Mechanisms of Predation Resistance and the Effect Ofpredation on the Pathogenicity of Marine Bacteria

Mechanisms of predation resistance

and the effect of predation on the pathogenicity of marine bacteria

Parisa Noorian

A thesis in fulfilment of the requirements for the degree of

Doctor of Philosophy

School of Biotechnology and Biomolecular Sciences

Faculty of Science

The University of New South Wales

Sydney, Australia

July 2018

Thesis/Dissertation Sheet

Surname/Family Name :Noorian Given Name/s :Parisa Abbreviation for degree as give in the University calendar :Ph.D. Faculty :Science School :BABs :Mechanisms of predation resistance and the effect of predation on Thesis Title the pathogenicity of marine bacteria

Abstract 350 words maximum: (PLEASE TYPE) Vibrio spp. are an autochthonous inhabitant of coastal marine environments where predation by protozoa is a shaping force leading to the evolution of antiprotozoal mechanisms that may also function as virulence factors in animal and human hosts. Vibrio cholerae and Vibrio vulnificus were used in this study in order to further elucidate bacterial adaptations against different model protozoa and the potential role of these antiprotozoal factors in providing fitness in the environment and in a host.

The transcriptomic profile of established V. cholerae biofilms to predation by the amoeba, Acanthamoeba castellanii, was investigated. Genes that were significantly differentially expressed between grazed and ungrazed cultures were identified. Tyrosine metabolic genes were among the down-regulated transcripts in the grazed population. Homogentisic acid is the main intermediate of the L-tyrosine catabolic pathway, and is known to auto-oxidize, leading to the formation of the pigment, pyomelanin. Indeed, a pigmented mutant, disrupted in the gene encoding homogentisate 1, 2 - dioxygenase (hmgA) was more resistant to grazing by A. castellanii than the wild type.

Grazing resistance of V. vulnificus of different genotypes and places of isolation were evaluated using the protozoan predators, Tetrahymena pyriformis and A. castellanii, but no significant correlation was found in relation to grazing resistance. However, an oyster isolate, V. vulnificus Env1, showed significant grazing resistance and toxicity towards T. pyriformis. The whole genome sequence of Env1 was completed, annotated and compared to grazing sensitive strains to identify Env1 unique features.

Further studies revealed one of the antiprotozoal mechanisms of Env1 was secreted and iron-dependent. The transcriptomic profile of V. vulnificus Env1 under iron-replete and -deplete conditions was characterised. A master virulence regulator, arcA, was up-regulated when iron was readily available and an arcA mutant, showed a significant decrease in grazing resistance. Therefore, ArcA is a novel global regulator controlling the grazing resistance of V. vulnificus. In summary, this project revealed new defence systems against protozoan grazing expressed by V. cholerae and V. vulnificus that also play dual roles in environmental survival and pathogenicity, accentuating how protozoan grazing drives the evolution of pathogenicity in bacteria in the environment.

Declaration relating to disposition of project thesis/dissertation

I hereby grant to the University of New South Wales or its agents the right to archive and to make available my thesis or dissertation in whole or in part in the University libraries in all forms of media, now or here after known, subject to the provisions of the Copyright Act 1968. I retain all property rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation.

I also authorise University Microfilms to use the 350 word abstract of my thesis in Dissertation Abstracts International (this is applicable to doctoral theses only).

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Signature Witness Signature Date The University recognises that there may be exceptional circumstances requiring restrictions on copying or conditions on use. Requests for restriction for a period of up to 2 years must be made in writing. Requests for a longer period of restriction may be considered in exceptional circumstances and require the approval of the Dean of Graduate Research.

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Originality statement

‘I hereby declare that this submission is my own work and to the best of my knowledge it contains no materials previously published or written by another person, or substantial proportions of material which have been accepted for the award of any other degree or diploma at UNSW or any other educational institution, except where due acknowledgement is made in the thesis. Any contribution made to the research by others, with whom I have worked at UNSW or elsewhere, is explicitly acknowledged in the thesis. I also declare that the intellectual content of this thesis is the product of my own work, except to the extent that assistance from others in the project's design and conception or in style, presentation and linguistic expression is acknowledged.’

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‘I hereby grant the University of New South Wales or its agents the right to archive and to make available my thesis or dissertation in whole or part in the University libraries in all forms of media, now or here after known, subject to the provisions of the Copyright Act 1968. I retain all proprietary rights, such as patent rights. I also retain the right to use in future works (such as articles or books) all or part of this thesis or dissertation. I also authorise University Microfilms to use the 350 word abstract of my thesis in Dissertation Abstract International (this is applicable to doctoral theses only). I have either used no substantial portions of copyright material in my thesis or I have obtained permission to use copyright material; where permission has not been granted I have applied/will apply for a partial restriction of the digital copy of my thesis or dissertation.'

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AUTHENTICITY STATEMENT

‘I certify that the Library deposit digital copy is a direct equivalent of the final officially approved version of my thesis. No emendation of content has occurred and if there are any minor variations in formatting, they are the result of the conversion to digital format.’

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Acknowledgements

I would like to thank to all those who made the completion of this work possible. First and foremost, I wish to express my immense gratitude to my supervisor Associate Professor Diane McDougald; without whom none of this would have been possible. Thank you for everything. I could have not imagine a better mentor.

I would like to extend my appreciation to Professor Ruiting Lan for his generous support and accepting me as his student. I would like to express my gratitude to Professor Liz Harry for her continuous and warm support. It was a great experience to work in ithree institute. I would also like to thank Associate Professor Scott Rice, for all the help, advice and generous support.

Many people helped me along the way and I want to thank them for all their help, support, interest and valuable hints. Especially, I am indebted to thank Dr. Shuyang (Garfy) Sun. You are one of the first people who trained me back in my Mphil when I had just started real research and continued to support and help me in developing real research to this day. I would like to thank Gustavo Espinoza for all those long talks on science, experiments and life. I would like to express my heart-felt gratitude to Nasim Shah Mohammadi. You have been a great friend and support along the way.

I would like to thank Dr. Sean booth for collaboration on extraction of the toxic factor even if it still needs work. I received generous support from Dr. Florentin Constancias and would like say thank you for all your help in RNA-seq. We traded so many emails to make it work. I am happy to acknowledge Dr. Zhilliang Chen, Dr. Tonia Russel and Dr. Nandan Deshpande for all the help in technical support.

There are not enough words to describe how much I am indebted to my parents. You have continuously supported me towards my goals and have provided me with everything in life. There is no way for me to thank you enough. I would like to specially thank my brother Farzad. You are my pillar of support and strength, without you, I would have been lost long ago.

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Abstract

Vibrio spp. are an autochthonous inhabitant of coastal marine environments where predation by protozoa is a shaping force leading to the evolution of antiprotozoal mechanisms that may also function as virulence factors in animal and human hosts. Vibrio cholerae and Vibrio vulnificus were used in this study in order to further elucidate bacterial adaptations against different model protozoa and the potential role of these antiprotozoal factors in providing fitness in the environment and in a host.

Further studies revealed one of the antiprotozoal mechanisms of Env1 was secreted and iron-dependent. The transcriptomic profile of V. vulnificus Env1 under iron-replete and -deplete conditions was characterised. A master virulence regulator, arcA, was up- regulated when iron was readily available and an arcA mutant, showed a significant decrease in grazing resistance. Therefore, ArcA is a novel global regulator controlling the grazing resistance of V. vulnificus. In summary, this project revealed new defence systems against protozoan grazing expressed by V. cholerae and V. vulnificus that also play dual roles in environmental survival and pathogenicity, accentuating how protozoan grazing drives the evolution of pathogenicity in bacteria in the environment.

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List of contents

Contents

Thesis/Dissertation Sheet ...... II

Originality statement ...... III

Acknowledgements ...... V

Abstract ...... VI

List of contents ...... VII

List of figures ...... XII

List of tables ...... XV

List of appendices ...... XVI

List of abbreviations ...... XVII

1 Chapter 1: General introduction and literature review ...... 1

1.1 General introduction ...... 1

1.2 Description of Vibrio vulnificus and Vibrio cholerae ...... 3

1.2.1 Vibrio vulnificus ...... 3

1.2.2 V. vulnificus genotypes and biotypes ...... 4

1.2.3 Distribution of V. vulnificus ...... 6

1.2.4 Pathogenicity factors ...... 8

1.2.5 Vibrio cholerae ...... 21

1.2.6 Serogroups and biotypes ...... 21

1.2.7 Distribution of V. cholerae...... 22

1.2.8 Pathogenicity factors ...... 24

1.3 Predation by protozoa ...... 34

1.4 Bacterial adaptations against predation by protozoa...... 37

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1.5 Vibrio spp. use factors with dual roles in providing fitness in the environment and pathogenicity ...... 42

1.6 Chapter synopsis ...... 47

2 Chapter 2: Vibrio cholerae hmgA-mediated pyomelanisation confers resistance to predation by A. castellanii ...... 48

2.1 Introduction ...... 48

2.2 Materials and methods ...... 51

2.2.1 Strains and growth conditions ...... 51

2.2.2 Transcriptomic profiling of continuous-culture biofilms ...... 52

2.2.3 Transcriptome data analysis ...... 53

2.2.4 Early and late biofilm grazing assay with A. castellanii ...... 54

2.2.5 Quantification of pyomelanin and reactive oxygen species...... 55

2.2.6 H2O2 treatment of V. cholerae biofilms ...... 55

2.2.7 Catalase treatment of V. cholerae biofilms ...... 56

2.2.8 Experimental validation - qRT-PCR ...... 56

2.2.9 T. pyriformis grazing assays...... 57

2.2.10 V. cholerae - A. castellanii intracellular survival assay ...... 57

2.2.11 Data analysis ...... 58

2.3 Results and discussion ...... 59

2.3.1 RNA-seq revealed differences in the transcriptomes of grazed and ungrazed biofilms ...... 59

2.3.2 Pyomelanin production increases the grazing resistance of V. cholerae biofilms ...... 62

2.3.3 Pyomelanin and reactive oxygen species (ROS) concentrations ...... 71

2.3.4 Hydrogen peroxide addition increased the grazing resistance of V. cholerae 74

2.3.5 Conclusion ...... 76 VIII | P a g e

2.4 Acknowledgements ...... 76

3 Chapter 3: Characterisation of the antiprotozoal activity of Vibrio vulnificus ...... 77

3.1 Introduction ...... 77

3.2 Materials and methods ...... 80

3.2.1 Strains and growth conditions ...... 80

3.2.2 Assessment of resistance of planktonic bacteria to T. pyriformis predation 82

3.2.3 Assessment of resistance of biofilms to predation by T. pyriformis and A. castellanii ...... 82

3.2.4 Selective grazing assay ...... 83

3.2.5 Whole genome sequencing ...... 84

3.2.6 Comparative genomics ...... 84

3.2.7 Supernatant toxicity assay ...... 85

3.2.8 Biosurfactant emulsification assay...... 86

3.2.9 Data analysis ...... 86

3.3 Results and discussion ...... 87

3.3.1 Grazing resistance of V. vulnificus of different genotypes and from different sources...... 87

3.3.2 Grazing resistance of V. vulnificus biofilms ...... 87

3.3.3 Grazing resistance of planktonic cells of V. vulnificus ...... 93

3.3.4 V. vulnificus Env1 ...... 96

3.3.5 Features of the Env1 genome ...... 96

3.3.6 Comparative genomics ...... 99

3.3.7 Env1 unique genes encoding virulence and defence factors...... 108

3.3.8 Genes encoding iron uptake and utilisation proteins ...... 112

3.3.9 Supernatant toxicity assay ...... 118

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3.3.10 Presence of biosurfactants ...... 122

3.3.11 Conclusion ...... 123

3.4 Acknowledgements ...... 123

4 Chapter 4: Investigation of the secreted toxic factor produced by Vibrio vulnificus Env1 ...... 124

4.1 Introduction ...... 124

4.2 Materials and methods ...... 126

4.2.1 Strains and growth conditions ...... 126

4.2.2 V. vulnificus growth under iron-replete and -deplete conditions and toxicity of supernatants to T. pyriformis ...... 127

4.2.3 Transcriptome of V. vulnificus Env1 grown in iron-replete and -deplete conditions...... 128

4.2.4 Transcriptome data analysis ...... 128

4.2.5 Ammonia toxicity assay ...... 129

4.2.6 Biotin add-back experiments ...... 130

4.2.7 Electroporation and conjugation conditions ...... 130

4.2.8 Site-directed mutagenesis and complementation ...... 131

4.2.9 Assessment of resistance of V. vulnificus mutants to T. pyriformis predation 131

4.2.10 Data analysis ...... 132

4.3 Results and discussion ...... 133

4.3.1 V. vulnificus growth under iron-replete and -deplete conditions and toxicity of supernatants to T. pyriformis ...... 133

4.3.2 RNA-seq revealed differentially expressed genes under iron-deplete compared to -replete conditions...... 134

4.3.3 Effect of ammonium on T. pyriformis growth ...... 135

4.3.4 Genes involved in biotin biosynthesis ...... 139

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4.3.5 Genes involved in pathogenicity and virulence ...... 141

4.3.6 Conclusion ...... 151

4.4 Acknowledgements ...... 151

5 Chapter 5: Summary, general discussion and future work ...... 152

5.1 HmgA-mediated pyomelanisation protects V. cholerae against grazing by A. castellanii ...... 153

5.2 Place of isolation and genotype did not correlate with protozoan grazing resistance ...... 155

5.3 Master virulence regulator, ArcA, controls antiprotozoal activity of an iron- dependent toxic factor expressed by V. vulnificus ...... 156

5.4 Concluding remarks ...... 157

5.5 Future work ...... 159

References ...... 160

Appendices ...... 196

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List of figures

Figure 1-1 Model of the V. vulnificus quorum-sensing system ...... 15

Figure 1-2 The regulation of virulence genes by SmcR and Fur in response to iron and population density in V. vulnificus ...... 18

Figure 1-3 V. cholerae quorum sensing systems, comprised of three parallel circuits .. 30

Figure 1-4 Diagram depicting the regulatory pathways involved in virulence of Vibrio spp...... 31

Figure 1-5 Potential bacterial adaptations against predation by protozoa ...... 38

Figure 2-1 Differentially expressed transcripts in grazed compared to ungrazed biofilms involved in tyrosine degradation in V. cholerae ...... 61

Figure 2-2 Biofilm biomass of early (A) and late (B) biofilms of V. cholerae A1552 exposed to grazing by A. castellanii for 72 hours ...... 64

Figure 2-3 Biofilm biomass of V. cholerae wild type and hmgA mutant grown for 1 or 3 d as determined by CV staining ...... 65

Figure 2-4 The number of A. castellanii trophozoites (A) and cysts (B) after 0 and 24 - hour incubation in cell-free supernatants of wild type and hmgA mutant strains of V. cholerae ...... 66

Figure 2-5 Early biofilms of V. cholerae A1552 exposed to T. pyriformis for 72 hours. Biofilm biomass was determined by CV staining (A) and the planktonic cells in the supernatant enumerated by the drop plate method (B) ...... 67

Figure 2-6 Amount of pyomelanin produced by ungrazed or grazed established biofilms after 3 d exposure to A. castellanii (A) or T. pyriformis (B)...... 68

Figure 2-7 Pyomelanin production by V. cholerae wild type (A) and hmgA mutant (B) biofilms ...... 70

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Figure 2-9 Effect of cell-free supernatants (A), H2O2 (B) and catalase (C) on grazing resistance to A. castellanii ...... 73

Figure 2-10 Number of intracellular V. cholerae after 24-hour co-incubation with A. castellanii ...... 75

Figure 3-1 A selective grazing assay on day 7...... 83

Figure 3-2 Biofilm biomass of V. Vulnificus clinical C-genotype (MO6-24, CMCP6, C7184, YJ016), clinical E-genotype (E64MW, LSU2098, LSU1657, LSU549) and environmental E-genotype (JY1305, JY1701, Env1, SS108-A3A) strains exposed to grazing by T. pyriformis for 72 hours ...... 89

Figure 3-3 Biofilm biomass of clinical C-genotype (MO6-24, CMCP6, C7184, YJ016), clinical E-genotype (E64MW, LSU2098, LSU1657, LSU549) and environmental E- genotype (JY1305, JY1701, Env1, SS108-A3A) V. vulnificus strains exposed to grazing by A. castellanii for 72 hours ...... 90

Figure 3-5 Grazing resistance of clinical C-genotype (MO6-24, CMCP6, C7184, YJ016), clinical E-genotype (E64MW, LSU2098, LSU1657, LSU549) and environmental E- genotype (JY1305, JY1701, Env1, SS108-A3A) V. vulnificus isolates exposed to T. pyriformis for 24 hours ...... 94

Figure 3-6 The graphical map of Env1 genome ...... 99

Figure 3-7 Phylogenetic tree of sequenced V. vulnificus genomes ...... 101

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Figure 3-8 Circular maps that compare the genomes of three grazing sensitive V. vulnificus strains, MO6-24, CMCP6 and YJ016 with V. vulnificus Env1 ...... 103

Figure 3-9 Whole-genome alignments of V. vulnificus Env1, CMCP6, MO6-24/O and YJ016 ...... 108

Figure 3-10 Genome of V. vulnificus Env1 was compared to V. cholerae N16961, V. vulnificus CMCP6, and V. vulnificus YJ016 ...... 109

Figure 3-11 T. pyriformis after co-incubation with V. vulnificus Env1 cell-free supernatant ...... 119

Figure 3-12 Effect of cell-free supernatants from V. vulnificus Env1 on the health of T. pyriformis and A. castellanii ...... 120

Figure 4-1 Growth of V. vulnificus Env1 ...... 134

Figure 4-2 Effect of ammonia on survival of T. pyriformis ...... 138

Figure 4-4 Biomass of V. vulnificus Env1 wild type and purA and copA mutants exposed to grazing by T. pyriformis for 24 hours ...... 145

Figure 4-5 Biomass of V. vulnificus Env1 wild type and arcA and smcR mutants exposed to grazing by T. pyriformis for 24 hours ...... 149

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List of tables

Table 1-1 A summary of the virulence factors reviewed in this chapter...... 32

Table 2-1 Strains and plasmids used in this study...... 52

Table 2-2 Primers used in this study...... 57

Table 3-1. List of bacterial and protozoal strains...... 81

Table 3-2 Position and length of genes potentially involved in virulence, disease and defence in V. vulnificus Env1 ...... 112

Table 3-3 Proteins for iron uptake and utilisation...... 114

Table 3-4 Health of T. pyriformis, 1 hour after the addition of each treated cell-free supernatant ...... 121

Table 3-5 Emulsification activity of cell free supernatant of V. vulnificus Env1 using n- hexadecane ...... 122

Table 4-1 List of bacterial and protozoal strains...... 127

Table 4-2 Genes involved in ammonia accumulation, production and transport ...... 137

Table 4-3 Amount of ammonia produced by V. vulnificus under the following conditions ...... 138

Table 4-4 Genes involved in biotin biosynthesis ...... 140

Table 4-5 Genes involved in pathogenicity and virulence regulation ...... 142

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List of appendices

Appendix 1 List of differentially expressed transcripts in Chapter 2 ...... 196

Appendix 2 List of strain and NCBI reference sequence used in the genomic analysis ...... 202

Appendix 3 Whole-genome alignments of V. vulnificus Env1, CMCP6, MO6-24/O and YJ016, JY1305, JY1701 and E64MW ...... 204

Appendix 4 List of differentially expressed transcripts in Chapter 4 ...... 205

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List of abbreviations

AI Autoinducer

ANOVA Analysis of variance

ATCC American type culture collection

BLAST Basic local alignment search tool

C Celsius

cAMP cyclic adenosine monophosphate

CDS Coding DNA sequence

CFU Colony forming unit

cm Centimetre

Cm Chloramphenicol

CPS Capsular polysaccharide

CT Cholera toxin

CV Crystal violet

EPS Extracellular polymeric substances

Fur Ferric uptake regulator

g Gram

GFP Green fluorescent protein

h Hour

HGT Horizontal gene transfer

LB Lysogeny broth

LD50 50% Lethal Dose

min Minute

ml Millilitre

µl Microlitres

XVII | P a g e mm Millimetre mM Millimolar

µM Micromolar

MSHA Mannose-sensitive haemagglutinin

MARTX Multifunctional autoprocessing repeats-in-toxin nM Nanomolar

NSS Nine salts solution

OD Optical density

O Opaque

ORF Open reading frame

PCR Polymerase chain reaction rpm Revolutions per minute

ROS Reactive oxygen species

RT Room temperature

SD Standard deviation

T Translucent

T1SS Type I secretion system

T6SS Type VI secretion system

TCP Toxin-coregulated pili

VAS Virulence associated secretion

VBNC Viable but nonculturable

VNSS Väätänen nine salts solution

WT Wild type

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1 Chapter 1: General introduction and literature review

1.1 General introduction

Our understanding of extracellular pathogens is blurred due to a tendency to focus on a pathogen’s life cycle and evolution in the human host rather than in their natural environmental niche. The evolutionary timeline of life on earth has yet to be accurately determined; so far, according to current scientific theory, life on earth began with simple prokaryotic cells about 3.6 billion years ago. The first eukaryotic cells evolved about two billion years ago, while only in the last 2.5 million years, has the genus Homo existed. This history of bacteria and protozoa interacting for millions of years led to coevolution of both predator and prey (Matz & Kjelleberg, 2005, McFall-Ngai et al., 2013).

The interactions between bacteria and protozoa are complex. Heterotrophic protists are the biggest consumers of bacteria in the environment, and a major mortality factor for bacteria (Jürgens & Matz, 2002). In contrast, protozoan grazing is a major contributor to nutrient cycling in the microbial food web, thus contributing to microbial growth in the environment (Sherr et al., 1982). Due to the long history of co-evolution, bacteria have developed many different mechanisms that allow for protection against grazing by protozoa, leading to the prevalence of bacteria that are grazing resistant (Matz & Kjelleberg, 2005). The coincidental evolution hypothesis states that many pathogenic bacteria have acquired anti-protozoal mechanisms in their natural niche that may also function as virulence factors in plant and animal hosts.

In this study, the aim was to assess these grazing resistance mechanisms in model members of the genus Vibrio, and to evaluate the role of pathogenicity factors in environmental survival with respect to protozoan grazing. Vibrio is a genus of Gram- negative bacteria typically found in brackish waters that have great impact on human health and aquaculture. Here, two of the most important members of this genus, Vibrio vulnificus and Vibrio cholerae were selected for further investigation. V. vulnificus is an estuarine, opportunistic and ferrophilic pathogen responsible for diarrhoea, wound infections and septicaemia following ingestion of contaminated seafood or through

1 | P a g e exposure of a wound to seawater. This bacterium has the highest reported mortality rate of any seafood-related disease (Jones & Oliver, 2009). V. cholerae, the causative agent of cholera, persists in brackish and estuarine water systems. Cholera is an acute diarrhoeal illness caused by ingestion of contaminated food or water. Even with the recent developments in public health management, cholera remains a global threat due to poor hygiene and lack of basic health infrastructure in impoverished countries or after natural disasters (Huq et al., 1990, Colwell & Huq, 1994, Lutz et al., 2013, Ali et al., 2015, Sun et al., 2015).

Vibrio species are natural inhabitants of coastal marine environments where they are exposed to predation by heterotrophic protozoa. This chapter reviews the organism’s physiological characteristics as well as the wide array of virulence factors used by the Vibrio spp. (Jones & Oliver, 2009). Protozoan predators and effects of predation on bacterial populations will be introduced and discussed. The mechanisms by which bacteria are protected against protozoa predation will be investigated.

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1.2 Description of Vibrio vulnificus and Vibrio cholerae

1.2.1 Vibrio vulnificus

V. vulnificus is an autochthonous inhabitant of coastal marine environments globally and has been isolated from variety of sources including water, sediments, and a range of animals, including shrimp, fish, oysters, clams and zooplankton (crustaceans, mainly copepods) (Oliver et al., 1982, Tamplin et al., 1982, Myatt & Davis, 1989, DePaola et al., 1994, Wright et al., 1996, Høi et al., 1998, Bisharat et al., 1999, Baffone et al., 2006, Mahmud et al., 2008, Ji et al., 2011). This opportunistic pathogen can enter the human body through open wounds when exposed to water or by puncture from fish spines and crustacean shells, or by consumption of contaminated seafood (especially raw or undercooked oysters) (Johnston et al., 1985, DePaola et al., 1994, Jones & Oliver, 2009).

The first documented case of V. vulnificus may date back to the 5th century BC when Hippocrates described a fatal foot wound infection of a man living by the Aegean Sea. Symptoms included a swollen foot with black blisters on the skin leading to sepsis and death within 48 hours of the onset of infection (Baethge & West, 1988).

The first modern clinical case of V. vulnificus infection was reported in 1970 and the organism responsible was erroneously diagnosed as Vibrio parahaemolyticus. The patient developed endotoxin-related septic shock after exposure to New England coastal waters (Roland, 1970). The patient experienced vomiting and diarrhoea, and had a generalised papular haemorrhagic rash and a leg wound infection that became gangrenous and required amputation. Hollis et al. (1976) described the laboratory characteristics of V. vulnificus while the clinical characteristics were reported by Blake et al. (1979). In 1979 Farmer officially named the pathogen V. vulnificus as vulnificus is a Latin word meaning "inflicting wounds" (Farmer, 1979).

Ingestion of V. vulnificus can cause vomiting, diarrhoea, and abdominal pain in healthy individuals while in immunocompromised patients, particularly those with chronic liver disease, V. vulnificus can infect the bloodstream, causing a severe and potentially fatal systemic infection characterised by fever and chills, decreased blood pressure (septic

3 | P a g e shock), and blistering skin lesions. Wound infections may progress rapidly to cellulitis, ecchymosis, and bullae, which can result in necrotising fasciitis at the site of infection. Even with aggressive antibiotic chemotherapy, mortality rates for septicaemia can be as high as 75%, and for wound infection can be as high as 50% (Blake et al., 1979, Klontz et al., 1988, Hlady & Klontz, 1996).

Data collected between 1988 and 1996 from 422 reported infections (Shapiro et al., 1998) and 459 reported cases between 1992 and 2007 (Belkin & Colwell, 2006) revealed a 51.6 - 61% fatality rate from primary septicaemia. Interestingly 85.6 - 86 % of patients were male, and 96% with primary septicaemia from the consumption of raw oysters. Underlying liver disease was associated with fatal outcome of the patients as 95.3% had some pre-existing disease(s), such as cirrhosis, hepatitis and hemochromatosis (all of these result in excess iron in blood) (Wright et al., 1981, Belkin & Colwell, 2006, Jones & Oliver, 2009).

Considering that 85% of patients that develop endotoxic shock from V. vulnificus are males, the role of oestrogen in advancement of the infection was investigated. It was shown that gonadectomy of female rats resulted in increased mortality when challenged with V. vulnificus, and oestrogen replacement resulted in decreased mortality in gonadectomised females as well as males. These data indicate that oestrogen provides protection against V. vulnificus lipopolysaccharide-induced endotoxic shock (Merkel et al., 2001).

1.2.2 V. vulnificus genotypes and biotypes

V. vulnificus strains exhibit considerable variation in genotypes as well as phenotypes, hence various attempts have been made to categorise them using several biotypic and genotypic classifications. V. vulnificus strains are classified into three biotypes based on their biochemical characteristics. Strains belonging to biotype 1 are mostly responsible for human infections, while biotype 2 strains are primarily eel pathogens (Tison et al., 1982, Amaro & Biosca, 1996). Biotype 3 causes human wound infections and seems to

4 | P a g e be geographically limited to Israel. Genomic analysis indicated that biotype 3 is a hybrid of biotypes 1 and 2 (Bisharat et al., 1999, Naiel et al., 2005).

Biotype 2 expresses only one type of lipopolysaccharide (LPS)-based homogeneous O serogroup (serovar E) (Biosca et al., 1996, Biosca et al., 1997), while serologically different types of LPS are detected among biotype 1 strains (Shimada & Sakazaki, 1983, Martin & Siebeling, 1991). An investigation of the toxicity of LPS showed binding affinity for both eel and human erythrocyte membranes that led to agglutination of the cells. The injection of pure LPS only caused endotoxic effects and death in rats but not in eels (Biosca et al., 1999).

Another attempt to categorise V. vulnificus isolates, grouped them into two lineages based on multilocus sequence typing of six housekeeping genes, and this division was confirmed using 16S rRNA and VV0401 gene sequencing. Lineage I contained only biotype 1 isolates and had a higher proportion of clinical isolates, whereas lineage II contained some biotype 1 and all biotype 2 isolates. Lineage I isolates carried a 33- kbp genomic island (region XII), one of three regions identified by genome comparisons as being unique to the species. This genomic island may give isolates a fitness advantage in the human host or in the aquatic environment, or both (Cohen et al., 2007).

Another classification scheme was based on a 200 bp randomly amplified polymorphic DNA (RAPD) PCR amplicon that is associated with V. vulnificus. Each V. vulnificus strain produced a unique banding pattern, indicating that the members of this species are genetically quite heterogeneous (Warner & Oliver, 1999). The sequence of the virulence correlated gene (vcg) locus distinguishes environmental (from oysters, clams and shrimp, sea water and sediment) and clinical isolates. The “environmental isolates” possess the vcgE allele (E-genotypes) while the “clinical isolates” have the vcgC allele (C- genotypes). It was shown that 90% of the C-genotype strains were clinical isolates, while 93% of environmental isolates were classified as E-genotype (Rosche et al., 2005, Warner & Oliver, 2008a). A closer look at V. vulnificus populations showed that 84.4% of isolates recovered from oysters contained the vcgE allele. In contrast, isolates from waters surrounding the oyster sites revealed an almost equal distribution of the two genotypes. Interestingly, the percentage of C-genotype strains from both sources increased when the water temperatures rose (Warner & Oliver, 2008b). 5 | P a g e

Further work based on alignment of eight housekeeping and virulence loci of V. vulnificus clustered strains based on their genotype (C- or E-genotype), suggesting possible different ecotypes. The authors speculated that the E-genotype strains could grow better under conditions present in oysters, whereas the C-genotype strains may be favoured during the stressful transition from seawater/oyster to humans. Therefore, evolution of strains in different niches gave rise to the two genotypes of V. vulnificus (Rosche et al., 2010).

One of the significant differences in the physiology of the E and C-genotypes strains is the ability of C-genotypes to resist the bactericidal effects and even grow in human serum, whereas E-genotypes strains are sensitive to the bactericidal effects (Bogard & Oliver, 2007). However, genotypes do not strictly predict the pathogenicity of V. vulnificus biotype 1 strains (Thiaville et al., 2011).

Recently, a phylogenomic analysis of the core genome of 80 strains was undertaken in order to improve classification of V. vulnificus. This resulted in the identification of five phylogenetic groups or lineages (L). The L1 lineage contains biotype 1 strains of both clinical and environmental origin which are most responsible for cases infection in humans. The L2 lineage contains a mixture of Biotype 1 and 2 strains from various sources relating to disease in eels. The L3 lineage includes only biotype 3 strains and L4 and L5 contains limited numbers of biotype 1 associated with specific geographical areas (Roig et al., 2018)

1.2.3 Distribution of V. vulnificus

V. vulnificus has been isolated from estuarine waters, sediments and animals, including crustacea, molluscan shellfish, and from the intestines of finfish obtained from the East and Gulf Coasts of the United States (Oliver et al., 1983, O'Neill et al., 1992, Tamplin & Capers, 1992, DePaola et al., 1997).

The occurance of V. vulnificus is strongly correlated with temperature, turbidity, levels of dissolved oxygen and numbers of estuarine and coliform bacteria. V. vulnificus has been recovered from water with salinities between 0.4 and 3.7% with an optimum between 1.0

6 | P a g e and 2.5% and a temperature range of 7 to 36°C with an optimum of 20°C (Høi et al., 1998, Motes et al., 1998, Pfeffer et al., 2003).

V. vulnificus numbers increase in seawater in the temperature range of 13 to 22°C but at temperatures below 8.5°C, its survival decreases (Kaspar & Tamplin, 1993). V. vulnificus can survive in shucked Pacific (Crassostrea gigas) and Eastern (Crassostrea virginica) oysters stored at 10°C and below, demonstrating that they survive refrigeration (Kaysner et al., 1989). V. vulnificus was consistently detected in fish intestines, but infrequently detected at lower levels in oysters and during the less favourable periods (Givens et al., 2014).

V. vulnificus isolation is correlated with lower salinities and with isolation from samples collected closer to the bottom (Wright et al., 1996). V. vulnificus was not recovered from deionized water, indicating lysis, and highlighting the requirement for salts for maintenance of osmotic balance (Kaspar & Tamplin, 1993). In a recent study, pathogenic V. vulnificus was detected in 14% of samples from the Sydney harbour estuary, with its occurrence restricted to the late summer and a salinity range of 5 to 26 ppt (Siboni et al., 2016).

V. vulnificus is known to enter a viable but nonculturable (VBNC) state, wherein the cells are no longer culturable on routine media but can be shown to be viable and metabolically active. V. vulnificus remains virulent, at least for some time, when in the VBNC state following in vivo resuscitation (Oliver & Bockian, 1995). This is in contrast to starved bacteria which can grow in or on normal media after a period of inactivity (Colwell et al., 1985). The expression of pathogenicity factors still occurs in VBNC cells. For example, constitutive transcription of the gene encoding a haemolysin, vvhA, was detected in VBNC V. vulnificus cells (Saux et al., 2002).

V. vulnificus expresses the global stress regulator, RpoS (σS) in the VBNC state (Smith & Oliver, 2006), and perhaps other stress-related genes that can provide cross protection against various other stresses (Nowakowska & Oliver, 2013). VBNC cells of V. vulnificus show increased resistance to high temperature, low and high pH, oxidative and osmotic stress, and exposure to ethanol, zinc, chloramphenicol and ampicillin when compared to culturable cells (Nowakowska & Oliver, 2013).

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1.2.4 Pathogenicity factors

Extracellular pathogens such as V. vulnificus that mainly live in natural non-infective ecosystems have acquired classical virulence determinants through either mutation or horizontal gene transfer (HGT). The fact that these factors are maintained in the genome, suggests that they are likely to play dual roles in both of the pathogen’s life styles. For example, factors such as iron uptake and metabolism can be advantageous inside the human body as well as the aquatic environment (Martínez, 2013, Martínez, 2014). Here, the factors that V. vulnificus employs to cause infection in humans are described, followed by a discussion of the potential role that these factors may play in successful survival in aquatic environments.

V. vulnificus factors that have been implicated as possible virulence determinants include the ability to form biofilms, expression of capsular polysaccharide, motility, iron acquisition systems, degradative enzymes and cytotoxic factors (including haemolysin, elastase, collagenase, DNase, lipase, phospholipase, mucinase, chondroitin sulfatase, hyaluronidase, fibrinolysin and proteases with activity against native serum albumin) (Linkous & Oliver, 1999, Tolker-Nielsen et al., 2000, Jones & Oliver, 2009).

1.2.4.1 Cell surface factors

V. vulnificus evades host defences by expression of an antiphagocytic capsular polysaccharide (CPS) (Kreger et al., 1981). The virulent, encapsulated strains have an opaque (O) colony morphology whereas the non-virulent translucent (T) colonies produce less or no CPS (Hayat et al., 1993). The lethal dose 50% (LD50) for iron overloaded mice is reported to be a single cell for O strains, while increasing to 6.5 × 107 cells for non- encapsulated cells (Wright et al., 1981). It should be noted that the T colonies of biotype 2 (eel pathogen) are still lethal in the mouse model (Simpson et al., 1987).

V. vulnificus O colonies may switch to T colony types that express less CPS and are less virulent (Simpson et al., 1987). Irreversible phase variation of O to T variants has been observed to be dependent on the amount of aeration and temperature as well as mutations in rpoS (Hilton et al., 2006). The presence of cations in the environment (e.g. calcium

8 | P a g e and manganese) substantially increases O to T phenotypic switching (Garrison-Schilling et al., 2011, Kaluskar et al., 2015). Moreover, E-genotypes show much higher conversion rates than C-genotypes (Hilton et al., 2006). However, oxidative stress, catalase and antioxidants, quorum sensing, growth in cell-free spent media, nutrient rich or poor media does not affect conversion rates (Hilton et al., 2006).

The rfaH gene encodes a protein important for antitermination control that promotes RNA polymerase to proceed past Rho-dependent termination sites without interrupting transcription. The operons under RfaH control possess an operon polarity suppressor (ops) element usually found upstream of the first gene of the operon (Bailey et al., 1997). Previously an ops element was identified within the group I CPS operon of V. vulnificus (Wright et al., 2001). A mutant in rfaH was observed to have reduced capacity to switch phases. Moreover, CPS production and serum resistance was greatly reduced (Garrett et al., 2016).

The production of CPS in V. vulnificus is dependent upon a variety of genetic loci. The V. vulnificus CPS locus is composed of wza, wzb and wzc genes, which are highly conserved among the Enterobacteriaceae (Whitfield & Paiment, 2003). The wza gene encodes a CPS transporter and does not have a role in CPS biosynthesis while the wzb gene encodes a phosphatase required for CPS expression (Wright et al., 2001). The three genes, wcvA, wcvF and wcvI are responsible for biosynthesis, polymerisation, and transport of CPS and are clustered on a single chromosomal fragment, while a fourth gene (ORF4) is encoded on an integron-gene cassette (Smith & Siebeling, 2003).

A third, intermediate (Int) colony morphotype, in which the colonies are less opaque than O colonies but are not fully translucent, results from decreased wzb expression. Int strains can switch to a T (wzb negative) morphotypes (Rosche et al., 2006). A rugose morphotype of V. vulnificus has been reported to occur when cells are cultured repeatedly at temperatures below 37°C and which, once formed, persists even at 37°C. The rugose morphotype forms aggregated pellicles that sink to the bottom of test tubes in liquid culture. A mutation in wcvA that is essential for capsule production, did not affect the rugose morphotype which indicates that the rugose morphotype is independent of CPS production (Grau et al., 2005). Both O rugose and Int rugose phenotypes form biofilms with large amounts of biomass (Grau et al., 2008).

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It has been demonstrated that capsule production negatively affects the ability of V. vulnificus biotype 1 to attach and form biofilms in a microtiter plate assay. This is in contrast to many other species, including V. cholerae, where rugose morphotypes form more dense biofilms. Thus, it appears that the V. vulnificus CPS inhibits surface attachment and therefore, biofilm formation and indeed, the T variants produce significantly more biofilm biomass under all conditions tested than O variants (Joseph & Wright, 2004).

