negative (control) toadlets (Kaplan-Meier, log-rank test, χ2=106, df=1, p<0.001; Fig. 1-2A). None of the 45 toadlets serving as Bd-negative controls developed signs and symptoms of chytridiomycosis, nor did they die over the four-week period of the susceptibility trial. In contrast, all 44 toadlets exposed to Bd became infected, developed clinical signs of severe chytridiomycosis, and died within 25 days of exposure to the pathogen (Fig. 1-2A). Progression of disease from onset of visible clinical signs to death was rapid, with individuals dying a mean of three days after first signs of lethargy or inappetence. One individual died within seven days of exposure with a load of 4.2 x

104 ZE (zoospore equivalents), and the remaining toadlets died within 11-25 days of exposure (median=15 days) with zoospore loads ranging from 1.5 x 105 to 2.8 x 107 ZE with a median zoospore equivalent of 7.5 x 106 ZE (Fig. 1-2B).

Figure 1-2. Survival and Bd load in juvenile A. canorus exposed to Bd inoculum or Bd-negative control solution. A) Bd-exposed toads (red solid, n=44) experienced 100% mortality 25 days after exposure, while the Bd-negative toad group (grey dashed, n=45) had zero mortality over the same four weeks. Kaplan-Meier, log-rank test, p<0.001. B) Mortality in Bd-exposed toads was associated with increases in Bd load over time, with Bd loads reaching a median 7.5 x 106 zoospore equivalents (ZE). Bd-positive qPCR results from skin swabs in Bd-negative control toads (grey), is likely due to contamination; Bd-negative control toads all had Bd-negative qPCR results at 4 weeks post initiation of the susceptibility trial. 20

Toadlets exposed to Bd that survived the first two weeks post exposure lost body mass during that time interval (paired t-test, t=4.95, df=36, p<0.001), while toadlets in the

Bd-negative control group neither gained nor lost mass during the first two weeks of the susceptibility trial (paired t-test, t=1.89, df=44, p=0.065). This difference in mass between Bd-exposed and Bd-negative control individuals is significant (Welch’s t-test, t=3.56, df=54, p<0.001), with Bd-exposed toadlets losing 0.04 + 0.0081g in two weeks, approximately 10% of their mass at the initiation of the susceptibility trial.

Quantitative PCR results suggest that some toadlets in the Bd-negative control group had Bd-positive qPCR results two weeks after initiation of the susceptibility trial, with a mean 471 + 92 ZE (Fig. 1-2B). However, all Bd-negative control toadlets had Bd- negative qPCR results four weeks post exposure at the termination of the susceptibility trial and at follow-up swabbing events over the following month. These Bd-positive qPCR results from Bd-negative control toadlets may be false positives for Bd infection due to contamination during swabbing or the qPCR processing of swabs. Alternatively, these results may represent true low infection loads of control toadlets, possibly resulting from contamination during husbandry.

Effects of axial deformity and larval development under drying conditions on Bd susceptibility. For toads exposed to Bd, I did not find a difference in survivorship (Wald test=1.52, df=2, p=0.469; Fig. 1-3A) or Bd load (ANOVA, F(2,41)=0.668, p=0.518) among mild, severe, and control drought treatments. I also did not find a singular effect of axial deformity on time to death (Wald test=2.2, df=1, p=0.138; Fig. 1-3B) or Bd load at death

(Wilcoxon rank sum, W=173, p=0.476). However, the additive effect of reduced 21 hydroperiod and development of severe axial deformities reduced survival for toadlets exposed to Bd (Wald test=6.14, df=2, p=0.046; Fig. 1-4). For individuals with axial malformations, risk of death was 2.17 + 0.39 times higher for juveniles that developed under drought conditions than in control drought tanks (z= 1.967, p=0.049). The additive cox proportional hazards model including drought and scoliosis terms was the most parsimonious and best fit using stepwise model selection with AIC criterion (Table 1-1).

Incidence of axial deformities was not related to drought treatment (χ2=2.27, df=4, p=0.685).

Figure 1-3. Differences in survival probability of Bd-exposed juveniles (A) among drought treatments and (B) between toads with and without axial deformities. A) Toads reared under control (blue solid, n=13), mild drought (green dashed, n=15), or severe drought (yellow dashed, n=16) water conditions did not differ in survival probability (p=0.469). B) Toads with axial deformities present (black, n=13) or absent (grey, n=31) did not differ significantly in survival probability (p=0.138). Inset: examples of a healthy tadpole and a tadpole with severe axial deformity.

Figure 1-4. Additive effects of scoliosis and drought on survival time of Bd-exposed toadlets. Individuals that developed under drought conditions and had severe axial deformities (red dashed, n=7) had a lower survival probability than individuals that developed under drought conditions but were not deformed (yellow dashed, n=24), or than individuals with (dark blue solid, n=7) or without (light blue solid, n=6) axial deformities that developed under stable water conditions (p=0.046).

Table 1-1. Cox proportional hazards model selection for survival of Bd-exposed toads using a stepwise approach with AIC. Predictor variables of scoliosis, mass, snout-vent length (SVL), and age refer to these measurements taken at day 0 of the susceptibility trial before Bd exposure.

Predictor variables AIC Drought:Scoliosis + Drought + Scoliosis + Mass + SVL + Age 252.9 Drought:Scoliosis + Drought + Scoliosis + SVL + Age 251.0 Drought + Scoliosis + SVL +Age 249.4 Drought + Scoliosis + SVL 249.2 Drought + Scoliosis 248.4 Drought 251.2 Scoliosis 250.6

Discussion

Changes in the environment can alter host morphology, physiology, and behavior

(Pigliucci 2001; Dewitt 2004). For amphibian hosts with distinct habitat needs across life stages, an environmental change encountered during the aquatic larval stage may alter disease outcomes in adulthood. In this study, I found that A. canorus juveniles are highly susceptible to Bd infection and develop lethal disease. Simulated drought conditions shortened time to metamorphosis but did not affect individual body size at metamorphosis or Bd infection load and Bd-implicated mortality. While drought did not alter susceptibility to Bd, the additive effect of axial deformities and development under drying conditions increased the risk of death. This result suggests that complex interactions and compounding stressors from a host’s environment may alter disease outcomes.

Larval response to decreased hydroperiod. Reduced hydroperiod accelerated time to metamorphosis by 2-3 days but did not affect body size during development or at completion of metamorphosis. As ephemeral pool breeders, A. canorus tadpoles may exhibit developmental plasticity in the wild resulting in earlier metamorphosis and smaller body size under drying conditions. A recent study suggests that A. canorus tadpoles in meadows with lower than median precipitation are smaller throughout development than tadpoles found in meadows with higher precipitation (Maier et al.

2016). If A. canorus do in fact show a decrease in size as a tradeoff with earlier development, I may not have observed this response due to the timing or intensity of prescribed drought treatments. I began drought treatments during prometamorphosis, a 24 larval phase during which tadpoles invest more in development than growth (Wilbur and

Collins 1973). Initiating drought treatments during premetamorphosis when tadpoles are growing exponentially may have a greater impact on tradeoffs between larval period and body size, while drying initiated during prometamorphosis may have a greater effect on larval period alone (Gervasi and Foufopoulos 2008). In addition, while A. canorus tadpoles can be found utilizing the margins of deep pools or occasionally lakes

(Karlstrom 1962; personal observation), they are most often found in shallow pools or flooded areas with a mean water depth of 4-6cm (Liang et al. 2017; personal observation), and it is not uncommon to observe Yosemite toad tadpoles in water depths of 1-2cm (Karlstrom 1962; Sherman 1980; personal observation). Given an ending depth of 2 cm in experimental tanks and a drought treatment initiated during prometamorphosis, the experimental drying regime may not may not have been severe enough to elicit a response for this species.

