Factors Influencing the Gut Microbiota of Antarctic Seals Tiffanie Nelson

Home , Allofustis, Atopobacter, Fusobacterium, Southern elephant seal

Factors Influencing the Gut Microbiota of Antarctic Seals

Tiffanie Nelson

A thesis in fulfilment of the requirements for the degree of Doctor of Philosophy

Evolution & Ecology Research Centre School of Biological, Earth & Environmental Sciences University of New South Wales, Sydney, Australia April 2012

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Abstract

A mammal’s gastro-intestinal tract houses trillions of bacteria, known as the gut microbiota, which is first acquired during birth. This community provides a number of beneficial functions to the host, including breakdown of food and creation of energy, immunity regulation and cellular development. To date, studies on the mammalian gut microbiota have focused primarily on captive or laboratory-reared terrestrial mammals and there have been a limited number of studies on mammals living under natural conditions or from marine environments. This thesis aimed to address this knowledge gap through a study on the gut microbiota of the southern elephant seal, Mirounga leonina, and the leopard seal, Hydrurga leptonyx, inhabiting Antarctica. The study was designed to understand how time, diet, phylogeny, age, sex, maternal relationship, sewage contamination and captivity influence composition. DNA extracted from faecal samples was analysed via multiple molecular methods to identify an individual’s gut microbiota. The community was dominated by four phyla: Firmicutes, Fusobacteria, Proteobacteria and Bacteroidetes and was identified as stable over a period of months. Diet-driven differences were noted between individuals as a result of species, captivity, sex and age. Specifically, nutrient composition of the diet and bacteria associated with the diet were identified as strong influences on the gut microbiota. Length of the gut, social interactions and antibiotic use were also identified as contributing factors. Despite the observed differences in the gut microbiota between hosts, a ‘core’ microbiota was identified between Antarctic seal species and also between three related species of Arctic seals . This is suggested to be a result of co-evolution which is maintained over generations via transfer from mother to pup. Bacterial isolates possessing virulence factors and resistance to antimicrobial agents were also identified in the gut of these hosts suggesting they may be vulnerable to sewage contamination containing novel bacteria. This research provides a baseline for future work regarding bacterial associations in these hosts. It identifies the complex community inhabiting the gut of mammals and the factors which contribute to its composition. Additionally, it indicates the differences that exist between mammals living in captivity and under natural conditions and has implications for the future health of these hosts. iii

Acknowledgements and Thanks

There are numerous people who have helped me through this journey. I’d like to thank my partner, Tom Rayner, who has been extremely loving and supportive, particularly during the last two years. Specifically, he has read most of my work and discussed it at length with me, which was extremely helpful in improving my skills. My supervisors, Mark Brown and Tracey Rogers, have been very supportive. Mark has always had patience for my endless questions about microbiology, bacteria, DNA sequencing etc. Tracey together with Michaela Ciaglia facilitated many parts of my research and Tracey always provided me with a practical approach to research.

Huge thanks goes to Alejandro Carlini, who taught me so much while literally on the ice and his sudden passing in late 2010 saddened many people. I will never forget his advice on accomplishing tasks in Antarctic conditions, often while holding a cigarette. Funding for field support was provided by the Secretaría de Ciencia y Tecnología with the Dirección Nacional del Antártico (PICTO No. 36054), the Australian Research Council, Winifred Scott Foundation, and the Evolution and Ecology Research Centre, University of New South Wales (UNSW). Research trips involved many transfers between planes, ships, helicopters, and boats and it also included a comfortable stay while at base. In the field, my research would not have been possible without the support provided by Michaela Ciaglia, Javier Negrete, Nigel Edwards, Larry Vogelnest, Nadine Constantinou, David Slip, Matias Baviera, Martin Montes, Karina Smit, Mercedes (Mecha) Santos, Martin Gray, Marie Attard and Mariana Juares. Javier Negrete was an excellent team leader. Nigel, Nadine, Michaela, Mecha and I shared an enjoyable and, sometimes, language-barrier-confusing experience in Primavera. Members of the Argentinean fieldwork team whilst in Jubany were really fun and supportive, especially with my “Spanish”.

Many Taronga Zoo staff supported me with the collection of samples from the captive leopard seals. Thanks go to David Slip, Kaye Humphreys, Rebecca Spindler, Larry Vogelnest, Vanessa Di Giglio, Jane Hall, Karrie Rose and Cheryl Sangster who were

always on hand to answer questions and provide necessary information. Thank you also for assistance from Jose Altunas, Danielle Fox and marine mammal keepers.

Laboratory work is never easy when entering a new field and was heavily supported at UNSW by Bill Sherwin and his lab, including Lee Ann Rollins, Emily Miller, Jackie Chan and Jenny Sinclair. Brett Neilan and members of his lab were always helpful in providing access to equipment and answering questions. Special thanks go to Ivan Wong who has been very helpful. The Ramaciotti Centre was very supportive in the processing of samples and particular thanks go to Tonia Russell. Sequencing completed by the Research and Testing laboratories in Lubbock, Texas was made easier with the help of Scot Dowd. Some specific laboratory tests were carried out at the University of Montreal in Saint-Hyacinthe with John Fairbrother, who was warm and supportive. Whilst there, laboratory work was supported by Clarisse Desautels and members of the Reference Laboratory for Escherichia coli. Special thanks for administrative tasks in Canada go to Jacinthe Lachance.

At UNSW many people supported me. They included, Jonathan Russell, Matt Hunt, Gemma Smart, Stephen Bonser, Rob Brooks, Alan Wilton, Mark Tanaka, Emma Johnston, Bill Sherwin, Alistair Poore, Richard Kingsford, Firoza Cooper, David Eldridge and Angela Moles. Support from David Cohen has also been exceptional. All research related to animal handling was approved by the University of New South Wales Animal Care and Ethics Committee (08/83B and 03/103B).

A number of colleagues provided excellent reviews of chapters and also general advice and support throughout this whole process (emotional and otherwise). These people were Maria Luisa Gutierrez Zamora, Emily Miller, Lee Ann Rollins, Nadine Constantinou, Nigel Edwards, Melanie Sun, Nicole Lima, Jutta Zwielehner, Margo Adler, Alex James, Anna Kopps, James Smith, Jessica Roe, Chris Hellyer, and Patty Zenonos. Lastly but most definitely not least, the Nelson and Rayner families have been very encouraging and loving through this entire process.

Table of Contents

Originality Statement ...... i Copyright Statement ...... ii Authenticity Statement ...... ii Abstract ...... iii Acknowledgements and Thanks ...... iv Table of Contents ...... vi List of Figures ...... x List of Tables ...... xi Chapter 1 ...... 1 Investigating the gut microbiota of wild marine mammals ...... 1 1.1 The mammalian gut microbiota ...... 1 1.1.1 A distinct and rich symbiotic community ...... 1 1.1.2 Factors influencing community composition of the gut microbiota ...... 1 1.1.3 The gut microbiota and host health ...... 2 1.1.4 The gut microbiota of non-human hosts ...... 2 1.2 Analysing microbial communities ...... 4 1.2.1 Microbial analysis and taxonomy ...... 4 1.2.2 Community fingerprinting ...... 4 1.2.3 Next-generation sequencing ...... 5 1.3 Life history traits and status of marine mammals, specifically phocid seals, in relation to current understanding of the gut microbiota ...... 6 1.3.1 Exceptional traits of marine mammals ...... 6 1.3.2 Phocid taxonomy and distribution ...... 7 1.3.3 The southern elephant seal ...... 8 1.3.4 The leopard seal ...... 10 1.3.5 Current status of marine mammals with a focus on phocid seals ...... 12 1.3.6 Current understanding of the gut microbiota of phocid seals ...... 12 1.4 The Antarctic environment ...... 13 1.5 Thesis organisation ...... 17 Chapter 2 ...... 18 Stability of the gut microbiota of captive leopard seals: dietary influence and longitudinal change ...... 18 2.1 Abstract ...... 18 2.2 Introduction ...... 19 2.3 Materials and methods ...... 20 2.3.1 Sample collection from captive leopard seals and fish ...... 20

2.3.2 Genomic DNA extraction and community fingerprinting using ARISA ...... 21 2.3.3 Data analyses ...... 22 2.4 Results ...... 23 2.4.1 Stability of the gut microbiota ...... 23 2.4.2 The microbiota associated with the intestinal tract and the faeces ...... 27 2.4.3 Transfer of gut microbiota from fish to seal ...... 28 2.5 Discussion...... 29 2.5.1 The gut microbiota of captive leopard seals is stable over time ...... 29 2.5.2 The gut microbiota represents the lower intestinal microbiota...... 31 2.5.3 Diet introduces bacteria into the gut ...... 32 2.6 Conclusions ...... 33 Chapter 3 ...... 34 Patterns influencing the gut microbiota of Antarctic seals ...... 34 3.1 Abstract ...... 34 3.2 Introduction ...... 35 3.3 Materials and methods ...... 36 3.3.1 Sample collection from Antarctic seals ...... 36 3.3.2 Genomic DNA extraction and community fingerprinting using ARISA ...... 38 3.3.3 Data analyses ...... 38 3.4 Results ...... 39 3.4.1 Influence of host species and captivity on the gut microbiota...... 39 3.4.2 Influence of age-class and maternal transfer on the gut microbiota of southern elephant seals ...... 42 3.4.3 Influence of sex on the gut microbiota ...... 44 3.5 Discussion...... 46 3.5.1 Gut morphology, physiology and diet shape the gut microbiota of wild southern elephant seals and leopard seals ...... 46 3.5.2 Captivity and illness exert a strong affect on the gut microbiota of mammals ...... 47 3.5.3 Age-class and maternal transfer shape the gut microbiota of southern elephant seals ...... 48 3.5.4 Sex-driven dietary differences influences the gut microbiota of southern elephant seals ...... 49 3.6 Conclusions ...... 50 Chapter 4 ...... 51 The gut microbiota of wild and captive Antarctic seals as identified using 16S rRNA gene pyrosequencing ...... 51 4.1 Abstract ...... 51 4.2 Introduction ...... 52 4.3 Materials and methods ...... 53 4.3.1 Sample collection and DNA extraction from Antarctic seals ...... 53

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4.3.2 Community sampling via 16S rRNA gene pyrosequencing ...... 54 4.3.3 Data analyses ...... 54 4.4 Results ...... 55 4.4.1 16S rRNA gene pyrosequencing of the seal gut microbiota ...... 55 4.4.2 Influence of age-class on the gut microbiota of southern elephant seals . 56 4.4.3 Influence of host species and captivity on the gut microbiota...... 59 4.4.4 Shared OTUs within and between seal hosts...... 62 4.4.5 The influence of sex and the external environment on the gut microbiota 67 4.5 Discussion...... 68 4.5.1 ARISA and 16S rRNA gene pyrosequencing methods are compatible .... 68 4.5.2 Diet and gut physiology drive differences in the gut microbiota...... 69 4.5.3 A conserved core gut microbiota exists in Antarctic seal hosts ...... 70 4.5.4 Antibiotic-use and captivity may exert a strong influence on the gut microbiota ...... 71 4.5.5 Social interactions increase the transmission of OTUs between hosts .... 72 4.5.6 Negligible contributions to the gut microbiota from the external environment ...... 73 4.5.7 Sexual dimorphism drives dietary influences on the gut microbiota ...... 74 4.6 Conclusions ...... 74 Chapter 5 ...... 76 Comparing the gut microbiota of mammals from marine and terrestrial habitats ...... 76 5.1 Abstract ...... 76 5.2 Introduction ...... 77 5.3 Materials and methods ...... 78 5.3.1 Data acquisition ...... 78 5.3.2 Data analyses ...... 79 5.4 Results ...... 79 5.4.1 The gut microbiota of Antarctic and Arctic phocid seals ...... 79 5.4.2 The gut microbiota of marine and terrestrial mammals ...... 82 5.5 Discussion...... 87 5.5.1 A shared core gut microbiota in Antarctic and Arctic phocid seals ...... 87 5.5.2 A distinct marine mammal gut microbiota ...... 88 5.5.3 Richness of the gut microbiota is influenced by diet type ...... 89 5.6 Conclusion ...... 90 Chapter 6 ...... 91 Investigating Escherichia coli pathotypes and antimicrobial resistance patterns in wild southern elephant seals, Antarctica ...... 91 6.1 Abstract ...... 91 6.2 Introduction ...... 92 6.3 Materials and methods ...... 93 viii

6.3.1 Study site ...... 93 6.3.2 Sample collection and bacterial growth from southern elephant seals ..... 95 6.3.3 Sample collection and bacterial growth from seawater ...... 95 6.3.4 Sample processing from rectal swabs ...... 96 6.3.5 Identification of virulence factors by multiplex PCR ...... 96 6.3.6 Isolation of bacteria positive for virulence factors ...... 99 6.3.7 Antimicrobial resistance (AMR) testing ...... 99 6.3.8 Data analyses ...... 99 6.4 Results ...... 100 6.4.1 Prevalence of E. coli virulence factors ...... 100 6.4.2 Prevalence of AMR ...... 102 6.5 Discussion...... 104 6.5.1 E. coli pathotypes are present in southern elephant seals ...... 104 6.5.2 Human sewage increase the presence of AMR isolates in seawater ..... 104 6.5.3 Sewage-contaminated seawater is a possible transmission route to southern elephant seals ...... 105 6.6 Conclusion ...... 106 Chapter 7 ...... 107 The gut microbiota of Antarctic seals ...... 107 7.1 Introduction ...... 107 7.2 A distinct and conserved phocid seal gut microbiota ...... 107 7.3 Antarctic phocids have a close relationship with the Fusobacteria ...... 108 7.4 Introductions into the gut from prey but few from the external environment ...... 109 7.5 Captivity has a strong influence on the gut microbiota of mammals .... 110 7.6 Future directions ...... 110 APPENDIX 1 ...... 111 APPENDIX 2 ...... 115 APPENDIX 3 ...... 117 APPENDIX 4 ...... 121 APPENDIX 5 ...... 135 APPENDIX 6 ...... 143 References ...... 146

List of Figures

Figure 1.1 Descriptive phylogeny of families from the order Carnivora...... 9 Figure 1.2 Descriptive phylogeny of the family Phocidae...... 10 Figure 1.3 Southern elephant seals and leopard seals in Antarctica...... 11 Figure 1.4 Location of Antarctica...... 15 Figure 1.5 Location of the western Antarctic Peninsula highlighting sampling sites . 16 Figure 2.1 Similarity of female and male leopard seal gut microbiota over time...... 25 Figure 2.2 Relative abundance of characteristic OTUs in the gut microbiota of leopard seals through time...... 26 Figure 2.3 Relative abundance of shared and unique OTUs in the gut microbiota of captive leopard seals...... 27 Figure 2.4 Similarity of the microbiota of intestinal sections and faeces...... 28 Figure 2.5 Relative abundance of characteristic OTUs along the intestinal tract and in the faeces...... 28 Figure 2.6 Shared OTUs between leopard seals and their fish diet...... 29 Figure 3.1 Similarity of the gut microbiota of seal host species groups...... 40 Figure 3.2 Shared OTUs between the gut microbiota of individuals within seal host species groups...... 42 Figure 3.3 Stability and similarity of the gut microbiota of captive leopard seals...... 43 Figure 3.4 Similarity and shared OTUs in the gut microbiota of mother-pup pairs. .. 45 Figure 3.5 Similarity of the gut microbiota of seal host species based on sex...... 46 Figure 4.1 Average relative abundance of major phyla in the gut microbiota of seal host species groups...... 58 Figure 4.2 Microbial richness of the gut microbiota of seal host species groups...... 58 Figure 4.3 Similarity of the gut microbiota of seal host species groups...... 60 Figure 4.4 Characteristic OTUs in the gut microbiota of seal host species groups. . 61 Figure 4.5 Changes in the gut microbiota of captive leopard seals through time. .... 62 Figure 4.6 Similarity and shared OTUs in the gut microbiota of mother-pup pairs. .. 64 Figure 4.7 Shared OTUs within and between host species groups...... 66 Figure 4.8 Average relative abundance of major phyla in the gut microbiota of southern elephant seal adult and sub-adult males and females...... 68 Figure 5.1 Influence of diet and habitat on the gut microbiota of mammals...... 83 Figure 5.2 Average relative abundance of major phyla in the gut microbiota of mammals based on groupings of location and diet...... 85 Figure 5.3 Similarity of the gut microbiota between hosts from the order Carnivora. 86 Figure 5.4 Microbial richness of the mammalian gut microbiota based on groupings of location and diet...... 87 Figure 6.1 Potter Cove, 25 de Mayo / King George Island displaying sampling locations...... 94 Figure A3.1 Relationship between characteristic OTUs and mass of southern elephant seal adults...... 120 Figure A4.1 Relationship between microbial richness, characteristic phyla and OTUs with mass of southern elephant seal adults and sub-adults...... 133 Figure A5.1 Phylogeny of the family Phocidae...... 139 Figure A5.2 Similarity of the gut microbiota of host mammals with captive and wild representatives...... 139 Figure A5.3 Similarity of the gut microbiota of mammals grouped by diet and location...... 140 x

List of Tables

Table 2.1 PERMANOVA on the gut microbiota of male and female captive leopard seals over time...... 24 Table 2.2 Similarity of the gut microbiota of captive leopard seals over time...... 25 Table 3.1 PERMANOVA on the gut microbiota of hosts based on groupings of species, location, age-class and sex...... 39 Table 3.2 Similarity of the gut microbiota of hosts based on groupings of species, age-class and sex...... 41 Table 4.1 PERMANOVA on the gut microbiota of hosts based on groupings of species, location, age-class and sex...... 57 Table 4.2 Shared OTUs in the gut microbiota of mother-pup pairs...... 65 Table 4.3 Shared OTUs in the gut microbiota of host species groups...... 66 Table 4.4 Shared OTUs in the gut microbiota of southern elephant seals and wild leopard seals...... 67 Table 5.1 Shared genera in the gut microbiota of Antarctic and Arctic seals...... 80 Table 5.2 Nearest seal and non-seal sources of shared sequences in the gut microbiota of phocid seals...... 81 Table 5.3 ANOSIM of the gut microbiota of host mammals based on groupings of diet, location, phylogeny and gut morphology...... 84 Table 5.4 Student’s t-test of the main phyla in the gut microbiota of mammals based on groupings of location and diet...... 85 Table 6.1 Multiplex PCR and virulence factors used to detect E. coli pathotypes. ... 97 Table 6.2 Multiplex PCR conditions and control strains used for detection of virulence factors in E. coli isolates...... 98 Table 6.3 Antimicrobial agents and their diameter sensitivity interpretation...... 100 Table 6.4 Relationship between the presence of virulence factors of E. coli and site of origin in faecal samples from seals of different age-classes and seawater...... 101 Table 6.5 Occurrence of E. coli pathotypes in southern elephant seal faecal samples grouped by age-class...... 102 Table 6.6 Relationship between AMR and site of origin of E. coli isolates...... 103 Table A1.1 Characteristics of Antarctic seals, fish and seawater samples...... 111 Table A1.2 Clinical notes on the status of the female captive leopard seal during sample collection...... 114 Table A2.1 Characteristic OTUs in the gut microbiota of captive leopard seals grouped by sex and month...... 115 Table A3.1 Characteristic OTUs in the gut microbiota of seal hosts grouped by species, age-class, sex and location...... 117 Table A3.2 Shared OTUs between most southern elephant seal mother-pup pairs...... 119 Table A4.1 Abundance of representative phyla and classes in the gut microbiota of wild and captive Antarctic seals...... 121 Table A4.2 Characteristic OTUs in the gut microbiota and their taxonomy grouped by species, age-class, sex and location...... 122 Table A4.3 Gut microbial richness of captive leopard seals over time...... 124 Table A4.4 Shared OTUs in the gut microbiota of most southern elephant seal adults and sub-adults...... 125 Table A4.5 Shared OTUs in the gut microbiota of most wild leopard seals...... 128

Table A4.6 Shared OTUs in the gut microbiota of captive leopard seals over time...... 130 Table A4.7 Shared OTUs in the gut microbiota of most southern elephant seal adults and sub-adults and captive leopard seals...... 132 Table A4.8 PERMANOVA on gut microbiota of southern elephant seal adults and sub-adults with mass...... 132 Table A4.9 Nearest sequence matches to the genus Cetobacterium in GenBank. 134 Table A5.1 Characteristics of mammalian hosts used in the study...... 135 Table A5.2 Characteristic OTUs in the gut microbiota of mammal hosts grouped by diet and location...... 141 Table A6.1 Prevalence of E. coli pathotypes in seawater...... 143 Table A6.2 Prevalence of E. coli pathotypes in faecal samples from mother-pup pairs...... 143 Table A6.3 Relationship between intermediate AMR and site of origin of E. coli isolates in faecal samples from seals of different age-classes and seawater samples...... 144 Table A6.4 Information pertaining to E. coli control strains used in multiplex PCRs...... 145

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Chapter 1

Investigating the gut microbiota of wild marine mammals

1.1 The mammalian gut microbiota

1.1.1 A distinct and rich symbiotic community

Most animals maintain symbiotic partnerships with bacteria. Vertebrate animals form associations with highly complex communities of bacteria. In these, apparently symbiotic, associations the bacterial communities often outnumber the host’s own body cells [1]. For instance in humans, bacterial cells outnumber host body cells by a factor of ten, equivalent to 100 trillion cells [2,3]. These communities inhabit the skin, oral cavity and gastro-intestinal tract among others and are distinct to the habitat, performing numerous functions for the host [4].

The gastro-intestinal tract, known henceforth as the gut, of mammals can have an extremely dense population of bacteria, more so than other host habitats [1]. Functionally this community is akin to a hidden organ with a metabolic activity in excess of the liver [5,6] and with an estimated 150-fold more genes than the human genome [7-9]. These functions expand the hosts’ physiological capabilities to harvest energy from food items, develop and regulate the immune system and aid in cellular maturation of the gut [3,10,11]. Studies in humans have elucidated a highly diverse community at the species level with most bacteria belonging to just a few dominant phyla, primarily the Firmicutes and the Bacteroidetes [12,13]. Adults display remarkable inter-individual differences in community composition [12] and estimates of microbial richness range from 150 to 500 bacterial species [13,14].

1.1.2 Factors influencing community composition of the gut microbiota

Mammals acquire their first gut microbiota during transfer through the birth canal [15- 17]. The infant’s gut is marked with a dynamic microbial community less diverse than that inhabiting an adult gut [15]. Composition is heavily influenced by parturition and the events immediately following; differences, for instance, have been observed as a result of mode of delivery (i.e. vaginal or caesarean), and type of feeding (i.e. breast

milk or formula) [16,18-20]. The gut microbiota of humans stabilises to resemble an adult community at around two years of age coinciding with the introduction of solid foods [15].

Throughout life, hosts are continuously exposed to bacteria from their surroundings, primarily through the ingestion of food and drink, but also via a promiscuous process of transmission from multiple sources [12,21,22]. The host influences the gut microbial composition through diet type, genotype, physiology, gut morphology and disease or immune status [10,23].

1.1.3 The gut microbiota and host health

The gut microbiota plays an important role in host health. The microbial community primes the immune system from early associations and provides a barrier to infection [24]. When host defences are compromised or disease ensues, shifts in composition of the gut microbiota have been identified with pathologies, such as colon cancer [25], obesity [26] and metabolic disorders [27]. Antimicrobial treatments also alter the gut microbiota considerably, predominantly in the short-term [28] although evidence suggests effects may remain for extended periods [29].

Understanding the influence of the gut microbiota on host health requires an initial understanding of the background ‘normal’ community and the factors influencing its composition. To date, few ‘top-down’ nor ‘bottom-up’ influences of the gut microbiota and host pathologies have been clearly deciphered [30,31]. Current research projects such as, The Human Microbiome Project, are focused on resolving these complexities [5].

1.1.4 The gut microbiota of non-human hosts

Knowledge of the gut microbiota of mammal hosts has come from studies using captive or laboratory-reared mammals [13,32-34] and also through the use of domesticated hosts associated with agriculture [e.g. 14,35]. These studies have provided enormous insight into complex host-microbial relationship, yet the relevance of this information in its application to non-human mammals is unclear.

In a pioneering study conducted by Ley et al. [23] the faecal microbiota from a wide variety of terrestrial mammals were compared. This study was (and remains) the largest of its kind, including 59 mammalian species living either in zoos or in their wild ecosystem. The study resolved that the gut microbiota were broadly influenced by phylogeny, diet and gut morphology and that the bacterial community has co- diversified with their hosts. Subsequent studies have built on this knowledge, identifying that the gut microbiota of different hominid species is linked to their phylogenetic lineages [36,37]. In a more recent study, Muegge et al. [38] suggested that diet is the convergence of gut microbiota between hosts and this is not a result of evolutionary lineage.

Overall, these studies have provided significant advances in our understanding of gut microbial associations with mammals. However, in many of these studies, faecal samples were sourced from mammals living in a zoo or captive environment [23,38]. The inherent variability present in natural systems (familiar to ecologists) suggests investigating the gut microbiota of wild mammals may provide further insight into this functional relationship. It would allow identification of crucial processes influencing the gut microbiota relevant to their natural lifestyle. A number of studies on birds and primates have highlighted the differences in the gut microbiota of hosts in a captive environment compared with living in the wild [39-43].

Current understanding of the gut microbiota of non-human hosts has been further limited by the omission of marine mammals, on which very few studies have been carried out [44-46]. The distinct lifestyle traits and potentially important differences in anatomy, phylogeny, and diet of marine mammal hosts compared with terrestrial hosts is reason enough to initiate further studies including these mammals. The comprehension of the evolution of the gut microbiota relies on the inclusion of as many distinct dietary and phylogenetic groups as possible, and as many marine mammals represent unique lineages [47], this may provide further insight into the co- evolution of the gut microbiota of mammals.

1.2 Analysing microbial communities

1.2.1 Microbial analysis and taxonomy

Since the late 19th century culture-dependent techniques have been employed to identify bacterial species. With the development of the polymerase chain reaction (PCR) and its application to the amplification of 16S ribosomal RNA (rRNA) genes from both cultures and DNA extracted directly from environmental samples, the perception of microbial diversity has changed considerably [48]. Implementation of these methods has led to the identification of the Archaea as one of three Domains of life as well as the existence of many phyla, genera and species that were previously unknown [49,50].

Today, there are numerous ways to approach microbial community analyses, and, as with many studies in ecology, the choice of method depends on the ecological question being asked [51]. Molecular methods are routinely employed to explore the composition of microbial communities, and have been used to define patterns in microbial community structure and function across a wide range of environments including soils [52,53], air [54,55], deep oceans [56,57], contaminated lands [58,59], polar environments [60,61] and in host-associations [4,37]. This has provided significant advances in our understanding of processes on earth and allowed the construction of conceptual frameworks to identify patterns. Here, I outline two relevant community analysis methods.

1.2.2 Community fingerprinting

Community or DNA fingerprinting techniques have been designed to provide a ‘snapshot’ of the microbial community from any given sample. A variety of methods exists including denaturing gradient gel electrophoresis (DGGE) [62], terminal restriction fragment length polymorphism (T-RFLP) [63], and automated ribosomal intergenic spacer analysis (ARISA) [64]. All use PCR for the amplification of ribosomal genes, followed by a method to detect the variety present within the amplified fragments. The methods are well established and generally require no significant investment in terms of development. They provide fast, relatively inexpensive and reproducible estimates of community composition in the form of 4

DNA fragment profiles which can be considered analogous to community composition data gathered by ‘classical’ ecologists [65].

ARISA utilises the inherent length heterogeneity of the intergenic transcribed spacer (ITS) region between prokaryotic 16S and 23S rRNA genes [64]. PCR-amplification across this region provides a characterisation of the taxa present in the sample [65]. Although DNA fingerprinting techniques are susceptible to methodological artefacts which may occur during amplification, ARISA appears to provide robust and repeatable profiles [64,66] and where tested, ARISA data has correlated strongly with actual cellular abundance data [65]. Numerous studies have proved the usefulness of ARISA in comparing and understanding microbial communities in a diverse range of environments [e.g.67,68-72].

Limitations to the ARISA method include overlap in the size of the ITS region; one ARISA fragment does not always equal one ‘species’ of bacteria [73]. Conversely, some bacteria with multiple rRNA operons (groups of genes) may produce multiple ARISA peaks [65,74]. The ARISA method may also miss the detection of some members of the community which are present in very low abundances [75] as well as certain bacteria, including the phylum of aquatic bacteria, Planctomycetes, which may not contain linked 16S-23S rRNA operons and therefore lack an ITS region [76]. Regardless, differences in ARISA profiles reflect true differences in community composition and thus can be correlated to abiotic factors [77,78]. The method allows for a high throughput of samples, facilitating spatial and temporal analysis of microbial communities and adherence to experimental design principles [79,80].

1.2.3 Next-generation sequencing

Sequence-based methods, as indicated in reviews by numerous authors [e.g. 81,82- 84], have revolutionised the study of the ecology and evolution of Bacteria and Archaea [85,86]. Sequencing of the 16S rRNA gene universal in all bacteria allows taxonomic relationships to be identified [48,87,88]. The 16S rRNA gene is ~1,550 base pairs (bp) long and is composed of both variable and conserved regions [89]. Sequencing hyper-variable regions (~400 bp long) of the gene instead of the whole gene often provides sufficient information to resolve the taxonomic relationships of 5

an organism down to the species level [90,91]. Publicly-available databases, such as the Ribosomal Database Project v.10 (RDP) [92] and SILVA [93], which house many thousands of well curated, full length 16S sequences (e.g. 1,921,179 sequences in release 10.27 of the RDP) allow for the rapid and accurate identification of taxa.

Sequencing methods are generally favoured because they allow investigators to detect and quantify organisms that may be difficult to culture leading to an improved understanding of prokaryotic richness [88]. There are some limitations to these methods, such as the accurate differentiation between closely related species [94,95]. There is also a substantial cost, both in money and post-production data analysis, associated with sequencing methods [90]. The recent surge in accessible, lower cost sequencing, such as that provided by Roche 454 and illumina next generation sequencing platforms, has resulted in significant data generation. However, for many studies there is still limited replication of samples, thereby reducing the power to answer scientific questions [96]. Hence, the opportunity of cost-effective data generation, is not necessarily aligned with efficient knowledge generation [97].

1.3 Life history traits and status of marine mammals, specifically phocid seals, in relation to current understanding of the gut microbiota

1.3.1 Exceptional traits of marine mammals

Marine mammals are a relatively small taxonomic group, yet given their biomass and position in the global food web; they are a biologically important part of marine biodiversity [98,99]. Marine mammals are comparably different in both their physiology and anatomy to terrestrial mammals. However, compared to terrestrial mammals, marine mammals are less well studied: 38% compared with 15% data deficient on the IUCN Red List (www.iucnredlist.org) [100]. This is largely due to the aquatic environment, where marine mammals spend most of their time, making observing or testing individuals comparatively more challenging [101,102]. However, novel methods and the application of technologies in novel ways, such as accelerometers, critter cams and GPS, are addressing this inability to observe marine mammals [103-105].

The constraints of an underwater lifestyle have caused marine mammals to undergo considerable physiological, anatomical and ecological adaptations during their evolution. Adaptations have evolved primarily to maximise underwater foraging times [106]. These include systems to thermoregulate, osmoregulate, breathe and communicate [107-109]. In particular, there is significantly greater potential for heat loss in marine mammals, particularly in polar and deepwater environments (the specific heat loss of water is 25 times that of air), and therefore species have evolved adaptations to compensate for this metabolic expense [106,110,111]. Another example is the comparatively longer intestinal lengths and increased metabolic rate of marine compared with terrestrial carnivores which is thought to allow for adequate digestion of prey [112].

1.3.2 Phocid taxonomy and distribution

The family Phocidae (class Mammalia, order Carnivora) are marine mammals representing the ‘true’ or earless seals and contain 15 extant species [113,114]. Together with the Otariidae (eared seals: sea lions and fur seals) and the Odobenidae (walrus) they constitute 34 extant members of a higher-level group of aquatic carnivores termed Pinnipedia (Figure 1.1) [113]. Rough estimates of pinniped abundance suggest there are around 30 million individuals [115], which are concentrated at higher latitudes, especially near Antarctica [116,117]. Debate has ensued over whether pinnipeds originated from one (monophyletic) or two (diphyletic) ancestral stocks [118], yet several recent studies have used molecular techniques to identify the Pinnipedia as a monophyletic group [114].

The last common ancestor of pinnipeds is thought to have been shared with an arctoid carnivore more than 25 million years ago (mya) making their nearest relative either mustelids (weasels) or ursids (bears) (Figure 1.1) [113,114]. Within the family Phocidae, there are two subfamilies: Monachinae or ‘southern’ hemisphere seals (comprising Antarctic, elephant and monk seals) and Phocinae or ‘northern’ hemisphere seals (10 species that inhabit the Arctic and sub-Arctic regions) (Figure 1.2) [47]. These two families are thought to have split into their respective groups between 15 to 23 mya [114,119].

1.3.3 The southern elephant seal

Southern elephant seals, Mirounga leonina, are wide-ranging predators breeding in colonies on ice-free land, primarily islands, up to 20° latitude in the Antarctic and Southern Ocean (Figure 1.3A-C) [103,120-122]. The total number of animals is estimated to be around 600,000 [115] making them a significant contributor to Antarctic biomass. In recent decades they have suffered and recovered from declines in abundance as a result of shifts in resource availability, a consequence of the animals fidelity to foraging and breeding sites [120,121,123,124]. Breeding female adults aggregate in large groups (harems) to give birth and males compete for access to mate with them [124-126]. After breeding they spend around 70 days at sea and then come ashore again for one month to molt (seasonal shedding of fur) [122].