One of the most important factors in V. vulnificus infection in humans is LPS-associated endotoxic shock. Intravenous injections of V. vulnificus LPS into mice caused a decrease in the mean arterial pressure (McPherson et al., 1990). LPS is a known pyrogen that elicits a cytokine response in mice and causes the release of tumour necrosis factor alpha (TNF-α) which in turn controls the level of type 1 immunity that contributes to sepsis and refractory septic shock (McPherson et al., 1990, Powell et al., 1997). Low-density lipoprotein (LDL), cholesterol and oestrogen reduce the effects of LPS, as injection of LDL prior to injection of LPS increased mouse survival and delayed death in fatal cases (Park et al., 2007). This is similar to observations that oestrogen protects females against the endotoxic activity of V. vulnificus LPS (Merkel et al., 2001). To date, no mechanism has been described for the decreased effect of LPS when LDL and oestrogen are present (Jones & Oliver, 2009).

Initial attachment of bacterial cells to surfaces, which is often dependent on pili, is an important step in the colonisation of host surfaces (Strom & Lory, 1993). In V. vulnificus, a type IV prepilin peptidase/N-methyltransferase encoded by pilD (previously known as vvpD), is responsible for the maturation of the type IV pre-pilin. Mutation of pilD results in reduced adherence to human epithelial (HEp-2) cells, decreased virulence and a block in secretion of several exoenzymes, including cytolysin, metalloprotease and chitinases which are secreted via a type II secretion pathway (Paranjpye et al., 1998).

Another type IV pilin, PilA, is encoded in an operon consisting of three other pilus biogenesis genes, pilBCD. A mutant in pilA showed a significant reduction in the formation of biofilms on borosilicate glass and a slight reduction in biofilm formation on polystyrene and polyvinyl chloride surfaces, as well adherence to HEp-2 cells. This mutant also exhibited decreased virulence in the mouse model of infection (Paranjpye &

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Strom, 2005) and in colonisation of oysters (Paranjpye et al., 2007). It should be noted that pili are still present on the surface of the pilA mutant strain with no visible difference in the total number or morphology of the pili as compared to those expressed by the wild- type cells. This could be due to compensation by another type IV pili, such as the mannose-sensitive haemagglutinin (MSHA) pilus (Paranjpye & Strom, 2005).

E-genotype strains attach to chitin significantly better than C-genotype strains at 20°C (environmental temperature) while at 37°C (human body temperature) C-genotypes attach better. The expression of the type IV pilin genes pilA, pilD, and mshA which are implicated to be involved in chitin adherence, were shown to be higher in E-genotype strains compared to C-genotype strains even in the absence of chitin. In contrast, the level of expression of the gene encoding N-acetylglucosamine binding protein A (gbpA) was significantly higher in C-genotype strains (Williams et al., 2014). Attached C-genotypes cells detach significantly more from chitin compared to E-genotypes cells after 24 hours and type IV pilin production was significantly down-regulated in C-genotype strains compared to E-genotype strains. In addition, gbpA was up-regulated in the C-genotypes during detachment (Phippen & Oliver, 2015a). E-genotype strains differentially express capsule and attachment genes compared to C-genotypes, both aerobically and anaerobically. A lack of oxygen results in reduced CPS and biofilm formation as result of down regulation pilA, pilD, and mshA genes in both E- and C- genotype strains (Phippen & Oliver, 2015b).

The recently described Tad type IV pilus is frequently found in Vibrio spp.. The V. vulnificus genome encodes three distinct tad loci. In V. vulnificus, tad-3 expression correlated with increased biofilm formation, auto-aggregation, and oyster colonisation. It was shown that tad-3 promoted initial surface attachment, auto-aggregation and resistance to mechanical clearance of V. vulnificus biofilms (Pu & Rowe-Magnus, 2018b, Pu & Rowe-Magnus, 2018a).

Flagellin, the flagellar structural subunit, binds to Toll-like receptor 5 (TLR5) and thus activates immune responses (Lee et al., 2006). A non-motile flagellum-deficient flgE mutant, which encodes the flagellum component, and a flgC mutant, which encodes a flagellar basal body showed reduced adherence to mammalian cells and abiotic surfaces as well as reduced lethality to mice (Ran Kim & Haeng Rhee, 2003, Lee et al., 2004).

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The reduction in the pathogenicity was attributed to the loss of motility, which leads to decreased adhesion and inhibition of cytotoxin delivery (Ran Kim & Haeng Rhee, 2003, Lee et al., 2004, Jones & Oliver, 2009).

1.2.4.2 Biofilm formation

Many bacteria favour a surface-associated lifestyle due to its protective nature. Vibrio spp. have the ability to form biofilms, surface-associated bacterial communities that are embedded in extracellular polymeric substances (EPS) (Donlan & Costerton, 2002, Yildiz & Visick, 2009). Although V. vulnificus has been reported to form biofilms on a variety of biotic and abiotic surfaces, the role of biofilms in pathogenicity of this species has not been extensively studied. V. vulnificus biovar 2 (serotype E) has been reported to form biofilms on eel skin and epidermal cells, as well as hydrophobic and hydrophilic surfaces when eel mucus was used as a growth medium. Furthermore, the strain formed biofilms at the air-liquid interface and at the bottom of the wells or tubes (Marco-Noales et al., 2001). V. vulnificus biofilm formation and development on abiotic polystyrene microtiter plates is affected by nutrient and glucose concentrations, but not by salt concentration or temperature (McDougald et al., 2006). Moreover, a quorum sensing (QS) mutant showed rapid biofilm development with a different biofilm architecture from the wild type strain (McDougald et al., 2006).

Many regulatory pathways control biofilm formation. The RNA-binding protein carbon storage regulator (CsrA) is a global regulatory protein that inhibits biofilm formation by V. vulnificus. V. vulnificus csrA mutant strains have been isolated from shellfish, and these strains form more biofilm biomass compared to csrA-positive strains (Jones et al., 2008). In contrast, elevated levels of the secondary messenger cyclic di-GMP (c-di- GMP) promote the production of EPS that is distinct from the CPS, and that leads to an enhancement in biofilm formation and rugose colony morphology in a CPS-independent manner. However, the production of EPS does not compensate for the loss of CPS production in virulence assays (Nakhamchik et al., 2008, Park et al., 2015a).

CabA is an extracellular matrix calcium-binding protein crucial for the structural integrity of robust biofilms in flow cells and on oyster shells. This protein was identified to be a

12 | P a g e component of the biofilm matrix of V. vulnificus that contributes to the development of biofilms and rugose colony morphology under elevated c-di-GMP conditions and is secreted through functional CabB and CabC proteins (Park et al., 2015b).

Expression of the brp exopolysaccharide locus containing nine genes responsible for mediating surface adherence, is controlled by c-di-GMP, as well as the regulator BrpT. C-di-GMP and BrpT regulate the expression of the cabABC operon, brpT and another VpsT-like transcriptional regulator, brpS (Guo & Rowe-Magnus, 2010, Chodur et al., 2017, Chodur & Rowe-Magnus, 2018).

1.2.4.3 Quorum sensing (QS)

QS systems control gene expression in a density-dependent manner, and were originally described in Vibrio fischeri by Nealson et al. (1970). QS acts on the production, release, accumulation, and detection of extracellular signal molecules called autoinducers. At low cell density when levels of autoinducers are low, bacteria express phenotypes that are beneficial to individual cells, but at high cell density when levels of autoinducers are high, bacteria express phenotypes that are beneficial for the population (Hurley & Bassler, 2017). One of the most studied examples of this phenomenon is bioluminescence by V. fischeri at high cell density in the light organ of the squid host (Nealson & Hastings, 1979, Verma & Miyashiro, 2013). Another example of this phenomenon is in infection by V. cholerae where at low cell density, virulence factors, such as the toxin co-regulated pilus (TCP), are expressed while at high cell density, TCP is repressed. This facilitates the V. cholerae cells leaving the human host in preparation for returning to the aquatic niche (Higgins et al., 2007).

Some Gram-negative bacteria use the acylated homoserine lactone (AHL) (Swift et al., 1993, Cha et al., 1998) QS system, where the LuxI autoinducer synthase and the LuxR response regulator activate target genes (Engebrecht & Silverman, 1984, Ng & Bassler, 2009). In the autoinducer 2 (AI-2) system, LuxS is the synthase for the AI-2 signal, a furanosyl borate diester. AI-2 binds to the periplasmic LuxP protein, which can then interact with the hybrid sensor/kinase, LuxQ. At high cell density, LuxU and LuxO are deactivated due to loss of phosphate and LuxR is activated (Cámara et al., 2002, Milton,

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2006). In the V. vulnificus QS system the luxR homologue is called smcR (vvpR) (McDougald et al., 2000). A mutant in smcR showed defects in starvation survival (McDougald et al., 2001) and oxidative stress resistance (McDougald et al., 2003). Loss of smcR resulted in increased expression of stationary phase phenotypes and increases in exoproteases, motility, fimbria production and biofilm formation (McDougald et al., 2001). In contrast to hapR in V. cholerae, smcR is involved in the repression of protease expression during exponential growth rather than its induction and is not responsible for the induction of AI-2 signalling product (McDougald et al., 2001). Biofilm of the smcR mutant were impaired in virulence and colonisation capacity during infection of mice, resulting in decreased histopathological damage to mouse jejunum tissue (Kim et al., 2013b).

There is no evidence that there is an AHL QS system in V. vulnificus but V. vulnificus has signalling activity that can induce luminescence expression in Vibrio harveyi through the AI-2 signalling system. The AI-2 synthase gene (luxSVv) is 80% identical to that of V. harveyi (luxSVh) at the amino acid level (Kim et al., 2003a). This suggests the V. vulnificus QS system as described in Figure 1-1.

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Figure 1-1 Model of the V. vulnificus quorum-sensing system (Milton, 2006). See text for explanation.

1.2.4.4 Iron acquisition

V. vulnificus human infections are highly correlated with elevated serum iron levels. Virulence assays in mice have shown that the infectious dose as well as time of death after infection, is directly correlated with the level of serum iron (Wright et al., 1981). Although C-genotype strains are superior in human serum survival compared to the E- genotype strains, the addition of exogenous iron allows both genotypes to survive equally well (Bogard & Oliver, 2007).

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The role of iron in V. vulnificus pathogenicity has been the subject of many studies as iron limitation is one of the main obstacles for growth of V. vulnificus in human serum. The 6 addition of excess iron to mice in infection models lowered the LD50 from 10 cells to 1.1 cell (Wright et al., 1981). One study showed that excess iron results in decreased neutrophil activity in the host and thus, leads to a compromised immune response (Hor et al., 2000). Another study demonstrated that the presence of excess iron dramatically increased the growth rate of clinical strains, suggesting that excess iron enhances growth of this pathogen (Starks et al., 2006). These findings indicate multiple roles for elevated serum iron levels in enhancing V. vulnificus infection (Jones & Oliver, 2009).

One of the most important iron acquisition mechanisms employed by V. vulnificus is the siderophore, vulnibactin (Simpson & Oliver, 1983, Okujo et al., 1994). Vulnibactin in addition to an exocellular protease are involved in the utilisation of transferrin and lactoferrin-bound iron (Okujo et al., 1996) as it acquires iron from transferrin (Kim et al., 2006). Mutation in vuuA, an iron-regulated outer membrane vulnibactin receptor, resulted in loss of uptake and use of transferrin or vulnibactin as well as reduced virulence in an infant mouse model (Webster & Litwin, 2000). V. vulnificus L-180, a strain isolated from the blood of a septic human patient (Miyoshi et al., 1987), produces a significant amount of metalloprotease that, in addition to vulnibactin, is effective in the utilisation of Fe2+ bound to transferrin and lactoferrin. Protease-deficient mutants are not able to assimilate iron bound to the proteins and addition of the protease to the mutant rescued the phenotype (Nishina et al., 1992, Okujo et al., 1996).

The haem uptake receptor (HupA) has a role in virulence in mice and in tissue culture cells (Oh et al., 2009). A mutant in hupA, similar to TonB-dependent outer membrane receptors, did not produce a 77-kDa protein and could not utilise haemin or haemoglobin as a source of iron (Litwin & Byrne, 1998). The hupR gene encodes a ferric uptake regulator (Fur) binding site and acts as a positive regulator of hupA under low iron conditions when haemin is present (Litwin & Quackenbush, 2001). Changes in growth temperature as well as iron availability acts as environmental cues controlling the expression of the hupA gene. It has been shown that the cyclic AMP receptor protein (CRP) activates the expression of hupA at the transcriptional level (Oh et al., 2009).

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Fur is a metal-dependent regulator found in many Gram-negative bacteria that controls expression of many iron-responsive genes (Escolar et al., 1999). A fur deletion mutant overexpressed iron-regulated outer membrane proteins (Litwin & Calderwood, 1993a). In an study by Kim, et al. (2006) a mutation in the fur gene de-repressed expression of the vulnibactin-mediated iron uptake system and vulnibactin production, and facilitated iron assimilation from transferrin (Kim et al., 2006). The Fur-box is located within the promoter region and the Fur-binding site was shown to be 5’- AATGANAATNATTNTCATT-3’ (Ahmad et al., 2009).

Fur boxes are associated with a small regulatory RNA gene, ryhB in Vibrio spp. Three Fur boxes were predicted to occur in the promoter region of the ryhB in Vibrio salmonicida. RyhB in V. cholerae negatively regulates a number of genes involved in various metabolic processes while ryhB is itself repressed by Fur (Davis et al., 2005, Mey et al., 2005). In V. vulnificus, Fur is a positive regulator and binds to an AT-rich sequence upstream of fur under iron-limited conditions, as demonstrated by DNase I foot-printing and mutagenesis analyses (Lee et al., 2007a). Kim et al. (2013a) suggested that two biologically important environmental signals, Fur iron-sensing and SmcR QS coordinates the expression of virulence factors as illustrated in Figure 1-2.

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Figure 1-2 The regulation of virulence genes by SmcR and Fur in response to iron and population density in V. vulnificus (Kim et al., 2013a).

The gene, vvpE, encoding the virulence factor elastase is a part of the QS regulon in V. vulnificus and is repressed under iron-rich conditions. Repression is mediated by Fur- dependent repression of smcR, the QS master regulator. The Fur-iron complex binds reversibly to two regions upstream of smcR to allow expression of QS-regulated genes at high cell density under iron-rich conditions. Under iron-limiting conditions, Fur fails to bind either region and the expression of smcR is regulated solely by QS (Kim et al., 2013a).

1.2.4.5 Degradative toxins and enzymes

V. vulnificus can secrete multifunctional autoprocessing repeats-in-toxin (MARTX) toxins, MARTXVv which are involved in the pathogenesis of Vibrio spp. The V. vulnificus

18 | P a g e rtx gene cluster is comprised of rtxA1, rtxC, rtxB1, rtxD, rtxE and one hypothetical ORF. RtxB1, RtxD and RtxE are hypothesised to act as an ABC transporter while RtxC seems to be the activator of the RtxA1 toxin molecule (Kim et al., 2008, Lee et al., 2008b).

V. vulnificus produces at least four different types of MARTX (types I–IV) where the plasmid-encoded MARTX type III (or RtxA13) is structurally and evolutionarily different from MARTX types I and II (Kwak et al., 2011, Roig et al., 2011). The V. vulnificus biotype 2 MARTX type III is involved in the lysis of a wide range of eukaryotic cells, including amoebae, erythrocytes, epithelial cells and phagocytes after bacterium–cell contact (Lee et al., 2013a).

V. vulnificus RtxA1 can cause contact-dependent acute cytotoxicity and haemolysis of host cells (Lee et al., 2007b). RtxA1 can cause damage in a variety of ways, including causing cytoskeleton rearrangement (Kim et al., 2008), apoptotic cell death (Lee et al., 2008a), generating of reactive oxygen species (ROS) (Chung et al., 2010), protecting against phagocytosis (Lo et al., 2011) and modification of cytosolic Ca2+ levels (Kuo et al., 2015).

The MARTXVv toxin functions along with the VvhA cytolytic/haemolysin pore-forming toxin in the gut to promote early in vivo growth and dissemination of V. vulnificus from the small intestine to other organs. After cytoskeletal rearrangements and cytotoxicity caused by RtxA1, VvhA, is produced leading to accelerated cell death (Kim et al., 2008, Jeong & Satchell, 2012, Gavin & Satchell, 2015). VvhA is a 51-kDa extracellular cytolysin in V. vulnificus homologous to the V. cholerae El Tor haemolysin. A study using a mutant that no longer produced cytolysin showed no affect in virulence in the mouse model without iron loading (Yamamoto et al., 1990, Wright & Morris, 1991).

Serum from infected mice and from patients recovering from acute infections contain specific antibodies against the cytolysin (Gray & Kreger, 1986) but its role in V. vulnificus infections was not clear since there was no difference in the virulence of wild-type and isogenic strains unable to express the cytolysin (Wright & Morris, 1991). However, in recent studies it has been demonstrated that VvhA significantly upregulates autophagy flux and induces necrotic cell death and apoptosis in human intestinal epithelial cells (Lee et al., 2015, Song et al., 2016).

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The QS regulator, LuxO, controls the cytotoxicity of V. vulnificus. SmcR has been shown to repress the expression of both rtxA1 and vvhA. It seems that SmcR mediates cytotoxicity by repressing hlyU, an activator of rtxA1 and vvhA (Shao et al., 2011). HlyU is a master regulator of in vivo virulence expression (Kim et al., 2003b) and in V. vulnificus up-regulates the expression of rtxA1 with two other genes, a peptide chain release factor 1 and a haemolysin acyltransferase (Liu et al., 2007).

In addition to regulation by SmcR (Shao et al., 2011), the haemolysin is also regulated by the ToxR master regulator, as a mutation in toxR resulted in decreased haemolysin production (Lee et al., 2000). In addition, Fur repressed the transcription of the vvhHA operon but the haemolysin content and haemolytic activity were lowered in cell-free supernatants of the fur mutant as result of the exoproteolytic activity of the elastase, VvpE, and a metalloprotease, VvpM, which were also regulated by Fur (Lee et al., 2013b).

V. vulnificus produces a 45 kDa thermolysin-like zinc metalloprotease (Vvp) as an important virulence factor that increases vascular permeability and causes serious haemorrhagic damage (Miyoshi et al., 1993, Miyoshi & Shinoda, 2000). The metalloprotease digests the vascular basement membrane, and in particular the type IV collagen forming the framework of the membrane (Miyoshi et al., 2001). It also causes oedema through induction of exocytotic histamine release by the mast cells (Miyoshi et al., 2003) and activation of the factor XII-plasma kallikrein-kinin cascade which results in enhancing the vascular permeability (Miyoshi et al., 2004).

Both V. vulnificus biotypes 1 and 2 have the vvp gene but only the strains belonging to biotype 1 can produce the metalloprotease and grow in human serum (Watanabe et al., 2004). A V. vulnificus vvp mutant was as virulent as the parental strain in mice infected intraperitoneally and was more virulent in mice infected via the oral route. However, the cytolysin activity in the culture supernatant of the mutant was found to be higher than the wild type (Shao & Hor, 2000).

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1.2.5 Vibrio cholerae

V. cholerae is a natural inhabitant of brackish and saltwater systems (Colwell et al., 1977). It can enter the human body through consumption of contaminated water or food and cause cholera, a type of severe rice water diarrhoea. Infection can lead to death due to dehydration, electrolyte imbalance and shock (Carpenter, 1971). Cholera is still a global concern, but mostly occurs in impoverished communities or when other calamities such war or natural disasters destroy critical infrastructure, limiting access to clean water and hygienic facilities.

Cholera-like diseases were described in ancient times by Hippocrates and Buddha but cholera outbreaks have only been documented since 1817 (Barua, 1992). In 1849 Sir John Snow, one of the founding fathers of modern epidemiology, correlated cholera outbreaks with contaminated water (Snow, 1855). However, it was not until 1854 that Filippo Pacini, a famous anatomist, first described V. cholerae (Pacini, 1854). In 1883, Robert Koch, a pioneering microbiologist, rediscovered the same organism in a separate study on an expedition to Egypt, and presumably unaware of Pacini’s work, named it Kommabazillen, overshadowing Pacini’s discovery for several decades (Koch, 1884, Lippi & Gotuzzo, 2014).

Seven distinct cholera pandemics have been recorded since 1817, from which, the sixth and most probably fifth pandemics were caused by V. cholerae O1 of the Classical biotype. The seventh pandemic caused by the El Tor biotype extends to the present day (Kaper et al., 1995), and includes the recent outbreak of cholera after the earthquake in Haiti. The disease had been absent for over a century in Haiti, but the current outbreak resulted in more than 531,000 cases (5 percent of the population) and more than 7,050 deaths as of March 2012 (Chin et al., 2011, Sontag, 2012).

1.2.6 Serogroups and biotypes

V. cholerae has more than 200 serogroups identified to date, from which only serogroups, O1 and O139 are shown to be responsible for pandemic cholera (Chatterjee & Chaudhuri, 2003). The O1 serogroup can be divided into Classical and El Tor biotypes as well as

21 | P a g e two major serotypes; Ogawa and Inaba, and a third rarely isolated serotype Hikojima (Feeley, 1965). The O139 serogroup has recently emerged and has alterations in both phenotypic and genetic characteristics when compared to V. cholerae O1 serogroup that was responsible for previous epidemics (Swerdlow, 1993). The O1 El Tor biotype acquired the 0139 O antigen by horizontal gene transfer, becoming the current O139 strain (Waldor et al., 1994, Elisabeth M.Bik et al., 1995). The O1 and O139 strains possess either the O1 or O139 antigens and produce cholera toxin (CT), an enterotoxin responsible for rapid fluid loss from the intestinal epithelium (Kaper et al., 1995). However, some V. cholerae O1 strains have been found to be CT negative (Kaper et al., 1981, Kaper et al., 1995). Interestingly, it has been shown that phage transduction with CT-encoding phage CTXφ can convert the non-toxigenic environmental strains to CT positive strains in the gastrointestinal environment (Waldor & Mekalanos, 1996).

The genes encoding TCP are clustered on the V. cholerae pathogenicity island (VPI) which is present in O1 (El Tor and Classical) and O139 V. cholerae strains, but lacking in the majority of environmental isolates (Karaolis et al., 1998). In addition to VPI (Faruque & Mekalanos, 2003), toxigenic O1 and O139 serogroups possess Vibrio pathogenicity island-2 (VPI-2) which encodes a neuraminidase (nanH), part of a mucinase complex that degrades the mucin layer of the gastrointestinal tract (Crennell et al., 1994), and genes required for the utilisation of amino sugars (Jermyn & Boyd, 2005). El Tor and related O139 strains also possess Vibrio seventh pandemic islands, VSP-1 and VSP-2, encoding mostly hypothetical open reading frames that are presumed to be involved in the fitness and epidemic spread of the seventh pandemic strains (O'Shea et al., 2004).

1.2.7 Distribution of V. cholerae

Both toxigenic and non-toxigenic strains of V. cholerae, like many other members of the genus Vibrio, are autochthonous inhabitants of brackish and estuarine water systems from the tropics to temperate waters world-wide (Lutz et al., 2013). The strains isolated from the environment outside epidemic regions are usually non-toxigenic (Colwell & Huq,

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1994), although non-toxigenic V. cholerae non-O1/O139 strains have been isolated from patients with gastritis (Hasan et al., 2015).

V. cholerae can survive for at least 70 days at 25°C at salinities between 1.5 and 3%. However, at 4°C and an optimal salinity of 2.0%, the cells lose viability after 45 days, regardless of nutrient availability. In contrast, V. cholerae survives for a long period of time in nutrient-deplete warm water at salinities between 0.25 and 3% at a pH of 8.0 (Miller et al., 1984). V. cholerae has also been shown to endure low nutrient and low temperature conditions (i.e. 10°C) for long periods if supplemented with sodium (Singleton et al., 1982).

V. cholerae, like V. vulnificus, has the ability to enter the VBNC state under stressful conditions (i.e. nutrition deprivation, high salinity and low temperature). When entering the VBNC state, the cells exhibit a reduction in size and cannot be grown on normal culture media (Colwell et al., 1985). V. cholerae biofilms have been shown to enter a VBNC state and to resuscitate to a cultarable state after being passaged in animals (Alam et al., 2007). It has been shown by Huq et al. (1990) in a rabbit ileal loop infection model as well as in human volunteers that VBNC cells are capable of resuscitation, resulting in cholera. Thus, VBNC cells of V. cholerae are a potential health threat (Huq et al., 1990, Colwell & Huq, 1994).

V. cholerae has the ability to attach to a variety of animal and plant surfaces, including but not limited to chitinous exoskeletons of zooplankton and phytoplankton (Vezzulli et al., 2010). It colonises the surface of dead plankton more readily than live plankton and attached cells may act as natural reservoirs for V. cholerae in the aquatic environment (Mueller et al., 2007). V. cholerae shows higher survival rates when attached to crustaceans such as copepods, cyanobacteria and chironomid egg masses in comparison to free-living planktonic cells (Huq et al., 1983, Halpern et al., 2004, Islam et al., 2004). In places where access to clean water is limited, it has been proposed that filtration of contaminated water through used sari cloth removes 99% of the plankton and consequently attached V. cholerae cells, resulting in a reduction in the incidence of cholera (Huq et al., 1996).

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1.2.8 Pathogenicity factors

V. cholerae genomes exhibit plasticity and possess a variety of virulence factors that differ between biotypes. From more than 200 recognised serogroups, only serogroup O1 and O139 have been associated with severe disease and cholera pandemics. Although non-O1/O139 V. cholerae strains still carry certain virulence genes and cause sporadic disease, the severity of disease is less than infections caused by O1/ O139 serogroups (Reidl & Klose, 2002).

1.2.8.1 Toxins

Like other pathogenic bacteria, V. cholerae possesses several toxins that play a role in pathogenesis. CT is the most important factor causing the signature symptoms of cholera; secretion of water, electrolytes and mucin into the bowel resulting in profuse diarrhoea known as rice water stools (Forstner et al., 1981). The gene encoding the A subunit of CT, ctxA, is responsible for the diarrhoea but does not have a role in colonisation of the intestinal epithelium. The ctxA gene is positively regulated by the response regulator, ToxR. The tcpA gene encodes the major subunit of TCP, and is also regulated by ToxR. TCP is involved in the colonisation of epithelial cells (Herrington et al., 1988). Pathogenic strains acquire CT by acquisition of a filamentous bacteriophage, CTXφ, which uses the TCP as a receptor. The ctxAB genes encoding CT are arranged in a cluster of genes that were first described as the CTX genetic element (Pearson et al., 1993) until this region was shown to be the genome of the CTX phage (Waldor & Mekalanos, 1996).

CT is a member of the AB toxin family. The A-subunit is a catalytically active heterodimeric linked to a homopentameric B-subunit. CT binds to the cell membrane receptor, GM01, and moves from the plasma membrane through the trans-Golgi and endoplasmic reticulum (ER) to the cytosol. Here CT targets the basolaterally located adenylate cyclase which becomes constitutively activated after toxin-induced mono- ADP-ribosylation of the regulating GS-protein. Elevated intracellular cAMP levels lead to severe loss of water and electrolytes (Lencer & Tsai, 2003, Vanden Broeck et al., 2007). The expression of the TCP and CT, are regulated by the LysR master regulator,

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AphA/B which activates the expression of the membrane-bound transcription factors TcpP and TcpH (Skorupski & Taylor, 1999, Kovacikova & Skorupski, 2001).

Many V. cholerae El Tor O1 and non-O1 strains express haemolysin (HlyA), a water- soluble cytolytic protein (Yamamoto et al., 1984, Kaper et al., 1995) that permeabilises a wide spectrum of eukaryotic cells, including human and rabbit erythrocytes (Yamamoto et al., 1984), by forming transmembrane pentameric pores (Zitzer et al., 1999). The purified toxin causes secretion of fluid in a rabbit ligated ileal loop, suggesting its involvement in the pathogenesis of cholera (Ichinose et al., 1987).

At least four haemagglutinins (HAs) have been described in V. cholerae to have a role in intestinal colonisation (Hanne & Finkelstein, 1982). An extracellular Zn-dependent metalloprotease, haemagglutinin protease (HAP, vibriolysin), is potentially involved in a broad range of pathogenic activities. In tissue culture cells and animal infection models, it has been shown to cause the covalent modification of other toxins, degradation of the protective mucus barrier and disruption of intestinal tight junctions (Benitez & Silva, 2016). The major extracellular protease HAP, encoded by the hapA gene, functions as a detachase that activates the release of attached cells into the intestinal lumen where they are ultimately secreted. HAP is regulated by QS where at high cell density, HapR activates the expression of hapA (Finkelstein et al., 1992).

V. cholerae also carries the RtxA toxin that is activated by RtxC and an associated ABC transporter system, RtxB and RtxD. In V. cholerae strains of the Classical biotype, a deletion within the gene cluster results in loss of rtxC, eliminating cytotoxic activity. In contrast, the El Tor O1 and O139 strains contain a functional gene cluster and display cytotoxic activity (Lin et al., 1999). In V. cholerae, RtxA causes the depolymerisation of actin stress fibres resulting in the rapid rounding of cells in culture (Fullner & Mekalanos, 2000).

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1.2.8.2 Colonisation factors

Expression of multiple factors is a necessary requirement for colonisation of the small intestine (Kaper et al., 1995). TCP is critical for colonisation of epithelial cells (Attridge et al., 1993, Kovacikova & Skorupski, 2002), as a V. cholerae strain lacking TcpA, a subunit of TCP, is incapable of causing cholera in humans volunteers (Herrington et al., 1988). TCP also acts as the receptor for the CTX phage (Waldor & Mekalanos, 1996) and in V. cholerae O1 and O139, has a role in the formation of three dimensional biofilms (Yildiz & Visick, 2009) in addition to its role in colonisation of the human intestine (Herrington et al., 1988). TCP promotes microcolony formation by mediating pilus-pilus interactions between bacteria (Taylor et al., 1987, Kirn et al., 2000).

There are several other factors that, in addition to TCP, are important for the colonisation of human epithelial cells (Herrington et al., 1988). One of the factors involved in intestinal colonisation is an accessory colonisation factor encoded by acf located immediately adjacent to the tcp and toxT loci (Peterson & Mekalanos, 1988). In addition, V. cholerae requires a cell-associated mannose and fucose resistant haemagglutinin (MFRHA) to persist in the intestines (Franzon et al., 1993). V. cholerae also uses several pili for colonisation, including a MSHA type IV pilus (Jonson et al., 1991) and a core- encoded pilus (Pearson et al., 1993). Several outer membrane proteins (Webb et al., 2003) like the iron-regulated virulence determinant (IrgA) (Goldberg et al., 1990), OmpU (Sperandio et al., 1995) and LPS (Chitnis et al., 1982), are also involved in this complex process.

Pili are used for adhesion of bacterial cells to surfaces as well as to other bacteria and target cells in response to various environmental cues. The attachment of the cell by the pili represses the rotation of flagella (Yildiz & Visick, 2009). In V. cholerae O1 El Tor strains the MSHA type IV pilus accelerates the formation of early stage biofilms and plays an important role in attachment to nutritive substrates (Watnick et al., 2001). A V. cholerae O139 mshA mutant that cannot produce pili was still able to form biofilms and aggregate in the planktonic phase, indicating that the MSHA pilus is not necessary for early biofilm formation in this strain (Moorthy & Watnick, 2004). Another factor affecting biofilm formation is the chitin-regulated pilus (ChiRP), formerly known as PilA,

26 | P a g e which has been found to be involved in biofilm formation on chitin surfaces (Meibom et al., 2004).

1.2.8.3 Biofilm formation

Biofilms are communities of microorganisms associated with biotic or abiotic surfaces and their protective nature allows V. cholerae to survive and thrive in aquatic and intestinal environments (Teschler et al., 2015). V. cholerae is able to form biofilms both in its environmental niche as well as in the human host as evidenced by the aggregates observed in stool samples from cholera patients (Yildiz & Schoolnik, 1999, Faruque et al., 2006). The biofilm lifestyle protects V. cholerae against stresses such as nutrient deprivation, changes in salinity, pH and temperature, exposure to UV, antibiotics, chlorine and predators such as protozoa (Hall-Stoodley & Stoodley, 2005). Biofilm formation can be a successful anti-predation mechanism. For example physically large cell clumps are beyond the size smaller predators can ingest and biofilms have been shown to have coordinated release of antiprotozoal factors (Hahn & Höfle, 2001, Jürgens & Matz, 2002). Antiprotozoal mechanisms will be discussed in detail in the next section.

The flagellum is important in initial stages of biofilm formation by V. cholerae. The loss of flagellar genes generally leads to decreases in attachment and hence biofilm formation, although this varies between strains (Yildiz & Visick, 2009). For example, a mutant in flaA aggregated in liquid media due to excess production Vibrio polysaccharide (VPS). This strain took much longer to form a biofilm than the wild type strain as the reversible attachment stage was prevented (Tolker-Nielsen et al., 2000). The motB and motY mutants of V. cholerae that possess a complete but paralysed flagellum did not form a biofilm in an 18 hour assay, clearly exhibiting a defect in attachment to surfaces (Watnick et al., 2001).

VPS is the exopolysaccharide produced by V. cholerae and it contributes to the extracellular matrix which also includes extracellular DNA (eDNA), polysaccharides, proteins, amyloid fibres and bacteriophage (Flemming et al., 2007, McDougald et al., 2012). When the cells become completely sessile and permanently attached to the

27 | P a g e substratum, the extracellular material adheres the bacterial cells to each other as well as to the surface (Yildiz & Visick, 2009).

Like V. vulnificus, V. cholerae has two morphotypes that are easily distinguished as wrinkled (rugose) and smooth colonies. Rugose cells express higher amounts of VPS and typically form thicker biofilms in comparison to smooth cells. There are two mostly conserved loci amongst Vibrios that are involved in the production of VPS and therefore, biofilm formation, the vps-I and vps-II operons (Yildiz & Visick, 2009). VpsR and VpsT are positive master regulators of VPS biosynthesis and biofilm formation in V. cholerae. Mutants in vpsR revert irreversibly to a smooth morphotypes and vpsR mutants and vpsR vpsT double mutants are unable to form microcolonies (Yildiz et al., 2001, Casper- Lindley & Yildiz, 2004, Beyhan et al., 2007). The master virulence regulator AphA also binds to the vpsT promoter and positively regulates its expression (Yang et al., 2010).

The intracellular secondary messenger molecule c-di-GMP (Jenal & Malone, 2006) is synthesised by diguanylate cyclases, a class of enzymes containing GGDEF domains, while hydrolysis is due to phosphodiesterases containing EAL domains (Jenal & Malone, 2006). The over expression of proteins with GGDEF domains results in upregulation of vpsR and vpsT (Beyhan et al., 2006) which leads to increased rugose colony formation (Lim et al., 2006) and a decrease in flagellar motility (Römling & Amikam, 2006). Some of the genes encoding GGDEF and EAL domain proteins are regulated by QS. A reduction in c-di-GMP levels at high cell density results in decreased biofilm formation. In addition, the QS response regulator, HapR, can directly bind to vpsT and repress its expression (Waters et al., 2008). This negatively regulates biofilm formation by reducing c-di-GMP levels when the cell density is high, repressing the vpsA and vpsL operons further reducing biofilm formation (Hammer & Bassler, 2003).

1.2.8.4 Quorum sensing

V. cholerae possesses multiple QS pathways to control virulence and biofilm formation. There are at least three parallel QS circuits that transmit information through four redundant regulatory small RNAs (sRNAs) called quorum regulatory RNAs (Qrrs)

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(Figure 1-3) (Lenz et al., 2005). System 1 has a two-component system composed of the synthase, CqsA, that produces the CAI-1 autoinducer, (S)-3-hydroxytridecan-4-one. CAI-1, is recognised by the cognate receptor, CqsS, a transmembrane histidine kinase. System 2 uses the LuxS enzyme that synthesises the furanosyl borate diester, (2S,4S)-2- methyl-2,3,3,4-tetrahydroxytetrahydrofuran borate autoinducer (AI-2), a periplasmic binding protein, LuxP, and a two component sensor, LuxQ (Bassler et al., 1994, Chen et al., 2002, Miller et al., 2002). System 3 is the VarS/VarA two-component sensory system, which controls transcription of three sRNAs, CsrB, CsrC and CsrD. These three sRNAs control the activity of the global regulatory protein, CsrA. Regulation of the expression of the Qrr sRNAs by VarS/VarA-CsrA/BCD system leads to regulation of the entire QS regulon (Lenz et al., 2005). Both systems 1 and 2 transduce signals through the LuxU phosphotransfer protein to LuxO, while system 3 feeds directly to LuxO. Ultimately, all the information from the three pathways is transduced through LuxO to the response regulator, HapR (Miller et al., 2002, Lenz et al., 2005).

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Figure 1-3 V. cholerae quorum sensing systems, comprised of three parallel circuits (Ng & Bassler, 2009). See text for explanation.

LuxO~P activates expression of genes encoding Qrr1-4 (Lenz et al., 2004). Qrr sRNAs negatively regulate the mRNA of luxO, establishing a feedback loop to control Qrr production (Svenningsen et al., 2009), and to activate production of the master transcription regulator, AphA (Rutherford et al., 2011). In response to environmental cues, QS regulates the ToxR, TcpP, and ToxT signal transduction cascade, which in turn controls the virulence regulon, including CT and TCP. At low cell densities, LuxO

30 | P a g e upregulates the expression of virulence factors and mutation of luxO represses the ToxR regulon. However, LuxO represses hapR expression in early log-phase growth as HapR represses expression of genes required for virulence factor production and biofilm formation. At high cell densities expression of hapR blocks ToxR-regulon expression, and in turn represses the expression of the essential virulence regulator, TcpP. In addition, LuxO and HapR are responsible for regulation of a variety of other cellular functions such as motility, protease production, and biofilm formation (Kovacikova & Skorupski, 2002, Zhu et al., 2002).

Figure 1-4 Diagram depicting the regulatory pathways involved in virulence of Vibrio spp.. The QS pathway regulates the expression of VpsR, VpsT, AphA and HapR that leads to induction of virulence factors such as TCP and CTX. The second messenger cAMP is generated by adenylate cyclase CyaA, and in complex with cAMP receptor protein (CRP), upregulate HapR production, which regulates transcription of genes involved in biofilm formation, possibly by promoting VpsT-mediated transcriptional activation of vps genes.

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Table 1-1 A summary of the virulence factors reviewed in this chapter.