Selecting ecologically relevant timing and intensity of drying regimes is likely important in eliciting age-body size tradeoffs in controlled experiments. In addition, the effect of shortened hydroperiod on age and size at metamorphosis may be linked to species life history and breeding strategy, as species exposed to highly variable environments are more likely to express high levels of plasticity (Bradshaw 1965; Van

Buskirk 2002). Past studies present a range of outcomes resulting from experimentally reduced hydroperiod, including reduced time to and decreased size at metamorphosis

(Scaphiopus couchii, Newman 1989; S. hammondii, Denver 1998), reduced time to but no effect of size at metamorphosis (Anaxyrus americanus, Wilbur 1987; Rana sylvatica,

Gervasi and Foufopolous, 2008), or no effect on either time to or size at metamorphosis 25

(R. sylvatica, Wilbur 1987). These differences may be attributed to interspecific differences in plasticity range or may be due to experimental timing and severity of hydroperiod. I suggest follow-up experiments measuring pool depth and other factors

(e.g., predation, temperature, density, food availability; Altwegg and Reyer 2003; reviewed in Edge et al. 2016) in natural breeding pools to determine ecologically relevant drivers of phenotypic plasticity and growth-development tradeoffs.

While smaller body sizes have been associated with decreased survival and increased disease risk (Carey et al. 2006; Garner et al. 2009), other physiological costs, such as changes in immune and endocrine systems, may also be associated with earlier metamorphosis and altered susceptibility to disease. Individuals developing under drought conditions may have weak immune responses or elevated stress hormones irrespective of body size at metamorphosis (Gervasi and Foufopolous 2008; Crespi and

Warne 2013; Rollins-Smith 2017). Assessing immune function and glucocorticoid levels both during development under drought stress and during disease progression post exposure may help assess the role that larval stress and developmental programming play in disease susceptibility at later life stages.

Juvenile susceptibility to Bd. I found that A. canorus juveniles are highly susceptible to chytridiomycosis in a controlled setting. Juveniles exhibited clinical signs of disease, lost body mass over the course of disease progression, and carried high fungal loads consistent with lethal chytridiomycosis at time of death. Exposure to Bd led to 100% mortality within 25 days. The high susceptibility to Bd that I observed, paired with evidence of chytridiomycosis in museum specimens collected during the first witnessed 26 decline of A. canorus in the 1970s (Green and Sherman 2001), suggest that Bd played a role in historic declines of A. canorus and may still serve as a stressor or direct cause of mortality in wild populations.

The Bd-positive qPCR results from Bd-negative control toadlets two weeks into the susceptibility trial may be false positives for Bd infection due to contamination during swabbing, DNA extraction, or qPCR. Alternatively, if these results represent true low infection loads on control toadlets resulting from inadvertent exposure, A. canorus may be able to resist Bd infection if exposed to low zoospore loads. Future susceptibility trials exposing A. canorus juveniles to low Bd zoospore concentrations (i.e., using a dose- response approach; Carey et al. 2006), could resolve whether juvenile A. canorus can resist Bd infection at low exposure doses.

Juvenile A. canorus were highly susceptible to Bd in a controlled setting, but disease dynamics in the wild may be quite different for this species. Bd requires moisture and cool temperatures between approximately 2-27°C to grow and reproduce (Johnson et al. 2003; Piotrowski et al. 2004; Johnson and Speare 2005; Woodhams et al. 2008;

Stevenson et al. 2013; Voyles et al. 2017). Bd viability drops dramatically above 27°C

(Stevenson et al. 2013) and the fungus dies after eight days at 30°C in vitro (Piotrowski et al. 2004). While A. canorus spends most of its time operating within the Bd thermal optimum, adult body temperatures have been recorded well above the Bd thermal optimum (28-33°C; Mullally and Cunningham 1956; Cunningham 1963), suggesting that toads may be able to avoid or behaviorally regulate Bd infection by seeking out warmer microhabitats and maintaining high body temperatures (Richards-Zawacki 2009; Murphy et al. 2011; Hossack et al. 2013). Future studies tracking Bd prevalence and intensity on 27 individuals and populations over time in conjunction with measuring local temperatures and water availability will help define the roles that Bd infection and the environment play in A. canorus declines.

Effects of larval development under drying conditions on Bd susceptibility. While I observed an effect of drought treatment on time to metamorphosis, I did not find that reduced hydroperiod altered toadlet susceptibility to Bd. My results do not suggest a difference among drought treatments in toadlet survival, survival time, or Bd load at death. Reduced hydroperiod may not influence Bd susceptibility for this species, drought treatments may not have been severe enough to impose physiological costs, or high inoculation doses may have masked potential drought treatment effects. Lower inoculation doses paired with more extreme hydroperiod scenarios may tease apart the additive effects of drought and disease on survival rate.

Regardless of its effects on disease, drought is likely a direct cause of mortality for A. canorus tadpoles (Sherman 1980; Sherman and Morton 1993; Jennings and Hayes

1994; Brown et al. 2013). Hydroperiod of A. canorus breeding pools is likely to decrease due to a changing climate (IPCC 2014; Erwin 2009; Viers et al. 2013; Ryan et al. 2014;

Lee et al. 2015). Climate projections for this region include increased drought frequency and intensity, decreases in annual snowpack, earlier snow melt-out dates, and reduced water availability in late summer (Hamlet et al. 2007; PRBO 2011; Godsey et al. 2014).

This shift toward drier habitats may lead to increases in mortality rates due to larval death by desiccation. Although my study did not detect an effect of reduced hydroperiod on disease, A. canorus tadpoles in the wild may incur physiological or energetic costs 28 associated with developing under drying conditions. In addition to tracking the effects of drought on disease post-metamorphosis in the field under natural conditions, studies monitoring the effect of drought on larval mortality, recruitment, and population growth rates independent of disease will likely be important in understanding declines for this species.

Although reduced hydroperiod alone may not have changed disease outcomes for toads exposed to Bd, developing under drying conditions increased risk of death for toads with axial deformities. These results must be interpreted with caution due to low sample sizes but suggests that compounding stressors from a host’s environment can alter disease outcomes. While the mechanisms driving the axial deformities in my cohort of A. canorus are currently unknown, environmental factors including low oxygen, metals, pollutants, calcium deficiency, and UVB can trigger these malformations in laboratory studies (Hardy 1964; Worrest and Kimeldorf 1976; Cooke 1981; Hecnar 1995; Hays et al. 1996; Unrine et al. 2004; Budischak et al. 2009; Sayed et al. 2012). Agrochemicals and heavy metals have also been shown to alter immune function in some amphibians and increase infection (Gilbertson et al. 2003; Kiesecker 2002; Christin et al. 2003;

Christin et al. 2004). Scoliosis is rare in the wild (Reeves et al. 2010) and an unlikely factor exacerbating disease in natural systems. However, the additive effect of drought and deformity on risk of death illustrates that compounding costs incurred from changes in the environment can alter disease outcomes.

