Bacterial Communities Within the Microbiome of Three Shark Species in South Florida Waters

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UNIVERSITY OF MIAMI

BACTERIAL COMMUNITIES WITHIN THE MICROBIOME OF THREE SHARK SPECIES IN SOUTH FLORIDA WATERS

Chelsea Leigh Black

A THESIS

Submitted to the Faculty of the University of Miami in partial fulfillment of the requirements for the degree of Master of Science

Coral Gables, Florida

May 2020

UNIVERSITY OF MIAMI

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

BACTERIAL COMMUNITIES WITHIN THE MICROBIOME OF THREE SHARK SPECIES IN SOUTH FLORIDA WATERS

Chelsea Leigh Black

Approved:

Neil Hammerschlag, Ph.D. Maria Estevanez, M.A, M.B.A. Research Associate Professor Senior Lecturer Marine Ecosystems and Society Marine Ecosystems and Society

Liza Merly, Ph.D. Guillermo Prado, Ph.D. Senior Lecturer Dean of the Graduate School Marine Biology and Ecology

BLACK, CHELSEA LEIGH (M.S., Marine Ecosystems and Society) (May 2020) Bacterial Communities Within the Microbiome of Three Shark Species in South Florida Waters

Abstract of a thesis at the University of Miami.

Thesis supervised by Professor Neil Hammerschlag No. of pages in text: (81)

Assessing shark health can provide information on species and environmental condition. One emerging measure of health status is the microbiome. The assemblage of bacterial communities within the microbiome of a host can provide insight into the environmental conditions and outside stressors a species is encountering. Anthropogenic pressures such as fishing and bycatch are well-known threats to global shark populations, however, habitat degradation and other environmental changes may actually contribute to their susceptibility to disease. Bacteria are known to have explicit roles in vertebrate tissues, so examining the bacterial communities across the microbiome may prove to be a useful tool for assessing the role of commensal bacteria in shark health and determining health status. This study characterized the bacterial communities within water samples and across several tissue sites of blacktip (Carcharhinus limbatus), bull (Carcharhinus leucas) and tiger (Galeocerdo cuvier) sharks off the coast of Miami, Florida through microbiological and molecular sequencing methods. Blacktip sharks displayed the highest bacterial diversity in terms of their microbiome. Planococcaceae bacteria possessing antibiotic compounds were unique to the skin of tiger sharks in this study, further highlighting the likelihood for species-specific bacteria. Further, known pathogens such as Vibrio and Staphylococcus were identified in the wounds of sharks in this study. While

water samples did possess unique bacteria not identified on the sharks, it also included pathogenic bacteria that was identified in the wounds of blacktip and bull sharks. The results indicate that while some bacteria may be normal residents of the microbiome, the loss of protective dermal denticles due to a wound may provide opportunistic bacteria a chance to colonize in areas where they would not normally be found. It will be important for future work to continue investigating the bacterial communities found within the microbiome of sharks to determine which bacteria are normal colonizers and which species may be pathogenic invaders. Determining the presence of microbes and their functionality will provide insight into the role of the microbiome to host health, as well as the environmental conditions that these sharks are encountering.

Acknowledgements

I would like to thank my advisor, Dr. Neil Hammerschlag, for opening his door to me and trusting me as a valued member of the Shark Research and Conservation Program.

The opportunities you have given me not only solidified my passion for marine biology, but instilled confidence to pursue this career. To my co-advisor, Dr. Liza Merly, thank you for your encouragement and guidance over the years as I learned microbiology. You made me believe in myself when I needed it most and I will always appreciate your patience and support. To Maria Estevanez, I appreciate your immeasurable kindness and encouragement through my entire academic career. Your smile was an illuminating force guiding me to the finish line. This thesis is a result of the lessons and encouragement provided to me by my committee and I will always value my time learning from them.

I must thank my fellow SRC members, past and present, for the countless laughs and memories along the way. To Stephen Cain, thank you for being my career-coach, mentor, barista, but most importantly my good friend. You have imparted wisdom on me that I can never repay. To Shannon Moorhead, I am forever appreciative of everything you have taught me and for being my confidant. To Laura McDonnell, thank you for being a shoulder to lean on, the best travel buddy, and an inspiration. To Mitchell Rider for fueling me with donuts and assisting with my final analysis. To Patricia Albano for lifting my spirit with laughter. I must also thank Andy Fragola for trusting me to build on her research project and her mentorship in the lab. To Allison Banas for being an amazing undergraduate assistant and Julia Saltzman for making quite possibly every agar plate in this study. To Elana Rusnak for her guidance and company in the lab. It is through all of these people, and more, that my research was possible.

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I would also like to thank the friends I have made here at the University of Miami for their support through the ups and downs. To Megan Huse, Erin Cain, and Kathryn Grazioso- your friendships kept me sane during this journey. Thank you for providing me with encouragement along the way and celebrating each milestone with me.

Last, but certainly not least, I would like to thank my family. Without my parents, a childhood dream would have remained just that. Thank you for instilling a passion for science within me and fostering my love for the ocean from a young age. It is because of your encouragement, love, and support that my hard work has come to fruition. There are no words to describe what you have given me, and I only hope to continue making you proud. This is dedicated to you.

Table of Contents

List of Figures...... vi

List of Tables...... vii

Chapters

Introduction...... 1

1. Marine Microbes...... 4

2. Microbes as Indicators for Shark Health...... 13

3. Methods...... 26

4. Results...... 34

5. Discussion...... 50

Conclusion...... 60

Literature Cited...... 62

Appendix A – Marine Agar Preparation...... 76

Appendix B – Slide Microscopy...... 77

Appendix C – DNA Sequencing...... 78

Appendix D – Statistics for Shannon’s Index Calculations...... 80

List of Figures

Figure 1. Fishing method for safe capture of sharks...... 26

Figure 2. Process of microbial sampling...... 27

Figure 3. Location of all sampled sharks within the study...... 28

Figure 4. Process of culturing each cell scrape...... 30

Figure 5. Distribution of Gram-negative and Gram-positive across tissue sites...... 35

Figure 6. Distribution of Gram-staining across all shark species and water samples...... 36

Figure 7. Distribution of bacteria morphology types across all shark species...... 38

Figure 8. Distribution of bacteria morphology shapes across all shark species...... 39

Figure 9. Summary of taxonomic groups identified through DNA sequencing...... 40

Figure 10. Total composition of bacteria phyla for sequenced samples...... 41

Figure 11. Distribution of phyla across species and tissue site...... 42

Figure 12. Venn diagram showing families of bacteria shared between sharks...... 44

Figure 13. Venn diagram showing bacteria families shared with water samples and sharks...... 45

Figure 14. Depiction of bacteria families found across the tissue sites of each species...... 46

Figure 15. Summary of bacteria families found across all water samples...... 47

List of Tables

Table 1. Distribution of bacterium genera in the phylum Actinobacteria...... 48

Table 2. Distribution of bacterium genera in the phylum Firmicutes...... 49

Table 3. Distribution of bacterium genera in the phylum Proteobacteria...... 49

Table B-1. Total number of microscope slides per species and tissue site...... 77

Table C-1. Samples submitted for DNA sequencing...... 78

Table D-1. Diversity of bacterial genera in blacktip sharks...... 80

Table D-2. Diversity of bacterial genera in bull sharks...... 80

Table D-3. Diversity of bacterial genera in tiger sharks...... 81

Table D-4. Diversity of bacterial genera in water samples...... 81

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Introduction

Health parameters can provide valuable information about wild shark populations and the environments they inhabit. One emerging measure of health status in animals is the characterization of their microbiomes (Hollister et al. 2014). The microbiome is defined as the community of microorganisms that associates with a host organism and has possibly co-evolved with it (Rosenberg et al. 2007). Members of this community are typically prokaryotes (bacteria and archaea), eukaryotes (fungi, protists, algae), and viruses

(Pogoreutz et al. 2019). Bacteria are known to have explicit benefits to their host, therefore, examining bacterial communities across a microbiome may prove to be a useful tool in assessing shark health. However, to measure these parameters, there must first be an effective means to characterize this microbiome.

In captive settings such as aquariums or research laboratories, animals can be sampled several times in controllable conditions to assess variations in normal parameters; however, even with access to study the microbiome of sharks, little research has been conducted. It is important to study shark populations to define a microbiome for each species before this can be used as a tool for health assessment in the future.

The objectives of the current study were to assess the bacteria associated with shark tissues and to conduct an initial characterization of shark microbiomes. To address the objectives of this study, three shark species were sampled in Biscayne Bay, Florida to identify bacterial communities within their microbiomes. The species of interest were the blacktip (Carcharhinus limbatus), bull (Carcharhinus leucas) and tiger (Galeocerdo

cuvier) sharks. These species were selected due to their common habitat use in Biscayne

Bay and unique life history traits.

1 2

Blacktip sharks display a migratory behavior, with gravid females along the US

Atlantic and Florida Gulf coasts migrating to nursery areas to give birth (Keeney et al.

2005). This migration exposes blacktip sharks to a wide range of environmental conditions throughout the east coast and Florida. The bull shark also exhibits a wide-ranging distribution in Florida and is known to travel long distances (Compagno 1984). This species is also unique in that it will aggregate in estuaries and freshwater, exposing them to numerous anthropogenic stressors occurring in nearshore areas (Karl et al. 2011). Bull sharks prove to be an interesting study subject as they likely experience the widest range of environmental conditions. Lastly, the tiger shark is a wide-ranging species that displays a variety of horizontal movement patterns, making use of both coastal and pelagic waters

(Vaudo et al. 2014). All three of the study species were targeted due to their habitat use patterns, in addition to displaying physical traits relevant to microbial research. Both blacktip and bull sharks encountered in Biscayne Bay often presented dark necrotic patches of skin behind the first dorsal fin and before the caudal fin. These compromised areas, referred to as “wounds”, have not been thoroughly investigated in previous studies. This study provided an opportunity to compare the bacterial communities present on uncompromised and compromised areas of the same individual, to assess how a shark’s microbiome may respond to physical insult and loss of integrity. Secondly, tiger sharks have a visibly thicker mucus layer covering their skin in comparison to bull and blacktip sharks. While this highly productive mucoid layer has not been studied, it triggered questions about how their microbiome may differ from the other species in the study. The combination of a thin mucus layer and the unique structure of dermal denticles may result in a selective skin microbiome for all three species (Doane et al. 2017).

To characterize bacterial communities across the microbiomes of these sharks, cell swabs were taken from the mouth, gills, dorsal skin (hereafter referred to only as “skin”), and compromised areas if present. Additionally, a water sample was collected at the location where each shark was sampled to identify microbes present in the surrounding environment. Sampling multiple tissue sites across the same individual provided a snapshot of the tissue-specific microbiome on the shark. From there, comparisons between tissue site, species, and study location could be made. The current study had four specific aims:

1) determine what bacterial communities were present across various tissue sites of the same species, 2) determine how species-specific these bacterial communities were for sharks occupying similar habitats, 3) determine if bacteria found within shark tissues were significantly different than those found in their immediate environment, and 4) determine what bacteria were present on compromised areas versus uncompromised areas of the same individual.