Virulence factor V. cholerae V. vulnificus Aggregates observed in Adherence to eel , protects Biofilm formation human stool from stress Capsular polysaccharide Observed in non-O1 strain, Antiphagocytic factor (CPS) poorly understood essential in pathogenicity Cell-associated mannose and Helps to persist in the fucose resistant intestines haemagglutinin (MFRHA) Secretion of water, Cholera toxin (CT) electrolytes and mucin into the bowel Upregulates autophagy flux Cytolytic/haemolysin pore- and induces necrotic cell forming toxin (VvhA) death and apoptosis in human intestinal epithelial cells Adherence to mammalian Attachment and biofilm Flagellin cells and abiotic surfaces, formation cytotoxin delivery Haem uptake receptor Haem uptake (HupA) Degradation of the protective Haemagglutinin protease mucus barrier and disruption (HAP, vibriolysin) of intestinal tight junction Haemagglutinins (HAs) Intestinal colonisation permeabilises eukaryotic Haemolysin (HlyA) cells Iron-regulated virulence Intestinal colonisation determinant (IrgA) Pyrogenic factor associated Lipopolysaccharides (LPS) with endotoxic shock Mannose-sensitive Intestinal colonisation, haemagglutinin (MSHA)type formation of early stage Chitin and oyster adherence IV pilus biofilms Damage cells by cytoskeleton rearrangement, apoptotic cell death, Multifunctional Cytotoxic activity, generating of reactive autoprocessing repeats-in- depolymerisation of actin oxygen species (ROS), toxin (MARTX) stress fibres protecting against phagocytosis and modification of cytosolic Ca2+ levels. N-acetylglucosamine binding Chitin adherence Chitin and oyster adherence protein A (gbpA) Outer membrane protein, Intestinal colonisation OmpU Formation of biofilms, biofilm formation on chitin PilA, type IV pilin adherence to human cells and surfaces colonisation of oysters

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PilD, type IV prepilin Adherence to human peptidase/N- epithelial cells and secretion methyltransferase of several exoenzymes Utilisation of Fe2+ bound to Siderophore, vulnibactin transferrin and lactoferrin Biofilm formation, auto- Tad, type IV pilin aggregation, and oyster colonization Increases vascular Thermolysin-like zinc permeability and causes metalloprotease (Vvp) serious haemorrhagic damage colonisation of epithelial Toxin-coregulated pili (TCP) cells Vibrio exopolysaccharide Biofilm Formation Biofilm formation (VPS or EPS)

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1.3 Predation by protozoa

Predator and prey responses of animals have been extensively studied, but predation in the microscopic scale is less well studied as observing these interactions in the environment is difficult. The natural predators of bacteria in the environment are viruses, protists and other bacteria. Lytic phage can effectively control bacterial populations. For example, in a three year study of cholera in interepidemic periods in Bangladesh, the water samples were shown to contain cholera phage and no viable bacteria were found (Faruque et al., 2005). Another known group of predators that can effectively feed on bacteria are protozoa which is the focus of this study. It is known that predation by protozoa is also a force controlling bacterial populations, genotypes and phenotypes (Jürgens & Matz, 2002).

Protists are a highly diverse group of unicellular eukaryotic microorganisms that were first discovered by Van Leeuwenhoek in 1674 as “animalcules”. The term protozoa was first introduced in 1818 and was derived from the Greek words “protohi” and “zoa” meaning “first animal” (Goldfuß, 1820). For simplicity, protozoa are grouped into flagellates, ciliates and amoeba based on their morphology, method of feeding and locomotion. However, this classification is not reflective of phylogenetic or taxonomical relationships (Levine et al., 1980, Parry, 2004).

Protozoa vary in size from 2 to 200 µm but can reach up to 2 mm. They are abundant in aqueous as well as soil environments and can inhabit a wide spectrum of trophic levels, however, a majority of protozoa are heterotrophic (Porter et al., 1985, Sherr & Sherr, 2007). Some protozoan species are autotrophic and there are species that have the ability to switch between autotrophic and heterotrophic feeding based on nutritional availability in the environment. There are reports of ciliate species taking up chloroplasts or phototrophic algae to form a symbiotic relationship (Gustafson et al., 2000). Most heterotrophic protozoa are either active raptorial predators or filter feeders that consume detritus or microorganisms via phagocytosis or by absorption of soluble dissolved organic matter via pinocytosis (Sherr et al., 1982, Porter et al., 1985, Sigee, 2005). Different environmental niches accommodate different prey. For example, protozoa that are mainly sessile can feed on attached bacteria while motile protozoa may be able to feed on

34 | P a g e planktonic bacteria or biofilms (Lynn, 2001, Parry, 2004, Sigee, 2005). These heterotrophic predators are the focus of this study.

Phagotrophic protozoa are competent grazers, consuming large numbers of prey sometimes several times their own body weight (Vickerman, 1992). Amoebae are surface attached organisms that mainly feed on sessile bacteria at rates of between 0.2 and 1465 bacteria amoeba-1 hour-1 but they cannot feed efficiently on planktonic cells (Parry, 2004, Huws et al., 2005). Flagellates are small predators with ingestion rates of 2 – 300 bacteria flagellate-1 hour-1. Ciliates can feed on both planktonic and attached microcolonies and as the major predators of bacteria in aquatic environments, can reach values of 1254 bacteria ciliate-1 hour-1 (Parry, 2004).

Flagellates are usually smaller than other protozoa (2 to 20 µm) and possess one or more flagella used for swimming and feeding. Many heterotrophic nanoflagellates (HNF) are bacterivores and ingest smaller sized prey, generally under 20 µm (Gonzalez et al., 1990, Parry, 2004). Flagellates have diverse feeding preferences, some can feed on both planktonic and sessile bacterial cells but they can’t consume bigger clumps of cells. Flagellates have been shown to select their prey based on size (Chrzanowski & Simek, 1990). They draw their prey towards the base of the flagellum and into an oral groove by creating a strong current using their flagella. Some filter-feeding flagellates use a collar of tentacles located at the base of the flagellum that allows the smallest prey particles to pass through (Chrzanowski & Simek, 1990, Matz et al., 2002, Parry, 2004). Some flagellates attach to surfaces and thereby create larger feeding currents in order to increases prey ingestion rates (Fenchel, 1982). The common suspension feeder, Cafeteria roenbergensis, is a motile marine bacterivorous flagellate that creates circular water currents to collect bacterioplankton (O'Kelly & Patterson, 1996, Boenigk & Arndt, 2000). The cosmopolitan, benthic flagellate Rhynchomonas nasuta, attaches to a surface by a posterior flagellum and feeds on surface-attached bacteria by raptorial feeding (Swale, 1973, Matz et al., 2002).

Ciliates are generally larger than flagellates (10 to 1000 μm) and can feed on larger prey including bacteria, algae, flagellates and even other ciliates (Epstein et al., 1992). The main mode of motility is by the beating of numerous cilia found on the cell surface. In

35 | P a g e some species of ciliates, these cilia cover the entire cell while in others, the cilia form cirri, which are compound ciliary organelles. Ciliates can be raptorial or filter feeders (Epstein et al., 1992, Finlay & Esteban, 1998). Filter-feeding is accomplished by the use of a dense row of membranelles resembling short hairs that generate a water current bringing prey into a cytostome that acts as the mouth. Ciliates can direct several larger prey cells into a single food vacuole (Parry, 2004). Prey-filled food vacuoles are trafficked through the cytosome toward the posterior end of the cell by attached actin filaments (Hosein et al., 2005). Ciliated protozoa are suitable for a variety of tests for environmental risk assessment since they occupy a wide range of trophic levels and play a key role in the microbial loop. For example, many studies use Tetrahymena spp., and in particular Tetrahymena pyriformis (Yoshioka et al., 1985, Sauvant et al., 1999) to assess toxicity of chemicals (Carter & Cameron, 1973, Yoshioka et al., 1985, Sauvant et al., 1999, Bogaerts et al., 2001).

Protozoan use chemical cues for finding prey. Experiments using the free-swimming filter feeder Tetrahymena sp. and the surface-associated predator Chilodonella sp., showed both a had preference for Pseudomonas costantinii biofilms when offered different food choices even though they could feed readily on both P. costantinii and Serratia plymuthica. Furthermore, using bacterial cell-free extracts in microcosm experiments, both ciliates were shown to use chemotaxis to find prey (Dopheide et al., 2011).

The term "amoebae" is used for a wide range of protists ranging in size from 20 µm to 2 mm that move by crawling on surfaces. Naked amoebae are raptorial feeders that use cytoplasmic extensions from the cell surface called pseudopodia to move. The pseudopodia also envelop prey, trapping them in a food vacuoles (Parry, 2004). Some groups of amoebae actively hunt for prey (Sigee, 2005), mostly bacteria or algae, but non- living organic particles are also often ingested (Finlay & Esteban, 1998). Amoebae can be found in a variety of environments, from the Antarctica where temperatures are extremely low to arid deserts with temperatures above 45°C. They are found in fresh and marine waters and soil making them one of the most successful thermotolerant protists (Bradley & Marciano-Cabral, 1996). Amoebae typically have different life stages; a feeding vegetative trophozoite stage and a highly resistant cyst stage whereby a thick wall

36 | P a g e enables the organism to survive during stresses such as starvation and dehydration (Bradley & Marciano-Cabral, 1996).

Acanthamoeba castellanii is a free-living amoeba that is also an opportunistic pathogen of humans (Rodríguez-Zaragoza, 1994). In aqueous environments, A. castellanii attaches to surfaces and feeds on biofilm communities (Huws et al., 2005). A. castellanii is an important model protozoan as it interacts with a wide variety of microorganisms such as fungi, bacteria and viruses. In addition, A. castellanii is physiologically similar to mammalian immune cells such as macrophages (Guimaraes et al., 2016).

1.4 Bacterial adaptations against predation by protozoa

The major constraint on bacterial biomass in the environment is grazing by bacterivorous protozoa (Sherr & Sherr, 1994). Predation by protists can lead to blooms or loss of certain bacterial species in particular niches, as different protozoa feed on different prey. Predation by bacterivorous protists in aquatic habitats can shape the taxonomic composition and physiological status of bacterial communities (Hahn & Höfle, 2001). Heavy grazing mortality has been shown to play a significant role in regulating numbers of V. cholerae in coastal marine waters (Worden et al., 2006).

Animals evolved in a world that has been inhabited by billions of bacteria, archaea, and protozoa (McFall-Ngai et al., 2013), and thus it is speculated that the rise of human pathogens is the result of the evolution of ancestors that were defended from protozoan grazing or in some cases were even able to parasitise protists (Strassmann & Shu, 2017). This is likely to be the case as there are several similarities between mammalian immune and protozoan cells such as prey recognition by cell surface receptors (Brown et al., 1975), prey killing by oxygen radicals (Davies et al., 1991) and similar digestive enzymes (Harb et al., 2000). Thus, mechanisms of bacterial survival and defence that work against protozoa, are likely to work against mammalian cells due to this conservation of targets. Protozoa can be considered to be a ‘biological gym’, where the intracellular bacterial pathogens become more adapted to mammalian cells as a consequence of interactions with protists (Harb et al., 2000).

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Bacteria use a number of defence mechanisms against predation, including pre- ingestional and post-ingestion adaptations (Matz & Kjelleberg, 2005, Pernthaler, 2005) that are employed during different stages of protozoan grazing, namely encounter, capture, ingestion and digestion stages (Figure 1-5).

Figure 1-5 Potential bacterial adaptations against predation by protozoa are depicted in two divergent groups of pre-ingestional (a-e) and post-ingestional (f-h) strategies (Matz & Kjelleberg, 2005).

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Some species of bacteria have been shown to change cell size or shape, and as mentioned earlier, this is effective since some protozoa feed size-selectively. Furthermore, in instances where protozoan predation is the major selective pressure, different groups of grazing-resistant filamentous bacteria can be established and become dominant in that niche (Jürgens et al., 1999). For example, under grazing pressure small cocci and rod shaped bacteria were shown to form star-like prosthecate bacteria and filaments that were resistant to grazing (Bianchi, 1989). In another example Comamonas acidovorans formed filamentous cells in response to exposure to grazing by the bacterivorous flagellate, Ochromonas sp. (Hahn & Höfle, 1998). Flectobacillus sp. is another genus demonstrated to form filamentous cells under grazing pressure (Hahn et al., 1999), increasing from an average size of 2 to 7 µm up to 40 µm under grazing pressure (Corno & Jürgens, 2006). Lower predation rates on filamentous cells is due to lower ingestion efficiency rather than being solely dependent on size selection (Wu et al., 2004).

Some bacteria have the ability to increase swimming speed to avoid being captured by protozoa (Matz & Jurgens, 2005). A grazing experiment using the nanoflagellate Spumella sp. as the predator showed that increases in bacterial swimming speed increased contact rates but since the capture efficiency was decreased, the overall ingestion of the prey was decreased (Matz et al., 2002). Another grazing experiment with three nanoflagellates and several bacterial strains with the same cell size but different motility rates, showed increases in swimming speed and hence lower ingestion rates (Matz & Jurgens, 2005).

Cell surface properties have also been shown to affect grazing resistance. For example, cell surface hydrophobicity has been shown to affect grazing of picoplankton cells by nanoflagellates (Monger et al., 1999). Moreover, Wildschutte et al. (2004) showed that differences in O-antigen are sufficient to allow for prey discrimination by protozoa grazing on different serotypes of Salmonella spp. Protozoa, especially amoeba, may be able to identify antigens and receptors on prey cell surfaces, thus allowing for discrimination of prey. A. castellanii possesses a mannose sensitive carbohydrate binding protein that is used for selecting prey based on cell contact (Brown et al., 1975). A Ca2+- dependent, mannose-binding lectin on the marine dinoflagellate Oxyrrhis marina, was shown to act as the feeding receptor for recognising prey and a strain lacking this lectin

39 | P a g e could not distinguish between different sugar-coated beads (Wootton et al., 2007). Production of CPS, LPS and outer membrane proteins protects Klebsiella pneumoniae from predation by Dictyostelium discoideum and macrophage, where K. pneumoniae mutants defective in the LPS core, lipid A palmitoylation, OmpA and OmpK36 are susceptible to phagocytosis (March et al., 2013).

Some bacteria use post-ingestional defence mechanisms to avoid being digested. For example, the cyanobacteria Synechococcus sp., were excreted from nanoflagellates a few minutes after being ingested, while heterotrophic bacteria in the same experiment were digested (Boenigk et al., 2001). Many bacteria are known to survive inside various strains of amoebae such as Acanthamoeba sp., Hartmanella and Naegleria sp. and ciliates such as Tetrahymena sp. (Greub & Raoult, 2004). Bacteria that survive intracellularly, including Legionella pneumophila (Bozue & Johnson, 1996), Campylobacter jejuni (Axelsson-Olsson et al., 2005), Francisella tuleransis (Abd et al., 2003), Escherichia coli K12 (Alsam et al., 2006), Burkholderia cepacia (Landers et al., 2000) and Mycobacterium avium (Steinert et al., 1998) employ the same mechanisms to survive inside amoebae and human macrophages, supporting the idea that development of resistance to protozoa has led to the evolution of pathogenicity in these bacteria (Barker & Brown, 1994).

V. cholerae O1 and O139 strains have been shown to survive inside various amoebae, including A. castellanii, Acanthemoeba polyphaga and Naegleria gruberi (Thom et al., 1992, Abd et al., 2005, Abd et al., 2007). Van der Henst et al. (2016) showed that V. cholerae can resist intracellular killing by A. castellanii where the non-digested bacteria were either released or replicated within a vacuole.

Copper and zinc efflux and iron and manganese uptake affect bacterial survival in protozoan and macrophage phagosomes. Iron, manganese, zinc and copper play important roles in the innate immune system and antimicrobial activity of macrophages in defence against invading microbial pathogens. Successful feeding and growth of protozoa is dependent on bacterial susceptibility to copper (White et al., 2009, Hood & Skaar, 2012, Samanovic et al., 2012). The cue system, responsible for copper efflux is main copper resistance mechanism in E. coli and CueR regulates the expression of two genes, cueO and copA (Grass & Rensing, 2001, Rensing & Grass, 2003). E. coli lacking 40 | P a g e the Cu(I)-translocating P1B-type ATPase (copA), iron uptake transporters (feoAB and entC) and manganese uptake transporter (mntH), and a Pseudomonas aeruginosa mutant lacking Cu(I)-translocating P1B-type ATPase (cueA) showed reduced grazing resistance against D. discoideum (Hao et al., 2016). Genes encoding the copper uptake transporter p80 and a triad of Cu(I)-translocating PIB-type ATPases such as copA, were upregulated after phagocytosis by D. discoideum (Hao et al., 2016).

Another important defence strategy is the production and secretion of toxins that can harm or kill the predator (Matz & Kjelleberg, 2005). Janthinobacterium lividum and Chromobacterium violaceum release violacein, an alkaloid purple pigmented secondary metabolite that causes apoptosis of nanoflagellates. The production of violacein is controlled by AHLs in these organisms, and a mutant in AHL production was sensitive to grazing pressure (Matz et al., 2004). Violacein is also produced by Peudomalteromonas tunicata, predominately within biofilms, and very low concentrations (nanomolar) inhibit protozoan feeding by inducing a conserved eukaryotic cell death program (Matz et al., 2008b).

Pseudomonas fluorescens uses the cyclic lipopeptide surfactants (CLPs), massetolide, viscosin (Mazzola et al., 2009), lipopeptide (LP), putrescine (Song et al., 2015), 2,4- diacetylphloroglucinol (DAPG), pyrrolnitrin , hydrogen cyanide (HCN), and pyoluteorin (Jousset et al., 2010) to protect itself against protozoan predators. These surfactants and toxic factors kill some protozoan predators, e.g. Naegleria americana. For example, CLPs have been shown to disrupt the protist cell membrane (de Bruijn et al., 2007). P. aeruginosa PAO1 also synthesises HCN that paralyses the nematode Caenorhabditis elegans (Gallagher & Manoil, 2001). Pantoea ananatis uses rhlA and rhlB involved in the biosynthesis of a biosurfactant glycolipid to defend against the amoebae D. discoideum. The biosurfactant is capable of compromising cellular integrity leading to cell lysis (Smith et al., 2016).

E. coli O157:H7 produces Shiga toxin which has been shown to kill Tetrahymena thermophila (Lainhart et al., 2009), and a type III secretion system (T3SS) has been shown to be involved in killing of A. castellanii (Matz et al., 2008a). Virulence of enterohemorrhagic E. coli (EHEC) strains depends on induction of Shiga toxin- converting prophages. T. thermophila is killed when co-cultured with bacteria bearing a 41 | P a g e

Shiga toxin prophage. This ciliate is mouthless and takes up Shiga toxin via pinocytosis. Bacteriophage-mediated lysis of Stx-encoding bacteria is necessary for Stx toxicity in protozoa whereas digestion of bacteria by T. thermophila is harmless to the cell. Mutations in the bacterial SOS response gene (recA) and enzymes involved in the breakdown of H2O2 (catalase) decrease grazing resistance against T. thermophila (Lainhart et al., 2009, Stolfa & Koudelka, 2013). Shiga toxin-converting prophages can be induced via treatment with H2O2. Oxidative stress as a consequence of H2O2 excretion by either neutrophils of infected humans or protist predators can potentially induce the production of Shiga toxin-converting prophages (Katarzyna et al., 2016).

1.5 Vibrio spp. use factors with dual roles in providing fitness in the environment and pathogenicity

The coincidental evolution hypothesis suggests that virulence factors evolved for functions other than virulence, such as adaptation to other ecological niches. As noted above, one particular selective pressure on bacteria is predation by the protozoa, thus grazing resistance may be an evolutionary force that drives the evolution of bacterial pathogenicity (Levin, 1996, Adiba et al., 2010, Erken et al., 2013).

There are many examples of virulence factors or their homologues that exist in environmental strains that may play another functional role in the environment, including but not limited to toxT, ctxA, rstR, tcpA, tcpl, acfB, rtxA, rtxC, hlyA, mshA and stn (heat- stable enterotoxin gene) (Mukhopadhyay et al., 2001, Faruque et al., 2003, Faruque et al., 2004). However, the functions of virulence genes or their homologues in the environment have not been extensively explored. This section will focus on factors that are potentially used by Vibrio spp. in a human host as well as in the aquatic environment.

The primary virulence factor of V. cholera, CT is not only responsible for the dehydrating diarrhoea in humans, it is also required for full lethal intoxication of the Drosophila melanogaster fly. In infected flies, exocyst-mediated junctional trafficking in epithelial cells is disrupted but only when co-administered with a pathogenic but not CT positive strain of V. cholerae (Blow et al., 2005, Guichard et al., 2013).

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Pathogens use proteases for a wide range of purposes during the infection process, as well as in the environment where they are important for acquisition of nutrients. The HAP protease destroys host cell receptors that recognise and bind V. cholerae adhesins and is also involved in the degradation of the gelatinous matrix of chironomid egg masses (Halpern et al., 2003). QS-regulated production of HAP is also important in defence against phages at high cell density (Hoque et al., 2016). Another protease, LonA regulates the levels of c-di-GMP and the type VI secretion system (T6SS), and deletion of lonA in V. cholerae leads to hypermotility and defects in both biofilm formation and colonisation of the infant mouse (Rogers et al., 2016). Mucinase, a soluble haemagglutinin protease, is important for the association and long-term survival of V. cholerae with mucilaginous blue-green algae (Anabaena sp). V. cholerae uses mucinase to degrade the mucin as a nutrient source (Islam et al., 2002). A V. cholerae extracellular protease, PrtV, causes instant cytotoxic effects during infection in human intestinal cell lines and degrades the extracellular matrix components fibronectin and fibrinogen in human plasma (Vaitkevicius et al., 2008). PrtV is regulated by QS, can kill predators such as the flagellate C. roenbergensis, the ciliate T. pyriformis and the nematode, C. elegans by colonisation of the worm’s intestine (Vaitkevicius et al., 2006).

Secretion systems delivering toxic factors to the target, not only are an important part of the infection process but also play a significant role in survival in the environment. An important virulence factor, MARTX type III, is involved in the lysis of a wide range of eukaryotic cells, including amoebae, erythrocytes, epithelial cells and phagocytes after bacterium–cell contact by V. vulnificus biotype 2. The amoebae, Neoparamoeba pemaquidensis, isolated and purified from the same niche as V. vulnificus, a turbot (Scophthalmus maximus) gill, grew significantly less in the presence of the wild-type strain compared to the rtxA13 mutant (Lee et al., 2013a).

The bacterial type VI virulence-associated secretion system (VAS) of V. cholerae first identified by Pukatzki et al. (2006) are nanomachines that inject proteins into host cells by the action of a dynamic intracellular tubular structure analogous to a contractile phage tail sheath (Basler et al., 2012, Ho et al., 2014). VAS genes in V. cholerae encode the T6SS where three proteins (VgrG-1, 2, 3) form a trimeric complex that punctures target cell membranes and delivers the effectors (VasX and TseL) into eukaryotic cells (Leiman

43 | P a g e et al., 2009, Dong et al., 2013). In V. cholerae non-O1/non-O139 strains, T6SS has been shown to have toxic effects towards the amoeba, D. discoideum, as well as mammalian macrophages (Pukatzki et al., 2006, Pukatzki et al., 2007, Miyata et al., 2011). In addition to the direct role of T6SS in pathogenesis through the injection of toxins responsible for actin crosslinking in eukaryotic host cells, the T6SS also has a role in competition with neighbouring bacteria (Ma et al., 2009). The T6SS of V. cholerae O1 strain A1552 was demonstrated to be functionally active under high-osmolarity and low temperature conditions, suggesting that the system may be important for the survival of the bacterium in the environment (Ishikawa et al., 2012). The V. cholerae cold shock gene, cspV, controls biofilm formation through modulation of the c-di-GMP and regulates T6SS-mediated interspecies killing in a temperature-dependent manner. A mutation in cspV resulted in significant defects in attachment to the surface of the aquatic crustacean Daphnia magna and T6SS-mediated killing (Townsley et al., 2016). In Vibrio anguillarum, the T6SS may serve as a sensor for an extra-cytoplasmic signal that results in transport of a solute that activates expression of RpoS and VanT, regulators of the general stress response which is important in environmental survival (Weber et al., 2009).

Many factors that have a role in attachment and colonisation of V. cholerae to human epithelial cells during infection also play a role in attachment to surfaces in the aquatic environment and biofilm formation. V. cholerae TCP is required for intestinal colonisation and acquisition of the CTX phage, and has been shown to mediate bacterial interactions required for biofilm differentiation on chitinaceous surfaces (Reguera & Kolter, 2005). GbpA (p53 protein) is involved in attachment to human intestinal cells as well as attachment to chitin particles and chitin-containing plankton organisms (Kirn et al., 2005, Zampini et al., 2005). In fact, V. vulnificus GbpA is also a mucin-binding protein and is essential for pathogenesis in a mouse model (Jang et al., 2016). ChiRP is expressed by chitin-attached V. cholerae and contributes to fitness during the colonisation and adherence to chitin (Meibom et al., 2004). Natural competence of V. cholerae was found to require a type IV pilus assembly complex (Meibom et al., 2005). In V. vulnificus, the type IV pilus structural protein PilA and, to a greater degree, the pre-pilin peptidase PilD, contribute to binding both to abiotic surfaces and to human epithelial cells (Paranjpye et al., 1998, Paranjpye & Strom, 2005). PilA and PilD are also necessary for V. vulnificus and Vibrio parahaemolyticus prolonged attachment to oysters (Paranjpye et

44 | P a g e al., 2007, Aagesen et al., 2013). A ChiRP mutant of V. parahaemolyticus attached to the surface of the coverslip, but did not form aggregates, suggesting that ChiRP plays a role in bacterial agglutination during biofilm formation (Shime-Hattori et al., 2006). V. cholerae primarily use two extracellular chitinases, ChiA1 and ChiA2 (Meibom et al., 2004) and the expression of chiA2 is maximal in the host intestine (Mondal et al., 2014).

The MSHA pilus of V. cholerae is involved in colonisation of zooplankton, chitin beads and the chitinous exoskeleton of the crustacean, Daphnia pulex, biofilm formation on non-nutritive abiotic surfaces, and interaction with bivalve haemolymph (Finn et al., 1987, Chiavelli et al., 2001, Meibom et al., 2004, Pruzzo et al., 2005). V. fischeri colonisation of the squid, Euprymna tasmanica, light organ is directly linked to the expression of mshA (Ariyakumar & Nishiguchi, 2009). However, a direct role in pathogenesis of V. cholerae is debated (Heidelberg et al., 2000). A V. parahaemolyticus MSHA pilin mutant formed aggregates and exhibited a reduction in attachment to abiotic surfaces (Shime-Hattori et al., 2006) and mshA acted as a significant factor in adherence to human intestinal epithelial cells enabling pathogenesis (O’Boyle et al., 2013).

Biofilm formation employed by many bacteria, including V. cholerae, is protective against predation, while planktonic cells are readily consumed (Matz & Kjelleberg, 2005). Predation by protozoa actually promotes biofilm formation by V. cholerae and the rugose morphotype which produces more VPS than the smooth morphotype, is more resistant to grazing (Matz et al., 2005). VPS protects both early and late stage biofilms of V. cholerae from predation by the surface-feeding nanoflagellate, R. nasuta and the amoeba A. castellanii (Sun et al., 2013). The rugosity and biofilm structure modulator A encoded by rbmA, is required for rugose colony formation and biofilm structural integrity in V. cholerae (Fong et al., 2006). A mutant of rbmA exhibited reduced VPS production and biofilm formation, and mutants in the vps gene clusters or rbmA exhibited a defect in intestinal colonisation (Fong et al., 2010).

QS systems in Vibrio spp. regulate a number of virulence factors including CT production, biofilm formation, protease secretion as well as protection from predators (Zhu et al., 2002). The V. cholerae QS mutant hapR exhibited lower survival rates in comparison to the wild type during exposure to protists (Matz et al., 2005). While the V.

45 | P a g e cholerae hapR QS mutant is grazed more than the wild type, it is not completely susceptible to grazing, which suggests that V. cholerae regulates antiprotozoal activities by a combination of QS-dependent and -independent regulation (Erken et al., 2011). It has also been shown that the supernatant from wild-type rugose and smooth V. cholerae A1552 led to decreases in R. nasuta numbers while supernatants from a hapR mutant biofilm supported growth of the flagellate. In a recent study in our lab, mixed biofilms containing V. cholerae rugose wild type and ΔhapR QS mutant strains were exposed to predation by R. nasuta and A. castellanii. The VPS and hapR mutants were grazed more than the wild type and the hapR mutant was the most grazing sensitive, especially in mature biofilms (Sun et al., 2015).

The stress response regulator, RpoS, of V. cholerae is important for environmental survival as it plays a role in tolerance of V. cholerae to high osmolarity, carbon starvation in seawater and most importantly to exposure to hydrogen peroxide (Yildiz & Schoolnik, 1998). RpoS is also involved in survival in the host and dispersal and exit from the intestines, as well as in increasing viability under stressful environmental conditions. HapR upregulates rpoS and positively affects the bacterium's response to oxidative and nutritional stresses (Joelsson et al., 2007). HapR is also required for V. cholerae cells to disperse from the epithelial surface of a rabbit ileal loop and this is positively regulated by RpoS. RpoS along with HapR controls the expression of genes involved in flagellar assembly and chemotaxis, where under high cell density and nutrient limitation leads to the mucosal escape response, which is a key event resulting in the return of the cells back to the aquatic environment (Nielsen et al., 2006).

Overall, the bacterial mechanisms of grazing resistance, environmental survival and virulence were described. Mechanisms such as toxins and biofilm formation are crucial for grazing protection and pathogenesis. This study aims to investigate the presence of antiprotozoal activities in V. cholerae and V. vulnificus biofilm and planktonic populations and to further elucidate the potential of these factors to be involved in virulence.

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1.6 Chapter synopsis

This works investigated the mechanisms of protozoan predation resistance in the marine bacteria, V. cholerae and V. vulnificus. In order to evaluate the coincidental evolution hypothesis, the potential dual role of factors possessed by these bacteria in virulence and predation resistance was studied.

In order to elucidate the response of V. cholerae biofilms to protozoan predation, in Chapter 2 (Noorian et al., 2017), whole transcriptome shotgun sequencing of V. cholerae biofilms during predation was employed to identify the transcripts that were significantly differentially expressed when compared to the ungrazed control. Differentially expressed transcripts were further investigated in order to study the response of V. cholerae biofilms to predation.

The hypothesis that grazing resistance may correlate with genotype and place of isolation was investigated in Chapter 3, by evaluation of grazing resistance of different genotypes of V. vulnificus isolated from clinical and environmental sources. Furthermore, to identify possible novel predation resistance mechanisms, this chapter investigates the defence mechanism of one grazing resistant environmental strain by whole genome sequencing and analysis. Genome comparison between the grazing sensitive and resistant strains was used and the involvement of pathogenicity factors were investigated.

Chapter 4, investigates the potential factors involved in the grazing resistance of V. vulnificus by analysing the transcriptome under grazing resistant conditions established in Chapter 3. The differentially expressed transcripts were further studied to elucidate the mechanism used by V. vulnificus in defence against protozoa.

Chapter 5 provides a summary of the work presented in the previous chapters and discusses the implications of this research. Considerations for future research are also suggested.

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2 Chapter 2: Vibrio cholerae hmgA-mediated pyomelanisation confers resistance to predation by A. castellanii

2.1 Introduction

Vibrio cholerae, the causative agent of cholera, persists in brackish and estuarine water systems (Colwell et al., 1977, Huq et al., 1990) where it is exposed to starvation conditions, fluctuations in temperature and salinity, and predators (Lutz et al., 2013). The continuous presence and growth of V. cholerae in the environment indicates it is able to respond to each one of these stresses (Lutz et al., 2013, Sun et al., 2015). Heterotrophic protists are the biggest consumers of bacteria in the environment and are thus a major mortality factor for bacteria (Jürgens & Matz, 2002).

In benthic marine, brackish, and freshwater sediments where V. cholerae naturally occurs, ciliates are the most abundant protists, while amoebae contribute most to predator biomass (Lei et al., 2014). V. cholerae shares an ecological niche with the model protozoa, Acanthamoeba castellanii and Tetrahymena pyriformis. The free-living amoeba, Acanthamoeba spp. have been isolated from various fresh and salt water sources (Khan, 2006) where they feed on bacterial biofilms. V. cholerae and Acanthamoeba spp. were detected in water samples collected from different cholera endemic areas in Sudan (Shanan et al., 2011). V. cholerae is often isolated from freshwater systems (Nair et al., 1988) where T. pyriformis typically occurs, feeding on bacterioplankton (Elliott, 1970). These predators are among the few axenic protozoan cultures available, making them ideal ecologically relevant model organisms.

Both clinical and environmental strains of V. cholerae have been shown to associate with a range of amoeba (Thom et al., 1992, Abd et al., 2005, Abd et al., 2007) and Van der Henst et al. (2016) showed that V. cholerae can grow inside A. castellanii. A study using laboratory microcosms of natural bacterioplankton communities from the Gulf of Mexico showed elimination of V. cholerae by ciliates and heterotrophic nanoflagellates (HNFs) (Martínez Pérez et al., 2004). In contrast, when V. cholerae biofilms were exposed to

48 | P a g e predation by flagellates, there was little effect on biofilm biomass indicating that biofilms are protected from predation (Matz et al., 2005).

Biofilms provide physical protection along with a high cell density population that enables cell-to-cell communication, or quorum sensing. Quorum sensing (QS) has been shown to regulate antiprotozoal activities in V. cholerae biofilms including Vibrio polysaccharide (VPS) that protects both early and late stage biofilms from predation by the surface feeding nanoflagellate, Rhynchomonas nasuta and the amoeba A. castellanii (Sun et al., 2013). The extracellular protease, PrtV is involved in grazing resistance against the flagellate Cafeteria roenbergensis and the ciliate, T. pyriformis (Vaitkevicius et al., 2006). The type VI secretion system (T6SS) uses virulence-associated secretion (VAS) proteins that are involved in cytotoxicity towards the amoebae, Dictyostelium discoideum and cultured mammalian macrophages (Pukatzki et al., 2006, Miyata et al., 2011). Despite the fact that the early and late biofilms of a V. cholerae QS mutant were more susceptible to grazing by A. castellanii, C. roenbergensis and R. nasuta than the wild type, it was not completely eliminated by predation (Erken et al., 2011, Sun et al., 2013), suggesting other anti-predation strategies could be present. Studies on bacterial prey and protozoan predators have shown several potential defences against grazing, including production of toxins, microcolony formation and changes in cell surface properties (Matz & Kjelleberg, 2005).

In order to study the factors contributing to grazing resistance of V. cholerae, the transcriptome of biofilms exposed to A. castellanii was analysed to identify genetic features that likely contribute to survival during predation. Here, the effect of down- regulation of genes involved in tyrosine degradation on grazing resistance of V. cholerae was examined. A decrease in the activity of homogentisate 1, 2 - dioxygenase (HmgA) leads to accumulation of homogentisic acid (HGA) which auto-oxidizes to form pyomelanin (Turick et al., 2010). Results show that the production of pyomelanin has a protective effect against predation by A. castellanii.

The work in this chapter has been published; Noorian Parisa, Hu Jie, Chen Zhilliang, Kjelleberg Staffan, Wilkins Mark, Sun Shuyang & McDougald Diane. Pyomelanin

49 | P a g e produced by Vibrio cholerae confers resistance to predation by Acanthamoeba castellanii. FEMS Microbiology Ecology 2017; 93: fix147 (Noorian et al., 2017).

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2.2 Materials and methods

2.2.1 Strains and growth conditions

Organisms used in this study are listed in (Table 2.1). Bacterial strains were routinely grown in Lysogeny broth (LB) and on agar plates (Sambrook et al., 1989) as appropriate, with carbenicillin (100 µg ml−1). A. castellanii was routinely passaged in 15 ml growth medium containing peptone-yeast-glucose (PYG) (20 g l-1 proteose peptone, 1 g l-1 yeast -1 -1 extract) supplemented with 1 litre 0.1 × M9 minimal medium (6 g l NaH2PO4, 3 g l -1 -1 2 K2PO4, 0.5 g l NaCl, 1 g l NH4Cl) and 0.1 M sterile-filtered glucose in 25 cm tissue culture flasks with ventilated caps (Sarstedt Inc., Nümbrecht, Germany) and incubated statically at 30°C. A. castellanii was passaged 3 days prior to harvesting for experiments and enumerated microscopically using a haemocytometer.

The browsing ciliate, T. pyriformis was maintained as above but incubated statically at room temperature (RT). Prior to experiments, 500 μl of T. pyriformis were passaged in -1 -1 20 ml of 0.5 × nine salts solution (NSS) medium (8.8 g l NaCl, 0.735 g l Na2SO4, 0.04 -1 -1 -1 -1 -1 g l NaHCO3, 0.125 g l KCl, 0.02 g l KBr, 0.935 g l MgCl2.6H2O, 0.205 g l -1 -1 CaCl2.2H2O, 0.004 g l SrCl2.6H2O and 0.004 g l H3BO3) (Mårdén et al., 1985) supplemented with 1% heat- killed P. aeruginosa PAO1 (HKB) in a 25 cm2 tissue culture flask, and further incubated at RT statically for 2 days before enumeration and use. This process is necessary to remove the nutrient media and to acclimatise the ciliate to phagotrophic feeding.

To prepare heat-killed bacteria (HKB), P. aeruginosa was grown overnight in LB at 37°C 9 -1 with shaking at 200 rpm and adjusted to (OD600=1.0; 10 cells ml ) in 0.5 × NSS. The tubes were then transferred to a water bath at 65°C for 2 hours, and then tested for viability by plating on LB agar plates at 37°C for 2 days. HKB stocks were stored at -20°C.

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Table 2-1 Strains and plasmids used in this study. Strain Properties Reference/Source Bacterial strains

V. cholerae A1552 Wild type, O1 El Tor, Inaba, smooth, Rifr (Valeru et al. 2009)

V. cholerae A1552 O1 El Tor, Inaba, smooth, ΔhmgA, Rifr, Kmr (Valeru et al. 2009) hmgA

V. cholerae A1552 O1 El Tor, Inaba, smooth, ΔhmgA, (Valeru et al. 2009) hmgA complement hmgA::pUC18, Rifr, Apr, Kmr

V. cholerae A1552 O1 El Tor, Inaba, smooth, pUC18, Rifr, Apr This study pUC18

V. cholerae A1552 O1 El Tor, Inaba, smooth, ΔhmgA, pUC18, This study hmgA pUC18 Rifr, Apr, Kmr

Plasmids

pUC18 Cloning vector, pMB1 ori, LAC pr, lacZ, APr (Yanisch-Perron et al. 1985) Protozoan strains

A. castellanii Wild type ATCC 30234

T. pyriformis Wild type ATCC 205063

2.2.2 Transcriptomic profiling of continuous-culture biofilms

For the transcriptomic analysis, 3-days-old V. cholerae biofilms were exposed to grazing by A. castellanii in a continuous flow system. Briefly, 3 biological replicates of V. cholerae biofilms were cultivated on the interior surfaces of Silastic® laboratory tubing (Dow Corning, Michigan, USA) (3.2 mm diameter; length, 14 cm) in 0.5 × Väätänen nine salts solution (VNSS) (1 g bacteriological peptone, 0.5 g yeast extract, 0.5 g D-glucose,

0.01 g FeSO4·7H2O and 0.01 g Na2HPO4) in 1 litre of 0.5 × NSS (Väätänen, 1977) and fed at a flow rate of 9 ml h-1 using a continuous flow system at RT. After 3 days, washed cells of A. castellanii were resuspended in 0.5 × VNSS, injected into tubing and incubated

52 | P a g e without flow for 2 hours. A protist free control biofilm was treated the same to exclude oxygen or starvation effects.