Conclusions and implications for conservation. In recent decades, A. canorus has experienced decreases in abundance at select sites and reductions in its distribution 29

(Sherman and Morton 1993; Jennings and Hayes, 1994; Drost and Fellers 1994; Green and Sherman 2001; Brown et al. 2012), leading to its current listing of threatened by the

United States Fish and Wildlife Service (USFWS) and endangered by the International

Union for the Conservation of Nature (IUCN). Despite efforts investigating invasive predators, pesticide use, and meadow grazing by pack stock and cattle as drivers of declines, no clear pattern has emerged (Sadinski 2004; Grasso et al. 2010; Roche et al.

2012; McIlroy et al. 2013).

Declines of A. canorus were first observed at a long-term study site near Tioga

Pass in Yosemite National Park, CA in the 1970s and 1980s, with declines attributed to drought and diseases including chytridiomycosis (Sherman and Morton 1993; Green and

Sherman 2001). The timing of the Tioga population’s decline coincides with the first observed declines for yellow-legged frogs (Rana muscosa and Rana sierrae), a high-

Sierra species complex that is sympatric with the A. canorus (Jennings 1996; Ouellet et al. 2005). Bd has caused severe declines of yellow-legged frogs (Briggs et al. 2005;

Rachowicz et al. 2006; Vredenburg et al. 2010), but A. canorus susceptibility to Bd has not been directly tested until now.

Understanding species-level susceptibility to Bd is important for conservation because not all amphibian species develop lethal Bd infections (Daszak et al. 2004;

Kriger and Hero 2007). My results show that A. canorus juveniles are susceptible to Bd and develop lethal chytridiomycosis in a controlled setting. This result suggests that chytridiomycosis could be a proximate cause of decline in this species. Understanding disease dynamics in the wild through long term disease monitoring of natural populations will be important to expand our knowledge of chytridiomycosis threats for A. canorus. 30

While reduced hydroperiod did not alter disease outcomes in this study, effects of drought on disease susceptibility and survival may be an important factor in the wild.

Individually, these challenges can threaten local populations, but additively, disease and climate change may exacerbate mortality events for certain life stages (Alford et al. 2007;

Hof et al. 2011; Rohr and Raffel 2010; Rohr et al. 2011). Continued research examining the independent and additive effects of reduced hydroperiod on disease outcomes and survival will be important, especially in the face of predicted increases in climate extremes.

Chapter 2: The effect of daily fluctuating temperatures on Bd growth and reproduction rate in vitro

Introduction

The environment can have profound effects on disease outcomes (Harvell et al.

2002; Altizer et al. 2006; Altizer et al. 2013). Temperature is an environmental factor that can alter disease due to the thermal sensitivity of host and pathogen physiological processes (Carey et al. 1999; Demas and Nelson 2012). Changes in environmental temperature can alter host immune function and, in turn, host susceptibility, especially in ectothermic hosts (Carey et al. 1999; Raffel et al. 2006; Rollins-Smith and Woodhams

2012; Greenspan et al. 2017). Viral, bacterial, fungal, and protozoan pathogens are also influenced by temperature because temperature can change growth patterns and reproductive rates of infective stages (Shapiro and Cowen 2012). By changing the rate of production of the transmissible stage of a pathogen (May and Anderson 1979), temperature may alter pathogenicity or intensity of infection (Woodhams et al. 2008).

Therefore, understanding pathogen growth and reproduction at different temperatures is important for predicting disease under varying thermal environments.

Temperature affects disease outcomes for amphibian hosts exposed to

Batrachochytrium dendrobatidis (Bd), a pathogenic fungus responsible for global amphibian declines (Woodhams et al. 2003; Lips et al. 2006; Richards-Zawacki 2009;

Rohr and Raffel 2010; Rowley and Alford 2013). Bd colonizes adult amphibian skin

(Berger et al. 1998; Longcore et al. 1999), and Bd growth and proliferation damage the epidermis, causing the disease chytridiomycosis (Voyles et al. 2009). Increases in Bd 32 load are correlated with the severity of chytridiomycosis, and in some species, Bd infection intensities must reach a threshold per host before causing mortality (Vredenburg et al. 2010; Voyles et al. 2012). Exposure to different temperatures affects Bd growth and reproduction (Woodhams et al. 2008; Voyles et al. 2012, Stevenson et al. 2013), such that temperature may alter the progression of disease by increasing or decreasing the time it takes for infection intensities to reach the threshold for host mortality.

The disease ecology of Bd is inextricably linked to its thermal environment

(Rollins-Smith and Woodhams 2012; Stevenson et al. 2013). Bd requires temperatures between approximately 2-27°C to grow and reproduce, with an isolate-dependent optimal temperature range falling between 15-25°C (Johnson et al. 2003; Piotrowski et al. 2004;

Johnson and Speare 2005; Woodhams et al. 2008; Stevenson et al. 2013; Voyles et al.

2017). Temperatures from 7-10°C, below the Bd thermal optimum, slow the Bd reproductive cycle but increase fecundity, allowing the pathogen to maintain high levels of virulence closer to its thermal minimum (Woodhams et al. 2008; Voyles et al. 2012).

At its upper thermal limits, Bd viability drops dramatically above 27°C in vitro, and the fungus dies after eight days at 30°C, four days at 32°C, and within four hours at 37°C

(Piotrowski et al. 2004; Stevenson et al. 2013).

The effect of temperature on Bd growth patterns may contribute to disease dynamics documented in wild amphibian populations (Berger et al. 2004; Daskin et al.

2011; Rowley and Alford 2013). Environmental temperatures that fall within the Bd thermal optimum (15°C-25°C) in tropical regions and within and below the thermal optimum in temperate regions are associated with high Bd prevalence and host mortality, while temperatures close to and above the Bd thermal maximum may provide refuge 33 from Bd infection (Woodhams et al. 2003; Lips et al. 2006; Knapp et al. 2011;

Puschendorf et al. 2011; Daskin et al. 2011). In nature, amphibian hosts that spend prolonged periods of time at temperatures above the Bd thermal maximum are less likely to be infected with Bd (Rowley and Alford 2013). In laboratory settings, frogs have cleared Bd infections within 16 hours at 37°C (Woodhams et al. 2003). Where climate permits, amphibians can avoid or behaviorally regulate Bd infection by seeking out these warmer temperatures, actively reducing Bd infection intensity and increasing host survival (Richards-Zawacki 2009; Murphy et al. 2011; Hossack et al. 2013).

To date, temperature studies have predominantly focused on host and pathogen responses at stable temperatures (Berger et al. 2004; Carey et al. 2006; Murphy et al.

2011). In contrast, our understanding of host-pathogen interactions at fluctuating temperatures is limited (Raffel et al. 2013; Greenspan et al. 2017). Expanding our knowledge of disease dynamics under varying temperatures is important because amphibians live in microhabitats with remarkable thermal heterogeneity across daily, seasonal, and annual cycles (Mullally and Cunningham 1956; Feder and Lynch 1982).

Both broad and fine scale temperature gradients across different spatial and temporal scales can alter host-pathogen interactions (Woodhams and Alford 2005; Kriger and

Hero, 2007; Lafferty 2009; Paaijmans et al. 2010; Puschendorf et al. 2011; Carrington et al. 2013; Sapsford et al 2013; Altizer et al. 2013). For amphibians exposed to Bd, daily heat pulses at 29°C reduce and even eliminate infection (Greenspan et al. 2017). In in vitro studies, brief freeze or heat shock treatments outside the Bd thermal range effectively decrease Bd growth rates (33°C, Daskin et al. 2011; -12°C and 28°C, Voyles 34 et al. 2017). These effects of fluctuating temperature on Bd growth may alter disease outcomes in amphibian hosts (Raffel et al. 2013; Greenspan et al. 2017).

