DETERMINING HOW ENVIRONMENTAL CHANGES IMPACT GROWTH OF BATRACHOCHYTRIUM DENDROBATIDIS USING A NOVEL IN VITRO SYSTEM By Amanda D. Layden, B.S. East Stroudsburg University of Pennsylvania A Thesis Submitted in Partial Fulfillment of the Requirements for the Degree of Master of Science in Biology to the office of Graduate and Extended Studies of East Stroudsburg University of Pennsylvania May 8, 2020 SIGNATURE/APPROVAL PAGE The signed approval page for this thesis was intentionally removed from the online copy by an authorized administrator at Kemp Library. The final approved signature page for this thesis is on file with the Office of Graduate and Extended Studies. Please contact Theses@esu.edu with any questions. ABSTRACT A Thesis Submitted in Partial Fulfillment of the Requirements for the Degree of Master of Science in Biology to the Office of Graduate and Extended Studies of East Stroudsburg University of Pennsylvania. Student’s Name: Amanda D. Layden, B.S. Title: Determining how Environmental Changes Impact Growth of Batrachochytrium dendrobatidis Using a Novel In Vitro System Date of Graduation: May 8, 2020 Thesis Chair: Joshua Loomis, Ph.D. Thesis Member: William Loffredo, Ph.D. Thesis Member: Emily Rollinson, Ph.D. Abstract Chytridiomycosis, caused by the etiologic agent Batrachochytrium dendrobatidis, affects the keratinocytes of the amphibian epithelium. While there have been several studies done on B. dendrobatidis both in vivo and in vitro, there is still little known about what environmental factors influence the growth of this fungus. To better understand such factors, a novel, high-throughput in vitro system was developed that utilized tissue culture plates as a submerged in vitro substrate. After analyzing B. dendrobatidis’s life cycle in this new system, studies were conducted to determine the impact of pH, phosphate and nitrate concentration, and protein concentration on its growth. Results showed that B. dendrobatidis completed its life cycle in submerged tissue culture wells and that growth rates were sensitive to concentrations of protein and environmental pH. Results suggest that B. dendrobatidis can regulate its growth kinetics depending on access to environmental nutrient sources. Acknowledgements I would like to acknowledge everyone that has helped me through my thesis. I acknowledge all my committee members, other faculty at East Stroudsburg University, Dr. Joyce E. Longcore from the University of Maine Chytrid Laboratory, Sigma XI, and my family and friends who have supported and guided me during this process. I thank my graduate committee members Dr. Joshua Loomis, Dr. William Loffredo, and Dr. Emily Rollinson for all their endless help and guidance through this process. Special thank you to Dr. Loomis for taking a chance on me and committing to work with me as my committee chair. Thank you to Dr. Loffredo for not only supporting me through my thesis project but also supporting me during my undergraduate years at East Stroudsburg University. Thank you to Dr. Rollinson for helping with my statistics and for teaching me to think about the big picture with my project. You all taught me lessons that go beyond the classroom, and I appreciate it whole-heartedly. Thank you to Dr. Joyce Longcore for sending me my original samples of Batrachochytrium dendrobatidis. Thank you to Sigma XI for providing me with funding for my project. Thank you to Dr. Thomas Tauer for letting me borrow materials for my project and thank you to Larry Laubach and Heather Dominguez for helping me order materials for my project. Lastly, I would like to thank all my friends and family who have helped me during this process. I’d especially like to thank my friends Kristine Bentkowski, Kacie Marcum, Eric Januszkiewicz, Melanie Quain, and Ryan McGonagle, as well as my incredible parents, William and Denise, my sister, Kiera, and my amazing boyfriend, Nate, for also offering endless support throughout this process. Table of Contents List of Figures .................................................................................................................. III List of Tables ...................................................................................................................... V Chapter I............................................................................................................................. 1 Introduction ......................................................................................................................1 What is a Wildlife Disease? ..................................................................................................... 6 Origin and Dissemination ........................................................................................................ 8 Life Cycle................................................................................................................................. 9 Overview of Morphology ...................................................................................................... 10 Optimal Growth Environment ............................................................................................... 12 Transmission and Clinical Signs ............................................................................................ 13 Pathology and Pathogenesis ................................................................................................... 15 Immune Defenses Against B. dendrobatidis.......................................................................... 16 Attachment and Colonization of Amphibian Skin ................................................................. 18 Environmental factors affecting growth of B. dendrobatidis................................................. 20 Nitrogen and Phosphorus ....................................................................................................... 21 Study Objectives .................................................................................................................... 23 Chapter II ......................................................................................................................... 25 Materials and Methods ................................................................................................... 25 Obtaining B. dendrobatidis Strain JEL 423 ........................................................................... 25 Cryo-preserving B. dendrobatidis Isolates............................................................................. 25 Thawing of Cryo-preserved B. dendrobatidis Isolates .......................................................... 26 Novel in vitro growth of B. dendrobatidis ............................................................................. 26 Crystal Violet Staining of B. dendrobatidis Isolates.............................................................. 27 Effect of pH on Growth of B. dendrobatidis ......................................................................... 27 Effect of Keratin on Growth of B. dendrobatidis .................................................................. 28 Effect of Nitrate on Growth of B. dendrobatidis ................................................................... 29 Effect of Phosphate on Growth of B. dendrobatidis .............................................................. 29 I Statistical Analysis ................................................................................................................. 30 Chapter III........................................................................................................................ 31 Results ............................................................................................................................ 31 Creation of a Novel In Vitro System...................................................................................... 31 Effects of pH on the Growth of B. dendrobatidis .................................................................. 36 Effects of Keratin on the Growth of B. dendrobatidis ........................................................... 37 Effect of Phosphate on the Growth of B. dendrobatidis ........................................................ 41 Effect of Nitrate on the Growth of B. dendrobatidis ............................................................. 42 Chapter IV ........................................................................................................................ 44 Discussion ....................................................................................................................... 44 Creation of a Novel in vitro System....................................................................................... 45 Effects of pH, Nitrate and Phosphate on the Growth of B. dendrobatidis ............................. 46 Effect of Keratin on the Growth of B. dendrobatidis............................................................. 49 Conclusions ............................................................................................................................ 52 Future Studies ........................................................................................................................ 52 Literature Cited ................................................................................................................ 54 Appendix A: Raw Data ................................................................................................... 69 Appendix B: R Code ....................................................................................................... 76 II List of Figures Figure Page Figure 1. Cladogram indicating taxonomy of B. dendrobatidis showing that it falls in the a) kingdom Fungi, b) phylum Chytridiomycota and c) order Rhizophydiales. (Adapted from Van Rooij et al. 2015: the topology is derived from Martel et al. 2013, Longcore et al. 1999 and Hibbett et al. 2007)45,66,69,113 ............................................................................................................. 2 Figure 2. Worldwide distribution of B. dendrobatidis. (Adapted from Fisher et al. 2009)32 ........... 3 Figure 3. Time bar showing the first occurrence of chytridiomycosis in Africa in 1938, the first occurrence outside of Africa in 1961, (Quebec, Canada, North America) and records outside of Africa following the 23-year gap. (Adapted from Weldon et al. 2004, Quellet 2003, Berger 1999, Speare 2001, Bonaccorso 2003, Rollins-smith 2002, Bosh 2000, Waldman 2001.)8,11,12,84,92,105,118,119 .................................................................................................................... 9 Figure 4. Life cycle of B. dendrobatidis in culture: A=zoospore, B=germling, C=mature zoosporangium, D=moncentric zoosporangium, E=colonial zoosporangium with a dividing septum. (Adapted from Berger et al. 2005)6 .................................................................................. 10 Figure 5. Image showing a formalin-fixed B. dendrobatidis zoospore with multiple small lipid droplets (L) taken from the skin of a Cane toad (Bufo marinus) (N = nucleus, R = ribosomes, Mb = microbody, L = lipid droplet) (Adapted from Berger et al. 2005)6............................................. 11 Figure 6. Clinical signs of chytridiomycosis. a) naturally infected moribund common midwife toad (Alytes obstetricans) with abduction of the hind legs and loose sloughed skin. b) section through the ventral skin (drink patch) of the same infected toad showing epidermal hyperkeratosis and hyperplasia combined with the presence of numerous zoosporangia. c) detail of intracellular septate zoosporangia. (Adapted from Pessier 2008)78 ................................................................... 15 Figure 7. Image showing the infection cycle of B. dendrobatidis in a susceptible host. The lifecycle includes invasion mediated by a discharge tube, establishment of intracellular thalli, spreading to the deeper skin layers, and upward migration by the differentiating epidermal cell to finally release zoospores at the surface of the skin (Adapted from Berger at al. 2005, Van Rooij et al. 2012, and Greenspan et al. 2012)6,40,112 ..................................................................................... 20 Figure 8. Life cycle of B. dendrobatidis as shown from A-H. A= day 1: motile zoospores. B= day 2: germlings. C= day 3: developing zoosporangia/germlings. D-H= days 4-8: developed zoosporangia with note of newly produced zoospores at day 5 shown by black line arrow (Photo Credit to Amanda Layden)............................................................................................................. 33 III Figure 9. Structures and stages of the life cycle of B. dendrobatidis as shown from A-D. A= zoomed in view of Day 5 from life cycle in tissue culture plates in vitro to show newly produced zoospore. B= left arrow shows a developing monocentric zoosporangium and right arrow shows a mature colonial zoosporangium with a septum dividing the thallus body into two compartments. C= germlings stained with crystal violet to show rhizoid structures noted by arrow. D= a clear, empty zoosporangium with a single discharge papillae (tube) noted by arrow (Photo Credit to Amanda Layden)............................................................................................................................ 34 Figure 10. Growth of B. dendrobatidis using this novel in vitro system. A= growth quantified using cell counts on a hemocytometer, B= growth quantified using absorbance (495 nm) values from a spectrophotometer .............................................................................................................. 35 Figure 11. Effect of pH values on growth of B. dendrobatidis in the novel in vitro system ......... 36 Figure 12. Effect of protein on growth of B. dendrobatidis. A= quantitative data showing effect of keratin on growth on B. dendrobatidis in novel in vitro system. B= quantitative data showing effect of keratin on growth on B. dendrobatidis in broth tubes. C= image showing effect of keratin on growth on B. dendrobatidis when added to 1% tryptone agar. 1% tryptone agar is shown on the top row and 1% tryptone + keratin agar is shown on the bottom row. D= image showing effect of bovine serum albumin on growth of B. dendrobatidis when added to 1% tryptone agar. (Photo Credit to Amanda Layden) .......................................................................... 40 Figure 13. Effect of phosphate concentration growth of B. dendrobatidis .................................... 42 Figure 14. Effect of nitrate concentration on growth of B. dendrobatidis ..................................... 43 IV List of Tables Table Page Table 1. Examples of affected wild amphibian species due to chytridiomycosis4 .......................... 3 Table 2. pH Analysis of Variance ................................................................................................. 37 Table 3. Keratin Broth Tubes Analysis of Variance ...................................................................... 