The V. cholerae biofilms on the walls of the tubing were washed by a flow of 2 volumes of 0.5 × VNSS to remove planktonic bacteria, and immediately resuspended in 2 volumes of RNAlater (Qiagen, Hilden, Germany) and harvested from the interior surface of the tubing by mechanical manipulation, i.e. by squeezing the biofilm out of the tubes. Total RNA was extracted by lysozyme digestion and use of the RNeasy® plus mini kit (Qiagen, Hilden, Germany) according to the manufacturer’s instructions. For the mRNA-Seq sample preparation, the Illumina standard kit was used, according to the manufacturer’s protocol (Illumina, USA).

2.2.3 Transcriptome data analysis

Prior to RNA-Seq analysis, filters were applied to remove low quality reads from all pair- end samples. Pair-end raw reads were trimmed with the BWA trimming mode at a threshold of Q13 (P = 0.05) as implemented by SolexaQA version 3.1.3 (Cox et al., 2010). Low-quality 3’ ends of each read were filtered and reads that were less than 25 bp in length were discarded.

The trimmed reads were subsequently depleted of ribosomal RNA with SortMeRNA version 1.8 (Kopylova et al., 2012). Trimmed reads (102 bp) were first mapped to the A. castellanii contigs (GenBank accession ID GCA_000826485.1) using Bowtie (version 2.2.3) (Langmead & Salzberg, 2012) with default parameters. Reads that were not mapped to amoeba contigs were then mapped to the reference genome (V. cholerae O1 biovar El Tor str. N16961) and V. cholerae A1552 indel correction table (http://microbes.ucsc.edu/lists/vibrChol1/StrainA1552-list.html) using Bowtie2 with parameters set to -N 1. Cuffdiff (Cufflinks version 2.2.1) with default parameters was finally used to identify differentially expressed transcripts of V. cholerae biofilms grazed by A. castellanii compared to ungrazed controls. Cuffdiff calculated the log fold change in FPKM, and then the significance of the fold change. A false discovery rate (FDR) adjusted p-value was calculated to give the statistical validity level of significance. Transcripts with an FDR-adjusted p-value of <0.05 were considered to be SDE genes.

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The lists of up and down regulated SDE genes were placed into COG categories by NCBI conserved domain search (Tatusov et al., 1997). With the assistance of the Database for Annotation, Visualisation and Integrated Discovery (DAVID) Bioinformatics Resources 6.7 (National Institute of Allergy and Infectious Diseases, NIH), the differentially expressed transcripts were further analysed using databases such as Gene Ontology Annotation Database (to analyse the biological processes, molecular functions and cellular components), KEGG Pathway (to analyse the metabolic pathways), and InterPro/UniProt (to analyse the protein domains).

2.2.4 Early and late biofilm grazing assay with A. castellanii

Overnight cultures of V. cholerae were adjusted so that 105 cells ml-1 in 0.5 × VNSS were added to 24-well microtitre plates (Falcon™, Becton Dickinson, New Jersey, USA) and incubated for 24 and 72 hours with shaking at 60 rpm at RT. After incubation, media was refreshed with or without A. castellanii (2 × 104 cells ml-1) and incubated at RT with shaking at 60 rpm for 3 days. The cell density of each well was measured by 2 spectrophotometry at OD600 nm (Wallac Victor 1420 Multilabel Counter, Perkin Elmer Life Sciences, Massachusetts, USA). In order to quantify the biofilm biomass, crystal violet (CV) assays were performed (O'Toole & Kolter, 1998). Briefly, all the planktonic cells were removed by was washing three times with 0.5 × NSS before adding crystal violet (0.3%) for 15 minutes. The wells were washed a further three time using 0.5 × NSS to remove the unbound CV and then the stain was solubilized using 96% ethanol and the OD490 nm was determined by spectrophotometry.

To determine if the cell-free supernatants from the hmgA mutant would provide protection against grazing by A. castellanii to the WT, the cell-free supernatant of 3-days-old established biofilms were acquired by centrifugation at 6000 × g for 10 minutes and filtration (0.22 mm filters, Millex-GP, Millipore, MA, USA). The cell free supernatant was then added to 3-days- old established biofilms of the WT strain at a ratio of 50% with fresh VNSS with or without A. castellanii (2 × 104 cells ml-1) and incubated at RT with shaking at 60 rpm for 3 days. The biofilm biomass was then quantified by CV assays.

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2.2.5 Quantification of pyomelanin and reactive oxygen species

The amount of pyomelanin in the aqueous phase was determined by spectrophotometry

(OD405 nm) of the cell-free supernatant acquired by centrifugation at 6000 × g for 10 minutes and filtration (0.22 mm filters, Millex-GP, Millipore, MA, USA). In order to study the effect of nutrients released from A. castellanii on the pyomelanin production by V. cholerae, A. castellanii was incubated in 0.5 × VNSS with or without 1% HKB for 3 days at RT. Furthermore, to assess if phagocytosis by the amoeba predator is required for induction of pyomelanin production, A. castellanii was heat inactivated at 65°C for 15 minutes. The trophozoites were confirmed to be intact using microscopy, and the viability checked by addition to PYG and incubation at RT for 3 days. The cell-free supernatant or heat-killed A. castellanii was added to the 3 days old established biofilm. The amount of pyomelanin was then determined in the cell free supernatant after incubation for 3 days at RT.

To assess the level of reactive oxygen species (ROS), 25 μM dihydroethidium (DHE) (Sigma-Aldrich, MO, USA), a fluorescent dye for detection of intracellular O2- was used (Owusu-Ansah et al., 2008). The biofilms were washed with 0.5 × NSS after which 25 μM DHE in fresh 0.5 × VNSS medium was added and incubated in the dark for 2 hours. After incubation, the cells were washed with 0.5 × NSS, and the ROS production was determined by spectrophotometer (518 and 605 nm for excitation and emission, respectively). The plates were incubated for 3 days before measurement of pyomelanin as described.

2.2.6 H2O2 treatment of V. cholerae biofilms

The overnight cultures were inoculated at a final concentration of 106 cells ml-1 in 0.5 × VNSS in 24-well plates incubated at RT with shaking at 60 rpm. After 3 days, the biofilms were treated with 30 mM H2O2 for 30 minutes, after which the H2O2 was removed and fresh 0.5 × VNSS medium with or without A. castellanii (8 × 103 cells ml- 1) was added. Protozoal controls were supplied with 1% HKB as food. After co-

55 | P a g e incubation for 3 days, the amoebae were enumerated by inverted microscopy (ECLIPSE TE2000-5, Nikon, Japan) (five images per well) and the V. cholerae biofilm biomass was quantified using the CV assay.

2.2.7 Catalase treatment of V. cholerae biofilms

The overnight cultures were inoculated at a final concentration of 105 cells ml-1 in 0.5 × VNSS in 24-well plates incubated at RT with shaking at 60 rpm. After 3 days, the media was refreshed with or without A. castellanii (2 × 104 cells ml-1) and 0.1 mg ml-1 catalase (Sigma-Aldrich, MO, USA) was added. After co-incubation for 3 days, the V. cholerae biofilm biomass was quantified using the CV assay.

2.2.8 Experimental validation - qRT-PCR

RNA was prepared from late biofilm grazing assay with A. castellanii in 24 well plates. After removal of supernatant, RNAlater (Qiagen, Hilden, Germany) was added and cells were harvested from the wells by mechanical manipulation. RNA extraction was then performed as described previously. The concentration was measured using a spectrophotometer (NanoDrop ND-1000; NanoDrop Technologies) after treatment with TURBO™ DNase (Ambion- Life Technologies). DNA was prepared from 500 ng RNA from each sample by iScript™ Reverse Transcription (Bio-Rad). Quantitative reverse transcriptase PCR (qRT-PCR) experiments were done using PowerUp™ SYBR™ Green Master Mix (Applied Biosystems) by QuantStudio™ 6 Flex Real-Time PCR System using the primers specific for VC1344, VC1345, VC1346 and VC1347 listed in (Table 2-2). The expression was determined relative to the expression of the endogenous control gene gyrA using the comparative Ct (DDCt) method of reverse-transcriptase PCR (RT- PCR).

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Table 2-2 Primers used in this study. Gene Forward Reverse VC1344-F, 5′- VC1344-R-5’- VC1344 TTATGGCGATCGCAGCATCT CACAGGTTCATGTTGCCACG VC1345-F, 5′- VC1345-R-5’- VC1345 GTCTAGAGCCGATTGAGCCC TTGATCGAAGGCACGTCCAA VC1346-F 5’- VC1346-R-5’- VC1346 ATGCGTGATGGTCAACTGGT TCCAACGCCTGCTGTAATGT VC1347-F, 5′- VC1347-R- 5′- VC1347 CCGCCTATCGAGTGCGTATT AACTCGCTAGGATTGAGGCG gyrA-F, 5′- gyrA-R, 5′- gyrA CACGAACTCTTGGCAGACCT CAATACCAGATGCGCCGTTG

2.2.9 T. pyriformis grazing assays

Microtitre plates containing 1-day-old biofilms were prepared as described above. After 24 hours, the supernatants were removed and fresh 0.5 × VNSS media with or without T. pyriformis was added (103 cells ml-1; determined by inverted microscopy) and the plates were incubated at RT with shaking at 60 rpm for 3 days. The cell density was measured by spectrophotometry at OD600 nm. Planktonic fractions were collected for determination of CFU ml-1 and biofilm biomass was determined by CV staining and spectrophotometry

(OD490 nm). Numbers of T. pyriformis were determined by microscopy at each sampling time and pyomelanin was measured in the cell-free supernatants as described previously.

2.2.10 V. cholerae - A. castellanii intracellular survival assay

To determine the role of hmgA in the intracellular survival of V. cholerae internalised by A. castellanii, number of internal V. cholerae was measured after a 24-hour time point. Briefly, A. castellanii (2 × 105 cells ml-1) in 0.5 × NSS and 1% HKB were seeded in 24 well microtitre plates one day prior to the start of the experiment until confluent on the bottom of the wells. On the day of experiment, the wells were washed gently with 0.5 × NSS and V. cholerae (107 cells ml-1) in 0.5 × NSS were added to wells containing A.

57 | P a g e castellanii. Plates were incubated for 1 hour statically at RT to allow internalisation of V. cholerae. To remove all the extracellular bacteria, the wells were washed three times with cells 0.5 × NSS and further treated with gentamicin (300 μg ml-1) for 1 hour at RT. The gentamicin was then removed by washing three time with 0.5 × NSS. The cells were then incubated in 0.5 × NSS at RT statically for 24 hours, after which the amoeba cells were lysed by addition of 1% Triton X-100 in 0.5 × NSS for 20 minutes. The numbers of bacteria were enumerated by drop plate colony counting.

2.2.11 Data analysis

Statistical analysis was performed using GraphPad Prism version 7.01 for Windows, GraphPad Software, La Jolla California USA, (www.graphpad.com). Data that did not follow Gaussian distribution was determined by analysing the frequency distribution graphs and was transformed using natural log. Two-tailed student’s t-tests were used to compare means between experimental samples and controls. For experiments including multiple samples, one-way or 2-way ANOVAs were used for the analysis and Sidak's or Dunnett's multiple comparison Test provided the post-hoc comparisons of means when appropriate.

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2.3 Results and discussion

The current study was designed to further elucidate antiprotozoal activities generated by V. cholerae biofilms. Heterotrophic protists are major predators of bacteria, and consequently, bacteria have evolved both pre- and post-ingestional defence strategies to resist predation (Matz & Kjelleberg, 2005). Such defence strategies employed by V. cholerae include biofilm formation (Matz et al., 2005), expression of the PrtV protease (Vaitkevicius et al., 2006) and the T6SS (Pukatzki et al., 2006). Although a QS mutant of V. cholerae was more sensitive to predation than the corresponding isogenic wild type, it was still partially resistant to grazing (Erken et al., 2011), implying the existence of other QS-independent anti-protozoal mechanisms.

2.3.1 RNA-seq revealed differences in the transcriptomes of grazed and ungrazed biofilms

In order to identify other anti-predation strategies employed by V. cholerae biofilms, RNA-Seq was performed. Total RNA isolated from 3 biological replicates of biofilms exposed to grazing by A. castellanii was subjected to Illumina HiSeq 2000 sequencing. Between 108 and 127 million pairs of reads were generated with approximately 5 million reads per sample removed after quality filtering and trimming. Between 98.31 and 99.42% of reads were mapped to the V. cholerae N16961 genome and approximately

0.13% were mapped to A. castellanii contigs. The log2 fold change in fragments per kilobase of exon per million fragments mapped (FPKM) varies from -1.964 to -0.724 for the down-regulated transcripts, and from 0.797 to 3.535 for the up-regulated transcripts.

Significantly differentially expressed genes were considered at fold-change of 2.0 and adjusted p-value of p<0.05. Cuffdiff analysis of the transcriptome revealed that 71 transcripts were significantly up-regulated and 60 were significantly down-regulated in the grazed biofilm compared with the ungrazed control (see Appendix 1 for the complete list of differentially expressed genes).

A relatively large fraction of the up-regulated transcripts corresponds to genes involved in metabolism, in particular nucleic acid, amino acid, lipid and carbohydrate transfer and

59 | P a g e metabolism. These transcripts encode proteins associated with amino acid biosynthesis and metabolism, such as VC0027 (threonine metabolism), VC1061 (cysteine biosynthesis), hisD, hisG, hisH, VC1134, VC1135, VC1137, VC1138 and VC1139 (histidine metabolism), trpA (tryptophan biosynthesis), gltD and VC2373 (glutamate biosynthesis), glnA (glutamine biosynthesis) argC, VC2617, VC2641, VC2642, VC2643, and VC2508 (arginine metabolism and biosynthesis), VC1704 (cysteine and methionine metabolism), VC0162, VC0031 and VC0028 (isoleucine biosynthesis) and VC0392, VCA0604 and VCA0605 (aminotransferases). The increase in metabolism and energy production might be related to an increase in available nutrient resources since feeding will result in the release of nutrients by protozoa, either due to ‘sloppy feeding’ or excretion of waste products (Wang et al., 2009) .

The genes in the tyrosine catabolic pathway (VC1344 to VC1347) were down-regulated in the grazed samples compared to ungrazed samples. These genes lead to the catabolism of tyrosine to fumarate and acetoacetate (Valeru et al., 2009, Wang et al., 2011), and are confirmed to be associated with a pigmented phenotype due to pyomelanin production (Ivins & Holmes, 1980, Ivins & Holmes, 1981, Ruzafa et al., 1995). The enzyme HmgA (VC1345) is involved in L-tyrosine catabolism in both prokaryotic and eukaryotic organisms (Fernández-Cañón & Peñalva, 1995, Kotob et al., 1995) and a null mutation in hmgA in V. cholerae leads to accumulation and auto-oxidation of HGA, which in turns will lead to production of pyomelanin (Figure 2-1). The down regulation of VC1344 -

VC1347 during predation was confirmed by qRT-PCR (log2 fold change of -1.51, -1.21, -1.67, and -1.45 respectively).

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Figure 2-1 Differentially expressed transcripts in grazed compared to ungrazed biofilms involved in tyrosine degradation in V. cholerae. FC represents log 2 fold changes. The pathway in V. cholerae is proposed by Valeru et al. (2009).

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Melanin production by auto-oxidation of HGA has been shown to occur in many organisms, ranging from bacteria to humans. The gene encoding HmgA as well as the rest of the pathway is well conserved, and a BLAST search of the NCBI database revealed that many Vibrio spp., including non-pathogenic environmental Vibrio spp. have this pathway. The protective mechanisms of melanin are unclear, but melanin (charged polymers) present in the cell wall may serve as a physical or chemical barrier (Nosanchuk & Casadevall, 1997, Jacobson, 2000, Eisenman et al., 2005). An hmgA mutant of V. cholerae exhibited greater UV and oxidative stress resistance, increased expression of a subunit of TCP and CT, and was enhanced in its ability to colonise the infant mouse (Valeru et al., 2009). In contrast, a Vibrio campbellii hmgA mutant did not show increased UV resistance and was less virulent than the wild type strain, although the wild type strain exhibited higher resistance to oxidative stress when incubated with supernatants from the hmgA mutant (Wang et al., 2013).

Pigment production has also been demonstrated to provide a range of functions in many different microorganisms. For example, melanin can protect the pathogenic fungus, Cryptococcus neoformans, from antibody-mediated phagocytosis by macrophages (Wang et al., 1995), as well as from digestion of phagocytosed cells by the amoeba A. castellanii (Steenbergen et al., 2001). Melanised C. neoformans are significantly less susceptible to hydrolytic enzymes commonly used by environmental predators than non- melanised cells (Rosas & Casadevall, 2001). Melanin production in the fungus, Paracoccidioides brasiliensis, increases protection from phagocytosis by macrophages and intracellular resistance, and decreased drug susceptibility (da Silva et al., 2006), while in the yeast Exophiala (Wangiella) dermatitidis, melanin production prevented killing by the phagolysosomal oxidative burst of human neutrophils (Schnitzler et al., 1999).

2.3.2 Pyomelanin production increases the grazing resistance of V. cholerae biofilms

In order to determine the role of pyomelanin in grazing resistance of biofilms, V. cholerae wild type and hmgA mutant strains were allowed to form biofilms and after 1 or 3 d, either

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T. pyriformis or A. castellanii were added. After 3 d of grazing, CFU and CV measurements determined planktonic cell and biofilm biomass, respectively.

The grazing resistance of early biofilms (1-day-old) of the V. cholerae hmgA mutant and wild type strains in the presence of A. castellanii were not significantly different (Figure 2-2A). In contrast, when late biofilms (3 d) were exposed to grazing by A. castellanii, the hmgA mutant was significantly more grazing resistant than the wild type. The biofilm biomass of the grazed wild type is reduced by 7.3% compared to the ungrazed control, whereas the hmgA mutant shows 16.5% increase in biofilm biomass after grazing (Figure 2-2B).

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r W c g c e u m u e m h p p le P T A p W g m m o h c A g m h Figure 2-2 Biofilm biomass of early (A) and late (B) biofilms of V. cholerae A1552 exposed to grazing by A. castellanii for 72 hours. Biofilm biomass was determined by CV staining. Data were natural log transformed and the percentage change of biofilm biomass was calculated by removing the biomass of ungrazed samples from the grazed samples divided by the ungrazed. The experiment was run in three replicates and repeated three times separately. Error bars represent standard deviation. Small letters indicate different statistical groups derived from 1-way ANOVA and Dunnett's multiple comparisons tests. Statistical significance is indicated by (** p < 0.001; ***, p < 0.0001 and Ns, not significant).

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Control ungrazed biofilms of the WT and hmgA mutant strains were not significantly different, indicating that the biofilm growth for both strains was similar (Figure 2-3). Furthermore, the cell-free supernatant of the hmgA mutant does not show toxicity towards A. castellanii trophozoites compared to the WT (Figure 2-4).

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Figure 2-3 Biofilm biomass of V. cholerae wild type and hmgA mutant grown for 1 or 3 d as determined by CV staining. The experiment was run in triplicate and repeated three times. Error bars represent standard deviation. Statistical analysis was performed using 1-way ANOVA and Sidak's multiple comparisons test which revealed no significant differences, p>0.2.

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0 h o u rs 2 4 h o u rs

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Figure 2-4 The number of A. castellanii trophozoites (A) and cysts (B) after 0 and 24 - hour incubation in cell-free supernatants of wild type and hmgA mutant strains of V. cholerae. Error bars represent standard deviation. Statistical analysis was performed using 2-way ANOVA and Sidak's multiple comparisons test which revealed no significant differences, p=0.8 (A) and p>0.9 (B).

The resistance of planktonic cells of the hmgA mutant to predation by T. pyriformis was also investigated, as A. castellanii cannot feed efficiently on planktonic cells (Huws et al., 2005). T. pyriformis is a filter-feeding ciliate that can feed effectively on early

66 | P a g e biofilms as well as planktonic cells (Parry, 2004). The use of a second type of grazer with different feeding mechanisms and niche is important for establishment of the generality of a grazing resistance mechanism. The early biofilm (1-day-old) biomass (Figure 2-5A) and numbers of planktonic cells (Figure 2-5B) of the hmgA mutant and wild type strains in the presence of T. pyriformis were not significantly different. Interestingly, a further increase in pyomelanin production by the hmgA mutant was observed after 3 d of grazing by A. castellanii but not when exposed to grazing by T. pyriformis (Figure 2-6), which is consistent with the increased grazing resistance of the mature biofilm against A. castellanii.

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P Figure 2-5 Early biofilms of V. cholerae A1552 exposed to T. pyriformis for 72 hours. Biofilm biomass was determined by CV staining (A) and the planktonic cells in the supernatant enumerated by the drop plate method (B). Data was natural log transformed and the percentage change in biofilm biomass was calculated by removing the ungrazed biomass from the grazed biofilm biomass divided by the ungrazed. The experiment was run in three replicates and repeated three times separately. Error bars represent standard deviation. Statistical analysis was performed using student's t-test which revealed no significant difference, p=0.3 (A) and p= 0.1 (B).

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o T A N W g m h Figure 2-6 Amount of pyomelanin produced by ungrazed or grazed established biofilms after 3 d exposure to A. castellanii (A) or T. pyriformis (B). Pyomelanin secreted by the biofilm into the supernatant was measured by optical density (OD 405 nm) of the cell-free supernatant obtained from the biofilms. Amount of pyomelanin was then normalised by using the corresponding biofilm biomass measured by CV assays (OD 490 nm). Experiments were run in triplicates and repeated 3 times on different days. Error bars represent the standard deviation of three replicates. Bars indicate different statistical groups derived from 2-way ANOVA and Sidak's multiple comparisons test. Statistical significance is indicated by (****, p < 0.0001).

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Addition of more nutrients to WT and hmgA mutant strains did not result in the same increase in pyomelanin levels as active grazing by A. castellanii. This indicates that the extra nutrients in the A. castellanii culture are not responsible for induction of pyomelanin production in the hmgA mutant when exposed to grazing by A. castellanii. Furthermore, the addition of cell-free supernatants from A. castellanii with or without HKB and heat- killed A. castellanii cells did not induce overproduction of pyomelanin, supporting our hypothesis that active phagocytosis by A. castellanii is required (Figure 2-7).

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o ii ii S B n B n N S K K la la N H l H l V te te % s % s 1 a 1 + + a c ii c S . . A n A S la N + l te d V S s le S a il N c k V . t A a e + H S S + N S V S N V

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N ii ii S B n B n S K K la la N H l H l V te te % s % s 1 a 1 + + a c ii c S . . A n A S la N + l te d V S s le S a il N c k V . t A a e + H S S + N S V S N V Figure 2-7 Pyomelanin production by V. cholerae wild type (A) and hmgA mutant (B) biofilms. Supernatants of A. castellanii were obtained after incubation in 0.5 × VNSS with or without 1% HKB for 72 hours at RT. Cell-free supernatants were added after 72 hours incubation of biofilms. Pyomelanin secreted by the biofilm into the supernatant was measured by optical density (OD 405 nm) of the cell-free supernatant obtained from the biofilms. Amount of pyomelanin was then normalised by using the corresponding biofilm biomass measured by CV assays (OD 490 nm). Experiments were run in triplicate and repeated three times. Error bars represent standard deviation. Statistical analysis was performed using 1-way ANOVA and Dunnett's multiple comparisons tests comparing to 0.5 × VNSS controls which revealed no significant differences, p>0.1 (A) and p>0.5 (B).

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2.3.3 Pyomelanin and reactive oxygen species (ROS) concentrations

The increases in grazing resistance of the hmgA mutant biofilms (Figure 2B) correlated with an increase in pigment production. On day one, the pyomelanin concentration in both the supernatant of wild type and hmgA mutant strains was low (normalised pigment production (OD405nm/Biofilm biomass) = 0.0013 and 0.0009 respectively) (Figure 2-8A). However, after 3 d the pyomelanin concentration in supernatants of the hmgA mutant was twenty-fold higher than those of the wild type (normalised pigment production

(OD405nm/Biofilm biomass) = 0.0016 and 0.033 respectively).

In order to further investigate the relationship between grazing and pigment production, the amount of O2- generated in biofilms of the V. cholerae A1552 wild type, hmgA mutant and complemented strain was monitored, as it has been suggested that ROS are generated during pigment production (Valeru et al., 2009). Notably, when pigment production increased on day 3 in the hmgA mutant, the ROS level also increased (Figure 2-8B). Biofilms of the hmgA mutant strain generated 79% more O2- than the A1552 biofilms (p =0.0259)

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Figure 2-8 Amount of pyomelanin produced by biofilms of V. cholerae A1552 wild type and hmgA mutant strains after 1 and 3 d (A). Amount of ROS in the cell-free supernatant of 3-days-old biofilms of V. cholerae A1552 wild type and hmgA mutant (B). Pyomelanin secreted by the biofilm into the supernatant was measured by optical density (OD 405 nm) of the cell-free supernatant obtained from the biofilms. Amount of pyomelanin was then normalised by using the corresponding biofilm biomass measured by CV assays (OD 490 nm). Error bars represent the standard deviation of three replicates. Statistical analysis was performed using 2-way ANOVA and Sidak's multiple comparisons test (A) and Student's t-test (B). Statistical significance is indicated by (**, p < 0.001, ****, p < 0.0001).

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Figure 2-9 Effect of cell-free supernatants (A), H2O2 (B) and catalase (C) on grazing resistance to A. castellanii. The cell-free supernatant of 3-days-old V. cholerae biofilms were added to 3-days-old biofilms at a concentration of 50% in fresh VNSS and incubated with A. castellanii for 3 d. Biofilm biomass was determined by CV staining. Experiments were run in triplicate and repeated 3 times on different days. Error bars represent the standard deviation of three replicates. Statistical analysis was performed using 1-way ANOVA and Dunnett's multiple comparisons test comparing all to the WT (A), Student's t-test (B) and 2-way ANOVA and Sidak's multiple comparisons test (C). Statistical significance is indicated by (*, p < 0.05; ***, p < 0.001 and ****, p < 0.0001 and Ns, not significant).

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To determine if the hmgA cell-free supernatant would provide protection against predation by A. castellanii to the WT strain, the cell-free supernatant of 3-day-old established biofilms were added to the A. castellanii grazing experiments. The addition of undiluted cell-free supernatants of V. cholerae to pre-established biofilms would lead to their dispersal due to lack of nutrients and accumulation of waste. Therefore, cell-free supernatants were diluted with fresh VNSS, and results showed that at a concentration of 50%, the cell-free supernatant of the hmgA strain significantly increased the WT biofilm resistance to grazing by A. castellanii (Figure 2-9A).

2.3.4 Hydrogen peroxide addition increased the grazing resistance of V. cholerae

Previous studies have shown that exposure of V. cholerae to H2O2 induced oxidative stress responses and virulence factor expression (Valeru et al., 2009). The auto-oxidation of HGA can generate superoxide radicals and H2O2 in eukaryotic cells at physiological pH (Martin Jr & Batkoff, 1987). Here, the effect of addition of H2O2 as a substitute for pyomelanin-associated ROS on resistance of V. cholerae biofilms to amoebae grazing was tested in order to further confirm the pyomelanin/ROS-mediated grazing resistance.

V. cholerae biofilms were pre-grown for 3 d, followed by exposure to H2O2 for 30 min

(Figure 2-9B). A. castellanii was added after H2O2 exposure and the culture incubated for 3 d. The health and number of A. castellanii was monitored and there was no difference between the total numbers of amoebae when co-incubated with the V. cholerae biofilms that had been exposed to H2O2 compared to the controls with HKB. After 3 d of grazing by A. castellanii, the biomass of the untreated V. cholerae biofilms were significantly reduced while biofilms that had been treated with H2O2 for 30 min, were not reduced (p = 0.0246) (Figure 2-9B). In addition, pre-grown 3-days-old biofilms were treated with 0.1 mg ml-1 catalase to reduce ROS of the hmgA mutant biofilm. After 3 d of grazing by A. castellanii, the biomass of the treated V. cholerae hmgA biofilm was significantly reduced compared to the untreated biofilm (p = 0.0176) (Figure 2-9C). Taken together, our results suggest that the production of pyomelanin results in production of ROS, which in turn, results in an increase in grazing resistance.

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The role of pyomelanin in survival of V. cholerae inside A. castellanii was investigated. The total number of V. cholerae cells associated with A. castellanii (extracellular and intracellular) as well as the number of intracellular V. cholerae was determined, and there was no difference between wild type and hmgA mutant strains (Figure 2-10).

3 0 0 0 0

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Figure 2-10 Number of intracellular V. cholerae after 24-hour co-incubation with A. castellanii. Error bars represent standard deviation. Statistical analysis was performed using student's t-test which revealed no significant differences, p> 0.9.

Pigmented hypertoxinogenic strains of V. cholerae have been previously reported both in random mutagenesis experiments (Mekalanos et al., 1979, Parker et al., 1979, Ivins & Holmes, 1980) as well as isolates obtained from the environment. For example, V. cholerae, ATCC 14035 serotype Ogawa serovar O1 strain isolated originally from a stool sample produced a reddish brown pigment when grown in low nutrient condition media supplemented with L-glutamic acid and L-tyrosine (Ruzafa et al., 1995). In addition, six nontoxigenic serogroup O139 (water isolates) and one toxigenic O1 (clinical isolate) strains isolated from different years and from different provinces of China were pigmented. All the O139 strains had the same 15-bp deletion in hmgA and a 10-bp deletion was found in the VC1345 gene of the O1 strain, indicating that the mutation of this gene may provide a fitness advantage in the environment (Wang et al., 2011).

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2.3.5 Conclusion

Overall this study demonstrates that V. cholerae O1 El Tor alters its transcriptome in the presence of the predator, A. castellanii. One metabolic pathway that was down-regulated under grazing pressure was the tyrosine catabolic pathway, resulting in accumulation of pyomelanin. Experiments with a pyomelanin-overproducing mutant demonstrate that it is more resistant to predation by A. castellanii than the isogenic wild type. Furthermore, the hmgA mutant produces more ROS, which may account for the increased grazing resistance of the hmgA mutant, as V. cholerae biofilms pre-treated with H2O2 were also more grazing resistant.

This chapter provides insight into the genes involved in defence against protozoan grazing of V. cholerae. Data presented here shows that the expression of pyomelanin aids in protection of V. cholerae from grazing in the environment and previous reports have shown that it also plays a role in virulence factor expression and colonisation ability (Valeru et al., 2009). This further supports our hypothesis that predation is a major selective factor for maintenance of virulence genes in the environment and thus melanin production may be one such dual use virulence factor.

2.4 Acknowledgements

Illumina HiSeq 2000 sequencing was performed in the Ramaciotti Centre for Genomics. I would like to thank Dr. Zhilliang Chen for help in RNA-seq analysis. I would also like to thank Hu Jie for help with RNA extraction and quantification of reactive oxygen species.

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3 Chapter 3: Characterisation of the antiprotozoal activity of Vibrio vulnificus

3.1 Introduction

When hurricane Katrina hit New Orleans, floodwaters exposed a large number of people to toxic and contaminated waters. Three months after the devastating storm, seven people were documented as having been infected by Vibrio vulnificus, four of whom were deceased (American Health Care Association/National Centre for Assisted Living (AHCA/NCAL) Update, 2005) (Rhoads, 2006). V. vulnificus is the deadliest seafood- borne pathogen, and infections caused by this organism have the worst prognosis. This organism is found in marine and estuarine waters throughout the world and thrives in warm waters. Most commonly, people with wound or breaks in their skin who are exposed to estuarine or seawater, or who ingest raw or undercooked seafood are at risk of infection (Oliver, 2013).

In order to survive in these aquatic ecosystems, V. vulnificus is equipped with an array of adaptive response mechanisms. V. vulnificus strains are phenotypically, genotypically and serologically heterogeneous and have been classified into different genotypes and biotypes (Tison et al., 1982, Bisharat et al., 1999). Many studies have attempted to find genetic markers for classification of V. vulnificus into groupings that predict environmental persistence, pathogenicity and hence what danger they may present to the human population and to aquaculture.

One such genotyping system is based on the virulence-correlated gene (vcg). Clinical isolates carry the vcgC gene (C-genotypes), while environmental strains (E-genotypes) contain vcgE (Rosche et al., 2005). A study by Warner & Oliver (2008b) revealed that 84.4% of V. vulnificus populations recovered from oysters belonged to the E-genotype whereas isolates recovered from the waters surrounding the oyster sites contain almost equal distributions of the two genotypes. Interestingly, the ratio of C-genotype strains from both sources increased with increasing water temperatures (Warner & Oliver,

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2008b). However, E-genotype isolates are not always less virulent, as some of the most virulent strains carry vcgE (Thiaville et al., 2011).

This presents the question as to which of these genotypes would have a fitness advantage in the aquatic environment under predation by protozoa. Are the environmental strains better equipped to survive the pressure of protozoan predation or does the array of virulence factors possessed by clinical strains provide better protection? It has been reported that environmental strains of E. coli O157:H7 (EcO157) from soil and feral pig faeces survived longer when co-incubated with the sessile ciliate, Vorticella microstoma and the ciliate, Colpoda aspera, isolated from dairy wastewater when compared to clinical isolates (Ravva et al., 2014). In contrast, Legionella pneumophila pathogenesis is associated with survival in both human macrophages and amoeba and strains that do not grow well in the amoebae, Acanthamoeba polyphaga, are rarely isolated from the environment and therefore less likely to cause disease in humans (Patrizia et al., 2013).

The C-genotype strains have higher survival rates in human serum. The presence of the siderophore-encoding gene, viuB (vibriobactin utilisation protein) in all C-genotypes strains but very few E-type strains, is correlated with this serum survival. The addition of iron to human serum restored the survival of viuB lacking E-genotype strains (Bogard & Oliver, 2007). In contrast E-genotype strains integrate better into aggregates (marine snow) that can boost their uptake by oysters (Crassostrea virginica) (Froelich et al., 2013).

In this chapter, the relationship between grazing resistance of V. vulnificus with genotype and place of isolation was examined. For this purpose, the grazing resistance of planktonic and biofilm phases of V. vulnificus cultures was tested against different model grazers. This included Acanthamoeba castellanii which feeds on biofilms and Tetrahymena pyriformis that feeds on planktonic cells as well as attached microcolonies (Parry, 2004, Huws et al., 2005).

As part of the natural bacterioplankton community, V. vulnificus is under constant predation pressure by phagotrophic protists and the long-term persistence of different strains is dependent on how they respond to this stress. One of the main goals of this study was to identify new mechanisms that may be used by V. vulnificus in defence 78 | P a g e against protozoa and that allow for survival in the environment. To date, the predation resistance mechanisms of V. vulnificus have not been comprehensively studied. One of the factors involved in defence of V. vulnificus biotype 2 against protozoa is the plasmid- encoded MARTX type III that has been shown to be involved in the plasmolysis of amoebae (Lee et al., 2013a). RtxA13 is structurally and evolutionarily different to MARTX types I and II found in other biotypes (Kwak et al., 2011, Roig et al., 2011, Lee et al., 2013a).

Here, an environmental strain of V. vulnificus, Env1 that is resistant to grazing by T. pyriformis was identified and characterised. To identify the mechanisms that allow adaptation to the marine environment, pathogenesis and grazing resistance, the genome of V. vulnificus Env1 was analysed. In this chapter, the genome sequence of V. vulnificus Env1 was compared to available genomes, as well as to strains shown to be grazing sensitive to T. pyriformis. Furthermore, factors previously identified as potential virulence factors and grazing resistance mechanisms were investigated. One of the antiprotozoal mechanisms used by Env1 was found to be a secreted, iron-dependent, heat- stable and proteinase resistant compound.

A part of the work in this chapter has been published; Noorian Parisa, Sun Shuyang and McDougald Diane (2018). Complete Genome sequence of oyster isolate Vibrio vulnificus Env1. Genome Announcements 6 (20) (Noorian et al., 2018).

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3.2 Materials and methods

3.2.1 Strains and growth conditions

Organisms used in this study are listed in (Table 3-1). Bacterial strains were routinely grown in LB supplemented with 2% NaCl. A. castellanii was routinely maintained in PYG as described in Section 2.2.1. A. castellanii was passaged 3 d prior to harvesting for experiments and enumerated microscopically (ECLIPSE TE2000-5, Nikon, Japan) using a haemocytometer. T. pyriformis was routinely maintained in PYG as described in Section 2.2.1. Prior to experiments, 500 μl of T. pyriformis were passaged in 20 ml of -1 -1 -1 0.5 × nine salts solution (NSS) medium (8.8 g l NaCl, 0.735 g l Na2SO4, 0.04 g l -1 -1 -1 -1 NaHCO3, 0.125 g l KCl, 0.02 g l KBr, 0.935 g l MgCl2.6H2O, 0.205 g l CaCl2.2H2O, -1 -1 0.004 g l SrCl2.6H2O and 0.004 g l H3BO3) (Mårdén et al., 1985) supplemented with 1% heat-killed P. aeruginosa PAO1 (HKB) (see section 2.2.1 for production of HKB) in a 25 cm2 tissue culture flask, and further incubated at RT statically for 2 d before enumeration and use. This process is necessary to remove the nutrient media and to acclimatise the ciliate to phagotrophic feeding.

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Table 3-1. List of bacterial and protozoal strains. All the strains listed here were kindly provided by J D Oliver, UNC Charlotte, Charlotte, NC, USA.