In the Sierra Nevada, amphibian microhabitat temperatures frequently fall within the thermal limits for Bd growth and reproduction, leaving Sierra alpine amphibians vulnerable to Bd infection (Knapp et al. 2011). This includes the Yosemite toad

(Anaxyrus [Bufo] canorus), a threatened California endemic living exclusively in high elevation Sierra Nevada meadows (Karlstrom 1962; Brown et al. 2015). In the summer months, A. canorus utilize shallow pools and flooded meadows that undergo large daily temperature fluctuations (Mullally and Cunningham 1956; Cunningham 1963). While post-metamorphic A. canorus spend most of their time living within the Bd thermal optimum, both larval and post-metamorphic A. canorus also spend time in microclimates above the Bd thermal maximum (~30°C; Mullally and Cunningham 1956; Cunningham

1963). The thermal gradient that A. canorus are exposed to during summer months allows for the possibility of reducing or even eliminating Bd loads when individuals occupy microhabitats with temperatures above the Bd thermal maximum. It is unclear how these differences in daily fluctuating temperatures may alter Bd growth rate and fecundity, and in turn, A. canorus infection.

Growth rate and fecundity of Bd can be measured by quantifying growth and reproduction of the different Bd life stages. Bd has a complex life history with stationary and dispersing stages (Longcore et al. 1999; Berger et al. 2005), both of which are affected by temperature. Motile Bd zoospores encyst in keratinized amphibian tissues found in larval mouthparts and adult skin and develop into zoosporangia (Berger et al.

1998; Pessier et al. 1999; Rachowicz and Vredenburg 2004). Zoosporangia produce the 35 next generation of zoospores and release them into the environment or back onto the amphibian host (Longcore et al. 1999; Berger et al. 2005). The relationship between zoosporangia growth and zoospore production is temperature-dependent, and increases in zoospore production and zoospore longevity are not always proportional to increases in zoosporangia growth rate, particularly at temperatures below the Bd thermal optimum (7-

10°C; Woodhams et al. 2008; Voyles et al. 2012). Therefore, it is important to quantify differences in growth and reproduction of specific life stages of these pathogens to understand tradeoffs between growth and reproduction under different environmental conditions (Woodhams et al. 2008).

I hypothesized that exposure to a temperature profile that diurnally fluctuated between 7.5°C and 27.5°C would reduce Bd growth in vitro. Using temperature data collected from pools occupied by A. canorus tadpoles in the Sierra Nevada (Maier 2018),

I grew Bd at an ecologically relevant fluctuating temperature profile to examine how daily fluctuating temperature affects Bd growth rate, Bd fecundity, zoosporangia viability, zoospore production, and time to peak zoospore release. I used multiple growth assays to quantify the effects of fluctuating temperatures on distinct life stages of Bd because the relationship between zoosporangia growth rate and zoospore production rate can change with temperature (Woodhams et al. 2008; Voyles et al. 2012). I predicted that

Bd grown at a fluctuating temperature profile with a maximum just below the Bd thermal maximum would have a reduced growth rate, reduced fecundity, reduced zoosporangia viability, and reduced zoospore production as compared to Bd grown at a constant temperature within its thermal optimum. Examining growth patterns of Bd in vitro provides an initial step to look at the effects of fluctuating temperatures on pathogen 36 growth and reproduction. This approach extends our understanding of Bd disease dynamics under natural fluctuating temperatures in ephemeral habitats.

Methods

Incubation temperature selection. I defined my experimental fluctuating temperature profile using daily temperature ranges experienced by A. canorus tadpoles in natural breeding pools. I used ibutton (Thermochron) temperature data collected at ten different toad-occupied meadows representative of toad habitat in Yosemite National Park, CA

(data collected by P. Maier, as in Maier 2018). Ibuttons placed in ephemeral pools containing A. canorus tadpoles between 07 June and 26 June 2016 recorded temperature every two hours until collection between 18 July and 07 Aug 2016 (Maier 2018).

Many temperature profiles of A. canorus pools exceeded the Bd thermal maximum (Fig. 2-1A). While ephemeral habitats with thermal conditions above the Bd thermal maximum may or may not exclude Bd growth in the wild, these high temperatures (up to 37°C) are likely to kill Bd after short periods of time in a laboratory setting (Piotrowski et al. 2004; Woodhams et al. 2008; Stevenson et al. 2013). Therefore,

I selected a fluctuating temperature profile that does not exceed the Bd thermal maximum or minimum but fluctuates within these bounds between 7.5°C and 27.5°C (Fig. 2-1B). In addition to the fluctuating temperature profile, I measured Bd growth and viability at constant temperatures set at the daily high (27.5°C), the daily low (7.5°C), and the daily mean (17.5°C) temperatures of the fluctuating profile. These stable temperatures provide relevant growth comparisons of the pathogen at its thermal maximum (27.5°C), within its thermal optimum (17.5°C), and close to its thermal minimum (7.5°C) (Fig. 2-1B). 37

Figure 2-1. Observed and experimental diurnal temperature fluctuations. A) Water temperature over one 24-hour period of 10 different pools containing Yosemite toad tadpoles (grey lines; yellow line represents pool fluctuating within 27.5 and 7.5°C). Data from Maier 2018. B) Incubator temperature profiles over one 24-hour period. Fluctuating temperature = black; constant temperature set to daily thermal maximum (27.5°C) = red; constant temperature set to daily thermal minimum (7.5°C) = blue; constant temperature set to daily thermal mean (17.5°C) = green. Bd thermal optimum range (15-25°C; Stevenson et al. 2013) = green shaded band; Bd thermal tolerance (4-28°C) = grey shaded band.

Isolate selection, culturing, and plate set-up. I selected Sierra Nevada isolate MYLF-

16343, collected from mountain yellow-legged frog (Rana muscosa) skin and known to cause chytridiomycosis in juvenile A. canorus in a controlled laboratory environment (see

Chapter 1). This Bd isolate was collected and cryo-archived under known protocols

(Boyle et al. 2003) and was passaged 22 times after revival from cryo-archive. I centrifuged MYLF-16343 cultures to loosely pellet zoosporangia and dead cells and allow motile zoospores to swim into the supernatant. By drawing off motile zoospores from the supernatant for culture passage, I eliminated most of the cellular debris and sporangia from the previous culture and ensured that new cultures were seeded with live zoospores (Voyles 2011). I cultured Bd in TGhL liquid growth media in 75cm2 tissue culture flasks (20mL TGhL, 4mL Bd culture) at 18°C for 7-9 days until peak zoospore release (Voyles 2011). 38

At peak zoospore release, I filtered cultures to harvest zoospores for the experiment (Voyles 2011). I filtered cultures through sterilized filter paper, counted zoospores using a hemocytometer (Ultra Plane Improved Neubauer, Hausser Scientific), and diluted zoospores with fresh TGhL to a concentration of 46 x 104 zoospores per mL

(Voyles 2011). I then inoculated zoospores into 12 sterile, flat-bottom 96-well plates, assigning three plates to each temperature treatment. For experimental wells, I pipetted

50 μL of standardized zoospore culture and 50 μL TGhL for a final concentration of 23 x

104 zoospores per mL per well. For negative controls, I pipetted 50 μL of heat-killed standardized zoospore culture (10-minute incubation at 40°C) and 50 μL TGhL for a final concentration of 23 x 104 heat-killed zoospores per mL per well. Finally, I pipetted 100

μL TGhL into perimeter wells to reduce evaporation. I assigned plates to either 7.5°C,

17.5°C, 27.5°C, or fluctuating temperature incubators (Conviron A1000; Haier). I monitored incubator temperatures using ibuttons.