Chapter 1: Marine Microbes

Background

Chondrichthyan fishes have survived mass extinction events over the past 400 million years (Lund & Grogan 2004). Sharks represent 41.7% of all extant chondrichthyan species (Lund & Grogan 2004). Most chondrichthyan species are characterized by low growth rates, late sexual maturity, and low fecundity when compared to bony fish (Ferretti et al. 2010). Due to these life history traits, sharks demonstrate twice the fishing extinction risk of bony fish with a poor ability to recover after population depletion (Myers & Worm

2005). Sharks have been evolving for millions of years, but today over 30% of shark species are classified as endangered, vulnerable, or near threatened by the IUCN Red List

(2019). Increased research is needed, not only inform conservation efforts for these important species, but to better understand the current state of our oceans.

Anthropogenic pressures, such as fishing and bycatch, are well-known threats to global shark populations, however, an often-overlooked threat includes the susceptibility of sharks to bacterial infection or disease. Little is known about the physiological status of wild sharks, as the majority of research on health parameters has taken place in captive settings, such as aquariums and research laboratories. By studying wild populations, not only can we assess their health status, but also relate it to the quality of the environments in which they reside.

Role of Microbiomes in Host Health

The microbiome is a community of various microorganisms that are associated with their host. The term “community” can be defined as multispecies assemblages, in which organisms live together and interact with each other in a contiguous environment

(Boon et al. 2013). Members of this community are typically prokaryotes (bacteria and archaea), eukaryotes (fungi, protists, algae) and viruses (Pogoreutz et al. 2019). These communities of microorganisms are critical to the health of the host by being involved in host functions such as nutrient supplementation, successful development, and disease susceptibility (Doane et al. 2017). Host-microbiome dynamics are described in two categories: firstly, as symbiosis, in which the organisms are involved in the normal physiological functions and metabolic interactions, and secondly as dysbiosis, in which the relationship or interactions are heavily altered, possibly related to a major stress or infection event (Apprill 2017).

Investigating the microbiome of marine species is a relatively novel field, but previous research has paved the way for methodology and demonstrated the ability to use the microbiome for health assessment. Host-microbiome dynamics have recently received attention in human medicine as conditions, such as obesity and irritable bowel syndrome, have been linked to deviations from the normal gut microbiome in the host (Greenblum et al. 2012). However, these same concepts are also applicable to other organisms, including those found in the ocean. DNA staining methods have revealed that the number of bacteria in the open ocean exceeds 1029 cells, with average cell concentrations of 106 per milliliter of seawater (Whitman et al. 1998). Bacterial abundance is now known to vary at the milliliter scale, and this variability rises in response to increases in the concentration of particulate organic matter in seawater (Long & Azam 2001). Therefore, research investigating bacterial communities within the microbiome of aquatic species has begun to shed light on some of these host-microbiome dynamics. In coral species, different coral- associated bacteria are hypothesized to play varying roles in coral health, suggesting that

6 coral reef microbial communities may serve as indicators of environmental stress and individual health (McDevitt-Irwin et al. 2017). In aquaculture settings, studying fish microbiomes is important for assessing stock health and applying necessary treatments to disease (Olafsen 2001). In sharks, however, the role that microbiomes play in host health is poorly understood, and research on the topic is scarce. Mounting evidence from other marine groups indicates that a shift in the normal bacterial communities can leave their host vulnerable to disease or infection and that these shifts can be related to environmental conditions such as water quality, prey availability, and temperature change.

Marine Microbes

Marine animals share the sea with an assortment of microorganisms including protists, bacteria, archaea, fungi and viruses (Apprill 2017). The full wealth of microbial diversity in the ocean has yet to be revealed as a large fraction of marine bacteria have not been cultured, and the complexity of microenvironments existing in the water column is difficult to account for (Long & Azam 2001). A major challenge for exploring host- microbiome interactions is at the genetic level. Amplifying or shotgun sequencing microbial DNA with the presence of abundant host cells often requires an optimization or high sequencing output, but taxonomic databases generally contain very few microbial sequences from marine animals due to lack of research on the subject (Apprill 2017).

Another challenging issue is gaining information from unknown genes and gene families, which potentially make up over half of the environmental microbial genomes (Rocha et al.

2014).

Microbial Diversity in Marine Species

Microorganisms that associate with marine animals are part of the animal’s microbiome. Some of these microorganisms are thought to originate from the surrounding supply of seawater-associated cells (Nussbaumer et al. 2006), while others may have strict inheritance patterns that are passed on through generations (Sharp et al. 2007). The overall microbiome of an individual is likely the result of both horizontal transmission from the environment and vertical transmission through inheritance. Vertical transmission of associated microorganisms is a feature that has been observed across many metazoans, such as bivalves, ascidians, bryozoans, oligochaetes, insects, and fish (Sipkema et al.

2015). This transmission is seen as an indicator for ancestral symbioses between microbes and their host. Horizontal transmission is the recruitment of microorganisms from the environment (Sipkema et al. 2015). Marine sponges, for example, are particularly fit to acquire their associated microorganisms horizontally, due to their high-water pumping rates and filtration system that encounters numerous cells per day (Sipkema et al. 2015).

In the ocean’s upper photic zone, marine animals are exposed to variations in temperature and light, and host-symbiont interactions (especially in ectothermal animals) could vary across seasons (Apprill 2017). Normal variations in species-specific patterns can also alter host-microbiome relationships, including changes in diet, stress, or the production of hormones (Apprill 2017). Certain life events occurring over longer time periods, such as animal development, aging, and reproduction, also have the potential to trigger alterations in host-microbiome relationships (Apprill 2017). The microbiome of marine animals may harbor bacteria that are either directly antagonistic or capable of outcompeting potential pathogens (Olafsen 2001). In some species, the role of the

8 microbiome in organismal health and responsiveness to environmental change has been studied and can inform how this process may work in shark species.

Microbiome in Corals

Corals have been a recent topic for microbial research due to the increasing concern for their global decline. Reports of diverse protists, bacteria, archaea, and viruses in association with corals can provide insight into how these cells are fulfilling functional processes within their coral host (Thompson et al. 2015). One of the most visible signs of host-microbiome dysbiosis is within scleractinian corals, whose relationship with unicellular microalgae breaks down under certain environmental conditions such as long- term increases in seawater temperature, causing the coral to become bleached (Brown

1997).

The surface of living coral is covered by a mucoid material that provides a matrix for bacterial colonization that is often higher in bacterial abundance than the surrounding water (Reed et al. 1999). Coral-associated bacteria can be transferred either vertically or horizontally. Sharp et al. (2012) supported the first evidence of vertical transmission (from parent to offspring) of bacteria in corals. Their results showed that at least two groups of bacterial taxa were consistently associated with juvenile scleractinian corals that were important for host health and survival.

Previous research has also concluded that like other systems studied, the coral microbiome is likely to be variable until the colony reaches adulthood (Williams et al.

2015). Corals also possess a wide range of symbiotic eukaryotes within their community structure, and variations in their microbiome have also been linked to the by-products of photosynthesis from these eukaryotes (Hernandez-Agreda et al. 2017). In addition, coral

9 microbiomes have been shown to fluctuate seasonally, influenced by abiotic factors

(Hernandez-Agreda et al. 2017).

Different coral-associated bacteria are hypothesized to play varying roles in coral health, nutrition, and development, suggesting that coral reef microbial communities may serve as indicators of both environmental stress and individual coral health (McDevitt-

Irwin et al. 2017). Many coral-associated bacteria defend the coral by exuding antimicrobial compounds to prevent invasions by pathogens or other bacteria. Rypien et al. (2010) discovered that nearly 70% of cultivable isolates from the coral Montastrea annularis inhibited the growth of other bacteria, while 11.6% inhibited the growth of the known coral pathogen Vibrio shiloi. Morrow et al. (2012) discovered that corals closer to human disturbance have been shown to harbor higher bacterial diversity than those farther from the disturbance.

Microbiome in Sponges

Sponges are common members of the ocean’s benthic habitat and are one of the oldest lineages of animals with a simple body plan that commonly associates with bacteria, archaea, algal protists, fungi, and viruses (Apprill 2017). Hentschel et al. (2002) was the first to describe sponge-derived bacteria that cluster together, regardless of their origin

(host sponge and/or sampling location). These sponge-specific bacteria described a phylogenetically complex community that was repeatedly detected in sponges around the world but was different from microbial seawater communities (Hentschel et al. 2002).

Schmitt et al. (2012) proposes that, while different sponges contain bacterial communities consisting of mainly different bacterial species, they are still more closely related to each other than to surrounding seawater bacteria. Like corals, the microbiomes of some sponge

10 species appear to change in community structure in response to changing environmental conditions such as temperature, ocean acidification, and synergistic impacts (Apprill

2017).

Microbiome in Fishes

Previous research on the microbiome of teleosts has been slow, with a focus primarily on aquaculture species (Llewellyn et al. 2014). Modern aquaculture provides an effective means for studying microbiota under controllable conditions. During the intensive hatching of eggs and rearing of marine larvae, various forms of interactions between bacteria and biological surfaces occur. This may result in the formation of an indigenous microflora or be the first step of infection (Olafsen 2001). The diverse flora that eventually develops on the egg’s surface reflects the bacterial composition of the water, however, species-specific adhesion might also affect the composition of the epiflora

(Olafsen 2001). Romero and Navarrete (2006) pioneered the identification of dominant bacterial populations associated with the early life stages of Coho salmon (Oncorhynchus kisutch) and focused on three developmental stages (eggs, first-feeding, juvenile). They documented the environmental bacterial communities in both the surrounding water and the feed to determine the origin of dominant bacteria within the intestinal tract. The authors concluded a stable microbiota developed after the first feeding stages, with major components derived from surrounding water and egg epibiota.

The microbiome of the skin and gut regions of teleost fishes has been investigated in several species. Bacteria present on the skin of fast-swimming fish change the properties of the epithelial layer to reduce drag and assist in locomotion. The resident bacteria might have the ability to attach strongly to the skin, have high surface hydrophobicity, the ability

11 to release drag-reducing polymers, or mechanisms for overcoming antimicrobial agents

(Cahill 1990). The taxonomic composition of the skin microbiome on fishes is primarily driven by the host (Larsen et al. 2013). Extreme environmental changes induce a fluctuation in this composition, as previously observed in the Atlantic salmon (Salmo salar) microbiome during migrations from river to ocean habitats (Doane et al. 2017). Host factors selecting for symbiotic skin microorganisms on fishes include the skin mucus and the anti-microbial compounds within that mucus (Ingram 1980).

The fish gut microbiota contributes to digestion and can affect nutrition, growth, reproduction and vulnerability of the host fish to disease (Ghanbari et al. 2015). The digestive tract of fish encounters both food and water that contain microorganisms from the surrounding environment. The primary bacterial groups represented in the fish microbiota are Proteobacteria, Fusobacteria, Firmicutes, Bacteroidetes, Actinobacteria,

Cloostridia, Bacilli and Verrucomicrobia (Ghanbari et al. 2015). Symbiotic bacteria can help their host acquire nutrients from their diet, so gut bacteria often promote nutritional provisioning and influence the digestion of complex carbohydrates (Sullam et al. 2012).

Sullam et al. (2012) performed a meta-analysis of gene sequence data from fourteen published and two unpublished data sets, concluding that host trophic level, habitat, and possibly host phylogeny played a role in shaping fish gut microbial communities.