Strain Properties Source* Reference Bacterial strains

Human blood (Kim et al., 2003b) CMCP6 C-genotype, clinical isolate, WT Korea Human blood (Poole & Oliver, 1978) C7184 C-genotype, clinical isolate, WT Atlanta Human blood (Wright et al., 1990) MO6-24 C-genotype, clinical isolate, WT California Human blood (Hor et al., 2000) YJ016 C-genotype, clinical isolate, WT Taiwan Oyster (Rosche et al., 2005) JY1701 E-genotype, environmental isolate, WT Louisiana Oyster (Rosche et al., 2005) JY1305 E-genotype, environmental isolate, WT Louisiana Oyster (Rosche et al., 2005) Env1 E-genotype, environmental isolate, WT Nk Oyster (Rosche et al., 2005) SS108-A3A E-genotype, environmental isolate, WT Nk Human Wound (Rosche et al., 2005) LSU2098 E-genotype, clinical isolate, WT Nk Human Wound (Rosche et al., 2005) LSU549 E-genotype, clinical isolate, WT Nk Human Wound (Rosche et al., 2005) LSU1657 E-genotype, clinical isolate, WT Nk Human Wound (Rosche et al., 2005) E64MW E-genotype, clinical isolate, WT Nk Protozoan strains

A. castellanii Wild type ATCC 30234 T. pyriformis Wild type ATCC 205063

* NK= Not known

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3.2.2 Assessment of resistance of planktonic bacteria to T. pyriformis predation

To assess grazing resistance of planktonic V. vulnificus, overnight cultures were adjusted so that 106 cells ml-1 in 0.5 × Väätänen nine salts solution (VNSS) (1 g bacteriological peptone, 0.5 g yeast extract, 0.5 g D-glucose, 0.01 g FeSO4·7H2O and 0.01 g Na2HPO4) in 1 litre of 0.5 × NSS (Väätänen, 1977) were added to 24-well microtitre plates (BD Falcon™, Becton Dickinson, New Jersey, USA). T. pyriformis was added to each well (104 cell ml-1; determined by inverted microscopy) and the plates were incubated at RT with shaking at 60 rpm for 24 hours. The cell density of each well was measured by spectrophotometry at OD 600 nm (Eppendorf® PlateReader AF2200, Hamburg, Germany). Planktonic fractions were collected for enumeration of CFU ml-1. Numbers and health of T. pyriformis were determined by microscopy at each sampling time.

3.2.3 Assessment of resistance of biofilms to predation by T. pyriformis and A. castellanii

To assess grazing resistance of the V. vulnificus biofilms, overnight cultures were adjusted so that 105 cells ml-1 in 0.5 × VNSS were added to 24-well microtitre plates (BD Falcon™, Becton Dickinson, New Jersey, USA). Microtitre plates were incubated at RT with shaking at 60 rpm for 24 hours. After 24 hours, the supernatants were removed and fresh 0.5 × VNSS media with or without A. castellanii or T. pyriformis was added to each well (104 cell ml-1; determined by inverted microscopy) and the microtitre plates were incubated at RT with shaking at 60 rpm for 3 d. In order to quantify the biofilm biomass, crystal violet (CV) assays were performed (O'Toole & Kolter, 1998). Briefly, all the planktonic cells were removed by was washing three times with 0.5 × NSS before adding CV (0.3%) for 15 minutes. The wells were washed a further three times with 0.5 × NSS to remove the unattached CV and then the stain was solubilised using 96% ethanol and the OD490 nm was determined by spectrophotometry (Eppendorf® PlateReader AF2200, Hamburg, Germany).

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3.2.4 Selective grazing assay

In order to test the relative grazing resistance of various strains of V. vulnificus, a selective grazing assay was performed. Overnight cultures of V. vulnificus were adjusted so that 105 cells ml-1 were resuspended in 0.2 × Marine broth 2216 (MB) (BD Difco™, Becton Dickinson, New Jersey, USA) and inoculated 1 cm from the centre of a 0.2 × Marine agar 2216 (MA) plate outward (Figure 3-1). Each strain was plated in replicates of eight on different plates. A total of 6 × 104 cells of 3-days old culture of A. castellanii in 15 μl of 50% NSS were added to the centre of each plate. Plates were incubated at RT for 10 d. The distance of the grazing front of each streak was measured.

Figure 3-1 A selective grazing assay on day 7. A. castellanii was inoculated at the centre of the agar plate (+) and streaks of different bacterial isolates were inoculated 1 cm from the centre (-) on 0.2 × MA plates. The final result of the selective grazing experiment was measured as the distance of the grazing front of each streak (red lines) on day 10 of incubation.

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3.2.5 Whole genome sequencing

High-molecular-weight genomic DNA (gDNA) was extracted from overnight cultures grown in LB broth by lysozyme digestion and use of the QIAamp® DNA Mini Kit (QIAGEN®, Hilden, Germany) according to the manufacturer’s instructions for Gram- negative bacterial cultures. Integrity of high molecular weight gDNA was evaluated by Pippin Pulse Gel and the concentration was determined by fluorometry (Qubit™ 3.0, Thermo Fisher Scientific Inc., Massachusetts, United States). Genomic DNA libraries (20 Kb) were generated by the Ramaciotti Centre for Genomics and sequenced using PacBio RS II, Continuous Long Read Sequencing- 1 SMRT cells (Pacific Biosciences, Menlo Park, CA) using previously established methods and commercially available chemistries. De novo assembly of the resulting reads was performed using the hierarchical genome assembly process (HGAP.3 Pacific Biosciences). The chromosomes were then submitted to the Rapid Annotation using Subsystem Technology server (RAST) (Aziz et al., 2008) as well as well as NCBI prokaryotic genome annotation pipeline (https://www.ncbi.nlm.nih.gov/genome/annotation_prok/) (Tatusova et al., 2016). CGView Server (http://stothard.afns.ualberta.ca/cgview_server/) was used to visualise the genome (Grant & Stothard, 2008).

3.2.6 Comparative genomics

The graphical map of Env1 genome was generated using the CGView Server (Grant & Stothard, 2008). Reference isolate Env1 was mapped against the resulting translated coding sequences with BLASTx with a percentage identity cutoff value of 70% and an expected cutoff value of 1 × 10–10 for high-scoring segment pairs >100 aa. The results were visualised by using GView (Petkau, Stuart-Edwards et al. 2010). The phylogeny tree was constructed using reference sequence alignment-based phylogeny builder (REALPHY) online tool (Bertels, Silander et al. 2014). The assembled Env1 genome from RAST was used as the reference and available genomes in NCBI database were used for comparison. The tree was rooted to Env1. The strains and NCBI reference sequences used in the genomic analysis are listed in the Appendix 2. The assembled Env1 genome from RAST was used as the reference and aligned against complete genomes available in

84 | P a g e the NCBI database using the progressive alignment algorithm implemented in MAUVE version 2.4.0. The output of this alignment was then used to check for rearrangements in each genome (Darling et al., 2010). The sequence-based comparison of Env1 to that of other available Vibrio spp. was carried on by using the SEED Viewer (Overbeek et al., 2014). The virulence factor database (VFDB) (Chen et al., 2011) that contains gene annotations related to virulence determinants in bacteria was used to identify virulence genes (either validated or predicted). The SEED Viewer was used to browse the annotated ORFs. Chromosomal regions containing conserved predicted proteins identified by these analyses were further investigated using the Compare Regions function of the SEED viewer (Overbeek et al., 2014).

3.2.7 Supernatant toxicity assay

To determine if factors secreted by V. vulnificus Env1 are toxic to protozoa, overnight cultures of V. vulnificus Env1 were adjusted to 106 cells ml-1 in 0.5 × VNSS and incubated for 24 hours. Cell-free supernatants were collected by centrifugation at 3220 × g for 5 minutes and filtered through a 0.22 µm filter (Millipore; Bedford, MA, USA). The supernatants were then tested for toxicity towards T. pyriformis. Treatments of the supernatant consisted of heat (95°C for two hours), freeze/thaw (-20°C for 24 hours), ultrafiltration (Amicon® Ultra-0.5-10, 000NMWL), exposure to proteinase K (200 μg ml-1) from Tritirachium album (Sigma-Aldrich, MO, USA), proteinase (1 mg ml-1) from Streptomyces griseus (Pronase E) (Sigma-Aldrich, MO, USA) and the iron chelator 2-2’ dipyridyl (100 μM) (Sigma-Aldrich, MO, USA). After each treatment, the supernatant was added to 24-well microtitre plates (BD Falcon™, Becton Dickinson, New Jersey, USA) containing T. pyriformis (104 cell ml-1) and numbers and health of T. pyriformis determined by microscopy. Media controls (0.5 × VNSS) from each treatment were also added to T. pyriformis to make sure none of the treatments caused toxicity. Moreover, to assess if the cell-free supernatants were toxic to A. castellanii, 3-day-old A. castellanii trophozoites were seeded in 0.5 × NSS for 24 hours before addition of cell-free supernatants. Numbers and health of A. castellanii were determined by microscopy.

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3.2.8 Biosurfactant emulsification assay

To determine if V. vulnificus Env1 produced biosurfactants that were responsible for the toxic effect, overnight cultures were adjusted to 106 cells ml-1 in 0.5 × VNSS and incubated for 24 hours. Cell-free supernatants were prepared as above (see section 3.2.7) and used to measure emulsification using n-hexadecane. Two-millilitre aliquots of cell free supernatant were mixed with 2 ml of n-hexadecane and vortexed at high speed for 2 minutes. The emulsion was observed after letting the tubes stand at room temperature for 10 minutes. The lipophilic dye Sudan black was added to the n-hexadecane to increase contrast. To calculate the emulsion index, the height of the stable emulsion layer was measured. The emulsion index was then calculated as the percentage of ratio of the height of the emulsion layer to the total height of the sample (Zavala-Moreno et al., 2014).

3.2.9 Data analysis

Statistical analyses were performed using GraphPad Prism version 7.03 for Windows, (GraphPad Software, La Jolla California USA) (www.graphpad.com). Data that did not follow Gaussian distribution as determined by the frequency distribution graphs was natural log transformed. Two-tailed student’s t-tests were used to compare means between experimental samples and controls. For experiments including multiple samples, one-way or 2-way ANOVAs were used for the analysis and Sidak's or Dunnett's multiple comparison Test provided the post-hoc comparison of means when appropriate.

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3.3 Results and discussion

3.3.1 Grazing resistance of V. vulnificus of different genotypes and from different sources

V. vulnificus resides in the aquatic environment in both planktonic and surface-attached communities. As previously described in Chapter 1, planktonic and biofilm bacteria employ different mechanisms to defend against protozoan grazers. A. castellanii is surface-attached and will exclusively feed on biofilms attached to a substratum, while T. pyriformis swims with great speed and filter feeds on particles in its path. It can efficiently feed on both attached micro-colonies and planktonic cells (Parry, 2004, Huws et al., 2005). Even though bacteria encased in the biofilm matrix are protected physically from smaller predators, grazers that can eat larger particles of food have the capacity to consume them. Consequently, in order to survive bacteria may further protect themselves by production of antiprotozoal factors (Lutz et al., 2013).

In order to determine whether isolates from clinical and environmental samples differ in resistance to predation, 12 well-characterised strains of C- and E-genotypes of V. vulnificus were exposed to predation by either T. pyriformis or A. castellanii. The strains were categorised into three groups based on their source of isolation and genotype; clinical C-genotypes (MO6-24, CMCP6, C7184, and YJ016), clinical E-genotypes (E64MW, LSU2098, LSU1657, and LSU549) and environmental E-genotypes (JY1305, JY1701, Env1, and SS108-A3A).

3.3.2 Grazing resistance of V. vulnificus biofilms

In order to assess the grazing resistance of V. vulnificus biofilms to grazing by different predators, 1-day-old biofilms were established in 24-well microtitre plates. The biofilms were then exposed to grazing by either T. pyriformis or A. castellanii for 3 d at RT and the biofilm biomass was determined by CV staining. The health of the grazers was monitored by inverted light phase microscopy.

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Firstly, to evaluate the grazing resistance of each strain, the grazed biomass was compared to ungrazed controls (Figure 3-2 A and 3-3 A). V. vulnificus biofilms are sensitive to grazing by the filter feeder T. pyriformis, as evidenced by the reduction in the biomass of almost all strains after grazing compared to the ungrazed strains. The biofilm biomass of the majority of the strains was reduced by 75%. The percentage change in biofilm biomass was calculated by removing the biomass of ungrazed samples from the grazed samples and divided by the ungrazed biofilm biomass (Figure 3-2 A). The only strain that did not show a significant reduction in biomass after grazing was the environmental strain, Env1. However, this strain does not form robust biofilms under the conditions used here, as evidenced by the low amount of CV stain taken up (OD490 = 0.04) by the ungrazed control (Figure 3-2 A). Upon exposure to Env1, T. pyriformis stopped swimming and feeding and sank to the bottom of the well, behaviour that is consistent with introduction to an undesirable environment. After 3 d, T. pyriformis was not detected in the wells containing the Env1 biofilm suggesting that this strain might be toxic towards T. pyriformis. V. vulnificus Env1 interactions with T. pyriformis will be further described in the next sections.

In contrast to grazing by T. pyriformis, all the biofilms were protected against grazing by A. castellanii as evidenced by the increase in the biofilm biomass after grazing in comparison to the ungrazed controls (Figure 3-3 A). On average, the biofilm biomass of most strains increased by 50% after grazing. The relationship between source of isolation and genotype of the isolates with grazing resistance was tested by comparing the percentage change in biofilm biomass of all the strains in each category (Figure 3-2 B and Figure 3-3 B). There was no significant difference in grazing resistance of the V. vulnificus biofilms of clinical C-genotype, clinical E-genotype and environmental E- genotype strains to grazing by T. pyriformis or A. castellanii, (p-value= 0.051 and 0.056 respectively.

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In order to further investigate the grazing resistance of biofilms to A. castellanii predation, a plate-based selective grazing assay was also employed. In this assay, the susceptibility of each strain was determined by the reduction of pre-established late-stage biofilms (streaks) on low nutrient (0.2 × MA) agar plates (Figure 3-4). There was no significant difference in grazing resistance of the V. vulnificus clinical C-genotype, clinical E- genotype and environmental E-genotype isolates to grazing by A. castellanii (p-value= 0.249).

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Figure 3-4 A. castellanii selective grazing assay with clinical C-genotype (MO6-24, CMCP6, C7184, YJ016), clinical E-genotype (E64MW, LSU2098, LSU1657, LSU549) and environmental E-genotype (JY1305, JY1701, Env1, SS108-A3A) V. vulnificus isolates. The amount of biofilm grazed (A) and mean value of each group (B) was determined. The plates were incubated for 10 d at RT. The distances from the starting position to grazing front reflected the relative grazing resistance of each strain. Error bars represent the standard error of the mean for four strains with four replicates each. The experiment was run in replicates of four and repeated three times. Statistical analysis was performed using 1-way ANOVA which revealed no significant differences.

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3.3.3 Grazing resistance of planktonic cells of V. vulnificus

The resistance of planktonic cells of V. vulnificus isolates to predation by T. pyriformis was investigated by determining the numbers of planktonic cells after exposure to T. pyriformis. Both C- and E- genotype strains exposed to predation by T. pyriformis were efficiently grazed, showing a significant decrease in CFUs over time, with the exception of one environmental E- genotype strain, Env1, which was resistant to elimination by the filter-feeding ciliate (Figure 3-5 A).

The total numbers of T. pyriformis increased significantly when feeding on the grazing sensitive planktonic cells when compared to the media control. The increase in T. pyriformis numbers was correlated with the decrease in the biomass of the 11 grazing sensitive strains. In contrast, the total number of T. pyriformis cells was not significantly different to the initial inoculum and media control when exposed to the grazing resistant strain, Env1 (Figure 3-5 B). There was no significant difference in grazing resistance of the clinical C-genotype, clinical E-genotype or environmental E-genotype V. vulnificus planktonic cells to grazing by T. pyriformis (p-value= 0.474).

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P p p p ty ty ty o o o n n n e e e -g -g -g C E E l l l a a ta ic ic n n n e i li l m C C n o ir v n E Figure 3-5 Grazing resistance of clinical C-genotype (MO6-24, CMCP6, C7184, YJ016), clinical E-genotype (E64MW, LSU2098, LSU1657, LSU549) and environmental E- genotype (JY1305, JY1701, Env1, SS108-A3A) V. vulnificus isolates exposed to T. pyriformis for 24 hours. The biomass of planktonic V. vulnificus cells in the supernatant

94 | P a g e was determined by spectrophotometry (OD 600 nm) of grazed and ungrazed strains (A), and total numbers of T. pyriformis was enumerated by inverted microscopy (B) and percentage change in biomass of planktonic V. vulnificus cells in each group +/- SD was determined (C). The experiment was run in 3 replicates and repeated 3 times separately. Error bars represent standard deviation. Statistical analysis was performed using 2-way ANOVA and Sidak's multiple comparisons test (A) and 1-way ANOVA and Dunnett's multiple comparisons test comparing all strains with the VNSS media control (B) where statistical significance is indicated by ****, p < 0.0001 and Ns, P > 0.05. One-way ANOVA and Dunnett's multiple comparisons tests revealed no significant differences (C).

The findings of this study demonstrate that genotype and place of isolation does not affect the grazing resistance of V. vulnificus under the experimental conditions used here. V. vulnificus biofilms are protected against grazing by the amoeba, A. castellanii and biofilm biomass increases after exposure to the predator. In Chapter 2, transcripts related to metabolism and energy production in V. cholerae increased after exposure to predation, and this might be due to increases in available nutrients caused by ‘sloppy feeding’ or excretion of waste products by the protozoa (Wang et al., 2009). Here, an increase in nutrient levels in the grazed samples may have contributed to growth in comparison to ungrazed cultures where there was no feeding. The speed of growth of the bacteria was much faster than the consumption rate by A. castellanii.

In contrast, both planktonic and biofilm cells of V. vulnificus were sensitive to grazing by the ciliate, T. pyriformis, with the exception of the environmental strain, Env1. Many bacteria have been shown to express antiprotozoal factors that can affect the health of the protozoa. The focus of the rest of this study is to investigate the antiprotozoal mechanisms that Env1 potentially uses in order to defend itself against predation.

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3.3.4 V. vulnificus Env1

V. vulnificus Env1 is an environmental strain isolated from an oyster in Louisiana, United States of America (Rosche et al., 2005) and was previously designated as SS109B-3B2 (Warner & Oliver, 2007). In the previous section, it was shown that Env1 was resistant to grazing by the filter-feeding ciliate, T. pyriformis. T. pyriformis is unable to feed efficiently on Env1 and exhibits sign of being exposed to unfavourable conditions such an antiprotozoal toxin. Here, the characteristics of Env1 to understand what factors are involved in efficient defence were investigated.

In a previous study of resistance to human serum of C- and E genotype isolates, four of the five E-genotype strains lacked resistance but Env1 was strongly resistant and exhibited serum survival comparable to that of C-type strains. Interestingly, Env1 was one of only two E-type strains tested that possessed the viuB gene, a siderophore-encoding gene, which influenced serum survival of all isolates of V. vulnificus tested (Bogard & Oliver, 2007). V. vulnificus Env1 also consistently expressed capsular genes, wza and wzb, during both in vitro and in situ studies at all time points, even when the cells were in the viable but nonculturable (VBNC) state and following resuscitation. In contrast, the C-genotype isolates down-regulated capsule genes during in situ incubation and in the VBNC state (Smith & Oliver, 2006).

The mechanisms that facilitate resistance to predation of planktonic V. vulnificus are not fully understood; therefore, the genome of this isolate was investigated to gain insight into the genes that potentially facilitate this resistance. Here, the fully sequenced genome of V. vulnificus Env1 is reported and its potential antiprotozoal factors are analysed. The genome of Env1 is used for comparison with genomes of other sequenced strains that were used in the grazing resistance experiments.

3.3.5 Features of the Env1 genome

The complete genome sequence of V. vulnificus Env1 was generated using the PacBio RS platform with single molecule, real-time (SMRT) sequencing with 140× coverage. The genome consists of 4,954,048 bp with an average guanine-plus-cytosine (G+C) content

96 | P a g e of 46.7%; and like other known Vibrio spp. contains two circular chromosomes of 3,241,343 and 1,712,705 bp (Figure 3-6). There was no evidence of a plasmid in this isolate. The genome sequence of V. vulnificus Env1 has been deposited in the GenBank database under the accession numbers CP017635 for chromosome I and CP017636 for chromosome II.

The sequences were submitted to the RAST (Aziz et al., 2008) as well as NCBI prokaryotic genome annotation pipeline (Tatusova et al., 2016) and results show that there are a total of 4,378 coding sequences (CDSs) and 157 RNAs predicted. Approximately 28.4% of the CDSs were annotated as hypothetical proteins, including proteins that are conserved in other bacteria. The majority of the CDSs were related to functions in subsystems including amino acids and derivatives (515 genes), carbohydrates (456), protein metabolism (310) and the synthesis of cofactors, vitamins, prosthetic groups, and pigments (303 genes). There were 9 genes related to prophage, 52 genes to iron acquisition and metabolism, 164 genes related to stress responses and 92 to virulence, disease and defence as determined by the SEED viewer (an annotation environment that detects and compares genes between functional subsystems from groups of genomes) (Overbeek et al., 2014).

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Figure 3-6 The graphical map of Env1 genome was visualised using CGView Server, chromosome I (A) and chromosome II (B) (Grant & Stothard, 2008). The two chromosomes are depicted by circular representation. From the outside to the centre: CDS on the forward strand (one ring for each reading frame), CDS on the reverse strand, GC content, and GC skew.

3.3.6 Comparative genomics

To date, the genome sequences of 83 other V. vulnificus strains have been deposited in the GenBank database, 12 of which are complete genomes. A phylogenetic tree was 99 | P a g e constructed by mapping the available whole genome sequences as well as the partial genomes of the strains used in these grazing resistance experiments to the whole genome of Env1 (Figure 3-7). The tree was constructed by the REALPHY server which merges alignments from mappings to multiple full reference sequences, in order to include all the single nucleotide polymorphisms (SNPs) and nonpolymorphic sites in an alignment (Bertels et al., 2014).

The phylogenetic analysis performed here demonstrates the relatedness of the core V. vulnificus genome and further supports the accuracy of E- and C- genotyping. Branching suggests a fundamental divergence between the two genotypes, which correlates with the divergent lifestyle preferences of E- and C- genotypes of V. vulnificus. This is consistent with the finding of Morrison et al. (2012) who used a random sampling of 175 single copy genes in Vibrio spp. and found the of E- and C- genotypes cluster together in the phylogenetic tree. Env1 grouped with other E-genotype strains used in the grazing assays and hence the phylogenetic tree establishes that Env1 is related to other E-genotypes.

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Figure 3-7 Phylogenetic tree of sequenced V. vulnificus genomes. REALPHY was used to construct the tree using full sequenced genome alignment and combination of data from mapping to multiple references (Bertels et al., 2014). Red boxes indicate strains classified as C- genotypes and green boxes indicate strains classified as E- genotype. The ratio of mapped sites to the polymorphisms detected were reported in the black boxes. The genome sequence of V. vulnificus Env1 was used as the core genome.

The genes found in all investigated genomes (core genome) are responsible for the phenotypic similarities of the genus or species, while unique features are attributed to particular genes. For example, virulence factors found in the pathogenic strains of an environmental species occur only in some strains (Kahlke et al., 2012). To determine the genetic features that might play a role in a strain’s unique phenotypic behaviour, differential genome comparison was employed. Ideally such a comparison would be done

101 | P a g e between all represented strains but the lack of completed genomes limited the ability to perform a comprehensive analysis (Ehrlich et al., 2008).

GView was used to map the pangenome, which depicts the BLAST Atlas of Env1 against three grazing sensitive strains. Each Env1 chromosome was used as the seed genome to construct the core genome (Petkau et al., 2010). There were 3017 homologous genes identified as being present in all strains with 70% identity. In Vibrio spp., chromosome I carries most of genes necessary for essential cell functions, while most of the hypothetical genes are located in the smaller chromosome II (Heidelberg et al., 2000). Similarly, the Env1 chromosome I is more similar to the other tested strains than chromosome II which may have been the target for many HGT events (Figure 3-8).

It is important to note the genome size difference between compared strains. The Env1 total genome is 4.9 Mbp of which chromosome I accounts for 3.24 Mbp of the sequences and chromosome II for 1.71 Mbp (Figure 3-6). The grazing sensitive clinical strains, Yj016, CMCP6 and MO6-24/O have genomes of 5.26, 5.12 and 5.00 Mbp, respectively. Much of this difference comes from the size of chromosome II. The size of chromosome II in Yj016, CMCP6 and MO6-24/O is 1.85, 1.84 and 1.81 Mbp, respectively. This difference in size can be visualised by blast comparison in Figure 3-8 where a large gap in chromosome II of Env1 (dark green circle) corresponds to genes missing in Env1 compared to the other strains.

An in-depth comparison between the grazing resistant Env1 and other strains used in the grazing experiments revealed that Env1 harbors many virulence genes, including some not previously reported in V. vulnificus. In a sequence-based comparison, 387 genes were annotated as hypothetical that had identity matches of less than 30% with CMCP6 and YJ016. In addition, 22 ORFs were annotated as mobile elements with the same criteria.

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Figure 3-8 Circular maps that compare the genomes of three grazing sensitive V. vulnificus strains, MO6-24, CMCP6 and YJ016 with V. vulnificus Env1. Reference isolate Env1 was mapped against the CDSs using pangenome analysis with a percentage identity cutoff value of 70% and an expected cutoff value of 1 × 10–10 for high-scoring segment pairs >100 aa. The results were visualised by using GView (Petkau et al., 2010).

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To further investigate genomic differences between the sequenced V. vulnificus strains, Progressive Mauve alignment was used (Darling et al., 2010). Three strains, CMCP6, MO6-24/O and YJ016 were chosen for the comparison. Chromosomes I and II are illustrated separately here for better resolution (Figure 3-9). Appendix 3 shows the genomic comparison between V. vulnificus Env1, CMCP6, MO6-24/O, YJ016, JY1305, JY1701 and E64MW. None of these strains exhibited resistance to grazing by T. pyriformis. Although the sequences for JY1305, JY1701 and E64MW are available in the NCBI database, they are scaffolds and not closed genomes (Appendix 2). The only strain in this analysis that has been shown to carry a plasmid is YJ016. There is no evidence of plasmid sequences in the genome of Env1 or the other E-genotype strains.

Genome arrangement has profound effects on gene expression and phenotypic traits. Chromosomal rearrangements and inversions not only contribute to mutations in the genome, but are also involved in differential regulation of many genes (Mahan & Roth, 1991, Stover et al., 2000). The genomes of Vibrio spp. show multiple events of intra- and inter-chromosomal rearrangements, with the overall gene content and arrangement of the large chromosomes better conserved than the small chromosome that is divergent in size and gene content (Chen et al., 2003). Considerable genomic rearrangement in chromosome II accounts for evolution in the genus Vibrio, for example when Vibrio mimicus and V. cholerae diverged (Hasan et al., 2010). This is consistent with the overall pangenome analysis of each chromosome where chromosome II shows high divergence between Env1 and the clinical strains (Figure 3-8).

Locally collinear blocks (LCB) were defined by outlining a region of the genome sequence that aligned to part of other genomes and is considered to be homologous and internally free from genomic rearrangement (Figure 3-9). Each of these blocks has been designated a different colour (Darling et al., 2010). The analysis here shows high divergence and rearrangement mostly in chromosome I compared to other clinical and environmental strains. The rearrangements are shown by LCB connecting lines (Figure 3-9A). Such genome rearrangements can play important roles in the evolution of Vibrio spp. and the difference in the block interchanges can be used to predict evolutionary paths between species. For example, block-interchange distance between V. vulnificus and

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Vibrio parahaemolyticus is smaller than that between V. vulnificus and V. cholerae or V. parahaemolyticus and V. cholerae (Lin et al., 2005).

Many inversions are also observed in chromosome I of CMCP6, MO6-24/O and YJ016, in comparison to Env1 genome. These inversions are shown as LCBs below the genome centre line. An example of inversion in MO6-24/O compared to Env1 is shown with an arrow in Figure 3-9. However, in a previous study when the reference genome of V. vulnificus CMCP6 and those of JY1305, E64MW, and JY1701 were aligned using MAUVE, only one inversion in the E64MW was observed (Morrison et al., 2012). An example of the inversion has been made apparent in Figure 3-9 by use of an arrow, where inversions are depicted as blocks below a genome’s centre line.

Chromosome I and II both contain several gaps which are due to deletions or missing genes in Env1. The second and biggest alignment gap in chromosome I of Env1 has been made apparent in Figure 3-9A by use of an arrow. This gap also exists in other strains and in Env1 consists of 200000 bp. The gap starts with the integron integrase, IntI4, a hallmark of super-integrons that is also present in all other strains analysed here. This large gap contains many mobile elements and hypothetical genes unique to the Env1 genome. The following section focuses on the unique factors of Env1 that might contribute to grazing resistance.

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Figure 3-9 Whole-genome alignments of V. vulnificus Env1, CMCP6, MO6-24/O and YJ016, chromosome I (A) and chromosome II (B) were performed using the Progressive Mauve alignment (Darling et al., 2010). LCBs are represented by blocks of different colours. The LCB connecting lines depict rearrangements. The degree of similarity is indicated using white areas where the coloured areas have higher similarity and the areas that are completely white are not aligned and probably contain sequence elements specific to a particular genome. Inverted regions in the genome are clearly depicted as blocks below a genome’s centre line. The names of the strains are listed at the bottom of the blocks.

A recent publication by Roig et al. (2018) announced submission of additional genome sequences in the NCBI database that became available in January of 2018 just before this thesis was submitted and hence are not included here. The sequences of several grazing sensitive strains (SS108-A3A, LSU1657 and LSU2098) are also now available, which will enable further analysis for future work.

3.3.7 Env1 unique genes encoding virulence and defence factors

The unique genes (e.g. homologs of virulence-related genes) present in Env1 are initial targets for identification of the antiprotozoal factors that affect T. pyriformis. Although, as discussed in the previous section, rearrangements and inversions may also alter the expression of genes that are common between all the strains. Furthermore, even though a cutoff point for similarity comparisons was selected, even a single base pair change can alter the functionality of a gene.

The position of selected genes discussed in this section and the sequence identity of Env1 compared to V. cholerae N16961, V. vulnificus CMCP6, and YJ016 were compared using the RAST SEED viewer sequenced-based comparison tool (Figure 3-10). The position and length of each gene is summarised in Table 3-2.

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Figure 3-10 Genome of V. vulnificus Env1 was compared to V. cholerae N16961, V. vulnificus CMCP6, and V. vulnificus YJ016, from the outer ring to the inner ring respectively. Horizontal lines are drawn to separate chromosomes. Arrows indicates area of unique genes or genes of interest. Percent of gene sequence identity is shown by color coding provided in the legend. RAST SEED viewer sequenced-based comparison tool was used (Overbeek et al., 2014).

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A putative internalin adjacent to a putative rearrangement hotspot (Rhs)-related gene in the Env1 chromosome I was identified that does not exist in the grazing sensitive strains. Blast analysis revealed that one other strain of V. vulnificus, FORC-036, carries these genes and was isolated from a surf clam in South Korea (Han Young et al., 2017). Both internalin and Rhs-related proteins are known to be involved in virulence in many species of bacteria. For example, in Listeria monocytogenes the surface proteins, internalins InlA and InlB, interact with a host receptor, E-cadherin, to enter epithelial cells (Gaillard et al., 1991, Lecuit et al., 1997, Ferraris et al., 2010). Rhs proteins were first identified as sites that promote recombination in E. coli. They are widely distributed bacterial exotoxins that carry polymorphic C-terminal toxin domains that are horizontally acquired. Some Rhs genes encode surface exposed or secreted proteins such as rhsT in Pseudomonas aeruginosa that encodes an exposed toxic protein that is induced upon contact with phagocytic cells. These proteins translocate into phagocytic cells causing inflammasome- mediated death (Kung et al., 2012). There is potential for these known virulence determinants to be involved in intracellular survival of V. vulnificus in eukaryotic hosts.

Furthermore, an ankryin gene encoded on chromosome II is adjacent to an Rhs-related gene, a transposase and a VgrG gene belonging to the T6SS. Ankyrin repeat proteins have a series of 33 amino acid tandem repeats in the protein binding domain. Mutants of ankH or ankJ of L. pneumophila have defects in intracellular replication in phagosomes similar to dot/icm mutants. These L. pneumophila eukaryotic-like ankyrin proteins play a role in intracellular replication within amoeba, ciliated protozoa and human macrophages (Habyarimana et al., 2008). Ankyrin-repeat proteins in E. coli can interfere with amoebal phagocytosis and phagosome acidification leading to accumulation of bacteria in the phagosome (Nguyen et al., 2014).

The second largest gap belongs to a type I secretion system (T1SS) secreted agglutinin, RTX, on chromosome I from 996788 to 973842bp (Figure 3-9 and Figure 3-10). RTX proteins are virulence factors that are directly involved in pathological damage to the host, as described in Chapter 1. The RTX protein, exported by the T1SS, can cause damage in a variety of ways (Kim et al., 2008, Lee et al., 2008a, Chung et al., 2010, Lo et al., 2011, Kuo et al., 2015). The MARTX type III toxin of V. vulnificus biotype 2 is as an antiprotozoal factor (Lee et al., 2013a). However, V. vulnificus produces at least four

110 | P a g e different types of MARTX (types I–IV) (Kwak et al., 2011, Roig et al., 2011) and three CDSs annotated as RTX toxins, two of which are located on chromosome II (15612 and 13968 bp long) were found. These two RTX toxins have homologues across most of the other V. vulnificus strains. The putative type I secretion system (T1SS) secreted agglutinin, RTX, located on chromosome I was annotated by RAST as being 22947 bp long and encodes a large 7649 aa protein. The only other strain carrying this type I secretion system (T1SS) secreted agglutinin, RTX, is V. vulnificus 93U204 that was isolated from a moribund tilapia in Kaohsiung, Taiwan (Lo et al., 2014). RTX proteins are involved in toxicity against many types of cells and the RTX protein detected here in Env1 may be a potential antiprotozoal factor. This possibility will be further examined in the next chapter.

Copper is one of the essential metals in nature that organisms need for proper functioning of enzymes, but excess copper is toxic (Grass et al., 2011). Many mechanisms have been described that provide copper tolerance to bacteria inside host cells (White et al., 2009, Ladomersky & Petris, 2015). Ubiquitous copper exporting systems such as the P1B-type ATPases, including CopA in E. coli, P. aeruginosa and Salmonella typhimurium have been shown to protect bacteria in host phagosomes (Rensing & Grass, 2003, Bondarczuk & Piotrowska-Seget, 2013, Ladomersky & Petris, 2015), including in protozoa (Hao et al., 2016). Env1 encodes for a suppressor of copper-sensitivity (ScsD) membrane protein that is lacking in the other V. vulnificus isolates. Interestingly, the rest of the operon (scsA, scsB and scsC) was absent. The suppressor for copper sensitivity (scs) locus in Vibrio spp. was shown not to affect copper tolerance in a previous study, as the scs genes were deleted and rearranged in the genomes (Busch, 2012).

Many sequences encoding putative mobile elements are located throughout the Env1 genome and these elements carry many hypothetical genes. Genes encoding a retron- type RNA-directed DNA polymerase and prophage Lp2 protein 6 are examples of phage sequences found in the Env1 genome. Bacterial retron reverse transcriptases are unusual enzymes that are present in some bacteria, including V. cholerae, which utilise RNA molecules as a template (Matiasovicova et al., 2003). They are hypothesised to be involved in mutagenesis or regulation of gene expression by an antisense mechanism (Mao et al., 1995, Maas et al., 1996). Bacteria, especially vibrios have a range of

111 | P a g e processes for acquisition of exogenous genetic elements. Transfer of these elements can be through integrons, transposons, gene cassettes and plasmids or alternatively by conjugation between bacteria or transduction via phages. These events can occur in the host and in environmental niches.

Table 3-2 Position and length of genes potentially involved in virulence, disease and defence in V. vulnificus Env1.

NCBI RAST gene Length Function Location Start Stop gene ID Id (bp) Internalin, BJD94_0 fig|1246305. putative 6450 4.peg.1353 ChrI 1442310 1449341 7032

Rhs-related BJD94_0 fig|1246305. protein, putative 6455 4.peg.1354 ChrI 1449352 1452945 3594

BJD94_1 fig|1246305. Ankyrin 5070 4.peg.3014 ChrII 53750 52920 831

T1SS secreted BJD94_0 fig|1246305. agglutinin RTX 4465 4.peg.953 ChrI 996788 973842 22947

Membrane protein, BJD94_1 fig|1246305. suppressor for 5820 4.peg.3165 ChrII 238230 238721 492 copper-sensitivity ScsD

Prophage Lp2 BJD94_1 fig|1246305. protein 6 9405 4.peg.3909 ChrII 1109110 1108031 1080

Retron-type BJD94_0 fig|1246305. RNA-directed 3695 4.peg.796 ChrI 798030 798030 954 DNA polymerase

3.3.8 Genes encoding iron uptake and utilisation proteins

All organisms, with a few exceptions, need to acquire and utilise iron in order to grow (Chu et al., 2010). V. vulnificus is a ferrophilic bacterium and the role of iron in pathogenesis has been established (Wright et al., 1981). The pathogenicity of V.

112 | P a g e vulnificus is dependent on the uptake and transport of iron in the iron-deplete host or aquatic environment (Ratledge & Dover, 2000). Therefore, it is not surprising that there are many systems involved in iron uptake and transport in V. vulnificus. The role of most of these systems in growth and virulence has been well established. Therefore, it was hypothesised here that iron may be involved in defence against protozoan predation. Proteins involved in Env1 iron uptake and utilisation are listed in Table 3-3.

V. vulnificus has three TonB systems for transport of both hydroxamate-type siderophores and vulnibactin, as well as haeme and haemoglobin. V. vulnificus TonB1 and TonB2 systems are induced under iron-limiting conditions whereas TonB3 transcription does not change in response to iron concentration (Alice et al., 2008, Kustusch et al., 2012). The Env1 genome carries three ferric siderophore TonB transport proteins, one located on chromosome I and the other two on chromosome II. There are six TonB-dependent outer membrane receptors, including HutA, HutR and a zinc-regulated TonB-dependent outer membrane receptor. Two TonB receptors located on chromosome II are unique to Env1 and do not have homologues in the other V. vulnificus strains. It has been demonstrated that V. vulnificus CMCP6 has three cooperating TonB systems for iron assimilation, although it has been suggested that these systems might not be required for in vivo growth but are required for virulence (Duong-Nu et al., 2016).