Quantification of Bd growth. I used three methods to quantify Bd growth over time: (1) optical density (OD), (2) motile zoospore counts, and (3) zoosporangia viability assays.

Optical density readings measure whole culture growth, motile zoospore counts track zoospore production and timing of release, and zoosporangia viability assays quantify sporangia growth. In addition, I looked at the ratio of zoospores to culture OD as a metric for culture fecundity.

Optical Density (OD). I measured OD of individual wells to capture Bd logistic growth over time. OD measurements are a fast, easy, and inexpensive way to quantify culture 39 growth. I measured OD of individual wells daily at 490nm (Biotek ELx800 Absorbance

Reader). Before reading OD, I removed condensation from 96-well plate lids with a

KimWipe or used a separate sterile lid for plate reading if the lid was scratched. After using a well for a zoospore count or zoosporangia viability assay, thus changing the contents of the well, I eliminated that well from future OD readings.

Zoospore production. I counted motile zoospores from individual wells to approximate zoospore viability over time. Due to the differences in life cycle times for Bd cultures incubating at different temperatures, I counted zoospores from plates incubated at 17.5°C,

27.5°C, and fluctuating temperatures every other day and from plates at 7.5°C once every four days. At each time point, I randomly selected six wells and counted motile zoospores using a hemocytometer (Ultra Plane Improved Neubauer, Hausser Scientific; Voyles

2011). After I used a well for zoospore counts, I did not resample it for OD, zoospore counts, or zoosporangia viability assays for the remainder of the experiment.

Zoosporangia viability assay. I used an MTT assay to measure zoosporangia metabolic activity and viability. The assay uses the yellow tetrazolium salt MTT (3-(4,5- dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide), which is reduced to purple

MTT-formazan crystals in metabolically active cells (Mosmann 1983; Liu et al. 1997).

The formazan crystals can be solubilized in various solvents or detergents, and the color change can be quantified by reading culture absorbance at 570nm, the most sensitive wavelength for this assay (Altman 1976). The MTT assay is a reliable and inexpensive colorimetric assay of cell viability used frequently in mammalian cell lines and 40 occasionally used to measure cell viability of select unicellular fungi (Levitz and

Diamond 1985; Freimoser et al. 1999).

I incubated randomly selected wells (Bd, n=8; heat-killed, n=3, TGhL, n=3) with

20 μL 5mg/mL MTT (Mosmann 1983; Hansen et al. 1989) for two hours in the dark and then solubilized the formazan crystals using 140 μL 20%SDS/50%DMF with gentle homogenization (Hansen et al.1989). After dissolving the formazan product, I read culture absorbance at 570nm (Biotek ELx800 Absorbance Reader).

Statistical analysis. I conducted all analyses using R v3.4.3 (R Core Team 2017). Unless otherwise noted, summary statistics in figures and text represent mean + standard error

(SE).

I used nonlinear mixed effects models to fit three-parameter logistic growth curves to OD measurements of cultures grown at different temperature profiles. Logistic growth curves are classically used to model the lag phase, exponential phase, and stationary phase of microbial growth (Madigan and Martinko 2006). Three-parameter logistic models can be used to predict the culture stationary phase (i.e. asymptote), the time at which cultures are in exponential growth phase halfway to stationary phase (i.e. inflection point), and a scale parameter that determines the steepness of the growth curve.

In addition, these models can be used to find the maximum growth rate of a culture by finding the slope at the inflection point. A nonlinear modeling approach allows for meaningful parameter estimates describing logistic growth in a nutrient-limited setting while accounting for possible correlation among wells over time. In addition, nonlinear 41 models are often more parsimonious and allow for more reliable predictions than an analogous polynomial linear model (Pinheiro and Bates 2000).

I fit three-parameter logistic growth curves to 27.5°C, 17.5°C, and fluctuating temperature culture OD measurements using nonlinear mixed effects models with the

‘nlme’ package v3.1.131.1 in R (Pinheiro and Bates 2018). I included temperature treatment fixed effects and plate random effects on all parameters (asymptote, inflection point, and scale parameter). I addressed within-group error heteroscedasticity by adding a power variance function structure to the model (Pinheiro and Bates 2000). I corrected for initial zoospore inoculation and media color by subtracting OD values of heat-killed controls from OD values of wells containing Bd.

Growth was slow and minimal at 7.5°C such that OD readings lacked sensitivity to detect increases in culture growth over time. The pattern of growth was linear as opposed to logistic, and nonlinear mixed effects models fit to a three-parameter logistic curve would not converge for OD readings from 7.5°C cultures. As such, I fit a linear model to OD measurements from cultures grown at 7.5°C.

I compared differences in peak zoospore production among temperature treatments using ANOVA and Tukey’s HSD post hoc tests. I square root transformed zoospore counts to achieve homogeneity of variance among temperature treatments.

I used a Kruskal-Wallis test to compare differences in sporangia viability at peak culture metabolic activity. I looked at pairwise comparisons between temperature treatments using a Conover-Iman post-hoc test with a Bonferroni correction. I used a nonparametric analysis because applied transformations could not correct for heterogeneity of variance at peak sporangia viability. 42

I approximated culture fecundity as a ratio of zoospores produced to mean optical density of culture per day. This metric provides a proxy for culture fecundity, approximating the number of zoospores produced relative to sporangia growth. I log- transformed this fecundity metric and added a correction factor of 1 to allow for log- transformation of values from wells with zero fecundity (no zoospores produced). I compared differences in culture fecundity at peak zoospore production using ANOVA and Tukey’s HSD post-hoc tests. I back-transformed percent differences in fecundity reported in the results.

Results

Optical density growth measurements. Total culture growth measured by well optical density (OD490) differed among temperature treatments (Fig. 2-2). By fitting nonlinear mixed effects models to OD490 data for cultures grown at 17.5°C, 27.5°C, and fluctuating temperatures, I found that growth in each culture followed a three-parameter logistic growth curve, and that temperature altered Bd culture carrying capacity during stationary phase (asymptote; p<0.001), time to stationary phase (inflection point, p<0.001), and growth curve scale (scale; p<0.001; Fig. 2-2). The fitted model suggested that 17.5°C cultures had the highest maximum growth, reached stationary phase after 15 days, and achieved the fastest growth rate during the exponential growth phase of all temperature treatments (Table 2-1). Compared to cultures grown at 17.5°C, Bd grown at fluctuating temperatures had a lower maximum growth, took longer to reach carrying capacity, and had a lower maximum growth rate (Table 2-1). Cultures grown at 27.5°C had the lowest growth rate and lowest carrying capacity yet reached maximum growth faster than 43

optimal or fluctuating temperature cultures. Analysis of OD490 readings for 7.5°C cultures suggested little or no growth rather than logistic growth over time (F = 3.15, df = 13, intercept = 0.0012, p = 0.10).