41 Table 4. Keratin Analysis of Variance ........................................................................................... 41 Table 5. Phosphate Analysis of Variance ...................................................................................... 42 Table 6. Nitrate Analysis of Variance............................................................................................ 43 Table 7. Environmental Factors Raw Data .................................................................................... 69 V Chapter I Introduction Chytridiomycosis is an emerging infectious wildlife disease that affects the keratinized epidermal cells of the amphibian epithelium. The etiologic agent that causes this disease is Batrachochytrium dendrobatidis, a spherical-shaped fungus that is found in a variety of water sources and moist soil environments. B. dendrobatidis taxonomically falls in the Phylum Chytridiomycota, Class Chytridiomycetes, and Order Rhizophydiales (Figure 1). Chytridiomycota (chytrids) is the only phylum of true Fungi that reproduces with posteriorly uniflagellate, motile spores (zoospores)48,97.. The order Rhizophydiales was formed on the basis of molecular monophyly and zoospore ultrastructure, in which three new families and two new genera were delineated60. Of these three families, the Rhizophydiaceae includes a species known as Rhizophydium globosum, which has been included in numerous chytrid inventories35,55,59,60,72,104. R. globosum is sparsely described as having a spherical sporangium with 2-4 discharge papillae and occurs as a parasite on Closterium and other algal hosts16,60. Although B. dendrobatidis has not been officially assigned a taxonomic family, there are other chytrids in the order Rhizophydiales that act as a parasite on other organisms. 1 B. dendrobatidis is the only known parasitic chytrid fungus of vertebrates. It has been implicated as the main factor in severe amphibian population declines and has been confirmed on every major continent except Antarctica (where amphibian fauna are not present)32 (Figure 2). B. dendrobatidis infects over 350 amphibian species and has been implicated in driving the decline of over 200 of them32,103. Some of these affected species have been categorized as critically endangered (CR), endangered (EN) and in some cases extinct (EX) on the IUCN Red List as a result of this emerging disease4,5,7,13,37,62,63,73,86,89,100,110,111 (Table 1). Figure 1. Cladogram indicating taxonomy of B. dendrobatidis showing that it falls in the a) kingdom Fungi, b) phylum Chytridiomycota and c) order Rhizophydiales. (Adapted from Van Rooij et al. 2015: the topology is derived from Martel et al. 2013, Longcore et al. 1999 and Hibbett et al. 2007)45,66,69,113 2 Figure 2. Worldwide distribution of B. dendrobatidis. (Adapted from Fisher et al. 2009)32 Table 1. Examples of affected wild amphibian species due to chytridiomycosis4 Species Common Name Habitat Rheobatrachus vitellinus Eungella gastricbrooding frog Panamanian golden frog Australia Panama Pathogenic Sharpsnouted day frog Western toad Australia Pathogenic USA Pathogenic Chiriqui harlequin frog Sardinian brook salamander Costa Rica Pathogenic Italy Pathogenic Atelopus zeteki Taudactylus acutirostris Anaxyrus boreas Atelopus chiriquiensis Euproctus playtcephalus 3 Mechanism Conservation Reference Status (IUCN red list) Pathogenic Extinct (EX) Retallick, et al. between 2004 1985-1986 Still listed as Critically Endangered (CR); but most likely extinct (EX) Extinct: between 1993-1994 Near threatened (NT) Critically endangered (CR) Endangered (EN) Gewin, 2008 Schloegel et al. 2005 Muths et al. 2003 Berger et al. 1998; Lips, 1999 Bovero et al. 2008 Gastrotheca cornuta Horned marsupial frog Costa Rica, Panama, Ecuador, Columbia New Zealand Pathogenic Endangered (EN) Lips et al. 2006 Leiopelma archeyi Archey’s frog Pathogenic Bell et al. 2004 Mountain yellowlegged frog USA; California Pathogenic Critically Endangered (CR) Endangered (EN) Rana muscosa Eleutherodactylus jasperia Golden coqui Puerto Rico Pest and disease transmission Critically Endangered (CR) US Fish and Wildlife Service, 1999; Rachowisz et al. 2006 US Fish and Wildlife Service, 2013 The evidence implicating B. dendrobatidis in the amphibian declines is compelling. Firstly, chytrid fungus can be pathogenic to amphibians in both the field and the laboratory7,79. A study done by Berger et al. utilized experimental transmission of cutaneous chytridiomycosis on captive-bred sibling frogs (Mixophyes fasciolatus)7. The sample was taken from a dead frog of the same species that had naturally acquired the infection7. In the results, it was noted that chytrid sporangia were seen during histological examination of the captive-bred sibling M. fasciolatus frogs7. Furthermore, they concluded that chytrids are associated with a transmissible fatal disease of anurans in the field and in the laboratory7. Secondly, there is genetic evidence suggesting the emergence of a hypervirulent strain of chytrid fungus that shows genetic signal consistent with range expansion29,49,79. A study done by Farrer et al. collected samples of B. dendrobatidis isolates from locations on every continent except Antarctica and found that there was a much greater 4 diversity of B. dendrobatidis than was previously recognized29. They also noted that multiple lineages were being vectored between continents by the trade of amphibians29. One of those lineages, (BdGPL=global panzootic lineage) had been characterized with hypervirulence, suggesting that the emergence and spread of chytridiomycosis is largely due to the globalization of the recently emerged recombinant lineage29,31. Ultimately, the researchers concluded that the global trade in amphibians is resulting in contact and cross-transmission of B. dendrobatidis among previously naïve host species which resulted in intercontinental pathogen spread and an increase in recombinant genotypes generated29. Lastly, amphibian population declines appear to have followed a broad wave-like pattern consistent with the spread of a novel pathogen63,64,79. A study done by Lips et al. discussed analyses supporting a classical pattern of disease spread across naïve populations (at odds with the CLEH (climate-linked epidemic hypothesis) proposed by Pounds et al., 2006) and how their analyses cast doubt on CLEH64,81. In their results, they found evidence of directional spread of B. dendrobatidis along most of the principal cordilleras of Lower Central America and the Andean region, supporting the hypothesis that this is an exotic pathogen that was introduced into South America in the late 1970searly 1980s and has caused multiple amphibian declines in the past 30 years20,58,63,64,68,93. One of these declines (the 1987 amphibian decline at Monteverde Cloud Forest Reserve in Costa Rica) is widely assumed to have been caused by an outbreak of B. dendrobatidis; however, direct evidence does not exist64. Prevalence of B. dendrobatidis was noted in 2003, indicating that the pathogen is now endemic to that area64. The researchers examined museum specimens for evidence of B. dendrobatidis prior to 1986 5 and found that most of the specimens showed histological evidence of B. dendrobatidis infection64. Ultimately, their analyses supported a hypothesis that B. dendrobatidis is an introduced pathogen that spreads from its point of origin in a pattern typical of emerging infectious diseases64. What is a Wildlife Disease? A wildlife disease can be defined as a pathological condition occurring in a susceptible population in nature. Emerging infectious diseases (EIDs) of free-living wild animals can be classified into three major groups on the basis of key epizootiological criteria. The first group involves EIDs associated with “spill-over” from domestic animals to wildlife populations living in proximity. The second group involves EIDs related directly to human intervention, via host or parasite translocations. The final group of EIDs is related with no overt human or domestic animal involvement24. These diseases have two major biological implications: first, many wildlife species are reservoirs of pathogens that threaten domestic animal and human health, and second, wildlife EIDs pose a substantial threat to the conservation of global biodiversity24. The USGS National Wildlife Health Center (NWHC) works to safeguard our nation’s wildlife from diseases by studying their causes and by developing strategies to prevent and manage them75. Aside from chytridiomycosis, other wildlife diseases exist and have not only caused devastating declines in wildlife populations globally but have also caused issues in human populations. An example of a wildlife disease that has caused issues in human populations is Lyme disease. Lyme disease is spread by the 6 blacklegged tick (Ixodes scapularis) and the CDC estimates reports of approximately 30,000 confirmed cases each year25. There are many wildlife diseases aside from Chytridiomycosis that cause harm to populations found in nature; however, the top three are Chytridiomycosis, White-Nose Syndrome, and Snake Fungal Disease (SFD). WhiteNose Syndrome affects all life stages of hibernating bats, and mortality at newly-affected hibernacula can be very high, resulting in substantial and rapid decreases in bat abundance33,82. Millions of North American bats have died from this disease, and population declines for heavily impacted species could result in regional extirpation of some previously common species such as the little brown bat (Myotis lucifugus) and northern long-eared bat (M. septentrionalis)17,28,33,34,82,90,106. Snake Fungal Disease (SFD) has been confirmed in numerous species of snakes and is caused by the fungus Ophidiomyces ophiodiicola107. As of August 2017, this fungus has been detected in at least 23 states and one Canadian province; however, researchers suspect that SFD may be more widely distributed due to limitations in monitoring snake populations107. Studying disease ecology in wildlife can be challenging but understanding wildlife epidemiology is important for the benefit of human health, animal welfare, productivity in agricultural systems, and global biodiversity24,26,70,122. A similar factor between many emerging wildlife diseases is that the global trade of wildlife provides disease transmission mechanisms for these pathogens54. Outbreaks resulting from wildlife trade have caused hundreds of billions of dollars of economic damage globally54. For instance, white-nose syndrome is hypothesized to have been introduced to North America from Europe or Asia33,82. Since there is no bat migration occurring between North America and Europe, it is very likely that this fungus was 7 introduced to North America from global movement of humans, animals, and trade120. Similarly, examination of historical fungal isolates has demonstrated that O. ophiodiicola was present in captive snakes in the eastern USA since at least 198667,102. Furthermore, no wild snake isolates are known prior to 2008, indicating that introduction by spillover of O. ophiodiicola from captive to wild snake populations represents a plausible explanation for the sudden emergence of SFD67. In regard to Chytridiomycosis, the global trade of a specific species of anuran has enabled B. dendrobatidis to be transmitted throughout the world. Origin and Dissemination Discovering the origin of an infectious disease is critical for determining how to prevent and treat it. To date, the origin of B. dendrobatidis is something still argued by herpetologists, mycologists, and epidemiologists around the world. The earliest case of chytridiomycosis was recorded in 1938 from an African clawed frog (Xenopus laevis) in southern Africa119 (Figure 3). Chytridiomycosis was a stable, endemic infection in southern Africa for 23 years before any positive specimens were found outside of Africa119. Some emerging infectious diseases arise when pathogens that have been localized to a single host or small geographic region go beyond previous boundaries and according to research; it is highly likely that this is how B. dendrobatidis emerged as well119. African clawed frogs are considered natural carriers of B. dendrobatidis and are not overly susceptible to its disease symptoms. After 23 years of globally trading African clawed frogs for educational and research purposes, the first case of chytridiomycosis 8 outside of Africa was noted in North America in 1961, specifically in Quebec, Canada114,119. After the case in Canada, the earliest cases from other countries follow sequentially over a period of 38 years from 1961 to 1999119 (Figure 3). Figure 3. Time bar showing the first occurrence of chytridiomycosis in Africa in 1938, the first occurrence outside of Africa in 1961, (Quebec, Canada, North America) and records outside of Africa following the 23-year gap. (Adapted from Weldon et al. 2004, Quellet 2003, Berger 1999, Speare 2001, Bonaccorso 2003, Rollins-smith 2002, Bosh 2000, Waldman 2001.)8,11,12,84,92,105,118,119 Life Cycle The life cycle of B. dendrobatidis begins with a motile zoospore and is approximately 4-5 days. Once the zoospore attaches to a substrate, it morphologically changes into a growing organism called a thallus. Once matured, the thallus body grows into a single zoosporangium (container for zoospores)6 (Figure 4). The contents of the zoosporangium (also known as the sporangium) cleave into new zoospores which exit the sporangium through one or more discharge papillae (also called discharge tubes)6. While sexual reproduction has not been seen in this organism to date, there is another variation in the life cycle known as ‘colonial development’ resulting from the formation of more than one sporangium from one zoospore66. Zoosporangia undergoing colonial development have a septum dividing the contents of the zoosporangium. The life 9 cycle of this fungus has been found to be the same in culture (in vitro) as it is on amphibian skin (in vivo)9,66. Figure 4. Life cycle of B. dendrobatidis in culture: A=zoospore, B=germling, C=mature zoosporangium, D=monocentric zoosporangium, E=colonial zoosporangium with a dividing septum. (Adapted from Berger et al. 2005)6 Overview of Morphology Zoospore and Germling Zoospores are the waterborne, motile stage of the life cycle. Zoospores of B. dendrobatidis are unwalled and mostly spherical shaped but can also be elongate and amoeboid when they are first released from the zoosporangium6,66. The zoospores are approximately 3-5 µm in diameter with a posteriorly directed flagellum66. Zoospore ultrastructure is used to differentiate orders and genera among the Chytridiomycota. The features of the zoospore of B. dendrobatidis that are common to the order Chytridiales are that the nucleus and kinetosome are not associated, ribosomes are aggregated into a core surrounded by endoplasmic reticulum, the microbody partially surrounds the lipid 10 globules, and the nonflagellated centriole (NFC) is parallel and connected to the kinetosome65,66. A key feature of B. dendrobatidis is the numerous small lipid droplets with the microbodies that are associated with the edge of the ribosomal mass66 (Figure 5). Additionally, B. dendrobatidis is aneuploid, with copy numbers of