The siderophore, aerobactin, is responsible for growth of V. vulnificus under iron-limiting conditions and is comprised of a three-gene operon (vatCDB) which encodes an ATP- binding cassette transport component, iutA encoding a ferric aerobactin receptor and iutR which encodes a transcriptional repressor (Tanabe et al., 2005). Env1 possesses all these genes on chromosome II. Another system V. vulnificus uses to take up ferric iron is the siderophore, vibriobactin, and the vuuA gene encoding a vibriobactin receptor (Webster & Litwin, 2000). The ferric vibriobactin, enterobactin transport system is also located on the chromosome II. Vulnibactin is another siderophore present in V. vulnificus (Okujo et al., 1994). In Env1, vulnibactin utilisation proteins VuuA and VuuB are located on chromosome II and the catechol siderophore ABC transporter, substrate-binding protein is located between these two genes.

The ferric iron ABC transporter (Fbp) system for ferric iron uptake is a periplasmic binding protein-dependent ABC transporter that is used to transport the siderophore and 113 | P a g e haeme through the inner membrane (Mey et al., 2008). Two complete ferric iron ABC transporter systems exist in Env1, one on each chromosome. The one located on chromosome I has no known homologues in the NCBI database. The ABC transporter ATP-binding protein is located adjacent to a mobile element, which means it may be a recent acquisition. V. vulnificus Env1 has systems for transport of ferrous iron as well as the genes encoding ferrous iron transport proteins A, B and C are located on chromosome I.

All of these iron acquisition and utilisation systems have high affinity for iron and respond to low levels of iron available in the host and environmental niches. Most bacteria use iron as an essential factor involved in an array of metabolic and signalling pathways where hundreds of enzymes involved in primary and secondary metabolites need iron as a co-factor. Pathogens need iron in order to multiply in a host (Miethke & Marahiel, 2007). Furthermore, several transcriptional regulators, such as ferric uptake regulator (Fur) (Escolar et al., 1999), use iron as a regulatory signal, which in turn regulates many internal pathways. Iron could be a factor involved in several mechanisms that contribute to grazing resistance. It is possible that iron is involved in production of secondary metabolites or involved in the regulatory pathways in coordination of the antiprotozoal defences of Env1.

Table 3-3 Proteins for iron uptake and utilisation.

NCBI RAST gene Length Function gene Location Start Stop Id (bp) ID

Category Ferric siderophore transport system, BJD94 fig|1246305. ChrI 2276229 2276861 633 periplasmic binding _10340 4.peg.2131 protein TonB

Ferric siderophore transport system, BJD94 fig|1246305. ChrII 1242592 1242218 375 periplasmic binding _19930 4.peg.4015 protein TonB

TonB System TonB

BJD94 fig|1246305. ChrII 1681193 1680555 639 Ferric siderophore _21765 4.peg.4411 transport system,

114 | P a g e periplasmic binding protein TonB

TonB-dependent heme and hemoglobin BJD94 fig|1246305. ChrII 1624873 1622735 2139 receptor HutA, _21510 4.peg.4358 ferrichrome receptor

TonB-dependent BJD94 fig|1246305. heme receptor ChrII 1173215 1175350 2136 _19625 4.peg.3954 HutR

TonB-dependent BJD94 fig|1246305. ChrII 820339 818255 2085 receptor _18250 4.peg.3676

TonB-dependent receptor; Outer membrane receptor BJD94 fig|1246305. ChrI 2270398 2272986 2589 for _10315 4.peg.2126 ferrienterochelin and colicins

TonB-dependent receptor; Outer membrane receptor BJD94 fig|1246305. ChrII 1074941 1077007 2067 for _19245 4.peg.3876 ferrienterochelin and colicins

Zinc-regulated TonB-dependent ChrI 1766147 1765008 1140 outer membrane BJD94 fig|1246305. receptor _08000 4.peg.1651

Biopolymer BJD94 fig|1246305. transport protein ChrI 2275817 2276224 408 _10335 4.peg.2130 ExbD/TolR

Biopolymer BJD94 fig|1246305. transport protein ChrII 1681597 1681193 405 _21770 4.peg.4412 ExbD/TolR

MotA/TolQ/ExbB BJD94 fig|1246305. proton channel ChrI 2273837 2275198 1362 _10325 4.peg.2128 family protein

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MotA/TolQ/ExbB BJD94 fig|1246305. proton channel ChrI 331621 332304 684 _01510 4.peg.310 family protein

MotA/TolQ/ExbB BJD94 fig|1246305. proton channel ChrII 1683524 1682157 1368 _21780 4.peg.4414 family protein

Tol biopolymer BJD94 fig|1246305. transport system, ChrI 332304 332744 441 _01515 4.peg.311 TolR protein

BJD94 fig|1246305. TolA protein ChrI 332759 333826 1068 _01520 4.peg.312

tolB protein precursor, periplasmic protein BJD94 fig|1246305. involved in the ChrI 333839 335188 1350 _01525 4.peg.313 tonB-independent uptake of group A colicins

Ferric aerobactin ABC transporter, BJD94 fig|1246305. ChrII 626936 627703 768 ATPase component _17460 4.peg.3508 vatC

Ferric aerobactin ABC transporter, BJD94 fig|1246305. periplasmic ChrII 627721 628617 897 _17465 4.peg.3509 substrate binding protein vatB

Ferric aerobactin ABC transporter, BJD94 fig|1246305. ChrII 628614 630611 1998 permease _17470 4.peg.3510

Aerobactin component vatD

Aerobactin siderophore BJD94 fig|1246305. ChrII 632074 634230 2157 receptor iutA _17480 4.peg.3513

Transcriptional repressor of ChrII 630771 631733 963 aerobactin receptor BJD94 fig|1246305. iutR _17475 4.peg.3511

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Ferric vibriobactin, enterobactin BJD94 fig|1246305. transport system, ChrII 1428275 1429030 756 _20760 4.peg.4193 ATP-binding protein

Ferric vibriobactin, enterobactin BJD94 fig|1246305. transport system, ChrII 1426389 1427324 936 _20750 4.peg.4191 permease protein

vctD

Ferric vibriobactin, enterobactin BJD94 fig|1246305. transport system, ChrII 1427317 1428267 951 _20755 4.peg.4192 permease protein vctG

Catechol Vibriobactin/enterobactin siderophore ABC BJD94 fig|1246305. transporter, ChrII 435297 436205 909 _16670 4.peg.3340 substrate-binding protein

Vulnibactin BJD94 fig|1246305. utilisation protein ChrII 438348 436291 2058 _16675 4.peg.3341 VuuA

Vulnibactin BJD94 fig|1246305. utilisation protein ChrII 905875 905060 816 _16645 4.peg.3335 VuuB

Isochorismate pyruvate-lyase of siderophore BJD94 fig|1246305. ChrII 907258 906929 330 biosynthesi, _16655 4.peg.3337 Vulnibactin Vulnibactin- specific

Isochorismate synthase of BJD94 fig|1246305. ChrII 427773 428954 1182 siderophore _16635 4.peg.3332 biosynthesis

Ferric iron ABC BJD94 fig|1246305. transporter, ATP- ChrI 3061765 3062793 1029 _14145 4.peg.2818 binding protein

ABC

Ferric iron iron Ferric

transporter

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Ferric iron ABC BJD94 fig|1246305. transporter, ATP- ChrII 1297591 1298697 1107 _20195 4.peg.4071 binding protein

Ferric iron ABC BJD94 fig|1246305. transporter, iron- ChrI 3059027 3060040 1014 _14135 4.peg.2816 binding protein

Ferric iron ABC BJD94 fig|1246305. transporter, iron- ChrII 1295896 1296906 1011 _20190 4.peg.4069 binding protein

Ferric iron ABC BJD94 fig|1246305. transporter, ChrI 3060140 3061765 1626 _14140 4.peg.2817 permease protein

Ferric iron ABC BJD94 fig|1246305. transporter, ChrII 1298727 1300430 1704 _20200 4.peg.4072 permease protein

Ferrous iron BJD94 fig|1246305. ChrI 1549737 1549594 144 transport protein A _06925 4.peg.1444

Ferrous iron BJD94 fig|1246305. ChrI 1549594 1547318 2277 transport protein B _06920 4.peg.1443

Ferrous iron BJD94 fig|1246305. ChrI 1547321 1547091 23 Ferrous iron transport iron Ferrous transport protein C _06915 4.peg.1442

3.3.9 Supernatant toxicity assay

In order to further investigate the grazing resistance of V. vulnificus Env1, the presence of a secreted toxin was investigated. Bacteria may produce several toxins or antiprotozoal compounds. A rapid bioassay, BACTOX has been developed that detects the overall toxicity of bacterial strains that synthesise toxic secondary metabolites (Schlimme et al., 1999). The test uses T. pyriformis and bacterial suspensions, and the health of the ciliate is observed soon after co-incubation (Schlimme et al., 1999).

In order to determine if toxins are secreted by V. vulnificus Env1, a cell-free supernatant was obtained added to healthy T. pyriformis. V. vulnificus was grown in 0.5 × VNSS for 24 hours before the supernatant was filtered and T. pyriformis was added. After 10 minutes, T. pyriformis stopped swimming and sank to the bottom of the microtitre plates

118 | P a g e and the cytoplasm began leaking from cells. One hour after incubation at RT, 100% of the T. pyriformis were dead and after 24 hours, cells were completely degraded (Figure 3-11).

Figure 3-11 T. pyriformis after co-incubation with V. vulnificus Env1 cell-free supernatant. At 0 hours, T. pyriformis are healthy and swimming. After 1 hour, all the cells have stopped swimming and sunk to the bottom of the well. After two hours, the dead cells start lysing. Scale bar = 100 μm.

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Cell-free supernatants from Env1 are not, however, toxic to A. castellanii. A. castellanii trophozoites remain healthy even after 24 hours of incubation in Env1 cell-free supernatant (Figure 3-12). The cell membrane of T. pyriformis is different from A. castellanii, therefore, the amoeba is not sensitive to the same toxic factor or at the same dose as T. pyriformis. The grazing resistance of biofilms of V. vulnificus to A. castellanii may therefore be due to other protective mechanisms (Figure 3-3).

M e d ia c o n tro l

E n v 1 c e ll-fre e )

1 s u p e rn a ta n t

l m

1 5 0 0 0 * * * * N S

(

s l

l 1 0 0 0 0

l a

e 5 0 0 0

r e

b 0 m

u s ii i n

N m a r ll fo te ri s y a p c . . T A

Figure 3-12 Effect of cell-free supernatants from V. vulnificus Env1 on the health of T. pyriformis and A. castellanii. The cell-free supernatants were added to protozoan cultures, and the numbers and health of the protists determined by microscopy after 24 hours. Error bars represent the standard deviation of three replicates. Statistical analysis was performed using 2-way ANOVA and Sidak's multiple comparisons test. Statistical significance is indicated by (****, p < 0.0001 and NS, not significant).

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The previous section highlights that iron is important for V. vulnificus growth and pathogenicity, and that the genome of Env1 contains many genes involved in uptake and transport. Thus, the effect of iron depletion on the secretion of this factor was investigated by growing Env1 in 0.5 × VNSS supplemented with 100 μM of the iron chelator, 2-2’ dipyridyl. The survival of T. pyriformis increased more than 90% in the cell-free supernatant of V. vulnificus grown under iron-deplete conditions (Table 3-4).

For further experiments (Table 3-4), a toxicity scale similar to the BACTOX test to evaluate the toxicity of the cell-free supernatants after different treatments was used (Schlimme et al., 1999). The supernatant pH was 4.77, which T. pyriformis tolerates as demonstrated by the 0.5 × VNSS media control. The supernatant was treated with heat at 95 °C for 2 hours, frozen at -20 °C and treated with protease and proteinase K but none of these treatments were effective at reducing the toxicity of the cell-free supernatant. Ultrafiltration using Amicon® Ultra-0.5-10,000 NMWL filters demonstrated that the factor is under 10 KDa.

Table 3-4 Health of T. pyriformis, 1 hour after the addition of each treated cell-free supernatant. Treatment T. pyriformis health# Heat treatment (2 hours-95 °C) * Freeze/ Thaw (-20 °C) * Ultrafiltration (Amicon® Ultra-0.5-10,000 NMWL) * Protease * Proteinase K * Supernatant of ENV1 grown under iron-deplete conditions ****

# The protozoan concentration in the media control was considered to be 100% survival; the concentration in each treatment was compared with the control to determine the relative percent survival, (****) n >90% and (*) n <10%).

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3.3.10 Presence of biosurfactants

The death of T. pyriformis when exposed to the cell-free supernatant of V. vulnificus Env1 is similar to effect of biosurfactants that permeabilise eukaryotic cell membranes and cause lysis (Figure 3-11). For example, a strain of Pantoea ananatis exhibited grazing resistance against Dictystelium discoideum. Genes involved in the biosynthesis of a biosurfactant, rhlA and rhlB were involved in the cytotoxicity and a glycolipid with monohexose-C10-C10 was identified as the main factor responsible for compromising cellular integrity and leading to cell lysis (Smith et al., 2016). The damaging effects of non-ionic surfactants such as Triton X-100 to the T. pyriformis membrane has been previously demonstrated (Dias et al., 2003). In order to investigate if a biosurfactant is produced by Env1, emulsification activity of the cell-free supernatants was measured using n-hexadecane. Cell-free supernatants from V. vulnificus Env1 grown in 0.5 × VNSS and 0.5 × VNSS supplemented with 100 μM 2-2' dipyridyl were compared to media controls (0.5 × VNSS). The emulsion was observed after settling at RT for 10 minutes. The lipophilic dye Sudan black was added to the n-hexadecane to increase visibility of each layer. After 10 minutes, there was no mixing of layers in any sample, and thus no emulsifying activity was detected (Table 3-5). These results suggested there is no production of an extracellular biosurfactant that might be involved in Env1 toxicity against T. pyriformis. Thus, the toxic factor is a biomolecule that does not have surfactant properties, nor is it a large polypeptide (protein) that is sensitive to heat or proteolysis. This suggests it could be a small molecule or secondary metabolite.

Table 3-5 Emulsification activity of cell free supernatant of V. vulnificus Env1 using n- hexadecane. Sample Emulsion index# V. vulnificus ENV1 in VNSS 0.10% V. vulnificus ENV1 in VNSS + 100 μM dipyridyl 0.10% VNSS 0.01%

# The emulsion index was calculated as the percentage of ratio of the height of the emulsion layer to the total height of the sample (Zavala-Moreno et al., 2014).

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3.3.11 Conclusion

In this work, place of isolation and genotype did not correlate with grazing resistance of V. vulnificus. However, one of the environmental strains (Env1) showed grazing resistance against T. pyriformis. One of the defence mechanisms used by Env1 was a secreted toxic factor that kills T. pyriformis. The factor was determined to be iron- dependent, heat-stable, proteinase-resistant and had a molecular mass of <10 kDa and is not a biosurfactant. Potential grazing resistance mechanisms were further investigated by whole genome sequencing of the grazing resistant strain and comparison with genomes of grazing sensitive strains. The unique genes discussed here include a putative internalin, a putative rhs-related protein, an ankyrin protein, a T1SS secreted agglutinin RTX membrane protein and a suppressor for copper-sensitivity ScsD. Each of these factors may play an important role in virulence and defence in V. vulnificus and will be further investigated.

In order to investigate the potential iron-dependent factors involved in the toxicity of V. vulnificus to T. pyriformis, the transcriptomic responses of V. vulnificus under iron-replete and -deplete conditions were investigated and are presented in Chapter 4.

3.4 Acknowledgements

Whole genome sequencing data were generated by the Ramaciotti Centre for Genomics using PacBio RS II. I would like to thank Dr. Tonia Russel and Dr. Nandan Deshpande in help with the De novo assembly using HGAP.3 Pacific Biosciences.

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4 Chapter 4: Investigation of the secreted toxic factor produced by Vibrio vulnificus Env1

4.1 Introduction

Bacteria secrete exotoxins in order to damage surrounding cells that may pose a threat like immune cells. Some exotoxins are secondary metabolites that may cause growth inhibition, paralysis and cell lysis of target cells. The production of secondary metabolites also repels or inactivates predators such as protozoa. The release of nutrients by lysing predator cells will also provide nutrients to the bacterial community (Jousset et al., 2008).

Our group has previously shown that when Tetrahymena pyriformis is co-incubated with Vibrio fischeri biofilms, T. pyriformis undergoes morphological changes after 24 hours, and lysis of 60% of the T. pyriformis cells occurs. It is speculated that this lysis may be the result of the production of lytic compounds by V. fischeri (Chavez-Dozal et al., 2013). Co-culture of bacteria bearing a Shiga toxin (Stx)-encoding temperate bacteriophage or the purified Stx kills Tetrahymena thermophile (Lainhart et al., 2009). It is believed that the production of hydrogen peroxide by T. thermophile signals the bacteria to produce Stx. This is evident as mutants in the SOS response (recA mutations) or in enzymes that breakdown hydrogen peroxide (catalase) do not kill T. thermophile (Lainhart et al., 2009). Janthinobacterium lividum and Chromobacterium violaceum produce a purple pigment, violacein, which is toxic to nanoflagellates. Violecelin causes paralysis of the flagellum of the nanoflagellate, Ochromonas sp., after which the cells swell and burst in less than an hour (Matz et al., 2004).

Iron regulates many cellular functions and is necessary for the activity of variety of enzymes and virulence factors, and for DNA replication and energy production. For example, iron is the integral part of iron complexes in enzymes such as iron–sulphur clusters attached to functional proteins (Litwin & Calderwood, 1993b, Frazzon & Dean, 2003, Payne et al., 2016). Iron is also involved in cell signalling and regulatory pathways. One of most wide-spread regulators in bacteria is the ferric uptake regulator, Fur. Fur is known to regulate many systems (Hantke, 1987, Litwin & Calderwood, 1993a) including

124 | P a g e the hemolysin, vvhA (Lee et al., 2013b), the non-coding RNA, ryhB (Večerek et al., 2007) and superoxide dismutase, sodA (Mey et al., 2005).

An environmental strain of Vibrio vulnificus, Env1 was shown in the previous chapter to be resistant to grazing by T. pyriformis. One of the antiprotozoal factors used by Env1 is a secreted iron-dependent, heat stable, proteinase-resistant factor with a molecular mass of <10 kDa. In this chapter, the transcriptomic profile of V. vulnificus Env1 grown under iron-depleted conditions was compared to the iron-replete condition. A variety of differentially expressed genes were investigated further, including those involved in ammonia production, biotin and purine biosynthesis, copper transfer, virulence and transcriptional regulation. In order to investigate each of these genes, add back experiments, ammonia quantification and site-directed mutagenesis was performed. Here, a master regulator, arcA, which is shown to be involved in the grazing resistance of Env1 is described in terms of its regulatory role and its specific role in controlling predation defence.

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4.2 Materials and methods

4.2.1 Strains and growth conditions

Organisms used in this study are listed in (Table 4-1). Bacterial strains were routinely grown in LB broth supplemented with 2% NaCl and on agar plates (Sambrook et al., 1989), as appropriate, with carbenicillin (100 µg ml−1) and chloramphenicol (17 µg ml−1). Acanthemoeba castellanii was routinely maintained in PYG as described in section 2.2.1 and was passaged 3 d prior to harvesting for experiments and enumerated microscopically using a haemocytometer. T. pyriformis was routinely maintained in PYG as described in 2.2.1. Prior to experiments, 500 μl of T. pyriformis were passaged in 20 ml of 0.5 × nine -1 -1 -1 salts solution (NSS) medium (8.8 g l NaCl, 0.735 g l Na2SO4, 0.04 g l NaHCO3, 0.125 -1 -1 -1 -1 -1 g l KCl, 0.02 g l KBr, 0.935 g l MgCl2.6H2O, 0.205 g l CaCl2.2H2O, 0.004 g l -1 SrCl2.6H2O and 0.004 g l H3BO3) (Mårdén et al., 1985) supplemented with 1% heat- killed P. aeruginosa PAO1 (HKB) (see section 2.2.1 for production of HKB) in a 25 cm2 tissue culture flask, and further incubated at RT statically for 2 d before enumeration and use. This process is necessary to remove the nutrient media and to acclimatise the ciliate to phagotrophic feeding.

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Table 4-1 List of bacterial and protozoal strains. Strain Properties Reference Bacterial strains

E-genotype, Oyster environmental Env1 (Rosche et al., 2005) isolate, WT

Env1 ∆ arcA ∆arcA:pGP704, ampr This study

Env1 ∆ arcA complementation ∆ arcA, pLG401::arcA, ampr, Cmr This study construct

Env1 ∆copA ∆copA:pGP704, ampr This study

Env1 ∆purA ∆purA:pGP704, ampr This study

Env1 ∆smcR ∆smcR:pGP704, ampr This study

Plasmids

pGP704 Suicide vector, oriR6K, mob/RP4, Apr (Miller & Mekalanos, 1988)

pACYC Ori, Mob, promoterless gfp Lynn Gilson, University pLG401 r with multiple cloning site, Cm of Hawaii

Protozoan strains A. castellanii Wild type ATCC 30234

T. pyriformis Wild type ATCC 205063

4.2.2 V. vulnificus growth under iron-replete and -deplete conditions and toxicity of supernatants to T. pyriformis

6 -1 Overnight cultures of V. vulnificus Env1 were adjusted to 10 cells ml (OD600=0.001) in 0.5 × VNSS (iron-replete) (1 g bacteriological peptone, 0.5 g yeast extract, 0.5 g D- glucose, 0.01 g FeSO4·7H2O and 0.01 g Na2HPO4) in 1 litre of 0.5 × NSS (Väätänen, 1977) or 0.5 × VNSS supplemented with 100 μM of the iron chelator 2-2’ dipyridyl (iron- deplete) in 24 well microtitre plates (BD Falcon™, Becton Dickinson, New Jersey, USA).

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Plates were incubated at RT with shaking at 60 rpm for 24 hours and samples were taken each hour in replicates of six. The cell density of each sample was measured by spectrophotometry at OD 600 nm using a spectrophotometer (Halo xs-10; Dynamica, London, UK). Cell-free supernatants were collected at each time point by centrifugation at 3220 × g for 5 minutes and filtration through a 0.22 µm pore size filter (Millipore; Bedford, MA, USA) and used to test for toxicity to T. pyriformis.

4.2.3 Transcriptome of V. vulnificus Env1 grown in iron-replete and -deplete conditions.

6 -1 Overnight cultures of V. vulnificus were adjusted to 10 cells ml (OD600=0.001) in 0.5 × VNSS (iron-replete) or 0.5 × VNSS supplemented with 100 μM of the iron chelator 2-2’ dipyridyl (iron-deplete) in 24 well microtitre plates (BD Falcon™, Becton Dickinson, New Jersey, USA). Plates were incubated for 10 hours at RT with shaking at 60 rpm (early stationary phase) and the supernatant toxicity was confirmed. The samples were fixed in RNAprotect Bacteria Reagent (QIAGEN®). Total RNA was extracted by lysozyme digestion and the RNeasy® Plus Mini kit (QIAGEN®). Total RNA was extracted by the use of the RNeasy® Plus Mini kit according to the manufacturer’s instructions. RNA concentration and purity were determined using a spectrophotometer (NanoDrop ND-1000). RNA integrity of the samples was determined by agarose gel electrophoresis. The RNA was stored at -80°C. For the RNA-Seq sample preparation, the standard Illumina kit was used according to the manufacturer’s protocol (Illumina) and the samples were sequenced by paired-end sequencing on the Illumina Hi-Seq 2500 platform with reads of 100 bp length.

4.2.4 Transcriptome data analysis

The quality of the paired-end reads was initially checked using FastQC (http://www.bioinformatics.bbsrc.ac.uk/projects/fastqc). Illumina adaptors, short and low-quality reads were removed using cutadapt (version 1.11) (Chen et al., 2014). High- quality reads (97% to 98% of the raw reads, Appendix 4) were subjected to sortmeRNA for in silico rRNA depletion (version 2.0) (Kopylova et al., 2015). Messenger RNA reads

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(from 75,495 to 131,958 read pairs) were then mapped to the V. vulnificus Env1 genome (annotated genome of Env1 generated by the Rapid Annotation using Subsystem Technology (RAST) server using Bowtie2 (version 2.2.9). From 95% to 98% of the reads mapped to the reference genome and the numbers of reads mapping each gene were determined using HTSeq (version v.0.6.1p1) (Anders et al., 2015).

The raw count table of transcripts was used as an input for the Deseq2 R package for differential expression analysis (Love et al., 2014). Briefly, the raw counts were normalised according to the sample library size and a negative binomial test was performed to identify the differentially expressed genes. Genes were considered as differentially expressed if their absolute fold-change value was greater than two and the associated adjusted p-value was smaller than 0.05. The normalised transcripts were then log2 (N+1) transformed prior to principal component analysis and UPGMA hierarchical clustering for the sample dendrogram on the heatmap.

4.2.5 Ammonia toxicity assay

In order to determine the tolerance of T. pyriformis to ammonia, the growth of T. pyriformis in media containing a range of concentrations of ammonium chloride was tested. Briefly, 1000 cell ml -1 T. pyriformis in 0.5 × NSS supplemented with 1% HKB with a range of concentrations (0 to 30 mM) of NH4Cl, were incubated for 24 hours and enumerated by inverted microscopy. Overnight cultures of V. vulnificus were adjusted to 106 cells ml-1 in 0.5 × VNSS (iron-replete) or 0.5 × VNSS supplemented with 100 μM of the iron chelator, 2-2’ dipyridyl (iron-deplete) in 24-well microtitre plates (BD Falcon™, Becton Dickinson, New Jersey, USA). The amount of ammonia produced by V. vulnificus was measured after 24 hours of incubation by use of the Hach™ ammonia reagent set (Hach, Loveland, Colorado) according to the manufacturer’s instructions.

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4.2.6 Biotin add-back experiments

V. vulnificus overnight cultures were adjusted so that 106 cells ml-1 in 0.5 × VNSS with 0 to 100 µM biotin (Huang et al., 2008, Kassem et al., 2017) were added to 24-well microtitre plates (BD Falcon™, Becton Dickinson, New Jersey, USA). T. pyriformis was added to each well (104 cell ml-1; determined by inverted microscopy) and the plates were incubated at RT with shaking at 60 rpm for 24 hours. The cell density of each well was measured by spectrophotometry at OD600 nm (Eppendorf® PlateReader AF2200, Hamburg, Germany). Numbers and health of T. pyriformis were determined by microscopy at each time point.

4.2.7 Electroporation and conjugation conditions

To prepare elctrocompetent cells for electroporation, E. coli BW20767 was grown in LB broth overnight at 37 °C with shaking at 200 rpm. The overnight cultures were subcultured and incubated at 37 °C with shaking at 200 rpm until they reached an OD600 = 0.7. The cells were harvested by centrifugation at 4 °C, 3220 × g for 10 minutes. From this point, E. coli BW20767 cells were kept on ice. The pellets were washed twice with ice-cold 10% glycerol. The cells were resuspended in ice-cold 10% glycerol and stored at -80 °C until use in electroporation

Electroporation was performed by mixing 10 μl of ligation mixture with 100 μl of electrocompetent cells on ice. The mixture was then electroporated at 1.8 kV pulse (Program Ec1, MicroPulser™, Bio-Rad). After electroporation, cells were recovered by addition of 1 ml of SOC broth (tryptone 20 g l-1, yeast extract 5 g l-1, NaCl 0.5 g l-1and KCl 0.186 g l-1) and incubated at 37 °C with shaking at 200 rpm for 1 hour. After recovery, cells were plated onto LB medium supplemented with the appropriate antibiotic.

Conjugation was performed by washing the overnight cultures of donor and recipient cells in fresh LB. The donor and recipient cells were mixed at a ratio of 1:3 on 0.22 μm membrane filters (GSWP Millipore™) and placed on an LB agar plate. The plates were incubated for 18 hours at 37 °C. Transconjugants were selected by resuspending the

130 | P a g e resultant growth in 10 mM MgSO4 and plating serial dilutions on LB agar supplemented with the appropriate antibiotic.

4.2.8 Site-directed mutagenesis and complementation

To generate insertion mutations in smcR, purA, copA and arcA, a 200 bp internal fragment of each gene was amplified by PCR. The PCR products were then digested by restriction enzyme SalI and ligated into the SalI site of the suicide vector pGP704. The ligation was confirmed by PCR. The constructed plasmid was transformed into E. coli BW20767 by electroporation and transferred to V. vulnificus Env1 by conjugation as described above (section 4.2.7). Transconjugants were plated onto LB medium supplemented with polymyxin B (40 µg ml-1) to select for V. vulnificus and carbenicillin (100 μg ml-1) to select for the insertion. The mutation was confirmed by PCR and sequencing. In order to complement each mutation, the entire gene, including the promoter region, was PCR amplified. The amplicons were digested with restriction enzyme SalI or SphI and ligated into the corresponding restriction site of the cloning plasmid, pLG401. The constructed plasmid was transformed into E. coli BW20767 by electroporation and selected by plating on LB medium containing chloramphenicol (34 μg ml-1). PCR was used to confirm the ligation. Transconjugants were plated onto LB medium supplemented with polymyxin B (40 µg ml-1) to select for V. vulnificus strains and chloramphenicol (17 μg ml-1) to select for complementation cloning vector.

4.2.9 Assessment of resistance of V. vulnificus mutants to T. pyriformis predation

To assess grazing resistance of planktonic V. vulnificus mutants, overnight cultures were adjusted so that 106 cells ml-1 in 0.5 × VNSS were added to 24-well microtitre plates (BD Falcon™, Becton Dickinson, New Jersey, USA). T. pyriformis was added to each well (104 cell ml-1; determined by inverted microscopy; ECLIPSE TE2000-5, Nikon, Japan) and the plates were incubated at RT with shaking at 60 rpm for 24 hours. The cell density of each well was measured by spectrophotometry at OD 600 nm (Eppendorf® PlateReader

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AF2200, Hamburg, Germany). Planktonic fractions were collected for CFU ml-1 counts. Numbers and health of T. pyriformis were determined by microscopy at each sampling time.

4.2.10 Data analysis

Statistical analysis was performed using GraphPad Prism version 7.03 for Windows, GraphPad Software, La Jolla California USA, (www.graphpad.com). Data that did not follow Gaussian distribution was determined by analysing the frequency distribution graphs and was transformed using natural log. Two-tailed student’s t-tests were used to compare means between experimental samples and controls. For experiments including multiple samples, one-way or 2-way ANOVAs were used for the analysis and Sidak's or Dunnett's multiple comparison Test provided the post-hoc comparisons of means when appropriate.

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4.3 Results and discussion

V. vulnificus ENV1 was isolated from an oyster from Louisiana, USA (Smith & Oliver, 2006) and exhibits serum survival similar to clinical (C-type) strains (Bogard & Oliver, 2007). V. vulnificus infections are highly correlated with elevated serum iron levels. Virulence assays in mice have shown that the infectious dose and time of death post- infection both decrease as the serum iron levels increase (Wright et al., 1981).

In Chapter 3 it was shown that Env1 exhibits resistance against T. pyriformis predation. Furthermore, the cell-free supernatant from Env1 showed toxicity towards T. pyriformis. The toxic factor is secreted, heat-stable, proteinase-resistant and has a molecular mass of <10 kDa. Iron depletion significantly increased the survival of the T. pyriformis in the presence of the cell-free supernatant of Env1. In order to investigate the potential iron- dependent factors expressed by this strain, total RNA was extracted from Env1 grown under iron-replete and -deplete conditions and the transcriptomes was analysed.

4.3.1 V. vulnificus growth under iron-replete and -deplete conditions and toxicity of supernatants to T. pyriformis

In order to establish when the toxicity is expressed, Env1 was grown in 0.5 × VNSS (iron- replete condition) and 0.5 × VNSS (Väätänen, 1977) supplemented with 100 μM 2-2' dipyridyl (iron-deplete condition) for 24 hours and samples were taken hourly. The cell density was measured by spectrophotometry at OD 600 nm (Halo xs-10; Dynamica, London, UK) and toxicity of the cell-free supernatant of each time point was determined. Under iron-deplete conditions, the bacterial growth is less than the control in the presence of iron, but nonetheless, Env1 cells enter the stationary at the same time under both iron- replete and -deplete conditions. The cell-free supernatant of Env1 grown under iron- replete conditions becomes toxic to T. pyriformis after 10 hours growth, which is when the cells reach stationary phase. It is common that the production of secondary metabolites by bacteria occurs in stationary phase and not log phase cells. Thus, RNA extraction was carried out on early stationary phase cells.

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In order to confirm that the cell-free supernatants of cells grown in iron-deplete conditions are not toxic, supernatants from cells grown for 24 hours were tested and shown to be non-toxic. In order to confirm that the lack of toxicity was not due to the lower cell density, supernatants were prepared from cultures with a higher inoculum (107 cells ml - 1) but there was no significant difference in toxicity.

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l 0 .0 5

. V 0 .0 0 0 2 4 6 8 1 0 1 2 1 4 1 6 1 8 2 0 2 2 2 4 H o u r

Figure 4-1 Growth of V. vulnificus Env1 in 0.5 × VNSS (iron-replete) and 0.5 × VNSS supplemented with 100 μM 2-2' dipyridyl (iron-deplete). The time point at which the cell free supernatant of Env1 shows toxicity to T. pyriformis is indicated by the red line. Error bars represent standard deviation. The experiment was run in three replicates and repeated three times separately.

4.3.2 RNA-seq revealed differentially expressed genes under iron-deplete compared to -replete conditions.

The iron-dependent toxicity was further investigated by analysing the transcriptomic profile of V. vulnificus grown in 0.5 × VNSS (iron-replete condition) and 0.5 × VNSS supplemented with 100 μM 2-2' dipyridyl (iron-deplete condition). Total RNA was isolated from 3 biological replicates and subjected to Illumina HiSeq 2500 sequencing.

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Between 94.66% and 98.35% of reads were mapped to the V. vulnificus Env1 genome (Chapter 3) generated by RAST and between 82.93% and 87.89% were mapped against the genome of V. vulnificus Yj016.

Significantly differentially expressed genes were considered at fold-change of 2.0 and adjusted p-value of p<0.05. Here, the differential expression ranges from log2 fold change of -10.10 to 7.86 where 108 genes were significantly up-regulated and 140 genes were significantly down-regulated in iron-replete conditions in comparison to the iron-deplete condition. The complete list of the 248 differentially expressed genes can be found in the Appendix 4. The genes that could potentially be involved in grazing resistance based on their roles in survival and pathogenicity will be individually investigated. Many of these genes are hypothetical proteins and in the iron-replete condition 42 of the differentially expressed genes have no predicted function (Appendix 4).

As expected, most of the genes related to systems involved in iron uptake and transport are down-regulated in the iron-replete condition since iron is readily available in the environment. These genes include ABC transporters, catechol siderophore, isochorismate pyruvate-lyase and isochorismase of siderophore biosynthesis, TonB system, vulnibactin, aerobactin, ferric siderophore transport system and ferrous iron transport system (Appendix 4). The genes selected in the next sections for further analysis are based on unique genes in Env1 described in Chapter 3 and genes previously described as virulence and survival factors.

4.3.3 Effect of ammonium on T. pyriformis growth

Analysis of the transcriptome shows five genes related to ammonia production were differentially expressed. Three genes predicted to be involved in ammonia production were up-regulated and 2 genes involved in the consumption of ammonia were down- regulated under iron-replete conditions (Table 4-2). This would lead to secretion of ammonia when iron is abundant. Bacteria are known to use ammonia as a secondary metabolite for defence and virulence. For example, Vibrio shiloi produces a non- dialysable heat-resistant factor that when added to NH4Cl, inhibits photosynthesis of coral endosymbiotic zooxanthellae (Ben‐Haim et al., 1999). The chitin-dependent toxicity of 135 | P a g e

Vibrio cholerae biofilms towards the nanoflagellate, Rhynchomonas nasuta was shown to be attributed partly to ammonium (3.5 mM) detected in the supernatants of 3-day-old biofilms of V. cholerae grown on chitin flakes (Sun et al., 2015).

Several studies have reported toxicity of ammonia to protozoa, especially ciliates such as Paramecium bursaria and Euplotes vannus (Xu et al., 2004, Henglong et al., 2005). Various ciliates tolerate different concentrations of ammonia where the lethal concentration differs from 2.5 M for Stentor coeruleus to 25.9 M for Coleps hirtus (Klimek et al., 2012). Ammonia is abundant in the water column and can be the result of catabolism of organic material.

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Table 4-2 Genes involved in ammonia accumulation, production and transport.

Log2 NCBI RAST Locus Common Fold p- Annotation Locus ID ID name Chan adj

Category ge BJD94_05 1246305.4.peg Catalyses the 5.59 120 .1082 Hydroxylamine reduction of 7.86 611 reductase (hcP) hydroxylamine to form E-11 water and ammonia

BJD94_05 1246305.4.peg Catalyses the oxidative 455 .1152 1.68 cleavage of amine oxidase 2.71 364 alkylamines into E-14 aldehydes and ammonia

Ammonia production Ammonia BJD94_05 1246305.4.peg cytochrome C 0.00 Dissimilatory nitrate 275 .1114 nitrate 2.91 22 reduction to ammonia reductase

BJD94_07 1246305.4.peg Catalyses the ATP- 085 .1477 dependent conversion asparagine of aspartate into 0.00 synthetase B -2.23 asparagine, using 02 (asnB) glutamine or ammonia (lower efficiency) as a source of nitrogen

BJD94_10 1246305.4.peg Forms glutamine from 540 .2172 glutamine 9.20 ammonia and synthetase -4.89 03E- glutamate with the

Ammonia consumption Ammonia (glnA) 86 conversion of ATP to ADP and phosphate

To determine the tolerance of T. pyriformis to ammonia, the growth of T. pyriformis in the presence of different concentrations of ammonia were tested. As the concentration of ammonia increased in the medium, the growth of T. pyriformis decreased (Figure 4-2). At 20 mM ammonia, T. pyriformis growth was inhibited, although the protists were still viable and motile. However, at 350 mM ammonia, the killing of T. pyriformis was on par with the killing effect of the Env1 supernatant. To investigate whether ammonia was produced by V. vulnificus, the ammonia concentration in the 24-hour cell-free supernatant of Env1 was measured.

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0 0 .0 2 .5 5 .0 7 .5 1 0 .0 1 2 .5 1 5 .0 1 7 .5 2 0 .0 2 2 .5 2 5 .0 2 7 .5 3 0 .0 m M Figure 4-2 Effect of ammonia on survival of T. pyriformis. The red line represents the initial inoculum. The amount of ammonia produced by V. vulnificus after 24 hours in 0.5 × VNSS (iron-replete) (1.45mM) or 0.5 × VNSS supplemented with 100 μM of the iron chelator, 2-2’ dipyridyl (iron-deplete) (1.02 mM) are indicated by red arrows. Error bars represent standard deviation. The experiment was run in three replicates and repeated three times separately.