Microbiome in Rays

To date, there is very little research on the microbiome of ray species. Johny et al.

(2018) explored the gut microbiota in the dark blindray (Benthobatis moresbyi) from the

Arabian Sea and compared the results to existing datasets. The dominant phyla included

Actinobacteria, Proteobacteria and Acidobacteria.

The microbial communities on the dorsal and ventral surface of cownose rays

(Rhinoptera bonasus) in touch-tank aquariums were investigated by Kearns et al. (2017).

Their analysis revealed a unique microbial community associated with the skin of the rays that had a lower diversity than the surrounding habitat. Additionally, no human pathogens were detected, suggesting that human interactions did not significantly promote the growth of human-associated bacteria on ray skin.

McWhirt (2019) evaluated the microbiome of wild and captive spotted eagle rays

(Aetobatus narinari) by comparing the bacteria found on the dorsal, gills, and cloaca. The author concluded that the gill and skin microbiomes differed between captive and wild populations and were similar but distinct from the water column communities. However, the cloaca microbiomes were more divergent from that of the water samples.

Chapter 2: Microbes as Indicators for Shark Health

Shark Health

While host-microbiome dynamics have been investigated in marine species such as corals, sponges and teleosts, this area of research has not been fully explored in other marine animals. Similar to corals and teleosts, sharks also possess a mucoid matrix that provides a surface for bacterial colonization. It is likely that the same host-microbiome theories applied to their aquatic neighbors can also be applied to sharks and investigating bacterial communities within this microbiome may help us understand their role in host health.

Sharks have been popular attractions in public aquariums and zoos since the 1860s when they were first exhibited, so knowledge about their biology and husbandry has increased consistently for years. They have been used as model organisms to study physiology, however, knowledge about diseases is limited and only a few disorders have been reported in peer-reviewed literature (Florio et al. 2016). Reports of viral, bacterial, and fungal diseases are relatively scarce and often involve case reports of only a single animal; it is likely that this is due to a lack of diagnostic sampling rather than a true lack of outbreaks (Terrell 2014). Because stress-related disruptions to the microbiome are believed to be a precursor to disease in other vertebrates, it is important to begin investigating the role of symbiotic microorganisms in the health of sharks.

To date there is limited research on the microbiome composition of elasmobranchs, and even fewer studies on the microbiome of sharks. However, it is important to investigate how microbes influence shark health as both an ecological indicator of environmental quality and a factor affecting the abundance and trends of a species’

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population. The composition and abundance of the microbiome varies through both space and time in response to ecological interactions with the host and environment (Van Opstal

& Bordenstein 2015). Microbiomes have the potential to influence health, physiology, behavior and ecology of marine species. These findings could alter current understandings of how these species might adapt to anthropogenic and natural changes in their environment. It is presumed that symbiotic microbial associations in various shark tissues may contribute to protective mechanisms against pathogens and disease in these animals.

Bacterial Disease in Elasmobranchs

The most common elasmobranch species studied for diseases include: cownose rays (Rhinoptera bonasus), southern rays (Dasyatis americana), dusky smooth-hounds

(Mustelus canis), bonnethead sharks (Sphyrna tiburo), and bamboo sharks (Chiloscyllium plagiosum) (Garner 2013). In research by Garner et al. (2013), bacterial infections (sepsis, dermatitis, branchitis and enteritis) made up 15% of all cases. 9% of all cases were parasitic infections (nematodiasis, ciliate, trematodiasis, coccidiosis, myxozoanosis, amoebiasis and more), 20% of which were identified as trematodiasis.

Outbreaks of bacterial disease are relatively uncommon in elasmobranchs, but of those reported, the most common is that of Vibrio spp, which have emerged as the most significant group of marine pathogens across a wide range of metazoans (Bertone et al.

1996). V. alginolyticus, V. anguillarum, V. cholerae, V. damsela, V. ordalii, V. salmonicida, V. vulnificus, and V. carchariae have all been described as fish pathogens, some of which can also cause human infections. The most common Vibrio spp. isolated from sharks is Vibrio carchariae, which has been repeatedly implicated as the cause of meningitis in sand tiger (Carcharias taurus), lemon (Negaprion brevirostris) and sandbar

15 sharks (Carcharhinus plumbeus), as well as the spiny dogfish (Squalus acanthias) (Terrell

2014). The trematode Dermophthirius sp. was responsible for the transmission of V. carchariae between sharks and was associated with chronic skin ulcers and inflammation of the outer covering of the brain (Terrell 2014). V. carchariae was found to be the predominant strain in an area of necrotic skin on a sandbar shark kept in the Genoa

Aquarium, Italy. Bertone et al. (1996) concluded that while several strains of V. carchariae were identified on the shark’s ulcer, this does not mean that it was the causal agent of the ulcer. It is possible that the bacteria could have been an opportunistic or secondary invader, as the same bacteria has been identified in healthy sharks. This is yet another demonstration of the importance of understanding the normal microflora present in each shark species or population to determine the role of these various microbes in both shark health and ecosystem functioning.

Aeromonas salmonicida has been documented as a causal agent of hemorrhagic septicemia in the blacktip reef shark (Carcharhinus melanopterus), Serratia marcescens has been associated with ampullary system infection and septicemia in a bonnethead shark, and Mycobacterium avium has been isolated from granulomas in an epaulette shark

(Hemiscyllium ocellatum) (Florio et al. 2016). Skin lesions characterized by the abundance of white necrotic tissue was identified as an infection caused by Tenacibaculum maritimum on the skin of a captive sand tiger shark was identified by Florio et al. (2016). This pathogen has the ability to attach itself strongly to external fish surfaces and adhere to skin, including the mucus layer, due to its production of extracellular polymeric substances. The infection could have been triggered by mechanical injuries that occurred during mating or as a result of aggressive behavior between conspecific sharks (Florio et al. 2016).

A common bacterium, Carnobacterium spp., has been found inhabiting the intestine, urogenital tracts, and gills of both fresh and saltwater teleosts, both in aquaculture settings and in wild populations (Schaffer et al. 2013). While some species of

Carnobacterium are known to produce bacteriocins (proteins that kill or inhibit the growth of other bacteria), other Carnobacterium species, particularly C. maltaromaticum, have been associated with morbidity and mortality in a variety of fish species (Schaffer et al.

2013). Carnobacterium has not been reported as normal flora in sharks, but a series of juvenile salmon shark (Lamna ditropis) strandings along the California coast prompted researchers to examine the bacterial composition of 19 individuals. Schaffer and colleagues

(2013) discovered meningitis or meningoencephalitis in 18 of 19 brains, with bacterial cultures of 13 sharks growing the genus Carnobacterium- the first report of this bacteria in any shark species. The severity of the infection identified in this study suggested the inflammation of the brain altered the sharks’ behavior and ultimately resulted in their stranding. All stranded salmon sharks were juveniles, so the epidemiology of this disease would be better understood by investigating the source of exposure for juvenile salmon sharks and any predisposing factors such as age or environment. The researchers concluded the infections may be specific to juveniles because of unknown aspects of their life history, and it is possible that adults are exposed and infected but not reported because they inhabit the northern Pacific Ocean where reporting and retrieval of stranded individuals is less likely. In addition, adult salmon sharks are pelagic, and any ill or dead individuals are more likely to be preyed upon or scavenged in deep water as opposed to stranded on beaches

(Schaffer et al. 2013).

Antibiotic resistance was investigated by Blackburn et al. (2010) in seven shark species through cloacal swabs. Through gram staining and resistance tests, it was discovered that resistance to at least one antibiotic-resistant bacteria was present within all species tested. This pattern was found across both species and regions. It has been determined that different types of epidermal mucus have antibacterial effects against pathogenic bacteria. In a study by Yemiske et al. (2018), 164 bacteria were isolated from the epidermal mucus of three elasmobranch species in Turkey. Approximately 2% of isolated strains showed antibacterial activity against Escherichia coli and Enterococcus faecalis. In addition, Gram-negative strains such as Klebsiella pneumonia and

Pseudomonas aeruginosa were sensitive to an active compound isolated from fish mucus, possibly due to the antimicrobial peptides found in mucus.

Viral Diversity in Elasmobranchs

Two different viral diseases have been reported in captive elasmobranchs (Terrell

2014). The first disease, observed in the dusky smooth-hound, was characterized by areas of white to gray skin discoloration, which manifested following a stressful event and spontaneously resolved itself after a variable period of time (Terrell 2014). The second disease, observed in both the dusky smooth-hound and leopard sharks (Triakis semifasciata), was a viral infection of the red blood cells known as viral erythrocytic necrosis. The disease destroyed red blood cells and typically affected young animals with no immunity to the virus (Terrell 2014).

Epitheliocystis is an infectious disease of the gills and skin in many marine and freshwater species but was only recently identified in a leopard shark. Polkinghorne et al.

(2010) described the first molecular identification of a bacterial agent associated with

18 epitheliocystis in a non-teleost species. However, the sequences detected in the leopard shark were distinct from those identified in epitheliocystis in teleost species and was unresponsive to treatment which have been effective in other fish species.

Parasitic Diversity in Elasmobranchs

The body of an elasmobranch displays a diversity of sites that can be, and often are, occupied by other animals such as parasites. Parasites can steal nutrients, destroy or create obstructions within host tissues, negatively alter behavior, or predispose hosts to secondary infections of opportunist pathogens (Benz & Bullard 2004). Despite the relatively extensive literature on elasmobranch parasites, the diversity is underestimated as hundreds of species of elasmobranchs have yet to be examined for parasites (Carrier et al. 2004).

Certain organs and organ systems such as the skin, digestive system, and gills tend to host particularly diverse faunas of parasites (Carrier et al. 2004). Parasites in captive elasmobranchs have been routinely observed as they are a common health problem for species in captive settings. Parasites can be divided into three ecological groups according to their style of infection: ectoparasites, endoparasites, or mesoparasites (Benz & Bullard

2004). Ectoparasites inhabit any exterior site or orifice of the host and include leeches, arthropods and mollusks (Carrier et al. 2004). Endoparasites inhabit sites associated with interior cavities and organs, and the major groups include acanthocephalans, nematodes and platyhelminths (Carrier et al. 2004). While parasites as a group can infect any host tissue, almost all parasites tend to infect specific tissues or body regions. Parasites that attach to the skin of elasmobranchs are often specific to the skin of a particular region of the body, and this site specificity is commonly seen with copepods and monogeans (Carrier et al. 2004). Organisms parasitizing the skin of elasmobranchs possess appendages or

19 attachment structures that are useful for attaching below, around, or on top of the placoid scales of the elasmobranch (Carrier et al. 2004).

The invertebrate metazoans parasitizing elasmobranchs belong to six phyla:

Mollusca, Acanthocephala, Annelida, Nematoda, Arthropoda and Platyhelminthes (Carrier et al. 2004). The greatest diversity of parasites consists of arthropod and platyhelminth species (Carrier et al. 2004).

Fungal Diversity in Elasmobranchs

A wound or abrasion in the skin is a possible entry site for opportunistic pathogens, such as fungi. Fungal diseases are typically characterized in an individual animal rather than an explosive outbreak affecting multiple animals (Terrell 2014). A well characterized fungal disease is an infection by the fungus Funsarium solani (Terrell 2014). The disease has been described in bonnethead sharks and scalloped hammerhead sharks (Sphyrna lewini). The affected sharks develop white pustules along their lateral line and cephalofoil, and in some cases skin erosions or ulcers have been recorded with the potential for hemorrhages (Terrell 2014). Because fungal outbreak typically follows an environmental change, it has been noted that Funsarium spp. has developed after shallow warm-water sharks are kept in cold deep-water aquarium systems.