While at sea, southern elephant seals undergo long migrations, travelling distances up to 5,000 km and reaching depths greater than 1,500 m, staying submerged for up to 120 minutes [127-129]. During foraging trips, they feed on many different species of cephalopods (squids) and fish with compositional differences in prey reflected in the age and sex of the individual [130-133].

Figure 1.1 Descriptive phylogeny of families from the order Carnivora. Adapted from Agnarsson et al. [134]. Members of the Phocidae are marked with red circle.

Figure 1.2 Descriptive phylogeny of the family Phocidae. Adapted from Berta et al. [135] based on molecular data from Davis et al. [113].

1.3.4 The leopard seal

Leopard seals, Hydrurga leptonyx, are solitary animals that inhabit the pack ice surrounding the Antarctic continent (Figure 1.3D-F) [136-138]. Information on the ecology of these animals is limiting due to their elusive nature and sparse distribution [139,140]. Generally, they are thought to stay within a small radius (~50 km) of the Antarctic continent and on occasion move north [136,141,142]. They are completely dependent on ice floes for survival and different sized floes are selected for pupping, resting, molting and for protection against predation by the killer whale, Orcinus orca [136,138,143-145]. Estimates of population abundances on the eastern side of Antarctica, where most of the research on these animals has been conducted, suggest 12,000-15,000 individuals inhabit the region [139,146]. Total estimates surrounding the continent are thought to be in the order of 220,000 representing a relatively small portion of total Antarctic seals (~0.3-1.4%) [115,147]. 10

Regardless of the comparatively lower number of individuals, leopard seals, are an important apex predator in the Antarctic ecosystem [140,148,149]. They forage for a variety of prey, including seals, birds, penguins, fish, cephalopods and krill [148,150- 152]. Often referred to as a generalist predator, leopard seals are now thought to vary the composition of their prey according to regional and seasonal differences [148,153-156]. Some individuals may even become specialist hunters, concentrating on a particularly vulnerable and abundant prey in one region [152,153,157].

Figure 1.3 Southern elephant seals and leopard seals in Antarctica. Southern elephant seal females come ashore to birth in colonies forming harems (A) fought over by male adults for breeding rights (B). After breeding, during the molt, sub-adults are often observed in large groups on land (C). Leopard seals inhabit the pack ice and are almost always observed in isolation (D, E) except during breeding when they occur in mother-pup pairs (F). Photo credit B: Matias Baviera, all other photos: Tiffanie Nelson. 11

1.3.5 Current status of marine mammals with a focus on phocid seals

Marine mammals are of significant conservation concern with 23% of species currently threatened with extinction: 10% are considered vulnerable, 11% endangered and 3% critically endangered [98,116]. Due to the difficulty in gaining adequate data on marine species, it has been estimated that declining populations may go unnoticed at least 70% of the time [158]. Many of the traits exceptional to marine mammals, such as large body size, long gestation periods and low number of offspring, make them inherently vulnerable to extinction [98,159]. In addition, their distribution across international boundaries and in international waters, such as phocid seals in Antarctica, makes managing populations politically difficult [e.g. 160]. In general terms, there are numerous threats that have been linked to their decline and several excellent reviews have discussed these [see 161,162,163]. These threats include habitat degradation, harvesting (specifically hunting for food and materials), accidental mortality (from vessel strike and through fisheries by-catch) and pollution (including chemical contaminants, marine debris and noise) [98].

For phocid seals inhabiting polar regions, indirect effects from climate change will likely have the biggest impacts on habitat and resource availability, which is significantly important for ice-dependent species such as the leopard seal [164-166]. Disease too is predicted to increase as a result of changes in climate, and although rare in marine mammals, outbreaks have led to catastrophic declines in some species of phocid seals [98,167-170]. Colonising, social species such as southern elephant seals will be particularly vulnerable to these threats [169,171]. The ability to understand and manage the risk associated with disease in wild mammals can be greatly improved by understanding their relationship with microbial communities and advancing knowledge of the factors, which regulate the ecology of these hosts, such as the gut microbiota.

1.3.6 Current understanding of the gut microbiota of phocid seals

Current knowledge of bacteria associated with phocid seals is limited. Studies, to date, have targeted clinically relevant zoonotic organisms (organisms which can be transmitted between humans and wildlife) [172,173] or been focused on particular 12

groups of bacteria implicated in disease outbreaks in populations of wild seals, specifically species from the genera Brucella, Leptospira, Bartonella, Mycobacterium (tuberculosis) and Escherichia [174-180]. Knowledge of the commensal bacteria in marine mammals would provide a baseline against which future changes can be compared. To date, Glad et al. [44] have completed the most comprehensive study of the gut microbiota associated with phocid seals. In their study, they used 16S rRNA gene extracted from DNA to characterise the bacterial community from culled wild hooded seals, Cystophora cristata (number of individuals analysed, n = 9), harbour seals , Phoca vitulina (n = 1), and grey seals, Halichoerus grypus (n = 1), inhabiting the Arctic. The gut microbiota were found to be abundant and diverse, representing four bacterial phyla in similar abundances to recent studies investigating terrestrial mammals [44].

1.4 The Antarctic environment

The Antarctic continent is located south of the 60° latitude and together with the Southern Ocean covers 50 million km2 or about 10% of the Earth’s surface (Figure 1.4) [181,182]. Continental Antarctica is generally described as the coldest and windiest place on Earth. The surface is covered almost entirely with ice (97%), averaging over 2,000 m thick. These ice sheets hold ~90% of the Earth’s fresh water and their thermal mass is significant for global climate [183,184]. Sea ice also forms across vast regions of the Southern Ocean ranging from 3 million km2 in summer to 18 million km2 in winter [185]. Sea surface temperatures remain stable year-round at -1.9°C and air temperatures commonly remain below zero, with an extreme minimum of -89.2°C recorded in 1983 [181]. The dramatic seasonal changes which exists in the Antarctic and Southern Ocean Region are best suited to species which are mobile, therefore plant life is minimal and animal life is sea or airborne and migratory [186]. Compared with temperate ecosystems, the number of animal species is few but they are abundant and many of them are endemic [181,186,187].

Resource availability in the Southern Ocean is strongly seasonal and related to the cycle of sea ice. Life history traits of the dominant fauna, which includes marine mammals, penguins and various seabirds, are tightly synchronised with this phenomenon. However, in the past 50 years the Antarctic Peninsula has been 13

undergoing one of the most rapid rates of warming on the planet at the rate of 1.09°C per decade [184,188-190]. This is causing significant changes to the properties of the ecosystem including increasing atmospheric temperatures retreating glaciers, decreasing sea ice extent, and decreasing ice season [188,191- 196]. These changes threaten to unravel the current synchronicity impacting all levels of the food web and recent studies have already identified declines in polar species related to decreasing sea ice extent [183,190,197]. For apex predators, such as seals, changes in the abundance and availability of prey is set to have profound implications on their foraging behaviour and survival [198-201]. It is important to have a complete understanding of the ecology of these hosts, including their ability to respond to predicted changes. The lack of knowledge on the gut microbiota of these hosts, essential to energy harvest and healthy survival, reduces the ability to understand their potential to respond to changes and be managed.

The western Antarctic Peninsula is located 1,000 km from the tip of South America; it boasts warmer temperatures and increases in humidity and precipitation compared to the Antarctic continent [189,202]. It is abundant with flora and fauna with large populations of breeding wildlife, including seals [186,189]. Adjacent to the peninsula, along a 510 km south to northwest stretch sits an archipelago of 11 islands known as the South Shetland Islands (Figure 1.5). The largest of these islands is 25 de Mayo / King George Island. Up to 35 scientific stations are present in the region, belonging to 15 nationalities; nine of these occur on 25 de Mayo / King George Island [203,204]. Two sampling sites were chosen off the western Antarctic Peninsula to access populations of southern elephant and leopard seals. Southern elephant seals were located off 25 de Mayo / King George Island (62o14’S, 58o39’W) where they accumulate in large numbers to breed and molt. Leopard seals, which are common throughout the area and occur in greater abundances closer to the peninsula, were located off Cierva Point (64o09’S, 60o57’W), Danco Coast (Figure 1.5).

Figure 1.4 Location of Antarctica. Figure credit: Australian Antarctic Division [205]

Figure 1.5 Location of the western Antarctic Peninsula highlighting sampling sites Figures adapted from Casaux et al. [206] and Peck et al. [181].

1.5 Thesis organisation

The gut microbiota of mammals is an essential part of their ecology. This symbiotic relationship underpins the health and metabolic potential of the host, whilst providing insight into aspects of the host’s evolution and biology. The main objectives of this PhD research were to characterise the gut microbial community of Antarctic marine seals and assess the factors influencing community structure. This aimed to contribute more broadly to the understanding of gut microbiota of wild mammalian hosts with a specific focus on marine mammals. Every attempt was made to write this thesis as a series of stand-alone papers and some repetition occurs. Nelson is a senior author and major contributor of all chapters. References are combined at the end of the thesis for convenience.

Chapter 2 describes the development of sample methodology regarding these particular hosts. It identifies the stability of the gut microbiota through time, and the use of faecal samples as a proxy for the microbial community associated with regions of the gastro-intestinal tract. Chapters 3 and 4 characterise the gut microbiota of wild southern elephant and leopard seals with a comparison to captive leopard seals. Chapter 3 uses the community DNA fingerprinting technique ARISA to assess the faecal microbiota relative to the factors attributed to each host seal. Chapter 4 uses next-generation sequencing to characterise the taxonomy of the faecal microbiota of hosts sub-sampled from Chapter 3. Chapter 4 delves further into patterns identified in Chapter 3. Chapter 5 places the faecal microbiota of Antarctic marine seals into a global context by comparing them to current published literature of terrestrial and marine mammals in a meta-analysis. Chapter 6 examines the potential for transmission of bacteria and antimicrobial resistance (AMR) to occur between human wastes and nearby southern elephant seals surrounding 25 de Mayo / King George Island, Antarctica. An overall discussion of the research generated from this thesis is contained in Chapter 7. Information relating to sample collections from all seal hosts is listed in Appendix 1. Supplementary information for Chapters 2, 3, 4, 5 and 6 are contained in Appendices 2, 3, 4, 5 and 6, respectively.

Chapter 2

Stability of the gut microbiota of captive leopard seals: dietary influence and longitudinal change

2.1 Abstract

Understanding the stability of a community and the primary influences on its composition are necessary to provide a context for large-scale surveys. The microbial community within the gut of mammals is both numerous and diverse. To study aspects of the gut microbiota, methods are required which cause minimal harm to the host mammal. Composition of the gut microbiota is influenced by a variety of sources, most notably, dietary intake. This study aimed to identify the stability of the gut microbiota over time, establish the microbial community associated with sections along the gastro-intestinal tract and faeces and assess the influence of diet on the gut microbiota. The faecal microbiota of two captive leopard seals housed at Taronga Zoo, Sydney, Australia, and their dietary items (fish), were assessed over a three month period using the DNA fingerprinting method, automated ribosomal intergenic spacer analysis (ARISA). Microbial communities associated with the upper and lower intestines were collected opportunistically during necropsy from a leopard seal, which stranded off Sydney and was subsequently euthanased. The gut microbiota of the captive leopard seals remained stable for months at a time with fluctuations observed due to illness and modification of food intake. Regardless of illness in the female and not the male captive leopard seal they both shar ed 76% of operational taxonomic units (OTUs) in their faecal microbiota. The faecal microbiota was identified as being most similar to the lower gastro-intestinal tract and least similar to the upper gastro-intestinal tract in samples from the necropsied leopard seal. This study confirmed the value of faecal samples in understanding the ecology of the gut microbiota of these mammals. The gut microbiota of dietary fish shared an average relative abundance of 34.8 ± 4.9% of OTUs with the leopard seal gut microbiota at any one time. The shared enclosure space and diet of the captive leopard seals is a strong driver of the gut microbiota, more so than their unique responses to colonisation.

2.2 Introduction

To comprehend the structure and function of ecological communities, an understanding of their temporal stability is necessary. Studies in ecology often rely on representative sampling without temporal replication necessary to assess community composition. This is particularly true for host-associated bacterial communities. For the symbiotic gut microbiota of mammals, dietary items may considerably influence community composition, therefore, it is important to understand the stability within a host species and provide a context for future studies.

Studying the ecology of wild mammals is hindered by their relative inaccessibility. Specific to marine mammals, they often live in areas, such as the deep ocean or Polar Regions, which present considerable challenges for scientists. Once located, minimally invasive sampling is required to ensure ethical treatment of the mammal and limit any undue physical stress [102]. Thus, temporal repeat sampling in wild hosts is often either physically or ethically impractical. Therefore, representative sampling or sampling regimes with low replication are often employed and when possible it is important to ground-truth these methods to provide context for future field studies.

Dietary sources provide the gut habitat with the source bacterial community, whilst at the same time, provide sustenance for community members to survive [22]. The composition of dietary items can have a profound effect on gut bacterial community composition. Dietary fibre in the form of apples, for instance, caused an increase in numbers of Bifidobacteria and Lactobacillus resulting in a healthier intestinal environment in humans [207]. Specific bacterial species have also been introduced and sustained with particular dietary items, such as Camembert cheese [208].

It has been shown that faecal material is representative of the bacterial community present in the colon and lower intestines of mammals [13]. Hence, the collection and analysis of faeces provides a low-impact sampling regime to study the gut microbiota of mammals. However, most of our knowledge on the stability and composition of the gut microbiota has come from the study of humans and domesticated animals.

Experiments conducted on these subjects under laboratory conditions and in humans are likely to give different results to those conducted on wild mammals [209]. This is primarily due to the increasing understanding that the processes, which generate natural variability, operate over different scales of space and time [209].

The goal of this study was to gain an accurate understanding of the temporal bacterial composition of the gut microbiota of the leopard seal, Hydrurga leptonyx. The aims were to identify: 1) community dynamics of the faecal microbiota over three months in two individual leopard seals; 2) gut microbiota associated with sections along the intestinal tract and in the faeces; and 3) contribution of bacteria from dietary items to the faecal microbiota.

2.3 Materials and methods

2.3.1 Sample collection from captive leopard seals and fish

Faecal samples were collected opportunistically from two captive leopard seals housed at Taronga Zoo, Sydney, NSW, Australia between March and June 2010 (Table A1.1). The seals, one male and one female, were housed together in the same enclosure, after being rescued from the wild. Based on their morphology and behaviour at the time of rescuing, the seals were estimated to be sub-adults between two to four years old at the time of sampling. Faecal samples were frozen and stored at -20˚C after collection.

Samples along the intestinal tract and the faeces were collected opportunistically from a leopard seal during necropsy. This seal was a male sub-adult who stranded off the south east coast of NSW, Australia in July 2009 and was subsequently euthanased as a result of illness [210]. Samples were collected from the duodenum, jejunum, ileum, colon, rectum and faeces. The necropsy and sample collection was conducted approximately 20 h post-mortem and samples were frozen at -20˚C.

In captivity, leopard seals are fed a variety of fish species, sourced from coastal waters off Sydney and subsequently stored in frozen batches. Specimens of fish species used in this study included: yellowfin whiting (Sillago schomburgkii), southern herring, (Herklotsichthys castelnaui) and slimy mackerel, (Scomber 20

australasicus). Each fish within a batch are assumed to possess a microbiota, which is representative of every fish in that batch. Therefore, fish sampled prior to consumption provide an indication of the microbiota being transferred into the gut of the consumer, in this case, the leopard seal. At two points a sample of each fish species from their current batch was collected along with a faecal sample from each leopard seal (Table A1.1). Skin, muscle tissue and samples from the gut cavity of each fish were collected and frozen at -20˚C. Faeces was collected and treated as outlined above.

2.3.2 Genomic DNA extraction and community fingerprinting using ARISA

Total genomic DNA was extracted from faecal, intestinal and fish samples using QIAamp Stool DNA Mini Kit (Qiagen Pty Ltd., VIC, Australia) following the manufacturers protocol. DNA was extracted from 26 faecal and gastro-intestinal tract samples, including 10 from the male captive leopard seal, 10 from the female captive leopard seal and six from faeces and multiple sites along the gastro-intestinal tract from the necropsied male leopard seal. DNA was extracted from the skin, muscle and gut cavity of six fish. Only two quality samples from the gut cavity of two fish species, including one from yellowfin whiting and one from southern herring were successfully extracted (Table A1.1). These were analysed in conjunction with DNA extracted from two faecal samples from the male captive leopard seals and two faecal samples from the female captive leopard seal (Table A1.1).

ARISA [64] was performed on all DNA samples to assess microbial community composition. ARISA utilises length heterogeneity of the intergenic transcribed spacer (ITS) region between the bacterial 16S and 23S rRNA genes, a hyper-variable region that varies among species and among strains of bacteria [64]. This method fails to detect bacteria that lack a linked operon, including some Archaea and Planctomycetes [76]. However, Planctomycetes comprised only 0.08% of organisms previously identified from a wide range of mammalian gut samples using other methods [23]. In the same study Archaea were not detected at all within a range of terrestrial mammals [23] and were discovered to be low in wild Arctic and sub-Arctic seals [44]. Phylogenetic resolution is ≥ 98% 16S rRNA gene sequence identity, near the range widely considered to be bacterial ‘species’ or ‘phylotypes’ [74]. 21

Polymerase chain reaction (PCR) amplification of the ITS region was performed using the forward primer 1392F 5'-TGYACACACCGCCCGT-3' and the HEX-labelled reverse primer 23SR 5'-GGGTTBCCCCATTCRG-3' (Sigma-Aldrich Pty. Ltd, NSW) [64]. Reactions contained 12.5 l Mastermix (Promega, NSW), 0.5 l of each primer (20 pmol/l), 10.5 l of water and ~5 ng of genomic DNA. The reaction mixture was held at 94˚C for 10 min, followed by 30 cycles of amplification at 94˚C for 1 min, 54˚C for 1 min and 72˚C for 2 min with a final step of 72˚C for 5 min using a Mastercycler ep Realplex (Eppendorf South Pacific Pty Ltd., NSW). Fragment length analysis was carried out at the Ramaciotti Centre (University of NSW, NSW) using an Applied Biosystems 3730 DNA Analyser (Applied Biosystems, CA, USA) with a GS1200 LIZ internal size standard. Peak size and area data were extracted using GeneMapper software v 3.7 (Applied Biosystems). Given the approximate known lengths of the ITS regions included in the primer set, fragment lengths below 350 bp and above 1200 bp were eliminated from analyses.

Final peak thresholds were determined using the online analysis tool T-REX [211]. True peaks were defined from background noise as those with heights exceeding twice the standard deviation computed over all peaks within a sample. Data comprising ‘true’ peak sizes and peak areas were converted to abundance per binned operational taxonomic units (OTUs) using the custom binning script ‘interactive binner’ [212] in the R software package [213], with a relative fluorescence intensity cut-off of 0.09%, a window size of two and a shift size of 0.1.

2.3.3 Data analyses

Fourth-root-transformed abundance data were used to generate a resemblance matrix using the Bray-Curtis similarity algorithm [214]. Similarities between sample groups were visualised using non-metric multi-dimensional scaling (nMDS) [215]. The result of nMDS ordination is a map where the position of each sample is determined by its distance from all other points in the analysis. To test for differences in composition of the faecal microbiota between captive leopard seals through time a two factor permutational multivariate analysis of variance (PERMANOVA) [216] was carried out based on the similarity of ARISA profiles. Each test was done using 9,999 permutations under Type III sum of squares (SS) and a reduced model to generate a 22

permutated F statistic (F) and p-value (P). In the case of a significant interaction term and where the number of factors was greater than three, pair-wise a posteriori tests for all combinations of factors were conducted using the t-statistic. Results were considered significant where p-value = < 0.025. Homogeneous dispersion of samples within each factor was assessed by a test of multivariate dispersion (PERMDISP) [217]. In cases where a positive PERMDISP was identified indicating heterogeneous dispersion amongst groups, the nMDS was examined to observe separation of groups and in addition analysis of similarity (ANOSIM) was performed. ANOSIM is a non-parametric permutation procedure, which is applied to the similarity matrix and is more robust to heterogeneous dispersion. Only results pertaining to the PERMANOVA are reported where significance is confirmed with the use of ANOSIM.

The contribution of OTUs to the average dissimilarity between groups and, therefore characteristic of community structure was calculated using a similarity percentages procedure (SIMPER). SIMPER identifies which OTUs account for observed differences in microbiota between individuals. SIMPER also provides a within-group similarity based on OTU abundances in each individual within a group. OTUs were defined as ‘shared’ when present in more than half (most) of the samples being tested within a group. OTUs were defined as ‘unique’ when only present in one sample or group of samples compared with another sample or group of samples. All statistical tests were performed using the software PRIMER-E v6 [218].

2.4 Results

2.4.1 Stability of the gut microbiota

A total of 193 operational taxonomic units (OTUs) were found across all 20 samples. No significant differences were observed between the microbiota obtained from the two seals (F = 1.3512, PPERM = 0.179; Table 2.1). However, their microbiota did change significantly when samples from the two seals were grouped together by month (March, April and May), most notably between March and May (F = 1.6477,

PPERM = 0.0016), but also between March and April (F = 1.4764, PPERM = 0.0167) and

April and May (F = 1.4208, PPERM = 0.0214; Table 2.1). Samples collected during

April had the highest similarities for each individual and when combined (Table 2.2). Overall, a gradual and similar change in community was observed for each individual through time (Figure 2.1).

Characteristic OTUs in the faecal microbiota of captive leopard seals were identified using SIMPER analysis (Table A2.1). Fluctuations in the relative abundance of characteristic OTUs within an individual was ~6% or less per week (Figure 2.2). On occasion, during one week, such as OTU 471.8 during the period 20-27 April 2010 for the female captive leopard seal, the relative abundance fluctuated from 10% to 35% (Figure 2.2A). Fluctuations in abundance of characteristic OTUs greater than 15% were more common for the female captive leopard seal than the male (Figure 2.2).

Table 2.1 PERMANOVA on the gut microbiota of male and female captive leopard seals over time. PERMANOVA of gut microbial abundance data to generate a permutated F statistic (F) and permutated p-value (P) with calculated degrees of freedom (d.f.) and sums of squares (SS) noted. Pair-wise a posteriori tests between age-classes were conducted using the t-statistic (t). Significance level: ***P = ≤ 0.001, **P = 0.01, *P = 0.025. Source of d.f. SS F P Variation Individual 1 1572 1.3512 0.179 Month 2 2723 2.3399 0.0014** Individual x 2 1179 1.0131 0.4575 month Pair-wise comparison t P March, April 1.4764 0.0167* March, May 1.6477 0.0016** April, May 1.4208 0.0214*

The faecal microbiota of the male and female captive leopard seals possessed an almost identical within-group similarity as identified using SIMPER with 50.5% and 47.6%, respectively (Table 2.2). On average, the relative abundance of shared OTUs (OTUs identified in both individuals at any one time) in the faecal microbiota of the male was 73.7 ± 5.1% and 77.2 ± 4.2% in the faecal microbiota of the female (Figure 2.3). The relative abundance of shared OTUs decreased markedly on 27th March 2010 for each seal, especially for the male (Figure 2.3). During this sampling period the relative abundance of unique OTUs (OTUs occurring only in the male leopard seal and not the female) is 59.6%, which was far greater than the average of 26.3 ± 24

5.1% (Figure 2.3). Similarly the relative abundance of unique OTUs in the female’s faecal microbiota increased to 42.8% on the 26-May 2010, which was greater than the average relative abundance of 22.8 ± 4.1% (Figure 2.3).

Table 2.2 Similarity of the gut microbiota of captive leopard seals over time. Within-group similarity is based on the faecal microbiota of samples within a defined group using SIMPER analysis. Number of individuals analysed (n) within each group is noted. Captive Within-group Groupings n leopard seal similarity (%) Male 10 50.5 Male March 5 49.8 Male April 3 60.3 Male May 2 54.6 Female 10 47.6 Female March 5 51.4 Female April 3 55.6 Female May 2 43.0 March 10 50.2 Male and April 6 59.1 female May 4 46.4

Figure 2.1 Similarity of female and male leopard seal gut microbiota over time. nMDS ordination plot dislaying similarity of the faecal microbiota of each male and female captive leopard seal over time.

Figure 2.2 Relative abundance of characteristic OTUs in the gut microbiota of leopard seals through time. Relative abundance of characteristic operational taxonomic units (OTUs) in the gut microbiota of the male and female leopard seal on the same sampling date (Date). Characteristic OTUs are identified using SIMPER and are shown as follows: (A) OTU 471.8; (B) OTU 473.8; (C) OTU 685.8; and (D) OTU 709.8. A full list of characteristic OTUs for each individual is listed in Table A2.1.

Figure 2.3 Relative abundance of shared and unique OTUs in the gut microbiota of captive leopard seals. Relative abundance of shared and unique operational taxonomic units (OTUs) in the gut microbiota of the female and male leopard seal at each sampling date.

2.4.2 The microbiota associated with the intestinal tract and the faeces

A total of 91 OTUs were identified along the intestinal tract and in the faeces of a necropsied leopard seal. The bacterial community present in the upper parts of the tract differed substantially from the faecal microbiota (Figure 2.4). However, little difference was observed between the microbiota of the lower intestines and the faeces (Figure 2.4).

The relative abundance of some characteristic OTUs, such as OTU 397.2 and OTU 399.2, fluctuated by up to 35% in the small intestines (duodenum, jejunum and ileum) compared with the large intestines (colon and faeces) and the faeces (Figure 2.5). Characteristic OTUs are shared between adjacent sections of the intestinal tract, such as OTU 857.2 in the colon, rectum and faeces (Figure 2.4).

Figure 2.4 Similarity of the microbiota of intestinal sections and faeces. nMDS ordination plot based on the similarity of the bacterial communities collected from the leopard seal small intestines (duodenum, jejunum and ileum), large intestines (colon and rectum) and the faeces.

Figure 2.5 Relative abundance of characteristic OTUs along the intestinal tract and in the faeces. Relative abundance of the foremost characteristic operational taxonomic units (OTUs) identified using SIMPER analysis from the intestinal tract and faeces of a necropsied leopard seal. 2.4.3 Transfer of gut microbiota from fish to seal

Fish samples from the gut cavity of two prey species and faecal samples from each captive leopard seal yielded 175 OTUs. Both of the fish samples possessed OTUs, which were present in the faecal microbiota of each captive leopard seal (Figure 2.6). Many of the OTUs found in the leopard seals were not found in the gut cavity of

either fish and vice versa (Figure 2.6). The relative abundance of shared OTUs between fish and seals was higher for the male captive leopard seal with 40.0 ± 3.3% than for the female captive leopard seal with 27.6 ± 6.4% (Figure 2.6). OTUs from the whiting made up a greater proportion of the shared OTUs observed in seal faecal microbiota compared with OTUs from the herring (Figure 2.6).

Figure 2.6 Shared OTUs between leopard seals and their fish diet. Shared operational taxonomic units (OTUs) from the microbial community of the faeces of female and male leopard seals and of the gut cavity of representatives of their fish diet: southern herring (Herring), and yellowfin whiting (Whiting). Fish images credit: www.fishbase.org.

2.5 Discussion

2.5.1 The gut microbiota of captive leopard seals is stable over time

This is the first study that provides an analysis of the temporal stability of the faecal microbiota of leopard seals. The community was identified to be relatively stable over the three-month sampling period. In humans, the faecal microbiota has been identified to remain very stable over prolonged periods of time after two years of age [15,219]. Dissimilarity and differences in the faecal microbiota noted within individuals across the months of the present study were a result of changes in the relative abundances of OTUs, yet these were rarely greater than 10%. Microbial communities in the gut of mammals are known to respond to alterations in diet, such as the introduction of fibre or food types, which alter the physicochemical and nutrient properties in the intestines [35,220]. In this case, the physicochemical and nutrient properties of the leopard seals’ intestines, although not tested, can be 29

assumed to remain relatively constant. In captivity, the leopard seals experience regular and consistent activity within their enclosure and receive a steady flow of dietary items. The assumption is that the nutrient and physicochemical properties of the leopard seals’ intestines remains constant resulting in minimal fluxes in gut microbial composition.

Characteristic OTUs that make up the faecal microbiota of captive leopard seals remained present throughout the sampling period and only their relative abundance changed between sampling times. This pattern has been observed in human twins [12]. Causes for these shifts in abundance of bacteria in the gut are numerous. Influences of diet type and composition, antibiotics, illness and other lifestyle factors have been demonstrated in several studies [10,28,221]. The female leopard seal displayed a reduced within-group similarity for the months April and May and overall compared with the male captive leopard. The increased fluctuations in characteristic OTUs in the female captive leopard seal more so than the male captive leopard seal may have been a result of illness. In February 2010, the female captive leopard seal endured significant weight loss due to reduced eating and at one point was administered with an antibiotic treatment (Table A1.2). Fasting, altered eating and antibiotic treatment have been shown to greatly reduce abundances of bacterial species in the human intestinal community [28,222,223]. However, regardless of these influences, the faecal microbiota of the female captive leopard seal closely resembled the male captive leopard seal’s, even though the male was not undergoing weight loss or treatment with antibiotics.

Changes in the relative abundance of OTUs in the gut microbiota of the male and female captive leopard seals were consistent within and between individuals for each month. The high abundance of shared OTUs (73.7 ± 3.2%) in the faecal microbiota at each sampling time is common amongst social mammals and family members [12,224]. In adults, transmission occurs as a result of social interactions, mating, dietary exposures and other behaviours [225]. We would expect some general community similarity in these con-specific seals, even though they are not related in a population-ecology sense. In the captive enclosure of the zoo, the leopard seals share a space of approximately 500 m3, consume a similar diet and are exposed to

the same external environment. Previously, environmental effects caused by a shared space have been shown to increase the likelihood of sharing bacterial community members between hosts [15,226]. For example, in the case of blue (Parus caeruleus) and great tit (P. major) nestlings, the nest in which the individuals were raised had a greater influence on regulating nestling gut community structure than the nest of first origin or nestling genotype [226].

2.5.2 The gut microbiota represents the lower intestinal microbiota

Faecal material is an opportunistic sample capable of providing information about the intestinal community of mammals. Comparisons of the microbial community in the small intestines and the large intestines to the faeces are difficult to measure due to the nature of obtaining adequate, coinciding samples from these regions. Several methods have been tested [e.g. 227,228]; yet the difficulty involved in implementing these methods means there is a continued reliance on samples obtained post- mortem [229,230]. In this study, samples were obtained opportunistically from a leopard seal during necropsy. This particular leopard seal stranded off the south east coast of NSW, Australia in July 2009 and was subsequently euthanased due to illness. Leopard seals are known to venture north during winter in search of food and on occasion sub-adults, in particular, will venture as far north as the east coast of NSW [142]. When the seals occur this far north, they are often ill, potentially due to their inexperience in catching prey [142]. Prior to euthanasia, this seal was observed to be exceptionally thin, behaving abnormally, and possessed visible external wounds [210]. This provided a unique opportunity to collect samples and compare microbial communities along the intestinal tract and in the faeces.

The faecal microbiota of the leopard seal sampled in the present study was found to be more similar to the microbiota from sections of the large intestines (colon and rectum) than to the small intestines (duodenum, jejunum and ileum). OTUs were shared more readily with adjacent intestinal areas; therefore the faeces shared more OTUs with the large intestine than the small intestine. The gastro-intestinal tract of mammals extends from the mouth to the anus. In human studies, bacterial community members are commonly shared between adjacent sections of the intestinal tract, highlighting the translocation of upstream 'source' community 31

members to downstream 'sink' communities [229,231,232]. However, bacterial species are known to be specifically associated with particular regions, such as numerous species of the phylum Bacteroidetes occurring specifically in the large intestines [13].

The increase in shared OTUs between the colon, rectum and faeces suggests sampling faecal material provides an indication of the bacterial community present in the large intestines of these hosts. Physicochemical and nutrient properties vary along the gastro-intestinal tract [233], as does the relative absorption of fats, protein, glucose and water [233]. This, in turn, influences the abundance of individual bacterial types and the composition of the gut microbiota [234]. Oxygen availability decreases from the small to the large intestines and the bacterial community shifts from facultative anaerobes (bacteria that grow without oxygen but can use it if it is present) to obligate anaerobes (bacteria that cannot grow in the presence of oxygen). Obligate anaerobic bacteria have been found to be up to 100 times lower in the upper gut compared with the faeces [228,235]. The reduction in the relative abundance of shared OTUs in the small intestines and the faeces are likely due, in part, to differences in oxygen availability and other physicochemical factors.

2.5.3 Diet introduces bacteria into the gut

Dietary items contribute greatly to the composition of the gut microbiota. They serve as the primary mode of colonisation for bacteria into the gut and their physicochemical composition of becomes the substrate for bacteria inhabiting the gut [236]. This study confirmed that dietary items, in particular prey items with their own associated gut microbiota, are a dominant source of bacterial community members into the gut of the host mammal. The introduction of bacterial species from particular dietary products has been identified previously [208]. Indeed, the transmission of necessary bacterial members, such as cellulose-degrading bacteria, into the gut of infant herbivorous animals is considered essential to their future ability to harvest energy from their plant-based diet [237,238]. However, few studies have identified this transfer from prey items. Carnivorous hosts may also be reliant on the introduction of bacteria from their prey. These results suggest that the gut microbiota of a prey item will be a window into the gut microbiota of the predator. 32

2.6 Conclusions

Rarely do we have the opportunity to collect time-series data from the guts of wild marine mammals. As a result, the temporal data obtained from captive leopard seals can provide important contextual information on the structure and dynamics of the faecal microbiota. In this study, captive leopard seals maintained a stable gut microbiota through time, which was influenced by diet and environmental conditions. Diet was identified as a primary mode of transmission of bacterial community members to hosts, specifically from the gut of prey items. The study demonstrated the value of faecal material as a useful tool for representing the bacterial community of the large intestine. The study has important implications for future field-based research projects.