Figure 2-2. Daily fluctuating temperature reduces Bd growth compared to growth at a stable optimal temperature. Logistic growth curves (solid lines) and maximum growth rate (dashed lines) fit to daily optical density measurements of Bd grown at 17.5°C (green), 27.5°C (red), and fluctuating (black) temperatures. OD readings of cultures grown at 7.5°C (blue) did not follow a logistic growth curve. Points and error bars represent mean + standard deviation. Sample size decreases with time as wells were sacrificed for zoosporangia viability assays or zoospore counts. Day 0, n=120. Day 14, n=8.

Table 2-1. Parameter estimates for logistic growth models of Bd grown at stable (17.5°C and 27.5°C) and fluctuating temperatures (7.5°C – 27.5°C, “Flux”). Temp Max. Asymptote p-value Inflection p-value Scale p-value growth (+SE) point (+SE) (+SE) rate 17.5°C 0.036 0.152+0.01 <0.001 7.60+0.23 <0.001 1.07+0.07 <0.001

Flux 0.015 0.103+0.01 <0.001 8.71+0.51 0.0378 1.67+0.15 <0.001

27.5°C 0.005 0.028+0.01 <0.001 6.49+0.54 0.0489 1.45+0.19 0.053 44

Zoospore production. Bd cultures grown at 7.5°C, 17.5°C, 27.5°C, or a daily fluctuating temperature profile differed in time to and productivity at peak zoospore release (Fig. 2-

3; ANOVA, df=3, F=42.2, p<0.001). Between-group comparisons suggest that peak zoospore production in cultures grown at 17.5°C was higher than in cultures grown at any other temperature profile (Tukey HSD, p<0.001 for all pairwise comparisons with

17.5°C). The cultures grown at 17.5°C reached a mean maximum zoospore production of

20.7 + 3.0 x 104 zoospores/mL on day 10 with a zoospore burst beginning on day 6 and lasting until day 10. Zoospore production reached a mean maximum of 7.3 + 1.2 x 104 zoospores/mL in fluctuating temperature cultures on days 8-10. Peak zoospore release occurred over the same period, from days 6 to 10, in fluctuating temperature and 17.5°C cultures, but the mean maximum zoospore production was lower in fluctuating cultures than in 17.5°C cultures (Tukey HSD, p<0.001). Cultures grown at 7.5°C took the longest to reach peak zoospore production, with a mean maximum of 7.0 + 1.0 x 104 zoospores per mL on day 14. Although there was a difference in timing to zoospore peak, there was no difference in maximum zoospore production between Bd cultures grown at 7.5°C and at fluctuating temperatures (Tukey HSD, p=0.99). I did not observe new zoospore production in cultures grown at 27.5°C. Zoospore counts reached a mean maximum of

0.83 + 0.2 x 104 zoospores/mL on day 4 at 27.5°C; these zoospores are likely remnants from the initial inoculation on day 0.

Culture fecundity. I measured culture fecundity as the ratio of motile zoospores to total culture growth per day. I found that fecundity varied among different temperature 45

Figure 2-3. Zoospore production in Bd cultures grown at stable (7.5°C, 17.5°C, or 27.5°C) and fluctuating temperautres. Cultures grown at 17.5°C (green) had the highest maximum zoospore production (ANOVA, p<0.001). Cultures grown at 7.5°C (blue) had similar maximum zoospore production as cultures grown under fluctuating temperatures (black) but reached peak production later. Cultures grown at 27.5°C (red) did not produce zoospores. Points represent mean counts of motile zoospores (n=6), error bars represent standard error. Letters indicate signifcant differences among temperature treatments at peak zoospore release (Tukey HSD).

Figure 2-4. Fluctuating and optimal temperature cultures do not differ in fecundity. Culture fecundity is measured as the ratio of motile zoospores to total culture growth (OD) per day. Despite low OD readings, cultures grown at 7.5°C (blue, solid) had higher fecundity than cultures grown at 17.5°C (green, solid) or under fluctuating temperatures (black, solid; ANOVA, p<0.001). Cultures grown at 27.5°C did not produce zoospores. Dashed vertical lines correspond to time of peak zoospore production in fluctuating (black, day 8), optimal (green, day 10), and cold (blue, day 14) cultures. Points and error bars represent mean + standard error. Letters indicate signifcant differences among temperature treatments at peak zoospore release (Tukey HSD). 46 treatments (ANOVA, F=164.2, df=2, p<0.001), with cultures grown at 7.5°C having higher fecundity than cultures grown at 17.5°C or fluctuating temperatures (Fig. 2-4). At peak zoospore production, cultures grown at 7.5°C were 26% (15-43%) more productive per unit culture growth (OD490) than cultures grown at 17.5°C temperatures (Tukey’s

HSD, p<0.001), and 23% (14-39%) more productive than cultures grown at fluctuating temperatures (Tukey’s HSD, p<0.001). There was no difference in fecundity at peak zoospore production between cultures grown at 17.5°C and fluctuating temperatures

(Tukey’s HSD, p=0.87). Cultures grown at 27.5°C did not produce zoospores and therefore had a fecundity of zero.

Zoosporangia viability. I used an MTT assay to quantify differences in culture metabolic activity over time among temperature treatments. Temperature altered culture metabolic activity, with differences in maximum metabolic activity among all temperature treatments (Fig. 2-5; Kruskal-Wallis, χ2=28.75, df = 3, p < 0.001; Conover-Iman, p<0.001 for all pairwise temperature comparisons). Metabolic activity was highest in cultures grown at 17.5°C (Conover-Iman, p<0.001), with metabolic activity increasing rapidly on day 10 and peaking on day 14 (OD570 = 0.612 + 0.045). Cultures grown at fluctuating temperatures also showed a rapid increase in metabolic activity on day 10

(OD570 = 0.323 + 0.009) with a plateaued peak through day 14, but fluctuating temperature cultures never surpassed optimal temperature cultures in metabolic activity

(Conover-Iman, p<0.001). Peak metabolic activity was lower in cultures grown at cold temperatures (OD570 = 0.087 + 0.012) than in cultures grown at 17.5°C and at fluctuating temperatures, and reached maximum metabolic activity on day 16 well after 17.5°C and 47 fluctuating temperature cultures. Although zoosporangia grown at 27.5°C failed to produce zoospores, zoosporangia remained metabolically active at 27.5°C for 14 days and reached peak metabolic activity (OD570 = 0.045+ 0.001) on day 10.

Patterns in peak metabolic activity over time mimic patterns in zoospore production for 7.5°C, 17.5°C, and fluctuating temperature cultures, but the increases in metabolic activity lag behind peak zoospore production by two to four days. This result is likely due to the MTT assay detecting newly encysted zoosporangia resulting from the zoospore burst.

Figure 2-5. Differences in zoosporangia viability among Bd cultures grown at fluctuating and stable temperatures (7.5°C, 17.5°C, or 27.5°C). Cell viability was measured using a the colorimetric MTT assay that stains metabolically active zoosporangia purple (inset). Cultures grown under fluctuating temperatures (black) had lower zoosporangia viability than cultures grown 17.5°C (green) but higher viability than cultures grown at 7.5°C (blue) or at 27.5°C (red; Kruskal-Wallis, p<0.001). Points and error bars represent mean + standard deviation, n=8 per temperature per day. Letters indicate signifcant differences (Conover-Iman).

Discussion

Temperature fluctuations at different temporal scales can alter the thermal performance of an organism and in turn disease dynamics (Fargues and Luz 2000;

Paaijmins et al. 2010; Lambrechts et al. 2011; Carrington et al. 2013; Raffel et al. 2013;

Vasseur et al. 2014). Diurnal temperature fluctuations associated with shallow water body habitats may alter growth patterns of Bd with potential implications for amphibian disease. I examined the effect of an ecologically relevant daily temperature fluctuation on the growth of the amphibian-specific pathogenic fungus, Bd. My results suggest that daily temperature fluctuations reduce Bd growth in vitro, which may affect disease outcomes in wild amphibian populations.