Microbiome in Elasmobranchs

Many marine organisms appear to have an intrinsic, natural ability to prevent biofouling (the growth of unwanted biofilms on surfaces) through non-toxic mechanisms

(Sullivan & Regan 2011). The skin is the first line of defense for sharks and acts as a physical barrier to the surrounding environment. Like teleosts, sharks have a biologically active mucus layer, although more reduced in comparison (Meyer & Seegers 2012).

Wounds seen on sharks often heal quickly, and the surrounding skin retains normal functioning (Doane et al. 2017). Sharks have few documented infections and are commonly observed in the wild with open wounds, yet rarely do these present as infected (Doane et al. 2017). Skin may provide a habitat for host-associated bacteria that confer additional protections to the host against infection by pathogens. These skin-associated microbes must be able to thrive, however, in the unique environment of shark skin.

A shark’s skin is comprised of dermal denticles, which are rough in texture to aid in protection against injuries, as well as increase hydrodynamics for the species, and also act as a physical barrier to infection. Dermal denticles are a form of placoid scales and contain riblets that allow fast-swimming shark species an additional 8-10% swimming efficiency in comparison to a smooth surface (Sullivan & Regan 2011). Because of this, dermal denticles are of interest in antifouling material development, due to the dual functionality in both drag reduction and biofouling reduction. In the fields of medicine and environmental pollution, efficient methods to prevent bacteria from growing on surfaces are constantly being sought. One example is the Sharklet AF™, a novel antifouling material that mimics dermal denticles by consisting of a topographically textured surface with repeating patterns of odd-numbered riblets. It has been reported that the Sharklet AF™ is successful in inhibiting biofilm formation of the bacteria Staphylococcus aureus, as well as the migration of E. Coli (Sakamoto 2014). Even when the grooves of this pattern were shallow, it was shown that biofilm formation and swarming motility of bacteria were more effectively inhibited by this shark skin micropattern than by regular parallel ridges. The results of studying the Sharklet AF™ suggest that bacterial infections could be greatly

21 reduced if surfaces on medical instruments or within hospitals could be covered with this material.

Doane et al. (2017) characterized microbial communities from the epithelial surface of the thresher shark (Alopias vulpinus). The results suggested that thresher sharks shared common microbial genera and functions across individuals, which varied from those found in the water column. By comparing the microbial taxonomic diversity, the authors suggest that the shark, water, and algal microbial communities share some common community characteristics, while also exhibiting differences. At the gene level, the shark microbiome displayed greater diversity, richness, and evenness; this suggests varying mechanisms involved in regulating the skin microbial communities. The higher richness of the shark microbiome may reflect the shark skin encountering more microbial taxa due to their higher spatial habitat use. Pseudoalteromonas was overrepresented in the shark microbiome and is responsible for synthesizing anti-microbial compounds that hinder settlement by eukaryotic marine fouling organisms. Erythrobacter was also common and is responsible for oxygenic photosynthesis, which could supplement a variety of compounds to other members of the microbiome. Idiomarina spp. was also present and is responsible for detoxification of heavy metals that may leech onto the skin surface. Marinobacter was the fourth genus to distinguish thresher shark skin from the water column and is known to degrade hydrocarbon and produce lipopolysaccharides, which mediate host inflammatory response.

In addition to examining the skin, other areas of the shark, such as the mouth and gills, can serve as potential entry points for microbes due to the constant passage of water through the mouth and gills during ram ventilation. While sharks have been responsible for

22 fewer than 400 human fatalities around the world since first being recorded in the early

1900s (International Shark Attack File), there is a growing need to understand consequences from human-shark interactions, particularly in the medical community.

When shark bites occur, treatment for infection is usually carried out with non-specific antibiotics since the identification of bacterial pathogens in the oral cavity of sharks is rare

(Interaminense et al. 2010). In Recife, Brazil, between 1992 to 2009 sharks bit 53 people,

20 of whom died as a result of bleeding (Interaminense et al. 2010). The species identified in these cases were primarily bull (Carcharhinus leucas) and tiger sharks (Galeocerdo cuvier). Due to amputations, survivors were treated with antibiotic therapy to avoid wound infections. Interaminense et al. (2010) screened bacterial isolates from the oral cavity of captured bull and tiger sharks for antibiotic susceptibility to obtain data to support antibiotic therapies for shark bite victims. The microbial isolates included Gram-positive and Gram-negative bacteria previously associated with various illnesses in humans related to wound or soft-tissue infections: pyomyositis, osteomyelitis and bacteremia. Some bacterial species identified in the oral cavity have also been identified in seawater. The authors of this study compared the results of the bacterial isolates found in the oral cavity of the sharks to the medical records of the shark bite victims in Brazil. The wound culture of one patient showed the bacteria Pseudomonas aeruginosa and E. coli which were also identified in the oral cavity of tiger sharks. Common treatment for shark victims includes the antibiotics clindamycin for prevention of Gram-positive infections and ciprofloxacin for the remaining strains. However, in the study described above, 33% of Gram-positive strains identified in the oral cavity of sharks were resistant to clindamycin and 10.9% of the enterobacteria were resistant to ciprofloxacin. The authors concluded that antibiotics

23 such as levofloxacin and amikacin could be reliable alternatives as antibiotic therapy for shark bite victims.

In addition to the microbiome of the mouths of sharks, gut microbiota research has been conducted on select shark species to understand their potential influence on the health of their host. Givens et al. (2015) investigated the gut microbiome of spinner

(Carcharhinus brevipinna), Atlantic sharpnose (Rhizoprionodon terraenovae), and sandbar sharks and bony fish. The authors discovered that both the shark and bony fish guts harbored more diversity than suggested by earlier studies (Newman et al. 1972;

Grimes et al. 1985; MacFarlane et al. 1986; Spangaard et al. 2000; Verner-Jeffreys et al.

2003; Ransom 2008). Increased richness and diversity of the gut microbiome correlated with a more varied diet, and the host may contribute to the community assembly by selecting for microbial populations that include specialized bacteria to aid in the digestion and absorption of nutrients. It has been theorized that the presence of bacteria in sharks aids in the metabolism of urea produced as a normal byproduct of shark metabolism

(Terrell 2014).

Microbiome of Compromised Areas

The most recent study to investigate the microbiome of sharks was conducted by

Pogoreutz et al. (2019) in the Seychelles. This represents the first study to directly compare a healthy versus compromised area of skin on the same individual. The authors assessed whether bacterial community composition differed between visibly healthy and insulted

(injured) shark skin by comparing the bacterial assemblages covering the gills and the back from wild-caught blacktip reef sharks. Overall, 28 of the sampled sharks were visibly healthy and 16 displayed injuries around the gills. The majority of bacterial sequences on

24 the phylum level were assigned to Proteobacteria, Bacteroidetes, Actinobacteria, and

Firmicutes. Overall, the three most abundant bacterial families observed included the

Rhoodobacteraceae, Alteromonadaceae, and Halomonadaceae.

Results showed that shark skin-associated bacterial communities were diverse, and significant differences in the community composition were observed for sharks sampled from different locations. However, while the bacterial communities changed across location of capture, there were no significant differences found between the healthy versus injured areas on the same individual. This suggests the absence of severe bacterial infections after injury, and this overall conserved bacterial community implies that bacterial functions may be maintained in injured skin. The bacteria identified in this study were also previously identified to be a characteristic of shark skin, as discovered in thresher sharks by Doane et al. (2017). The bacteria may have a potentially critical role in the structure of the shark skin microbiome and protect against bacterial infection in injured areas. Thresher sharks and blacktip reef sharks exhibit very different ecological niches and lifestyles, suggesting the shared bacterial genera play a potentially conserved role in shark skin health.

Gaps in Knowledge

Commensal bacteria and other host-associated microbes perform specific roles that benefit the host animal, such as heavy metal detoxification, hindering the formation of biofilm, and preventing the invasion of fungal pathogens (Pogoreutz et al. 2019). This is particularly important for wound healing; whether or not bacterial colonization progresses to an infection depends on the host’s immune response (Pogoreutz et al. 2019).

Fundamentally, microbiome acquisition and stability are thought to be under the control of

25 receptors expressed on the surface of host cells, including those of the innate immune system. Chronic stress is known to impact the immune defense of several teleost species, so disruptions in that system that cause shifts in the microbiome may be the result of environmental stressors (Llewellyn et al. 2014). Ultimately, it has been determined that stress-related imbalances in the microbiome of teleosts may be a precursor to disease. This can be applied to not only teleosts, but to other marine species as well. Determining the bacterial communities present within the microbiome of sharks will be a useful tool in assessing how they contribute to host homeostasis in various tissues and to the overall health of sharks. Characterizing the microbiome in the context of the environment where a particular population resides may help in understanding the relationship between microbiomes and population health. Sharks face multiple stressors including climate change, urbanization, and deteriorating habitats. How their health is affected by these issues will be essential to understanding how they will survive and adapt in the future.

Chapter 3 - Methods

Sample collection

In total, 18 sharks were sampled for this study from three species: 5 blacktip sharks

(Carcharhinus limbatus), 6 tiger sharks, and 7 bull sharks. Sampling began October 2018 and ended in July 2019. Field sampling occurred with The University of Miami Shark

Research and Conservation (SRC) team during weekly shark tagging trips. The target species were captured using a drumline system as described in Figure 1. Each shark was reeled in by hand and either brought onto the deck of the boat, or a partially submerged modified jet ski platform. The shark “work-up” included measurements, blood collection, muscle biopsy, and the insertion of an ID tag. It was during the work-up that the microbe sampling was performed.

Figure 1. Fishing method for safe capture of sharks. (a) bullet floats; (b) large floating poly-ball; (c) rope attaching float to submerged weight; (d) ~18 kg cement weight; (e) hook timer; (f) main line of ~410 kg test monofilament; (g) 4m double-stranded leader of ~900 test monofilament; (h) 16/0 or 13/0 5°-offset circle hook (Gallagher et al. 2014).

26 27

Microbial Sampling

The sampling methodology included using a cell scraper to sample up to four areas of the shark: (1) the mouth, (2) the gills, (3) an uncompromised portion of skin, and (4) any lesions or wounds that were present (Figure 2). A water sample at the capture location was

also collected for each sampled shark. Cell scrapers with microbial samples were stored in a sterile, 30 mL conical tube labeled with the species, sex, date, and tag number and stored on ice until transported to the lab.

Figure 2. Process of microbial sampling. Cell scrapes were taken for each shark of the mouth (A), gills (B), skin (C), and wound (D) if present. Each cell scrape was then stored in a 30 ml conical tube on ice until transport to the lab. Photos by Josh Liberman, Gammon Koval, Nicole Lin, and Matt Bernanke.

Legend

Figure 3. Location of all sampled sharks within the study. Bull (blue), tiger (orange), and blacktip (gray) sharks were all encountered in Biscayne Bay, Florida. The majority of sharks were encountered in the Northern part of the bay.