Chapter 3

Patterns influencing the gut microbiota of Antarctic seals

3.1 Abstract

The gut microbiota of mammals is acquired initially during the birth process. As the infant mammal ages, the gut microbiota becomes increasingly diverse following dietary change from mother’s milk to solid food. Functionally, the gut microbiota extracts energy from the diet and regulates the immune system suggesting it is an essential component of host survival. Research into this symbiosis has been directed towards factors, which influence the composition of this microbial community. These factors are thought to include, diet, genotype, environment, health, weight, gut morphology and other lifestyle factors. While they are becoming increasingly well understood, there has been a reliance on human and laboratory-reared or captive mammals. In natural systems, which are inherently more variable, fundamental questions regarding these essential communities have not been identified. This study aimed to assess the gut microbiota of two species of marine mammals, the southern elephant seal (n = 41) and the leopard seal (n = 21), living in their natural ecosystem of Antarctica. Two captive leopard seals housed at Taronga Zoo, Sydney, Australia, were also analysed for comparison. The fingerprinting technique, ARISA, was used to identify the bacterial community in DNA extracted from faecal samples. Composition of the faecal microbiota of wild leopard seals was significantly different to southern elephant and captive leopard seals. Captive leopard and southern elephant seals did not display dissimilar faecal microbiota. Amongst southern elephant seals, adults and pups and also mothers and pups displayed significant differences in their faecal microbiota. Only between adult southern elephant seals were significant differences in the faecal microbiota observed due to sex. These results suggest that the consumption of different prey and dietary items between these host species, in the wild and in captivity, contributes to the composition of their faecal microbiota.

3.2 Introduction

Bacteria are symbiotically associated with the gut of mammals. In this habitat, they perform numerous beneficial functions for the host, such as harvesting energy, regulating immunity and aiding cellular maturation [3,10,239]. The gut microbiota first originates during passage through the birth canal. Subsequent feeding through lactation and dietary items and influences from the surrounding environment impacts the response of the infant gut to colonisation [15]. As the mammal ages, the community grows up to ~500 species [15,17].

Recent investigations into the gut microbiota of mammals have identified numerous factors, which influence the unique and variable community present in each individual. These include diet composition and type, host genotype, gut morphology and physiology, social interactions, exercise, health, host mass and antibiotic use [12,21,23]. Understanding these influences have come from studies on humans, laboratory-reared, agricultural and captive animals [e.g. 14,34,35,240,241]. Natural systems, however, are inherently more variable than these experimental environments and this is likely to result in different influences impacting colonisation to the gut or host responses, which are not adequately replicated in captivity, such as evolved social interactions or natural dietary items. Therefore, the current knowledge of factors influencing the gut microbiota may not adequately represent mammals inhabiting natural systems.

One of the most understudied groups of mammals living in natural systems are marine mammals. Few studies have looked at the gut microbiota of wild mammals [e.g. 43,242] and fewer have investigated marine mammals [i.e. 44,45]. This deficiency in is due, primarily, to the difficulty in obtaining adequate samples from wild mammals and, particularly marine mammals. Wild marine mammals represent an exceptional group of mammals as they have traits that are adapted to an aquatic or semi-aquatic lifestyle. This may have implications for the composition of the gut microbiota. For instance, the gastro-intestinal tract of marine mammals is comparatively longer than terrestrial mammals, which would impact the physicochemical properties, such as oxygen availability, of the habitat [243]. They also have unusual metabolic adaptability, in some cases, they possess elevated metabolic 35

rates due to their large size yet are able to reduce this considerably during foraging dives [112,244].

In general, comparisons of the gut microbiota of mammals inhabiting wild and captive environments have been modestly studied. Ley et al. [23] in their study on numerous terrestrial mammals, included some from captive and wild environments, and concluded that con-specific hosts were generally more similar to each other than to non-con-specific hosts. However, published studies, which have investigated these differences at a finer scale, have observed considerable differences between wild and captive individuals. To date, these studies have been limited in mammals to primate species [42,43,245] with several other studies focusing on birds [39-41] and fish [246].

The aim of this study was to investigate factors influencing the gut microbiota of wild marine mammals. Faecal samples were collected from two species of wild marine mammals, the southern elephant seal and the leopard seal, inhabiting Antarctica were analysed. These were compared to faecal samples from leopard seals inhabiting Taronga Zoo, Sydney, NSW, Australia. Faecal samples were previously identified as representative of the lower gastro-intestinal tract in leopard seals providing a less-invasive method of sample collection in wild hosts (Chapter 2). The community fingerprinting technique, ARISA, was used on DNA extracted from faecal samples. The aims were to: 1) compare the gut microbiota between southern elephant seals and leopard seals living under natural conditions, leopard seals living under natural conditions and in captivity, hosts of different age-classes and hosts of different sexes; and, 2) determine the influence of age-class and maternal transmission on the composition of the gut microbiota of southern elephant seals.

3.3 Materials and methods

3.3.1 Sample collection from Antarctic seals

Samples were collected from southern elephant and leopard seal populations in the western Antarctic between September 2008 and March 2009. Southern elephant seals were sedated using the protocol described by Carlini et al. [247] on 25 de Mayo / King George Island (62o14’S, 58o39’W), South Shetland Islands (Figure 1.5). 36

Leopard seals were sedated using the protocol described by Higgins et al. [248] on ice floes off Cierva Point (64o09’S, 60o57’W), Danco Coast, western Antarctic Peninsula (Figure 1.5). Seals were sampled within a ~5 km2 study area in each geographic region.

Faecal samples were collected from sedated seals by rectal swabbing using sterile swabs (Oxoid Australia Pty Ltd, SA, Australia), which were frozen and then stored at -20˚C. While the microbial community of mammals is known to vary along the length of the digestive system and in faeces [Chapter 2, 228,235], faecal material is generally indicative of the lower intestinal tract microbiota where bacterial biomass is greatest [13,235] and which was confirmed for leopard seals (Chapter 2).

At the time of sample collection, sex, weight, age-class, standard length (nose to tail) and axiliary girth were noted for each seal. Three distinct age-classes of southern elephant seals were estimated based on knowledge of the morphology and behaviour of an individual, which changes as the seal ages. Host age-classes of southern elephant seals were classified as follows: female adult (> 4 y), male adult (> 6 y), female sub-adult (1-4 y), male sub-adult (1-6 y), and pup (< 3 wks) (Table A1.1). Wild leopard seal adults only were sampled. A portion of southern elephant seals were weighed to obtain their mass using the method described in Carlini et al. [249]. Briefly, this involved suspending sedated seals in a net stretcher from a load cell hanging scale attached to an aluminium tripod. The mass of non-weighed southern elephant seals was estimated using the standard length and axiliary girth of the individual using model six and nine as described in Bell and Hindell [250]. In total, rectal swabs were collected from 41 southern elephant seals (five male adults, 10 female adults, 13 male sub-adults, three female sub-adults, eight male pups and two female pups) and 21 leopard seals (14 male adults and seven female adults).

Repeat faecal samples (n = 3) were collected from two captive leopard seals, one male and one female sub-adult, housed at Taronga Zoo (Table A1.1). During the sampling period, it was ascertained that the female captive leopard seal was ill, undergoing weight loss and requiring the administration of antibiotics (Table A1.2). Results are reported here, with and without the inclusion of affected samples.

3.3.2 Genomic DNA extraction and community fingerprinting using ARISA

Total genomic DNA was extracted and ARISA [64] was performed on 68 rectal swab and faecal samples (see Chapter 2 for details).

3.3.3 Data analyses

Square-root-transformed abundance data were used to generate a resemblance matrix and similarities between sample groups were visualised using nMDS as detailed in Chapter 2. Spearman’s rank correlation was used to identify which OTUs correlated strongly with gut microbial compositions and vectors were overlaid onto nMDS plots. In addition the OTUs characteristic of community structure were calculated using SIMPER as detailed in Chapter 2. To test for differences in composition of the faecal microbiota between seals based on a priori groupings of species, sex and age-class a one factor Permutational multivariate analysis of variance (PERMANOVA) [216] was carried out based on the similarity of ARISA profiles. The nature of sample collection did not provide adequate replication for the comparison of groups in a two factor PERMANOVA, such as sex and age-class. For some terms in the analysis, there were not enough permutable units to get a reasonable test by permutation, so a p-value was obtained using a Monte Carlo random sample from the asymptomatic permutation distribution [251]. Results were considered significant where p-value = < 0.025. Statistical tests were performed as detailed in Chapter 2 using the software PRIMER-E v6 [218].

The abundance of OTUs present within a group, e.g. southern elephant seals, were identified as a majority when present in more than half (most) of the individuals within that group. Significant differences in phyla abundances between groups were tested using a Student’s T-Test in Excel 2010 (Microsoft Pty Ltd). Linear regression was used to examine the relationship between host weight and OTU abundance and richness in faecal microbiota.

3.4 Results

3.4.1 Influence of host species and captivity on the gut microbiota

A total of 361 OTUs were identified in the faecal microbiota of seal hosts. Host species were significantly separated based on the composition of their faecal microbiota (F = 4.6538, PPERM = 0.0001; Table 3.1). Pair-wise testing identified dissimilarity between southern elephant seals and leopard seals (t = 2.0744, PPERM =

0.0001), wild leopard seals and captive leopard seals (t = 2.341, PPERM = 0.0001), and southern elephant seals and captive leopard seals (t = 2.1644, PPERM = 0.0001; Table 3.1).

Table 3.1 PERMANOVA on the gut microbiota of hosts based on groupings of species, location, age-class and sex. PERMANOVA of gut microbial abundance data to generate a permutated F statistic (F) and permutated p-value (P) with calculated degrees of freedom (d.f.) and sums of squares (SS) noted. P-values given in italics were obtained using Monte Carlo samples from the asymptotic permutation distribution. Pair-wise a posteriori tests between age-classes were conducted using the t-statistic (t). Significance level: ***P = ≤ 0.001, **P = 0.01, *P = 0.025. Source of Data d.f. SS F P variation Southern elephant seals, Species wild leopard seals, 2 28912 4.6538 0.0001*** and location captive leopard seals Pair-wise comparison t P Southern elephant seals, wild leopard seals 2.0744 0.0001*** Southern elephant seals, captive leopard seals 2.1644 0.0001*** Wild leopard seals, captive leopard seals 2.341 0.0001*** Southern elephant seals Age 2 19360 3.4439 0.0001*** Pair-wise comparison t P Adult, sub-adult 1.5097 0.0004*** Sub-adult, pup 1.923 0.0001*** Adult, pup 2.1841 0.0001*** Southern elephant seal Age 1 10231 4.1868 0.0032** mothers and pups Southern elephant seal Sex 1 6183.2 2.4441 0.004** adults Wild leopard seals Sex 1 4374 1.3353 0.2209 Captive leopard seals Individual 1 2372.4 1.3748 0.2889

Spearman’s correlation coefficient identified OTU 567.3 as correlating strongly with similarity of the faecal microbiota of southern elephant seals (Figure 3.1A). OTU 751.3 and 479.3 correlated strongly with similarity of faecal microbiota for captive

leopard seals (Figure 3.1B). These OTUs were identified as characteristic members of host species by SIMPER analysis (Table A3.1). OTU 629.3 was shared across most individuals in all each host species groups (i.e. southern elephant seals, wild leopard seals and captive leopard seals) contributing an average relative abundance of 4.3 ± 1.1% per individual (Figure 3.1A). This was the only OTU shared between wild and captive leopard seals. Southern elephant seals and wild leopard seals shared an additional seven OTUs (Figure 3.2A).

Figure 3.1 Similarity of the gut microbiota of seal host species groups. nMDS ordination plot dislaying similarity of the faecal microbiota of each individual of groups of (A) southern elephant seals and wild leopard seals and (B) wild leopard seals and captive leopard seals. OTUs identified by Spearman’s correlation coefficient with an influence on community similarity at the level of 0.5 are displayed. 40

The faecal microbiota of captive leopard seals showed the highest within-group similarity of 40% as identified using SIMPER analysis (Table 3.2). Wild seals displayed less within group similarity in their faecal microbiota (Table 3.2). Captive leopard seals possessed a greater abundance of shared OTUs shared with an average relative abundance of 52.6 ± 2.5% in each seal at most of the sample times (Figure 3.2). Southern elephant seals and wild leopard seals shared a similar average relative abundance of OTUs in most individuals (Figure 3.2). Within-group similarity of the faecal microbiota between the captive leopard seals increased with the removal of the sampling time when the captive female was ill (Table 3.2). During illness, the faecal microbiota of the female captive leopard seal was dissimilar to the other two time points and characteristic OTUs shifted substantially with an increased abundance of OTU 567.3 (Figure 3.3).

Table 3.2 Similarity of the gut microbiota of hosts based on groupings of species, age-class and sex. Within-group similarity is based on the gut microbiota of samples within a defined group using SIMPER analysis. Replicate number of individuals analyses (n) within each group is noted. Samples from captive leopard seals (n = 2) were collected at three times (x 3). Within- group Species group Groupings n similarity (%) Southern elephant seals 41 21 Adults 15 26 Female adults 10 29 Male adults 5 31 Sub-adults 16 22 Pups 10 31 Mother-pup pairs 9 17 Mothers 9 28 Pups 9 30 Wild leopard seals 21 19 Females 7 20 Males 14 18 Captive leopard seals 6 40 Female 1 x 3 36 Male 1 x 3 51 Female captive leopard seal (sampling during time of 5 45 illness omitted) Female 1 x 2 63

Figure 3.2 Shared OTUs between the gut microbiota of individuals within seal host species groups. Bar charts display the average relative abundance of shared operational taxonomic units (OTUs) ± standard error (SE) in most (more than half) individuals with host species groups.

3.4.2 Influence of age-class and maternal transfer on the gut microbiota of southern elephant seals

The faecal microbiota of southern elephant seals from three age-classes were significantly different (F = 3.4439, PPERM = 0.0001; Table 3.1). Pair-wise tests revealed significant differences in the faecal microbiota between each of the age- classes (Table 3.1). The faecal microbiota of milk-drinking pups differed significantly from mothers (F = 4.1868, PPERM = 0.0032; Table 3.1). Within each of the nine mother-pup pairs, similarity of the faecal microbiota ranged from 0 to 28% as identified by SIMPER analysis. The relative abundance of OTUs shared between mother-pup pairs ranged from contributing 0 to 83.9% in the pup’s faecal microbiota (Figure 3.4). On average, each mother-pup pair shared 10 OTUs (Table A3.2). Five OTUs (399.3, 629.3, 641.3, 655.3 and 721.3) were shared in four of the mother-pup pairs and one OTU (523.3) was shared in seven of the pairs (Table A3.2). A total of 57.9 ± 5.0% of OTUs were shared within pups, which was greater than any other age-class of southern elephant seals or other host species group (Figure 3.2). The

OTUs that were characteristic of the faecal microbiota of southern elephant seal pups, e.g. OTU 629.3, were present in lower abundances in southern elephant seals adults and sub-adults (Table A3.1).

Figure 3.3 Stability and similarity of the gut microbiota of captive leopard seals. Figure (A) displays nMDS ordination plot based on the similarity of the gut microbiota of the male and female captive leopard seals at each of the sampling times. Black dashed lines represent similarity of those samples at the level of 45%. Pie charts display the abundance of characteristic OTUs in the faecal microbiota of: (B) the male captive leopard seal during sampling times: 71, 77 and 78; (C) the female captive leopard seal during samplign times: 72 and 75; and, (D) the female captive leopard seal during illness, sampling time 73. 43

3.4.3 Influence of sex on the gut microbiota

Sex influenced the faecal microbiota of southern elephant seal adults (F = 2.4441,

PPERM = 0.004), but not wild leopard seals or captive leopard seals (Table 3.1; Figure 3.5). Strongly unbalanced sex ratios amongst southern elephant seal sub-adults and pups did not allow for significance testing. As adult southern elephant seals display sexual dimorphism (males are ~3,000 kg and females are ~600 kg) the association of host mass with OTUs identified as characteristic of the adult female and adult male faecal microbiota was explored further using linear regression (Figure A3.3). Linear regression yielded no significant result with host mass (Figure A3.3).

Figure 3.4 Similarity and shared OTUs in the gut microbiota of mother-pup pairs. Figure (A) displays nMDS ordination plot based on similarity of the faecal microbiota of each individual mother and milk-drinking pup. Figure (B) displays the relative abundance of shared OTUs between each mother-pup pair as indicated by lower case letters.

Figure 3.5 Similarity of the gut microbiota of seal host species based on sex. nMDS ordination plots based on similarity of gut microbiota of (A) wild leopard seal females and males; and, (B) southern elephant seal adult females and males.

3.5 Discussion

3.5.1 Gut morphology, physiology and diet shape the gut microbiota of wild southern elephant seals and leopard seals

This study identified considerable differences in the gut microbiota of southern elephant seals and leopard seals. Host-associated bacterial communities are 46

thought to have evolved with their host and this is also suggested for the gut microbiota of mammals [23,36]. This study confirms the previous findings that con- specific hosts display greater similarity in their gut microbiota than non-con-specific hosts, suggesting co-evolution of mammals and their gut microbial communities.

Differences observed in the faecal microbiota of wild southern elephant and leopard seals can be attributed to distinctions in their gut morphology and physiology. Southern elephant seals have a small intestine equivalent to 25 times their body length [252], whereas the leopard seals have a shorter small intestine which is ten times the body length and within the range of other phocid seals [243]. In each of these hosts, the gut has evolved with different physiological capabilities for food metabolism. For instance, the shorter gut length in leopard seals is compensated for with an extended food retention time of 28-99 h [253]; whereas southern elephant seals have a retention time of 12-29 h [254]. The extended food retention time in leopard seals is thought to allow for better digestion of substantially larger prey items [253]. Southern elephant seals, by contrast, receive food often and in relatively small amounts and so whilst the stomach is large, it will empty often [252]. These differences in length and retention times would contribute to the ability of colonising bacteria to persist in the gut.

Differences in dietary items have the potential to change the gut microbiota through the introduction of novel OTUs or alterations in available nutrients, and therefore substrate available to bacterial members. As identified in Chapter 2, carnivorous mammals acquire a portion of the OTUs in their gut (~35%) from the gut of their prey. Although, southern elephant and leopard seals are carnivores, which forage in the same region and have overlap in particular prey items, there are differences, which may contribute to the observed dissimilarity in their gut microbiota. Southern elephant seals forage for squid and fish [133,255]; whereas, wild leopard seals consume a variety of prey such as seals, krill, penguins and fish [148,152,154]. There is further evidence to suggest that leopard seals in this study region feed primarily on krill [137,150,151,256].

3.5.2 Captivity and illness exert a strong affect on the gut microbiota of mammals 47

Wild and captive leopard seals possessed significantly dissimilar faecal microbiota suggested to be a result of the hosts experiencing different physiological and dietary factors in captivity compared to the wild. Metabolic rate, body condition, hormonal production and other functions are known to change in animals when placed in a captive environment [257-259]. The external environment for these hosts differs considerably, which may influence other aspects of metabolism and thermoregulation. For instance, temperatures experienced by captive leopard seals in Sydney range from 5 to 41˚C (averaging 14 to 41˚C in summer) [260]; whereas wild leopard seals in Antarctica experience temperatures ranging from -16 to 3˚C (averaging -1 to 3˚C in summer) [202]. Also, and as mentioned above for wild southern elephant and leopard seals, dietary differences contribute to the introduction of bacteria and also the available substrate in the gut. Captive leopard seals are fed a diet of fish caught off Sydney and not krill, penguins or other Antarctic prey. As was noted in a study on Atlantic cod, Gadus morhua, changes in the gut microbiota from a wild to a captive environment were observed after six weeks in response to artificial feeding of the fish, i.e. a change in diet [246]. Other studies which have identified distinctions between captive and wild animals, primarily from studies on primates, birds and fish [39-43,245,246], have suggested that diet was responsible for these changes. This study gives a clear indication that the gut microbiota of captive mammals does not represent that of their wild counterparts.

Illness and antibiotic use may have caused a shift in the gut microbiota of captive leopard seals. During illness and after the administration of antibiotics (see Table A1.2) in the female captive leopard, the faecal microbiota shifts considerably suggesting these may be the cause. Several host pathologies and their symptoms such as reduced eating and weight loss has been observed to alter the hosts’ gut microbiota [222,261]. Shifts in bacterial composition and decreases in diversity post- antibiotic administration have been shown in human hosts [29] and was suggested in this leopard seal (Chapter 2).

3.5.3 Age-class and maternal transfer shape the gut microbiota of southern elephant seals

The faecal microbiota of southern elephant seal’s is distinct at each age-class. 48

Changes in the gut microbiota as a mammal ages is attributed to changes in gut tissue, immune regulation, gut maturation, absorptive function and diet [14,15,17,262]. Southern elephant seal pups feed on milk until they are weaned after approximately 30 d [263,264]. Adults and sub-adults forage for a variety of fish and squid species [133,255]. Different diets would introduce distinct bacteria into the gut and the functional capacity of particular bacteria would be more beneficial for some diets, e.g. the ability to break down milk in the gut of pups. Particularly in younger mammals, the marked differences in faecal microbiota with older mammals has been attributed to facultative bacterial species dominating the young gut before being outnumbered by anaerobic species in adulthood [15]. The distinct faecal microbiota of adult and sub-adult southern elephant seals is due to their differences in prey abundance and prey size, as a result of the reduced size and therefore diving ability and foraging range of sub-adults [132,265,266]. This has been suggested to be an evolved strategy to reduce intra-specific competition of similar prey resources between adults and sub-adults, particularly as they return [265]. The distinct faecal microbiota of each age-class indicates the influence of diet and the physiological stages of development on the composition of the gut microbiota.

Southern elephant seal mothers are a source of the gut microbiota of pups. However, significant differences exist between the faecal microbiota of mothers and pups, yet on average the relative abundance shared in the gut microbiota of each pup with the mother was 34.7 ± 9.3%. OTUs shared between the mother-pup pairs varied considerably supporting the theory that pups do not get their gut microbiota solely from their mother and potentially get none as seen in pup F. This suggests that colonisation is unique for each individual during the initial period after birth as has been identified in other infant mammals [14,15,267,268]. The high level of similarity between the pups suggests that they are exposed to common OTUs through their local habitat and diet, which colonise their gut and persist due to their similar developmental stages.

3.5.4 Sex-driven dietary differences influences the gut microbiota of southern elephant seals

The faecal microbiota of female and male adult southern elephant seals differs. As 49

the mass of an individual host has been linked to the composition of their gut microbiota [26,269,270], the dramatic sexual dimorphism displayed by adult southern elephant seals was suggested as the cause of observed differences in their faecal microbiota. Adult females commonly weigh ~600 kg and fully-grown adult males weigh ~3,000 kg [252] yet, no clear relationship was observed between an OTU and host mass. An increase in sample replication may provide greater detail on the specific influence of particular members of the microbiota with host mass. Physiological capabilities and metabolic differences due to different growth strategies exist between male and female adult southern elephant seals and these are considered to influence their respective faecal microbiota. In particular, as with different age-classes, different physiological capabilities of males and females govern their foraging strategies allowing them to access distinct prey resources [124,271]. Males forage more commonly on the benthos and their increased body size provides them with the added capability to handle larger prey items compared with females [129,271]. It is these differences in prey resources that influence the faecal microbiota between male and female adult southern elephant seals.

3.6 Conclusions

The results of this study suggest that from the earliest colonization events through to adulthood, the gut microbiota of wild marine mammals is determined by multiple factors. Host species, captivity, maternal relationship, age-class and sex are primary drivers in the diet available to an individual that strongly influences the gut microbiota, often as a result of altered gut morphology and physiological capabilities. In this study these factors were suggested as ways of describing the observed patterns. There is interplay between many of these factors, and also the phylogenetic history of the host as observed in the shared OTUs between wild southern elephant and leopard seals.

Chapter 4

The gut microbiota of wild and captive Antarctic seals as identified using 16S rRNA gene pyrosequencing

4.1 Abstract

Until recently, our knowledge of bacterial communities was limited by our ability to culture and grow colonies in order to identify them. This excluded ~99% of all bacterial species from discovery. Now advanced techniques, such as next- generation sequencing, are providing significant progress in our understanding of bacterial communities, particularly within the gastro-intestinal tract of mammals. The gut microbiota of mammals underpins the health and success of hosts; therefore characterising this community has implications for understanding aspects of their ecology and health. To further understand the bacterial taxa responsible for observed differences in wild mammals, 16S rRNA gene pyrosequencing was performed on the faecal microbiota of wild southern elephant and leopard seals inhabiting Antarctica and compared to faecal samples from leopard seals housed in Taronga Zoo, Sydney. Amongst all seals sampled (n = 38), four phyla dominated the gut microbiota: Firmicutes (41.5 ± 4.0%), Fusobacteria (25.6 ± 3.9%), Proteobacteria (17.0 ± 3.2%) and Bacteroidetes (14.1 ± 1.7%). The faecal microbiota of southern elephant seals and wild leopard seals differed significantly in relative abundances of Bacteroidetes and Proteobacteria. The faecal microbiota of wild leopard seals possessed a significantly increased abundance of operational taxonomic units (OTUs) from the phyla Proteobacteria compared with captive leopard seals. Several OTUs were present across most individuals within and between host species groups (southern elephant, wild leopard and captive leopard seals). In southern elephant seals, evidence of preferential transfer of ‘core’ OTUs between mother-pup pairs was observed. The presence of a ‘core’ microbiota within and across host species highlights the strong evolutionary history shared between wild mammals and their associated bacteria.

4.2 Introduction

It is estimated that > 99% of microorganisms observed in nature are not easily cultivated using traditional techniques [272]. A suite of culturing conditions can increase the range of microbial diversity discoverable in environmental samples [273]; yet, molecular methods provide the most time- and cost-efficient means of assessing microbial community structure. DNA fingerprinting methods, such as terminal restriction fragment length polymorphism [274], denaturing gradient gel electrophoresis [275] and ARISA (detailed in Chapters 2 and 3; [64]) have dramatically advanced our ability to efficiently and robustly identify microbial community structure [74,75,274]. However, these methods lack taxonomic sensitivity and generally have a reduced resolving power to detect numerically rare bacteria [276]. This can lead to a biased view of species richness within samples [276]. More advanced and informative techniques are providing an increased understanding of true bacterial richness in the natural world and the potential importance of these rare species [277].

Over the last 20 years, the routine use of PCR amplification of ‘housekeeping’ genes, directly from environmental samples, particularly those encoding the 16S rRNA gene [48], along with downstream cloning and sequencing, have allowed the characterisation of microbial communities in a wide range of habitats, from hydrothermal vents to disease-associated microbiota [56,278,279]. As a result, many novel evolutionary lineages have been identified, many of which are still known only from environmental sequences or non-cultivable representatives [48,50]. These ‘traditional’ molecular methods have resulted in a large, publicly available database of near full-length (~1,500 base pairs, bp) 16S rRNA gene sequences. The latest release of the Ribosomal Database Project v.10, a highly curated 16S rRNA gene repository, contains nearly two million sequences [280].

Next-generation sequencing technologies, applied to mining bacterial diversity include pyrosequencing, e.g. Roche 454 pyrosequencing, which involves tagged PCR amplification and sequencing of hyper-variable regions of the 16S rRNA gene. These regions are generally less than 400 bp in length, yet detection of taxonomic information is typically reliable, [277,281]. The first application of 16S rRNA gene 52

pyrosequencing to an environmental sample was in 2006 [276] and since then, this method has accounted for the discovery of almost 80% of non-cultivated species and an estimated 18,000 genera [13,50,277,278]. However, the gut microbiota of wild mammals remains relatively understudied using advanced sequencing methods. Studies of the gut microbiota of human and laboratory-reared mammals, in contrast, have identified generalities in the occurrence of taxonomic groups of bacteria with host traits [e.g. 220,231,282].

In Chapter 3, the composition of the faecal microbiota of wild and captive marine mammal hosts was identified using ARISA to be influenced by multiple factors including host diet and evolutionary history. This study aimed to further characterise and investigate the factors influencing composition of the faecal microbiota using methods with greater sensitivity and taxonomic resolution. Seal hosts used in Chapter 3 were sub-sampled and DNA extracted from faecal samples was analysed using next generation sequencing. Two hyper-variable regions of the 16S rRNA gene were sequenced and compared to the known database. The aims were to: 1) taxonomically identify and characterise the gut microbiota of these hosts; and 2) investigate microbial community structure in relation to the observed influences identified in Chapter 3.

4.3 Materials and methods

4.3.1 Sample collection and DNA extraction from Antarctic seals

Faecal samples were collected from southern elephant seals and leopard seals inhabiting the western Antarctic between September 2008 and March 2009 (see Chapter 3 for details). These were compared to faecal samples from leopard seals inhabiting Taronga Zoo, Sydney, NSW, Australia (see Chapter 2 and 3 for details). DNA was extracted from 40 samples (Table A1.1; see Chapter 2 for details) including 24 southern elephant seals (four male adults, seven female adults, four male sub-adults, three female sub-adults, four male pups and two female pups), 12 wild leopard seals (six male adults and six female adults) and two captive leopard seals (one male sub-adult and one female sub-adult, at two times).

4.3.2 Community sampling via 16S rRNA gene pyrosequencing

DNA from 40 samples was analysed using pyrosequencing of the 16S rRNA gene was carried out at the Research and Testing Laboratory (Lubbock, TX, USA). Primers 27F 5’-AGAGTTTGATCCTGGCTCAG-3’ and 519R 5’- GTATTACCGCGGCTGCTG-3’ were used to amplify by PCR a 500 bp product spanning the bacterial 16S rRNA gene hyper-variable regions V1-V3 [283]. Sequencing was carried out on a Roche 454 titanium sequencer. Initial quality control measures, used to ensure sequence fidelity, included: the removal of short sequences (<100 bp); removal of any sequence not perfectly matching the 5’ primer; and, removal of any sequence containing an unresolved nucleotide.

Sequence data were analysed using the Mothur v.1.13.0 suite of programs [284]. Potential chimeras were removed using the Chimera.slayer tool with the minsnp parameter set to 100. A 2% pre-clustering step was used to remove potential errors in sequence data [285]. Briefly this method assigns each sequence within 2% similarity to the cluster with most sequences [285]. Sequences were aligned to the SILVA database [93] and those that did not align in the appropriate zone were removed. Alignments were trimmed so that all sequences covered the entire alignment length. The final dataset comprised 251,648 sequences. Clustering was performed at a sequence similarity cut-off of 97%. Representatives of resultant OTUs were taxonomically identified using the SILVA taxonomy tool.

4.3.3 Data analyses

Square-root transformed abundance data were used to generate a resemblance matrix that was visualised using nMDS as detailed in Chapter 2. The OTUs characteristic of community structure were calculated using SIMPER as detailed in Chapter 2.To test for differences in composition of the faecal microbiota between seals based on a priori groupings of species, sex and age-class a one factor permutational multivariate analysis of variance (PERMANOVA) [216] was carried out as detailed in Chapters 2 and 3. Statistical tests were performed as detailed in Chapter 2 using the software PRIMER-E v6 [218]. As per Chapters 2 and 3, OTUs present in samples within a group, e.g. southern elephant seals, were identified as a

majority when present in more than half (most) of the individuals within that group. The sum of abundances of those OTUs were calculated and averaged as a relative abundance of the total to give the ‘shared’ OTUs within a group. Shared OTUs between groups were used to create a Venn diagram using an online applet [286]. The non-parametric estimator of species richness, Chao1 [287], was calculated using OTU abundance data for each individual using the online software program EstimateS [288]. Significant differences in microbial richness and phyla abundance between groups were tested using a Student’s T-Test in Excel 2010 (Microsoft Pty Ltd). Linear regression was used to examine the relationship between host weight and OTU abundance and richness in faecal microbiota.

To obtain contextual information regarding the provenance of any similar 16S rRNA gene sequences to our dataset, sequences were compared to the National Centre for Biotechnology Information non-redundant database (http://www.ncbi.nlm.nih.gov/) using the Basic Local Alignment Search Tool (BLAST) in GenBank [289]. The BLAST tool finds regions of local similarity between sequences, and compares nucleotide sequences to the sequence database to return a calculated statistical significance of matches. The sources of origin from highest scoring pairs were then classified as being from an external environment or ‘free-living’ (e.g. marine) or host- associated source (e.g. gut).

4.4 Results

4.4.1 16S rRNA gene pyrosequencing of the seal gut microbiota

A total of 251,648 sequences from the hyper-variable V1-V3 region of the 16S rRNA gene were obtained. This resulted in a total of 6,159 OTUs at ≥ 97% sequence similarity. Taxonomic characterization of the faecal microbiota identified representatives from 19 bacterial phyla (Table A4.1). The community was dominated by 5880 OTUs (98.2 ± 0.5% of total abundance) from four phyla: Firmicutes (41.5 ± 4.0%), Fusobacteria (25.6 ± 3.9%), Proteobacteria (17.0 ± 3.2%), and Bacteroidetes (14.1 ± 1.7%).