When compared with Bd grown at an optimal temperature of 17.5°C, cultures grown at fluctuating temperatures exhibited a decrease in maximum culture growth and growth rate and a decrease in zoospore production at peak zoospore release. Since increases in Bd load are correlated with chytridiomycosis and resulting mortality

(Vredenburg et al. 2010; Voyles et al. 2012), reductions in Bd growth and zoospore production rates under daily fluctuating temperatures may slow disease progression in amphibian hosts experiencing the same fluctuating temperature regime.

Examining both the zoosporangia and zoospore life stages of Bd over time offered additional insights into the effects of fluctuating temperature on pathogen growth. For example, cultures grown at fluctuating temperatures and at 17.5°C took the same number of days to complete one reproductive cycle and had similar fecundities, but overall Bd growth (measured by OD) was reduced for cultures grown at fluctuating temperatures as compared to cultures grown at the Bd thermal optimum. These results suggest that the 49 decrease in zoospore production of Bd grown at fluctuating temperatures is likely due to a decrease in successful zoospore encystation. The possibility of temperature-induced reductions in encystation is further supported by the zoosporangia viability assays. After peak zoospore release on days 6 through 10, cultures grown at 17.5°C show an increase in zoosporangia metabolic activity, reflecting successful encystation of the second generation of zoospores. However, cultures grown under fluctuating temperatures show lower zoosporangia metabolic activity after the period of peak zoospore release, suggesting that the second generation of zoospores grown under fluctuating temperatures were not as successful at encysting and forming new zoosporangia as those grown at an optimal stable temperature of 17.5°C. This pattern of reduced encystation suggests that daily fluctuating temperatures may reduce Bd infectivity by reducing the number of viable zoospores. These results also highlight the advantage of quantifying growth patterns of both zoosporangia and zoospore life stages to form a more complete understanding of the effects of environmental factors on pathogen reproduction.

Results from Bd grown at stable temperatures indicate that growth and fecundity patterns were consistent with past research (Voyles et al. 2012; Voyles et al. 2017).

Cultures grown at 7.5°C had lower overall growth and a longer reproductive cycle but the highest fecundity of all temperature treatments as measured by the ratio of zoospores to culture OD. These characteristics of Bd growth below the Bd thermal optimum, along with increases in zoospore longevity, have been observed in other studies (Woodhams et al. 2008; Voyles et al. 2012; Stevenson et al. 2013). Cultures grown at 27.5°C failed to produce a second generation of zoospores, but zoosporangia viability assays showed that zoosporangia grown at 27.5°C remained metabolically active for 14 days. This finding 50 adds to research suggesting that Bd can survive short periods above its thermal maximum and in some cases resume growth after a return to an optimal temperature range

(Longcore et al. 1999; Piotrowski et al. 2004; Daskin et al. 2011; Stevenson et al. 2013;

Voyles et al. 2017).

The diurnal fluctuating temperature treatment decreased zoosporangia growth and zoospore production as compared to cultures grown within the Bd thermal optimum.

Future studies may help to pinpoint what specific aspect of the fluctuating temperature profile reduced Bd growth and reproduction. The reductions in Bd growth in vitro under a daily fluctuating temperature profile could be attributed to (1) the degree of difference between the daily temperature maximum and the Bd thermal maximum, (2) the length of time Bd spends above the thermal optimum, or (3) exposure to a novel thermal environment after passage in a stable and optimal thermal environment.

Bd growth and productivity under fluctuating temperatures are likely affected by both the amount of time in a diurnal cycle that Bd is exposed to temperatures above its thermal optimum, and by the degree of difference between the daily temperature maximum and the Bd thermal optimum (Greenspan et al. 2017). Temperature fitness curves are asymmetric, with a gradual rise from the thermal minimum to the optimal temperature and an abrupt drop in performance as temperatures rise above the thermal optimum (Huey and Stevenson 1979; Martin and Huey 2008). Therefore, exposure to environmental temperatures multiple degrees above an organism’s thermal optimum is more likely to decrease performance and fitness as compared to a smaller degree increase in temperature above the thermal optimum (Martin and Huey 2008; Amarasekare and

Savage 2012). This result has been demonstrated for Bd in culture, with Bd showing 51 reduced growth at daily one-hour periods at 33°C (Daskin et al. 2011), daily four-hour periods at 29°C (Greenspan et al. 2017), and perishing after four hours at 37°C (Johnson et al. 2003). While Bd growth and productivity were reduced when grown in vitro under a temperature regime fluctuating between 7.5°C and 27.5°C, Bd growth rates and reproduction are likely to decrease further if the pathogen is exposed to fluctuating temperatures with a higher daily maximum and prolonged exposure to temperatures above the pathogen’s thermal optimum.

Exposure to a novel thermal environment can also alter growth patterns of Bd

(Voyles et al. 2012). However, previous experiments show that Bd can respond to selective pressures from different thermal environments over time (Voyles et al. 2012) and can grow faster under predictable rather than stochastic temperature fluctuations

(Raffel et al. 2013). Bd exposed to a consistent and predictable daily fluctuating temperature profile may increase growth or fecundity across multiple generations as the pathogen adapts to its new thermal environment. While pools utilized by A. canorus in the wild experience large daily temperature ranges, the pattern of diurnal temperature fluctuation is largely predictable in summer months (Maier 2018), allowing for the possibility of Bd to adapt to diurnal temperature fluctuations if the daily maximum isn’t prohibitive for reproduction. Comparing Bd cultures serially passaged under stable optimal and consistent fluctuating temperatures to track pathogen evolution over time could elucidate whether temperature fluctuations or simply novel thermal environments are what drive reductions in Bd growth in a controlled setting.

Exposure to daily fluctuating water temperatures with daily maxima above the Bd thermal maximum may alter disease dynamics in the wild (Richards-Zawacki 2009; 52

Rowley and Alford 2013; Greenspan et al. 2017). Pools occupied by Yosemite toad tadpoles are often warmed above 27.5°C for prolonged daily periods, up to seven hours a day, and in some instances reach temperatures of 37°C (Maier 2018). Tadpoles have even shown preference for daytime pool temperatures near 33°C, and adults have been found with body temperatures at 28-33°C (Mullally 1953; Mullally and Cunningham 1956;

Cunningham 1963). Therefore, pool temperatures that exceed the Bd thermal maximum daily may reduce or in some cases eliminate Bd growth, and in turn, alter infection and disease development in amphibian hosts. Under projected temperature increases for the

Sierra Nevada (Viers et al. 2013), daily temperature maxima of shallow water bodies may provide refuge from Bd infection for amphibians.

Environmental daily temperature maxima and the predictability of daily temperature fluctuations are important drivers of disease for amphibians exposed to Bd

(Raffel et al. 2006; Rohr and Raffel 2010; Raffel et al. 2013; Greenspan et al. 2017).