Laboratory Methods

Marine Bacteria Cultures

Cell scrape samples were used to inoculate marine agar plates to culture bacteria.

Marine agar was prepared (HiMedia Zobell Marine Agar, Catalog number 95021-752) and sterilized before being transferred to sterile petri dishes in 25 mL increments (100mm

29 diameter, VWR, Catalog number 470151-288). After cooling for 15 minutes, plates were inverted and stored at 4oC until use (Appendix A).

A small amount of sterile DI water (5 mL) was added to tubes containing cell scrape samples and gently mixed. An inoculation loop was sterilized using a Bunsen burner and inserted into the sample tube to make contact with the cell scraper before being used to streak the sample onto ¾ of a marine agar plate. The inoculation loop was sterilized a second time and then sample was streaked in a different direction across the plate. This was repeated at least twice. Two plates per sample were prepared and placed in an incubator at 28°C for 48 hours to allow bacterial colonies to grow (Figure 4A).

Subculture of Bacterial Colonies

Following the 48-hour incubation period, plates are examined for bacterial colonies. The abundance and richness of colonies were counted and recorded for each plate.

For each (presumably) different colony, a new subculture plate was prepared using the same method as the initial sample. These subcultures were incubated at 28°C for another

48 hours for additional colony growth (Figure 4B).

Gram Staining

One microscope slide was prepared for each bacterial colony isolated and subsequently stained to identify Gram-negative or Gram-positive bacteria. Briefly, the inoculation loop was sterilized and dipped into a small beaker of sterile DI water to transfer a small water droplet onto a clean microscope slide. The inoculation loop was sterilized again and used to transfer bacterial colonies from the agar plate onto the water droplet. Slides were left to air dry for several minutes before staining (Figure 4C). For gram staining, slides were transferred to a crystal violet stain for one minute and then

30 transferred to a beaker of 60 ml of water for 30 seconds. This was repeated with iodine stain, D-stain, and safranin stain. The remaining colonies on each plate were aseptically collected and transferred to a sterile 1.5 ml microcentrifuge tube and stored in a -20°C freezer until further analysis (Figure 4D).

Figure 4. Process of culturing each cell scrape sample. Each sample is cultured for 48 hours until initial colony richness and abundance can be counted (A). Then, each distinct bacteria is subcultured for another 48 hours (B). Bacterial colonies are collected and placed onto microscope slides (C) and then stored in1.5 ml microcentrifuge tubes until future use.

DNA Extraction

To perform DNA extractions on each bacterial colony, frozen samples were first thawed for five minutes before following the protocol for the Qiagen QIAamp DNA Mini

Kit (catalog number 51304). Briefly, an average of 20 µl of sample was transferred into a new sterile 1.5 ml microcentrifuge tube and 180 µl of Buffer ATL and 20 µl of Proteinase

K was added. The samples were vortexed briefly before being placed in a heating block and incubated at 56°C for 3 hours. Following the incubation, 200 µl Buffer AL was added to each sample and vortexed. The samples were then incubated at 70°C for 10 minutes.

200 µl of 96% ethanol was added to each sample and vortexed for 15 seconds. The solution was then transferred into the QIAamp Mini spin column in a 2 ml collection tube and centrifuged at 6000 x g for 1 minute before the flow-through was discarded. The spin column was washed using centrifugation with Buffers AW1 and AW2 several times before a final wash with Buffer AE, according to the manufacturer’s instructions. Samples were then centrifuged at 6000 x g for 1 minute to elute the DNA. The subsequent DNA sample was tested for purity and concentration using a Thermo Scientific NanoDrop

Spectrophotometer. Samples were then stored at -20°C until PCR was performed.

PCR Reaction

To create a Polymerase Chain Reaction (PCR) product, DNA extracts were first thawed to room temperature. Gene-specific primers for the 16s rRNA gene were used in

PCR reactions for the isolation and amplification of bacterial genes (Forward primer CCT

ACG GGA GGC AGC AG; Reverse primer GGA CTA CHV GGG TWT CTA AT). PCR protocol for the Advantage 2 PCR Kit by Clontech (category number 639206) was followed. The PCR reaction mix was made up of 1 µl of DNA sample, 1 µl of the forward

32 primer, 1 µl of the reverse primer, 5 µl of the 10x PCR buffer, 1 µl dNTP, 1 µl of polymerase, and 40 µl of water. Samples were then transferred to an Eppendorf

Mastercycler (catalog number 6325000013) for PCR using the following cycle parameters: 94°C for 1 minute, followed by 30 cycles of 94°C for 1 minute, 30 seconds at

47°C and 30 seconds at 72°C.

Gel Visualization

To visualize amplification products, 1 µl of each PCR product was run on 1% agarose gel containing SYBR-Safe DNA gel stain (Invitrogen, catalog number S33111).

Each PCR product was placed into the gel’s wells with 6X TrackIt Cyan/Orange Loading buffer (Thermo Fisher Scientific, catalog number 10482028) along with a wide range

DNA ladder in the first well for reference (Takara Bio, code number 3415A). The gel was run at 130v for 40 minutes and was then visualized for bands under UV transillumination in a Gel Doc system.

DNA Preparation

DNA samples that yielded bands of the anticipated product size were cleaned following the Qiagen QIAquick Gel Extraction Kit (category number 28704) protocol.

Briefly, 3 volumes of Buffer QC to 1 volume of DNA sample was added and vortexed.

Isopropanol was added to the sample and mixed to increase the yield of DNA fragments.

The sample was then placed in a QIAquick spin column with a 2 ml collection tube and centrifuged for 1 minute. The flow-through was then discarded and the QIAquick column was placed back into the same collection tube. 0.5 ml of Buffer QG was added to the

QIAquick column and centrifuged for 1 minute. The spin column was washed several times in buffer, according to manufacturer’s instructions. To elute DNA, 50 µl of Buffer

EB (10 mM Tris-Cl, pH 8.5) was added and incubated for 4 minutes at room temperature before centrifuged for 1 minute. The QIAquick column was discarded and the remaining

DNA in the 1.5 ml microcentrifuge tube was stored at -20°C until being sent off to

Eurofins Genomics USA for sequencing.

DNA Sequencing

A subset of bacteria samples for each species of shark underwent DNA extraction to determine which groups of bacteria were present. DNA isolated from cultured bacteria samples from two individuals of each species were submitted for sequencing. Sequencing results were compared to the NCBI BLAST database for microbes. Search parameters for alignments were limited to an E value of < 0.0 and > 95% identified to the genus level.

Data Analysis

Shannon’s Index for biodiversity (H) was calculated for the three shark species in relation to bacterial genera present across the shark. The “count” of genera was calculated by number of tissue sites that bacteria were present on the individual. Shannon’s Index was also calculated for bacteria genera identified in water samples.

Chapter 4 – Results

Microscopy

All cultured bacteria samples were placed on microscope slides and stained to reveal morphology types and Gram-negative versus Gram-positive bacteria. Multiple morphology types were possible within each sample. A total of 188 microscope slides

(61= bull, 66 = tiger, 32 = blacktip, 29 = water) were made of bacterial colonies that were cultured from field samples and stained to reveal Gram-negative and Gram-positive bacteria within each culture.

Gram-negative vs Gram-positive Bacteria

Blacktip and tiger sharks had a higher abundance of Gram-positive bacteria in their mouths, while bull sharks possessed more Gram-negative bacteria in their mouths. The bacteria in the gills and skin tissue sites of all three species were closely distributed, with slightly higher abundances of Gram-negative bacteria in the gills.

Gram-negative bacteria were more abundant than Gram-positive bacteria across all sharks sampled (Figure 6). Gram-negative and Gram-positive bacteria were almost evenly distributed across all water samples (Figure 5).

34 35

Figure 5. Distribution of Gram-negative and Gram-positive bacteria across tissue sites and water samples. Blacktip (A), bull (B) and tiger (C) shark samples tested. D represents all combined water samples. Y axis represents the number of samples in which the Gram type was identified and X axis represents the tissue site.

15 Count of Count Samples 10

0 Blacktip Bull Tiger

Gram-Negative Gram-Positive

Figure 6. Distribution of Gram-staining across all shark species. Y axis represents total number of samples across all tissue sites for the species. X axis represents the shark species.

Distribution of Bacterial Morphology Types

All bacteria samples observed on slides were categorized by morphology type.

Three main morphology types were present across all samples: bacillus, coccus, and spiral.

Each morphology type was also assigned either positive or negative staining attributes for a total of 6 morphotypes: Gram-positive bacilli, Gram-negative bacilli, Gram-positive cocci, Gram-negative cocci, Gram-positive spirilli, and Gram-negative spirilli.

Mouth

The distribution of Gram-negative and Gram-positive coccus and bacillus bacteria was even across all three shark species. Across all species, the mouth had lower abundance of bacteria than the gills and skin tissue sites.

Gills

The distribution of morphology types and Gram-staining was even across all three shark species for the gills. The composition of morphology types was both Gram-negative and Gram-positive coccus and bacillus bacteria.

Skin

Spirilla bacteria was only identified in the skin of bull and tiger sharks. Gram- negative and Gram-positive spirilla were present in skin of tiger sharks, but only Gram- positive spirilla was present in the skin of bull sharks. Blacktip sharks possessed Gram- negative and Gram-positive coccus and bacillus bacteria. For all three species, the distribution of Gram-negative and Gram-positive coccus and bacillus bacteria was evenly distributed.

Wound

Bull sharks possessed higher counts of bacteria morphology types in comparison than blacktips. The distribution of Gram-negative and Gram-positive coccus and bacillus bacteria in the bull shark wounds was even. Blacktip sharks possessed primarily Gram- negative bacillus and Gram-negative coccus bacteria, with lower amounts of Gram- positive bacillus and Gram-positive coccus bacteria.

Water

The two morphology types present in the water samples included coccus and bacillus bacteria, which were almost evenly distributed across samples (Figure 8). No spirilla bacteria was identified within the water samples.

Figure 7. Distribution of bacteria morphology types across all shark species. Gram-negative bacillus is represented as “B-”, Gram-positive as “B+”. Gram-negative coccus is represented as “C-” and Gram- positive as “C+”. Gram-negative spirilla is represented as “S-” and Gram-positive as “S+”. Y axis is number of samples in which the morphology type was present. X axis is the different tissue sites in which samples were collected.

Figure 8. Distribution of bacterial morphology shapes across blacktip (A), bull (B), tiger (C) sharks, and all water samples (D). Y axis represents the tissue sites and X axis represents the number of samples in which the morphology type was present.

Identification of Bacterial Groups

Bacterial Taxonomy Across Tissue Sites

In total, 48 DNA samples were sequenced: 13 from blacktip shark tissue sites, 13 from bull shark tissue sites, 12 from tiger shark tissue sites, and 10 from water samples.

From the 48 DNA samples sequenced, 63 genera were identified in the NCBI database.

The 63 matches in the NCBI Database belonged to 3 phyla, 4 classes, 12 families, and 33 genera (Figure 9).

Legend

Phyla Class Family

Figure 9. Summary of taxonomic groups identified through DNA sequencing. This represents all samples submitted for sequencing for blacktip, bull, tiger sharks, and water.