4.4.2 Influence of age-class on the gut microbiota of southern elephant seals

The gut microbiota of southern elephant seal pups was different to that of adults and sub-adults (t = 1.9902, PPERM = 0.0001 and t = 1.8357, PPERM = 0.001, respectively; Table 4.1). The gut microbiota of adults and sub-adults was identified as not significantly different (t = 1.2799, PPERM = 0.0398; Table 4.1) and therefore considered alike in future analyses. The gut microbiota of adults and sub-adults possessed a significantly greater abundance of the phylum Firmicutes (43.3 ± 5.4%) compared with pups (18.0 ± 2.7%; Figure 4.1). Instead pups possessed a significantly greater abundance of the phylum Fusobacteria (62.0 ± 4.4%) in their gut microbiota compared with adults and sub-adults (20.3 ± 3.9%; Figure 4.1). These OTUs included representatives from the genera Ilyobacter, Fusobacterium and Psychrilyobacter (Table A4.2). In addition, there was an increasing trend observed in the microbial richness with age-class, with pups having a significantly reduced gut microbiota when compared with adults and sub-adults (Figure 4.2).

Table 4.1 PERMANOVA on the gut microbiota of hosts based on groupings of species, location, age-class and sex. PERMANOVA of gut microbial abundance data to generate a permutated F statistic (F) and permutated p-value (P) with calculated degrees of freedom (d.f.) and sums of squares (SS) noted. P-values given in italics were obtained using Monte Carlo samples from the asymptotic permutation distribution. Pair-wise a posteriori tests between host species groups and age-classes were conducted using the t-statistic (t). Significance level: ***P = ≤ 0.001, **P = 0.01, *P = 0.025. Source of Data d.f. SS F P variation Southern elephant seals Age 2 15870 2.8356 0.0001*** Pair-wise comparison t P Adult, sub-adult 1.2799 0.0398 Sub-adult, pup 1.8357 0.001** Adult, pup 1.9902 0.0001*** Southern elephant seals (pups omitted), wild Species 2 30241 4.6477 0.0001*** leopard seals, captive and location leopard seals Pair-wise comparison t P Southern elephant seals, wild leopard seals 2.3479 0.0001*** Southern elephant seals, captive leopard seals 1.983 0.0003*** Wild leopard seals, captive leopard seals 1.9878 0.0004*** Southern elephant seal Age 1 10930 4.7383 0.0019 mothers and pups Southern elephant seal Sex 1 7151.8 2.3829 0.0033** adults and sub-adults Wild leopard seals Sex 1 4567.4 1.3732 0.1209 Captive leopard seals Individual 1 2751.4 1.079 0.4199

Figure 4.1 Average relative abundance of major phyla in the gut microbiota of seal host species groups. Average relative abundance of operational taxonomic units (OTUs) for each major phyla in the faecal microbiota of each host species group. Southern elephant seals were separated to reflect similarity in faecal microbiota. Error bars represent standard errors (SE). Student’s paired t-test were conducted between groups of individuals for each phylum. Significance level: ***P = ≤ 0.001, **P = 0.01, *P = 0.025.

Figure 4.2 Microbial richness of the gut microbiota of seal host species groups. Microbial richness of the faecal microbiota as measured using Chao 1 mean with error bars representing standard errors (SE). Southern elephant seals were separated to reflect similarity in faecal microbiota. Student’s paired t-test conducted between seal groups. Significance level: ***P = ≤ 0.001, **P = 0.01, *P = 0.025.

4.4.3 Influence of host species and captivity on the gut microbiota

Community composition of the gut microbiota displayed significant separation between host species groups, i.e. southern elephant seal adults and sub-adults, wild leopard seals and captive leopard seals (F = 4.6477, PPERM = 0.0001; Table 4.1; Figure 4.3). Pair-wise tests revealed each host species group was significant from one another (Table 4.1). Abundances of characteristic OTUs as identified by SIMPER analysis differed considerably between hosts species groups (Figure 4.4). The phylum Proteobacteria contributed more to the faecal microbiota of wild leopard seals (31.1 ± 6.9%) than to captive leopard seals (3.6 ± 1.8%) or southern elephant seals (14.7 ± 4.2%) although these were not significant (Figures 4.1 and 4.4). The phylum Bacteroidetes contributed significantly more to the faecal microbiota of southern elehant seals (20.7 ± 2.9%) than wild leopard seals (8.3 ± 2.6%) or captive leopard seals (2.2 ± 1.0%) (Figures 4.1 and 4.4). Microbial richness of the gut microbiota was significantly higher for southern elephant seal adults and sub-adults compared to wild leopard seals (Student’s t-test P = < 0.0001; Figure 4.2). However, captive leopard seals possessed a gut microbiota with richness significantly higher than southern elephant seal adults and sub-adults and wild leopard seals (Student’s t-test P = 0.0196 and 0.0001, respectively; Figure 4.2).

Dissimilarities in the gut microbiota of wild leopard and southern elephant seal adults and sub-adults compared with captive leopard seals were observed as a result of high abundances of characteristic OTUs identified using SIMPER analysis (Figure 4.4). In captive leopard seals the dominant characteristic OTUs were from the genera Subdoligranulum and Sporobacter from the phylum Firmicutes, and the genera Fusobacterium and Cetobacterium from the phylum Fusobacteria (Table A4.2). In southern elephant seal adults and sub-adults characteristic OTUs that were relatively more abundant than in wild leopard seals or captive leopard seals were from the genus Ilyobacter from the phylum Fusobacteria, and the genus Subdoligranulum from the phylum Firmicutes (Table A4.2). Characteristic OTUs present in high abundance in wild leopard seals included representatives from the genus Oceanospirillales from the phylum Proteobacteria and from the genus Clostridiales XI from the phylum Firmicutes (Table A4.2).

Figure 4.3 Similarity of the gut microbiota of seal host species groups. Dissimilarity of the faecal microbiota of host species groups displayed in nMDS plot.

In general, the female and male captive leopard seal shared many of the same characteristic OTUs in their faecal microbiota, except during one time point when the female captive leopard seal was ill (Figure 4.5). At the time of illness, the gut microbiota of the female captive leopard seal was dominated by two OTUs from the genus Cetobacterium from the phylum Fusobacteria, which contributed a relative abundance of 77.7% (Figure 4.5). The nearest GenBank sequence similarities of these OTUs were from faeces of captive polar bears, Ursus maritimus, housed in San Diego Zoo (Table A4.9). The relative abundance of OTUs from the genus Cetobacterium in the male captive leopard seal and the female captive leopard seal (during good health) was on average 0.1 ± 0.1%. During the period of illness, microbial richness of the female captive leopard seal’s gut microbiota also decreased although the male leopard seal also showed a decrease in richness and the low replication did not allow for significance testing (Table A4.3).

Figure 4.4 Characteristic OTUs in the gut microbiota of seal host species groups. Pie charts display the foremost 35% of characteristic operational taxonomic units (OTUs) in faecal microbiota as identified using SIMPER analysis of host species groups: (A) southern elephant seals (pups removed); (B) wild leopard seals; and (C) captive leopard seals. Unnamed members are marked (*) and their highest known classification is noted. 61

Figure 4.5 Changes in the gut microbiota of captive leopard seals through time. Pie charts display the foremost 80% of operational taxonomic units (OTUs) as identified using SIMPER analysis in female and male captive leopard seals over time: female time point one (during illness), 19-Jan-10 (A); female time point two, 25-Feb-2010 (B); male time point one, 12-Jan-10 (C); and male time point two, 23-Feb-10 (D). Information regarding sample collection is listed in Table A1.1. Unnamed members are marked (*) and their highest known classification is noted.

4.4.4 Shared OTUs within and between seal hosts

Shared OTUs occurred in highest abundances within host species groups rather than between them. Within southern elephant seal adults and sub-adults, 95 OTUs were present in most individuals with an average relative abundance of 56.4 ± 3.7% per individual (Table A4.4). One OTU, OTU10, from the family Clostridiales XI from the phylum Firmicutes was present within all southern elephant seal adults and sub- adults (Table A4.4). Amongst wild leopard seals, 63 OTUs were present in most individuals with an average relative abundance of 54.6 ± 5.5% per individual (Table A4.5). One OTU, OTU17, from the genus Weissella from the phylum Firmicutes was present within all wild leopard seals (Table A4.5). In the two captive leopard seals, the number and abundance of OTUs present in both individuals was up to four times greater than those in southern elephant seal adults and sub-adults or wild leopard seals. Captive leopard seals shared 49 OTUs at each of the sampling times

contributing an average relative abundance of 40.5 ± 11.5% per individual (Table A4.6). With the removal of the sample collected during the female’s illness the number of shared OTUs at each sampling time increased to 121 with an average relative abundance of 65.8 ± 3.2% per individual. Shared OTUs amongst captive leopard seals were primarily represented by genera Subdoligranulum, Fusobacterium, Sporobacter, Ilyobacter and Oscillibacter from the phyla Firmicutes and Fusobacteria (Table A4.6). Only three OTUs were represented by different phyla: the genus Escherichia / Shigella from the phylum Proteobacteria, the genus Bacteroides from the phylum Bacteroidetes and the genus Collinsella from the phylum Actinobacteria.

The gut microbiota of southern elephant seal pups shared the most than any other host species groups, including captive leopard seals. They each shared 28 OTUs, which contributed an average relative abundance of 75.0 ± 2.9% per individual. In the gut microbiota of most pups a further 115 OTUs were shared contributing an average relative abundance of 13.6 ± 2.0% per individual. Although the gut microbiota of pups was significantly different to their mothers (F = 4.7383, PPERM = 0.0019; Table 4.1; Figure 4.6A) 20 OTUs contributing 53.3 ± 11.5% to total abundance in each pup was observed (Figure 4.6B and Table 4.3). In addition, 19 of these OTUs were identified as being shared in the gut microbiota of most southern elephant seal adults and sub-adults (Table 4.3). The greatest contributor to abundance was OTU1 from the genus Ilyobacter from the phylum Fusobacteria with an average presence of 30.9 ± 4.3% per pup (Table 4.3). The three other highest contributing OTUs in pups were also from the phylum Fusobacteria from the genera: Fusobacterium (OTU3), Psychrilyobacter (OTU8) and Ilyobacter (OTU13). Other genera included Bacteroides and Psychrilyobacter as well as an unnamed member of the family Peptostreptococcaceae (Table 4.3).

Figure 4.6 Similarity and shared OTUs in the gut microbiota of mother-pup pairs. nMDS ordination of southern elephant seals mother-pup pairs designated by letters (A). Relative abundance of operational taxonomic units (OTUs) shared between mother and pup in a pair. Lower case letters represent mother-pup pairs.

Table 4.2 Shared OTUs in the gut microbiota of mother-pup pairs. Average relative OTU abundance in the gut microbiota of mothers or pups ± standard error (SE). OTUs shared between most (more than half) of the individual: (*) southern elephant seal adults and sub-adults and wild leopard seals; and (^) southern elephant seals (mothers and pups omitted) are marked. Average relative abundance / individual ± OTU Phylum Genus SE (%) Mothers Pups OTU1*^ Fusobacteria Ilyobacter 2.93 ± 1.36 30.91 ± 4.32 OTU3*^ Fusobacteria Fusobacterium 5.56 ± 4.02 13.00 ± 3.85 OTU8*^ Fusobacteria Psychrilyobacter 2.13 ± 1.44 6.25 ± 0.44 OTU13^ Fusobacteria Ilyobacter 0.53 ± 0.27 6.08 ± 0.85 OTU11^ Bacteroidetes Bacteroides 0.98 ± 0.93 4.20 ± 2.87 OTU2^ Firmicutes Subdoligranulum 2.60 ± 1.21 1.70 ± 0.74 OTU27^ Firmicutes Sporobacter 3.65 ± 1.86 0.16 ± 0.07 OTU31^ Proteobacteria Sutterella 1.60 ± 1.53 1.94 ± 0.48 Unnamed member from the OTU75^ Firmicutes family 0.16 ± 0.07 1.51 ± 0.23 Peptostreptococcaceae OTU48^ Actinobacteria Collinsella 0.44 ± 0.20 1.10 ± 0.70 Unnamed member from the OTU287^ Firmicutes family 0.02 ± 0.00 0.67 ± 0.08 Peptostreptococcaceae OTU81 Proteobacteria Escherichia / Shigella 0.09 ± 0.05 1.06 ± 0.44 OTU151*^ Firmicutes Anaerococcus 0.69 ± 0.40 0.21 ± 0.09 OTU24^ Proteobacteria Campylobacter 0.12 ± 0.06 0.49 ± 0.26 Unnamed member from the OTU101^ Bacteroidetes 0.86 ± 0.27 0.38 ± 0.15 family Prevotellaceae OTU110 Firmicutes Sarcina 0.52 ± 0.44 0.34 ± 0.16 OTU388^ Fusobacteria Fusobacterium 0.09 ± 0.06 0.46 ± 0.08 OTU17*^ Firmicutes Weissella 0.32 ± 0.12 0.20 ± 0.06 OTU447^ Fusobacteria Ilyobacter 0.06 ± 0.02 0.32 ± 0.11 OTU33^ Bacteroidetes Alistipes 0.07 ± 0.01 0.16 ± 0.11

Three OTUs belonging to the genera Ilyobacter, Fusobacterium and Psychrilyobacter from the phylum Fusobacteria were present in most individuals in each host species group (Table 4.4). These OTUs contributed an average relative abundance of 9.7 ± 2.1% per individual (Figure 4.6). Most individual southern elephant seal adults and sub-adults and wild leopard seals shared a further 10 OTUs contributing an average relative abundance of 21.0 ± 2.4% per individual (Table 4.5; Figure 4.6) and including the top three most abundant OTUs shared in the gut microbiota of most southern elephant seal pups (OTU1, OTU3 and OTU8; Table 4.5). Captive leopard seals and southern elephant seal adults and sub-adults shared 21 OTUs contributing 22.2 ± 3.7% to total abundance compared with three OTUs contributing 4.3 ± 2.0% between captive and wild leopard seals (Figure 4.6 and 65

Table A4.7). These three OTUs were those previously observed as shared amongst most individuals in all host species groups (Table 4.4).

Table 4.3 Shared OTUs in the gut microbiota of host species groups. Average relative abundance of operational taxonomic units (OTUs) shared in most (more than half) of individuals in a host species group ± standard error (SE). Average relative abundance / individual ± SE (%) Southern OTU Phylum Genus elephant Wild Captive seals leopard leopard (pups seals seals removed) OTU3 Fusobacteria Fusobacterium 4.4 ± 1.8 4.4 ± 1.8 3.5 ± 2.2 OTU1 Fusobacteria Ilyobacter 8.2 ± 2.4 8.2 ± 2.4 0.5 ± 0.2 OTU8 Fusobacteria Psychrilyobacter 1.9 ± 0.6 1.9 ± 0.6 0.9 ± 0.5

Figure 4.7 Shared OTUs within and between host species groups. Venn diagram displays the average relative abundance of shared operational taxonomic units (OTUs) in the gut microbiota of most (more than half) individuals in a host species group’s ± standard error. Each circle represents the complete average abundance of OTUs in the gut microbiota of individuals within the designated host species group.

Table 4.4 Shared OTUs in the gut microbiota of southern elephant seals and wild leopard seals. Average relative abundance of operational taxonomic units (OTUs) shared in most (more than half) of individual southern elephant seal adults and sub-adults and wild leopard seals ± standard error (SE). OTUs occurring in the gut microbiota of more than three quarters of individuals are marked (). Average relative abundance / individual ± SE (%) OTU Phylum Genus Southern Wild elephant leopard seals (pups seals removed) OTU17 Firmicutes Firmicutes 0.2 ± 0.1 5.6 ± 2.2 OTU10 Firmicutes Clostridiales XI 1.1± 0.1 7.0 ± 3.1 OTU3 Fusobacteria Fusobacterium 4.4 ± 1.8 3.3 ± 2.1 OTU8 Fusobacteria Psychrilyobacter 1.9 ± 0.6 0.5 ± 0.4 OTU1 Fusobacteria Ilyobacter 8.2 ± 2.4 0.2 ± 0.1 OTU15 Bacteroidetes Petrimonas 4.5 ± 2.1 0.8 ± 0.4 OTU454 Firmicutes Roseburia 0.1 ± < 0.1 0.1 ± < 0.1 OTU151 Firmicutes Anaerococcus 0.3 ± 0.1 0.3 ± 0.3 OTU88 Firmicutes Leuconostoc < 0.1 ± < 0.1 1.5 ± 0.6 OTU167 Firmicutes Anaerococcus 0.1 ± 0.1 0.1 ± 0.1 Unnamed member of the OTU65 Proteobacteria 0.5 ± 0.3 0.1 ± < 0.1 family Pseudomonadaceae OTU999 Firmicutes Clostridiales XI < 0.1 ± < 0.1 0.1 ± 0.1 OTU145 Bacteroidetes Porphyromonas 0.2 ± 0.1 0.2 ±0.1

4.4.5 The influence of sex and the external environment on the gut microbiota

There was a significant difference in the gut microbiota of male and female southern elephant seal adults and sub-adults (F = 7151.8, PPERM = 0.0033; Table 4.1; Figure 4.7). This difference was not observed between male and female wild leopard seals or male and female captive leopard seals (F = 4567.4, PPERM = 0.1209 and F =

2751.4, PMC = 0.4199; Table 4.1). OTUs from the phylum Firmicutes were significantly more abundant in the gut of southern elephant seal females (56.0 ± 6.0% SE) compared with males (27.5 ± 5.9% SE). Abundances of OTUs from the phylum Fusobacteria were significantly higher in the gut of males (30.5 ± 6.6%) compared with adult females (12.2 ± 3.1% SE). The mass of sexually dimorphic southern elephant seal adults and sub-adults was included in a PERMANOVA with no significant effect (Table A4.8). In addition, linear regression analysis yielded no significant correlations with microbial community richness, characteristic phyla or OTUs (Figure A4.1).

Figure 4.8 Average relative abundance of major phyla in the gut microbiota of southern elephant seal adult and sub-adult males and females. Average relative abundance of operational taxonomic units (OTUs) for each major phyla in the gut microbiota of southern elephant seal adult and sub-adult males and females. Error bars represent standard errors (SE). Student’s paired t-test were conducted between groups of individuals for each phylum. Significance level: ***P = ≤ 0.001, **P = 0.01, *P = 0.025.

Across all seal hosts there was little difference in the abundance of OTUs in an individual’s gut microbiota deemed to be ‘free-living’ from an ‘external’ source. Free- living OTUs contributed an average relative abundance of 3.2 ± 1.3% in the gut microbiota of all seals. Free-living OTUs included representatives from the phyla Proteobacteria, Actinobacteria, Acidobacteria, Cyanobacteria, Deinococcus- Thermus, Chloroflexi, Gemmatimonadetes, Planctomycetes, Chlorobia, Spirochaetes, Verrucomicrobia and the candidate division TM7. Captive leopard seals displayed a reduced abundance of free-living OTUs (0.5 ± 0.4%), as did southern elephant seal pups (1.0 ± 0.5%) but there were no significant differences. Reanalysis of data excluding these ‘free-living’ sequences did not alter the findings described above.

4.5 Discussion

4.5.1 ARISA and 16S rRNA gene pyrosequencing methods are compatible

Differences in resolution between DNA fingerprinting using ARISA (Chapter 3) and 16S rRNA gene pyrosequencing are apparent. Pyrosequencing generated ~250,000 sequences yielding ~6,000 OTUs, whilst only 361 OTUs were identified using 68

ARISA. This difference is at least partially a result of the ARISA method, as with other DNA fingerprinting methods, the power of resolution is reduced [75,290]. Community fingerprinting may miss the majority of ‘rare’ species due to detection limits for resolving unique peaks or when different species generate the same size fragment resulting in them being binned into the same OTU [290,291]. The pyrosequencing method provides a means to observe ‘rarer’ OTUs which are potentially undetectable using the ARISA method [75]. However, the patterns influencing microbial community structure observed using both methods are comparable as has been previously observed in comparative studies [e.g. 274,292]. These are compatible methods, with pyrosequencing providing further insight into the taxonomy of OTUs driving observed differences in community structure, yet there is, at this stage, greater cost efficiency with the use of DNA fingerprinting methods such as ARISA.

4.5.2 Diet and gut physiology drive differences in the gut microbiota

Significant differences in the gut microbiota were observed between host species groups (southern elephant seal adults and sub-adults, wild leopard seals and captive leopard seals). These differences in community composition exist despite their broadly similar carnivorous diet, suggesting that colonisation of the gut is dramatically shaped by stochastic exposures during an individual's lifetime. They consume primarily proteins and lipids, which requires similar microbial functionality in their gut microbiota, such as, breaking down proteins for energy use. Due to the compositionally similar diets consumed by these hosts and the different OTU and phyla presence in their gut microbiota, this suggests that functionality of the gut microbiota is spread across bacterial taxa. Therefore, general functional tasks may be performed by numerous different OTUs and not any one particular OTU, known as functional redundancy. Previous studies have also identified the presence of functional redundancy in the gut habitat [e.g. 38] and on the surface of algae in marine environments [e.g. 293].

Composition of the gut microbiota of carnivorous hosts is the result of prey- associated OTUs. The introduction of prey-associated OTUs is an identified transmission pathway introducing OTUs into the gut microbiota of these hosts 69

(Chapter 2). Southern elephant seal adults and sub-adults and captive leopard seals shared 21 OTUs compared to three OTUs between wild and captive leopard seals. This is suggested to be due to the fish-dominated diets of southern elephant seal adults and sub-adults and captive leopard seals. OTUs from the phylum Proteobacteria, specifically the genus Psychrobacter, have been observed in the gut of Antarctic krill, Euphausia superba, Adelié penguins, Pygoscelis adeliae and gentoo penguins, P. papua [68,294], all of which are prey of wild leopard seals. The number of shared OTUs in the gut microbiota of southern elephant seal adults and sub-adults and captive leopard seals could be due, in part, to their consumption of a diet dominated by fish, more so than wild leopard seals.

Differences in the gut microbiota of hosts are also suggested to be due to selection from physiology and the available nutritional components influencing colonisers. The longer gut of the southern elephant seal provides a habitat that is reduced in available oxygen and they possess a greater abundance of members from the phylum Bacteroidetes, which are known to be obligate anaerobes. In contrast, the shorter gut length of leopard seals provides a habitat with increased oxygen availability and it is suggested that facultative anaerobes, such as those of the phylum Proteobacteria, particularly the class Gammaproteobacteria, are found in greater abundance in these hosts as a result. Bacterial composition due to oxygen availability as controlled by the gut length has been proposed as contributing to alterations in the gut microbiota of humans having undergone gastric bypass [295]. The proportions of available compounds and nutrients also influences the gut microbiota as has been recognised in components such as fibre in herbivores and omnivores [296]. There is limited knowledge available on the nutritional composition of prey items for Antarctic predators, although it could be suggested that wild leopard seals eat a diet which is higher in lipids (krill and seal pups) compared with the equivalent amount consumed by captive leopard and southern elephants seals [297- 299].

4.5.3 A conserved core gut microbiota exists in Antarctic seal hosts

Bacteria in the gut of mammals perform numerous functions that are beneficial to the host including: immune regulation, tissue maturation, intestinal development, and the 70

extraction of energy, nutrients and vitamins from dietary items [7,10]. The tight coupling of this symbiotic relationship suggests bacteria in the gut are evolutionarily linked to their mammalian host [23,36,37]. Southern elephant seal adults and sub- adult and wild leopard seals shared 16 OTUs in their gut, which is suggested to be the ‘core’ gut microbiota which has co-evolved with these mammalian hosts. Three of these ‘core’ OTUs, all from the phylum Fusobacteria, were shared with captive leopard seals, indicating the maintenance of the ‘core’ under altered conditions.

Nineteen core OTUs were identified as being transferred from southern elephant seal mothers to their pups, including three OTUs shared amongst all host species groups from the genus Ilyobacter and Psychrilyobacter. The gut microbiota of pups performs different functions compared to that required by older hosts. In young mammals, the gut microbial community is required to develop the immune system, mature the gut and break down the milk in order to provide energy [15,17,18]. The transfer of these OTUs into pups under three weeks of age and their apparent persistence through to adulthood, suggests they may provide a function which is not related directly to the diet, as adults and pups consume very different diets. Instead, they may serve a purpose of immune regulation as they are recognised by the innate immune system as harmless, which may be an indication of a long evolutionary association [300]. Rather than produce an aggressive immune response, which could result in harm to the new pup, these bacteria may actually prime the immune system in pups and later, in adults, regulate other aspects of the immune system [301]. If the role of core OTUs is linked to immune function of the hosts, the maintenance of the OTUs is ensured through vertical transmission (mother-to-infant transmission).

4.5.4 Antibiotic-use and captivity may exert a strong influence on the gut microbiota

The presence and persistence of Cetobacterium OTUs after antibiotic-use suggests the presence of antimicrobial resistance. When the female captive leopard seal was exposed to antibiotics, OTUs from the genus Cetobacterium from the phylum Fusobacteria increased dramatically. The proliferation of resistant strains of Cetobacterium, including species such as C. somerae, have been previously been 71

associated with the administration of antibiotics [302]. These Cetobacterium OTUs were identified in the gut microbiota of the captive leopard seals during good health, in wild leopard seals and in southern elephant seal adults and sub-adults. Other studies also report the presence of OTUs from this genus in the gut of freshwater fish [303,304]; the mouth of a minke whale (Balaenoptera acutorostrata); and the gut of a harbor porpoise (Phocoena phocoena) [305]. Additionally, the nearest GenBank sequence similarity to these OTUs was from captive polar bears faeces housed in San Diego Zoo. The presence of this genus and specifically these OTUs in seemingly healthy hosts suggests Cetobacterium is a member of the ‘normal’ leopard seal gut microbiota with the ability to flourish in response to antibiotic usage, possibly as a result of resistance.

The gut microbiota of captive and wild leopard seals is dissimilar as a result of the different exposures between these hosts. Con-specific hosts have been identified to display dissimilarities when sampled from captive and wild environments, although few have identified this in mammals, and this is the first study which identifies this incident in marine seals [e.g. 39,40,42,43,245]. Dietary items can drive differences in the gut microbiota through the introduction of OTUs associated with prey. Additionally, the influence of antibiotics results in a completely reduced faecal microbiota and although the community returns in a few weeks, it has been suggested that the community is incomplete [29]. The different faecal microbiota and low number of shared OTUs between wild and captive leopard seals suggests that the effects of antibiotics may be more long-term. Other studies have suggested that mammals in captivity serve as an indicator for wild species; however, the use of antibiotics appears to force changes in the gut microbiota due to re-colonisation and may result in the loss of some of the ‘core’ microbiota of these hosts.

4.5.5 Social interactions increase the transmission of OTUs between hosts

Southern elephant seal adults and sub-adults possessed a richer gut microbiota and a higher number of shared OTUs amongst individuals compared to wild leopard seals. This is a result of the social nature of southern elephant seal adults and sub- adults, compared to wild leopard seals. Southern elephant seals come ashore to give birth, breed and molt, congregating in colonies of 10s to 100s of individuals 72

[125,306]. The close proximity to large numbers of con-specifics has been identified as an added benefit of social behaviour in animals, providing increased transmission opportunities [225]. Wild leopard seals, by comparison, spend relatively little time with con-specifics and are solitary except for brief periods when mating and in mother-pup pairs post-partum [143,153,307]. The greater number of shared OTUs amongst southern elephant seal adults and sub-adults also suggests ongoing transfer of bacteria. This pattern has been observed in social bee species, where greater similarity was found in social Apis and Bombus species compared with the solitary Agapostemon species, indicating facilitated transmission of bacteria between individuals [224].

Captive leopard seals possessed a significantly richer gut microbiota and increased abundance of shared OTUs to one another, more so than southern elephant seal adults and sub-adult or wild leopard seals. As with southern elephant seal adults and sub-adults, the great number of shared OTUs and is due to the constant social interaction, co-habitation and ‘commonalities’ shared by the captive leopard seals, such as diet. In a previous comparative study, an increase in gut microbial richness was associated with a ‘contaminated’ and high-fibre diet [220]. It is suggested that the interaction of the captive leopard seals with human keepers, general public, other mammals (when kept in holding pens) and diet, which has been handled, provides increased opportunities for transmission of OTUs from host-associated sources capable of colonising the seals and resulting in a richer gut microbiota.

4.5.6 Negligible contributions to the gut microbiota from the external environment

The proportion of gut microbiota from an external or free-living environmental source (e.g. marine) was estimated to be negligible (3.2 ± 1.3%) across all seal hosts. Current global surveys suggest that dominant members of the gut microbiota, such as those form the phyla Firmicutes and Bacteroidetes, do not survive outside of their host [2]. Amongst the identified free-living OTUs, they included an OTU belonging to the group, SAR11, which is globally distributed in the marine water [308,309] and an OTU which was 99% identical to Prochlorococcus marinus, a bacterium never before identified in the Antarctic marine environment [310]. It is likely that these were 73

present in water droplets adhering to the skin of prey at the time of feeding or was introduced into the sample during handling and represent contamination.

Members of the phylum Fusobacteria, including those genera observed in the core gut microbiota, Ilyobacter and Psychyrilyobacter, have been identified in habitats such as anoxic sediments and anoxic enrichments of marine material [311]. Therefore, exposure from the environment into the gut of these hosts may have been underestimated. However, the persistent and high abundances of these OTUs in the gut across multiple species and age-classes suggests they are maintained in the gut habitat and presence between mother-pup-pairs is evidence for their vertical transmission.

4.5.7 Sexual dimorphism drives dietary influences on the gut microbiota

Sex related differences were only identified in the gut microbiota between southern elephant seal adults and sub-adults, which are sexually dimorphic. Fully grown, male adult southern elephant seals weigh ~3,000 kg, whereas females are commonly ~600 kg [252]. Studies of humans have identified shifts in the presence and abundance of phyla and overall richness in obese versus lean subjects which was suggested to be as a result of increase energy extraction from the gut microbiota [26,312]. It has since been discovered that the increase in dietary fats, not increased energy extraction capabilities, results in the observed composition of the gut microbiota [313]. For these hosts, it is likely that foraging depths and distances resulting in dietary differences of the males and females causes differences in their respective gut microbiota [271]. As identified in Chapter 3 physiological and metabolic differences due to different growth strategies exist between southern elephant seal adult and sub-adult males and females, including the ability to access distinct prey resources [124,271].

4.6 Conclusions

This study confirmed previous suggestions identified in Chapter 3 of a distinct gut microbiota in southern elephant seals, wild leopard seals and captive leopard seals. Diet type and composition, gut morphology and physiology, social behaviours and

antibiotics have primary roles in determining community composition. The external environment is thought to have a negligible role in shaping the community in these hosts. This study highlighted the strength of ‘rarer’ OTUs identified by pyrosequencing in the gut microbiota of an individual. A ‘core’ seal gut microbiota was present in these seals, established via mother-to-pup transmission and remains throughout the life of the host. These ‘core’ OTUs may have an essential role in the host and have been able to persist due to co-evolution. Suggestions of the potential role of core OTUs are to regulate the immune system or gut development. Experimental studies that include the alteration of diets and/or habitats would provide insight into the relative contribution of diet and evolutionary history in assembling the gut microbiota of these wild hosts.

Chapter 5

Comparing the gut microbiota of mammals from marine and terrestrial habitats

5.1 Abstract

After birth, mammals acquire a community of bacteria in their gastro-intestinal tract, which harvests energy and provides nutrients for the host. Comparative studies of numerous terrestrial mammal hosts have identified host phylogeny, diet and gut morphology as primary drivers of community composition. To date, marine mammals have been excluded from these comparative studies, yet they represent distinctive examples of evolutionary history, dietary and lifestyle traits. To provide a contextual understanding of the gut microbiota of marine mammals, 16S rRNA gene pyrosequencing of DNA extracted from faeces from southern elephant seals and leopard seals completed in Chapter 4 were compared to published studies on the gut microbiota (also based on faecal samples) of three Arctic seals, one dugong, Dugong dugon, and 109 terrestrial mammal species. Antarctic and Arctic seals shared 60 operational taxonomic units (OTUs), which displayed ≥ 99% sequence similarity, in their gut microbiota. These OTUs were highly specific to seals, enforcing the concept of a ‘core’ seal gut microbiota conserved in their phylogeny, regardless of their separation to each polar region ~15-25 million years ago (mya). When compared to other terrestrial mammals, Antarctic and Arctic seals were more similar to each other than to other species and possessed an increased presence of the phylum Fusobacteria than terrestrial mammals. The carnivorous marine mammals (Antarctic and Arctic seals) and the marine herbivore (dugong) possessed significantly richer gut microbiota than terrestrial carnivores and terrestrial herbivores, respectively. This suggests that evolutionary history and dietary items specific to the marine environment has resulted in an altered gut microbiota to that identified in terrestrial mammals.

5.2 Introduction

Bacteria inhabiting the gastro-intestinal tract of mammals expand their host’s metabolic potential by harvesting energy that would otherwise be inaccessible [3,8,10]. This symbiosis between mammals and bacteria has contributed, in part, to the success of the class Mammalia, allowing them to radiate in large numbers to occupy a variety of environmental niches [314]. For instance, herbivorous mammals were able to survive on plant material after acquiring gut microbiota with the capacity to digest cellulose in plant cell walls [315].