the chromosomal regions (contigs) within a single isolate running up to 530,94,101,113. After a period of motility and dispersal, the zoospore encysts, the flagellum is resorbed, and a cell wall forms6. Once the zoospore has encysted, fine branching rhizoids grow from one or more areas of the zoospore and it is then known as a germling6. Figure 5. Image showing a formalin-fixed B. dendrobatidis zoospore with multiple small lipid droplets (L) taken from the skin of a Cane toad (Bufo marinus) (N = nucleus, R = ribosomes, Mb = microbody, L = lipid droplet) (Adapted from Berger et al. 2005)6 Developing Zoosporangia As the germling develops, the thallus grows and the cytoplasm becomes more complex6. As this occurs, the thallus becomes multinucleate by mitotic divisions6. The contents then cleave and mature into rounded, flagellated zoospores6. At this point, the 11 swollen part of the thallus is now known as a zoosporangium6. Simultaneously, one or more discharge papillae (tubes that stick out away from the zoosporangium that aid in zoospore release) form. Some thalli that undergo colonial growth become divided by thin septa and each compartment grows into a separate sporangium with its own discharge tube6. These mature zoosporangia contain fully formed flagellated zoospores6. Zoospores are released when the plug blocking the discharge tube is dissolved. Once all the zoospores are released, it is considered an empty sporangium. The chitinous walls of the sporangia remain and may eventually collapse. Sometimes, zoospores do not escape and grow within the sporangia6. Optimal Growth Environment Growth and survival of B. dendrobatidis is dependent on many environmental factors. Optimal growth of B. dendrobatidis is observed between 17 and 25℃ and at pH 6-7 in vitro (agar and broth culture) which is similar to what is observed in amphibian skin in vivo and in the environment80,113. B. dendrobatidis grows slowly at 10℃ and ceases growth at 28℃ or higher50,80,113. Additionally, B. dendrobatidis zoospores are killed within four hours at 37 ℃50,80,113. Desiccation is poorly tolerated as this species requires wet or moist environments36,50,113. It has also been noted that 5% NaCl solutions are lethal to this pathogen36,50,113. In vitro, B. dendrobatidis has been shown to grow on a variety of keratin containing substrates such as autoclaved snakeskin, 1% keratin agar, frog skin agar, feathers and geese paws36,66,80,113. B. dendrobatidis can also grow on chitinous carapaces of crustaceans71,113. Although B. dendrobatidis grows well on these 12 substances, it grows best in tryptone or peptonized milk in both agar and broth in vitro66,113. The type of growth system used for studying B. dendrobatidis ultimately depends on the research questions under investigation. An in vitro system would be ideal for studying specific environmental factors on the growth of B. dendrobatidis because the variables can be easily manipulated. In contrast, an in vivo study involving specific environmental factors would be difficult because not all individual amphibians from the same species are exactly the same in regard to their immune system, skin microbiome, or other host defenses. Studies that require specific pathogen-host interactions can best be observed using an ex vivo or in vivo approach to obtain specific host defense data. Transmission and Clinical Signs In terms of virulence, B. dendrobatidis has an extremely broad host range, infecting at least 520 species of anurans (frogs and toads), urodeles (salamanders and newts) and caecilians39,113. Transmission among hosts is typically due to infection of motile waterborne zoospores or through direct contact with infected amphibians (ex. during mating)97,113. Another factor involving B. dendrobatidis’s virulence is that it can survive in water and moist soil for weeks up to several months, which makes it hard for amphibians to not become infected once they have entered an infected water source51,52,113. Additionally, B. dendrobatidis is able to saprobically grow on sterile bird feathers, arthropod exoskeletons, keratinous paw scales of waterfowl and can survive in the gastrointestinal tract of crayfish36,51,52,66,71,113. Being able to grow on many different 13 substances also increases this pathogen’s spreading capability and increases its chances of being transmitted to a new host. In anuran larvae, clinical signs of chytridiomycosis are generally limited to depigmentation of the mouthparts, low foraging, lethargy, and poor swimming abilities7,86,113. Although this does not cause mortality, chytridiomycosis can commonly contribute to reduction in anuran larvae body size43,113. In metamorphized amphibians, clinical signs are variable and range from significant skin disorder to sudden death without obvious disease symptoms113. The most common signs of chytridiomycosis are excessive shedding of the skin, erythema (redness), or discoloration of the skin78,113 (Figure 6). In frogs and toads, the skin of the ventral abdomen, especially the pelvic patch, feet and toes, are predilection sites of infection9,83,113. In contrast, salamanders are more prone to infection in the pelvic region, fore and hind limbs and the ventral side of the tail113,114. Other clinical signs of chytridiomycosis include lethargy, anorexia, abnormal posture, and neurological signs such as loss of righting reflex and flight response78,113. 14 Figure 6. Clinical signs of chytridiomycosis. a) naturally infected moribund common midwife toad (Alytes obstetricans) with abduction of the hind legs and loose sloughed skin. b) section through the ventral skin (drink patch) of the same infected toad showing epidermal hyperkeratosis and hyperplasia combined with the presence of numerous zoosporangia. c) detail of intracellular septate zoosporangia. (Adapted from Pessier 2008)78 Pathology and Pathogenesis In metamorphized amphibians, chytridiomycosis caused by B. dendrobatidis is diagnosed by the presence of immature chytrid thalli or maturing sporangia found intracellularly in the keratinized layers of the amphibian skin113. Infection is associated mainly with a mild to severe irregular thickening of the outermost keratinized layers of the epidermis (hyperkeratosis of the stratum corneum and stratum granulosum)113. Infection can also cause erosion of the stratum corneum and increased tissue growth (hyperplasia) of the stratum spinosum, which lies beneath the keratinized superficial skin layers113. Dissemination to the deeper layers of the skin or internal organs does not occur78,113. Instead, amphibian mortality is caused by B. dendrobatidis disrupting normal regulatory function of their skin96. Infection in anuran larvae is limited to the keratinized mouthparts78,113. It is only when the anuran larvae undergoes metamorphosis that the infection is able to spread to the epithelia of the body, limbs, and tail. 15 With the availability of B. dendrobatidis’s full genome, genetic studies have led to an improved understanding of host-pathogen dynamics and the identification of several putative pathogenicity factors with high specificity for skin-related substrates, facilitating colonization or causing host damage113. Nevertheless, processes that take place during the whole infection cycle at a molecular and cellular level such as cell signaling, induction of cytoskeletal change and so on are still barely understood and require more attention113. Immune Defenses Against B. dendrobatidis Innate and acquired immune components both contribute to the antimicrobial function of amphibian mucus113. Firstly, amphibians produce antimicrobial peptides in their dermal glands to act as an innate immune defense mechanism113. To date, approximately forty anuran antimicrobial peptides inhibiting B. dendrobatidis have been characterized91,113. Both purified and natural mixtures of these antimicrobial peptides effectively inhibit in vitro (broth and agar) growth of B. dendrobatidis zoospores and sporangia87,91,113,123. Although these antimicrobial peptides have been found to inhibit growth of B. dendrobatidis in vitro, it is unclear how these peptides provide protection against chytridiomycosis in vivo113. Another innate immune defense mechanism against chytridiomycosis is antifungal metabolites secreted by symbiotic bacteria present on amphibian skin113. To date, there have been only 3 inhibitory metabolites identified by the symbiotic bacterial species Janthinobacterium lividum, Lysobacter gummosus, and Pseudomonas fluorescens18,113. These natural metabolites are known as 2,4-DAPG (2,4diacetylphloroglucinol), indol-3-carboxaldehyde (I3C) and violacein18,113. These 16 metabolites can inhibit growth of B. dendrobatidis both in vitro and in vivo18,57,74,113. Myers et al. discovered that these metabolites work synergistically with antimicrobial peptides to inhibit growth of B. dendrobatidis at lowered minimal inhibitory concentrations necessary for inhibition by either metabolites or antimicrobial peptides74,113. In addition, 2,4-DAPG and I3C seem to repel B. dendrobatidis zoospores57,113. A final innate immune defense mechanism with fungicidal potential in amphibian skin mucus is lysozyme; however, this has not been studied in detail91,113. Bacterial cells contain two alternating amino acids sugars, N-acetylglucosamine (GlcNAc or NAGA) and N-acetylmuramic acid (MurNAc or NAMA), which are connected by a β1,4-glycosidic bond113. Lysozyme catalyzes bacterial cell lysing of the β-1,4 bonds of peptidoglycan, a polymer of N-acetylmuramic acid (GlcNAc) that is found in their cell wall113. Since the fungal cell wall consists mainly of chitin, a similar polymer consisting of β-1,4 linked GlcNAc units, it is also a potential target for lysozyme113. In contrast, the acquired immune system provides very specific protection against pathogens and involves both cell-mediated and humoral antibody responses. However, many researchers have become confused because of the apparent absence of a robust immune response in susceptible amphibian species113. So far, attempts to immunize frogs using subcutaneous or intraperitonial injection of formalin or heat-killed B. dendrobatidis failed to elicit an acquired immune response113. Only in X. laevis, B. dendrobatidis specific IgM, IgX (mammalian IgA-like) and IgY (mammalian IgG-like) antibodies were found in skin mucus after injection with heat-killed zoospores87,113. According to Ramsey et al., the mucosal antibodies elicited in X. laevis frogs bind with B. dendrobatidis zoospores in vitro and are suggested to limit colonization of the skin to mild and non17 lethal infections; however, their contribution to actual protection is still undetermined87,113. Rollins-Smith et al. observed that as B. dendrobatidis infections naturally occur in the skin, it seems likely that introduction of B. dendrobatidis antigens directly into the skin may be more effective, but more research needs to be done on this topic91,113. Despite this, susceptible amphibians still acquire this disease indicating that this fungus can withstand the host immune defenses. Attachment and Colonization of Amphibian Skin B. dendrobatidis infection of amphibian skin begins with the attachment of motile zoospores to the host’s skin (Figure 7). It is at this step when B. dendrobatidis interacts with the amphibian’s mucus barrier (mucosome). The main components of mucus are mucins or mucin glycoproteins113. The mucosome may be able to reduce the infection load on the skin during the first 24 hours of exposure, which is critical for colonization and establishing skin infection112,113. At this point, the zoospores germinate and adhere to the host surface. To date, the mechanisms and kinetics of adhesion of B. dendrobatidis to amphibian skin have only received limited attention113. Adhesion has been documented to occur approximately 2-4 hours after exposure to zoospores112. After the zoospores have attached, they mature into thick walled cysts on the host epidermis and often cluster in foci of infection113. The cysts are anchored into the skin via fine fibrillar projections, rhizoids and some adhesion not yet determined. These fibrillar projections and adhesions are similar to fibrillar adhesins documented for pathogenic fungi affecting human skin (Trichophyton mentagrophytes)113. Several genes encoding proteins involved in cell 18 adhesion such as vinculin, fibronectin, and fasciclin have been identified in the B. dendrobatidis genome and are expressed more in sporangia than in zoospores95,113. B. dendrobatidis is also equipped with a chitin binding module (CBM18) that is hypothesized to facilitate survival on its amphibian host113. It is suggested that a key role of CBM18 involves pathogenesis and protection against host-derived chitinases113. CBM18 also allows B. dendrobatidis to attach to non-host chitinous structures (insect or crustacean exoskeletons) allowing vectored-disease spread1,71,113. Once the zoospore has encysted, invasion of the epidermis begins. In general, B. dendrobatidis develops endobiotically, with sporangia located inside the host cell. This is generally achieved within 24 hours after initial exposure113. Colonization is established from the extension of a germ tube (discharge papillae) arising from the zoospore cyst that penetrates the host cell membrane and enables transfer of genetic material (zoospore nucleus and cytoplasm) into the host cell112,113. The distal end of the germ tube becomes swollen and gives rise to a new intracellular chytrid thallus113. B. dendrobatidis continues to use this mechanism to spread to deeper skin layers. Older thalli develop rhizoid-like structures that spread to deeper skin layers113. At this point, they form a swelling inside the host cells in the deeper skin layers and give rise to new daughter thalli113. 