All samples that were submitted for DNA sequencing contained bacteria belonging to three phyla: Actinobacteria (40%), Proteobacteria (36%) and Firmicutes (24%) (Figure

10). Bacteria belonging to the phylum Proteobacteria were present across all tissue sites and all shark species. Bacteria belonging to phyla Actinobacteria and Firmicutes were only present in blacktip and tiger sharks. While Actinobacteria were present in the skin and gills

41 of blacktip sharks, members of this phylum were only identified in the mouth of tiger sharks

(Figure 11). Firmicutes were present in the gills, mouth, and wound of blacktip sharks and in the gills, mouth, and skin of tiger sharks. Of interest are the bull shark samples, which were exclusively bacteria from the phylum Proteobacteria. All bacteria identified in water samples belonged to the phylum Proteobacteria as well.

Proteobacteria Actinobacteria 36% 40%

Firmicutes 24%

Figure 10. Total composition of bacteria phyla for sequenced samples. Percentage of sequenced samples by phylum for all isolated DNA samples submitted for sequencing. This includes samples from shark tissue sites and water samples.

Percentage Percentage of Samples

Actinobacteria Firmicutes Proteobacteria Figure 11. Distribution of phyla across species and tissue site. Distribution of bacteria phyla for each shark species and tissue sample site, proportional to the number of DNA matches in the NCBI database.

Bacterial DNA isolated from blacktip sharks belonged to 8 families:

Microbacteriaceae, Bacillaceae, Micrococcaceae, Rhodospirillaceae, Staphylococcaceae,

Vibrionaceae, Moraxellaceae and Pseudoalteromonadaceae. The wounds of the blacktip sharks were comprised of 3 families not present on the other tissue sites of the animal

(Staphylococcaceae, Vibrionaceae and Pseudoalteromonadaceae) (Figure 14A). Bacterial

DNA isolated from bull sharks belonged to 3 families: Vibrionaceae, Xanthomonadaceae and Moraxellaceae (Figure 14B). The mouth, gills, and skin were nearly identical in family composition and distribution. However, the cultured samples from the wound only belong to one family, Vibrionaceae.

Bacterial DNA isolated from tiger sharks belonged to 6 families. The samples cultured from the mouth were the most diverse, showing four different families of bacteria

(Figure 14C). The samples cultured from the gills shared the families Moraxellaceae and

Pseudoalteromonadaceae, but the family Staphylococcaceae was unique to this tissue site.

The samples cultured from the skin of these tiger sharks harbored bacteria from two families; Bacillaceae was also present in the mouth, but the Planococcaceae family was unique to the skin.

Blacktip and tiger sharks shared bacteria families Bacillaceae, Micrococcaceae,

Staphylococcaceae, and Pseudoalteromonadaceae (Figure 12). Blacktip and bull sharks shared the family Vibrionaceae, and it was only identified in the wounds. Each shark species also had unique bacteria families; Rhodospirillaceae and Microbacteriaceae was unique to blacktip sharks, Xanthomonadaceae unique to bull sharks, and Planococcaceae unique to tiger sharks. All three shark species shared the family Moraxellaceae. Bacteria

44 belonging to this family were identified in the mouth of all shark species, the gills of bull and tigers, and the skin of blacktip and bulls (Figure 14).

Legend

Figure 12. Venn diagram showing the families of bacteria shared between sharks. Blacktip sharks are represented in gray, bull sharks are represented in blue, and tiger sharks are represented in orange.

Legend

Figure 13. Venn diagram showing bacteria families shared with water samples and sharks. Water samples were combined and not specific to location of sharks.

Legend

Figure 14. Depiction of bacteria families found across tissue sites of each species. Blacktip sharks (A),

bull sharks (B), and tiger sharks (C). Pie charts are not proportional and only indicate presence of bacteria identified through DNA sequencing.

Collectively, bacterial DNA isolated from the various water samples belonged to 5 families: Shewanellaceae, Vibrionaceae, Pseudomonadaceae, Pseudoalteromonadaceae and Rhodospirillaceae. Shewanellaceae and Pseudomonadaceae were not identified in the bacteria samples from tissue sites on any shark species. The most abundant family identified in the water samples, Vibrionaceae, was only also identified in the wounds of both blacktip and bull sharks. The second most abundant family, Pseudoalteromonadaceae, was present in the mouth and gills of tiger sharks as well as the wound of blacktip sharks.

Rhodospirillaceae was also only seen in the gills of blacktip sharks.

Figure 15. Summary of bacteria families found across all water samples. Pie chart is not proportional and only indicates presence of bacteria.

Bacterial Diversity Indices

Bacterial DNA sequencing showed that 33 different genera were present across all samples collected. Isolated bacteria from blacktip sharks belonged to 22 genera, bull sharks belonged to 6 genera, and tiger sharks to 13 genera. Shannon’s Index for biodiversity (H) was calculated for the three shark species. The shark species with the most bacterial diversity was the blacktip shark (H=2.95), followed by the tiger shark (H=2.47) and bull shark (H=1.59). The water samples were less diverse (H=1.83) than blacktip and tiger sharks but more diverse than the bull sharks.

Table 1. Distribution of bacterium genera in the phylum Actinobacteria. Check marks indicate presence of bacteria per tissue site and species is represented by color.

Table 2. Distribution of bacterium genera in the phylum Firmicutes. Check marks indicate presence of bacteria per tissue site and species is represented by color.

Table 3. Distribution of bacterium genera in the phylum Proteobacteria. Check marks indicate presence of bacteria per tissue site and species is represented by color.

Blacktip Bull Tiger

Chapter 5 - Discussion

The bacteria discovered across the microbiome of blacktip, bull, and tiger sharks of this study align with findings of previous marine microbial research. All sequenced samples in this study belonged to the Actinobacteria, Firmicutes, and Proteobacteria phyla.

These phyla have been identified as major bacteria taxa in the gut microbiome of fishes and the skin of sharks and rays (Ghanbari et al. 2015; Johny et al. 2018; Pogoreutz et al.

2019).

Blacktip Sharks

The results of this study indicate that blacktip sharks had the highest bacterial diversity among the species tested (H=2.95). Blacktip sharks are a cosmopolitan species that inhabit both tropical and warm temperate waters around the world. In the eastern

United States, this species ranges from New England to the Florida Keys and Gulf of

Mexico (Castro 2011). Each winter in southeast Florida, thousands of blacktip sharks aggregate in shallow waters close to shore during their annual migration (Castro 2011).

The reason for this migration is most likely temperature, however, other environmental factors may contribute to their movement (Schlaff et al. 2014). All blacktip sharks in this study were sampled during this migration in October and December of 2018.

In the current study, blacktip sharks possessed more Gram-negative bacteria in the gills, skin, and wounds in comparison to gram-positive bacteria. Gram-negative bacteria have long been associated with pathogens in fish, as they are prevalent in the ocean. Gram- negative bacteria appear a reddish-pink color under light microscopy due to the structure of their cell wall, which contains less peptidoglycan than Gram-positive bacteria (Steward

2019). Marine-derived Gram-negative bacteria have been shown to have unique

50 51 and diversely structured lipopolysaccharide residues on their cell surface, which may be related to both recognition and virulence patterns in their hosts (Anwar & Choi 2014).

Examples of common Gram-negative bacteria include enterococci, salmonella, and pseudomonas species. Gram-negative bacteria have features that make them resistant to antibiotic drugs (Peleg & Hooper 2010). In fact, more than 30% of hospital-acquired infections in humans are caused by Gram-negative bacteria (Peleg & Hooper 2010).

Of the bacterial families identified in blacktip sharks, Rhodospirillaceae and

Microbacteriaceae were unique to blacktip sharks and were not identified in bull or tiger sharks. Not only were these two bacterium specific to the species, but they were also only isolated from one tissue site on the sharks: the gills. Microbacteriaceae belong to the phylum Actinobacteria. Actinobacteria bear characteristics of both bacteria and fungi, making them medicinally important due to their ability to produce various bioactive compounds (Verma et al. 2018). Members of this phylum are Gram-positive and represent the most efficient group of prokaryotes with capabilities of producing metabolites (Verma et al. 2018). These metabolites exhibit inhibitory effects against various pathogens and numerous antimicrobial compounds have been isolated from several Actinobacteria and are used as drugs to control human disease (Verma et al. 2018). In fact, approximately two- thirds of naturally occurring antibiotics have been isolated from Actinobacteria species

(Solanki et al. 2008). As the frequency of novel bioactive compounds from terrestrial

Actinobacteria has been decreasing over time, diverse environments like the ocean have been increasingly screened for their ability to produce new secondary metabolites (Kumar et al. 2011).

Members belonging to the Bacillaceae family were identified in the mouth and gills of blacktip sharks. Bacillus was identified in both tissue sites. Like other bacterium,

Bacillus species can either have beneficial or detrimental benefits to their host. Some marine species previously isolated from sponges produce compounds with antimicrobial, antifungal, and antibacterial activity (Devi et al. 2010). Furthermore, species belonging to this diverse genus have demonstrated the ability to inhibit the growth of biofilm on zebrafish (Chu et al. 2014). However, other species belonging to Bacillus have caused splenic rupture in humans and have also been identified in fish tumors (Aoyogi et al. 2009;

Vijayakumar et al. 2015).

Bacilli-shaped bacteria were abundant in the wounds of blacktip sharks. A well- known bacillus includes Vibrio cholerae, which is known to cause cholera (Rogers &

Kadner 2019). One of the most commonly identified group of pathogens belonging to

Gram-negative bacteria are bacteria of the genus Vibrio (Thune et al. 1993). This group of

Gram-negative bacteria can cause high mortality rates in marine fish that manifests as a hemorrhagic septicemia with extensive hemorrhaging and skin lesions (Thune et al. 1993).

DNA sequencing confirmed the presence of Vibrio in samples from the wounds of blacktip sharks. Vibrio species are diverse in function, as some members of the genus are known pathogens, while others are associated with normal tissue. Members of this genus have been known to cause bacterial infections in corals and fish (Thompson et al. 2005; Yilmaz et al. 2018).

Bacterial disease is considered relatively uncommon in elasmobranchs given the small number of documented cases. Of those, Vibrio species have emerged as the most significant group of marine pathogens (Bertone et al. 1996). Vibrio carchariae has been

53 repeatedly implicated as the cause of meningitis in several shark species in previous studies

(Bertone et al. 1996; Terrel 2014) While many species of Vibrio are known pathogens, some isolates from the marine environment have also shown antibacterial compounds

(Mansson et al. 2011). While Vibrio carchariae was identified in the ulcer of a sandbar shark by Bertone et al. (1996), the authors could not determine if the bacteria were the causal agent of the ulcer. In the present study, Macrococcus was also identified in the wounds of blacktip sharks. Macrococcus has been previously associated with tumors in fish by Vijayakumar et al. (2015). Staphylococcaceae was also identified in the wounds of blacktip sharks. Members of this family have been named as the main cause for cardiovascular infections such as Toxic Shock Syndrome in humans (Vanita & Jhansi

2011). Staphylococcus epidermidis has previously reported to cause sporadic epizootics in farmed fishes (Huang et al. 1999). Recently, members of Staphylococcaceae were identified in the intestinal microbiome of juvenile scalloped hammerhead sharks (Sphyrna lewini) in Fiji by Juste-Poinapen et al. (2019). It is not known what role Staphylococcaceae plays in the intestinal tract of sharks, as very little research has been performed on the subject.