Mammalian hosts first acquire their gut microbiota during transport through the birth canal and subsequently through maternal, social and environmental transmissions [15,225]. Genetic factors within the host also shape the gut microbiota, a result of their long history of co-evolution [32,34,36]. This is evident in the strong physiological effects which the gut microbiota can exert on the host mammal, such as modulating the immune response system or affecting brain development [316,317].

In a pioneering study, Ley et al. [23] compared the faecal microbiota, as proxy for the gut microbiota, of a variety of terrestrial mammals and identified that host phylogeny, diet and, to a lesser extent, gut morphology influenced the composition of the gut microbiota [12,23]. Studies have shown that the composition of the gut microbiota follows along evolutionary lineages [36,37] yet recently, it has been suggested that diet is the primary driver of functional capacity in the gut, resulting in a convergence of communities between phylogenetically related hosts [38].

Further insight could be gained by comparing a diverse range of extant mammals with differing life history traits. One group of mammals that have been relatively understudied is marine mammals. Their comparatively recent evolution and differing life history traits, adapted to a marine habitat [113], suggest they are a necessary addition to the current understanding of mammal faecal microbiota.

To identify patterns in the composition of the faecal microbiota of terrestrial and marine mammals, the gut microbiota of the Antarctic southern elephant and leopard seals from the family Phocidae generated using 16S rRNA gene pyrosequencing of

DNA extracted from faeces in Chapter 4 were compared to published gut microbiota data from three phocid seal species inhabiting the Arctic. A meta-analysis was conducted including this dataset, data from one dugong (a marine herbivore), and data from several published studies from a variety of terrestrial mammals (n = 109). The aims were to: 1) investigate if the gut microbiota of the family Phocidae is evolutionarily conserved; and 2) identify broad scale patterns of mammalian gut microbiota with terrestrial and marine dietary groups.

5.3 Materials and methods

5.3.1 Data acquisition

Pyrosequencing of the 16S rRNA gene was performed on DNA extracted from faeces of southern elephant seals (n = 24) and leopard seals (n = 14) (Table A1.1; see Chapter 4 for details). Data were compared to a published study of marine seals [44]: harbour seals, Phoca vitulina (n = 1), grey seals, Halichoerus grypus (n = 1), and hooded seals, Cystophora cristata (n = 9; Table A5.1). The dataset from Antarctic and Arctic seals was then compared to published studies from a range of terrestrial mammals (n = 109) and the dugong (n = 1) (Table A5.1). Studies were selected on the basis that analysis methods for microbial community composition were similar to 16S rRNA gene pyrosequencing and involved sequencing regions of the 16S rRNA gene for taxonomic identification. Individual sequence data were obtained from the National Centre for Biotechnology Information website (http://www.ncbi.nlm.nih.gov/) or directly from the source article. Sequence taxonomy was assigned using the Ribosomal Database Project (RDP) v.10 Classifier tool (http://rdp.cme.msu.edu/classifier/classifier.jsp) [280]. Sequences from Antarctic seals were compared to 319 full length 16S rRNA gene sequences from Arctic seals using Basic Local Alignment Search Tool (BLAST) (http://blast.ncbi.nlm.nih.gov/Blast.cgi) [318]. To normalise relative abundances of sequences from multiple hosts, sequences were randomly sub-sampled to a maximum of 100 sequences using the software Daisy-Chopper (www.genomics.ceh.ac.uk/GeneSwytch/). The result was a total of 13,848 sequences from 151 mammalian hosts (Table A5.1).

5.3.2 Data analyses

Sequence data from multiple mammal hosts were standardised prior to transformation to remove differences in total abundance where sequence totals were less than 100. Briefly, standardisation involved turning abundance counts into relative percentages for each host. Data were square-root-transformed and used to generate resemblance matrix that was visualised using nMDS as detailed in Chapter 2. The OTUs characteristic of community structure were calculated using SIMPER as detailed in Chapter 2. To test for differences in the composition of the gut microbiota between hosts based on a priori groupings of diet, location, phylogeny and gut morphology. Analysis of similarity (ANOSIM) was used. ANOSIM is a non- parametric permutation procedure, which is applied to the similarity matrix and generates a test statistic, R. Compared to PERMANOVA, ANOSIM is more robust to heterogeneous dispersion of data which is common to this dataset [218]. The magnitude of R is indicative of the difference within groups and between groups and is scaled to lie between - 1 and + 1, a value of zero represents the null hypothesis (no difference between groups) and value towards one represents the alternative hypothesis (all similarities within groups are less than any similarity between groups). R-values > 0.50 (**) were interpreted as well separated; R > 0.30 (*) as overlapping but clearly different; and, R > 0.20 as barely separable at all. Results were considered significant where P-value = < 0.025. The non-parametric estimator of species richness, Chao1 [287], was calculated using OTU abundance data for each individual using the online software program EstimateS [288]. Significant differences in microbial richness and phyla abundance between groups were tested using a Student’s T-Test in Excel 2010 (Microsoft Pty Ltd). All other statistical tests were performed as detailed in Chapter 2 using the software PRIMER-E v6 [218].

5.4 Results

5.4.1 The gut microbiota of Antarctic and Arctic phocid seals

A total of 60 16S rRNA gene sequences, represented by 19 genera, from the gut microbiota of Arctic seals displayed ≥ 99% sequence similarity to the gut microbiota of Antarctic seals (Table 5.1). These hosts are phylogenetically related within the

phylum Phocidae (Figure A5.1). The most abundant of these genera were Alistipes and Bacteroides of the phylum Bacteroidetes; Butyricicoccus, Faecalibacterium, Oscillibacter, Sporobacter and Subdoligranulum of the phylum Firmicutes; Fusobacterium of the phylum Fusobacteria; and, Sutterella of the phylum Proteobacteria (Table 5.1).

Table 5.1 Shared genera in the gut microbiota of Antarctic and Arctic seals. Genera observed in the faecal microbiota of the harbour, hooded and grey seals inhabiting the Arctic and the relative average abundance ± standard error / individual (%) in host species groups of Antarctic seals. Genera present in (*) most (more than half) individuals within each host species group are marked. Average relative abundance ± SE / individual (%) Southern Phylum Genus elephant seal Wild leopard Captive adults and seals leopard seals sub-adults Bacteroidetes Bacteroides* 2.1 ± 0.6 2.0 ± 1.1 0.5 ± 0.2 Firmicutes Faecalibacterium* 2.2 ± 1.2 0.6 ± 0.2 0.3 ± 0.1 Fusobacterium Fusobacterium* 5.5 ± 1.9 7.6 ± 2.9 7.0 ± 2.7 Firmicutes Sporobacter* 7.2 ± 1.3 1.8 ± 1.1 19.1 ± 6.1 Firmicutes Oscillibacter* 1.5 ± 0.4 0.5 ± 0.3 9.0 ± 2.8 Firmicutes Subdoligranulum* 10.4 ± 4.1 0.5 ± 0.2 22.2 ± 7.4 Bacteroidetes Alistipes* 2.7 ± 0.8 0.1 ± 0.1 0.4 ± 0.3 Firmicutes Butyricicoccus* 2.0 ± 1.7 0.6 ± 0.4 0.2 ± 0.1 Bacteroidetes Paraprevotella 0.4 ± 0.3 < 0.1 ± < 0.1 0.1 ± < 0.1 Proteobacteria Sutterella* 0.8 ± 0.3 0.3 ± 0.1 0.6 ± 0.3 Escherichia / Proteobacteria Shigella 0.3 ± 0.1 0.9 ± 0.9 1.1 ± 0.5 Proteobacteria Anaerobiospirillum 0.2 ± 0.1 0.1 ± < 0.1 < 0.1 ± < 0.1 Firmicutes Sporacetigenium 0.1 ± < 0.1 0.9 ± 0.9 0.3 ± 0.2 Firmicutes Clostridium 0.1 ± 0.1 0.2 ± 0.2 0.7 ± 0.3 Firmicutes Blautia < 0.1 ± < 0.1 < 0.1 ± < 0.1 < 0.1 ± < 0.1 Firmicutes Dorea < 0.1 ± < 0.1 < 0.1 ± < 0.1 0.1 ± < 0.1 Proteobacteria Lawsonia < 0.1 ± < 0.1 < 0.1 ± < 0.1 - Firmicutes Ethanoligenens - - < 0.1 ± < 0.1 Firmicutes Mahella - - -

Shared sequences in the gut microbiota of Antarctic and Arctic seals were compared to sequences in GenBank and found to be more similar across the sequence length to sequences sourced in seals than any other host or external environment (Table 5.2). Similarity across the sequence length is a measure used to compare similarity of previously sequenced bacterial strains or species. In this case, the average similarity of sequences from seal hosts was 99% and 96% from a non-seal host, which included humans, mice, dogs and polar bears. 80

Table 5.2 Nearest seal and non-seal sources of shared sequences in the gut microbiota of phocid seals. Sequences shared between the gut microbiota of the harbour (csh), hooded (wsp/s7), and grey seals (csg) from the Arctic and southern elephant and leopard seals from the Antarctic were compared to sequences in GenBank using the Basic Local Alignment Search Tool (BLAST) to find the nearest source host for each sequence. The uploaded sequence length is compared to the database of known and sequenced bacteria and produces a list of sequences and their sources with the percent similarity of gene sequence coverage between the two sequences (Max. ID %). In the first instance the list was searched for a seal host source and left blank when absent. Closest Max. Closest similarity to non- Max. Genus similarity to seal ID (%) seal host source ID (%) host source Fusobacterium Pig intestine 98 Fusobacterium Wild wolf faeces 99 Faecalibacterium csh45 99 Pig faeces 94 Fusobacterium Human faeces 98 Alistipes wsp107 99 human faeces 95 Faecalibacterium csh45 99 pig intestine 94 Faecalibacterium csh89 99 human faeces 95 Escherichia / Shigella human faeces 99 Bacteroides csh95 99 human faeces 95 Sutterella human faeces 96 Sporobacter wsp115 99 pig faeces 96 Subdoligranulum csg20 99 polar bear 99 Faecalibacterium csh33 99 human faeces 95 Bacteroides wsp135 99 human faeces 95 Subdoligranulum csg20 95 piglet faeces 94 Ethanoligenens csg58 99 mouse faeces 97 Butyricicoccus human colon 97 Anaerobiospirillum s738 99 human blood 98 Subdoligranulum csh82 99 polar bear faeces 99 Bacteroides wsp104 99 human faeces 95 Faecalibacterium csh28 99 pig faeces 94 Peptostreptococcus csg67 99 polar bear 99 Anaerobiospirillum s7 31 99 human blood 98 Oscillibacter wsp115 99 estuarine sediment 96 Ethanoligenens csg63 99 mouse faeces 97 Paraprevotella wsp131 99 human faeces 93 Subdoligranulum wsp79 99 geese colon 95 Bacteroides csh95 99 human faeces 95 Peptostreptococcus csg44 99 polar bear faeces 99 Alistipes wsp83 99 human faeces 95 Mahella csg34 99 human faeces 95 Escherichia/ Shigella s750 95 wastewater 90 Bacteroides csh67 99 human faeces 95 Butyricicoccus 99 human faeces 96 Faecalibacterium wsp128 99 pig faeces 94 Faecalibacterium csh45 99 pig faeces 94 Clostridium polar bear faeces 99 Bacteroides csh95 99 human faeces 95 Subdoligranulum csh70 99 rat faeces 95 81

Faecalibacterium csh52 99 dog colon 94 Bacteroides wsp32 99 human faeces 95 Blautia csg82 99 human faeces 97 Alistipes wsp109 99 human faeces 96 Lawsonia mouse faeces 95 Sporacetigenium mangrove sediment 99 Oscillibacter wsp112 99 wolf faeces 97 Dorea wsp93 99 human faeces 99 Alistipes wsp96 99 human faeces 95 Faecalibacterium csh89 99 human faeces 95 Blautia csg27 99 human faeces 97 Bacteroides csh95 99 human faeces 95 Bacteroides csh77 99 human faeces 95 Bacteroides csh32 99 human faeces 95 Faecalibacterium csh89 99 human faeces 95 Alistipes wsp107 99 human faeces 96 Alistipes wsp107 99 human faeces 96 Oscillibacter wsp26 99 human faeces 96 Bacteroides csg57 99 human faeces 95 Bacteroides csg62 99 geese colon 95 Faecalibacterium csh45 99 pig faeces 94

5.4.2 The gut microbiota of marine and terrestrial mammals

The gut microbiota of mammals was strongly influenced by diet type, with herbivores and carnivores displaying significant differences in gut community composition (ANOSIM: R = 0.49, p = < 0.01; Table 5.3; Figure 5.1). The gut microbiota of marine carnivorous mammals and terrestrial carnivorous mammals was significantly different (R = 0.69, p = < 0.01; Table 5.3). Con-specific hosts from the same family were more similar than non-con-specific hosts (R = 0.50, p = < 0.01). Across all mammals, the gut morphology of hosts did not result in significant differences of the gut microbiota (Table 5.3). Previously, the influence of captivity in leopard seals was identified as a strong driver of the gut microbiota (Chapters 3 and 4). In this study, six host species were sampled from both captive and wild environments and compared in an nMDS plot (Figure A5.2) with no significant effect noted.

Marine carnivorous mammals possessed a significantly lower average relative abundance of the phylum Firmicutes in their gut microbiota with 43.2 ± 6.7% compared to 65.6 ± 6.8% in the gut microbiota of terrestrial mammals and 68.9% in the gut microbiota of the herbivorous marine mammal (Figure 5.2). The phylum Proteobacteria was significantly more abundant in marine carnivores with an 82

average relative abundance of 15.6 ± 2.4% compared with 6.2 ± 0.6% in the gut microbiota of terrestrial mammals. The phylum Bacteroidetes was similar for each of the dietary groups of terrestrial and marine mammals with an average relative abundance of 19.2 ± 1.7% with the exception of terrestrial carnivores, which displayed a significantly reduced abundance of 3.7 ± 1.0%.

Figure 5.1 Influence of diet and habitat on the gut microbiota of mammals. nMDS ordination plot dislaying similarity of the gut host mammals grouped by diet and location.

Table 5.3 ANOSIM of the gut microbiota of host mammals based on groupings of diet, location, phylogeny and gut morphology. ANOSIM of gut microbial abundance data to generate a permutated Global R statistic (R) and permutated p-value (P). Significance level: **R = > 0.5, *R = 0.3 < R < 0.5. Source of variation Pair-wise comparisons R P Diet 0.39* < 0.01 Herbivore, omnivore 0.33 < 0.01 Herbivore, carnivore 0.49* < 0.01 Omnivore, carnivore 0.26 < 0.01 Diet and location 0.52** < 0.01 Terrestrial herbivore, terrestrial omnivore 0.33* < 0.01 Terrestrial herbivore, terrestrial carnivore 0.62** < 0.01 Terrestrial herbivore, marine carnivore 0.65** < 0.01 Terrestrial omnivore, terrestrial carnivore 0.32 < 0.01 Terrestrial omnivore, marine carnivore 0.50** < 0.01 Terrestrial carnivore, marine carnivore 0.69** < 0.01 Phylogenetic Order 0.19 < 0.01 Phylogenetic Family 0.50** < 0.01 Gut Morphology 0.11 < 0.01 Hindgut fermentor, simple 0.14 < 0.01 Hindgut fermentor, foregut fermentor 0.17 < 0.01 Simple, foregut fermentor 0.15 < 0.01

The phylum Fusobacteria was significantly greater in marine carnivores with an average relative abundance of 22.0 ± 3.4% compared with other dietary groups (2.1 ± 0.2%) (Figure 5.2). Domestic dogs, Canis lupus familiaris, were the only non- marine carnivore with a higher than average abundance of the phylum Fusobacteria with 24 ± 1.0%. The domestic dog clustered closest to the marine carnivorous mammals in the nMDS plot (Figure A5.3) and this is highlighted when compared amongst the order Carnivora (Figure 5.3). Similarity of the gut microbiota between members of the family Ursidae is also noted regardless of dietary preference with the clustering of pandas and bears (Figure 5.3).

Figure 5.2 Average relative abundance of major phyla in the gut microbiota of mammals based on groupings of location and diet. Average relative abundance of operational taxonomic units (OTUs) for each major phyla in the gut microbiota of mammal groups based on loaction and diet. Error bars represent standard errors (SE). The lack of replication in the marine herbivore grouping does not allow for estimation of SE or significance testing. Student’s paired t-test were conducted between groups and this data is in Table 5.3.

Table 5.4 Student’s t-test of the main phyla in the gut microbiota of mammals based on groupings of location and diet. Student’s paired t-test of gut microbial abundance data to generate a p-value (P). Significance level: ***P = ≤ 0.001, **P = 0.01, *P = 0.025. The lack of replication in the marine herbivore grouping does not allow for significance testing. P Comparison Firmicutes Bacteroidetes Proteobacteria Fusobacteria Terrestrial herbivore, 0.4733 0.9963 0.4233 0.0010 terrestrial omnivore Terrestrial herbivores, 0.0333 0.0015* 0.8280 0.0025 terrestrial carnivore Terrestrial herbivore, 0.0002*** 0.5020 0.0184* < 0.00001*** marine carnivore Terrestrial omnivore, 0.1652 0.0010** 0.2662 0.7354 terrestrial carnivore Terrestrial omnivore, 0.0003*** 0.5362 0.0038** 0.0004*** marine carnivore Terrestrial carnivore, 0.00002*** 0.0016** 0.1509 0.0050** marine carnivore

Figure 5.3 Similarity of the gut microbiota between hosts from the order Carnivora. nMDS plot displays faecal microbiota of all mammal host families in the order Carnivora. Host species labels are as follows: (RP) red panda; (BB) black bear; (PB) polar bear; (SB) spectacled bear; (GP) giant panda; (BEAR) bear from Norway; (CE) cheetah; (BDOG) bushdog; (DOG) domestic dog; (LI) lion; (HY) spotted hyena; (GREY) grey seal; (HOOD) hooded seal; (HARB) harbour seal; (ES) southern elephant seal; (LS) leopard seal.

Overlap in the presence of particular genera between host dietary groups was evident. Marine and terrestrial herbivores, as well as omnivores, displayed overlap in the genera Anaerotruncas, Ruminococcus and Roseburia from the phylum Firmicutes (Table A5.2). These were less abundant in marine or terrestrial carnivores. Some genera, including the genus Oscillibacter from the phylum Firmicutes, and the genera Prevotella and Bacteroides from the phylum Bacteroidetes were abundant in all groups except for terrestrial carnivores (Table A5.2). Likewise, some genera, such as Coprococcus and Blautia from the phylum Firmicutes were abundant across all groups with the exception of the marine carnivores. The genus Lactobacillus from the phylum Firmicutes was commonly shared between carnivorous hosts compared with other diet groups (Table A5.2).

Herbivores possessed a faecal microbiota significantly richer than that of carnivores or omnivores (Figure 5.4). The single marine herbivore displayed a gut microbiota richer (Chao 1 mean = 141.0) than that of terrestrial herbivores (Chao 1 = 51.0 ± 2.8) or marine carnivores (Chao 1 = 47.0 ± 7.4), although insufficient replication did not allow for significance testing of this pattern. The marine carnivores possessed 86

significantly richer faecal microbiota than the terrestrial carnivores (Figure 5.4). Additionally, hindgut fermentors displayed a significantly richer gut microbiota than hosts with simple gut morphology (T-Test: P = 0.0028).

Figure 5.4 Microbial richness of the mammalian gut microbiota based on groupings of location and diet. Microbial richness of the gut microbiota as measured using Chao 1 mean with error bars representing standard errors (SE). Student’s paired t-test conducted between groups. Significance level: ***P = ≤ 0.001, **P = 0.01, *P = 0.025. The lack of replication in the marine herbivore grouping does not allow for estimation of SE or significance testing.

5.5 Discussion

5.5.1 A shared core gut microbiota in Antarctic and Arctic phocid seals

This study indicated the co-occurrence of 60 OTUs representing 19 genera between five species of Antarctic and Arctic phocid seals, suggesting these OTUs are highly conserved within the phocid phylogenetic lineage and identify the existence of a ‘core’ gut microbiota. The Phocidae family made up of the sub-family Monachinae (Antarctic seals) and Phocinae (Arctic seals) are thought to have split ~15-25 mya to their current opposite polar distributions [114,119]. However, beyond this core group of bacteria, significant differences in the faecal microbiota have been observed between different phocid seal species (Chapter 4). Host-species-specific gut microbiota have been identified in wild hominids, where gut microbial composition is known to mirror host phylogeny [36]. While the Arctic seal dataset in the present 87

study was not sufficiently replicated to perform a similar analysis, the significant differences evident between gut microbiota of southern elephant seals and leopard seals suggest that seal phylogeny may similarly be reflected in gut community structure (Chapter 4).

5.5.2 A distinct marine mammal gut microbiota

The composition of the gut microbiota of marine mammals is clearly distinct from that of terrestrial mammals, despite the identification of distinctions within the Antarctic seals based on species, age-class and sex (Chapters 3 and 4). In addition, the differences observed in Chapters 3 and 4 relative to the influence of a captive environment were reduced when compared with numerous other mammals. The results indicate that the influences of host phylogeny and diet are stronger than the influence of a captive environment on a broad scale.

The gut microbiota of carnivorous marine mammals differs in their considerably reduced abundance of Firmicutes and increased abundance of Fusobacteria compared to terrestrial mammals. Members of the phylum Fusobacteria range from facultative anaerobes to obligate anaerobes that ferment carbohydrates or amino acids and peptides to produce various organic acids including acetic, formic and butyric acid, depending on the substrate and species [311,319,320]. Species occur in sediments as well as the oral or intestinal habitats of animals [311,321-323]. The presence of these species commonly in nature, suggests the close clustering of domestic dogs to seals in the nMDS plot may be due to the environmental exposure by both hosts to Fusobacteria, yet they exist in such a great abundance that their presence in the gut of dogs and seals could be a further indication of evolutionary links. The Canidae (dogs) are located with the Phocidae in the order Carnivora [134; Figure 1.1]. Canids and phocids possess shared immune system receptors and diseases, such as Morbillivirus, that have been passed between dog and seal hosts [324,325]. As suggested in Chapter 4, Fusobacteria may have a role in immune- system regulation. Conserved legacy effects are further indicated from the observed similarity of faecal microbiota between phylogenetic groups, including carnivores (polar bears) and herbivores (pandas) from the family Ursidae. This pattern was identified in the original study conducted by Ley et al. [23], although the present 88

study strengthens the finding with the inclusion of additional mammal hosts from the ursid family.

5.5.3 Richness of the gut microbiota is influenced by diet type

The non-parametric estimator of species richness, Chao 1, used in this study and also in Chapter 4, is considerably reduced in this study due to its reliance on counting individuals sampled from a community [326]. As this dataset was sub- sampled prior to analysis to allow for standardised comparisons of community members, the estimated richness is also reduced. As the purpose of Chao 1 estimation in this study was to compare between hosts and its use regardless of the reduction in values is relevant.

Several challenges are faced when consuming plant material as a primary food source due to the indigestible cell walls [315]. The need to access complex carbohydrates in plants, such as cellulose and starch, is thought to be the driver of a rich gut microbiota of herbivores [23]. In the present study, terrestrial and the marine herbivores were significantly richer than terrestrial omnivores and carnivores (confirming previous findings [23]). This suggests that marine mammals possess a richer gut microbiota than terrestrial mammals. In the marine environment, secondary metabolites produced by plants and other primary producers may be considerably higher [315,327,328], which could impact the faecal microbiota of herbivores or carnivores at higher trophic levels. Dugongs primarily consume seagrass and are known to occasionally supplement their diet with macro invertebrates and algae [329,330]. The dugong sampled in this case was in captivity during sampling and had been fed a diet of the eelgrass, Zostera marina [45]. Eelgrass has been identified as producing secondary chemical defences, specifically phenolic acids, which have the capacity to cause considerable reduction of bacteria at even low dosages in experimental studies [331,332]. The marine carnivores in this study are also known to feed directly on lower trophic levels, causing them to be exposed to secondary metabolites [333,334]. Marine mammals may require a larger community, one with additional functions enabling the break down of excess chemical compounds. Increased sampling of marine herbivores would enable us to unravel these processes. 89

5.6 Conclusion

The co-occurrence of many shared OTUs with ≥ 99% sequence similarity in Antarctic and Arctic phocid seals, despite their geographic separation for millions of years, indicates the highly conserved nature of the gut microbiota in these hosts. The shared OTU sequences are clearly distinct from their nearest matches in non-seal mammals, suggesting they represent a ‘core’ seal gut microbiota. The significant difference between marine and terrestrial mammals further suggests that different rules may govern the assembly of the gut microbiota of these mammals.

Chapter 6

Investigating Escherichia coli pathotypes and antimicrobial resistance patterns in wild southern elephant seals, Antarctica

6.1 Abstract

Human wastes have the potential to introduce agents that can cause disease in local animal populations. This is of particular concern in pristine wilderness regions, inhabited by populations of endemic wildlife. In Antarctica, human sewage has the potential to transfer novel bacteria or genetic elements into wild mammals. Samples of Antarctic seawater, incorporating contamination from a human sewage outfall, and faecal material from wild southern elephant seals were collected to identify human wastes as a potential transmission source. Virulence factors in Escherichia coli and antimicrobial resistance (AMR) were indexed for isolated bacteria. Potentially pathogenic extra-intestinal E. coli (ExPEC) and entero-pathogenic E. coli (EPEC) were identified in faecal samples from southern elephant seals. Additionally, for the first time in an Antarctic mammal, the occurrence of AMR was identified in bacteria isolated from the faeces of southern elephant seals. AMR in isolates from seawater correlated with human sewage contamination. This study identified southern elephant seals as a host of E. coli pathotypes, ExPEC and rarely EPEC and reports the occurrence of AMR in these hosts. AMR, including multi-resistant isolates, are prevalent in the seawater surrounding sewage outfalls from scientific stations, highlighting this as a possible route for gene transfer. Given the biodiversity value of the Antarctic environment and its extant fauna, the findings of this study warrant further investigation and management.

6.2 Introduction

The increase in global transport and human travel is contributing to the emergence of novel infectious diseases in humans and wildlife [167,335]. This may arise via direct input of pathogens or the transmission of novel genetic elements taken up and incorporated into endemic populations. In the marine environment, this is an acute issue. For many years, wastes such as human sewage have been dumped into the ocean environment, often untreated and in large volumes. In Antarctica, dumping of untreated human sewage into the marine environment is commonplace [336]. As the ability to travel to Antarctica has become easier with engineering advances, scientific research and visitations to the region have also increased [337]. There are approximately 100 research stations in Antarctica with populations that generate between 500 to 270,000 L of wastewater per day, often with little or no treatment and released into the marine coastal environment [336,338]. In addition to fixed research stations, tourist ship visits reach ~35,000 per annum which may also dispose of wastes overboard [336,339]. Due to the endemicity of the local fauna, the introduction of disease in this region could be damaging [169,336]. In addition, recent climatic predictions suggest Antarctic predators, such as seals, will suffer from a shift in resource availability (prey and habitat), with the potential to further compromise the ability of hosts to fight infections [340].

Identifying transmission pathways in wildlife populations is essential before the outbreak of disease. Wild mammals, particularly those with relatively limited prior human contact, maintain unique communities of host-associated bacteria (as was identified in Chapters 4 and 5). To identify routes of transmission between hosts is to ‘type’ well-known bacterial isolates based on the presence of genetic elements such as virulence factors and antimicrobial resistance (AMR). One common model organism for this is the bacterium, Escherichia coli. The main habitat of this bacterium is in the gastro-intestinal tract of humans and other vertebrates. Indexing patterns of genetic traits in E. coli, such as virulence factors, has provided an ability to distinguish between different host sources [341,342]. Cells of E. coli are released into the environment through faecal deposition, where they can survive for long periods, permitting infection between host species [343]. Previous studies have

yielded little occurrence of AMR or E. coli with and without virulence factors (pathotypes) in Antarctic fauna [180,344]. AMR has been observed in seawater contaminated with human sewage in the Antarctic suggesting this is a viable source in this environment [345-347]. Pathotypes of E. coli, notably EPEC, have been observed in Antarctic fur seal (Arctocephalus gazella) pups [180], although to date few studies have aimed to identify the occurrence of these in Antarctica and Antarctic mammal fauna.

This study aimed to investigate the occurrence of E. coli pathotypes and AMR in faeces of southern elephant seals and nearby sewage-contaminated seawater. The aims were to: 1) investigate the presence of E. coli pathotypes and AMR in bacterial isolates in faecal samples from southern elephant seals; and, 2) investigate the potential for human-contaminated seawater as a transmission source of E. coli pathotypes and AMR.

6.3 Materials and methods

6.3.1 Study site

The Island 25 de Mayo / King George Island, South Shetland Islands (6214’00”S, 5842’00”W) is in western Antarctica (Figure 6.1). Seawater samples were collected from Potters Cove, adjacent to the Argentinean research station of Jubany (Figure 6.1). Southern elephant seals were located on beaches off Potter Peninsula (Antarctic Specially Protected Area; ASPA No. 132) (Figure 6.1).

There are 17 major facilities located on 25 de Mayo / King George Island with representatives from eight countries. Jubany scientific station is under management of the Instituto Antártico Argentino (IAA) operating year-round [348]. The period March to December rarely sees more than 30 people, whereas during the period January to March, this number may reach a maximum of 100 [348]. Sewage and domestic wastes are discharged into the coastal waters of the cove (Figure 6.1) after primary treatment with sewage disposal averaging 365,000 L per annum [348]. Throughout the period of sampling, untreated sewage was released into Potter Cove due to malfunctioning plant operations, which had been ongoing for a period of 12

months. During this time, approximately 2,000 tourist cruise ships visited the site surrounding 25 de Mayo / King George Island and Potter Cove [339].

Figure 6.1 Potter Cove, 25 de Mayo / King George Island displaying sampling locations. Sites of seawater sampling are located at circled numbers 1, 2, 3, 4 and 5. Arrows (blue) represent the hydrodynamic flow of the bay as identified by Schloss et al. [349]. Sites of interest surrounding the Jubany base are: (A) glacier melt water stream; (B) sewage treatment outfall; (C) glacier melt water stream; and, (D) the area of southern elephant seal breeding colonies stretching ~3 km north east.

Potter Cove is described as ‘fjord-like’ with a mouth area of 3,500 m2 and an inner portion of 3,000 m2 [350]. Depths in the inner portion rarely exceed 30 m with a maximum of 50 m at the mouth and a maximum of 100-200 m in the middle [350]. 94

Circulation of Potter Cove follows a cyclonic pattern [349] (Figure 6. 1). Input from glacial ablation and melt water is thought to lower the temperature and salinity in the Cove often resulting in suspended sedimentation [349]. Sea ice rarely forms over the area [350].

6.3.2 Sample collection and bacterial growth from southern elephant seals

Rectal swabs were collected from southern elephant seals from the western Antarctic (see Chapter 3 for details) and stored at 1 to 4°C during transfer to the laboratory. In total, rectal swabs were collected from 41 southern elephant seals (five male adults, ten female adults, thirteen male sub-adults, three female sub-adults, eight male pups and two female pups; Table A1.1). Swabs were plated onto MacConkey’s and Tryptone Soy Agar plates (Oxoid Tube Media, Oxoid Australia, Pty Ltd) and grown for 24-48 h at 35°C. Following growth, colonies were scraped into cryopreservation media of brain heart infusion broth (Oxoid Australia, Pty Ltd) supplemented with 31% glycerol (Sigma-Aldrich Pty. Ltd, NSW). Samples were stored at -20°C until transport to Sydney.

6.3.3 Sample collection and bacterial growth from seawater

Water samples were collected from five sampling locations at three times (Figure 6.1; Table A1.1). Sampling locations were chosen to account for the sewage outfall and the hydrodynamic flow of the bay. Seawater was collected at an approximate depth of 30 cm using ethanol-sterilised containers. Two replicate samples of 500 ml were collected ~5 m apart at each sampling site. Samples were stored between 1- 4°C during transfer to the laboratory. Water samples were vacuum filtered onto 47 mm, 0.2 m nitrate filters (Sartorius Stedim Australia Pty. Ltd., VIC, Australia) at 40 kPa using a motorized vacuum pump (Silfab, Aspirador A Diafragma, model N33V-A, Industria Argentina). After filtration, the filter was transferred to a dehydrated selective differential medium, mEndo Nutrient Pad Sets (Sartorius Stedim Australia Pty. Ltd.) and incubated at 20-25°C for 24 to 48 h (Steridium Pty. Ltd., QLD, Australia). Lower incubation temperatures were employed to account for previous findings that Antarctic bacteria isolated from seawater grow under different conditions than their temperate counterparts [351]. However, growth of seawater

samples proved difficult due to failure of equipment and loss of colony formation for replicates of each site during time one and for all samples and replicates grown at time two. These were not included. This yielded a total of 11 samples, which were scraped and stored in cryopreservation media for transport.

6.3.4 Sample processing from rectal swabs

Samples were thawed in the laboratory and a swab of inoculum was plated onto MacConkey’s Agar and grown at 35°C for 40 h. Streaks of colonies were placed into tubes of 5 ml LB broth (BD Difco, ON, Canada) and enriched at 37°C for 10-18 h. DNA templates were prepared from the resulting enrichments by boiled cell lysis. Briefly, 1 ml of each enrichment was pelleted by centrifugation, resuspended in 0.5 ml of sterile deionised water and boiled for 10 min. The boiled-cell suspensions were centrifuged and the resulting lysates were stored at -20°C before use in PCR.