Table 4-3 Amount of ammonia produced by V. vulnificus under the following conditions. Conditions Ammonia concentration V. vulnificus ENV1 in VNSS 1.45 mM V. vulnificus ENV1 in VNSS + 100 μM dipyridyl 1.02 mM V. vulnificus ENV1 and T. pyriformis in VNSS 1.60 mM V. vulnificus ENV1 and T. pyriformis in VNSS + 100 μM dipyridyl 1.35 mM VNSS 0.16 mM VNSS + 100 μM dipyridyl 0.21 mM

The amount of ammonia produced by V. vulnificus grown under a variety of conditions was measured after 24 hours of incubation using the Hach™ ammonia reagent set (Table 4-3). As there was no ammonia used in the preparation of 0.5 × VNSS and these results

138 | P a g e show that the background ammonia in the media is very low (0.1 - 0.2 mM), the ammonia in the supernatant would be due to catabolism of proteins by V. vulnificus. The amount of ammonia produced by V. vulnificus in 0.5 × VNSS with or without the grazer after 24 hours was 1.5 mM. This amount of ammonia would not be enough to affect the viability of T. pyriformis (see red arrows in Figure 4-2). Although ammonia could be a contributing or enhancing factor, it is not likely to be the main grazing resistance factor.

4.3.4 Genes involved in biotin biosynthesis

Three genes involved in biotin biosynthesis were up-regulated in iron-replete conditions (Table 4-4). Biotin is an essential vitamin (B7) which bacteria can synthesise using a two-stage pathway. The first step in conversion of fatty acids into biotin is the conversion of the ω-carboxyl group of malonyl-CoA into a methyl ester by BioC (O- methyltransferase). Strains with an inactive bioC require exogenous biotin for growth. BioD (dethiobiotin synthetase) is second last enzyme in the pathway that the synthesises dethiobiotin from 7,8-diaminononanoate. BioB (biotin synthase) in the final step in the pathway and forms biotin from dethiobiotin (Lin & Cronan, 2011). Interestingly bioB has been reported to be induced in response to low iron but independently of Fur in a V. cholerae O395 classical strain (Mey et al., 2005).

A Mycobacterium tuberculosis bioA (biotin biosynthetic enzyme 7,8-diaminopelargonic acid synthase) mutant is unable to establish infection in mouse models (Woong Park et al., 2011). BioA was also shown to be an essential factor for intracellular survival and escape of Francisella tularensis from macrophage phagosomes (Napier et al., 2012). A virulence factor of F. tularensis, BioJ determines the chain length of the biotin valeryl side-chain (Feng et al., 2014).

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Table 4-4 Genes involved in biotin biosynthesis.

Log2 NCBI Locus RAST Locus Common Fold p-adj Annotation ID ID name Chang

Category e Catalyses the 1246305.4.peg bioB 0.000 formation of BJD94_05490 (biotin 4.72 .1160 6 biotin from synthase) dethiobiotin

bioC 1246305.4.peg (malonyl- Pimeloyl-ACP .1158 [acyl-carrier 0.005 biosynthesis in BJD94_05480 4.76 protein] O- 9 biotin methyltransfer production as)

Biotin biosynthesis Biotin BioD 1246305.4.peg Synthesises (ATP- 7.661 biotin from 7,8- BJD94_05475 .1157 dependent 6.64 24E- diaminononanoa dethiobiotin 06 te synthetase)

In order to investigate the role of biotin biosynthesis in the grazing resistance of Env1, exogenous biotin was added to iron-deplete media (100 nm to 100 µM) in planktonic grazing experiments. However, addition of biotin to the iron-deplete condition did not restore the grazing resistance phenotype (Figure 4-3). If disruption of biotin biosynthesis was the reason for grazing sensitivity, addition of the biotin should have restored the grazing resistance phenotype of Env1.

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l u

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. M M M M M V µ µ µ µ µ 0 .1 1 0 0 0 5 0 in in 1 t n t in o i o t in i t i o t B io B i o B i B B

Figure 4-3 V. vulnificus ENV1 in iron-deplete (0.5 VNSS supplemented with 100 μM 2- 2' dipyridyl) condition was supplemented with different concentrations of biotin (100 nm to 100 µM) and exposed to grazing by T. pyriformis. The experiment was run in three replicates and repeated three times separately. Error bars represent standard deviation. Statistical analysis was performed using 2-way ANOVA and Sidak's multiple comparisons test. Statistical significance is indicated by (****, p < 0.0001).

4.3.5 Genes involved in pathogenicity and virulence

Several genes previously indicated to be involved in virulence in Vibrio spp. were differentially expressed in the presence of iron when compared to growth in the iron- depleted media (Table 4-5). In this section, each one of these genes is further examined and site-directed mutagenesis is used in order to assess the potential role of each gene in the grazing resistance against T. pyriformis.

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Table 4-5 Genes involved in pathogenicity and virulence regulation.

NCBI RAST Log2 Locus ID Locus Common name Fold p-adj Annotation ID Change

Category BJD94_01050 1246305 copper- 1.91 0.01631 Copper- .4.peg.2 transporting transporting 18 ATPase (copA) ATPase

BJD94_07380 1246305 CopG protein 1.55 2.55263 CopG protein .4.peg.1 (copG) E-07 539

Copper resistance Copper

BJD94_16325 1246305 adenylosuccinat 3.74 0.00052 Purine

.4.peg.3 e synthetase biosynthesis, 271 (Adss) or (purA) catalyses the first step in the de Purine novo biosynthesis of AMP BJD94_15350 RTX toxins 1246305 RTXA 1.05234 determinant A .4.peg.3 -3.03 determinant E-06 and related Ca2+- 070 binding proteins

BJD94_04465 1246305 T1SS secreted 8.67073 T1SS secreted .4.peg.9 -3.00 agglutinin RTX E-37 agglutinin RTX 53

MARTX BJD94_19450 1246305 RTX toxins and .4.peg.3 RTX toxins -1.41 0.04002 related Ca2+- 920 binding proteins

BJD94_08750 1246305 Anaerobic 1.83 2.71518 Master regulator

.4.peg.1 respiration E-08 under both 799 control protein aerobic and (arcA) anaerobic conditions BJD94_12730 1246305 Transcriptional 1.74 0.02886 Master virulence .4.peg.2 regulator (aphA) regulator

Master regulators Master 571

Host immune systems utilise the antibacterial properties of soft metals such as copper and iron in the phagosome (Hood & Skaar, 2012, Samanovic et al., 2012). Therefore, many bacteria possess metal efflux pumps (such as copper tolerance systems) in order to survive

142 | P a g e in the host (German et al., 2013). It has been shown that V. cholerae copA, together with copG (VC2216), is required for copper tolerance but not for intestinal colonisation of the infant mouse. However, copG did not play a role in copper tolerance under aerobic conditions (Marrero et al., 2012). Deletion of copA in E.coli, iron uptake transporters (feoAB and entC) and manganese uptake transporter (mntH), and Cu(I)-translocating P1B-type ATPase (cueA) in Pseudomonas aeruginosa, resulted in decreases in grazing resistance against Dictystelium discoideum (Hao et al., 2016).

Here copA and copG, were up-regulated in presence of iron. The insertion mutant of copA (Cu(I)-translocating P-type ATPase) was examined in planktonic grazing assays with T. pyriformis but there was no significant difference when compared to the wild type (Figure 4-4). Copper resistance is involved in survival of bacteria in phagosomes (German et al., 2013), and thus, the data here suggests that copper efflux is not an essential mechanism for surviving predation.

Previously, it was demonstrated that proteins involved in purine and pyrimidine biosynthesis were induced in V. vulnificus grown under iron-rich conditions, probably reflecting that rapid synthesis of nucleic acids was needed for replication of the cells in the presence of iron (Alice et al., 2008). The purA gene, catalyses the first step in the de novo biosynthesis of AMP and is up-regulated in the transcript data (Table 4-5). Many bacteria including, Vibrio spp., have been shown to require purine and pyrimidine biosynthesis for successful host colonisation, e.g. V. cholerae requires purC, purF, purK and purL (Merrell et al., 2002) as well as purD, purH and purK gene for colonisation of intestinal epithelial cells (Chiang & Mekalanos, 1998), as demonstrated in different large- scale signature-tagged mutagenesis screenings. Furthermore, V. vulnificus mutants of pyrH and purH wer impaired for in vivo growth. These mutants had decreased cytotoxic activity against HeLa cells as well as increased intraperitoneal LD50 when tested in mice virulence assays (Kim et al., 2003b). De novo nucleotide biosynthesis is required for bacterial growth in vivo, especially in blood (Samant et al., 2008). However, data presented here demonstrate that there is no difference in grazing resistance of the purA mutant and the wild type (Figure 4-4), and thus, purA is not involved in the grazing resistance of V. vulnificus Env1.

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Purine biosynthesis and copper tolerance may still play role in the survival of V. vulnificus Purine biosynthesis and copper tolerance may still play roles in the survival of V. vulnificus Env1 in the phagosomes of protozoa. Future experiments are required to investigate the intracellular survival of mutants in purine biosynthesis (purA), copper tolerance (copA and copG) as indicated in RNA-seq data (Table 4-5) and scsD (Table 3-2), a unique suppressor of copper-sensitivity in the genome of V. vulnificus Env1.

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Figure 4-4 Biomass of V. vulnificus Env1 wild type and purA and copA mutants exposed to grazing by T. pyriformis for 24 hours. (A) Planktonic V. vulnificus cells in grazed and ungrazed cultures were enumerated by spectrophotometry (OD 600 nm) (A), and T. pyriformis was enumerated by inverted microscopy (B). The experiment was run in three replicates and repeated three times separately. Error bars represent standard deviation. Statistical analysis was performed using 2-way ANOVA and Sidak's multiple comparisons test (A) and 1-way ANOVA and Sidak's multiple comparisons test comparing all strains with the VNSS media control (B). Statistical analysis revealed no significant differences.

RtxA1 is a known virulence factor in V. vulnificus. RTX toxins are under regulation of a cascade of transcriptional regulators. In V. vulnificus CMCP6, the rtxA1 gene is

145 | P a g e positively regulated by the HlyU protein (Liu et al., 2007) while SmcR represses the expression of rtxA1 by repressing hlyU (Shao et al., 2011). In V. vulnificus MO6-24, under high cell densities and iron-limiting conditions, vvpE was highly expressed while rtxA1 and vvhA were highly expressed at low cell densities regardless of iron levels (Figure 1-2) (Kim et al., 2013a). Likewise in the biotype 2 strain, CECT4999 where rtxA13 has been shown to effect health of amoebae, rtxA13 was not over-express in iron- rich or -poor conditions and was only induced when exposed to eukaryotic cells, namely eel erythrocytes and phagocytes (Lee et al., 2013a). It has been previously hypothesised that rtxA1 and vvhA further destroy eukaryotic targets after iron has been scavenged during initial colonisation (Kim et al., 2013a).

There are various recombinations of the MARTXVv toxin in V. vulnificus, some of which were probably acquired from other marine pathogens by horizontal gene transfer (HGT). Involvement of rtxA1 in pathogenicity has been determined to differ in various strains, depending on the type of MARTX contained. For example, intra-gastric inoculation of an rtxA1 deletion mutant of the clinical strain CMCP6 resulted in a 2,600-fold increase in LD50 while resulting in a 180-fold increase in LD50 in MO6-24/O (Kwak et al., 2011).

In Chapter 3, three coding sequences (CDS) annotated as RTX toxins in the Env1 genome were identified. The 5,206-amino acid MARTXVv in the clinical grazing sensitive strains V. vulnificus CMCP6 and YJ016 (Kwak et al., 2011), as well as the RTXA determinant (1246305.4.peg.3070) in Env1 are 94% identical (Table 4-5). In addition, the RTX toxins and related Ca2+-binding proteins (1246305.4.peg.3920) have homologues that are more than 94% identical in the grazing sensitive strains such as MO6-24, CMCP6 and Yj016. In contrast, a type I secretion system (T1SS) secreted agglutinin, RTX, on chromosome I (1246305.4.peg.953) (Figure 3-10) does not have homologues in any of the grazing sensitive strains.

In the transcriptome of Env1 grown under iron-replete conditions, all of the RTX toxins and related Ca2+-binding proteins were down-regulated compared to the iron-depleted condition (Table 4-5). While the production of RTX toxins are under a complex signalling and regulatory system, they do not appear to be involved in the iron-dependent

146 | P a g e antiprotozoal defence of Env1 since Env1 is not protected against predation by T. pyriformis under iron limitation (Figure 4-5).

Bacteria utilise complex gene regulatory systems to coordinate cellular functions. In Vibrio spp. some of these intricate pathways have been described in detail. Here the genes encoding the virulence regulators anaerobic respiration control protein A (arcA) and a transcriptional regulator (aphA) were upregulated under iron-replete conditions (Table 4-5). In V. cholerae, virulence genes are regulated by a cascading system where a master regulatory system leads to expression of CT, TCP, and other virulence genes (Matson et al., 2007). These virulence genes include arcA and aphA.

AphA together with AphB activates the transcription of the membrane-bound transcription factors, TcpP and TcpH (Kovacikova & Skorupski, 2001, Kovacikova et al., 2004). TcpP and TcpH, together with ToxR/ToxS activate the expression of toxT, which results in the expression of virulence factors such as CT and TCP (DiRita & Mekalanos, 1991, DiRita et al., 1996, Kovacikova & Skorupski, 2001). The expression of aphA is repressed by the quorum sensing (QS) master regulator HapR (Kovacikova & Skorupski, 2002). AphA in turn, regulates several other genes including repression of alsR and alsS that are responsible for the biosynthesis of acetoin, a product that prevents intracellular acidification in presence of glucose, and two putative signal transduction proteins with EAL and GGDEF domains that oppositely influence motility and biofilm formation (Kovacikova et al., 2005).

Previously known as fexA, arcA from V. vulnificus it is 84% to 97% similar to ArcA, the anaerobic respiration control global regulator of other Enterobacteriaceae. A V. vulnificus arcA mutant previously exhibited a substantial decrease in motility and cytotoxicity toward intestinal epithelial cell lines in vitro (Ju et al., 2005). In V. cholerae, the global anaerobic response regulator, ArcA, functions as a positive regulator of toxT expression (Sengupta et al., 2003). In V. fischeri the luxICDABEG operon responsible for bioluminescence is repressed by ArcA (Bose et al., 2007).

The QS response regulator, HapR has been shown to affect the interactions of V. cholerae and A. castellanii by regulating antiprotozoal activities in biofilms (Matz et al., 2005, Erken et al., 2011, Sun et al., 2015). In V. vulnificus the homologue of the QS master 147 | P a g e regulator, HapR is called SmcR (McDougald et al., 2000). In the V. vulnificus clinical strain, MO6-O, under iron-limiting conditions the five quorum regulatory RNAs (Qrrs) additively repress SmcR (Wen et al., 2016). Since the role of QS master regulators such as HapR has been shown to be important in antiprotozoal activity (Matz et al., 2005, Erken et al., 2011, Sun et al., 2015) and both arcA and aphA interact with QS regulators in V. cholerae and V. fischeri, the Env1 smcR mutant was also tested for grazing resistance.

Although smcR and arcA mutants were easily generated in V. vulnificus Env1, mutagenesis of aphA was not successful, even though aphA mutants have been generated in other strains of V. vulnificus using the same method of mutagenesis (Ju et al., 2005). The smcR mutant in V. vulnificus Env1 did not show any significant difference in grazing resistance against T. pyriformis compared to the wild type. In contrast, the arcA mutant was statistically significantly more sensitive to grazing by T. pyriformis and the predator numbers increased significantly compared to the wild type (Figure 4-5). The grazing resistance phenotype was successfully restored by complementing the arcA mutant with a functional arcA.

Although the arcA mutant was statistically significantly grazed compared to the wild type, the biomass was only reduced by 37% compared to the non-grazed control. Furthermore, a connection between ArcA and toxin production could not be established. Although the planktonic cells of the arcA mutant were more sensitive to predation by T. pyriformis, a reduction in toxicity of the cell-free supernatant of arcA mutant was not observed. These results indicate that there are multiple antiprotozoal defense mechanisms used by V. vulnificus Env1.

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l t y A R 1 c n c v id r e n r a m y m s E 1 e ip v l 1 D n p v ' E m n 2 o E - c 2 S A M S c  N r a 0 V 0 n 1 1 i v 1 n v E n E Figure 4-5 Biomass of V. vulnificus Env1 wild type and arcA and smcR mutants exposed to grazing by T. pyriformis for 24 hours. (A) The planktonic V. vulnificus cells in grazed and ungrazed cultures were enumerated by spectrophotometry (OD 600 nm) (A), T. pyriformis was enumerated by inverted microscopy (B). The experiment was run in three replicates and repeated three times separately. Error bars represent standard deviation. Statistical analysis was performed using 2-way ANOVA and Sidak's multiple comparisons test (A) and 1-way ANOVA and Sidak's multiple comparisons test comparing all strains with the VNSS media control (B). Statistical significance is indicated by (****, p < 0.0001 and Ns, P > 0.4).

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Bacteria are known to produce secondary metabolites to defend against predators, destroy host cells, to obtain resources and to kill competitors. Production of these high energy cost metabolites are often under complex regulatory control. Here, a toxic factor produced by Env1 under iron-rich conditions and that is regulated by the master regulator, ArcA was described. This function of ArcA in controlling predator defence has not been previously reported. Interestingly, a mutant in the QS master regulator, smcR, did not show any difference in grazing resistance when compared with the wild type. In contrast, the QS master regulator in V. cholerae, HapR, has been shown to be involved in many grazing resistance mechanisms in V. cholerae (Matz et al., 2005, Erken et al., 2011, Sun et al., 2015).

ArcA is known to be involved in the regulation of many genes depending on oxygen levels. Genes regulated by ArcA and orthologues can be identified by conserved upstream promoter sequences that bind ArcA (ArcA-boxes) in many Gammaproteobacteria including E. coli, Yersinia pestis, Pasteurella multocida, and V. vulnificus (Gerasimova et al., 2003). The interconnected regulation of iron and ArcA, consistent with our findings has been previously described. For example, in Shigella flexneri arcA, was shown to be involved in virulence and plaque formation in anoxic environments together with the ferrous iron transport system Feo, Fur and fumarate and the nitrate reduction regulatory protein (FNR). The anaerobic transcription regulators ArcA and Fnr activate expression of feoABC while ArcA represses iutA, aerobactin siderophore synthesis. Fur is also repressed anaerobically in an ArcA-dependent manner (Boulette & Payne, 2007). The interconnection between FNR and ArcA has been shown, where ArcA represses fnr expression under microaerobic conditions (Sawers & Suppmann, 1992, Shalel-Levanon et al., 2005). FNR is known to have iron-sulfur (4Fe- 4S) clusters but the change in expression of FNR is not statistically significant in our data.

Although ArcA has been described to be involved in virulence, the large number of genes regulated by ArcA makes the identification of the antiprotozoal factor difficult. For example, in avian pathogenic E. coli (APEC) strains, ArcA is in involved in the regulation of enzymatic defences against reactive oxygen species (ROS) and virulence (Jiang et al., 2015). An RNA-Seq analysis comparing the wild type and arcA mutant showed 129 genes were differentially regulated, including genes involved in citrate transport and

150 | P a g e metabolism (citCEFXG), flagellum synthesis (flgB-K, motA and motB) and chemotaxis (cheA) (Jiang et al., 2015).

4.3.6 Conclusion

Overall, this chapter demonstrated that environmental V. vulnificus alters its transcriptome when iron is readily available and expresses many genes that could potentially be involved in grazing resistance. In total 255 genes were differentially expressed including genes related to ammonia production, biotin and purine biosynthesis and copper transfer and virulence master regulators. In order to assess their role in grazing resistance site-directed mutagenesis was performed. A mutant in arcA, the anaerobic respiration control global regulator showed a significant decrease in grazing resistance compared to the wild type. This is the first time ArcA, which is responsible for controlling numerous genes in many bacteria, is demonstrated to play a role in defence against protozoan grazing.

In the next chapter the implications of the antiprotozoal factors demonstrated by this work will be discussed and future experiments that will provide a better understanding of the complex mechanisms and factors governing these defence systems suggested.

4.4 Acknowledgements

Illumina HiSeq 2500 sequencing was performed in was performed in the Singapore Centre for Environmental Life Sciences Engineering (SCELSE). I would like to thank Dr. Florentin Constancias for help in RNA-seq analysis.

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5 Chapter 5: Summary, general discussion and future work

The aim of this project was to elucidate the defence mechanisms that Vibrio spp. employ to survive grazing by predatory protozoa, and to assess the role of pathogenicity factors in environmental survival with respect to protozoan grazing. Vibrio spp. are commonly found in the aquatic environments, both as surface-attached biofilms and suspended as aggregates or on particles in the water column (Colwell et al., 1977, Yildiz & Visick, 2009). The long history of interactions between bacteria and single celled eukaryotes has resulted in co-evolution and development of strategies regulating these interactions (Matz & Kjelleberg, 2005).

The coincidental evolution hypothesis states that ecologically relevant pathogens have acquired anti-protozoal mechanisms that may also function as virulence factors in plant and animal hosts (Levin, 1996, Adiba et al., 2010, Erken et al., 2013). This hypothesis posits that evolution of pathogenicity in some extracellular pathogens is a by-product of bacteria surviving in an environmental niche. Microorganisms must constantly adapt, evolve, and proliferate in order to avoid extinction in the highly competitive environment where food is scarce and predators abound.

Thus, the interactions between bacteria and protozoa are not limited to predator-prey interactions. Protozoa may also serve as a host and vector for bacteria. For example, many bacteria including Vibrio cholerae are known to survive intracellularly in amebae such as Naegleria and Acanthamoeba (Thom et al., 1992), although strains have varying ability to survive internally (Shanan et al., 2016). Living inside these “Trojan horses” (Barker & Brown, 1994) or training grounds for pathogens, not only protects them from adverse environmental stresses but equips them with traits that also cause infections in accidental human and animal hosts (Harb et al., 2000). The intracellular environment of free-living amoebae resembles that of mammalian immune cells such as macrophages and neutrophils. The mechanisms used to kill phagocytised bacteria and the antimicrobial peptides produced are similar. For example, during phagocytosis, Acanthamoeba and macrophages employ similar oxidative stress (oxygen and nitrogen radicals) as a mechanism for killing ingested bacteria (Siddiqui & Khan, 2012). Although the

152 | P a g e differences between the environmental niche of macrophages and amoebae should also be noted (e.g. temperature) (Strassmann & Shu, 2017).

Bacteria may also use pre-intestinal strategies to avoid capture by protozoa (Matz & Kjelleberg, 2005). Some of these mechanisms have been previously described in Vibrio spp. Known grazing resistance mechanisms of V. cholerae include biofilm formation (Matz et al., 2005), expression of the PrtV protease (Vaitkevicius et al., 2006) and the type 6 secretion system (T6SS) (Pukatzki et al., 2006). The quorum sensing (QS) master regulator, HapR in V. cholerae controls many such antiprotozoal activities (Matz et al., 2005, Erken et al., 2011, Sun et al., 2015) although the hapR mutant is not completely sensitive to predation, which suggests that other QS-independent anti-protozoal mechanisms exist (Erken et al., 2011). In contrast, less is known about the mechanisms of grazing resistance of Vibrio vulnificus. The only antiprotozoal activity previously described in V. vulnificus is in biotype 2 where a plasmid-encoded MARTX type III was shown to be involved in the plasmolysis of amoebae (Lee et al., 2013a).

Some strategies are used by the bacteria to avoid being grazed on while others rely on destroying the predator and possibly using them as a nutrient source. In this work, two examples of such strategies are presented. In Chapter 2, an hmgA-mediated pyomelanisation that confers resistance to predation by Acanthamoeba castellanii to an O1 El Tor V. cholerae strain was described. In Chapters 3 and 4, a toxin produced by an environmental E-genotype isolate of V. vulnificus that is capable of killing Tetrahymena pyriformis was investigated and it was established that this mechanism is regulated by the master virulence regulator, ArcA.

5.1 HmgA-mediated pyomelanisation protects V. cholerae against grazing by A. castellanii

V. cholerae biofilms are better protected than their planktonic counterparts, and grazing by nanoflagellates drives the conversion of the smooth morphotype to the biofilm- forming rugose morphotype (Matz et al., 2005). One of the main components of the biofilm matrix, vibrio polysaccharide (VPS) has been shown to be involved in protection

153 | P a g e of V. cholerae biofilms against protozoan predators (Sun et al., 2013). In addition to the physical protection provided by the biofilms, the high density of cells would lead to expression of QS-regulated antiprotozoal mechanisms (Lutz et al., 2013).

In Chapter 2, 3-day-old V. cholerae biofilms were exposed to grazing by A. castellanii in a continuous flow system. This system provides a continuous supply of fresh medium and air that would accurately mimic natural environments. RNA-seq of V. cholerae biofilm under grazing pressure showed 131 transcripts were significantly differentially expressed when compared to the ungrazed control. This shows that gene expression of V. cholerae changes after exposure to protozoan predator. Differentially regulated transcripts included transcripts involved in biosynthetic and metabolic pathways, flagellar assembly and biosynthesis, iron transport, outer membrane proteins, and transcriptional and translational regulators (Appendix 1).

The transcripts of genes involved in tyrosine metabolism were down-regulated in the grazed population, which indicates that the tyrosine metabolic regulon (VC1344-1347) may have a role in the response of V. cholerae biofilms to A. castellanii predation. In the catabolic pathway of L-tyrosine, accumulation of the natural intermediate homogentisic acid (HGA) would lead to its auto-oxidization, leading to the formation of the pigment, pyomelanin. This process is known to generate reactive oxygen species (ROS).

In order to test the grazing resistance of a pigmented mutant disrupted in the gene that converts HGA to maleylacetoacetate (hmgA), biofilms established in microtitre plates were tested for resistance to predation by T. pyriformis and A. castellanii. Using qRT- PCR the down-regulation of tyrosine metabolic regulon in biofilms established in the microtitre plates was also confirmed. Interestingly, the hmgA mutant was more resistant to amoebae predation than the wild type. Increased grazing resistance was correlated with increased production of pyomelanin and thus ROS. Indeed, treatment of the biofilm with H2O2 as well as cell-free supernatant of hmgA mutant provided protection against grazing by A. castellanii to the wild type. The amount of pyomelanin produced when under grazing pressure by A. castellanii increased as well.

Although the hmgA mutant biofilm was protected against grazing by A. castellanii compared to the wild type biofilm, the planktonic cells were not protected against grazing

154 | P a g e by T. pyriformis. This once more supports the fact that different bacterial antiprotozoal mechanisms are effective against different predators, and this is due to differences in niche, feeding mechanisms and cell physiology of the protozoan predator.

5.2 Place of isolation and genotype did not correlate with protozoan grazing resistance

V. vulnificus has been isolated from various clinical and environmental sources. The natural reservoirs for this facultative pathogen include water, sediments, oysters, clams and fish (Oliver et al., 1982, Tamplin et al., 1982, Myatt & Davis, 1989, DePaola et al., 1994, Wright et al., 1996, Bisharat et al., 1999, Harwood et al., 2004). Clinical isolates obtained from infected patients are often from wounds or blood.

The need to categorise and predict the pathogenic potential of this bacterium has led to genotyping systems, such as use of the virulence-correlated gene (vcg) where clinical isolates carry vcgC (C-genotypes) and environmental strains (E-genotypes) contain vcgE (Rosche et al., 2005). Although, it has also been report that some of the virulent strains carry vcgE (Thiaville et al., 2011). Generally, the C-genotype strains have higher survival rates in human serum, which is correlated with possession of the sidephore, viuB that exists in all C-genotypes strains but in very few E-genotype strains (Bogard & Oliver, 2007). Furthermore, E-genotype strains integrate better into aggregates and are taken up by oysters more readily (Froelich et al., 2013).

In Chapter 3 the relationship between place of isolation and genotype with the grazing resistance of V. vulnificus strains was investigated. Twelve isolates were divided into three groups of clinical C-, clinical E- and environmental E-genotypes and the grazing resistance of each group was analysed by exposure of bacterial planktonic cells or biofilms to T. pyriformis and A. castellanii. However, no significant correlation was found between genotype or place of isolation and grazing resistance.

The diversity in pathogenesis and survival mechanisms of Vibrio spp. is well known and attributed to huge genetic diversity and the capacity to uptake exogenous DNA by

155 | P a g e horizontal gene transfer. As a consequence, the pathogenic potential and the presence of specific virulence factors cannot be predicted by taxonomic relations (Roux et al., 2015).

Consistent with our results, in a recent publication, whole genomic analysis of 80 strains of V. vulnificus showed that chromosomal rearrangements in chromosome I and II are not consistent and thus, probably occurred multiple times during the evolution of V. vulnificus. The authors suggested five separate lineage base on phylogenomic analysis, regardless of biotype which do not correspond to pathogenesis towards humans (Roig et al., 2018)

5.3 Master virulence regulator, ArcA, controls antiprotozoal activity of an iron-dependent toxic factor expressed by V. vulnificus

In Chapter 3, 12 strains of V. vulnificus were screened for protozoan grazing resistance, and only 1 of the E-genotype strains isolated from an oyster showed significant grazing resistance and toxicity to T. pyriformis. The phylogenic analysis of the V. vulnificus Env1 and the grazing sensitive strains revealed that Env1 is closely grouped with the rest of E- genotypes and further away from C-genotypes. However, many unique genes that may be involved in pathogenesis or protozoan grazing resistance mechanisms are located on the Env1 genome. These genes were probably acquired by horizontal gene transfer, as evidenced by the presence of mobile elements, and contribute to the fitness of the bacterium and thus are maintained in the genome.

These factors include a putative internalin, a putative Rhs-related protein, a putative ankyrin protein, a type 1 secretion system (T1SS) secreted agglutinin RTX and suppressor for copper-sensitivity, scsD. Although further experiments on the cell-free supernatant of the V. vulnificus Env1 revealed one of the antiprotozoal mechanisms to be a secreted, iron-dependent, heat-stable, proteinase-resistant factor, roles of each of these genes in survival and grazing resistance can be further studied using site-directed mutagenesis.

Chapter 4 was focused on the iron-dependent toxic factor secreted by V. vulnificus Env1. Analysis of changes in gene expression in V. vulnificus Env1 was performed under iron-

156 | P a g e replete and -deplete conditions. There were 255 genes differentially expressed, including genes related to ammonia production, biotin and purine biosynthesis, copper tolerance and virulence determinants. Protozoan grazing experiments showed site directed mutagenesis of arcA, the anaerobic respiration control protein, significantly decreased grazing resistance compared to the wild type. Furthermore, the QS master regulator, SmcR was not involved in grazing resistance of V. vulnificus.

The master regulator, ArcA has been previously shown to be involved in motility and cytotoxicity toward intestinal epithelial cell lines in V. vulnificus (Ju et al., 2005). The Arc two-component system is a common master regulator found in ɣ-proteobacteria which, consists of ArcA, a global response regulator of OmpR family and ArcB, a hybrid sensor histidine kinase (Krell et al., 2010). This master regulator has been shown to be involved in regulation of a diverse set of genes in different bacteria, including genes involved in cytotoxicity and motility of V. vulnificus (Ju et al., 2005), virulence in V. cholerae (Sengupta et al., 2003) and bioluminescence in Vibrio fischeri (Bose et al., 2007). A genomic study of E. coli K12 predicted that 1139 genes are regulated by the arcA in a direct or indirect manner (Salmon et al., 2005). A transcriptomic analysis of an arcA overexpression strain of Mannheimia succiniciproducens showed 79 genes, mostly related to energy metabolism, carbohydrate transport and metabolism being repressed, while most of 82 upregulated transcripts were hypothetical genes (Yun et al., 2012). In results presented here, 42 hypothetical genes were shown to be differentially expressed in the iron-replete condition that may potentially be involved in antiprotozoal activity. Further research could provide more insight into the function of each of these genes and the possibility of arcA being involved in their regulation

5.4 Concluding remarks

The ever-changing conditions of aquatic environments drives the evolution of marine bacteria. The interactions of bacterial populations and the biotic/abiotic environments they live in, is complex and dictates the evolution of these populations (Materna et al., 2012). Vibrio spp. in particular, have a high capacity to evolve via acquisition of new

157 | P a g e genetic information that may potentially increase their survival in the environment (Seitz & Blokesch, 2013).

The newly emerging strains of V. cholerae and V. vulnificus are not only proof that these bacteria are constantly evolving, adapting and proliferating in the environment, but that the environment is driving these adaptations. Furthermore, these changes are not always benign with respect to their interactions with human hosts (Igbinosa & Okoh, 2008) as with the case of the newly emerging V. cholerae O139 strains (O'Shea et al., 2004) and V. vulnificus biotype 3 (Bisharat et al., 1999, Naiel et al., 2005).

Evaluation of pathogenesis of Vibrio spp., as well as their ability to survive in the environment is crucial to better understanding these changes and what drives them. For example, how the genomic content of V. vulnificus allows for its transmission from brackish water systems into filter feeding oysters and into humans where it causes disease after consumption of the oyster (Phillips & Satchell, 2017).

Overall, in this study two new grazing resistance mechanisms were found in V. cholerae and V. vulnificus. In V. cholerae El Tor O1 hmgA-mediated pyomelanisation increased grazing resistance against A. castellanii and in an environmental V. vulnificus, an iron- dependent toxin capable of killing T. pyriformis was identified. Furthermore, the master virulence regulator, ArcA was shown to be involved in regulation of a grazing resistance mechanism.

This project identified genes involved in defence against protozoan grazing of V. cholerae and V. vulnificus, some of those genes are involved in pathogenesis as well as environmental survival. This work further supports the hypothesis that predation acts as a selective force for uptake and maintenance of factors with dual potential in clinical and environmental settings. Recognition of the environment as an important driver of pathogenicity is crucial and much more work needs to be done in this area to improve our understanding of its role in bacterial evolution.

158 | P a g e

5.5 Future work

Future work will be required to identify the roles of each gene identified in Chapter 3 by whole genome comparison in resistance to protozoan grazing and more importantly for potential roles in pathogenicity.

Future work to investigate the secreted antiprotozoal factor includes analysis of the compounds using high-performance liquid chromatography (HPLC). Preliminary data suggests that the compound can be extracted in ethyl acetate as the organic phase retained the toxicity against T. pyriformis.

159 | P a g e

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Appendices

Appendix 1 List of differentially expressed transcripts in Chapter 2.