While the wounds of blacktip sharks displayed potential pathogens, commensal bacteria were also present. Pseudoalteromonas was identified in the wounds of blacktip sharks. Species belonging to this genus are known to play a variety of beneficial roles to their host, including producing antimicrobial compounds, aiding in prevention of biofouling, and inhibiting the biofilm of human pathogens (Holmstrom et al. 2002; Papa et al. 2015; Offret et al. 2016). Pseudoalteromonas has previously been identified as a core member of the microbiome of other shark species. As mentioned earlier, Pogoreutz et al.

(2019) identified this genus on the gills of blacktip reef sharks. The authors noted that the genus was present in both healthy and insulated areas of the sharks. Additionally, Doane et al. (2017) identified Pseudoalteromonas as being overrepresented in the skin of thresher sharks.

Both Gram-staining and the identification of morphology types demonstrates a pattern of potential pathogenic bacteria in the wounds of blacktip sharks, but it also revealed bacteria that provide benefits to their host. It is likely that Pseudoalteromonas is a normal member of the microbiome of blacktip sharks, as identified in other species by previous studies. While the cause of wounds on the sharks in this study is unknown, it can be theorized that the loss of integrity of the normal epithelial structure could have allowed pathogenic bacteria not otherwise associated with the shark to invade. However, potential pathogens, such as Bacillus, were present in other tissue sites of blacktip sharks as well.

Bull Sharks

Bull sharks are also a cosmopolitan species and are found in warm subtropical and tropical coastal, estuarine, and riverine waters (Bass et al. 1973; Compagno 1984). They are one of the few elasmobranch species capable of moving into freshwater for extended periods of time and are a common inhabitant of Florida’s coastal waters (Ortega et al.

2009). Previous studies have suggested that bull shark movement is influenced by a mix of prey abundance and environmental conditions; bull sharks likely optimize energy allocation through either passive transport, via tidal and river movement, or maintenance of favorable environmental and foraging conditions (Simpfendorfer et al. 2005; Ortega et al. 2009). While bull sharks may not display the same highly migratory patterns that

55 blacktip sharks do, they certainly experience a wide range of environmental conditions throughout their life.

Bull sharks in this study displayed the least amount of bacterial diversity (H=1.59) in comparison to the other two species. Bacteria belonged to three families,

Xanthomonadaceae, Moraxellaceae, and Vibrionaceae, among the tissue sites tested. The mouth, gills, and skin of bull sharks were comprised of Moraxellaceae and

Xanthomonadaceae. Moraxellaceae was confirmed to be present in all three shark species for this study. Members of this family appeared in the mouths of all three species. Within

Moraxellaceae is the genus Psychrobacter. Species of this genus are not novel to the marine environment and have been identified in numerous studies.

The most commonly identified genus was Psychrobacter. This genus was present in the mouth of all three shark species, the gills of bull and tiger sharks, and the skin of blacktip and bull sharks. This genus has been identified on the skin of whales, bony fish, and blacktip reef sharks (Pogoreutz et al. 2019). In the skin of bony fishes, isolates of these bacteria have been shown to inhibit the growth of aquatic fungal pathogens (Lowrey et al.

2015). Notably, the presence of Psychrobacter in the intestinal tract of scalloped hammerhead sharks was used as an indicator for pollution (Juste-Poinapen et al. 2019).

This is due to the observed diversity trends of Psychrobacter that may be related to environmental factors such as salinity and/or anthropogenic pressures (Azevedo et al.

2013).

Xanthomonadaceae was unique to bull sharks and did not appear in blacktip or tiger shark samples, nor any of the water samples. This family includes both helpful and potentially pathogenic bacteria species. Some species belonging to the family are known

56 pathogens that have caused infections in other marine species and even humans. For example, members of Stenotrophomonas can be antibiotic resistant pathogens causing infections such as pneumonia and endocarditis in humans (Brooke 2012).

Stenotrophomonas was identified in the mouth, gills, and skin of bull sharks in the current study. But, several genera of the Xanthomonadaceae family have potential benefits to their host. Pseudoxanthomonas species have the ability to degrade organic pollutants and even mercury (Mahbub et al. 2016). Bacteria belonging to Pseudoxanthomonas were identified in the mouth and skin of bull sharks.

Vibrio was the only genus identified in the wounds of bull sharks, which was also identified in the wounds of the blacktip sharks. This aligns with the findings from the wounds of blacktip sharks in this study. While the present study was unable to determine the cause of the wounds, it is interesting that this genus was not identified on other tissue sites of either species. There is a strong likelihood that Vibrio may be an opportunistic invader to these compromised areas.

While the bull sharks of this study appeared to have the lowest bacterial diversity in comparison to the other two species, there may have been bias during the culturing method as the protocol was selective for marine bacteria species. It is possible that bull sharks’ microbiome may include euryhaline bacteria that was missed during culturing.

Tiger Sharks

Tiger sharks are a large semi-coastal and oceanic species that inhabit temperate and tropical waters (Compagno et al. 2005). These animals are capable of migrating large distances across the globe. Adults have demonstrated annually repeated, round-trip migrations of over 7,500km in the northwest Atlantic (Lea et al. 2015). Due to their wide-

57 ranging diet, tiger sharks are highly connected in marine food webs and can display a wide niche, attributed to variation in prey consumption and depth utilization (Matich et al. 2011;

Vaudo et al. 2014). Previous satellite tracking data indicates that tiger sharks migrate north in spring and summer, as sea surface temperatures rise, but another motivation for seasonal migrations may be foraging opportunities (Lea et al. 2015). Similar to the other two species in the current study, tiger sharks have the flexibility to occupy a variety of oceanic environments, and their movements are influenced by both prey abundance and environmental factors. Tiger sharks in this study displayed the second highest diversity of bacteria (H=2.47), which is reflective of their broad spatial and temporal habitat use.

In the current study, spirilla bacteria were only identified in the skin of tiger and bull sharks. Spiral bacteria are bent and re-bent, forming a shape similar to a corkscrew.

Spirilla are Gram-negative bacteria with a rigid cell wall. There are two important human pathogens within spirilla: Campylobacter jejuni (the bacterial cause of diarrhea) and

Helicobacter pylori (the cause of peptic ulcers) (Todar 2004).

Planococcaceae was specific to tiger sharks and unique to only one tissue site: the skin. One genus belonging to this family identified in the present study, Planococcus, includes some marine species capable of producing antibiotic compounds and have been isolated from marine sponges (Austin & Billaud 1990; Kaur et al. 2012). Some

Planococcus species isolated from beach sediment have the ability to degrade hydrocarbons (Engelhardt et al. 2001). Planococcus can often be found in sea ice, a rich source of diverse bacteria (Bowman et al. 1997; Zhang et al. 2008). As previously mentioned, tiger sharks possess an highly productive mucoid layer in comparison to the other species in this study, which potentially serves as a source of diverse bacteria. As some

58 members of Planococcaceae have demonstrated the ability to produce antibiotic compounds and degradation of hydrocarbons, it is likely an important component of their skin microbiome.

The other families identified in the tissue sites of tiger sharks were shared with blacktip and bull sharks. Tiger sharks shared four families also identified in blacktip sharks:

Bacillaceae, Micrococcaceae, Staphylococcaceae, and Pseudoalteromonadaceae. While

Pseudoalteromonadaceae and Staphylococcaceae were identified in only the wounds of blacktip sharks, these two families were identified in the mouth and gills of tiger sharks.

As previously mentioned, members of these families have been identified in sharks of previous studies.

Water

Pseudomonadaceae and Shewanellaceae were unique to the water samples and did not present in any of the samples from the tissue sites of the three shark species. Members of Pseudomonadaceae have been discovered in marine sponges and display antimicrobial activity against Bacillus, Staphylococcus, and Micrococcus strains (Jayatilake et al. 1996).

Others, such as, Pseudomonas, have been linked to multiple nosocomial infections in humans (Gales et al. 2009). The same bacteria causing infections in humans was identified in the oral cavity of tiger and bull sharks in Brazil by Interaminense et al. (2011). While this bacteria was not identified on the tissue sites of the sharks of the current study, it was cultured from a water sample taken at the location of one of the sampled bull sharks.

Shewanella is the sole genus within the Shewanellaceae family. Shewanella species are commonly isolated from marine environments and like Pseudomonas, Shewanella species have been isolated from marine sponges (Lee et al. 2006; Rachanamol et al. 2014).

Three families identified in the water samples were also identified within shark tissue samples: Vibrionaceae, Pseudoalteromonadaceae, and Rhodospirillaceae.

Vibrionaceae was only identified in the wounds of blacktip and bull sharks. It is possible that the normal microbiome or structure of the epithelia of sharks would normally prevent the intrusion of this pathogen, so the loss of integrity in these compromised areas of the shark allowed the opportunity for Vibrio in the water column to transfer to the sharks.

Pseudoalteromonadaceae was also identified in the mouth and gills of tiger sharks, but within blacktip sharks this bacteria only presented itself within the wounds. This finding suggests that Vibrionaceae and Pseudoalteromonadaceae may not be normal inhabitants of the microbiome of blacktip and bull sharks, but opportunistic invaders from the surrounding water that have entered through a compromised area. However, future work would need to expand water sampling to look for differences between bacteria in the water column and bacteria on the microbiome of sharks before a definitive conclusion can be drawn.

Conclusion

Many of the bacterial communities identified across the microbiome of sharks in this study align with findings of previous research on marine species microbiota. However, to date, Planococcaceae and Rhodospirillaceae has not been previously identified as a member of a shark’s microbiome. The high bacterial diversity identified across blacktip sharks of this study is supportive of their highly migratory behavior through Florida waters.

Tiger sharks displayed the second highest bacterial diversity, which aligns with their wide oceanic migrations and their unique highly productive mucoid layer. While bull sharks displayed the least amount of bacterial diversity but are known to move through wider ranging habitats in terms of salinity, this may have been skewed by only culturing for marine bacteria.

While some families such as Staphylococcaceae and Pseudoalteromonadaceae were only identified on the wounds of blacktip sharks, members of this bacteria showed up in the mouth and gills of tiger sharks. This highlights the potential for bacteria to be normal assemblages in one shark species, but opportunistic invaders to others. The known pathogen, Vibrio, was identified in wounds on both blacktip and bull sharks, but also identified within the water samples. It is unknown whether this bacteria caused the wound, but it is likely a secondary invader after an initial insulation to the dermal denticles. This supports theories that the combination of dermal denticles and mucus production may cause a selective microbiome in sharks that varies by species. This study represents only a snapshot of bacteria characterized across the microbiome of blacktip, bull, and tiger sharks in South Florida waters. Previous research has determined that stress-related imbalances in the microbiome of teleosts could be a precursor to disease and new research suggests that

60 61 this is applicable to many marine species, such as sharks. It is important to continue to investigate bacterial communities within the microbiome of sharks for future comparisons in regard to potential diseases. Certain bacteria can also be used as indicators of pollutants or other anthropogenic stressors that sharks may be encountering. Establishing the core microbiome for shark species will be important for future work in order to differentiate between normal colonizers and pathogens, and to understand how the microbiome may shift in response to environmental changes.