19 Figure 7. Image showing the infection cycle of B. dendrobatidis in a susceptible host. The lifecycle includes invasion mediated by a discharge tube, establishment of intracellular thalli, spreading to the deeper skin layers, and upward migration by the differentiating epidermal cell to finally release zoospores at the surface of the skin (Adapted from Berger at al. 2005, Van Rooij et al. 2012, and Greenspan et al. 2012)6,40,112 Environmental factors affecting growth of B. dendrobatidis Changes to the chemical composition of an environmental water source have the potential to drastically alter the growth of microorganisms like B. dendrobatidis. For instance, the sudden introduction of nutrients such as nitrogen, phosphorus, and organic waste can trigger massive increases in microbial populations, which can have deleterious effects on the other aquatic life in that water source. Such changes can be caused by sewage infiltration, human pollution, or runoff. According to the USGS, runoff can be defined as the part of the precipitation, snow melt, or irrigation water that appears in uncontrolled water sources98. These water sources, surface streams, rivers, drains, or sewers, can be classified according to speed of appearance after rainfall or melting snow as direct runoff or base runoff98. Additionally, they can be classified according to source as surface runoff, storm interflow, or groundwater runoff98. When rain falls onto the landscape, it doesn’t wait to be evaporated by the sun or used as a drinking source by the local wildlife. Instead, it begins to move slowly due to gravity98. Some of the rainwater 20 seeps into the ground to refresh groundwater, but most of it flows down gradient. This is known as surface runoff98. As watersheds are urbanized and much of the vegetation is replaced by impervious surfaces, groundwater infiltration is reduced and stormwater runoff increases98. Stormwater runoff must be collected by drainage systems and storm sewers that carry the runoff directly to streams. Stormwater runoff that flows over the land surface can pick up potential pollutants that may include sediments, nutrients (from lawn fertilizers – nitrogen (N) and phosphorus (P)), bacteria (from animal and human waste), pesticides (from lawn/garden chemicals), metals (from rooftops and roadways), and petroleum by-products (from leaking vehicles)98. Nitrogen and Phosphorus Nitrogen (N) and Phosphorus (P) are two important and essential nutrients for healthy soil and aquatic environments. According to the Environmental Protection Agency (EPA), nitrogen is generally used and reused by plants within natural ecosystems, with minimal “leakage” into surface or groundwater, where nitrogen concentrations remain very low109,117. However, when nitrogen is applied to the land in amounts greater than can be incorporated into crops or lost in the atmosphere through volatilization or denitrification, concentrations in soil and streams can cause environmental issues109. The major sources of excess nitrogen in streams and other agricultural watershed sources are fertilizer and animal waste109. Excess nitrate is not toxic to aquatic life, but increased nitrogen may result in overgrowth of microorganisms 21 like soil bacteria, soil fungi, and algae (known as algal blooms)108. This can decrease the dissolved oxygen content of the water, thereby harming or killing fish and other aquatic species108. Phosphorus is also an essential nutrient for all life forms, but at high concentrations the most biologically active form of phosphorus, phosphate, can cause water quality problems by also overstimulating the growth of microorganisms (similar to nitrogen). Elevated levels of phosphorus in streams can result from fertilizer used, animal wastes, and wastewater109. The EPA states that freshwater streams and ponds fall under one of five categories when looking at nitrate levels (mg/L): <1 mg/L, 1-2 mg/L, 2-6 mg/L, 6-10 mg/L, and 10 mg/L or more109. The EPA also states that for phosphate levels, freshwater streams and ponds fall under one of four categories: <0.1 mg/L, 0.1-0.3 mg/L, 0.3-0.5 mg/L, and 0.5 mg/L or more109. According to the EPA, the recommended water quality for freshwater ponds and streams consists of <1 mg/L nitrogen and <0.1 mg/L phosphorus109. Increased levels of nitrogen and phosphorus can also impact many soil microorganisms. Long-term application of fertilizers can affect the plant-soil-microbe system by changing the composition and structure of plant and soil microbial communities47. Increasing the availability of these nutrients can also cause changes in soil pH. This change can affect species richness by causing a decline of plants and soil microbes47. These effects can eventually cause issues with some of the nutrient cycles many organisms rely on. Nitrogen cycling in natural ecosystems and traditional agricultural production relies on biological nitrogen fixation primarily by diazotrophic bacteria and sometimes, under specific conditions, free-living bacteria such as cyanobacteria, Pseudomonas, Asozpirillum, and Azobacter19,53,77. Diazotrophic 22 community structure and diversity have been shown to respond to changes in the nature of nitrogen added and are also especially sensitive to chemical inputs such as pesticides76,77. Although there has been a lot of research that focuses on the effects of nitrogen and phosphorus on water chemistry, algae, and bacteria, little has been done to study the effects of these elements on soil fungi and B. dendrobatidis in particular. Study Objectives Today, we know that B. dendrobatidis has a complex interaction with amphibians and that the response of amphibians to this pathogen depends on many ecological, environmental, and genetic factors. While these early studies have shed some light on the pathogenesis of B. dendrobatidis, they have provided only a limited understanding of its basic physiological processes. One major limitation is that most experiments with B. dendrobatidis have been conducted either using a complex and relatively expensive ex vivo system that typically involves the use of isolated frog skin or in vivo experiments on amphibians themselves. This study will be the first to utilize a tissue culture system as a novel and cheaper alternative to growing the fungus ex vivo or in vivo and it will be the first to test the effect of nitrogen and phosphate levels on the growth rate of B. dendrobatidis. The objectives were to: 1. Create a new in vitro system using tissue culture plates that will attempt to simulate a submerged growth substrate 2. Validate the in vitro system using already published data from other in vitro and ex vivo studies 23 3. Determine if addition of protein or an excess of nitrogen or phosphorus have an effect on the growth rate of B. dendrobatidis using the new in vitro system 24 Chapter II Materials and Methods Obtaining B. dendrobatidis Strain JEL 423 The original sample of B. dendrobatidis was obtained from Dr. Joyce Longcore from the University of Maine Chytrid Laboratory. Isolates of B. dendrobatidis were aseptically transferred from 1% tryptone agar plates to 100mL of 1% tryptone broth media. The culture was placed at room temperature (21-23℃) for two weeks and was then stored at 4℃ for prolonged usage. Cryo-preserving B. dendrobatidis Isolates Isolates of B. dendrobatidis were cryo-preserved following the procedure by Boyle et al. 200314. Freezing media was composed of 10% dimethyl sulfoxide (DMSO) and 10% Fetal Bovine Serum (FBS) in 1% tryptone broth. The culture used contained actively released zoospores and sporangia that were grown in 100mL of 1% tryptone media for 2 weeks at room temperature (21-23℃). Two milliliters of the actively growing culture was added to 13 mL of fresh 1% tryptone broth and spun in a centrifuge 25 at 1700 RPM for 10 minutes. In a Biosafety cabinet, the supernatant was discarded, and the sporangia pellet was resuspended in 1mL 10% DMSO+10% FBS in 1% tryptone broth and transferred to a 1mL cryotube. This was repeated to make 6 cryotubes. All 6 cryotubes were placed in a -80℃ freezer for long-term storage. Thawing of Cryo-preserved B. dendrobatidis Isolates Each time a cryotube was thawed, it was removed from the -80℃ freezer and placed at 37℃ for 1-2 minutes. Once thawed, the entire contents of the tube were put into 100 mL of fresh 1% tryptone broth. The newly inoculated culture was placed at room temperature (21-23℃) for 2 weeks without shaking to allow for growth. Novel in vitro growth of B. dendrobatidis One milliliter of inoculated culture was aseptically spread onto a 1% tryptone agar plate and placed at room temperature (21-23℃) for 8 days. On day 8, the agar plate was flooded with 5 mL 1% tryptone broth to lift zoospores. The zoospore suspension was then diluted 1:10 in fresh 1% tryptone. Cell density of the 1:10 dilution suspension was then determined using a hemocytometer and the following equation: Total number of cells/number of 1 mm2 squares counted x 10,000/mL x dilution factor After determining cell density, the 1:10 dilution was further diluted in order to achieve a final cell density of 165,000-330,000 zoospores/3mL of media (3mL of media was used in each well). The diluted zoospore suspension was then aseptically transferred 26 into the wells of sterile, 12-well cell culture plates and allowed to incubate at room temperature for a total of 12 days. Every 3 days, cell density was determined by scraping the cells off of the wells using a rubber policeman and measuring absorbance of the suspension at 495 nm. Wells were always scraped in triplicate in order to achieve more accurate data. Crystal Violet Staining of B. dendrobatidis Isolates B. dendrobatidis isolates were aseptically stained with crystal violet in the novel in vitro system to determine if rhizoid structures were present. One milliliter of culture was transferred into multiple wells in the 12-well culture plate and placed at room temperature (21-23℃) overnight to give the fungus time to adhere to the plastic wells. After 24 hours, the culture was pipetted out of the wells and a 0.5% crystal violet (in 1% formaldehyde) stain was placed into each well for approximately 1-2 minutes. Each stained well was then washed with distilled water to discard any residual stain. Once washed, the plate was observed under an inverted phase-contrast microscope using the 40x objective lens (400x total magnification) to determine presence of rhizoid structures. Effect of pH on Growth of B. dendrobatidis B. dendrobatidis was grown on 1% tryptone agar plates and zoospores were harvested after 8 days of growth as described above. Zoospores were diluted to a density of 165,000-330,000 cell/3 mL using 1% tryptone media that had been adjusted to various pHs (5-9) using HCl and NaOH. Diluted cell suspensions were applied to cell culture 1227 well plates and cell growth was monitored every 3 days for a total of 12 days. Similar to the previous experiment, growth was measured by absorbance at 495 nm using approximately 2-3mL of the media harvested from each well. Each measurement was obtained from harvesting wells in triplicate and each experiment was repeated three times. Effect of Keratin on Growth of B. dendrobatidis Two different experiments were conducted in order to determine the effect of keratin on the attachment and growth of B. dendrobatidis. In the first experiment, B. dendrobatidis was grown on two 1% tryptone agar plates and zoospores were harvested after 8 days of growth as described above. After the first plate was harvested, zoospores were diluted to a density of 165,000-330,000 cells/3 mL using 1% tryptone media and those cells were added to normal cell culture wells or wells that had been pre-coated with a 1% keratin solution for 1 hour. Similarly, zoospores were harvested from a second 1% tryptone agar plate and were diluted to a density of 165,000-330,000 cells/3 mL using 1% tryptone + 1% keratin media and those cells were added to normal cell culture wells. Cell growth was monitored every 3 days for a total of 12 days. Similar to the previous experiment, growth was measured by absorbance at 495 nm using approximately 2-3mL of the media harvested from each well. Each measurement was obtained from harvesting wells in triplicate and each experiment was repeated three times. In the second experiment, B. dendrobatidis was grown on two 1% tryptone agar plates and zoospores were harvested after 8 days as described above. Zoospores were diluted to a density of 165,000-330,000 cells/3 mL using 1% tryptone and 1% tryptone + 28 1% keratin media and those cells were added to broth tubes. Cell growth was again measured every 3 days for a total of 12 days using spectroscopy. Each measurement was obtained from harvesting broth tubes in triplicate and each experiment was repeated three times. Effect of Nitrate on Growth of B. dendrobatidis B. dendrobatidis was grown on 1% tryptone agar plates and zoospores were harvested after 8 days of growth as described above. Zoospores were diluted to a density of 165,000-330,000 cell/3 mL using 1% tryptone media that had been adjusted to various concentrations of NO3- [0 mg/L (1% tryptone), 5 mg/L, 10 mg/L, and 25mg/L] using solid NaNO3. Diluted cell suspensions were applied to cell culture 12-well plates and cell growth was monitored every 3 days for a total of 12 days. Similar to the previous experience, growth was measured by absorbance at 495 nm using approximately 2-3mL of the media harvested from each well. Each measurement was obtained from harvesting wells in triplicate and each experiment was repeated three times. Effect of Phosphate on Growth of B. dendrobatidis B. dendrobatidis was grown on 1% tryptone agar plates and zoospores were harvested after 8 days of growth as described above. Zoospores were diluted to a density of 165,000-330,000 cell/3 mL using 1% tryptone media that had been adjusted to various concentrations of PO4-3 [0 mg/L (1% tryptone), 0.05 mg/L, 0.2 mg/L, 0.4 mg/L, and 1mg/L] using solid Na2HPO4. Diluted cell suspensions were applied to cell culture 1229 well plates and cell growth was monitored every 3 days for a total of 12 days. Similar to the previous experience, growth was measured by absorbance at 495 nm using approximately 2-3mL of the media harvested from each well. Each measurement was obtained from harvesting wells in triplicate and each experiment was repeated three times. Statistical Analysis Statistical analysis was conducted using the statistical computing program R46,88,121. A linear model (LM) for each environmental factor (keratin in the in vitro system, keratin in in vitro broth tubes, pH, phosphate concentration, nitrate concentration) was used to test for the effect between each level of that environmental factor on the absorbance of the sample at day 12. The R code used for the analyses can be seen in Appendix B. For each linear model, all triplicate runs for each environmental factor (Appendix A) was utilized. For this analysis, the level of significance was set to α = 0.05. 30 Chapter III Results Creation of a Novel In Vitro System In order to determine if B. dendrobatidis can attach to and grow within submerged cell culture wells, zoospores were applied to the wells and growth was monitored for eight days. As shown by microscopy, zoospores successfully attached to the wells and transformed into germlings within the first 2 days (Figure 8A and B). From days 3-5. the newly-formed germlings transformed into zoosporangia and the zoosporangia produced new zoospores (Figure 8C-E). Zoospores continued to be produced and reattach over the next several days (Figure 8F-H). In all, these data suggest that B. dendrobatidis is able to complete its life cycle when grown in this submerged in vitro system. To further determine this organism’s success in completing its life cycle in this submerged in vitro system, specific structures and stages of the life cycle were identified using microscopy. On day 5, newly produced zoospores were observed in the tissue culture wells (Figure 9A). From days 3-8 when zoosporangia were maturing, both types of zoosporangia were observed (Figure 9B). The left arrow shows a developing ‘ 31 monocentric’ zoosporangium and the right arrow shows a developing ‘colonial’ zoosporangium. The colonial zoosporangium contained a septum, which divided the thallus body into two compartments for new zoospores. At days 3-5, rhizoid structures were formed by germlings and maturing zoosporangia (Figure 9C). At any time from days 4-8, zoospores were released from the mature zoosporangia and all that was left was a clear, empty zoosporangia with one (or multiple) discharge tube(s) from one side of the zoosporangia’s chitinous wall (Figure 9D). Different volumes of culture were tested to determine if inoculum size would make any difference in growth rate. Data was collected and quantified every 3 days for 6 days during B. dendrobatidis’s log growth phase in culture. Data was quantified by using hemocytometer cell counting (Figure 10A) and by measuring light absorbance of the culture at 495 nm (Figure 10B). Ultimately, volume did not make any significant differences in absorbance. Also, microscopic cell-counting and spectrophotometry produced very similar results. Since spectrometry allows for faster, more high-throughput acquisition of data, it was used for all further experiments. 