6.3.5 Identification of virulence factors by multiplex PCR

Extracted DNA was assayed via multiplex PCR for the presence of eleven virulence factors defining the most important E. coli pathotypes found in animals and humans (Table 6.1). The virulence factors, amplification primers, multiplex PCR and control strains are listed in Table 6.2. Multiplex PCRs one and two were completed using the following thermal cycling conditions: one cycle consisting of 2 min at 95°C, 25 cycles consisting of 30 s at 94°C, 30 s at 60°C, and 30 s at 72°C, and held at 4°C. Multiplex PCR three was completed using the following thermal cycling conditions: one cycle consisting of 5 min at 95°C, 25 cycles consisting of 30 s at 94°C, 30 s at 55°C, and 5 min at 72°C, and held at 4°C. All PCRs were carried out on a Biometra Tpersonal thermocycler using negative control E. coli O115 (ECL3463), positive control strains (Table 6.2) and following Reference Laboratory for Escherichia coli (EcL) laboratory procedures (www.apzec.ca) [352]. The PCR products were visualized by ethidium bromide staining after agarose gel electrophoresis.

Table 6.1 Multiplex PCR and virulence factors used to detect E. coli pathotypes. Virulence factors identifying pathotypes are based on current knowledge taken from the Global APZEC (Animal Pathogenic and Zoonotic E. coli) Project being undertaken at the OIE Reference Laboratory for Escherichia coli, University of Montreal, Saint-Hyacinthe, Canada (www.ecl-lab.com). Multiplex PCR PCR Virulence factors Escherichia coli pathotypes virulence factors eae EPEC Entero-pathogenic E. coli 1 eae:Stx1:Stx2 Shiga toxin producing E. stxA, stx2A STEC coli 2 STa:STb:LT:F4 estA, estB, eltB, faeG ETEC Entero-toxigenic E. coli PapC:CNF1/2:Tsh: 3 papC, cnf, tsh, iucD ExPEC Extra-intestinal E. coli Aerobactin

Table 6.2 Multiplex PCR conditions and control strains used for detection of virulence factors in E. coli isolates. Positive control strains were sourced from Escherichia coli Laboratory (EcL), University of Montreal, Saint-Hyacinthe, Canada (www.ecl- lab.com). References (Ref.) for PCR amplification primers are listed. Further information on EcL control strains are listed in Table A6.4. Final Virulence factor Primer Sequence primer Positive control strain serotype Ref. conc. EAE (Intimin) eae F: CAT TAT GGA ACG GCA GAG GT 10 pM E. coli O111 (EcL6611) [353] EAE (Intimin) eae R: ATC TTC TGC GTA CTG CGT TCA 10 pM E. coli O111 (EcL6611) [353] Stx1 stxA F: TTA GAC TTA TCG ACT GCA AAG 10 pM E. coli O111 (EcL6611) [354] Stx1 stxA R: TGT TGT ACG AAA TCC CCT CTG 10 pM E. coli O111 (EcL6611) [354] Stx2 all Shiga-like toxin type II subunit A stx2A F: TTA TAT CTG CGC CGG GTC TG 10 pM E. coli O111 (EcL6611) [354] & B Stx2 all Shiga-like toxin type II subunit A stx2A R: AGA CGA AGA TGG TCA AAA CG 10 pM E. coli O111 (EcL6611) [354] & B STa estA F: TCC CCT CTT TTA GTC AGT CAA CTG 5 pM E. coli O149:H10 [355,356] (EcL7805) [356] R: GCA CAG GCA GGA TTA CAA CAA [355,356] STa estA 5 pM E. coli O149:H10 (EcL7805) [356] AGT STb estB F: GCA ATA AGG TTG AGG TGA T 10 pM E. coli O149:H10 [355,356] (EcL7805) [357] STb estB R: GCC TGC AGT GAG AAA TGG AC 10 pM E. coli O149:H10 [355,356] (EcL7805) [357] LT eltB F: TTA CGG CGT TAC TAT CCT CTC TA 10 pM E. coli O149:H10 [355,356] (EcL7805) [358] LT eltB R: GGT CTC GGT CAG ATA TGT GAT TC 10 pM E. coli O149:H10 [355,356] (EcL7805) [358] F4 K88ab1 & K88ab2 faeG F: ATC GGT GGT AGT ATC ACT GC 10 pM E. coli O149:H10 [355,356] (EcL7805) [359] F4 K88ab1 & K88ab2 faeG R: AAC CTG CGA CGT CAA CAA GA 10 pM E. coli O149:H10 [355,356] (EcL7805) [359] P (PapC) papC F: TGA TAT CAC GCA GTC AGT AGC 10 pM EcL13421 [360] P (PapC) papC R: CCG GCC ATA TTC ACA TAA C 10 pM EcL13421 [360] CNF1/2 cnf F: TCG TTA TAA AAT CAA ACA GTG 10 pM EcL13421 [360] CNF1/2 cnf R: CTT TAC AAT ATT GAC ATG CTG 10 pM EcL13421 [360] Tsh tsh F: GGT GGT GCA CTG GAG TGG 10 pM E. coli O78 (EcL3110) [361] Tsh tsh R: AGT CCA GCG TGA TAG TGG 10 pM E. coli O78 (EcL3110) [361] Aerobactin iucD F: AAG TGT CGA TTT TAT TGG TGT A 10 pM E. coli O78 (EcL3110) [362] Aerobactin iucD R: CCA TCC GAT GTC AGT TTT CTG 10 pM E. coli O78 (EcL3110) [362] 98

6.3.6 Isolation of bacteria positive for virulence factors

Three or more isolated colonies from each enrichment found positive for virulence factors, were examined by multiplex PCR. Eleven virulence factors were targeted (eae, stxA, stx2A, estA, estB, eltB, faeG, iucD, tsh, papC, CNF).

6.3.7 Antimicrobial resistance (AMR) testing

Antimicrobials tested were those used in the Canadian Integrated Program for Antimicrobial Resistance Surveillance (CIPARS) [363]. AMR testing was carried out on one or two isolates selected randomly from each sample using the standard Kirby-Bauer disk diffusion method [364]. Briefly, a swab of inoculum was plated onto MacConkeys Agar and grown at 35°C for 24-40 h. A single colony was selected with a toothpick and plated onto blood agar base (BD Difco, ON, Canada) supplemented with 5% whole bovine blood and incubated at 37°C for 10-18 h. A suspension of the isolate was made in 10 ml sterile water to a turbidity of 0.5 MacFarland Standards using an optical density meter (Oxoid Turbidimeter, BD Difco). Sterile swabs were used to streak the suspension onto 150 mm Mueller-Hinton agar plates (BD Difco) and left to dry for approximately five minutes. Fifteen commercially prepared antimicrobial agent disks (BD BBL, ON, Canada) (Table 6.3) were placed onto the inoculated agar surface using a disk dispenser and flame-sterilized forceps. Plates were inverted and incubated at 37°C for 18-20 h.

6.3.8 Data analyses

Attempts were made to assign each E. coli isolate to a particular pathotype according to its set of virulence factors (Table 6.1). Data for AMR of each bacterial isolate were reported as resistant (R) or susceptible (S, not resistant) depending on the diameter of the zone of inhibition (in millimetres; Table 6.3) in relation to known values of resistance from approved standards [364].

Table 6.3 Antimicrobial agents and their diameter sensitivity interpretation. Antimicrobial agents were chosen based on their importance in treating human or animal E. coli infections and on the basis of their ability to provide diversity for representation of different antimicrobial agent classes [341]. Cephalosporins are categorised as (a) second- generation antibiotics or (b) third-generation antibiotics. Values of resistance are based on approved standards from the Clinical and Laboratory Standards Institute [364]. Disk Drug Resistant Intermediate Susceptible Antimicrobial agent potency code (mm) (mm) (mm) (µg) Aminoglycosides Amikacin AMK 30 ≤ 14 15-16 ≥ 17 Gentamicin GEN 120 ≤ 12 13-14 ≥ 15 Streptomycin STR 300 ≤ 11 12-14 ≥ 15 Kanamycin KAN 30 ≤ 13 14-17 ≥ 18 Phinicols Chloramphenicol CHL 30 ≤ 12 13-17 ≥ 18 Quinolones and fluoroquinolones Nalidixic Acid NAL 30 ≤ 13 14-18 ≥ 19 Ciprofloxacin CIP 5 ≤ 15 16-20 ≥ 21 Tetracyclines Tetracycline TET 30 ≤ 14 15-18 ≥ 19 Sulfonamides and potentiated sulfonamides Sulfisoxazole FIS 250 ≤ 12 13-16 ≥ 17 Trimethoprim- SXT 23.75 ≤ 10 11-15 ≥ 16 sulphamethaoxazole Beta-lactams Amoxicillin / AMC 20/10 ≤ 13 14-17 ≥ 18 clavulanic acid Ampicillin AMP 10 ≤ 13 14-16 ≥ 17 Cephalosporin Cefoxitin (a) FOX 30 ≤ 14 15-17 ≥ 18 Ceftriaxone (b) CRO 30 ≤ 13 14-20 ≥ 21 Ceftiofur (b) TIO 30 ≤ 17 18-20 ≥ 21

6.4 Results

6.4.1 Prevalence of E. coli virulence factors

Sample DNA from the faeces of 41 southern elephant seals were analysed for the presence of 11 virulence factors (Table 6.4). Only factors defining ExPEC were observed. The most prevalent factor was iucD, followed by papC and CNF. There was a trend towards higher prevalence of virulence factors in pups compared with older animals. Seawater results were negative for all but one factor, iucD, which occurred at site three during time one and site five during time three (Table 6.4).

100

Table 6.4 Relationship between the presence of virulence factors of E. coli and site of origin in faecal samples from seals of different age-classes and seawater. All samples were negative for eae, Stx1, Stx2, Sta, Stb, LT, F4 and Tsh. Seawater sampling sites (S) and time point (T) in Potter Cove are listed in Figure 6.1. iucD papC CNF Group n No. % No. % No. % Southern Adults 16 9.0 56.3 4.0 25.0 1.0 6.3 Elephant Sub-adults 16 6.0 37.5 3.0 18.8 0.0 0.0 Seal Pups 10 7.0 70.0 7.0 70.0 2 20.0 S1, T1 1 0.0 0.0 0.0 0.0 0.0 0.0 S2, T1 1 0.0 0.0 0.0 0.0 0.0 0.0 S3, T1 1 1.0 100.0 0.0 0.0 0.0 0.0 S4, T1 1 0.0 0.0 0.0 0.0 0.0 0.0 S5, T1 1 0.0 0.0 0.0 0.0 0.0 0.0 Seawater S1, T3 2 0.0 0.0 0.0 0.0 0.0 0.0 S2, T3 2 0.0 0.0 0.0 0.0 0.0 0.0 S3, T3 2 0.0 0.0 0.0 0.0 0.0 0.0 S4, T3 2 0.0 0.0 0.0 0.0 0.0 0.0 S5, T3 2 2.0 100.0 0.0 0.0 0.0 0.0

The presence of a positive virulence factor in the sample DNA prompted analysis of isolated colonies from the sample enrichment. Virulence factors were present in isolated colonies from 17 southern elephant seals. Overall, the most common positive pathotypes were iucD (81.5%), papC (48.1%) and iucD:papC (35.2%) (Table 6.5). Other pathotypes included iucD:cnf (5.6%), eae (3.7%), tsh (3.7%), iucD:tsh (1.9%) and iucD:papC:cnf (1.9%) (Table 6.5). The prevalence of papC was least abundant in the adult southern elephant seals. The eae factor was found on two occasions, isolated once from a male adult southern elephant seal and once from a female adult southern elephant seal. Prevalence of the different pathotypes appeared to be similar in the three age-class groups of adults, sub-adults, and pups. Repeat analysis of isolates from these individuals did not yield further positive isolates for the eae gene. In the two seawater locations with positive samples, isolates were tested for the presence of tsh, papC, CNF and iucD. The only positive pathotype was iucD (50.0%) at site five during time three (Table A6.1). In three of the mother-pup pairs, virulence factors iucD and papC present in the mother were also present in the milk-drinking pup (Table A6.2).

101

Table 6.5 Occurrence of E. coli pathotypes in southern elephant seal faecal samples grouped by age-class. All samples were tested and negative for Stx1, Stx2, Sta, Stb, LT, and F4. Replicate number of samples (n) tested are shown. No. of isolates (%) positive for: EPEC ExPEC iucD: iucD: Group n iucD: iucD: eae iucD tsh papC papC: cnf papC tsh cnf Adults 30 6.7 83.3 3.3 6.7 36.7 3.3 0.0 6.7 Sub- 0.0 11 0.0 72.7 0.0 27.3 18.2 0.0 0.0 adults Pups 13 0.0 84.6 0.0 15.4 38.5 0.0 4.3 4.3 Total 54 3.7 81.5 3.7 48.1 35.2 1.9 1.9 5.6

6.4.2 Prevalence of AMR

Anti-microbial resistance to 15 different antimicrobials (Table 6.3) were analysed for 56 isolates from faeces from southern elephant seals and 11 isolates from seawater (Table 6.6). A total of three (5.6%) resistant isolates were found in southern elephant seals for the antibiotics chloramphenicol, nalidixic acid, amoxicillin and cefoxitin (Table 6.6). No AMR was observed in southern elephant seal pups. A single multi- resistant isolate with resistance to two or more classes of antimicrobials (4.0%) was observed in an adult female southern elephant seal (Table 6.6). Further trends were noted for intermediate resistance of isolates from southern elephant seals, particularly for streptomycin, ampicillin and tetracycline (Table A6.3).

Anti-microbial resistance was observed at sites three, four and five at both times and sites one and two during time three. All of the isolates from seawater found to have AMR (73.0%) were also multi-resistant (Table 6.6). Isolates from seawater commonly possessed AMR for the antibiotics amoxicillin, ampicillin, cefoxitin and tetracycline. Multi-resistance patterns commonly consisted of amoxicillin and cefoxitin, which regularly occurred with ampicillin (Table 6.6). Multi-resistance isolates with resistance to three or more antimicrobials occurred at sites three and four of Potter Cove and were found at time three. In one isolate, resistance was found for eight antimicrobials, including all tested cephalosporins, suggesting the presence of extended-spectrum beta-lactamases.

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Table 6.6 Relationship between AMR and site of origin of E. coli isolates. Abbreviations of antimicrobial agents which were used in this study are: (AMC) amoxicillin/cavulanic acid; (TIO) ceftiofur; (CRO) ceftriaxone; (CIP) ciprofloxacin; (AMK) amikacin; (AMP) ampicillin; (FOX) cefoxitin; (GEN) gentamicin; (KAN) kanamycin; (NAL) nalidixic acid; (STR) streptomycin; (SXT) trimethoprim-sulphamethaoxazole; (CHL) chloramphenicol; (FIS) sulfisoxazole; and, (TET) tetracycline. Results are shown for southern elephant seals (ES) grouped by age-class and seawater sampling sites (S) and times (T) in Potter Cove. No. of No. of No. of multi- Total no. of Group susceptible Resistant resistant AMK GEN STR KAN CHL NAL CIP TET FIS SXT AMC AMP FOX CRO TIO isolates

Isolates isolates isolates Adults 25 24 1 (4%) 1 (4%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1.0 0.0 1.0 0.0 0.0 eals

Sub-adults 15 13 2 (13.3%) 0 (0%) 0.0 0.0 0.0 0.0 1.0 1.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0

Southern Southern Pups 16 16 0 (0%) 0 (0%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 elephant s elephant Summary ES 56 53 3 (5.6%) 1(1.7%) 0 0.0 0.0 0.0 0.0 1.0 0.0 0.0 0.0 0.0 1.0 0.0 1.0 0.0 0.0

S1, T1 0 0 0 (0%) 0 (0%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0

S2, T1 0 0 0 (0%) 0 (0%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1.0 0.0 0.0 0.0

S3, T1 1 0 1 (100%) 1 (100%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1.0 0.0 1.0 0.0 0.0

S4, T1 1 0 1 (100%) 1 (100%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1.0 0.0 1.0 0.0 0.0

S5, T1 1 1 1 (100%) 1 (100%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1.0 0.0 1.0 0.0 0.0

S1, T3 1 0 1 (100%) 1 (100%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1.0 0.0 0.0 0.0 1.0 0.0 0.0 0.0 Seawater

S2, T3 2 1 1 (50%) 1 (50%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1.0 0.0 0.0 0.0 1.0 0.0 0.0 0.0

S3, T3 1 0 1 (100%) 1 (100%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 1.0 1.0 1.0 0.0 0.0

S4, T3 2 0 2 (100%) 2 (100%) 0.0 0.0 1.0 1.0 0.0 0.0 0.0 1.0 0.0 0.0 2.0 2.0 2.0 1.0 1.0

S5, T3 2 2 0 (0%) 0 (0%) 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 0.0 Summary 11 4 8 (72.7%) 8 (72.7%) 0.0 0.0 1.0 1.0 0.0 0.0 0.0 3.0 0.0 0.0 6.0 6.0 6.0 1.0 1.0 seawater Total 67 57 11 (16.4%) 9 (13.4%) 0.0 0.0 2.0 2.0 1.0 1.0 0.0 3.0 0.0 0.0 7.0 6.0 7.0 1.0 1.0

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6.5 Discussion

6.5.1 E. coli pathotypes are present in southern elephant seals

The E. coli pathotypes ExPEC and to a lesser extent, EPEC, are common in the faeces of southern elephant seals in Antarctica. The pathotypes ETEC and STEC were not detected. ExPEC pathotypes are thought to be harmless while they are in the intestinal tracts, but can cause serious disease such as neonatal meningitis/sepsis or urinary tract infections [365]. Likewise, EPEC strains can cause severe disease in humans, pets and livestock. They are considered to be opportunistic members of the normal gut microbiota. Amongst wild and stranded pinniped species, gastroenteritis and other gastroenterological-related problems have been frequently diagnosed, although limited studies have shown this to be due to pathogenic E. coli [179,366,367]. In this study, seals with ExPEC and EPEC isolates showed no overt disease symptoms and the presence of these pathotypes, particularly the dominant occurrence of ExPEC (82.0%) across all age-classes of southern elephant seals, suggests it may be a part of the normal gut microbiota in these animals. In Chapter 4, 16S rRNA gene pyrosequencing revealed 25 different OTUs from the genus Escherichia, with an average abundance of four OTUs per seal and an abundance range from zero to ten per individual. Human hosts typically harbour one to three strains of E. coli, but ≥ 10 strains may be recovered at any one time [368,369]. Escherichia coli is unevenly distributed across non-domesticated mammal hosts, but has been isolated from a wide range of mammals, including leopard seals, Hydrurga leptonyx, and hooded seals, Cystophora cristata and Antarctic fur seals, Arctocephalus gazella [Chapter 4,370,371].

6.5.2 Human sewage increase the presence of AMR isolates in seawater

The presence of multi-resistant isolates in seawater proximal to the sewage outfall and surrounding sites in Potter Cove suggests human wastes are the cause of the observed AMR in the environment. AMR was isolated from 72.0% of seawater samples with each of those isolates displaying multi-resistance. One of these isolates was found to be resistant to eight antimicrobials, including all tested cephalosporins, suggesting the presence of extended-spectrum beta-lactamases

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which are enzymes responsible for widespread resistance to antibiotics [372,373]. This is indicative of pressure from antibiotic use and not expected to occur in natural and unexposed environments or bacteria [374]. Multi-resistance has been observed commonly in aquatic systems as a direct result of sewage input [375,376]. In the colder seawater of Antarctica, sewage has been shown to introduce human- associated and multi-resistant bacteria, which increases with the number of people at nearby scientific stations [377-379].

Fewer E. coli pathotypes were recovered from seawater samples compared with southern elephant seals. This may be a result of growth conditions insufficient to revive any potential bacteria from filtered samples. Recovery of E. coli from seawater conditions ranging in temperature from -1 to -2°C requires extra nourishment and lower incubation temperatures to retain colony-forming ability [351]. In this case, it is presumed that E. coli isolation from seawater was underestimated due to the use of a dehydrated medium suitable for temperate water, combined with the 20-25°C incubation temperatures, it is suggested this may not have provided sufficient nutrients to cells [351].

6.5.3 Sewage-contaminated seawater is a possible transmission route to southern elephant seals

Resistance to antimicrobials observed in isolates from southern elephant seals and seawater samples, suggests that human sewage into Potter Cove is a possible route for the transfer of AMR. This transfer pathway may through the direct uptake of isolates with AMR or via virulence factor transfer between similar cells in the gut of seal hosts. Seals are known to take up bacteria with AMR as a result of increased interactions with anthropogenic wastes, as has been identified in northern elephant seals (Mirounga angustirostris) and bottlenose dolphins (Tursiops truncatus) [162]. Transmission would normally occur when animals feed around high nutrient areas, such as those where sewage input is occurring, thereby exposing individuals to novel bacteria with virulence factors. As southern elephant seals are thought to obtain a portion of their gut microbiota from the microbes associated with their prey (Chapter 2), actively feeding on prey that has been feeding in high nutrient sewage waters is another possible pathway of transmission. Antimicrobial resistant or E. coli 105

pathotypes isolates may flourish in the gut environment or as AMR and virulence factors are often located on transmissible units, transmission between species already present in the gut may also occur [380]. The lack of multi-resistant bacteria in seal samples suggests that bacteria originating from human sewage contamination have not colonised the intestinal tract of seals in great abundance. Previous investigations in Antarctica have not observed AMR in wildlife, as was investigated recently in Adélie (Pygoscelis adeliae) and gentoo penguins (Pygoscelis papua) off islands south of 25 de Mayo / King George Island even when multi- resistance was identified in nearby sewage-contaminated seawater [346,347]. This could suggest that sewage contamination influencing wildlife may be highly localised. As the area surrounding 25 de Mayo / King George Island has relatively high human occupation and is visited more regularly by tourists than surrounding areas providing substantially more opportunities for wildlife to come into contact with sewage contaminated areas.

6.6 Conclusion

This study identifies southern elephant seals as a host of E. coli pathotypes ExPEC EPEC and reports, for the first time, the occurrence of AMR in these hosts. This study confirms the likelihood that the input of untreated sewage in Antarctica has the potential to transfer virulence and/or AMR factors from introduced bacteria to indigenous microbiota. AMR, including multi-resistant isolates, are prevalent in the seawater surrounding sewage outfalls from some scientific stations and highlights this as a possible route for gene transfer. The lack of multi-resistant bacteria in seal samples suggests that bacteria originating from human sewage contamination have not yet colonised in the intestinal tract of seals to any significant degree. Given the biodiversity value of the Antarctic environment and its extant fauna, the findings of this study warrant further investigation and management. This has implications for all Antarctic fauna, particularly those surrounding 25 de Mayo / King George Island given its significance as an area of high biodiversity and sensitivity. The results of this study warrant further attention and provide strong evidence for the restriction of dumping human sewage in Antarctica untreated.

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Chapter 7

The gut microbiota of Antarctic seals

7.1 Introduction

The primary aim of this thesis was to characterise the ecology of the gut microbiota of wild marine mammals by focusing on influences on Antarctic seal species. This research is, to my knowledge, the first study to investigate the ecology of the gut microbiota of seals from the Antarctic and Southern Ocean Region. Further, it is one of few studies investigating the mammalian gut microbiota of non-human hosts living in wild ecosystems with substantial replication. Interest in the origin, taxonomic composition and function of microbes inhabiting gut environments has increased markedly in recent years. A number of studies have sought to identify the gut microbiota of fish [e.g. 303,381], birds [e.g. 40,68], humans [e.g. 20,382], non-human terrestrial mammals [e.g. 23,242] and also marine mammals [e.g. 44,45]. These studies are beginning to provide an understanding of the complex and essential relationship that exists between microbial communities and their hosts. In mammals, the gut microbial community is known to influence the ecology of the host. Studies have identified microbially mediated mechanisms that directly affect host health and the metabolic capabilities of the host [269,383]. However, linking the composition of the gut microbiota to host functions is hampered by a lack of knowledge concerning the baseline community present under natural conditions. For wild mammals endemic to regions such as Antarctica, anthropogenic impacts and alterations in local climatic conditions are resulting in large-scale shifts in resource availability, often forcing them to adapt to new scenarios, including changes in diet.

Therefore, this thesis was completed to provide greater understanding of the ecology of these hosts, and more broadly to mammals and their associated bacteria. This discussion outlines the general conclusions of the research in this thesis and how this has built on previous knowledge.

7.2 A distinct and conserved phocid seal gut microbiota

The gut microbiota of Antarctic southern elephant seals and leopard seals is 107

dominated by few phyla. Although a total of 19 phyla were observed, 98.2 ± 0.5% of OTUs identified belonged to one of four phyla: Firmicutes, Bacteroidetes, Fusobacteria and Proteobacteria (Chapter 4). In line with other studies investigating the gut environment of mammals, high levels of richness were observed at the strain or species level with a total of 6,159 OTUs observed in the 40 seal hosts (Chapter 4). Broadly, these patterns are similar to those observed in the gut microbiota of other mammals, including humans [e.g. 13,221,231].

The gut microbiota of Antarctic seals appears to be driven strongly by two main factors: host phylogeny and diet. Both of these factors have been identified as drivers of the gut microbiota of mammals [23,38], yet there has been discussion over which of these factors is a stronger driver. This research suggests there is interplay between both factors. Core OTUs are shared between phocid seals regardless of diet or geographical differences, suggesting they have been maintained in each host species as a result of co-evolution (Chapters 4 and 5). Maintenance of this core gut microbiota is through vertical transmission from mother to pup and, in later life, through social interactions (Chapter 3 and 4). It is suggested that the core has been conserved in these hosts as they serve a function specific to the host that is broadly beneficial, such as regulating immune function or aiding development of the gut. The diet influences the gut microbiota of these hosts in two ways: via the direct introduction of microbes associated with their prey (Chapter 2) and also via the nutritional composition of dietary items which, with the gut morphology, controls the physicochemical habitat suitable to particular microbial species (Chapters 3 and 4).

7.3 Antarctic phocids have a close relationship with the Fusobacteria

Members of the phyla Firmicutes and Bacteroidetes are the most commonly associated phyla within the gut of mammals [13,23] and our findings expand this association to seal hosts. However, the phylum Fusobacteria was identified as a prominent member of the gut microbiota of Antarctic seals, occurring in higher abundances in these hosts compared with other mammals (Chapters 4 and 5). Similarly, the phylum Proteobacteria has been observed in the gut of mammals [36], but contributes a greater proportion to the community composition of seals than to any other host previously examined. 108

The relatively high abundance of the phylum Fusobacteria in seal hosts is of interest. Previous studies have recorded much lower abundances of members of the phylum Fusobacteria in the gut of mammal hosts with the exception of the Arctic phocid seals and members of the Canidae family (dogs and wolves) [Chapter 5; 32,261,302,303,384,385-388]. Fusobacteria are part of the core microbiota of phocid seals and their presence may indicate a tightly coupled relationship with an important functional role such as host immune response. In light of the shared immunogenetic responses between dogs and seals [324,325], the presence and relatively high abundance of members of the Fusobacteria in these hosts supports the hypothesis of evolutionary conservation of microbial-host associations.

7.4 Introductions into the gut from prey but few from the external environment

Previous work has suggested that the gut microbiota of carnivorous mammals may be sourced primarily from ‘free-living’ communities [8]. However, herein I have identified that bacteria are more likely to be introduced from the gut of prey items or host-associated sources (Chapters 2 and 4). By estimating the abundance of free- living OTUs based on the environmental associations of the closest relatives in GenBank, I suggest free-living bacteria make only a minimal contribution to the gut microbiota of Antarctic seal hosts (Table 7.1). I suggest that OTUs putatively identified as coming from external sources may be transient members of the community that are unlikely to establish themselves in the internal gut environment (for example, representatives from the Prochlorococcus and SAR11 marine lineages). Members of the gut microbial community is thought to require forms of ‘stress resistance’ in order to transition between hosts via a largely dry, saline and toxic ‘ex-host’ environment [2]. However, I did identify the potential for AMR and E. coli pathotypes to transition from the water column into the guts of southern elephant seals (Chapter 6). Interestingly, horizontal gene transfer from free-living organisms into the genomes of existing members of the gut microbiota has been identified previously [8,389]. Thus horizontal gene transfer, rather than the establishment of novel microbes within the gut environment, may play an important role in the ability of the gut microbiome (the full complement of factors in the gut microbial community) to acquire traits required to efficiently transition to a novel diet.

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7.5 Captivity has a strong influence on the gut microbiota of mammals

It has been reported previously that the gut microbiota of captive mammals is representative of their wild con-specifics [23,38]. However, my study has identified that captivity appears to have a significant impact on the gut microbial composition in leopard seals (Chapters 3 and 4). Wild systems are inherently more variable and their inhabitants are subject to a wide range of different exposures compared to captive or laboratory-reared individuals. For leopard seal hosts sampled here, dietary differences appear to be the primary influence on the gut microbiota, yet, it is still somewhat unclear as to how much the captive environment and antibiotic use has altered microbial functions in these hosts and if there are any ill effects of this shift.

7.6 Future directions

Future investigations into the gut microbiota of these hosts would help to tease apart functionality of the gut microbiota and quantify the influencing factors of phylogeny and diet. One of the primary difficulties with working with large vertebrate mammal hosts is that relationships must be inferred, as very little opportunity exists to experimentally test hypotheses. I suggest that monitoring the presence of OTUs, particularly those identified as ‘core’ members, from one host and matching this with immune function would provide insight. Functional capacity of the gut microbiota of these hosts would also prove useful and insightful. The evolutionary conservation of the gut microbiota could be addressed by further analysing the gut microbiota of a number of other phylogenetically related carnivore hosts with different diets from wild locations. Further quantification and understanding of the transmission of gut microbiota from prey to host may have the potential to remedy individuals in captivity during ill health, or post-antibiotic treatments. Finally, the ability of potentially novel bacterial members or potentially damaging genetic material to move from human sewage contamination of one host to the intestines of another in Antarctica is cause for concern in endemic populations such as this and requires further attention.

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APPENDIX 1

Table A1.1 Characteristics of Antarctic seals, fish and seawater samples. Sample ID refers to the identification number, location (Loc.) refers to the region inhabited by the host when sampled. Table data are as follows: (ES) southern elephant seal; (LS) leopard seal; (M) male; (F) female; (KGI) 25 de Mayo / King George Island, South Shetland Islands, western Antarctic; (DC) Danco Coast, western Antarctic; (TZ) Taronga Zoo, Sydney, NSW, Australia; (S1, S2...) sampling sites for seawater; and (rep.) replicate samples. Sample use in: (*) Chapter 2 for bacterial community stability analysis; (‡) Chapter 2 for community representatives along the intestines; (†) Chapter 2 for transfer of fish to leopard seal gut; (^) Chapter 3; (**) Chapters 4 and 5.