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Appendix A – Marine Agar Preparation

Marine Agar Recipe

27.625g of Himedia Zobell Marine Agar (Catalog number 95021-752) was placed in a 1000 ml Erlenmeyer flask on a stirring heat plate. 300 ml of deionized (DI) water and a stir bar were added while slowly increasing the heat for three to five minutes. After five minutes, 75 ml of DI water was added and the heat increased. The resulting fluid was transferred to a 500 ml volumetric flask and stirred without heat (Figure 7). DI water was added until it reaches 500 ml and stirred for several minutes before being transferred back to an Erlenmeyer flask. It was brought to a boil for at least one minute before being placed into an autoclave safe glass bottle and sterilized using a wet cycle (30 minutes sterilize, 20 minutes heat).

Appendix B – Slide Microscopy

Table B-1. The total number of microscope slides per species and tissue site.

TISSUE SITE BULL TIGER BLACKTIP TOTAL

MOUTH 13 15 7 34

GILLS 15 26 7 48

SKIN 11 25 8 44

WOUND 22 0 11 33

WATER 16 8 5 29

TOTAL 77 74 37 188

Appendix C – DNA Sequencing

Table C-1. Samples submitted for DNA sequencing and results from NCBI BLAST tool. Tag number represents each individual, while sample represents the tissue site and subculture number.

Tag Species Sample Family Genus % Query E Max Number Identified Cover Value Score N392645 Blacktip G1S1 Bacillaceae Fictibacillus 95.22 59 0.00 664 N392645 Blacktip G1S1 Bacillaceae Bacillus 95.19 59 0.00 662 N392645 Blacktip M1S1 Bacillaceae Bacillus 97.43 61 0.00 732 N392645 Blacktip M2S1 Bacillaceae Exiguobacterium 97.42 34 0.00 726 N393575 Tiger M1S2 Bacillaceae Bacillus 98.17 37 0.00 756 N393619 Tiger D2S1 Bacillaceae Lysinibacillus 96.68 35 0.00 701 N393619 Tiger D2S1 Bacillaceae Bacillus 95.75 35 0.00 682 N393619 Tiger D2S2 Bacillaceae Exiguobacterium 98.82 31 0.00 754 N392645 Blacktip G1S1 Microbacteriaceae Curtobacterium 99.02 34 0.00 730 N392645 Blacktip G1S1 Microbacteriaceae Clavibacter 98.77 34 0.00 719 N392645 Blacktip G1S1 Microbacteriaceae Zimmermannella 98.52 34 0.00 713 N392645 Blacktip G1S1 Microbacteriaceae Mycetocola 98.52 34 0.00 713 N392645 Blacktip G1S1 Microbacteriaceae Herbiconiux 97.79 34 0.00 702 N392645 Blacktip G1S1 Microbacteriaceae Cryobacterium 98.02 34 0.00 702 N392645 Blacktip G1S1 Microbacteriaceae Leifsonia 97.79 34 0.00 702 N392645 Blacktip G1S1 Microbacteriaceae Agrococcus 97.79 34 0.00 702 N392645 Blacktip D1S2 Micrococcaceae Kocuria 99.01 57 0.00 728 N392645 Blacktip D1S2 Micrococcaceae Arhtrobacter 98.28 57 0.00 710 N392645 Blacktip D1S2 Micrococcaceae Nesterenkonia 97.78 57 0.00 682 N392645 Blacktip D1S2 Micrococcaceae Rothia 97.04 57 0.00 682 N392645 Blacktip D1S2 Micrococcaceae Micrococcus 96.8 57 0.00 676 N393575 Tiger M1S1 Microococcaceae Kocuria 97.58 52 0.00 706 N393575 Tiger M1S1 Microococcaceae Arthrobacter 97.09 52 0.00 695 N393575 Tiger M1S1 Microococcaceae Nesterenkonia 95.88 52 0.00 667 N393575 Tiger M1S1 Microococcaceae Rothia 95.4 52 0.00 656 N393575 Tiger M1S1 Microococcaceae Micrococcus 95.16 52 0.00 651 N389792 Bull G1S1 Moraxellaceae Psychrobacter 96.45 37 0.00 695 N389792 Bull H2S2 Rhodospirillaceae Thalassospira 95.62 25 5E- 473 130 N389792 Bull H2S2 Pseudomonadaceae Pseudomonas 95.27 25 2E- 468 128 N389751 Bull D2S1 Moraxellaceae Psychrobacter 96.23 35 0.00 691 N389751 Bull M2S1 Moraxellaceae Psychrobacter 95.28 35 0.00 669 N392645 Blacktip D2S1 Moraxellaceae Psychrobacter 96.06 35 0.00 699 N392645 Blacktip M1S1 Moraxellaceae Psychrobacter 96.29 35 0.00 704 N393575 Tiger G1S2 Moraxellaceae Psychrobacter 96.46 35 0.00 699 N393575 Tiger M1S3 Moraxellaceae Psychrobacter 95.96 35 0.00 680

N389792 Bull D1S1 N/A N/A

N389792 Bull M1S1 N/A N/A

N389792 Bull M1S3 N/A N/A

78 79

N389792 Bull W2S1 N/A N/A

N389751 Bull W2S2 N/A N/A

N389751 Bull W2S3 N/A N/A

N392645 Blacktip D1S3 N/A N/A

N392645 Blacktip H1S1 N/A N/A

N393575 Tiger D1S1 N/A N/A N393619 Tiger D2S1 Planococcaceae Paenisporosarcina 96.23 35 0.00 693 N393619 Tiger D2S1 Planococcaceae Planococcus 95.99 35 0.00 688 N392645 Blacktip W1S2 Pseudo- Pseudo- 98.85 36 0.00 773 alteromonadaceae alteromonas N392645 Blacktip W1S3 Pseudo- Pseudo- 99.07 97 0.00 773 alteromonadaceae alteromonas N392645 Blacktip W2S1 Pseudo- Pseudo- 99.08 37 0.00 780 alteromonadaceae alteromonas N393619 Tiger D2S3 Pseudo- Pseudo- 99.3 37 0.00 776 alteromonadaceae alteromonas N393619 Tiger G1S1 Pseudo- Pseudo- 99.08 37 0.00 784 alteromonadaceae alteromonas N393619 Tiger G1S2 Pseudo- Pseudo- 97.94 36 0.00 754 alteromonadaceae alteromonas N393619 Tiger M2S2 Pseudo- Pseudo- 98.4 37 0.00 763 alteromonadaceae alteromonas N392645 Blacktip G1S1 Rhodospirillaceae Thalassospira 95.06 59 0.00 641 N393619 Tiger H1S1 Shewanellaceae Shewanella 96.48 36 0.00 702 N392645 Blacktip W1S1 Staphylococcaceae Macrococcus 99.3 36 0.00 769 N393575 Tiger G1S3 Staphylococcaceae Salinicoccus 98.35 35 0.00 745 N389751 Bull W2S1 Vibrionaceae Vibrio 96.30 40 0.00 706 N392645 Blacktip H1S1 Vibrionaceae Vibrio 98.83 36 0.00 763 N392645 Blacktip H2S1 Vibrionaceae Enterovibrio 97.41 38 0.00 713 N392645 Blacktip W1S1 Vibrionaceae Vibrio 98.84 41 0.00 767 N393575 Tiger H1S1 Vibrionaceae Vibrio 96.5 36 0.00 706 N393575 Tiger H1S2 Vibrionaceae Vibrio 96.29 37 0.00 706 N393619 Tiger H1S2 Vibrionaceae Vibrio 98.08 36 0.00 778 N389792 Bull D1S2 Xanthomonadaceae Stenotrophomonas 96.76 37 0.00 719 N389792 Bull D1S2 Xanthomonadaceae Xanthomonas 96.76 37 0.00 719 N389792 Bull D1S2 Xanthomonadaceae Pseudo- 96.53 37 0.00 713 xanthomonas N389792 Bull M1S2 Xanthomonadaceae Stenotrophomonas 97.22 37 0.00 730 N389792 Bull M1S2 Xanthomonadaceae Xanthomonas 97.22 37 0.00 730 N389792 Bull M1S2 Xanthomonadaceae Pseudo- 96.99 37 0.00 725 xanthomonas N389792 Bull M1S2 Xanthomonadaceae Luteimonas 95.37 37 0.00 686 N389751 Bull G1S2 Xanthomonadaceae Xanthomonas 97.00 96 0.00 728 N389751 Bull G1S2 Xanthomonadaceae Luteimonas 95.16 96 0.00 684 N389751 Bull G1S2 Xanthomonadaceae Stenotrophomonas 97.00 96 0.00 728 N389751 Bull H2S1 Xanthomonadaceae Photobacterium 99.05 40 0.00 758 N389751 Bull H2S2 Pseudo- Pseudo- 99.08 37 0.00 780 alteromonadaceae alteromonas

Appendix D – Statistics for Shannon’s Index Calculations

Table D-1. Genera and counts used to calculate Shannon’s Index for Biodiversity for all blacktip shark tissue sites.

Genus Count Shannon Index Kocuria 1 -0.1363258 Arthrobacter 1 -0.1363258 Nesterenkonia 1 -0.1363258 Rothia 1 -0.1363258 Micrococcus 1 -0.1363258 Fictibacillus 1 -0.1363258 Bacillus 2 -0.212378 Macrococcus 1 -0.1363258 Pseudoalteromonas 2 -0.212378 Psychrobacter 2 -0.212378 Curtobacterium 1 -0.1363258 Clavibacter 1 -0.1363258 Zimmermannella 1 -0.1363258 Mycetocola 1 -0.1363258 Herbiconiux 1 -0.1363258 Cryobacteria 1 -0.1363258 Leifsonia 1 -0.1363258 Agrococcus 1 -0.1363258 Exiguobacterium 1 -0.1363258 Vibrio 1 -0.1363258 Sum 23 2.95467321

Table D-2. Genera and counts used to calculate Shannon’s Index for Biodiversity for all bull shark tissue sites.

Genus Count Shannon Index Psychrobacter 2 -0.3099542 Xanthomonas 2 -0.3099542 Luteimonas 2 -0.3099542 Stenotrophomonas 3 -0.3543499 Pseudoxanthomonas 2 -0.3099542 Sum 11 1.5941667

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Table D-3. Genera and counts used to calculate Shannon’s Index for Biodiversity for all tiger shark tissue sites.

Genus Count Shannon Index Psychrobacter 2 -0.2517725 Salinicoccus 1 -0.1666596 Kocuria 1 -0.1666596 Arthrobacter 1 -0.1666596 Nesterenkonia 1 -0.1666596 Rothia 1 -0.1666596 Micrococcus 1 -0.1666596 Bacillus 2 -0.2517725 Lysinibacillus 1 -0.1666596 Paenisporosarcina 1 -0.1666596 Planococcus 1 -0.1666596 Exiguobacterium 1 -0.1666596 Pseudoalteromonas 3 -0.3061061 Sum 17 2.47624713

Table D-4. Genera and counts used to calculate Shannon’s Index for Biodiversity for all water samples.

Genus Count Shannon Index Photobacterium 1 -0.2441361 Pseudoalteromonas 1 -0.2441361 Thalassospira 1 -0.2441361 Pseudomonas 1 -0.2441361 Vibrio 3 -0.3662041 Enterovibrio 1 -0.2441361 Shewanella 1 -0.2441361 Sum 9 1.83102048