32 Figure 8. Life cycle of B. dendrobatidis as shown from A-H. A= day 1: motile zoospores. B= day 2: germlings. C= day 3: developing zoosporangia/germlings. D-H= days 4-8: developed zoosporangia with note of newly produced zoospores at day 5 shown by black line arrow (Photo Credit to Amanda Layden) 33 Figure 9. Structures and stages of the life cycle of B. dendrobatidis as shown from A-D. A= zoomed in view of Day 5 from life cycle in tissue culture plates in vitro to show newly produced zoospore. B= left arrow shows a developing monocentric zoosporangium and right arrow shows a mature colonial zoosporangium with a septum dividing the thallus body into two compartments. C= germlings stained with crystal violet to show rhizoid structures noted by arrow. D= a clear, empty zoosporangium with a single discharge papillae (tube) noted by arrow (Photo Credit to Amanda Layden) 34 A 4,000,000 1mL/ well 3,500,000 3,000,000 2mL/ well Number of Cells 2,500,000 2,000,000 1,500,000 1,000,000 500,000 0 -500,000 0 -1,000,000 3 6 Days B 2.50 1mL/ well Optical Density 2.00 2mL/ well 1.50 1.00 0.50 0.00 0 3 6 Days Figure 10. Growth of B. dendrobatidis using this novel in vitro system. A= growth quantified using cell counts on a hemocytometer, B= growth quantified using absorbance (495 nm) values from a spectrophotometer 35 Effects of pH on the Growth of B. dendrobatidis Different pH values were tested to validate whether B. dendrobatidis would grow similarly in this novel in vitro system when compared to other in vitro models (1% tryptone broth and agar), in vivo models (host amphibians), and the natural environment 80,113 . Five pH values were chosen based on previously published data about this organism’s optimal growth environment 80,113. Growth of B. dendrobatidis in this novel in vitro system was observed in pH values ranging from approximately 5-9 (Figure 11). There was a significant difference among pH treatments in growth of B. dendrobatidis (LM; df=4,10; F=29.23; P=0) (Table 2). Overall, B. dendrobatidis grew well in pH values of 6 and 7 in this system, similarly to what it favors in the environment and in Optical Desntiy other in vitro systems. 1.40 5.24 1.20 6.11 1.00 7.24 0.80 8.1 9.23 0.60 0.40 0.20 0.00 -0.20 -0.40 0 3 6 9 12 Days Figure 11. Effect of pH values on growth of B. dendrobatidis in the novel in vitro system 36 Table 2. pH Analysis of Variance ID Residuals Df 4 10 Sum Sq 3.9771 0.3402 Mean Sq 0.99427 0.03402 F value 29.23 Pr (>F) 1.702e-05 *** Effects of Keratin on the Growth of B. dendrobatidis Previous in vitro studies with B. dendrobatidis suggest that its growth may be impacted by increased concentrations of tryptone80. Since tryptone is a stable product of protein digestion, other proteins were tested to determine if they have any effects on the growth of B. dendrobatidis. Addition of keratin to the 1% tryptone media and as a precoat on the tissue culture wells was tested to determine whether higher levels of protein affect the growth of B. dendrobatidis in our system (Figure 12A). Overall, B. dendrobatidis favored 1% tryptone media for growth. The 1% keratin pre-coat slightly decreased growth and the 1% tryptone + keratin media showed little to no growth of B. dendrobatidis. There was a significant difference among keratin novel, in vitro system treatments in growth of B. dendrobatidis (LM; df=2,6; F=22.608; P=0.001) (Table 3). Additionally, keratin added to 1% tryptone broth tubes (Figure 12B) and 1% tryptone agar plates (Figure 12C) showed similar inhibitory effects. There was a significant difference among keratin in vitro broth tube treatments in growth of B. dendrobatidis (LM; df=1,4; F=63.141; P=0.001) (Table 4). A second, unrelated protein (bovine serum albumin) was also added as a supplement to 1% tryptone agar to determine whether protein concentration in general is impacting fungal growth (Figure 13D). Similar to what was seen for keratin, the addition of bovine serum albumin showed little 37 to no growth of B. dendrobatidis when compared to the standard 1% tryptone broth and agar. BSA was also tested in the novel in vitro system one time (data not shown) but results for this were inconclusive and needs further investigation. These preliminary results might suggest that increased concentrations of protein may indeed inhibit growth of B. dendrobatidis, but they are inconclusive and further studies will need to be performed to verify the effect or lack of effect of protein concentration on growth of B. dendrobatidis. 38 A 1.60 1%T (control) 1.40 Pre-coat 1%K 1.20 1%T+K Optical Density 1.00 0.80 0.60 0.40 0.20 0.00 -0.20 0 3 -0.40 6 9 12 9 12 Days B 0.20 1%T 1%T+K Optical Density 0.15 0.10 0.05 0.00 0 -0.05 3 6 Days 39 C D Figure 12. Effect of protein on growth of B. dendrobatidis. A= quantitative data showing effect of keratin on growth on B. dendrobatidis in novel in vitro system. B= quantitative data showing effect of keratin on growth on B. dendrobatidis in broth tubes. C= image showing effect of keratin on growth on B. dendrobatidis when added to 1% tryptone agar. 1% tryptone agar is shown on the top row and 1% tryptone + keratin agar is shown on the bottom row. D= image showing effect of bovine serum albumin on growth of B. dendrobatidis when added to 1% tryptone agar. (Photo Credit to Amanda Layden) 40 Table 3. Keratin Broth Tubes Analysis of Variance ID Residuals Df 1 4 Sum Sq Mean Sq 0.0184704 .0184704 0.0011701 .00002925 F value 63.141 Pr (>F) 0.001358 ** F value 22.608 Pr (>F) 0.001608 ** Table 4. Keratin Analysis of Variance ID Residuals Df 2 6 Sum Sq 3.9274 00.5212 Mean Sq 1.96370 0.08686 Effect of Phosphate on the Growth of B. dendrobatidis Different concentrations of phosphate were tested to see their effect on the growth of B. dendrobatidis. Concentrations tested were 0 mg/L (1% tryptone), 0.05 mg/L, 0.2 mg/L, 0.4 mg/L, and 1.0 mg/L (Figure 13). Similar growth patterns were observed with all concentrations; however, at day 6, the 1.0 mg/L concentration showed a steeper spike in growth when compared to the other concentrations. Growth of the 1.0 mg/L concentration remained steady between days 6-9 until day 12 when there was a second spike in growth observed. There was no significant difference among phosphate concentration treatments in growth of B. dendrobatidis (LM; df=4,10; F=2.0192; P=0.1) (Table 5). Overall, data showed that higher concentrations (>1 mg/L) of phosphate led to increased growth of B. dendrobatidis when compared to the traditional 1% tryptone broth. 41 0 mg/L (1% tryptone) 0.05 mg/L 1.70 Optical Density 1.20 0.2 mg/L 0.4 mg/L 0.70 1.0 mg/L 0.20 -0.30 0 3 6 -0.80 9 12 Days Figure 13. Effect of phosphate concentration growth of B. dendrobatidis Table 5. Phosphate Analysis of Variance ID Residuals Df 4 10 Sum Sq 1.9829 2.4551 Mean Sq 0.49572 0.24551 F value 2.0192 Pr (>F) 0.1676 Effect of Nitrate on the Growth of B. dendrobatidis Different amounts of nitrate were tested to see their effect on the growth of B. dendrobatidis. Concentrations tested were 0 mg/L (1% tryptone), 5 mg/L, 10 mg/L, and 25 mg/L (Figure 14). Similar growth patterns were observed between all concentrations; however, at day 6 the 25 mg/L concentration showed a steeper spike in growth when compared to the other concentrations. After day 9, it was noted that the optical density of the 25 mg/L concentration decreased. Similarly, by day 12, all concentrations tested had decreased from the day 9 observations. There was no significant difference among nitrate concentration treatments in growth of B. dendrobatidis (LM; df=3,8; F=0.0805; P=0.1) 42 (Table 6). Overall, data showed that higher concentrations (> 25 mg/L) of nitrate may cause an initial increase of growth during log phase, and then lead to a decrease over time. 0 mg/L (1% tryptone) 0.60 5 mg/L 0.50 10 mg/L 25 mg/L Optical Density 0.40 0.30 0.20 0.10 0.00 0 3 6 -0.10 9 12 Days Figure 14. Effect of nitrate concentration on growth of B. dendrobatidis Table 6. Nitrate Analysis of Variance ID Residuals Df 3 8 Sum Sq 0.01142 0.37847 Mean Sq 0.003808 0.047309 43 F value 0.0805 Pr (>F) 0.9688 Chapter IV Discussion Chytridiomycosis is an emerging infectious wildlife disease that is continuing to cause massive declines in amphibian populations on a global scale. As mentioned, B. dendrobatidis infects over 350 amphibian species and has been implicated in driving the decline of over 200 of these species32,103. B. dendrobatidis induced chytridiomycosis was first described 20 years ago and several studies have documented B. dendrobatidis growth and development at morphological and ultrastructural levels6,7,40,112. Understanding what environmental factors affect the growth of B. dendrobatidis is important in figuring out how to treat and prevent this disease. Aside from this, having the ability to utilize a novel, high-throughput in vitro system would enable researchers to study these factors more efficiently and in more detail by being able to look more closely at what factors effect this pathogen’s life cycle. This is the first study utilizing tissue culture plates as a novel, submerged in vitro growth system to test different environmental factors on the growth of this emerging environmental pathogen. 44 Creation of a Novel in vitro System B. dendrobatidis has been studied for decades; however, there is still much that is not known about its basic biology. To date, in vivo experimentation is still widely utilized in B. dendrobatidis research in order to understand host-pathogen interactions116. Others have turned to various types of ex vivo systems that involve inoculated amphibian skin explants. A study done by Verbrugghe et al. discussed pathogen-host interactions using primary amphibian keratinocytes, followed by internalization of B. dendrobatidis in these host cells116. They also developed an invasion model using X. laevis kidney epithelial cell line A6 mimicking the complete B. dendrobatidis colonization cycle in vitro116. That said, although in vivo research has tremendous value for understanding disease processes, the availability of a cost-effective in vitro system could provide a first line tool to gain insight into host-pathogen interactions and understanding the pathogen itself, which will reduce the number of animals used in infection experiments99,116. Understanding what factors can affect a pathogen’s life cycle is important in understanding how it’s able to cause disease. Infectious diseases are commonly studied in vitro by assessing the interaction of a pathogen with host cells116; however, this study showed that B. dendrobatidis is capable of completing its life cycle in a submerged, in vitro environment without the use of host cells. Since this pathogen has a stage of its life cycle where it is not attached to amphibian skin, understanding its growth behavior outside of host cells is extremely important. In vitro studies offer the advantage of being simplistic and easy to perform and repeat when studying a pathogen’s behavior in a specific environment or answering unknown questions regarding a pathogen. Also, it is relatively simple to determine if there are any environmental factors (temperature, pH, 45 salinity, etc), biotic triggers, or even purified host defenses that affect its life cycle or cell structure. As mentioned earlier, the amphibian host has both innate and acquired immune components that contribute to fighting chytridiomycosis infection113. To date, approximately 40 anuran antimicrobial peptides inhibiting B. dendrobatidis have been discovered and both purified and natural mixtures of these antimicrobial peptides have effectively inhibited in vitro broth growth of B. dendrobatidis113. Although these antimicrobial peptides have shown results in in vitro broth studies, this method does not allow one to determine how these antimicrobial peptides are inhibiting B. dendrobatidis’s life cycle or at what stage the life cycle is being affected. Additionally, having a novel, submerged in vitro assay similar to how this pathogen would grow in vivo on amphibian skin would be time efficient, cost efficient, and require no animal test subjects or field studies that could lead to low prevalence data and potential bias. This study showed that not only did B. dendrobatidis attach and grow successfully in this novel, submerged in vitro system, it proved that this system can be successfully utilized to test environmental factors, other aspects of B. dendrobatidis’s life cycle, genetic factors that control its life cycle, attachment proteins, antifungal drugs, water quality parameters, and a variety of other factors on the growth of this pathogen. Effects of pH, Nitrate and Phosphate on the Growth of B. dendrobatidis Understanding associations between B. dendrobatidis infection dynamics and environmental factors is important for mitigating adverse effects of the chytrid on amphibian populations56. The data in this study showed that B. dendrobatidis favored an 46 environmental pH of roughly 6-7. A study done by Karvemo et al. discussed that pond pH was strongly positively associated with B. dendrobatidis infection prevalence, particularly when pH was higher than 6.556. This is consistent with observations of increases in B. dendrobatidis growth rates with increases in pH in previous experimental and field studies10,22,56,80. Environmental pH is influenced by abiotic (ex. acidneutralizing capacity) and biotic (ex. organic carbon, aquatic plant community) characteristics of a system22,42,44,61. A lower pH can inhibit microbial metabolism21,22 and changes in pH are related to the acid-neutralizing capacity in a system. This is strongly tied to the amount of organic carbon present22,44,61 which in turn, is an important nutrient for aquatic fungi22,38. Although it is not clear as to why B. dendrobatidis does not grow well in a lower pH, Chestnut et al. suggested that it may be due to reductions in metabolic rates of the fungus and organic carbon substrates, which are important nutrients for aquatic fungi in low pH environments22,56. Aside from pH playing a major role in metabolic rates of fungi, it’s possible that B. dendrobatidis may favor an environmental pH of roughly 6-7 because that may be the external pH of amphibian skin. However, further investigation on this topic is needed to confirm or deny this hypothesis. As mentioned above, changes to the chemical composition of the environmental water source has the potential to drastically alter the growth of microorganisms like B. dendrobatidis. Nitrogen (N) and phosphorus (P) are two important and essential nutrients for healthy soil and aquatic environments. In addition, nitrogen and phosphorus are two of the many additives found in traditional fertilizers used for lawn care and plant food and fertilizers used to enhance agricultural productivity. Fertilizers influence both the aboveground biomass and the belowground microbial biomass124. Soil microbial 47 communities consist mainly of bacteria, fungi, and archaea and play critical roles in ecosystem function and regulate key processes such as carbon and nitrogen cycles3,15,124. Determining whether nitrogen and phosphorus fertilization impacts a microbial community is difficult because the soil microbe communities in various ecosystems are different and thus their responses to similar fertilizations might also be different124. A well-known outcome of an increase of nitrogen and phosphorus in aquatic environments is known as an algal bloom27. Algae that undergo these algal blooms are classified as microalgae, which includes dinoflagellates and bacillariophyta (diatoms)27. In the past several decades, a growing number of studies concerning the environmental factors of algal bloom outbreaks and decline have been explored125. According to Zhang et al., excessive exogenous nitrogen and phosphorus, high temperature, and adequate light intensity have been identified as major abiotic triggers of algal blooms125. Although algal blooms are seen only in aquatic environments, there are other microorganisms (bacteria, fungi, archaea) that coexist in these ecosystems and may also be affected by these environmental factors. As mentioned, B. dendrobatidis is found in a variety of water sources and moist soil environments. Similar to algae, B. dendrobatidis’s growth is known to be affected by abiotic triggers such as temperature fluctuations80. That said, with chytridiomycosis infection and the use of fertilizers in agriculture increasing over the last few decades, understanding if a similar event occurs with B. dendrobatidis is important to determine if these abiotic factors cause changes in the growth of this pathogen in the environment. 