Mother Sample Collection Sample type Host Sex Age-class -pup Loc. ID date pair *105 Faeces 9-Mar-10 LS M Sub-adult TZ *107 Faeces 2-Mar-10 LS M Sub-adult TZ *108 Faeces 2-Mar-10 LS F Sub-adult TZ *109 Faeces 13-Mar-10 LS M Sub-adult TZ *111 Faeces 13-Mar-10 LS F Sub-adult TZ *115 Faeces 6-Mar-10 LS M Sub-adult TZ *118 Faeces 27-Mar-10 LS M Sub-adult TZ *119 Faeces 10-Apr-10 LS M Sub-adult TZ *120 Faeces 17-Apr-10 LS M Sub-adult TZ *121 Faeces 25-Apr-10 LS M Sub-adult TZ *†122 Faeces 8-May-10 LS M Sub-adult TZ *†123 Faeces 26-May-10 LS M Sub-adult TZ *124 Faeces 6-Mar-10 LS F Sub-adult TZ *125 Faeces 9-Mar-10 LS F Sub-adult TZ *127 Faeces 27-Mar-10 LS F Sub-adult TZ *129 Faeces 10-Apr-10 LS F Sub-adult TZ *130 Faeces 17-Apr-10 LS F Sub-adult TZ *131 Faeces 25-Apr-10 LS F Sub-adult TZ *133 Faeces 8-May-10 LS F Sub-adult TZ †134 Faeces 25-May-10 LS F Sub-adult TZ *135 Faeces 26-May-10 LS F Sub-adult TZ †136 Faeces 1-Jun-10 LS F Sub-adult TZ †139 Intestines 20-May-10 Whiting - - - - †147 Intestines 20-May-10 Herring - - - - ‡84 Duodenum 12-Jul-09 LS M Sub-adult - TZ ‡83 Jejunum 12-Jul-09 LS M Sub-adult - TZ ‡82 Ileum 12-Jul-09 LS M Sub-adult - TZ ‡86 Colon 12-Jul-09 LS M Sub-adult - TZ ‡85 Rectum 12-Jul-09 LS M Sub-adult - TZ ‡79 Faeces 12-Jul-09 LS M Sub-adult - TZ **01 Swab 27-Nov-08 ES M Sub-adult - KGI ^02 Swab 05-Dec-08 ES M Sub-adult - KGI ^**04 Swab 24-Nov-08 ES F Sub-adult - KGI ^**10 Swab 07-Oct-08 ES M Adult - KGI ^**11 Swab 20-Oct-08 ES F Adult Mum A KGI ^**12 Swab 15-Oct-08 ES F Adult Mum B KGI 111

^13 Swab 29-Oct-08 ES M Pup - KGI ^**14 Swab 03-Nov-08 ES F Adult Mum C KGI ^**15 Swab 07-Nov-08 ES F Adult Mum D KGI ^**16 Swab 09-Oct-08 ES M Adult - KGI ^**17 Swab 15-Oct-08 ES F Adult Mum E KGI ^18 Swab 22-Oct-08 ES F Adult Mum F KGI ^**20 Swab 03-Nov-08 ES M Pup Pup C KGI ^**21 Swab 28-Nov-08 ES F Sub-adult - KGI ^**22 Swab 15-Oct-08 ES F Pup Pup B KGI ^23 Swab 22-Oct-08 ES M Pup Pup F KGI ^**24 Swab 22-Oct-08 ES M Pup Pup G KGI ^**25 Swab 11-Nov-08 ES M Adult - KGI ^**26 Swab 28-Nov-08 ES M Sub-adult - KGI ^27 Swab 02-Dec-08 ES M Sub-adult - KGI ^**28 Swab 09-Oct-08 ES M Adult - KGI ^**29 Swab 14-Oct-08 ES F Adult - KGI ^**30 Swab 15-Oct-08 ES M Pup Pup E KGI ^**31 Swab 20-Oct-08 ES M Pup Pup A KGI ^32 Swab 23-Oct-08 ES M Pup Pup H KGI ^**33 Swab 24-Nov-08 ES M Sub-adult - KGI ^**34 Swab 22-Oct-08 ES F Adult Mum G KGI ^35 Swab 23-Oct-08 ES F Adult Mum H KGI ^36 Swab 07-Nov-08 ES M Pup Pup J KGI ^**37 Swab 27-Nov-08 ES M Sub-adult - KGI ^38 Swab 28-Nov-08 ES M Sub-adult - KGI ^39 Swab 02-Dec-08 ES M Sub-adult - KGI ^**40 Swab 07-Nov-08 ES F Pup Pup D KGI ^41 Swab 07-Nov-08 ES F Adult Mum J KGI ^42 Swab 18-Nov-08 ES M Adult - KGI ^43 Swab 22-Nov-08 ES M Sub-adult - KGI ^44 Swab 29-Nov-08 ES M Sub-adult - KGI ^**45 Swab 30-Nov-08 ES F Sub-adult - KGI ^46 Swab 02-Dec-08 ES M Sub-adult - KGI ^47 Swab 06-Dec-08 ES M Sub-adult - KGI ^48 Swab 08-Dec-08 ES M Sub-adult - KGI ^**49 Swab 11-Feb-09 LS F Adult - DC ^**50 Swab 19-Feb-09 LS M Adult - DC ^**51 Swab 20-Feb-09 LS M Adult - DC ^**52 Swab 20-Feb-09 LS M Adult - DC ^**53 Swab 21-Feb-09 LS M Adult - DC ^**54 Swab 26-Feb-09 LS F Adult - DC ^**56 Swab 18-Feb-09 LS F Adult - DC ^**57 Swab 21-Feb-09 LS M Adult - DC ^**58 Swab 26-Feb-09 LS F Adult - DC ^**60 Swab 27-Feb-09 LS F Adult - DC ^61 Swab 10-Feb-09 LS M Adult - DC ^62 Swab 11-Feb-09 LS M Adult - DC ^**63 Swab 12-Feb-09 LS M Adult - DC ^64 Swab 16-Feb-09 LS M Adult - DC ^65 Swab 16-Feb-09 LS M Adult - DC ^66 Swab 18-Feb-09 LS M Adult - DC

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^**67 Swab 18-Feb-08 LS M Adult - DC ^68 Swab 01-Feb-08 LS F Adult - DC ^69 Swab 02-Feb-08 LS M Adult - DC ^70 Swab 18-Feb-08 LS M Adult - DC ^71 Faeces 25-Feb-10 LS M Sub-adult TZ ^**72 Faeces 25-Feb-10 LS F Sub-adult TZ ^**73 Faeces 19-Jan-10 LS F Sub-adult TZ ^75 Faeces 23-Feb-10 LS F Sub-adult TZ **76 Faeces 23-Jan-10 LS M Sub-adult TZ ^**77 Faeces 12-Jan-10 LS M Sub-adult TZ ^78 Faeces 12-Sep-09 LS M Sub-adult TZ Sample Collection Sample type Loc. ID date 90 Seawater 10-Nov-08 KGI S1 91 Seawater 10-Nov-08 KGI S2 92 Seawater 10-Nov-08 KGI S3 93 Seawater 10-Nov-08 KGI S4 94 Seawater 10-Nov-08 KGI S5 95 Seawater 2-Dec-08 KGI S1 Rep. A 96 Seawater 2-Dec-08 KGI S1 Rep. B 97 Seawater 2-Dec-08 KGI S2 Rep. A 98 Seawater 2-Dec-08 KGI S2 Rep. B 99 Seawater 2-Dec-08 KGI S3 Rep. A 100 Seawater 2-Dec-08 KGI S3 Rep. B 101 Seawater 2-Dec-08 KGI S4 Rep. A 102 Seawater 2-Dec-08 KGI S4 Rep. B 103 Seawater 2-Dec-08 KGI S5 Rep. A 104 Seawater 2-Dec-08 KGI S5 Rep. B

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Table A1.2 Clinical notes on the status of the female captive leopard seal during sample collection. Date Clinical notes Administered 3-Jan-10 - Two bouts of diarrhoea 4-Jan-10 - Anorexia - Began excenel (cetiofur hydrochloride) - Decreased alertness antibiotic treatment - Not eating - Fed electrolytes and sugars: vytrate 5-Jan-10 - Anorexia - Excenel (cetiofur hydrochloride) antibiotic - Decreased alertness treatment - Not eating 6-Jan-10 - Anorexia - Fed electrolytes and sugars: vytrate - Decreased alertness - Not eating 7-Jan-10 - Anorexia - Stop excenel treatment - Decreased alertness - Not eating 8-Jan-10 - Anorexia - Fed electrolytes and sugars: vytrate - Decreased alertness - Began rilexine (cephalaxin) antibiotic - Not eating treatment 15-Jan-10 - Anorexia - Stop rilexine treatment - Decreased alertness - Not eating 19-Jan-10 - Anorexia - Decreased alertness - Not eating - Sample collection (ID 73) 23-Jan-10 - Anorexia - Decreased alertness - Not eating - Sample collection (ID 75) 1-Feb-10 - Eye problem causing opacity - Increasing food intake slowly - Weight loss - Daily food intake ~3.5 kg 15-Feb-10 - Eye problems continuing - Began orbenin eye ointment, an antibiotic - Weight loss agent 17-Feb-10 - Eye problems - Stop orbenin treatment - Weight loss 25-Feb-10 - Sample collection (ID 72)

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APPENDIX 2

Table A2.1 Characteristic OTUs in the gut microbiota of captive leopard seals grouped by sex and month. Foremost ten characteristic operational taxonomic units (OTUs) present in the gut microbiota of female and male captive leopard seals as identified by SIMPER analysis. OTU Average abundance Contribution to total (%) 471.8 3.66 12.76 473.8 2.88 8.44 709.8 2.42 5.63 685.8 1.48 5.27 715.8 1.65 3.55 Female 623.8 1.54 2.93 625.8 1.75 2.66 479.8 0.73 2.54 469.8 1.11 2.43 531.8 1.11 2.15 709.8 3.35 10.44 471.8 3.03 9.31 623.8 2.32 6.48 715.8 2.27 6.08 685.8 1.7 5.88 Female March 629.8 2.17 5.25 473.8 2.25 4.8 843.8 1.39 3.07 531.8 1.56 2.81 721.8 1.48 2.78 471.8 3.24 11.53 473.8 3.57 11.18 491.8 2.18 5.22 709.8 2.54 4.29 475.8 1.99 4.27 Female April 685.8 1.31 4.23 469.8 1.19 3.28 625.8 2.09 2.3 531.8 0.96 1.99 623.8 1.52 1.93 471.8 5.47 22.7 625.8 4.05 14.31 473.8 3.25 8.57 685.8 1.25 3.69 479.8 0.79 2.94 Female May 539.8 1.97 2.36 717.8 1.32 2.25 407.8 0.53 2.23 387.8 0.6 2.19 377.8 0.57 2.15 473.8 3.33 9.41 Male 471.8 2.83 8.58 685.8 2.49 6.69 115

709.8 2.39 4.96 623.8 1.33 2.69 479.8 0.84 2.34 469.8 1.22 2.33 629.8 1.57 2.15 531.8 0.97 2.09 715.8 1.33 1.95 471.8 2.73 7.72 473.8 2.91 6.32 685.8 2.49 6.03 709.8 2.63 5.38 469.8 1.34 3.86 Male March 629.8 2.13 3.33 531.8 1.37 3.28 623.8 1.58 3.06 715.8 1.55 2.85 505.8 1.62 2.69 473.8 3.98 11.78 471.8 2.74 6.8 709.8 3.06 6.55 475.8 2.28 6.16 623.8 1.45 4.23 Male April 685.8 2.18 4.14 479.8 1.07 2.6 569.8 1.21 2.37 577.8 1.04 2.24 855.8 0.7 1.9 473.8 3.8 13.75 625.8 3.93 11.4 471.8 3.31 9.66 685.8 2.98 9.63 717.8 1.8 5.2 Male May 629.8 1.96 3.78 641.8 1.12 3.68 569.8 0.98 3.32 595.8 0.88 3.26 577.8 0.85 3.02

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APPENDIX 3

Table A3.1 Characteristic OTUs in the gut microbiota of seal hosts grouped by species, age-class, sex and location. Foremost ten characteristic operational taxonomic units (OTUs) in the gut microbiota of hosts identified using SIMPER analysis. Average Contribution Groups OTU abundance to total (%) 629.3 1.53 5.86 523.3 1.24 5.16 545.3 1.17 4.85 457.3 1.02 4.44 567.3 1.12 3.55 Southern elephant seals 721.3 0.86 2.92 749.3 1.00 2.87 399.3 0.98 2.56 493.3 0.90 2.53 641.3 0.80 2.52 457.3 1.72 9.55 545.3 1.87 7.36 523.3 1.53 5.84 399.3 1.65 4.53 513.3 1.32 3.95 Southern elephant seal adults 821.3 1.10 3.94 641.3 1.06 3.50 633.3 1.17 3.22 749.3 0.89 2.73 629.3 0.90 2.69 457.3 1.68 7.96 399.3 2.28 7.20 633.3 1.75 6.70 545.3 1.48 5.17 523.3 1.39 4.87 Southern elephant seal adult females 641.3 1.36 4.34 505.3 1.16 4.03 821.3 1.20 3.86 485.3 0.73 3.36 599.3 1.16 3.00 513.3 2.80 12.14 545.3 2.65 10.13 457.3 1.81 9.07 629.3 1.82 8.20 721.3 1.16 6.33 Southern elephant seal adult males 523.3 1.81 4.89 833.3 1.81 4.08 687.3 0.82 3.12 451.3 0.68 2.67 749.3 0.80 2.49 523.3 1.40 5.07 Southern elephant seal sub-adults 749.3 1.49 4.36 625.3 1.10 4.18 117

545.3 0.88 3.90 853.3 1.19 3.84 521.3 1.05 3.46 567.3 0.85 3.11 457.3 0.83 2.76 451.3 0.70 2.72 587.3 0.90 2.39 629.3 3.51 17.15 567.3 2.99 14.74 493.3 2.37 9.72 721.3 1.91 7.69 525.3 1.18 5.24 Southern elephant seal pups 655.3 0.61 3.22 515.3 0.82 2.64 377.3 0.86 2.28 549.3 1.16 2.26 559.3 0.63 2.18 523.3 2.48 14.48 655.3 1.55 8.8 521.3 1.64 7.16 525.3 1.08 5.19 587.3 1.42 4.48 Wild leopard seals 581.3 0.70 3.13 585.3 1.07 2.9 629.3 0.76 2.84 545.3 0.64 2.66 637.3 0.52 2.17 587.3 2.07 8.70 521.3 1.93 8.56 523.3 2.14 8.52 655.3 1.34 7.03 603.3 0.96 5.62 Wild leopard seal females 525.3 1.01 5.46 629.3 0.84 5.29 625.3 1.11 4.14 637.3 0.75 3.88 475.3 0.97 3.31 523.3 2.66 16.86 655.3 1.65 9.41 585.3 1.54 6.09 521.3 1.49 5.87 525.3 1.11 4.8 Wild leopard seal males 581.3 0.79 3.76 897.3 1.06 2.99 559.3 0.87 2.62 587.3 1.09 2.48 545.3 0.6 2.19 481.3 3.15 13.73 751.3 2.84 12.62 Captive leopard seals 629.3 2.08 11.17 479.3 2.44 9.41

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511.3 1.56 8.5 477.3 1.68 5.15 533.3 1.3 4.53 721.3 1.13 4.42 493.3 1.08 2.85 857.3 0.46 1.85 751.3 2.66 12.06 629.3 2.17 10.31 479.3 2.23 10.19 481.3 2.43 10.03 711.3 2.37 7.37 Captive leopard seal female 511.3 1.3 7.26 715.3 1.72 6.66 625.3 1.86 6.09 477.3 1.77 5.92 847.3 1.7 5.12 481.3 4.29 17.41 751.3 3.76 12.93 479.3 3.21 8.42 629.3 1.89 5.83 511.3 1.73 5.49 Captive leopard seal male 721.3 1.55 5.05 533.3 2.01 4.88 457.3 1.25 4.46 777.3 1.62 4.34 843.3 1.06 2.88

Table A3.2 Shared OTUs between most southern elephant seal mother-pup pairs. Average relative abundance of operational taxonomic units (OTUs) ± standard error (SE) shared between each mother and pup within a pair in most (more than half) of the mother- pup pairs. In the case of (*) OTUs were shared between more than two thirds of mother-pup pairs. Average relative abundance / OTU individual ± SE (%) Mothers Pups 523.3* 3.4 ± 1.5 0.5 ± 0.1 399.3 9.6 ± 5.1 1.0 ± 0.5 655.3 0.2 ± 0.7 0.5 ± 0.1 641.3 3.7 ± 1.5 1.3 ± 0.8 721.3 0.3 ± 0.1 5.7 ± 2.5 629.3 0.6 ± 0.3 16.4 ± 5.8

119

Figure A3.1 Relationship between characteristic OTUs and mass of southern elephant seal adults. Linear regression models the strength of the relationship between the mass of southern elephant seal adults with the abundance of: (A) OTU399.3; (B) OTU505.3; (C) OTU513.3; (D) OTU545.3; (E) OTU629.3; (F) OTU 687.3; (G) OTU641.3; (H) OTU633.3; (I) OTU721.3; (J) OTU833.3.

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APPENDIX 4

Table A4.1 Abundance of representative phyla and classes in the gut microbiota of wild and captive Antarctic seals. Average relative abundance of each phylum and class or division representated in the seal gut microbiota ± standard error (SE). Average relative Average relative Phylum abundance ± SE Class / division abundance ± SE / / individual (%) individual (%) Clostridia 35.9 ± 3.8 Bacilli 5.1 ± 1.9 Firmicutes 41.5 ± 3.9 Erysipelotrichi 0.3 ± 0.1 Unnamed member of the 0.5 ± 0.1 phylum Firmicutes Fusobacteria 25.6 ± 3.8 Fusobacteria 25.6 ± 3.8 Gammaproteobacteria 14.4 ± 3.2 Epsilonproteobacteria 1.3 ± 0.3 Proteobacteria 17.0 ± 3.2 Betaproteobacteria 1.1 ± 0.2 Alphaproteobacteria 0.2 ± 0.1 Deltaproteobacteria 0.1 ± < 0.1 Bacteroidia 13.4 ± 1.9 Unnamed member of the 0.3 ± 0.3 Bacteroidetes 14.1 ± 1.9 phylum Bacteroidetes Flavobacteria 0.2 ± 0.1 Sphingobacteria 0.1 ± 0.1 Actinobacteria 0.8 ± 0.1 Actinobacteria 0.8 ± 0.2 Tenericutes 0.7 ± 0.5 Mollicutes 0.7 ± 0.5 Spirochaetes 0.1 ± 0.1 Spirochaetes 0.1 ± 0.1 Unnamed member of the SR1 0.1 ± < 0.1 0.1 ± < 0.1 division SR1 Cyanobacteria < 0.1 ± < 0.1 Cyanobacteria < 0.1 ± < 0.1 Acidobacteria < 0.1 ± < 0.1 Acidobacteria Gp1 < 0.1 ± < 0.1 Unnamed member of the TM7 < 0.1 ± < 0.1 < 0.1 ± < 0.1 division TM7 Unnamed member of Unnamed member of the < 0.1 ± < 0.1 < 0.1 ± < 0.1 the kingdom Bacteria kingdom Bacteria Deferribacteres < 0.1 ± < 0.1 Deferribacteres < 0.1 ± < 0.1 Synergistetes < 0.1 ± < 0.1 Synergistia < 0.1 ± < 0.1 Deinococcus-thermus < 0.1 ± < 0.1 Deinococci < 0.1 ± < 0.1 Chloroflexi < 0.1 ± < 0.1 Chloroflexi < 0.1 ± < 0.1 Planctomycetes < 0.1 ± < 0.1 Planctomycetacia < 0.1 ± < 0.1 Gemmatimonadetes < 0.1 ± < 0.1 Gemmatimonadetes < 0.1 ± < 0.1 Verrucomicrobia < 0.1 ± < 0.1 Verrucomicrobiae < 0.1 ± < 0.1 Chlorobi < 0.1 ± < 0.1 Chlorobia < 0.1 ± < 0.1

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Table A4.2 Characteristic OTUs in the gut microbiota and their taxonomy grouped by species, age-class, sex and location. Foremost ten characteristic operational taxonomic units (OTUs) in the gut microbiota of hosts identified using SIMPER analysis. Average Contribution Group OTU Phylum Genus abundance to total (%) OTU1 Fusobacteria Ilyobacter 15.74 5.22 OTU2 Firmicutes Subdoligranulum 11.74 3.12 OTU3 Fusobacteria Fusobacterium 10.11 2.9 Southern OTU8 Fusobacteria Psychrilyobacter 7.32 2.54 elephant OTU13 Fusobacteria Ilyobacter 7.2 2.37 seal adult OTU10 Firmicutes Clostridiales XI 5.56 2.21 and sub- OTU27 Firmicutes Sporobacter 6.46 2.18 adults OTU15 Bacteroidetes Petrimonas 8.29 2.16 OTU63 Firmicutes Sporobacter 4.2 1.43 OTU35 Bacteroidetes Petrimonas 4.48 1.34 OTU1 Fusobacteria Ilyobacter 10.01 2.72 OTU10 Firmicutes Clostridiales XI 6.37 2.46 Southern OTU2 Firmicutes Subdoligranulum 12.69 2.34 elephant OTU15 Bacteroidetes Petrimonas 9.02 2.34 seal OTU20 Firmicutes Tissierella 9.4 1.91 female OTU3 Fusobacteria Fusobacterium 7.51 1.9 adult and OTU27 Firmicutes Sporobacter 6.43 1.84 sub-adults OTU8 Fusobacteria Psychrilyobacter 5.07 1.49 OTU35 Bacteroidetes Petrimonas 5.19 1.38 OTU60 Firmicutes Sporanaerobacter 4.45 1.35 OTU1 Fusobacteria Ilyobacter 22.91 7.22 OTU13 Fusobacteria Ilyobacter 10.35 3.37 OTU3 Fusobacteria Fusobacterium 13.36 3.27 OTU8 Fusobacteria Psychrilyobacter 10.12 3.19 Southern OTU2 Firmicutes Subdoligranulum 10.54 2.98 elephant OTU27 Firmicutes Sporobacter 6.49 1.83 seal male Unnamed member adult and OTU52 Bacteroidetes of the family 5.81 1.78 sub-adults Prevotellaceae OTU16 Proteobacteria Kushneria 9.91 1.59 OTU37 Proteobacteria Psychrobacter 7.32 1.55 OTU55 Bacteroidetes Odoribacter 5.14 1.52 OTU1 Fusobacteria Ilyobacter 35.42 14.16 OTU3 Fusobacteria Fusobacterium 20.71 6.91 OTU8 Fusobacteria Psychrilyobacter 15.99 6.51 OTU13 Fusobacteria Ilyobacter 15.91 6.29 OTU11 Bacteroidetes Bacteroides 17.5 6.02 Unnamed member Southern of the family elephant Firmicutes OTU75 Peptostreptococca- 7.99 2.79 seal pups ceae Unnamed member of the family Firmicutes OTU287 Peptostreptococca- 5.61 1.94 ceae OTU66 Fusobacteria Ilyobacter 4.56 1.67 122

OTU388 Fusobacteria Fusobacterium 4.76 1.64 OTU53 Firmicutes Butyricicoccus 7.47 1.6 OTU17 Firmicutes Weissella 9.43 6.44 Unnamed member OTU4 Proteobacteria of the order 15.35 5.47 Oceanospirillales OTU14 Proteobacteria Psychrobacter 8.95 3.92 Wild OTU10 Firmicutes Clostridiales XI 10.7 3.88 leopard OTU18 Proteobacteria Moraxella 9.04 3.36 seals OTU88 Firmicutes Leuconostoc 4.86 3.13 OTU19 Proteobacteria Psychrobacter 7.1 2.32 OTU140 Firmicutes Lactococcus 3.24 2.15 OTU94 Firmicutes Lactococcus 3.18 1.83 OTU90 Firmicutes Citrobacter 3.39 1.71 OTU14 Proteobacteria Psychrobacter 13.05 6.65 Unnamed member OTU4 Proteobacteria of the order 14.91 4.36 Oceanospirillales Wild OTU17 Firmicutes Weissella 6.83 4.34 leopard OTU29 Firmicutes Tissierella 9.37 4.18 seal OTU15 Bacteroidetes Petrimonas 5.98 2.91 females OTU40 Firmicutes Soehngenia 6.12 2.67 OTU18 Proteobacteria Moraxella 7.98 2.36 OTU64 Bacteroidetes Petrimonas 4.96 2.27 OTU88 Firmicutes Leuconostoc 3.55 2.13 OTU213 Firmicutes Helcococcus 3.57 2.04 OTU17 Firmicutes Weissella 12.02 8.08 OTU10 Firmicutes Clostridiales XI 14.16 6.98 Unnamed member OTU4 Proteobacteria of the order 15.78 4.92 Wild Oceanospirillales leopard OTU88 Firmicutes Leuconostoc 6.17 3.83 seal OTU19 Proteobacteria Psychrobacter 11.57 3.81 males OTU18 Proteobacteria Moraxella 10.11 3.27 OTU140 Firmicutes Lactococcus 3.77 2.35 OTU90 Proteobacteria Citrobacter 4.48 2.23 OTU253 Firmicutes Lactococcus 2.56 2.15 OTU205 Firmicutes Streptococcus 3.14 1.91 OTU12 Firmicutes Subdoligranulum 28.24 6.18 OTU34 Firmicutes Sporobacter 11.05 3.12 OTU30 Firmicutes Sporobacter 12.12 2.52 OTU58 Fusobacteria Fusobacterium 11.29 2.27 Captive OTU13 Fusobacteria Ilyobacter 8.76 1.89 leopard OTU137 Firmicutes Sporobacter 6 1.69 seals OTU70 Firmicutes Oscillibacter 8.16 1.58 OTU191 Firmicutes Oscillibacter 7.14 1.52 OTU3 Fusobacteria Fusobacterium 10.99 1.49 OTU32 Firmicutes Sporobacter 10.57 1.43 Captive OTU58 Fusobacteria Fusobacterium 16.69 8.46 leopard OTU34 Firmicutes Sporobacter 8.6 5.07 seal OTU12 Firmicutes Subdoligranulum 19.33 3.7 female OTU32 Firmicutes Sporobacter 8.21 3.64 123

OTU137 Firmicutes Sporobacter 4.58 2.81 OTU115 Firmicutes Oscillibacter 6.51 2.41 OTU191 Firmicutes Oscillibacter 3.6 2.23 OTU30 Firmicutes Sporobacter 9.12 2.13 OTU5 Fusobacteria Cetobacterium 32.44 1.82 OTU179 Firmicutes Sporobacter 3.59 1.82 OTU12 Firmicutes Subdoligranulum 37.14 6.96 OTU34 Firmicutes Sporobacter 13.49 2.56 OTU70 Firmicutes Oscillibacter 12.55 2.31 OTU30 Firmicutes Sporobacter 15.11 2.18 Captive OTU191 Firmicutes Oscillibacter 10.68 1.94 leopard OTU50 Fusobacteria Fusobacterium 10.58 1.51 seal male OTU13 Fusobacteria Ilyobacter 9.95 1.48 OTU135 Firmicutes Coprobacillus 7.89 1.47 OTU137 Firmicutes Sporobacter 7.41 1.44 OTU144 Firmicutes Oscillibacter 8.62 1.4

Table A4.3 Gut microbial richness of captive leopard seals over time. Richness of gut microbiota estimated by Chao 1 [287] of the female and male captive leopard seals at each sampling time. Time of illness is marked (*). Details of sample collection are listed in Table A1.1. Sample Chao 1 Seal sex Date number mean *LS73 Female 19-Jan-10 544 LS72 Female 25-Feb-10 1077 LS77 Male 12-Jan-10 1171 LS76 Male 23-Feb-10 1571

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Table A4.4 Shared OTUs in the gut microbiota of most southern elephant seal adults and sub-adults. Average relative abundance of operational taxonomic units (OTUs) shared in the gut microbiota of most (more than half) individual southern elephant seal adults and sub- adults ± standard error (SE). OTUs occurring in the gut microbiota of () all individuals and (*) more than three quarters of individuals are marked. Average relative OTU Phylum Genus abundance / individual ± SE (%) Unnamed member from the OTU10 Firmicutes 1.14 ± 0.28 family Clostridiales XI OTU8* Fusobacteria Psychrilyobacter 1.94 ± 0.62 OTU24* Proteobacteria Campylobacter 0.56 ± 0.18 OTU17* Firmicutes Weissella 0.23 ± 0.09 OTU1* Fusobacteria Ilyobacter 8.21 ± 2.45 OTU15* Bacteroidetes Petrimonas 4.49 ± 2.06 OTU3* Fusobacteria Fusobacterium 4.37 ± 1.77 OTU13* Fusobacteria Ilyobacter 1.66 ± 0.48 OTU27* Firmicutes Sporobacter 1.46 ± 0.44 OTU63* Firmicutes Sporobacter 0.72 ± 0.24 OTU2* Firmicutes Subdoligranulum 4.96 ± 2.17 OTU35* Bacteroidetes Petrimonas 1.01 ± 0.37 OTU33* Bacteroidetes Alistipes 0.63 ± 0.44 OTU263* Proteobacteria Campylobacter 0.28 ± 0.08 Unnamed member from the OTU171* Firmicutes 0.19 ± 0.06 family Ruminococcaceae Unnamed member from the OTU52* Bacteroidetes 0.85 ± 0.37 family Prevotellaceae OTU151* Firmicutes Anaerococcus 0.29 ± 0.10 OTU454* Firmicutes Roseburia 0.07 ± 0.03 OTU55 Bacteroidetes Odoribacter 0.50 ± 0.26 OTU167 Firmicutes Anaerococcus 0.15 ± 0.08 Unnamed member from the OTU378 Firmicutes 0.03 ± 0.01 family Ruminococcaceae OTU16 Proteobacteria Kushneria 2.57 ± 2.14 OTU39 Firmicutes Sporobacter 0.75 ± 0.25 OTU11 Bacteroidetes Bacteroides 0.58 ± 0.23 Unnamed member from the OTU101 Bacteroidetes 0.51 ± 0.13 family Prevotellaceae OTU98 Proteobacteria Psychrobacter 0.41 ± 0.17 OTU131 Proteobacteria Bergeriella 0.26 ± 0.13 OTU48 Actinobacteria Collinsella 0.24 ± 0.08 Unnamed member from the OTU277 Bacteroidetes 0.18 ± 0.08 family Prevotellaceae OTU89 Firmicutes Oscillibacter 0.17 ± 0.07 OTU200 Firmicutes Peptoniphilus 0.15 ± 0.06 OTU444 Firmicutes Peptoniphilus 0.09 ± 0.03 OTU447 Fusobacteria Ilyobacter 0.08 ± 0.02 OTU38 Firmicutes Sporobacter 0.85 ± 0.44 OTU82 Bacteroidetes Unnamed member from the 0.67 ± 0.24 125

family Prevotellaceae Unnamed member from the OTU65 Proteobacteria 0.51 ± 0.28 family Pseudomonadaceae OTU221 Bacteroidetes Alistipes 0.33 ± 0.17 OTU238 Firmicutes Anaerococcus 0.25 ± 0.11 OTU103 Proteobacteria Anaerobiospirillum 0.24 ± 0.08 Unnamed member from the OTU75 Firmicutes 0.23 ± 0.08 family Peptostreptococcaceae OTU234 Firmicutes Anaerococcus 0.18 ± 0.08 OTU219 Firmicutes Dialister 0.08 ± 0.03 OTU128 Proteobacteria Edwardsiella 0.07 ± 0.03 OTU757 Fusobacteria Ilyobacter 0.03 ± 0.01 OTU28 Bacteroidetes Alistipes 1.27 ± 0.62 OTU60 Firmicutes Sporanaerobacter 0.49 ± 0.18 OTU31 Proteobacteria Sutterella 0.48 ± 0.27 OTU104 Firmicutes Subdoligranulum 0.22 ± 0.11 OTU241 Fusobacteria Sneathia 0.20 ± 0.10 OTU207 Firmicutes Fastidiosipila 0.17 ± 0.07 OTU76 Proteobacteria Succinivibrio 0.14 ± 0.09 OTU250 Proteobacteria Psychrobacter 0.14 ± 0.06 OTU187 Firmicutes Papillibacter 0.13 ± 0.06 OTU133 Proteobacteria Campylobacter 0.13 ± 0.06 OTU223 Bacteroidetes Petrimonas 0.12 ± 0.05 OTU81 Proteobacteria Escherichia / Shigella 0.11 ± 0.06 OTU66 Fusobacteria Ilyobacter 0.11 ± 0.05 OTU236 Firmicutes Acetanaerobacterium 0.09 ± 0.04 OTU357 Firmicutes Sporobacter 0.08 ± 0.04 OTU218 Firmicutes Faecalibacterium 0.06 ± 0.04 OTU408 Firmicutes Subdoligranulum 0.05 ± 0.02 OTU88 Firmicutes Leuconostoc 0.05 ± 0.02 OTU638 Firmicutes Faecalibacterium 0.04 ± 0.01 OTU388 Fusobacteria Fusobacterium 0.04 ± 0.01 OTU333 Fusobacteria Ilyobacter 0.04 ± 0.01 Unnamed member from the OTU999 Firmicutes 0.03 ± 0.01 family Clostridiales XI OTU813 Fusobacteria Ilyobacter 0.02 ± 0.01 OTU6 Firmicutes Subdoligranulum 2.82 ± 1.51 OTU22 Firmicutes Butyricicoccus 1.20 ± 1.07 OTU36 Proteobacteria Psychrobacter 0.83 ± 0.33 OTU44 Proteobacteria Psychrobacter 0.48 ± 0.22 OTU54 Fusobacteria Fusobacterium 0.47 ± 0.24 OTU80 Firmicutes Helcococcus 0.45 ± 0.23 OTU127 Firmicutes Subdoligranulum 0.32 ± 0.16 OTU112 Proteobacteria Psychrobacter 0.26 ± 0.24 OTU335 Bacteroidetes Petrimonas 0.26 ± 0.17 OTU96 Firmicutes Anaerovorax 0.23 ± 0.15 OTU85 Firmicutes Allofustis 0.21 ± 0.09 OTU145 Bacteroidetes Porphyromonas 0.20 ± 0.08 OTU177 Bacteroidetes Porphyromonas 0.19 ± 0.09 OTU256 Firmicutes Anaerococcus 0.17 ± 0.08 OTU107 Firmicutes Sporanaerobacter 0.17 ± 0.06 OTU336 Firmicutes Anaerococcus 0.11 ± 0.05 126

OTU91 Fusobacteria Ilyobacter 0.10 ± 0.05 OTU168 Firmicutes Sporobacter 0.10 ± 0.05 OTU315 Firmicutes Sporobacter 0.08 ± 0.03 OTU769 Bacteroidetes Alistipes 0.07 ± 0.04 OTU585 Firmicutes Sporobacter 0.06 ± 0.02 Unnamed member from the OTU287 Firmicutes 0.04 ± 0.02 family Peptostreptococcaceae OTU373 Firmicutes Peptoniphilus 0.04 ± 0.02 OTU56 Fusobacteria Psychrilyobacter 0.04 ± 0.02 OTU307 Actinobacteria Arcanobacterium 0.03 ± 0.01 OTU421 Firmicutes Phascolarctobacterium 0.03 ± 0.01 OTU570 Fusobacteria Ilyobacter 0.02 ± 0.01 OTU673 Fusobacteria Psychrilyobacter 0.02 ± 0.01