Understanding pathogen growth in vitro under fluctuating temperatures provides an important first step in predicting disease outcomes under more realistic thermal environments (Voyles et al. 2017). Important next steps include investigating how A. canorus utilizes its thermal environment in regard to disease, and how daily fluctuating temperatures impact disease in vivo. Bd growth patterns can differ on amphibian hosts and in culture because the pathogen must cope with host behavioral thermoregulation, host microbiome, and host immune defenses in vivo (Woodhams et al. 2005; Woodhams et al. 2007; Richards-Zawacki 2009; Myers et al. 2012; Rowley and Alford 2013). These differences in pathogen growth in vivo and in vitro can be affected by fluctuating temperatures, as fluctuating temperatures can alter both pathogen growth and amphibian 53 immune responses (Plytycz and Jozkowicz 1994; Raffel et al. 2006; Raffel et al. 2013; reviewed in Rollins-Smith and Woodhams 2012; Cohen et al. 2017). Further investigations into the role of realistic temperature fluctuations in relation to the thermal performance of both host and pathogen will improve understanding of disease outcomes under natural thermal environments and better help to predict disease outcomes for imperiled amphibians.

Husbandry Supplement

Survival and mortality. Past researchers have had difficulty rearing Yosemite toads from egg or tadpole stages through completion of metamorphosis (pers.comm. R. Grasso and

C. Dodge). Considering past difficulties with this species, husbandry measures associated with Chapter 1 were largely successful. I had 75% survival during the egg stage, 89% survival during the tadpole stage, and 89% survival during the metamorph stage. Overall,

I had 60% survival from egg collection to successful metamorphosis and 79% survival from hatching to successful metamorphosis.

Egg mortality. Between June 22, 2017 and June 26, 2017, I observed mortality in

50 eggs, likely from a Saprolegnia fungus (water mold) commonly found on

Yosemite toad eggs in the wild (personal observation). The eggs were surrounded

in white cottony growth, characteristic of Saprolegnia colonization.

Tadpole mortality. Between June 27, 2017 and July 27, 2017, I counted mortality

in 16 tadpoles. One tadpole death was a confirmed cannibalism. The remaining 15

died of unknown causes, likely due to poor development.

Metamorph mortality. Between July 29, 2017 and August 15, 2017, I noted

mortality in 15 metamorphs. Metamorphosis is a difficult transition for

amphibians and is often associated with some mortality. Cause of death was

unknown for all metamorphs, although drowning was a likely cause of death for 55

some individuals. Incidence of drowning may be decreased in the future by

providing more or easier access to dry or moist habitat during metamorphosis.

Juvenile mortality - Susceptibility Trial. During the susceptibility trial from

August 20, 2017 to September 12, 2017, 44 juvenile toads from the group

exposed to Bd died from chytridiomycosis. No toads died from the Bd-negative

experimental control group during this time, and all appeared healthy.

Juvenile mortality – Post-susceptibility Trial. Beginning on September 25, 2017,

and continuing through the end of October, juvenile toads died at a rate of about

one a day, cause unknown. These animals were Bd-negative at time of death

(qPCR results). The remaining juvenile toad died on May 15, 2018, cause

unknown. Preserved specimens can be examined for causes of death.

Table S1. Yosemite toad mortality by life stage and experimental stage with presumed cause of death.

Date Life Stage Experimental Individuals Individuals Cause of death Stage Dead Remaining 6/21/2017 Egg Collection 0 200 NA 6/22/2017 – Egg Rearing 50 150 Saprolegnia 6/26/2017 6/27/2017 – Tadpole Rearing 16 134 Unknown; 7/27/2017 development 7/29/2017 – Metamorph Drought 15 119 Unknown; drowning, 8/15/2017 Treatment development 8/19/2017 – Juvenile Bd Exposure 4 115 Drowning 8/20/2017 8/20/2017 – Juvenile Bd 44 71 Chytridiomycosis 9/12/2017 Susceptibility Trial 9/25/2017 – Juvenile Rearing 70 1 Unknown 10/25/2017 5/15/2018 Juvenile Rearing 1 0 Unknown 56

Axial deformities. Many of the tadpoles I reared experienced some form of scoliosis or crooked or kinked tail. Causes of axial deformities in our tadpoles are currently unknown.

Although axial malformations can happen in wild populations, I believe that the abnormalities I observed in the tadpoles was due to husbandry measures. I speculate that water quality parameters caused the axial deformities. Possible guesses include water that was too soft (lacking in calcium and magnesium), the presence of fluoride or contaminants in unfiltered aged tap water, and the lack of a flow-through filter system with biological, chemical, and physical filters to improve aged tap water quality.

Tadpoles appeared healthy by appearance and behavior through 8 July 2017, approximately two weeks after hatching.

Water quality. Tank water quality parameters fell within suggested ranges for tadpoles, except for water hardness (Poole and Grow 2012). Nitrates, nitrites, and chlorine routinely read at 0ppm, alkalinity from 20-40ppm, hardness from 50-75ppm, and pH from 6.8-7.2. At 50ppm, water hardness was initially lower than the suggested range of

75-100ppm (Poole and Grow 2012). After tadpoles started to develop axial deformities, I added a Ca;Mg solution (6:7 ratio by weight; Odum and Zippel 2011) to the water with each water change to increase the water hardness to 75ppm. I attempted to keep ammonia levels below 0.5 at all times; in the few instances when ammonia levels exceeded 0.5ppm per tank, they never exceed 1ppm, an ammonia level that should not be exceeded for amphibian husbandry. Tank water temperatures remained around 20°C for the entire experiment.

Diet. I provided tadpoles with a diverse diet of Sera Micron, Tadpole Powder, Mazuri gel, Tadpole Smoothie, and romaine lettuce (Scherff-Norris et al. 2002; Ted Smith,

Colorado Parks & Wildlife Native Aquatic Species Restoration Facility, pers.comm;

Jessie Bushell, San Francisco Zoo, pers.comm). Tadpoles ate all food items, with less interest in romaine lettuce. In future experiments, I suggest a less complex diet for ease of husbandry purposes. Given the success of a diet composed of Sera Micron and an amphibian gel (Repashy or Mazuri brands) for Yosemite toad tadpoles and other Sierra amphibian species tadpoles at the San Francisco Zoo (pers.comm. Jessie Bushell), I recommend feeding Yosemite toad tadpoles these food items.

Tadpole Powder

Grind Hikari algae wafers, spirulina aquarium flake food, and shrimp flakes in a

coffee grinder in approximately equal amounts by mass. Store in a closed

container in the refrigerator.

Mazuri Gel Strips

Use approximately 50:50 ratio by weight of Mazuri dry gel to hot (>200°F) water

(eg., 250g of gel to 250mL water). Stir with a spoon and pour to a depth of one

inch in a flat pan. Chill overnight in a refrigerator. Cut gel into desired mass cubes

or strips. Coat each strip with Tadpole Powder and place strips in tank. Store

unused gel covered in a refrigerator for no more than two weeks or cut into

desired mass pieces and freeze. Note: The Boreal Toad Husbandry Manual

(Scherff-Norris et al. 2002) suggests feeding a 1”x1”x0.25” strip for every 3 58

Anaxyrus [Bufo] boreas tadpoles. I found that Yosemite toad tadpoles largely ignored these strips, and the large strips fouled the water quickly, so I decreased

Mazuri Gel Strip quantities per tank.

Tadpole Smoothie

150 g Collard greens

150 g Mustard greens (use turnip greens if not available)

300 g Yellow Squash

300 g Green zucchini

300-500 mL water

Weigh dry vegetables, blend thoroughly in blender. Add only as much water as needed to blend vegetables. Fill silicon mini ice cube trays with smoothie and freeze. Small cubes = 1.8g.

References

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