48 Overall, the data in this study showed that the lower concentrations of nitrate and phosphate added to 1% tryptone growth media did not have any effect on the growth of B. dendrobatidis. The higher concentrations tested (> 25 mg/L NO3 and > 1.0 mg/L PO4) indicated slight increased growth of B. dendrobatidis, but not enough to make a statistical significance. According to the EPA, naturally occurring amounts of nitrogen and phosphorus vary substantially between water sources109. Appropriate reference levels for normal water quality range from 0.12 to 2.2 mg/L total nitrogen and 0.01 to 0.075 mg/L total phosphorus109; however, nuisance algal growths are not uncommon in rivers and streams below the low reference level (0.1 mg/L) for phosphorus. Additionally, the EPA noted that excess nitrate is not toxic to aquatic life, but increased nitrogen may result in overgrowth of algae, which can decrease the dissolved oxygen content of the water, thereby harming or killing fish and other aquatic species108,109. Furthermore, this indicates that nitrate and phosphate may not cause a significant impact on the growth of this pathogen, but further exploration of this hypothesis is needed. To do so, a wider range of concentrations of nitrate and phosphate should be tested on the growth of this pathogen to determine if these environmental factors have an effect on the growth of this pathogen. Effect of Keratin on the Growth of B. dendrobatidis As mentioned, chytridiomycosis is an emerging infectious wildlife disease that affects the keratinized epidermal cells of the amphibian epithelium, which is an extremely important organ in amphibians. In infected amphibians, B. dendrobatidis is found in the cells of the epidermis and pathological abnormalities include a thickening of 49 the outer layer of the skin7,96. Cutaneous fungal infections in other vertebrates are not usually lethal, but amphibian skin is unique because it is physiologically active, tightly regulating the exchange of respiratory gases, water, and electrolytes96. That said, the physiological importance of the skin makes amphibians particularly vulnerable to skin infections96. Since this is a cutaneous infection of amphibians, a major theory regarding B. dendrobatidis is that it utilizes keratin as a nutrient source23. This is a major topic of discussion because this pathogen infects the keratinocytes of the stratum corneum and can only infect the keratinized mouthparts of tadpoles23. This study showed that keratin being added to 1% tryptone broth tubes in vitro, 1% tryptone agar in vitro, and to 1% tryptone in the novel, submerged in vitro system had a statistically-significant negative impact on the growth of B. dendrobatidis. Ultimately, adding keratin to the 1% tryptone media resulted in a decrease in growth of B. dendrobatidis when compared to the standard 1% tryptone media. It’s possible that keratin might be impacting the growth of B. dendrobatidis by altering cell signaling, cellto-cell communication, or some other unknown mechanism. It is also possible that the free form of keratin is not a viable nutrient source as compared to keratinized skin cells. Despite the increasing scientific attention to chytridiomycosis, mechanisms that influence host characteristics and B. dendrobatidis population densities still remain poorly understood115. Quorum sensing (QS) is a mechanism of cell-to-cell communication that allows unicellular organisms to determine their population density in order to regulate their population behavior, including growth2,115. A study done by Verbrugge et al. showed that B. dendrobatidis is capable of controlling its cell 50 populations, in which individual cells communicate with each other by secreting tryptophol in order to assess the population density and to coordinate their growth response115. When a certain density is achieved, they start producing tryptophol with an autostimulatory mode of action, and when tryptophol reaches high concentrations in the exponential/stationary phase of growth, this results in growth reduction115. According to Verbrugge et al., it could be suggested that nutrient limitation occurs during these growth phases, leading to growth decreases115. That said, when keratin was added to 1% tryptone media, there were no indications of a log growth phase. This suggests the possibility that keratin may not be a necessary nutrient for B. dendrobatidis. It’s also possible that B. dendrobatidis did not recognize the added keratin, due to it not being expressed within or on cells of the amphibian nonprofessional immune cells (keratinocytes, fibroblasts)41. Similarly, it could also be possible that an increase of protein concentration could potentially be causing B. dendrobatidis to halt its growth, cause cell death, or act as a signal for this fungus to switch from growth to invasion mechanisms. B. dendrobatidis is capable of growing on a variety of growth media in vitro such as 1% keratin agar, frog skin agar, feathers, geese paws, and chitinous carapaces of crustaceans36,66,71,80,112,113. Although B. dendrobatidis grows well on these substances, it grows best in tryptone or peptonized milk in both agar and broth in vitro66,113. Tryptone and peptonized milk are both digests of casein, a protein readily found in mammalian milk. A study done by Piotrowski et al. discussed that B. dendrobatidis does not require sugars other than those that were added to the 1% tryptone and that high percentages of sugar or tryptone (greater than 2%) hinder growth80. Similarly, throughout our study, it 51 was observed that 1% tryptone media with the addition of 1% keratin (a roughly 2% protein-rich growth media) hindered growth of this pathogen. Conclusions The overall increase in chytridiomycosis over the last few decades has had a severe impact in amphibian populations globally. In vivo studies on chytridiomycosis are valuable to obtain pathogen-host interactions; however, in vitro studies provide a faster, inexpensive, high-throughput way to test multiple environmental factors at once that could potentially be impacting growth of B. dendrobatidis. The results discussed in this paper and others suggest that B. dendrobatidis may be impacted by abiotic factors such as temperature, environmental pH, and increased protein concentrations. Other microorganisms, such as microalgae, are affected by similar abiotic factors and it is important to understand whether these abiotic factors also cause an impact on this pathogen as well. Future Studies As this was the first study utilizing tissue culture plates as a novel, submerged in vitro assay, there is ample opportunity to continue using this assay to test similar and new environmental factors that could potentially impact the growth of B. dendrobatidis. Although nitrogen, phosphorus, and pH are important components of water quality parameters, there are others that play a significant role as well. Future studies could look at some of these other important water quality parameters such as ammonium, dissolved 52 oxygen, alkalinity, and water hardness. The most interesting discovery of these results was that keratin concentration seemed to have a negative effect on the growth of B. dendrobatidis. Future studies could utilize this new-found information and determine at what stage the life cycle is being altered, or test if there are other amphibian surface proteins or generic proteins that show a similar result. With the availability of this system and the results of this study, it is important to continue researching what environmental factors, other aspects of B. dendrobatidis’s life cycle, genetic factors that control its life cycle, attachment proteins, antifungal drugs, water quality parameters, and a variety of other factors that have an effect on the growth of this fungus. Additionally, it is also important to continue observing a wider range of concentrations of nitrogen, phosphorus, and proteins to determine if any other concentrations outside of what was observed in this study show a similar or adverse effect on the growth of this pathogen. 53 Literature Cited 1. Abramyan J, Stajich JE. Species-specific chitin binding module 18 expansion in the amphibian pathogen Batrachochytrium dendrobatidis. MBio. 2012;3:e00112–e00150 2. Albuquerque P, Casadevall A. Quorum sensing in fungi – a review. Med. Mycol. 2012;50:337-345. doi: 10.3109/13693786.2011.652201 3. Balser TC, Firestone MK. Linking microbial community composition and soil processes in a California annual grassland and mixed-conifer forest. Biogeochemistry. 2005;73(2):395-415. doi: 10.1007/s10533-004-0372-y 4. Batrachochytrium dendrobatidis (Bd). (n.d.). Retrieved from https://www.cabi.org/isc/datasheet/109124#todistribution 5. Bell BD, Carver S, Mitchell NJ, Pledger S. The recent decline of a New Zealand endemic: how and why did populations of Archey’s frog Leiopelma archeyi crash over 1996-2001? Biological Conservation. 2004;120(2):189-199 6. Berger L, Hyatt AD, Speare R, Longcore JE. Life cycle stages if the amphibian chytrid Batrachochytrium dendrobatidis. Diseases of Aquatic Organisms. 2005;68:51-63. doi: 10.3345/dao068051 7. Berger L, Speare R, Daszak P, Green DE, Cunnungham AA, Goggin CL, Slocombe R, Ragan MA, Hyatt AD, McDonald KR, Hines HB, Lips KR, Marantelli G, Parkes H. Chytridiomycosis causes amphibian mortality associated with population declines in the rain forests of Australia and Central America. Proc Natl Acad Sci U S A. 1998;95:9031-9036. doi: 10.1073/pnas.95.15.9031 8. Berger L, Speare R, Hyatt A. Chytrid fungi and amphibian declines: Overview, implications and future directions. In: Campbell A, editor. Declines and disappearances of Australian frogs. Canberra, Australia: Biodiversity Group Environment Australia; 1999. p. 23–33 9. Berger L, Speare R, Skerratt L (2005) Distribution of Batrachochytrium dendrobatidis and pathology in the skin of green tree frogs (Litoria caerulea) with severe chytridiomycosis. Dis Aquat Organ 68:65–70 54 10. Blooi M, Laking AE, Martel A, Haesebrouck F, Jocque M, Brown T, Green S, Vences M, Bletz MC, Pasmans F. Host niche may determine disease-driven extinction risk. PloS one. 2017; 12: e0181051 https://doi.org/10.1371/journal.pone.0181051 PMID: 28704480 11. Bonaccorso E, Guayasamin JM, Méndez D, Speare R. Chytridiomycosis in a Venezuelan amphibian (Bufonidae: Atelopus cruciger). Herpetol Rev. 2003;34:331–4 12. Bosh J, Martínez-Solano I, García-Prís M. Evidence of a chytrid fungus infection involved in the decline of the common midwife toad (Alytes obstetricans) in protected areas of central Spain. Biol Conserv. 2000;97:331–7 13. Bovero S, Sotigiu G, Angelini C, Doglio S, Gazzaniga E, Cunningham AA, Garner TWJ. Detection of chytridiomycosis caused by Batrachochytrium dendrobatidis in the endangered Sardinian newt (Euproctus platycephalus) in southern Sardinia, Italy. Journal of Wildlife Diseases. 2008;44(3):712-715. http://www.wildlifedisease.org 14. Boyle DG, Hyatt AD, Daszak P, Berger L, Longcore JE, Porter D, Hengstberger SG, Olsen V. Cryo-archiving of Batrachochytrium dendrobatidis and other chytridiomycetes. Dis Aquat Organ. 2003;56(1):59-64 15. Bragazza L, Bardgett RD, Mitchell EA, Buttler A. Linking soil microbial communities to vascular plant abundance along a climate gradient. New Phytol. 2015;205(3):1175-1182 16. Braun A. Uber Chytridium, eine Gattung einzelliger Schmarotzergewӓchse auf Algen und Infusorien. Deutsche Akademie der Wissenschaften. 1855;1855: 21–83 17. Brooks RT. Declines in summer bat activity in central New England 4 years following the initial detection of white-nose syndrome. Biodiversity and Conservation. 2011;20(11):2537-2541. http://www.springerlink.com/content/9716632784163261/ 18. Brucker RM, Harris RN, Schwantes CR, Gallaher TN, Flaherty DC, Lam BA, Minbiole KPC. Amphibian chemical defense: antifungal metabolites of the microsymbiont Janthinobacterium lividum on the salamander Plethodon cinereus. J Chem Ecol. 2008;34:1422–1429 55 19. Burgmann HF, Widmer VW, Zeyer, J. New molecular screening tools for analysis of free-living diazotrophs in soil. Appl. Environ. Microbiol. 2004;70:240–247 20. Bustamante MR, Ron SR, Coloma LA. Cambios en la diversidad en siete comunidades de anuros en los Andes de Ecuador. Biotropica. 2005;37:180– 189 21. Chamier AC. Effect of pH on microbial degradation of leaf litter in seven streams of the English Lake District. Oecologia. 1987;71: 491–500 22. Chestnut T, Anderson C, Popa R, Blaustein AR, Voytek M, Olson DH, et al. Heterogeneous occupancy and density estimates of the pathogenic fungus Batrachochytrium dendrobatidis in waters of North America. PLoS One. 2014; 9: e106790 https://doi.org/10.1371/journal.pone.0106790 PMID: 25222122 23. Daszak, P, Berger L, Cunningham AA, Hyatt AD, Greene DE, Speare R. Emerging infectious diseases and amphibian population declines. Emerging Infectious Diseases. 1999;5:735-748 24. Daszak, P. Cunningham AA, Hyatt AD. Emerging Infectious Diseases of Wildlife-Threats to Biodiversity and Human Health. Science. 