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Table A4.5 Shared OTUs in the gut microbiota of most wild leopard seals. Average relative abundance of operational taxonomic units (OTUs) shared in the gut microbiota of most (more than half) individual wild leopard seals ± standard error (SE). OTUs occurring in the gut microbiota of () all individuals and (*) more than three quarters of individuals are marked. Average relative OTU Phylum Genus abundance / individual ± SE (%) OTU17 Firmicutes Weissella 5.63 ± 2.24 Unnamed member from the OTU4* Proteobacteria 9.57 ± 4.01 order Oceanospirillales OTU88* Firmicutes Leuconostoc 1.53 ± 0.64 OTU140* Firmicutes Lactococcus 0.68 ± 0.24 OTU14* Proteobacteria Psychrobacter 3.78 ± 1.41 OTU18* Proteobacteria Moraxella 3.36 ± 1.23 OTU3* Fusobacteria Fusobacterium 3.34 ± 2.13 OTU19* Proteobacteria Psychrobacter 2.71 ± 1.38 OTU90* Proteobacteria Citrobacter 1.01 ± 0.61 OTU94* Firmicutes Lactococcus 0.82 ± 0.37 OTU51* Proteobacteria Psychrobacter 0.68 ± 0.47 OTU183* Proteobacteria Acinetobacter 0.53 ± 0.25 OTU205* Firmicutes Streptococcus 0.46 ± 0.20 OTU213* Firmicutes Helcococcus 0.43 ± 0.26 OTU124* Firmicutes Peptoniphilus 0.34 ± 0.20 OTU389* Firmicutes Lactococcus 0.25 ± 0.10 OTU2371* Firmicutes Leuconostoc 0.07 ± 0.03 Unnamed member from the OTU10 Firmicutes 6.97 ± 3.12 family Clostridiales XI OTU25 Fusobacteria Fusobacterium 1.81 ± 1.27 OTU225 Firmicutes Lactococcus 0.20 ± 0.10 OTU549 Proteobacteria Psychrobacter 0.16 ± 0.06 OTU279 Proteobacteria Stenotrophomonas 0.11 ± 0.06 OTU1785 Firmicutes Weissella 0.09 ± 0.04 OTU108 Firmicutes Atopobacter 0.84 ± 0.68 OTU174 Firmicutes Helcococcus 0.76 ± 0.62 OTU92 Firmicutes Atopobacter 0.68 ± 0.55 OTU153 Firmicutes Peptostreptococcus 0.53 ± 0.34 OTU253 Firmicutes Lactococcus 0.19 ± 0.08 OTU1040 Proteobacteria Psychrobacter 0.15 ± 0.06 Unnamed member from the OTU999 Firmicutes 0.15 ± 0.06 family Clostridiales XI OTU365 Proteobacteria Acidovorax 0.12 ± 0.06 OTU429 Firmicutes Enterococcus 0.11 ± 0.04 OTU370 Firmicutes Peptoniphilus 0.09 ± 0.04 Unnamed member from the OTU815 Firmicutes 0.07 ± 0.03 family Clostridiales XI Unnamed member from the OTU65 Proteobacteria 0.07 ± 0.03 family Pseudomonadaceae OTU2357 Firmicutes Weissella 0.06 ± 0.02 OTU454 Firmicutes Roseburia 0.05 ± 0.02 OTU2359 Firmicutes Weissella 0.05 ± 0.02 OTU3506 Proteobacteria Psychrobacter 0.03 ± 0.01 128

OTU121 Firmicutes Weissella 0.89 ± 0.42 OTU15 Bacteroidetes Petrimonas 0.85 ± 0.40 OTU139 Firmicutes Weissella 0.80 ± 0.38 OTU64 Bacteroidetes Petrimonas 0.74 ± 0.38 OTU8 Fusobacteria Psychrilyobacter 0.54 ± 0.36 OTU151 Firmicutes Anaerococcus 0.34 ± 0.28 OTU188 Firmicutes Faecalibacterium 0.26 ± 0.23 OTU209 Firmicutes Peptoniphilus 0.22 ± 0.18 OTU1 Fusobacteria Ilyobacter 0.21 ± 0.10 OTU145 Bacteroidetes Porphyromonas 0.19 ± 0.10 OTU397 Proteobacteria Microvirgula 0.17 ± 0.08 OTU536 Proteobacteria Pectobacterium 0.15 ± 0.09 OTU167 Firmicutes Anaerococcus 0.15 ± 0.12 OTU519 Firmicutes Enterococcus 0.08 ± 0.04 OTU392 Proteobacteria Acinetobacter 0.08 ± 0.04 OTU298 Firmicutes Anaerococcus 0.07 ± 0.03 OTU404 Proteobacteria Psychrobacter 0.06 ± 0.03 OTU959 Proteobacteria Arcobacter 0.06 ± 0.03 OTU546 Firmicutes Peptoniphilus 0.06 ± 0.03 OTU5961 Firmicutes Lactococcus 0.05 ± 0.03 OTU1220 Proteobacteria Campylobacter 0.04 ± 0.02 OTU1074 Proteobacteria Psychrobacter 0.04 ± 0.01 OTU3599 Proteobacteria Citrobacter 0.03 ± 0.01 OTU2005 Bacteroidetes Petrimonas 0.03 ± 0.01

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Table A4.6 Shared OTUs in the gut microbiota of captive leopard seals over time. Average relative abundance of operational taxonomic units (OTUs) shared in the gut microbiota of both individual captive leopard seals during both sampling times ± standard error (SE). Average relative OTU Phylum Genera abundance / individual ± SE (%) OTU12 Firmicutes Subdoligranulum 15.90 ± 5.23 OTU3 Fusobacteria Fusobacterium 3.50 ± 2.16 OTU30 Firmicutes Sporobacter 3.02 ± 1.28 OTU34 Firmicutes Sporobacter 2.01 ± 0.50 OTU13 Fusobacteria Ilyobacter 1.57 ± 0.66 OTU70 Firmicutes Oscillibacter 1.34 ± 0.67 OTU2 Firmicutes Subdoligranulum 1.34 ± 0.98 OTU50 Firmicutes Sporobacter 1.05 ± 0.60 OTU191 Firmicutes Oscillibacter 0.99 ± 0.46 OTU89 Firmicutes Oscillibacter 0.66 ± 0.37 OTU144 Firmicutes Oscillibacter 0.64 ± 0.38 OTU137 Firmicutes Sporobacter 0.61 ± 0.29 OTU254 Firmicutes Oscillibacter 0.59 ± 0.15 OTU56 Fusobacteria Psychrilyobacter 0.56 ± 0.29 OTU211 Firmicutes Oscillibacter 0.53 ± 0.23 OTU1 Fusobacteria Ilyobacter 0.52 ± 0.22 Unnamed member of the OTU129 Firmicutes 0.48 ± 0.24 family Ruminococcaceae OTU169 Firmicutes Subdoligranulum 0.46 ± 0.28 OTU179 Firmicutes Sporobacter 0.42 ± 0.12 OTU142 Firmicutes Papillibacter 0.36 ± 0.18 OTU313 Firmicutes Oscillibacter 0.33 ± 0.26 OTU266 Firmicutes Subdoligranulum 0.31 ± 0.12 OTU162 Proteobacteria Escherichia / Shigella 0.31 ± 0.22 OTU193 Firmicutes Subdoligranulum 0.25 ± 0.15 Unnamed member of the OTU235 Firmicutes 0.24 ± 0.13 order Clostridiales OTU48 Actinobacteria Collinsella 0.21 ± 0.14 OTU333 Fusobacteria Ilyobacter 0.19 ± 0.08 OTU412 Fusobacteria Ilyobacter 0.18 ± 0.07 OTU330 Firmicutes Oscillibacter 0.16 ± 0.08 OTU306 Firmicutes Oscillibacter 0.14 ± 0.07 OTU1249 Firmicutes Sporobacter 0.14 ± 0.06 OTU578 Firmicutes Oscillibacter 0.14 ± 0.03 Unnamed member of the OTU483 Firmicutes family 0.14 ± 0.04 Peptostreptococcaceae OTU1142 Firmicutes Oscillibacter 0.12 ± 0.05 OTU329 Firmicutes Papillibacter 0.11 ± 0.03 OTU345 Firmicutes Sporobacter 0.10 ± 0.07 OTU751 Firmicutes Anaerovorax 0.10 ± 0.02 Unnamed member of the OTU773 Firmicutes 0.10 ± 0.06 family Lachnospiraceae OTU150 Firmicutes Coprobacillus 0.09 ± 0.05 OTU1430 Bacteroidetes Bacteroides 0.08 ± 0.05 130

OTU966 Firmicutes Oscillibacter 0.07 ± 0.02 OTU731 Firmicutes Sporobacter 0.07 ± 0.02 OTU1500 Firmicutes Sporobacter 0.07 ± 0.03 Unnamed member of the OTU709 Firmicutes 0.07 ± 0.03 order Clostridiales OTU1088 Firmicutes Soehngenia 0.06 ± 0.03 OTU487 Firmicutes Acetanaerobacterium 0.06 ± 0.02 Unnamed member of the OTU3327 Firmicutes 0.05 ± 0.02 order Clostridiales OTU613 Firmicutes Sporobacter 0.04 ± 0.01 OTU1051 Firmicutes Subdoligranulum 0.04 ± 0.02

131

Table A4.7 Shared OTUs in the gut microbiota of most southern elephant seal adults and sub-adults and captive leopard seals. Average relative abundance of operational taxonomic units (OTUs) shared in the gut microbiota of southern elephant adult and sub-adult seals and captive leopard seals ± standard error (SE). Average relative abundance ± SE / individual (%) Southern OTU Phylum Genus elephant seal Captive adults and leopard seals sub-adults OTU8 Fusobacteria Psychrilyobacter 1.94 ± 0.62 0.93 ± 0.54 OTU3 Fusobacteria Fusobacterium 4.37 ± 1.77 3.50 ± 2.16 OTU1 Fusobacteria Ilyobacter 8.21 ± 2.45 0.52 ± 0.22 OTU13 Fusobacteria Ilyobacter 1.66 ± 0.48 1.57 ± 0.66 OTU27 Firmicutes Sporobacter 1.46 ± 0.44 0.19 ± 0.11 OTU2 Firmicutes Subdoligranulum 4.96 ± 2.17 1.34 ± 0.98 Unnamed member from OTU378 Firmicutes the family 0.03 ± 0.01 0.04 ± 0.02 Ruminococcaceae OTU48 Actinobacteria Collinsella 0.24 ± 0.08 0.18 ± 0.07 OTU89 Firmicutes Oscillibacter 0.17 ± 0.07 0.64 ± 0.38 Unnamed member from OTU75 Firmicutes the family 0.23 ± 0.08 0.06 ± 0.06 Peptostreptococcaceae OTU333 Fusobacteria Ilyobacter 0.04 ± 0.01 0.21 ± 0.14 OTU128 Proteobacteria Edwardsiella 0.07 ± 0.03 1.18 ± 1.17 OTU81 Proteobacteria Escherichia / Shigella 0.11 ± 0.06 0.12 ± 0.06 OTU638 Firmicutes Faecalibacterium 0.04 ± 0.01 0.03 ± 0.02 OTU56 Fusobacteria Psychrilyobacter 0.04 ± 0.02 0.61 ± 0.29 OTU813 Fusobacteria Ilyobacter 0.02 ± 0.01 0.01 ± 0.01 OTU66 Fusobacteria Ilyobacter 0.11 ± 0.05 0.07 ± 0.05 OTU31 Proteobacteria Sutterella 0.48 ± 0.27 0.08 ± 0.05 Unnamed member from OTU287 Firmicutes the family 0.04 ± 0.02 0.02 ± 0.01 Peptostreptococcaceae OTU91 Fusobacteria Ilyobacter 0.10 ± 0.05 0.04 ± 0.03 OTU127 Firmicutes Subdoligranulum 0.32 ± 0.16 0.01 ± < 0.01

Table A4.8 PERMANOVA on gut microbiota of southern elephant seal adults and sub-adults with mass. PERMANOVA of gut microbial abundance data with known mass of seals used as co- variables to generate a permutated F statistic (F) and permutated p-value (P) with calculated degrees of freedom (d.f.) and sums of squares (SS) noted. Significance level: ***P = ≤ 0.001, **P = 0.01, *P = 0.025. Source of Data d.f. SS F P Variation Southern Mass 1 4437.9 1.4757 0.0602 elephant seal Sex 1 6555.5 2.1798 0.0031** adults and sub-adults Mass x sex 1 2076.1 0.69036 0.9135

132

Figure A4.1 Relationship between microbial richness, characteristic phyla and OTUs with mass of southern elephant seal adults and sub-adults. Linear regression models the strength of the relationship between the mass of southern elephant seal adult and sub-adults and (A) Chao 1; the abundance of phyla: (B) Firmicutes; (C) Fusobacteria; (D) Proteobacteria; (E) Bacteroidetes; and the abundance of operational taxonomic units (OTUs): (F) OTU52; (G) OTU1; (H) OTU3; (I) OTU13; (J) OTU15.

133

Table A4.9 Nearest sequence matches to the genus Cetobacterium in GenBank. Two operational taxonomic units (OTUs) from the genus Cetobacterium occurred in the gut microbiota of the female captive leopard seal during illness. These sequences were compared to sequences in GenBank using the Basic Local Alignment Search Tool (BLAST) to find the nearest source host for each sequence. The uploaded sequence length is compared to the database of known and sequenced bacteria and produces a list of sequences and their sources with the percent similarity of gene sequence coverage between the two sequences (Max. ID %). First nearest Max. ID Second nearest Max. ID OTU source (%) source (%) OTU5 Polar bear faeces 99 Zebrafish colon 99 OTU9 Polar bear faeces 98 Freshwater 97

134

APPENDIX 5

Table A5.1 Characteristics of mammalian hosts used in the study. Abbreviated table data is as follows: number of sequences used (No. of seq.); gut morphology (Gut morph.); hindgut fermentor (HG); foregut fermentor (FG); simple gut (S); marine (M); terrestrial (T); carnivore (C); herbivore (H); omnivore (O); and article reference (Ref.). No. Gut Sample name of Species name Common name Order Family Habitat Diet Ref. morph. seq. AE1 92 Elephas maximus Asian Elephant Proboscidae Elephantidae HG T H [23] AE2 101 Elephas maximus Asian Elephant Proboscidae Elephantidae HG T H [23] AE3 97 Elephas maximus Asian Elephant Proboscidae Elephantidae HG T H [23] AFBAB 100 Papio hamadryas Hamadryas Baboon Primates Cercopithecidae S T O [23] BAZ 100 Papio hamadryas Hamadryas Baboon Primates Cercopithecidae S T O [23] AFEL 100 Loxodonta africana African Elephant Proboscidae Elephantidae HG T H [23] AFEL2 100 Loxodonta africana African Elephant Proboscidae Elephantidae HG T H [23] AFEL3 100 Loxodonta africana African Elephant Proboscidae Elephantidae HG T H [23] AFYEL 100 Loxodonta africana African Elephant Proboscidae Elephantidae HG T H [23] AFZEB 100 Equus zebra hartmannae Hartmanns Mountain Zebra Perissodactyla Equidae HG T H [23] ARMA 100 Tolypeutes matacus Southern Three Branded Armadillo Xenarthra Dasypodidae S T I [23] AS1 100 Ovis ammon Argali Sheep Artiodactyla Bovidae FG T H [23] AS2 100 Ovis ammon Argali Sheep Artiodactyla Bovidae FG T H [23] AS3 100 Ovis ammon Argali Sheep Artiodactyla Bovidae FG T H [23] BARB 100 Babyrousa babyrussa Barbirusa Artiodactyla Suidae FG T O [23] BAT 100 Carollia perspicillata Sebas Short-tailed Bat Chiroptera Phyllostomidae S T O [23] BB1 100 Ursus americanus North American Black Bear Carnivora Ursidae S T O [23] BB2 100 Ursus americanus North American Black Bear Carnivora Ursidae S T O [23] BDOG1 100 Speothos venaticus Bushdog Carnivora Canidae S T C [23] BDOG3 34 Speothos venaticus Bushdog Carnivora Canidae S T C [23] BEAR 40 Ursus sp. Bear from Norway Carnivora Ursidae S T O [390] BG 97 Bos javanicus Banteng Artiodactyla Bovidae FG T H [23] BH1 100 Ovis canadensis Bighorn Sheep Artiodactyla Bovidae FG T H [23] BH2 100 Ovis canadensis Bighorn Sheep Artiodactyla Bovidae FG T H [23] BHSD 100 Ovis canadensis Bighorn Sheep Artiodactyla Bovidae FG T H [23] BKLE 100 Eulemur macaco Black Lemur Primates Lemuridae S T O [23] BNO 88 Pan paniscus Bonobo Primates Hominidae S T O [23] CAL 100 Callimico goeldii Goeldis Marmoset Primates Cebidae S T O [23] CAP 100 Hydrochoerus hydrochaeris Capybara Rodentia Caviidae HG T H [23] CATT 100 Bos taurus Holstein Cattle Artiodactyla Bovidae FG T H [391] CE2 100 Acinonyx jubatus Cheetah Carnivora Felidae S T C [23] CE3 90 Acinonyx jubatus Cheetah Carnivora Felidae S T C [23] CHIMP1 100 Pan troglodytes Chimpanzee Primates Hominidae S T O [23] CHIMP12 80 Pan troglodytes Chimpanzee Primates Hominidae S T O [23] 135

CLS1 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study CLS2 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study COL 101 Colobus guereza kikuyuensis Eastern Black and White Colobus Primates Cercopithecidae FG T H [23] DL 100 Pygathrix sp. Douc Langur Primates Cercopithecidae FG T H [23] DOG_ 100 Canis lupus familiaris Dog Carnivora Canidae S T O [384] DOG 100 Canis lupus familiaris Dog Carnivora Canidae S T O [384] DOG 100 Canis lupus familiaris Dog Carnivora Canidae S T O [384] DOG 100 Canis lupus familiaris Dog Carnivora Canidae S T O [384] DOG 100 Canis lupus familiaris Dog Carnivora Canidae S T O [384] DUG 90 Dugong dugong Dugong Sirenia Dugonidae M HM [45] EAC 100 Colobus angolensis East Angolan Colobus Primates Cercopithecidae FG T H [23] ECH 100 Tachyglossus aculeatus Short Beaked Echidna Monotremata Tachyglossidae S T CI [23] ELAND 64 Taurotragus oryx Eland Artiodactyla Bovidae FG T H [242] ES01 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES04 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES10 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES11 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES12 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES14 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES15 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES16 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES17 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES20 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES21 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES22 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES24 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES25 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES26 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES28 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES29 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES30 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES31 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES33 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES34 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES37 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES40 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study ES45 100 Mirounga leonina Southern Elephant Seal Carnivora Phocidae S M CM This study FF 100 Pteropus scapulatus Flying Fox Chiroptera Pterodidae S T H [23] FL 100 Trachypithecus francoisi Francois Langur Primates Cercopithecidae FG T H [23] GAZ_GRA 60 Nanger granti Grants Gazelle Artiodactyla Bovidae FG T H [242] GAZ_THO 77 Eudorcas thomsonii Thomsons Gazelle Artiodactyla Bovidae FG T H [242] GIR 100 Giraffa camelopardalis reticulata Reticulated Giraffe Artiodactyla Giraffidae FG T H [23] GOR 100 Gorilla gorilla gorilla Western Lowland Gorilla Primates Hominidae HG T H [23] GORSD 100 Gorilla gorilla gorilla Western Lowland Gorilla Primates Hominidae HG T H [23] 136

GP 100 Ailuropoda melanoleuca Giant Panda Carnivora Ursidae S T H [23] GREY_GL 76 Halichoerus grypus Grey Seal Carnivora Phocidae S M CM [44] GZ 100 Equus grevyi Grevys Zebra Perissodactyla Equidae HG T H [23] HAMS 100 Mesocricetus auratus Hamster Rodentia Cricetidae HG T O [223] HARB_GL 77 Phoca vitulina Harbour Seal Carnivora Phocidae S M CM [44] HH 100 Erinaceus albiventris Hedgehog Insectivora Erinaceidae S T CI [23] HOOD_GL 100 Cystophora cristata Hooded Seal Carnivora Phocidae S M CM [44] HORSEJ 100 Equus ferus caballus Horse Perissodactyla Equidae HG T H [23] HORSEM 100 Equus ferus caballus Horse Perissodactyla Equidae HG T H [23] HRX 100 Procavia capensis Rock Hyrax Hyracoidea Procaviidae FG T H [23] HUM_ECK 100 Homo sapiens sapiens Human Primates Hominidae S T O [13] HUM_FAT 100 Homo sapiens sapiens Human Primates Hominidae S T O [26] HUM_OLD 88 Homo sapiens sapiens Human Primates Hominidae S T O [229] HUM_VEG 61 Homo sapiens sapiens Human Primates Hominidae S T O [241] HY1 100 Crocuta crocuta Spotted Hyena Carnivora Hyaenidae S T C [23] HY2 100 Crocuta crocuta Spotted Hyena Carnivora Hyaenidae S T C [23] IR 100 Rhinoceros unicornis Indian Rhino Perissodactyla Rhinocerotidae HG T H [23] KO1 100 Macropus rufus Red Kangaroo Diprotodontia Macropidae FG T H [23] KO2 97 Macropus rufus Red Kangaroo Diprotodontia Macropidae FG T H [23] LI1 80 Panthera leo Lion Carnivora Pantherinae S T C [23] LI2 100 Panthera leo Lion Carnivora Pantherinae S T C [23] LI3 100 Panthera leo Lion Carnivora Pantherinae S T C [23] M 100 Callithrix geoffroyi Geoffreys Marmoset Primates Callitrichidae S T O [23] ML 100 Eulemur mongoz Mongoose Lemur Primates Lemuridae S T O [23] MOLERAT 102 Heterocephalus glaber Naked Molerat Rodentia Bathyergidae HG T H [23] OK1 100 Okapia johnstoni Okapi Artiodactyla Giraffidae FG T H [23] OK2 100 Okapia johnstoni Okapi Artiodactyla Giraffidae FG T H [23] OK3 100 Okapia johnstoni Okapi Artiodactyla Giraffidae FG T H [23] ORANG1 100 Pongo pygmaeus Orangutan Primates Hominidae HG T H [23] ORANG2 100 Pongo pygmaeus Orangutan Primates Hominidae HG T H [23] PB1 100 Ursus maritimus Polar Bear Carnivora Ursidae S M CM [23] PB2 100 Ursus maritimus Polar Bear Carnivora Ursidae S M CM [23] PBF_GL 100 Ursus maritimus Polar Bear Carnivora Ursidae S M CM [392] PBM_GL 24 Ursus maritimus Polar Bear Carnivora Ursidae S M CM [392] RA 100 Oryctolagus cuniculus European Rabbit Lagomorpha Leporidae HG T H [23] RABB 100 Oryctolagus cuniculus European Rabbit Lagomorpha Leporidae HG T H [393] RAT 80 Rattus norvegicus Norway Rat (Wistar) Rodentia Muridae HG T O [33] REIN_ NORNAT 34 Rangifer tarandus Norway Reindeer Artiodactyla Cervidae FG T H [394] REIN_ NORPEL 57 Rangifer tarandus Norway Reindeer Artiodactyla Cervidae FG T H [394] REIN_SVAL 31 Rangifer tarandus platyrhynchus Svalbard Reindeer Artiodactyla Cervidae FG T H [395] RH 100 Diceros bicornis Black Rhino Perissodactyla Rhinocerotidae HG T H [23] RHSD 100 Procavia capensis Rock Hyrax Hyracoidea Procaviidae FG T H [23] RP 100 Ailurus fulgens Red Panda Carnivora Ailuridae S T H [23] RPSD 100 Ailurus fulgens Red Panda Carnivora Ailuridae S T H [23] 137

RRH 100 Potamochoerus porcus Red River Hog Artiodactyla Suidae FG T O [23] RT 100 Lemur catta Ring Tailed Lemur Primates Lemuridae S T O [23] SAKI 100 Pithecia pithecia White Faced Saki Primates Pitheciidae S T O [23] SB 100 Tremarctos ornatus Spectacled Bear Carnivora Ursidae S T O [23] SBK 32 Antidorcas marsupialis Springbok Artiodactyla Bovidae FG T H [23] SBSD 100 Antidorcas marsupialis Springbok Artiodactyla Bovidae FG T H [23] SP2 100 Gazella spekei Spekes Gazelle Artiodactyla Bovidae FG T H [23] SP3 100 Gazella spekei Spekes Gazelle Artiodactyla Bovidae FG T H [23] SPIM 100 Ateles geoffroyi Black Handed Spider Monkey Primates Atelidae S T O [23] SQ 100 Callosciurus prevostii Prevosts Squirrel Rodentia Sciuridae S T O [23] TAK 100 Budorcas taxicolor Takin Artiodactyla Bovidae FG T H [23] TU1 100 Ovis orientalis arkal Transcaspian Urial Sheep Artiodactyla Bovidae FG T H [23] TU2 100 Ovis orientalis arkal Transcaspian Urial Sheep Artiodactyla Bovidae FG T H [23] VWP 100 Sus cebifrons Visayan Warty Pig Artiodactyla Suidae FG T H [23] WA 101 Equus africanus somaliensis Somali Wild Ass Perissodactyla Equidae HG T H [23] WLS49 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS50 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS51 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS52 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS53 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS54 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS56 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS57 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS58 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS59 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS60 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study WLS63 100 Hydrurga leptonyx Leopard Seal Carnivora Phocidae S M CM This study ZEB_NELS 41 Equus quagga Plains Zebra perissodactyla Equidae HG T H [242] ZEBU 30 Bos primigenius African Zebu Cattle Artiodactyla Bovidae FG T H [242]

138

Figure A5.1 Phylogeny of the family Phocidae. Adapted from Berta et al. [135] based on molecular data from Davis et al. [113]. Seals included in this study inhabiting the Antarctic () or Arctic () are marked.

Figure A5.2 Similarity of the gut microbiota of host mammals with captive and wild representatives. nMDS ordination plot displays similarity of the gut microbiota of host mammals with representatives from a: (c) captive and (w) wild environment.

139

Figure A5.3 Similarity of the gut microbiota of mammals grouped by diet and location. nMDS ordination plot displays similarity of the gut microbiota of host mammals with particular hosts of interest highlighted.

140

Table A5.2 Characteristic OTUs in the gut microbiota of mammal hosts grouped by diet and location. Foremost ten characteristic operational taxonomic units (OTUs) in the gut microbiota of hosts identified using SIMPER analysis. Hosts are grouped based on diet and location. Average Contribution Groups Phylum Genus abundance to total (%) Firmicutes Oscillibacter 2.4 10.4 Firmicutes Coprococcus 2.0 9.4 Bacteroidetes Rikenella 1.6 6.8 Firmicutes Papillibacter 1.5 6.4 Terrestrial Firmicutes Robinsoniella 1.3 4.9 herbivores Firmicutes Acetivibrio 1.3 4.7 Firmicutes Sporobacter 1.2 4.6 Firmicutes Ruminococcus 1.3 4.5 Firmicutes Ethanoligenens 1.0 3.2 Firmicutes Anaerotruncus 0.9 3.0 Bacteroidetes Prevotella 2.2 11.9 Firmicutes Coprococcus 1.7 10.9 Firmicutes Blautia 1.9 10.4 Firmicutes Streptococcus 1.5 4.8 Terrestrial Firmicutes Oscillibacter 1.2 4.6 omnivores Bacteroidetes Bacteroides 1.5 4.5 Firmicutes Robinsoniella 0.9 3.8 Firmicutes Faecalibacterium 1.0 3.6 Bacteroidetes Barnesiella 0.9 3.5 Proteobacteria Hallella 0.7 3.3 Firmicutes Peptostreptococcus 3.1 19.3 Firmicutes Clostridium 2.7 16.6 Firmicutes Sporacetigenium 2.1 11.7 Firmicutes Blautia 2.4 11.5 Firmicutes Coprococcus 1.5 7.8 Terrestrial Actinobacteria Collinsella 1.5 5.7 carnivores Firmicutes Lactobacillus 1.8 5.6 Escherichia / Proteobacteria Shigella 1.2 5.0 Firmicutes Robinsoniella 0.8 2.6 Firmicutes Enterococcus 0.8 2.5 Fusobacteria Fusobacterium 3.0 16.8 Firmicutes Faecalibacterium 1.9 9.8 Fusobacteria Cetobacterium 1.9 8.9 Firmicutes Oscillibacter 1.6 7.4 Marine Proteobacteria Psychrobacter 1.9 6.2 carnivores Bacteroidetes Bacteroides 1.5 5.5 Firmicutes Butyricicoccus 0.9 3.6 Firmicutes Sporanaerobacter 1.0 3.2 Firmicutes Sporobacter 0.7 3.0 Bacteroidetes Porphyromonas 1.1 2.9 Firmicutes Clostridium 10.0 11.1 Firmicutes Coprococcus 7.0 7.8 Marine Bacteroidetes Bacteroides 6.0 6.7 herbivore Bacteroidetes Prevotella 6.0 6.7 Firmicutes Oscillibacter 5.0 5.6 141

Verrucomicrobia Akkermansia 4.0 4.4 Firmicutes Anaerotruncus 4.0 4.4 Bacteroidetes Alistipes 3.0 3.3 Firmicutes Roseburia 3.0 3.3 Firmicutes Ruminococcus 3.0 3.3 Firmicutes Clostridium 10.0 11.1 Firmicutes Coprococcus 7.0 7.8 Bacteroidetes Bacteroides 6.0 6.7 Bacteroidetes Prevotella 6.0 6.7 Firmicutes Oscillibacter 5.0 5.6 Verrucomicrobia Akkermansia 4.0 4.4

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APPENDIX 6

Table A6.1 Prevalence of E. coli pathotypes in seawater. All samples were tested and negative for tsh, papC and CNF. Sampling sites (S) of locations in Potter Cove and sampling time (T) are listed in Figure 6.1. Number of Seawater site and time n isolates positive for iucD (%) S2, T1 3 0.0 S5, T3 6 50.0 Total 9 33.3

Table A6.2 Prevalence of E. coli pathotypes in faecal samples from mother-pup pairs. All samples were negative for Eae, Stx1, Stx2, Sta, Stb, LT, F4 and Tsh. Mother-pup pairs are designated by lower case letter. Number of positive isolates (%) Mother-pup pair n eae iucD papC CNF e Adult 5 20.0 60.0 20.0 0.0 Pup 5 0.0 60.0 20.0 0.0 h Adult 3 0.0 0 66.7 0.0 Pup 3 0.0 100 100 33.3 k Adult 5 0.0 100 100 0.0 Pup 6 0.0 50.0 0.0 0.0

143

Table A6.3 Relationship between intermediate AMR and site of origin of E. coli isolates in faecal samples from seals of different age- classes and seawater samples. Abbreviations of antimicrobial agents used to measure anitmicrobial resistance (AMR) in this study are: (AMC) Amoxicillin/Clavulanic acid; (TIO) Ceftiofur; (CRO) Ceftriaxone; (CIP) Ciprofloxacin; (AMK) Amikacin; (AMP) Ampicillin; (FOX) Cefoxitin; (GEN) Gentamicin; (KAN) Kanamycin; (NAL) Nalidixic acid; (STR) Streptomycin; (SXT) Trimethoprim-Sulphamethaoxazole; (CHL) Chloramphenicol; (FIS) Sulfisoxazole; (TET) Tetracycline. Information on (S) sampling sites and (T) times are noted in Figure 6.1 and Table A1.1. Total Number of number intermediate Group AMK GEN STR KAN CHL NAL CIP TET FIS SXT AMC AMP FOX CRO TIO of resistant isolates isolates Adults 25 4 (16.0%) 0 0 2 0 0 0 0 1 0 0 0 2 0 0 0

Sub- 15 4 (26.7%) 0 0 1 0 0 0 0 1 1 0 0 1 0 0 0 eals

s adults lephant lephant outhern outhern e S Pups 16 6 (37.5%) 0 0 2 0 1 1 0 1 0 0 1 1 0 0 0 Summary southern 56 14 (25.0%) 0 0 5 0 1 1 0 3 0 0 1 4 0 0 0 elephant seals S1, T1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

S2, T1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

S3, T1 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

S4, T1 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

S5, T1 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

S1, T3 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 Seawater

S2, T3 2 1 (9.1%) 0 0 1 0 0 0 0 0 0 0 0 0 0 0 0

S3, T3 1 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

S4, T3 2 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

S5, T3 2 1 (9.1%) 0 0 0 0 1 0 0 0 0 0 0 0 0 0 0 Summary 11 2 (18.2%) 0 0 1 0 1 0 0 0 0 0 0 0 0 0 0 seawater Total 67 16 (23.9%) 0 0 6 0 2 1 0 3 0 0 1 4 0 0 0

144

Table A6.4 Information pertaining to E. coli control strains used in multiplex PCRs. Control strains from the Escherichia coli Laboratory (EcL) used in this study were tested by colony hybridization for the following virulence factors: LT, STa, STb, Stx1, Stx2, eae, P, CNF, Aerobactin, EAST1, AFA, Paa, Aida, Tsh, F4, F18, F5, F6, F41, F17. Geographical Growth EcL strain Serotype Pathotype Isolated from History Reference origin conditions 1 Pig, two weeks, Canada, 1982, EcL Tryptic soy EcL3463 O115 Negative [396] ileum, diarrhoea Quebec Laboratory broth Stx1: Stx2: eae: EAST1: Bovine, unknown, Canada, 1994, EcL Tryptic soy EcL6611 O111 1 Paa: EhxA: EFA-1 ileum, unknown Quebec Laboratory broth LT: STa: STb: EAST1: Pig, six weeks, Canada, 1997, EcL Tryptic soy EcL7805 O149:H10 1 [355,356] Paa: F4(K88) ileum, diarrhoea Quebec Laboratory broth 1 Pig, four weeks, Canada, 2005, EcL Tryptic soy EcL13421 P: CNF unknown, unknown Quebec Laboratory broth

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