2000;287(5452), 443–449. doi: 10.1126/science.287.5452.443 25. Data and Surveillance. (2019, November 22). Retrieved from https://www.cdc.gov/lyme/datasurveillance/index.html?CDC_AA_refVal=https://ww w.cdc.gov/lyme/stats/index.html 26. Delahay RJ, Smith, GC, Hutchings MR. (2009). The science of wildlife disease management. Management of Disease in Wild Mammals. 2009;(pp.1–8). Tokyo,Japan: Springer 27. El Gamal AA. Biological importance of marine algae. Saudi pharmaceutical journal: SPJ: the official publication of the Saudi Pharmaceutical Society. 2010;18(1):1-25. doi: 10.1016/j.jsps.2009.12.001 56 28. Erickson RA, Thogmartin WE, Diffendorfer JE, Russel RE, Szymanski JA. Effects of wind energy generation and white-nose syndrome on the viability of the Indiana bat. Peerj. 2016;4:e2830 29. Farrer RA et al. Multiple emergences of genetically diverse amphibian-infecting chytrids include a globalized hypervirulent recombinant lineage. Proc. Natl Acad. Sci. USA. 2011;108(18):732 – 18 736. (doi:10.1073/pnas.1111915108) 30. Farrer RA, Henk DA, Garner TW, Balloux F, Woodhams DC, Fisher MC. Chromosomal copy number variation, selections and uneven rates of recombination reveal cryptic genome diversity linked to pathogenicity. PloS Genet. 2013;9: e1003703 31. Fisher MC, Garner TWJ. The relationship between the introduction of Batrachochytrium dendrobatidis, the international trade in amphibians and introduced amphibian species. Fungal Biol Rev. 2007;21:2–9. 22 32. Fisher MP, Garner TWJ, Walker SF. Global emergence of Batrachochytrium dendrobatidis and amphibian chytridiomycosis in space, time, and host. Annual Review of Microbiology. 2009;63: 291ñ310 33. Frick WF, Pollock JF, Hicks AC, Langwig KE, Reynolds DS, Turner CG, Butchkoski CM, Kunz TH. An emerging disease causes regional population collapse of a common North American bat species. Science (Washington). 2010;329(5992):679682. http://www.sciencemag.org 34. Frick WF, Puechmaille SJ, Hoyt JR, Nickel BA, Langwig KE, Foster JT, Barlow KE, Bartonicka T, Feller D, Haarsma AJ, Herzog C, Horàcek I, Kooij J van der, Mulkens B, Petrov B, Reynolds R, Rodrigues L, Stihler CW, Turner CG, Kilpatrick AM. Disease alters macroecological patterns of North American bats. Global Ecology and Biogeography. 2015;24(7):741-749. http://onlinelibrary.wiley.com/journal/10.1111/(ISSN)1365-2468 35. Gaertner A. U¨ ber das Vorkommen neiderer Erdphycomyceten in Afrika, Schweden und an einigen mitteleuropӓischen Standorten. Archiv für Mikrobiologie 1954;21: 4– 56 57 36. Garmyn A, Van Rooij P, Pasmans F, Hellebuyck T, Van Den Broeck W, Haesebrouck F, Martel A (2012) Waterfowl: potential environmental reservoirs of the chytrid fungus Batrachochytrium dendrobatidis. PLoS One 7:e35038 37. Gewin V. 2008. Riders of a modern-day ark. PLoS Biol. 6:e24 38. Gleason FH, Kagami M, Lefevre E, Sime-Ngando T. The ecology of chytrids in aquatic ecosystems: roles in food web dynamics. Fungal Biol Rev. 2008;22: 17–25 39. Gower DJ, Doherty-Bone T, Loader SP, Wilkinson M, Kouete MT, Tapley B, Orton F, Daniel OZ, Wynne F, Flach E, Müller H, Menegon M, Stephen I, Browne RK, Fisher MC, Cunningham AA, Garner TW (2013) Batrachochytrium dendrobatidis infection and lethal chytridiomycosis in caecilian amphibians (Gymnophiona). EcoHealth 10:173–183 40. Greenspan SE, Longcore JE, Calhoun AJ. Host invasion by Batrachochytrium dendrobatidis: fungal and epidermal ultrastructure in model anurans. Dis Aquat Organ. 2012;100:201–210 41. Grogan LF, Robert J, Berger L, Skerratt LF, Scheele BC, Castley JG, Newell DA, McCallum HI. Review of the Amphibian Immune Response to Chytridiomycosis, and Future Directions. Front. Immunol. 2018;9:2536. doi:10.3389/fimmu.2018.02536 42. Halstead BG, Tash JC. Unusual diel pHs in water as related to aquatic vegetation. Hydrobiologia. 1982;96: 217–224 43. Hanlon SM, Lynch KJ, Kerby J, Parris MJ (2015) Batrachochytrium dendrobatidis exposure effects on foraging efficiencies and body size in anuran tadpoles. Dis Aquat Organ 112:237–242 44. Herlihy AT, Kaufmann PR, Mitch ME. Stream chemistry in the eastern United States: 2. Current sources of acidity in acidic and low acid-neutralizing capacity streams. Water Resour Res. 1991;27: 629–642 58 45. Hibbett DS, Binder M, Bischoff JF, Blackwell M, Cannon PF, Eriksson OE, Huhndorf S, James T, Kirk PM, Lücking R, Thorsten Lumbsch H, Lutzoni F, Matheny PB, McLaughlin DJ, Powell MJ, Redhead S, Schoch CL, Spatafora JW, Stalpers JA, Vilgalys R, Aime MC, Aptroot A, Bauer R, Begerow D, Benny GL, Castlebury LA, Crous PW, Dai YC, Gams W, Geiser DM et al. (2007) A higher-level phylogenetic classification of the Fungi. Myc Res 3:509–547 46. Hope RM. Rmisc: Rmisc: Ryan Miscellaneous. 2013 47. Huang R, McGrath SP, Hirsch PR, Clark IM, Storkey J, Wu L, Zhou J, Liang Y. Plant-microb networks in soil are weakened by century-long use of inorganic fertilziers. Microbial Biotechnology. 2019;12(6):1464-1475. doi: 10.1111/17517915.13487 48. James TY, Letcher PM, Longcore JE, Mozley-Standridge SE, Porter D, Powell MJ, Griffith GW, Vilgalys R. A molecular phylogeny of the flagellated fungi (Chytridiomycota) and description of a new phylum (Blastocladiomycota). Mycologia. 2007;98(6):860-871. doi: 10.3852/mycologia.98.6.860 49. James TY, Litvintseva AP, Vilgalys R, Morgan JAT, Taylor JW, Fischer MC, et al. Rapid Global Expansion of the Fungal Disease Chytridiomycosis into Declining and Healthy Amphibian Populations. PLoS Pathog. 2009;5(5): e10000458. doi: 10.1371/journal.ppat.1000458 50. Johnson ML, Berger L, Philips L, Speare R. Fungicidal effects of chemical disinfectants, UV light, desiccation, and heat on amphibian chytrid Batrachochytrium denedrobatidis. Dis Aquat Organ. 2003;57:255-260. doi: 10.3354/dao057255 51. Johnson ML, Speare R (2003) Survival of Batrachochytrium dendrobatidis in water: quarantine and disease control implications. Emerg Infect Dis 9:922–925 52. Johnson ML, Speare R (2005) Possible modes of dissemination of the amphibian chytrid Batrachochytrium dendrobatidis in the environment. Dis Aquat Organ 65:181–186 59 53. Kahindi JHP., et al. Agricultural intensification, soil biodiversity and ecosystem function in the tropics: the role of nitrogen-fixing bacteria. Appl. Soil Ecol. 1997;6:55–76 54. Karesh WB, Cook RA, Bennett EL, Newcomb J. Wildlife Trade and Global Disease Emergence. Emerging Infectious Diseases. 2008;11(7):1000–1002. doi: 10.3201/eid1107.020194 55. Karling JS. Some zoosporic fungi of New Zealand. III. Phlyctidium, Rhizophydium, Septosperma, and Podochytrium. Sydowia. 1967;20: 74–85 56. Kärvemo S, Meurling S, Berger D, Höglund J, Laurila A. Effects of host species and environmental factors on the prevalence of Batrachochytrium dendrobatidis in northern Europe. PLoS ONE. 2018;13(10): e0199852. https://doi. org/10.1371/journal.pone.0199852 57. Lam BA, Walke J, Vredenburg VT, Harris RN. Proportion of individuals with antiBatrachochytrium dendrobatidis skin bacteria is associated with population persistence in the frog Rana muscosa. Biol Conserv. 2010;143:529–531 58. LaMarca E, Lips KR, Lötters S, Puschendorf R, Ibáñ ez R, et al. Catastrophic population declines and extinctions in Neotropical harequin frogs (Bufonidae: Atelopus). Biotropica. 2005;37:190–201 59. Letcher PM, McGee PA, Powell MJ. Zoosporic fungi from soils of New South Wales, Australia. Australasian Mycologist 2004a;22: 99–115 60. Letcher PM, Powell MJ, Churchill PF, Chambers JG. Ultrastructural and molecular phylogenetic delineation of a new order, the Rhizophydiales (Chytridiomycota). Mycological Research. 2006;110:898-915 61. Liator IM, Thurman EM. Acid neutralizing processes in an alpine watershed front range, Colorado, U.S.A.—1: Buffering capacity of dissolved organic carbon in soil solutions. Appl Geochem. 1988;3: 645–652 60 62. Lips KR. Mass mortality and population declines of anurans at an upland site in western Panama. Conservation Biology. 1999;13(1):117-125 63. Lips KR, Brem F, Brenes R, Reeve JD, Alford RA, Voyles J, Carey C, Livo L, Pessier AP, Collins JP. Emerging infectious disease and the loss of biodiversity in a Neotropical amphibian community. Proceedings of the National Academy of Science of the United States of America. 2006;103(9):3165-3170. http://www.pnas.org/ 64. Lips KR, Diffendorfer J, Mendelson JR III, Sears MW. Riding the wave: Reconciling the roles of disease and climate change in amphibian declines. PLoS Biol. 2008;6(3): e72. doi:10.1371/journal.pbio.0060072 65. Longcore JE. Morphology and zoospore ultrastructure of Lacustromyces hiemalis gen. et sp. nov. (Chytridiales). Can J Bot. 1993;71:414-425 66. Longcore JE, Pessier AP, Nichols DK. Batrachochytrium dendrobatidis gen. et sp. Nov., a Chytrid Pathogenic to Amphibians. Mycologia. 1999;91(2):219. doi: 10.2307/3761366 67. Lorch JM et al. Snake fungal disease: an emerging threat to wild snakes. Phil. Trans. R. Soc. B. 2016;371: 20150457. http://dx.doi.org/10.1098/rstb.2015.0457 68. Lynch JD, Grant T. Dying frogs in western Colombia: catastrophe or trivial observation? Rev Acad Colomb Cienc. 1988;22:149–152 69. Martel A, Spitzen-van der Sluijs A, Blooi M, Bert W, Ducatelle R, Fisher MC, Woeltjes A, Bosman W, Chiers K, Bossuyt F, Pasmans F (2013) Batrachochytrium salamandrivorans sp. nov. causes lethal chytridiomycosis in amphibians. Proc Natl Acad Sci U S A 110:15325–15329 70. Mcdonald, JL, Robertson A, Silk MJ. Wildlife disease ecology from the individual to the population: Insights from a long‐term study of a naturally infected European badger population. Journal of Animal Ecology. 2017;87(1), 101–112. doi: 10.1111/1365-2656.12743 61 71. McMahon TA, Brannelly LA, Chatfield MWH, Johnson PTJ, Joseph MB, McKenzie VJ, Richards-Zawacki CL, Venesky MD, Rohr JR. Chytrid fungus Batrachochytrium dendrobatidis has nonamphibian hosts and releases chemicals that cause pathology in absence of infection. Proc Natl Acad Sci U S A. 2013;110:210–215 72. Miller CE. Annotated list of aquatic phycomycetes from Mountain Lake Biological Station, Virginia. Virginia Journal of Science. 1965;14: 219–228 73. Muths E, Corn PS, Pessier AP, Green DE. Evidence for disease-related amphibian decline in Colorado. Biological Conservation. 2003;110(3):357-365 74. Myers JM, Ramsey JP, Blackman AL, Nichols EA, Minbiole KPC, Harris RN. Synergistic inhibition of the lethal fungal pathogen Batrachochytrium dendrobatidis: the combined effect of symbiotic bacterial metabolites and antimicrobial peptides of the frog Rana muscosa. J Chem Ecol. 2010;8:958–965 75. National Wildlife Health Center. (n.d.). Retrieved from https://www.usgs.gov/centers/nwhc/science/emerging-wildlife-diseases 76. Omar SA, Abd-Alla MH. Effect of pesticides on growth, respiration and nitrogenase activity of Azotobacter and Azospirillum. WorldJ. Microbiol. Biotechnol. 1992;8:326–328 77. Orr CH, James A, Leifer C, Cooper JM, Cummings SP. Diversity and Activity of Free-Living Nitrogen-Fixing Bacteria and Total Bacteria in Organic and Conventionally Managed Soils. Applied and Environmental Microbiology. 2011;77(3):911-919. doi: 10.1128/AEM.01250-10 78. Pessier AP. Amphibian chytridiomycosis. In: Fowler ME, Miller ER (eds) Zoo and Wild Animal Medicine. 2008;Current therapy, vol 6. Saunders Elsevier, St. Louis, pp 137–143 79. Phillips BL, Puschendorf R. Do pathogens become more virulent as they spread? Evidence from the amphibian declines in Central America. Proceedings of the Royal Society B: Biological Sciences. 2013;280(1766), 20131290. doi: 10.1098/rspb.2013.1290 62 80. Piotrowski JS, Annis SL, Longcore JE. Physiology of Batrachochytrium dendrobatidis, a chytrid pathogen of amphibians. Mycologia. 2004;96(1):9-15. doi: 10.1080/15572536.2005.11832990 81. Pounds JA, Bustamante MR, Coloma LA, Consuegra JA, Fogden MPL, et al. Widespread amphibian extinctions from epidemic disease driven by global warming. Nature. 2006;39: 161–167. 82. Pseudogymnoascus destructans (white-nose syndrome fungus) (n.d.) Retrieved from https://www.cabi.org/isc/datasheet/119002#todiseasesTable 83. Puschendorf R, Bolaños F (2006) Detection of Batrachochytrium dendrobatidis in Eleutherodactylus fitzingeri: effects of skin sample location and histologic stain. J Wildl Dis 42:301–306 84. Quellet M, Mikaelian I, Pauli BD, Rodrique J, Green DM. Historical evidence of widespread chytrid infection in North American amphibian populations. 2003 Joint Meeting of Ichthyologists and Herpetologists, 26 June–1 July 2003, Manaus, Amazonas, Brazil [cited 2004 April 10]. Available from http://lists.allenpress.com/ asih/meetings/2003/abstracts_IV_2003.pdf 85. Rachowicz LJ, Knapp RA, Morgan JAT, Stice MJ, Vredenburg VT, Parker JM, Briggs CJ. Emerging infectious disease as a proximate cause of amphibian mass mortality. Ecology. 2006;87(7):1671-1683 86. Rachowicz LJ, Vredenburg VT (2004) Transmission of Batrachochytrium dendrobatidis within and between amphibian life stages. Dis Aquat Organ 61:75–83 87. Ramsey JP, Reinert LK, Harper LK, Woodhams DC, Rollins-Smith LA. Immune defenses against Batrachochytrium dendrobatidis, a fungus Linked to global amphibian declines, in the South African Clawed Frog, Xenopus laevis. Infect Immun 2010;78:3981–3992 88. R Core Team. R: A Language and Environment for Statistical Computing. R Foundation for Statistical Computing; 2018. 63 89. Retallick RWR, McCallum H, Speare R. 2004. Endemic infection of the amphibian chytrid fungus in a frog community postdecline. PLoS Biol. 2:1965–71 90. Reynolds RJ, Powers KE, Orndorff W, Ford WH, Hobson CS. Changes in rates of capture and demographics of Myotis septentrionalis (Northern Long-eared Bat) in western Virginia before and after onset of white-nose syndrome. Northeastern Naturalist. 2016;23(2):195-204. http://www.bioone.org/loi/nena 91. Rollins-Smith LA. The role of amphibian antimicrobial peptides in protection of amphibians from pathogens linked to global amphibian declines. Biochim Biophys Acta. 2009;1788:1593–1599 92. Rollins-Smith LA, Reinert LK, Miera V, Conlon JM. Antimicrobial peptide defenses of the Tarahumara frog, Rana tarahumarae. Biochem Biophys Res Comm. 2002;297:361–7 93. Ron SR, Duellman WE, Coloma LA, Bustamante MR. Population declines of the Jambato toad Atelopus ignescens (Anura: Bufonidae) in the Andes of Ecuador. J Herpetol. 2003;37: 116–126. 94. Rosenblum EB, James TY, Zamudio KR, Poorten, TJ, Ilut D, Rodriquez D, Eastman JM, Richards-Hrdlicka K, Joneson S, Jenkinson TS, Longcore JE, Parra Olea G, Toledo LF, Arellano ML, Medina EM, Restrepo S, Flechas SV, Berger L, Briggs CJ, Stajich JE. Complex history of the amphibian-killing chytrid fungus revealed with genome resequencing data. Proc Natl Acad Sci USA. 2013;110:9385-9390 95. Rosenblum EB, Stajich JE, Maddox N, Eisen MB. Global gene expression profiles for life stages of the deadly amphibian pathogen Batrachochytrium dendrobatidis. Proc Natl Acad Sci U S A. 2008;105:17034–17039 96. Rosenblum EB, Voyles J, Poorten TJ, Stajich JE. The deadly chytrid fungus: a story of an emerging pathogen. PloS pathogens. 2010;6(1) e10000550. http://doi.org10.1371/journal.ppat.1000550 97. Rowley JJL, Alford RA (2007) Behaviour of Australian rain forest stream frogs may affect the transmission of chytridiomycosis. Dis Aquat Organ 77:1–9 64 98. Runoff: Surface and Overland Water Runoff. (n.d.). Retrieved from https://www.usgs.gov/special-topic/water-science-school/science/runoff-surface-andoverland-water-runoff?qt-science_center_objects=0#qt-science_center_objects 99. Russel WMS, Burch RL. The principles of Humane Experimental Technique. London, UK: Methuen; 1959. 100. Schloegel LM, Hero JM, Berger L, Speare R, McDonald K, Daszak P. 2005. The decline of the sharpsnouted day frog (Taudactylus acutirostris): the first documented case of extinction by infection in a free ranging wildlife species? EcoHealth 3:35–40 101. Schloegel LM, Toledo LF, Longcore JE, Greenspan SE, Vieira CA, Lee M, Zhao S, Wangen C, Ferreira CM, Hipolito M, Davies AJ, Cuomo CA, Daszak P, James TY. Novel, panzootic and hybrid genotypes of amphibian chytridiomycosis associated with the bullfrog trade. Mol Ecol. 2012;21:5162-5177 102. Sigler L, Hambleton S, Pare´ JA. Molecular characterization of reptile pathogens currently known as members of the Chrysosporium anamorph of Nannizziopsis vriesii complex and relationship with some human-associated isolates. J. Clin. Microbiol. 2013;51:3338– 3357. doi:10.1128/JCM. 01465-13 103. Skerratt LF, Berger L, Speare R, Cashins S, McDonald KR, et al. 2007. Spread of chytridiomycosis has caused the rapid global decline and extinction of frogs. EcoHealth 4:125–34 104. Sparrow FK. A contribution to our knowledge of the aquatic phycomycetes of Great Britain. Botanical Journal of the Linnean Society. 1936;50: 417–478 105. Speare R, Core Working Group of Getting the Jump on Amphibian Disease. Nomination for listing of amphibian chytridiomycosis as a key threatening process under the Environment Protection and Biodiversity Conservation Act 1999. In: Speare R, Steering Committee of Getting the Jump on Amphibian Disease, editors. Developing management strategies to control amphibian diseases: decreasing the risks due to communicable diseases. Townsville, Australia: School of Public Health and Tropical Medicine, James Cook University; 2001. p. 163–84. Available from http://www.jcu.edu.au/school/phtm/PHTM/frogs/adms/attach7.pdf 65 106. Thogmartin WE, Sanders-Reed CA, Szymanski JA, McKann PC, Pruitt L, King RA, Runge MC, Russell RE. White-nose syndrome is likely to extirpate the endangered Indiana bat over large parts of its range. Biological Conservation. 2013;160:162-172 107. Thompson NE, Lankau EW, Rogall GM. Snake fungal disease in North America—U.S. Geological Survey updates: U.S. Geological Survey Fact Sheet. 2018;2017–3064, 4 p., https://doi.org/10.3133/fs20173064 108. U.S. EPA (United States Environmental Protection Agency). 2005. National estuary program—challenges facing our estuaries. Key management issues: Nutrient overloading. https://www.epa.gov/nep/how-national-estuary-programs-addressenvironmental-issues 109. U.S. EPA (United States Environmental Protection Agency). Nitrogen and Phosphorus in Agricultural Streams. Report on the Environment. http://www.epa.gov/roe/ 110. US Fish and Wildlife Service. Endangered and Threatened Wildlife and Plants: Proposed Endangered Status for the Southern California Distinct Vertebrate Population Segment of the Mountain Yellow-Legged Frog. In: Endangered and Threatened Wildlife and Plants: Proposed Endangered Status for the Southern California Distinct Vertebrate Population Segment of the Mountain Yellow-Legged Frog: 1999; US Fish and Wildlife Service. 9 pp. 111. US Fish and Wildlife Service. Golden Coqui (Eleutherodactylus jasperi). 5-Year Review: Summary and Evaluation. In: Golden Coqui (Eleutherodactylus jasperi). 5Year Review: Summary and Evaluation: US Fish and Wildlife Service. 2013;18 pp. http://ecos.fws.gov/docs/five_year_review/doc4276.pdf 112. Van Rooij P, Martel A, D’Herde K, Brutyn M, Croubels S, Ducatelle R, Haesebrouck F, Pasmans F. Germ tube mediated invasion of Batrachochytrium dendrobatidis in amphibian skin is host dependent. 2012;PLoS One 7:e41481 113. Van Rooij, P, Martel, A, Haesebrouck, F. et al. Amphibian chytridiomycosis: a review with focus on fungus-host interactions. Vet Res. 2015;46:137. doi:10.1186/s13567-015-0266-0 66 114. Van Rooij P, Martel A, Nerz J, Voitel S, Van Immerseel F, Haesebrouck F, Pasmans F. Detection of Batrachochytrium dendrobatidis in Mexican bolitoglossine salamanders using an optimal sampling protocol. EcoHealth. 2011;8:237–243 115. Verbrugghe E, Adriaensen C, Martel A, Vanhaecke L, Pasmans F. Growth Regulation in Amphibian Pathogenic Chytrid Fungi by the Quorum Sensing Metabolite Tryptophol. Front. Microbiol. 2019;9:3277 doi: 10.3389/fmicb.2018.03277 116. Verbrugghe E, Van Rooij P, Favoreel H, Martel A, Pasmans F. In vitro modeling of Batrachochytrium dendrobatidis infection of the amphibian skin. PloS ONE. 2019;14(11): e0225224. http://doi.org/10.1731/journal.pone.0225224 117. Vitousek P, Mooney H, Olander L, Allison S. Nitrogen and nature. Ambio 2002;31:97-101 118. Waldman B, van de Wolfshaar KE, Klena JD, Andjic V, Bishop PJ, Norman RJdeB. Chytridiomycosis in New Zealand frogs. Surveillance. 2001;28:9–11 119. Weldon C, Preez LH, Hyatt AD, Muller R, Speare R. Origin of the Amphibian Chytrid Fungus. Emerging Infectious Diseases. 2004;10(12):2100-2105 120. White-nose Syndrome A Deadly Disease (n.d.). Retrieved from http://www.batcon.org/white-nose-syndrome 121. Wickham H. tidyverse: Easily Install and Load the “Tidyverse.” 2017. 67 122. Wiethoelter AK, Beltrán-Alcrudo D, Kock R, Mor SM. Global trends in infectious diseases at the wildlife–livestock interface. Proceedings of the National Academy of Sciences of the United States of America. 2015;(112):9662–9667 123. Woodhams DC, Ardipradja K, Alford RA, Marantelli G, Reinert LK, RollinsSmith LA. Resistance to chytridiomycosis varies among amphibian species and is correlated with skin peptide defenses. Anim Conserv. 2007;10:409–417 124. Yao L, Wang D, Kang L, Wang D, Zhang Y, Hou X, Guo Y. Effects of fertilizations on soil bacteria and fungi communities in a degraded arid steppe revealed by high through-put sequencing. PeerJ. 2018;6:e4623. http://doi.org/10.7717/peerj.4623 125. Zhang H, Jia J, Chen S, Huang T, Wang Y, Zjao Z, Feng J, Hao H, Li S, Ma X. Dynamics of Bacterial and Fungal Communities during the Outbreak and Decline of an Algal Bloom in a Drinking Water Reservoir. International journal of environmental research and public health. 2018;15(2):361. http://doi.org/10.3390/ijerph15020361 68 Appendix A: Raw Data Table 7. Environmental Factors Raw Data Variable pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH Run 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 ID 5.24 5.24 5.24 5.24 5.24 5.24 5.24 5.24 5.24 5.24 5.24 5.24 5.24 5.24 5.24 6.11 6.11 6.11 6.11 6.11 6.11 6.11 6.11 6.11 6.11 6.11 6.11 6.11 6.11 6.11 7.24 7.24 7.24 7.24 7.24 7.24 7.24 Days 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 Absorbance 0.0067 0.0141 0.0521 0.0210 0.0417 0.0156 0.0427 0.0546 0.0416 0.0748 0.0254 0.0344 0.0273 0.0424 0.0639 0.0120 0.0107 0.0755 0.2253 0.0530 0.0095 0.0145 0.0804 0.7199 0.4832 0.0116 0.1165 0.6705 0.7845 0.5769 0.0138 0.1025 0.4597 1.2923 1.2273 0.0099 0.0699 69 pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH pH KeratinBroth KeratinBroth KeratinBroth KeratinBroth 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 1 1 1 1 7.24 7.24 7.24 7.24 7.24 7.24 7.24 7.24 8.10 8.10 8.10 8.10 8.10 8.10 8.10 8.10 8.10 8.10 8.10 8.10 8.10 8.10 8.10 9.23 9.23 9.23 9.23 9.23 9.23 9.23 9.23 9.23 9.23 9.23 9.23 9.23 9.23 9.23 T T T T 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 0.2803 0.7826 1.1688 0.0182 0.1968 1.7243 1.8168 1.7162 0.0162 0.0221 0.0099 0.0262 0.0265 0.0178 0.0240 0.0590 0.0337 0.0163 0.0083 0.0196 0.0520 0.0719 0.0525 0.0089 0.0249 0.0111 0.0363 0.0000 0.0163 0.0088 0.0050 0.0390 0.0645 0.0083 0.0043 0.0352 0.0000 0.0335 0.0000 0.0106 0.0949 0.1360 70 KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth KeratinBroth Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin 1 1 1 1 1 1 2 2 2 2 2 2 2 2 2 2 3 3 3 3 3 3 3 3 3 3 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 2 T TK TK TK TK TK T T T T T TK TK TK TK TK T T T T T TK TK TK TK TK T T T T T PK PK PK PK PK TK TK TK TK TK T 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 0.1479 0.0313 0.0396 0.0480 0.0087 0.0720 0.0000 0.0288 0.1081 0.1068 0.1596 0.0000 0.0000 0.0423 0.0337 0.0347 0.0000 0.0156 0.1043 0.0903 0.1769 0.0000 0.0000 0.0394 0.0290 0.0448 0.0179 0.0385 0.2992 1.6870 1.9600 0.0179 0.0330 0.1815 0.9960 1.4600 0.0428 0.0376 0.0479 0.0641 0.0525 0.0368 71 Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin Keratin PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 2 2 2 2 2 2 2 2 2 2 2 2 2 2 3 3 3 3 3 3 3 3 3 3 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 T T T T PK PK PK PK PK TK TK TK TK TK T T T T T PK PK PK PK PK TK TK TK TK TK 0 0 0 0 0 0 0 0 0 0 0 0 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 0.0572 0.4578 1.5590 1.2971 0.0368 0.0560 0.2325 0.4575 0.8586 0.0316 0.0368 0.0291 0.0585 0.0395 0.0480 0.0871 0.6643 1.5490 1.7090 0.0480 0.0594 0.3953 0.7762 0.7419 0.0597 0.0087 0.0521 0.0558 0.0548 0.0106 0.0208 0.1425 0.4015 0.4976 0.0000 0.0784 0.3273 0.5628 0.7029 0.0290 0.0765 0.3407 72 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 0 0 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.05 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.2 0.4 0.4 0.4 0.4 0.4 0.4 0.4 0.4 0.4 0.4 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0.8284 1.0512 0.0077 0.0311 0.0904 0.3719 0.2279 0.0132 0.0940 0.3483 0.5199 0.5275 0.0247 0.0579 0.2857 1.4483 0.6164 0.0087 0.0492 0.2021 0.4472 0.3831 0.0118 0.1216 0.4792 1.1607 1.2352 0.0050 0.0249 0.1776 1.2437 1.1340 0.0151 0.0692 0.2298 0.3847 0.3172 0.0003 0.1799 0.5249 0.7961 1.0033 73 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 PO4 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 0.4 0.4 0.4 0.4 0.4 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 1.0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 5 5 5 5 5 5 5 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 0.0025 0.0531 0.2930 0.6028 0.9047 0.0044 0.0476 0.1686 0.3981 0.6620 0.0044 0.1808 0.6067 0.8047 2.3970 0.0037 0.1169 1.0044 0.7505 1.5823 0.0267 0.1437 0.3668 0.3887 0.3663 0.0179 0.0393 0.1037 0.3531 0.3287 0.0151 0.0105 0.1930 0.2362 0.4034 0.0362 0.1095 0.4006 0.3211 0.2287 0.0042 0.0418 74 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 NO3 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 1 1 1 1 1 2 2 2 2 2 3 3 3 3 3 5 5 5 5 5 5 5 5 10 10 10 10 10 10 10 10 10 10 10 10 10 10 10 25 25 25 25 25 25 25 25 25 25 25 25 25 25 25 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0 3 6 9 12 0.1939 0.6997 0.5815 0.0355 0.1862 0.2494 0.2342 0.2870 0.0114 0.0570 0.0873 0.1581 0.0683 0.0059 0.0866 0.2345 0.6871 0.6909 0.0043 0.1568 0.2591 0.3000 0.1594 0.0071 0.0609 0.3150 0.1733 0.2084 0.0000 0.0506 0.2654 0.7975 0.5293 0.0177 0.1384 0.4344 0.2196 0.1694 75 Appendix B: R Code The R function utilized in the analyses: ```{r} install.packages("tidyverse") library("tidyverse") library(readxl) #KeratinBrothLM Kbroth<- read_excel("KeratinBrothData.xlsx") > View(Kbroth) > Kbroth$Days<- as.factor(Kbroth$Days) > Kbroth$ID<- as.factor(Kbroth$ID) > Kbroth$Run<- as.factor(Kbroth$Run) > str(Kbroth) Classes ‘tbl_df’, ‘tbl’ and 'data.frame': 30 obs. of 4 variables: $ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 1 1 1 1 1 ... $ ID : Factor w/ 2 levels "T","TK": 1 1 1 1 1 2 2 2 2 2 ... $ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ... $ ABS : num 0 0.0106 0.0949 0.136 0.1479 ... > Kbrothfin<- Kbroth %>% filter(Days=="12") > Kbrothfin$Days <- factor(Kbrothfin$Days) > View(Kbrothfin) > Kbrothfinaov<- lm(ABS ~ ID, data=Kbrothfin) > anova(Kbrothfinaov) 76 Analysis of Variance Table Response: ABS Df Sum Sq Mean Sq F value Pr(>F) ID 1 0.0184704 0.0184704 63.141 0.001358 ** Residuals 4 0.0011701 0.0002925 --Signif. codes: 0 ‘***’ 0.001 ‘**’ 0.01 ‘*’ 0.05 ‘.’ 0.1 ‘ ’ 1 #KeratininvitroLM Kvitro<- read_excel("KeratinData.xlsx") > View(Kvitro) > Kvitro$Run<- as.factor(Kvitro$Run) > Kvitro$Days<- as.factor(Kvitro$Days) > Kvitro$ID<- as.factor(Kvitro$ID) > str(Kvitro) Classes ‘tbl_df’, ‘tbl’ and 'data.frame': 45 obs. of 4 variables: $ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 1 1 1 1 1 ... $ ID : Factor w/ 3 levels "PK","T","TK": 2 2 2 2 2 1 1 1 1 1 ... $ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ... $ ABS : num 0.0179 0.0385 0.2992 1.687 1.96 ... > Kvitrofin<- Kvitro %>% filter(Days=="12") > Kvitrofin$Days <- factor(Kvitrofin$Days) > View(Kvitrofin) > Kvitrofinaov<- lm(ABS ~ ID, data=Kvitrofin) > anova(Kvitrofinaov) 77 Analysis of Variance Table Response: ABS Df Sum Sq Mean Sq F value Pr(>F) ID 2 3.9274 1.96370 22.608 0.001608 ** Residuals 6 0.5212 0.08686 --Signif. codes: 0 ‘***’ 0.001 ‘**’ 0.01 ‘*’ 0.05 ‘.’ 0.1 ‘ ’ 1 #pHLM pH <- read_excel("pHData.xlsx") > pH$Run<- as.factor(pH$Run) > pH$Days<- as.factor(pH$Days) > pH$ID<- as.factor(pH$ID) > str(pH) Classes ‘tbl_df’, ‘tbl’ and 'data.frame': 75 obs. of 4 variables: $ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 2 2 2 2 2 ... $ ID : Factor w/ 5 levels "5.24","6.11",..: 1 1 1 1 1 1 1 1 1 1 ... $ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ... $ ABS : num 0.0067 0.0141 0.0521 0.021 0.0417 0.0156 0.0427 0.0546 0.0416 0.0748 ... > pHfin <- pH %>% filter(Days=="12") > pHfin$Days <- factor(pHfin$Days) > View(pHfin) > pHfinaov <- lm(ABS ~ ID, data=pHfin) > anova(pHfinaov) 78 Analysis of Variance Table Response: ABS Df Sum Sq Mean Sq F value ID Pr(>F) 4 3.9771 0.99427 29.23 1.702e-05 *** Residuals 10 0.3402 0.03402 --Signif. codes: 0 ‘***’ 0.001 ‘**’ 0.01 ‘*’ 0.05 ‘.’ 0.1 ‘ ’ 1 #PO4LM PO4 <- read_excel("PO4Data.xlsx") > View(PO4) > PO4$Run<- as.factor(PO4$Run) > PO4$ID<- as.factor(PO4$ID) > PO4$Days<- as.factor(PO4$Days) > str(PO4) Classes ‘tbl_df’, ‘tbl’ and 'data.frame': 75 obs. of 4 variables: $ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 2 2 2 2 2 ... $ ID : Factor w/ 5 levels "0","0.05","0.2",..: 1 1 1 1 1 1 1 1 1 1 ... $ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ... $ ABS : num 0.0106 0.0208 0.1425 0.4015 0.4976 ... > PO4fin <- PO4 %>% filter(Days=="12") > PO4fin$Days <- factor(PO4fin$Days) > view(PO4fin) > PO4finaov <- lm(ABS ~ ID, data=PO4fin) > anova(PO4finaov) 79 Analysis of Variance Table Response: ABS Df Sum Sq Mean Sq F value Pr(>F) ID 4 1.9829 0.49572 2.0192 0.1676 Residuals 10 2.4551 0.24551 #NO3 NO3 <- read_excel("NO3data.xlsx") > View(NO3) > NO3$Run<- as.factor(NO3$Run) > NO3$ID<- as.factor(NO3$ID) > NO3$Days<- as.factor(NO3$Days) > str(NO3) Classes ‘tbl_df’, ‘tbl’ and 'data.frame': 60 obs. of 4 variables: $ Run : Factor w/ 3 levels "1","2","3": 1 1 1 1 1 2 2 2 2 2 ... $ ID : Factor w/ 4 levels "0","5","10","25": 1 1 1 1 1 1 1 1 1 1 ... $ Days: Factor w/ 5 levels "0","3","6","9",..: 1 2 3 4 5 1 2 3 4 5 ... $ ABS : num 0.0267 0.1437 0.3668 0.3887 0.3663 ... > NO3fin<- NO3 %>% filter(Days=="12") > NO3fin$Days <- factor(NO3fin$Days) > View(NO3fin) > NO3finaov<- lm(ABS ~ ID, data=NO3fin) > anova(NO3finaov) 80 Analysis of Variance Table Response: ABS Df Sum Sq Mean Sq F value Pr(>F) ID 3 0.01142 0.003808 0.0805 0.9688 Residuals 8 0.37847 0.047309 81