NCBI Log2 Gene Fold NCBI Locus ID Name Regulation Change Annotation Protein ID VCA086 VCA0868 Down -1.96453 8 VCA057 heme transport protein NP_232966. VCA0576 Down -1.7162 6 HutA 1 NP_230398. VC0749 VC0749 Down -1.55355 scaffold protein 1 NP_230399. VC0750 VC0750 Down -1.47972 HesB family protein 1 NP_231335. VC1699 VC1699 Down -1.46148 hypothetical protein 1 NP_231934. VC2303 VC2303 Down -1.41095 hypothetical protein 1 NP_230396. VC0747 VC0747 Down -1.36977 hypothetical protein 1 NP_230397. VC0748 VC0748 Down -1.33128 cysteine desulfurase 1 NP_231210. VC1570 VC1570 Down -1.29949 quinol oxidase subunit II 1 NP_231211. VC1571 VC1571 Down -1.29949 quinol oxidase subunit I 1 multidrug resistance NP_231258. VC1618 VC1618 Down -1.27825 protein 1 VCA101 NP_233400. VCA1016 Down -1.25793 hypothetical protein 6 1 VCA019 NP_232595. VCA0195 Down -1.22807 hypothetical protein 5 1 4-hydroxyphenylpyruvate NP_230988. VC1344 VC1344 Down -1.22178 dioxygenase 1 NP_230989. VC1345 VC1345 Down -1.22178 oxidoreductase 1 flagellar hook-associated NP_231822. VC2191 flgK Down -1.21458 protein FlgK 1 NP_231639. VC2005 VC2005 Down -1.21331 hypothetical protein 1 NP_230400. VC0751 hscB Down -1.20539 co-chaperone HscB 1 NP_230763. VC1118 VC1118 Down -1.2041 transcriptional regulator 1 NP_230764. VC1119 VC1119 Down -1.2041 short chain dehydrogenase 1 NP_230990. VC1346 VC1346 Down -1.19862 hypothetical protein 1 NP_230991. VC1347 VC1347 Down -1.19862 glutathione S-transferase 1

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NP_230828. VC1183 VC1183 Down -1.19095 hypothetical protein 1 NP_230829. VC1184 VC1184 Down -1.19095 NifS-like protein 1 VCr022 5Sh Down -1.16157 ribosomal RNA NP_231324. VC1688 VC1688 Down -1.15853 hypothetical protein 1 VCr012 5Sd Down -1.1453 ribosomal RNA NP_231838. VC2207 VC2207 Down -1.14507 hypothetical protein 1 NP_231446. VC1812 VC1812 Down -1.13615 hypothetical protein 1 NP_231028. VC1384 VC1384 Down -1.12961 hypothetical protein 1 VCr003 5Sa Down -1.12623 ribosomal RNA VCr019 5Sg Down -1.12124 ribosomal RNA VCr025 VCr025 Down -1.11939 VCr015 5Se Down -1.11364 ribosomal RNA VCr016 5Sf Down -1.10739 ribosomal RNA VCr006 5Sb Down -1.1047 ribosomal RNA succinate dehydrogenase NP_231720. VC2088 sdhB Down -1.10289 iron-sulfur subunit 1 VCr009 5Sc Down -1.09949 ribosomal RNA phenylalanine 4- NP_233214. VCA0828 phhA Down -1.09867 monooxygenase 2 pterin-4-alpha- NP_233213. VCA0827 phhB Down -1.09867 carbinolamine dehydratase 1 flagellar hook-length NP_231759. VC2128 VC2128 Down -1.0984 control protein FliK 1 VCA090 NP_233287. VCA0902 Down -1.09554 hypothetical protein 2 1 NP_230908. VC1263 ribA Down -1.08297 GTP cyclohydrolase II 1 NP_230914. VC1269 VC1269 Down -1.06585 hypothetical protein 1 flagellar basal body rod NP_231827. VC2196 flgF Down -1.06305 protein FlgF 1 integration host factor NP_231548. VC1914 ihfB Down -1.05991 subunit beta 1 NP_231774. VC2143 VC2143 Down -1.05573 flagellin 1 flagellar basal body rod NP_231829. VC2198 flgD Down -1.05407 modification protein 1 VCA080 NP_233195. VCA0809 Down -1.05294 hypothetical protein 9 1 NP_231519. VC1885 VC1885 Down -1.0475 hypothetical protein 1

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flagellar hook-associated NP_231821. VC2190 flgL Down -1.04508 protein FlgL 1 flagellar basal-body rod NP_231831. VC2200 flgB Down -1.04321 protein FlgB 1 sodium-type flagellar NP_230654. VC1008 VC1008 Down -1.03887 protein MotY 1 NP_230078. VC0424 VC0424 Down -1.03848 hypothetical protein 1 NP_229684. VC0025 VC0025 Down -1.0305 hypothetical protein 1 NP_231213. VC1573 fumC Down -1.00813 fumarate hydratase 1 flagellar basal body L-ring NP_231825. VC2194 flgH Down -1.00179 protein 1 NP_230768. VC1123 VC1123 Down -1.00035 hypothetical protein 1 NP_230769. VC1124 VC1124 Down -1.00035 hypothetical protein 1 NP_230770. VC1125 VC1125 Down -1.00035 hypothetical protein 1 NP_229686. VC0027 VC0027 UP 1.01499 threonine dehydratase 1 ATP synthase F0F1 NP_232394. VC2768 VC2768 UP 1.02304 subunit B 1 NP_229958. VC0303 VC0303 UP 1.03064 sensor histidine kinase 1 NP_233131. VCA0744 glpK UP 1.04501 glycerol kinase 1 VCA060 ABC transporter substrate- NP_232992. VCA0603 UP 1.04796 3 binding protein 1 VCA068 NP_233077. VCA0689 UP 1.06273 hypothetical protein 9 1 cytochrome c-type protein NP_231585. VC1951 VC1951 UP 1.06664 YecK 1 NP_230761. VC1116 VC1116 UP 1.15436 hypothetical protein 1 glycerol-3-phosphate NP_233046. VCA0657 glpD UP 1.15619 dehydrogenase 1 NP_231584. VC1950 VC1950 UP 1.16059 biotin sulfoxide reductase 1 formate dehydrogenase NP_231153. VC1512 VC1512 UP 1.16496 iron-sulfur subunit 1 N-acetyl-gamma- NP_232272. VC2644 argC UP 1.17216 glutamyl-phosphate 2 reductase ATP synthase F0F1 NP_232391. VC2765 VC2765 UP 1.21346 subunit gamma 1 VCA069 NP_233079. VCA0691 UP 1.22042 acetoacetyl-CoA reductase 1 1 NP_230778. VC1133 hisD UP 1.2331 histidinol dehydrogenase 1

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tryptophan synthase NP_230814. VC1169 trpA UP 1.2333 subunit alpha 1 imidazole glycerol- phosphate NP_230780. VC1135 VC1135 UP 1.24944 dehydratase/histidinol 1 phosphatase imidazole glycerol NP_230781. VC1136 hisH UP 1.24944 phosphate synthase 1 subunit HisH VCA074 VCA0745 UP 1.25072 5 1-(5-phosphoribosyl-5- [(5- NP_230782. VC1137 VC1137 UP 1.27168 phosphoribosylaminometh 1 ylideneamino imidazole-4- carboxamide isomerase imidazole glycerol NP_230783. VC1138 VC1138 UP 1.27168 phosphate synthase 1 subunit HisF bifunctional phosphoribosyl-AMP NP_230784. VC1139 VC1139 UP 1.27168 cyclohydrolase/phosphori 1 bosyl-ATP pyrophosphatase protein ATP NP_230777. VC1132 hisG UP 1.28659 phosphoribosyltransferase 1 periplasmic nitrate VCA068 NP_233068. VCA0680 UP 1.42642 reductase cytochrome c- 0 1 type protein 5- methyltetrahydropteroyltri NP_231340. VC1704 VC1704 UP 1.45743 glutamate--homocysteine 1 S-methyltransferase glycerol-3-phosphate ABC NP_231191. VC1551 VC1551 UP 1.47283 transporter permease 1 glycerol-3-phosphate NP_231192. VC1552 ugpC UP 1.47283 transporter ATP-binding 1 subunit GntR family NP_230979. VC1335 VC1335 UP 1.47982 transcriptional regulator 1 NP_230980. VC1336 prpB UP 1.47982 2-methylisocitrate lyase 1 ketol-acid NP_229819. VC0162 VC0162 UP 1.4817 reductoisomerase 1 NP_230834. VC1189 VC1189 UP 1.496 hypothetical protein 1 NAD-dependent NP_231150. VC1509 VC1509 UP 1.50893 deacetylase 1 VCA069 acetyl-CoA NP_233078. VCA0690 UP 1.54663 0 acetyltransferase 1 VCA055 gamma- NP_232948. VCA0558 UP 1.57248 8 glutamyltranspeptidase 1

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dihydroxy-acid NP_229687. VC0028 VC0028 UP 1.5972 dehydratase 1 NP_230046. VC0392 VC0392 UP 1.63842 class V aminotransferase 1 acetolactate synthase 2 NP_229690. VC0031 VC0031 UP 1.6756 catalytic subunit 1 periplasmic nitrate VCA067 NP_233067. VCA0679 UP 1.67918 reductase cytochrome c- 9 2 type protein NP_230383. VC0734 VC0734 UP 1.79975 malate synthase 1 glycerol-3-phosphate ABC NP_231189. VC1549 VC1549 UP 1.84038 transporter substrate- 1 binding protein NP_230983. VC1339 VC1339 UP 1.84678 hypothetical protein 1 NP_230984. VC1340 VC1340 UP 1.84678 prpE protein 1 NP_232372. VC2746 glnA UP 1.84823 glutamine synthetase 1 histidinol-phosphate NP_230779. VC1134 VC1134 UP 1.86925 aminotransferase 1 phospho-2-dehydro-3- NP_230344. VC0695 VC0695 UP 1.94228 deoxyheptonate aldolase 1 NP_232270. VC2642 VC2642 UP 1.94676 argininosuccinate synthase 1 NP_232269. VC2641 VC2641 UP 1.95976 argininosuccinate lyase 1 NP_232271. VC2643 VC2643 UP 2.00503 acetylglutamate kinase 1 NP_230981. VC1337 VC1337 UP 2.0271 methylcitrate synthase 1 NP_230385. VC0736 VC0736 UP 2.05361 isocitrate lyase 1 ornithine NP_232137. VC2508 VC2508 UP 2.06797 carbamoyltransferase 1 NP_230982. VC1338 VC1338 UP 2.15753 aconitate hydratase 1 NP_231596. VC1962 VC1962 UP 2.31507 lipoprotein 1 glycerol-3-phosphate ABC NP_231190. VC1550 VC1550 UP 2.32933 transporter permease 1 DNA polymerase III NP_229954. VC0299 VC0299 UP 2.46775 subunit epsilon 1 NP_229955. VC0300 VC0300 UP 2.46775 hypothetical protein 1 NP_229956. VC0301 VC0301 UP 2.46775 hypothetical protein 1 glutamate synthase large NP_232003. VC2373 VC2373 UP 2.53906 subunit 1 NP_231230. VC1590 VC1590 UP 2.58425 acetolactate synthase 1

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short chain NP_231231. VC1591 VC1591 UP 2.58425 dehydrogenase/reductase 1 family oxidoreductase cysteine synthase/cystathionine NP_230706. VC1061 VC1061 UP 2.61109 beta-synthase family 1 protein VCA060 4-aminobutyrate NP_232994. VCA0605 UP 2.6204 5 aminotransferase 1 NP_232332. VC2705 VC2705 UP 2.78889 sodium/solute symporter 2 outer membrane protein NP_230962. VC1318 VC1318 UP 2.97125 OmpV 1 NP_230680. VC1035 VC1035 UP 3.03075 hypothetical protein 1 NP_232331. VC2704 VC2704 UP 3.0332 hypothetical protein 1 VCA060 2-aminoethylphosphonate- NP_232993. VCA0604 UP 3.07887 4 -pyruvate transaminase 1 NP_229953. VC0298 VC0298 UP 3.46211 acetyl-CoA synthetase 2 VC0607 VC0607 UP 3.50493 nitrogen regulatory protein NP_230256. VC0606 VC0606 UP 3.53499 P-II 1 * Up or down- regulation refers to grazed samples vs un-grazed controls

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Appendix 2 List of strain and NCBI reference sequence used in the genomic analysis. Size GC Strain Replicons WGS Scaffolds (Mb) % chromosomeI:NZ_CP014636.1/CP0146 36.1; chromo CECT 5.1631 46.53 someII:NZ_CP014637.1/CP014637.1; - 3 4999 3 plasmid pR99:NZ_CP014638.1/CP014638.1 chromosomeI:NC_005139.1/BA000037. 5.2600 2; chromosome YJ016 46.66 - 3 9 II:NC_005140.1/BA000038.2; plasmid pYJ016:NC_005128.1/AP005352.1 chromosome I:NC_004459.3/AE016795.3; CMCP6 5.1267 46.71 - 2 chromosome II:NC_004460.2/AE016796.2 chromosome MO6- 5.0077 I:NC_014965.1/CP002469.1; 46.95 - 2 24/O 7 chromosome II:NC_014966.1/CP002470.1 Chromosome I:NZ_CP009261.1/CP009261.1; 5.1273 chromosome 93U204 46.70 - 3 5 II:NZ_CP009262.1/CP009262.1; plasmid p93U204:NZ_CP009263.1/CP009263.1 chromosome FORC_00 1:NZ_CP009984.1/CP009984.1; 5.0607 46.74 - 2 9 chromosome 2:NZ_CP009985.1/CP009985.1 chromosome ATL 6- 4.9787 1:NZ_CP014048.1/CP014048.1; 46.84 - 2 1306 8 chromosome 2:NZ_CP014049.1/CP014049.1 chromosome FORC_01 5.0723 I:NZ_CP011775.1/CP011775.1; 46.74 - 2 6 7 chromosome II:NZ_CP011776.1/CP011776.1 chromosome 1:NZ_CP012739.1/CP012739.1; FORC_01 5.2292 46.60 chromosome - 3 7 3 2:NZ_CP012740.1/CP012740.1; plasmid unnamed:NZ_CP012741.1/CP012741.1 chromosome 1:NZ_CP015512.1/CP015512.1; FORC_03 6.0679 45.33 chromosome - 3 6 6 2:NZ_CP015513.1/CP015513.1; plasmid unnamed:NZ_CP015514.1/CP015514.1 chromosome FORC_03 5.1178 I:NZ_CP016321.1/CP016321.1; 46.83 - 3 7 9 chromosome II:NZ_CP016322.1/CP016322.1;

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plasmid unnamed:NZ_CP016323.1/CP016323.1 chromosome ATCC 5.0071 1:NZ_CP012882.1/CP012882.1; 46.70 - 2 27562 6 chromosome 2:NZ_CP012881.1/CP012881.1 4.9503 AFSW0 JY1305 46.7 - 153 3 1 4.9626 AFSX0 E64MW 46.7 - 269 8 1 4.9388 AFSY0 JY1701 46.5 - 329 4 1

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Appendix 3 Whole-genome alignments of V. vulnificus Env1, CMCP6, MO6-24/O and YJ016, JY1305, JY1701 and E64MW were performed using the Progressive Mauve alignment (Darling et al., 2010). Locally collinear block (LCB) are represented by blocks of different colours. The degree of similarity is indicated using white areas where the coloured area is higher where the similarity is high. Areas that are completely white within LCB are not aligned and probably contain sequence elements specific to a particular genome. The lines connecting similar blocks between genomes indicate which regions in each genome are homologous. Inverted regions in the genome are clearly depicted as blocks below a genome’s centre line. The names of the strains are listed at the bottom of the blocks.

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Appendix 4 List of differentially expressed transcripts in Chapter 4.

NCBI RAST Regulation* Log2 p-adj Annotation Locus ID locus ID Fold Change BJD94_166 1246305.4.peg.33 Down - 1.42896E- Non-ribosomal 20 29 10.1067 15 peptide synthetase/ siderophore biosynthesis BJD94_052 1246305.4.peg.11 Down -9.5466 8.0448E- Predicted manganese 45 08 14 transporter BJD94_164 1246305.4.peg.32 Down -8.9943 3.61952E- ABC transporter/ 30 92 12 ATP-binding protein BJD94_080 1246305.4.peg.16 Down -8.7055 1.50551E- Putative hemolysin 55 62 11 BJD94_055 1246305.4.peg.40 Down -8.4725 1.13132E- Hypothetical protein 55 16 10 BJD94_164 1246305.4.peg.32 Down -8.3031 3.24502E- Hypothetical protein 25 91 10 BJD94_121 1246305.4.peg.24 Down -7.9837 1.34087E- Manganese 50 67 09 superoxide dismutase BJD94_166 1246305.4.peg.33 Down -7.7794 8.54233E- 22C3- 40 33 09 dihydroxybenzoate- AMP ligase BJD94_080 1246305.4.peg.16 Down -7.5952 2.2023E- Hypothetical protein 65 64 08 BJD94_166 1246305.4.peg.33 Down -7.4700 1.95008E- Catechol siderophore 70 40 08 ABC transporter BJD94_166 1246305.4.peg.33 Down -7.4664 5.29743E- 22C3- 60 38 08 dihydroxybenzoate- AMP ligase BJD94_166 1246305.4.peg.33 Down -7.3110 4.48898E- Isochorismate 55 37 08 pyruvate-lyase of siderophore biosynthesis BJD94_080 1246305.4.peg.16 Down -7.2332 2.13675E- D-alanine-D-alanine 75 66 10 ligase BJD94_013 1246305.4.peg.28 Down -7.1921 2.32305E- COG0398: 70 3 07 uncharacterized membrane protein BJD94_166 1246305.4.peg.33 Down -7.1501 2.85749E- Aryl carrier domain 65 39 07 BJD94_107 1246305.4.peg.22 Down -7.1271 3.30185E- Putative signal peptide 80 16 07 protein BJD94_080 1246305.4.peg.16 Down -7.0649 7.74971E- Hypothetical protein 60 63 10 BJD94_166 1246305.4.peg.33 Down -6.9406 3.15329E- Isochorismatase of 50 36 07 siderophore biosynthesis BJD94_107 1246305.4.peg.22 Down -6.8989 5.66541E- Uncharacterized 85 17 07 protein conserved in bacteria

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BJD94_217 1246305.4.peg.44 Down -6.8693 1.39545E- TonB system 85 15 06 biopolymer transport component BJD94_080 1246305.4.peg.16 Down -6.8437 5.41365E- Hypothetical protein 70 65 07 BJD94_203 1246305.4.peg.40 Down -6.8229 1.98416E- Arginine/ornithine 05 95 06 antiporter ArcD BJD94_199 1246305.4.peg.40 Down -6.8221 3.15392E- Putative heme iron 40 18 09 utilisation protein BJD94_166 1246305.4.peg.33 Down -6.7275 1.01321E- 2-keto-3-deoxy-D- 25 30 06 arabino- heptulosonate-7- phosphate synthase I alpha BJD94_117 1246305.4.peg.24 Down -6.7161 1.02747E- O-methyltransferase- 55 01 06 related protein BJD94_166 1246305.4.peg.33 Down -6.6692 4.15232E- Vulnibactin utilisation 45 35 06 protein VuuB BJD94_166 1246305.4.peg.33 Down -6.6636 4.17521E- 22C3-dihydro-22C3- 30 31 06 dihydroxybenzoate dehydrogenase BJD94_194 1246305.4.peg.39 Down -6.6546 1.48808E- Arginine/ornithine 10 10 06 antiporter ArcD BJD94_166 1246305.4.peg.33 Down -6.6439 1.37054E- Isochorismate 35 32 08 synthase of siderophore biosynthesis BJD94_199 1246305.4.peg.40 Down -6.5268 9.47576E- Biopolymer transport 20 13 06 protein ExbD1 BJD94_107 1246305.4.peg.22 Down -6.2676 3.38164E- Glutamate synthase 90 18 05 [NADPH] large chain BJD94_057 1246305.4.peg.40 Down -6.2517 9.9435E- Peptide methionine 20 53 116 sulfoxide reductase MsrA/Peptide methionine sulfoxide reductase MsrB BJD94_023 1246305.4.peg.47 Down -6.1505 1.59345E- Hypothetical protein 15 4 05 BJD94_199 1246305.4.peg.40 Down -6.1427 1.1209E- 35 17 24 BJD94_055 1246305.4.peg.40 Down -6.1059 6.88264E- Ferric siderophore 50 15 05 transport system2C periplasmic binding protein TonB BJD94_023 1246305.4.peg.47 Down -6.0162 0.0001136 Hypothetical protein 10 3 13 BJD94_166 1246305.4.peg.33 Down -5.9230 2.97322E- Ferric vulnibactin 75 41 19 receptor VuuA BJD94_018 1246305.4.peg.37 Down -5.8087 1.48838E- Iron-regulated protein 45 7 06 A precursor BJD94_053 1246305.4.peg.11 Down -5.7917 0.0003540 Hypothetical protein 05 22 16

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BJD94_166 1246305.4.peg.33 Down -5.7728 0.0001021 Non-ribosomal 15 28 78 peptide synthetase modules2C siderophore biosynthesis BJD94_196 1246305.4.peg.39 Down -5.5547 0.0008587 TonB-dependent heme 25 54 48 receptor HutR BJD94_196 1246305.4.peg.39 Down -5.5530 0.0009885 Hypothetical protein 35 56 94 in cluster with HutR2C VCA0066 homolog BJD94_013 1246305.4.peg.28 Down -5.5140 0.0010025 Hypothetical protein 65 2 01 BJD94_215 1246305.4.peg.43 Down -5.4895 6.36755E- TonB-dependent heme 10 58 21 and hemoglobin receptor HutA %3B TonB-dependent hemin 2C ferrichrome receptor BJD94_217 1246305.4.peg.44 Down -5.4732 0.0012019 Ferric siderophore 65 11 98 transport system2C periplasmic binding protein TonB BJD94_179 1246305.4.peg.36 Down -5.4654 0.0013187 Peptide ABC 85 19 17 transporter2C ATP- binding protein BJD94_192 1246305.4.peg.38 Down -5.4644 0.0013816 Hypothetical protein 60 79 06 BJD94_158 1246305.4.peg.31 Down -5.4311 5.84446E- Glutaredoxin 40 70 30 BJD94_023 1246305.4.peg.47 Down -5.3859 0.0016065 Heat shock protein 30 7 03 22.5 BJD94_196 1246305.4.peg.39 Down -5.3703 0.0005606 Protease II 20 53 53 BJD94_199 1246305.4.peg.40 Down -5.3504 0.0005485 Periplasmic hemin- 15 12 48 binding protein BJD94_174 1246305.4.peg.35 Down -5.2842 0.0006876 Ferric aerobactin ABC 60 08 83 transporter2C ATPase component BJD94_199 1246305.4.peg.40 Down -5.2804 0.0008580 Ferric siderophore 25 14 95 transport system2C biopolymer transport protein ExbB BJD94_051 1246305.4.peg.10 Down -5.2780 0.0024007 ABC transporter2C 00 78 21 permease protein YnjC BJD94_196 1246305.4.peg.39 Down -5.2121 0.0036165 Hypothetical with 30 55 17 regulatory P domain of a subtilisin-like proprotein convertase BJD94_207 1246305.4.peg.41 Down -5.2041 2.85483E- Hypothetical protein 75 97 07

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BJD94_194 1246305.4.peg.39 Down -5.1804 0.0050982 Hypothetical protein 35 16 19 BJD94_199 1246305.4.peg.40 Down -5.1493 6.04592E- Pyridoxamine 5'- 45 19 18 phosphate oxidase- related putative heme iron utilisation protein BJD94_023 1246305.4.peg.48 Down -5.1419 0.0042817 Flp pilus assembly 50 1 98 protein TadA BJD94_174 1246305.4.peg.35 Down -5.0453 0.0067596 Ferric aerobactin ABC 65 09 8 transporter2C periplasmic substrate binding protein BJD94_080 1246305.4.peg.16 Down -5.0142 0.0001236 Hypothetical protein 80 67 27 BJD94_187 1246305.4.peg.37 Down -4.9985 0.0022104 Transcriptional 40 78 9 regulator2C AraC family BJD94_122 1246305.4.peg.24 Down -4.9427 0.0085998 Hypothetical protein 55 88 19 BJD94_157 1246305.4.peg.31 Down -4.9356 2.7119E- Phosphomannomutase 70 55 114 BJD94_007 1246305.4.peg.16 Down -4.9137 1.30755E- RNA polymerase 85 3 33 sigma-70 factor2C ECF subfamily BJD94_217 1246305.4.peg.44 Down -4.9127 0.0002191 MotA/TolQ/ExbB 80 14 4 proton channel family protein BJD94_196 1246305.4.peg.39 Down -4.9068 0.0096333 Hypothetical protein 40 57 93 in cluster with HutR2C VCA0067 homolog BJD94_107 1246305.4.peg.22 Down -4.8959 0.0128032 INTEGRAL 95 19 76 MEMBRANE PROTEIN (Rhomboid family) BJD94_105 1246305.4.peg.21 Down -4.8918 9.2003E- Glutamine synthetase 40 72 86 type I glnA BJD94_166 1246305.4.peg.33 Down -4.7064 0.0005738 Amide synthase 80 42 97 component of siderophore synthetase BJD94_093 1246305.4.peg.19 Down -4.4658 1.4029E- Uncharacterized 80 31 06 protein DUF547 BJD94_007 1246305.4.peg.16 Down -4.4589 2.81294E- Hypothetical protein 80 2 27 BJD94_141 1246305.4.peg.28 Down -4.4071 5.7733E- Ferric iron ABC 35 16 21 transporter2C iron- binding protein BJD94_061 1246305.4.peg.12 Down -4.3680 0.0022438 Hypothetical protein 75 98 43 BJD94_178 1246305.4.peg.35 Down -4.3277 0.0174520 Hypothetical protein 65 94 91

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BJD94_050 1246305.4.peg.10 Down -4.2040 2.03826E- ABC transporter2C 95 77 09 periplasmic substrate- binding protein YnjB BJD94_204 1246305.4.peg.41 Down -4.1459 0.0305584 Signal recognition 30 21 11 particle GTPase BJD94_069 1246305.4.peg.14 Down -3.9229 0.0010037 Ferrous iron transport 15 42 45 protein C BJD94_213 1246305.4.peg.43 Down -3.6724 0.0228541 Hypothetical protein 20 18 59 BJD94_187 1246305.4.peg.37 Down -3.6262 0.0037744 Ferrichrome-iron 35 77 2 receptor BJD94_217 1246305.4.peg.44 Down -3.5251 0.0087474 TPR domain 60 10 01 protein2C putative component of TonB system BJD94_023 1246305.4.peg.47 Down -3.3449 0.0124686 Flp pilus assembly 35 8 77 protein2C secretin CpaC BJD94_176 1246305.4.peg.35 Down -3.2568 1.86717E- Peptide methionine 05 38 11 sulfoxide reductase MsrB BJD94_044 1246305.4.peg.95 Down -3.1540 1.81919E- Type I secretion 70 4 08 system2C outer membrane component LapE BJD94_053 1246305.4.peg.11 Down -3.0430 0.0108799 Predicted signal 45 30 95 transduction protein BJD94_153 1246305.4.peg.30 Down -3.0391 1.05234E- RTX toxins 50 70 06 determinant A and related Ca2+-binding proteins rtxA BJD94_044 1246305.4.peg.95 Down -3.0026 8.67073E- T1SS secreted 65 3 37 agglutinin RTX BJD94_186 1246305.4.peg.37 Down -2.9442 0.0059386 Hypothetical protein 20 53 99 BJD94_176 1246305.4.peg.35 Down -2.9270 4.09663E- Peptide methionine 10 39 09 sulfoxide reductase MsrA BJD94_115 1246305.4.peg.23 Down -2.8592 4.95511E- Hypothetical protein 10 52 08 BJD94_070 1246305.4.peg.14 Down -2.5698 6.41662E- 65 73 13 BJD94_192 1246305.4.peg.38 Down -2.5684 1.90416E- Uncharacterized 65 81 21 conserved protein BJD94_157 1246305.4.peg.31 Down -2.3333 0.0093729 Outer membrane 25 45 23 protein A precursor BJD94_150 1246305.4.peg.30 Down -2.3094 0.0080750 Methyl-accepting 05 00 5 chemotaxis protein BJD94_126 1246305.4.peg.25 Down -2.3039 2.4911E- Bacterioferritin- 15 48 07 associated ferredoxin

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BJD94_192 1246305.4.peg.38 Down -2.3026 0.0024903 Chemotactic 80 84 84 transducer-related protein BJD94_069 1246305.4.peg.14 Down -2.2854 5.28817E- Ferrous iron transport 20 43 23 protein B BJD94_044 1246305.4.peg.95 Down -2.2767 0.0203347 T1SS peptidoglycan- 75 5 58 associated lipoprotein LapL BJD94_208 1246305.4.peg.42 Down -2.2640 4.56146E- Hypothetical protein 20 06 05 BJD94_025 1246305.4.peg.52 Down -2.2437 1.34087E- Hypothetical protein 35 0 09 BJD94_070 1246305.4.peg.14 Down -2.2347 0.0020064 Asparagine synthetase 85 77 23 [glutamine- hydrolyzing] BJD94_018 1246305.4.peg.37 Down -2.1417 0.0047798 Transcriptional 35 5 81 regulator for fatty acid degradation FadR2C GntR family BJD94_192 1246305.4.peg.38 Down -2.1121 2.69446E- Permease of the 75 83 18 drug/metabolite transporter (DMT) superfamily BJD94_109 1246305.4.peg.22 Down -2.0684 0.0043070 Predicted membrane 70 51 3 protein hemolysin III homolog BJD94_086 1246305.4.peg.17 Down -2.0511 1.44827E- Transcriptional 35 76 08 regulator VpsR BJD94_025 1246305.4.peg.51 Down -2.0381 0.0419598 Nicotinate 15 6 91 phosphoribosyltransfe rase BJD94_195 1246305.4.peg.39 Down -2.0250 0.0039709 Protease-related 15 33 7 protein BJD94_196 1246305.4.peg.39 Down -1.9812 0.0014704 Kinesin-related 00 49 27 protein K4 BJD94_066 1246305.4.peg.13 Down -1.8990 0.0181112 Cell division trigger 20 81 59 factor BJD94_002 1246305.4.peg.48 Down -1.8749 0.0151325 UPF0325 protein 20 44 YaeH BJD94_150 1246305.4.peg.30 Down -1.8569 0.0261202 Glycerol-3-phosphate 25 05 19 transporter BJD94_040 1246305.4.peg.87 Down -1.8119 1.01954E- Ribosome modulation 80 2 69 factor BJD94_005 1246305.4.peg.11 Down -1.7824 1.86717E- Flagellin protein FlaA 45 5 11 BJD94_140 1246305.4.peg.27 Down -1.7379 0.0348734 Dihydrolipoamide 15 92 09 dehydrogenase of pyruvate dehydrogenase complex BJD94_093 1246305.4.peg.19 Down -1.6960 0.0004314 Probable 40 23 transcriptional

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activator for leuABCD operon BJD94_201 1246305.4.peg.40 Down -1.6870 0.0229909 N-acetylglutamate 20 55 81 synthase BJD94_049 1246305.4.peg.10 Down -1.6576 0.0150976 Hypothetical protein 50 48 99 BJD94_053 1246305.4.peg.11 Down -1.6432 0.0261663 DNA-binding protein 00 21 83 inhibitor Id-2-related protein BJD94_071 1246305.4.peg.14 Down -1.6170 0.0069304 Thioredoxin domain- 75 97 5 containing protein EC- YbbN BJD94_198 1246305.4.peg.39 Down -1.5976 0.0208741 Maltose/maltodextrin 00 88 81 ABC transporter2C substrate binding periplasmic protein MalE BJD94_184 1246305.4.peg.37 Down -1.5862 0.0412228 Transcriptional 85 26 39 regulator2C LacI family BJD94_127 1246305.4.peg.25 Down -1.5791 0.0002398 Hydrogen peroxide- 70 79 69 inducible genes activator BJD94_193 1246305.4.peg.38 Down -1.5407 1.3183E- 2-amino-3- 60 99 06 ketobutyrate coenzyme A ligase BJD94_051 1246305.4.peg.10 Down -1.5219 0.0401803 Cystathionine beta- 55 89 53 lyase BJD94_147 1246305.4.peg.29 Down -1.5034 0.0021025 Glycerol kinase 10 32 82 BJD94_115 1246305.4.peg.23 Down -1.4458 3.48432E- Universal stress 20 54 22 protein A BJD94_194 1246305.4.peg.39 Down -1.4114 0.0400210 RTX toxins and 50 20 34 related Ca2+-binding proteins BJD94_049 1246305.4.peg.38 Down -1.3507 0.0010347 ATPase of the AAA+ 10 96 54 class BJD94_111 1246305.4.peg.22 Down -1.3199 0.0472062 ATP synthase protein 05 79 73 I2 BJD94_063 1246305.4.peg.13 Down -1.3113 0.0004144 Predicted metal- 80 39 46 dependent hydrolase with the TIM-barrel fold BJD94_048 1246305.4.peg.10 Down -1.3054 0.0491303 NAD-dependent malic 60 30 5 enzyme BJD94_074 1246305.4.peg.15 Down -1.2438 0.0013800 UDP-sugar hydrolase 35 48 35 %3B 5'-nucleotidase BJD94_065 1246305.4.peg.13 Down -1.2166 0.0036138 Peptidyl-prolyl cis- 95 76 82 trans isomerase PpiD BJD94_071 1246305.4.peg.14 Down -1.1132 0.0354037 Chaperone protein 15 84 HtpG

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BJD94_040 1246305.4.peg.87 Down -1.1131 0.0209486 ABC transporter ATP- 90 4 84 binding protein uup BJD94_041 1246305.4.peg.88 Down -1.1044 0.0035730 Membrane alanine 20 1 91 aminopeptidase N BJD94_047 1246305.4.peg.10 Down -1.0831 0.0024223 10 01 47 BJD94_217 1246305.4.peg.44 Down -1.0628 0.0007897 C4-dicarboxylate 55 09 31 transporter DcuB BJD94_021 1246305.4.peg.43 Down -1.0573 0.0014764 radical activating 05 0 69 enzyme BJD94_005 1246305.4.peg.11 Down -1.0053 0.0021377 Flagellin protein FlaG 50 6 33 BJD94_043 1246305.4.peg.92 Up 1.0058 5.2881E- Hypothetical protein 05 0 06 BJD94_098 1246305.4.peg.20 Up 1.0203 0.0282601 LSU ribosomal 20 22 44 protein L5p (L11e) BJD94_101 1246305.4.peg.20 Up 1.0319 0.0023523 UDP-N- 15 86 63 acetylglucosamine 42C6-dehydratase BJD94_073 1246305.4.peg.15 Up 1.0439 0.0020586 Flagellar basal-body 05 24 66 rod protein FlgB BJD94_113 1246305.4.peg.23 Up 1.0452 0.0369803 Ketol-acid 20 18 06 reductoisomerase BJD94_054 1246305.4.peg.11 Up 1.0523 0.0096550 DNA-binding protein 00 41 1 H-NS BJD94_094 1246305.4.peg.19 Up 1.1004 0.0318936 LSU ribosomal 45 44 06 protein L21p BJD94_073 1246305.4.peg.15 Up 1.1358 0.0165942 Flagellar basal-body 00 23 71 rod protein FlgB BJD94_203 1246305.4.peg.41 Up 1.1533 0.0106798 Predicted 50 04 46 phosphatases BJD94_105 1246305.4.peg.21 Up 1.1724 0.0002627 Phosphoenolpyruvate 00 64 73 carboxykinase [ATP] BJD94_162 1246305.4.peg.32 Up 1.2085 0.0010037 UPF0265 protein 00 46 45 YeeX BJD94_184 1246305.4.peg.37 Up 1.2118 1.41737E- Glutathione S- 05 10 05 transferase BJD94_131 1246305.4.peg.26 Up 1.2471 0.0003497 Predicted ATPase 75 47 91 related to phosphate starvation-inducible protein PhoH BJD94_093 1246305.4.peg.19 Up 1.2669 0.0011528 Pyruvate kinase 10 16 45 BJD94_072 1246305.4.peg.15 Up 1.2747 1.06107E- Phosphocarrier protein 20 06 05 of PTS system BJD94_006 1246305.4.peg.13 Up 1.2909 0.0066497 Flagellar biosynthesis 65 9 97 protein FlhF BJD94_095 1246305.4.peg.19 Up 1.3031 0.0279777 Putative alpha helix 85 73 88 protein BJD94_137 1246305.4.peg.27 Up 1.3450 0.0002797 Enolase 55 48 23

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BJD94_121 1246305.4.peg.24 Up 1.3475 0.0008423 Uncharacterized low- 95 76 19 complexity protein BJD94_025 1246305.4.peg.51 Up 1.3708 1.44827E- Integration host factor 30 9 08 alpha subunit BJD94_122 1246305.4.peg.24 Up 1.3784 6.15573E- 22C3- 90 95 10 bisphosphoglycerate- independent phosphoglycerate mutase BJD94_126 1246305.4.peg.25 Up 1.3790 1.12055E- Triosephosphate 30 51 07 isomerase BJD94_010 1246305.4.peg.21 Up 1.3861 7.86364E- Uridine phosphorylase 55 9 10 BJD94_005 1246305.4.peg.12 Up 1.4271 0.0013070 Flagellar regulatory 80 2 28 protein FleQ BJD94_137 1246305.4.peg.27 Up 1.4471 5.77629E- CTP synthase 50 47 05 BJD94_039 1246305.4.peg.85 Up 1.4620 5.32453E- Universal stress 85 3 05 protein E BJD94_073 1246305.4.peg.15 Up 1.4936 9.95076E- 75 38 15 BJD94_054 1246305.4.peg.11 Up 1.5061 0.0005975 Putative lipoprotein 30 47 88 precursor BJD94_017 1246305.4.peg.36 Up 1.5263 4.1998E- Acetate kinase 80 4 06 BJD94_093 1246305.4.peg.19 Up 1.5323 0.0321348 2-isopropylmalate 60 27 66 synthase BJD94_073 1246305.4.peg.15 Up 1.5529 2.55263E- CopG protein 80 39 07 BJD94_068 1246305.4.peg.14 Up 1.5840 0.0060163 UPF0325 protein 75 34 69 YaeH BJD94_076 1246305.4.peg.15 Up 1.5906 0.0005992 Lipoate synthase 60 79 49 BJD94_122 1246305.4.peg.24 Up 1.5948 4.9026E- Succinate 35 84 06 dehydrogenase flavoprotein subunit BJD94_127 1246305.4.peg.25 Up 1.6070 0.0138129 LSU ribosomal 00 65 86 protein L31p @ LSU ribosomal protein L31p2C zinc- dependent BJD94_109 1246305.4.peg.22 Up 1.6471 0.0037744 Protein yihD 80 54 2 BJD94_005 1246305.4.peg.12 Up 1.6941 0.0241955 Flagellar sensor 75 1 65 histidine kinase FleS BJD94_062 1246305.4.peg.13 Up 1.6943 1.14241E- Alcohol 20 07 36 dehydrogenase %3B Acetaldehyde dehydrogenase BJD94_124 1246305.4.peg.25 Up 1.7045 5.47861E- 3'-to-5' 50 16 18 exoribonuclease RNase R

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BJD94_127 1246305.4.peg.25 Up 1.7414 0.0288692 Transcriptional 30 71 67 regulator2C PadR family aphA BJD94_126 1246305.4.peg.25 Up 1.7484 0.0001001 Ribonuclease E 65 58 46 inhibitor RraA BJD94_121 1246305.4.peg.24 Up 1.8042 2.37449E- 6-phosphofructokinase 75 72 07 BJD94_147 1246305.4.peg.29 Up 1.8057 0.0326693 22C32C42C5- 20 34 69 tetrahydropyridine- 22C6-dicarboxylate N-succinyltransferase BJD94_087 1246305.4.peg.17 Up 1.8356 2.71518E- Anaerobic erobic 50 99 08 respiration control protein arcA BJD94_112 1246305.4.peg.23 Up 1.8437 0.0005700 NADH dehydrogenase 25 04 9 subunit II-related protein BJD94_001 1246305.4.peg.24 Up 1.9029 0.0016516 Na(+)-translocating 00 24 NADH-quinone reductase subunit F BJD94_008 1246305.4.peg.16 Up 1.9081 0.0005813 Phosphohistidine 05 8 57 phosphatase SixA BJD94_010 1246305.4.peg.21 Up 1.9129 0.0163185 Lead2C cadmium2C 50 8 51 zinc and mercury transporting ATPase %3B Copper- translocating P-type ATPase copA BJD94_128 1246305.4.peg.26 Up 1.9217 2.12475E- Glucose-6-phosphate 75 00 08 isomerase BJD94_188 1246305.4.peg.38 Up 1.9220 0.0009790 NAD-dependent 45 00 89 formate dehydrogenase alpha subunit fdhA BJD94_050 1246305.4.peg.10 Up 1.9572 6.51252E- Heat shock protein 90 76 13 HslJ BJD94_021 1246305.4.peg.44 Up 1.9610 2.63345E- Hypothetical protein 65 3 05 BJD94_214 1246305.4.peg.43 Up 1.9830 0.0177407 Uncharacterized 60 48 17 paraquat-inducible protein B BJD94_165 1246305.4.peg.33 Up 2.0361 3.50367E- Lipoprotein 40 13 09 BJD94_047 1246305.4.peg.10 Up 2.1130 0.0163185 Peptidase2C M20A 05 00 51 family BJD94_083 1246305.4.peg.17 Up 2.1216 5.50076E- Phosphate regulon 70 25 06 transcriptional regulatory protein PhoB (SphR) BJD94_098 1246305.4.peg.20 Up 2.1778 2.92628E- LSU ribosomal 30 24 05 protein L14p (L23e) BJD94_176 1246305.4.peg.35 Up 2.1821 5.06166E- Hypothetical protein 40 69 10 214 | P a g e

BJD94_122 1246305.4.peg.24 Up 2.2123 0.0418079 Transcriptional 50 87 39 regulator TetR family BJD94_078 1246305.4.peg.16 Up 2.2190 0.0005738 Aminoacyl-histidine 95 31 97 dipeptidase (Peptidase D) BJD94_068 1246305.4.peg.14 Up 2.2302 3.06612E- Starvation lipoprotein 70 33 05 Slp paralog BJD94_054 1246305.4.peg.11 Up 2.2395 7.2733E- Hypothetical 35 48 11 periplasmic protein BJD94_054 1246305.4.peg.11 Up 2.2587 4.13233E- Hypothetical protein 40 49 06 BJD94_046 1246305.4.peg.98 Up 2.2792 9.69999E- Catalase / Peroxidase 35 8 27 BJD94_086 1246305.4.peg.17 Up 2.2816 0.0010884 UPF0246 protein 85 86 68 YaaA BJD94_074 1246305.4.peg.15 Up 2.2856 0.0261663 Ribosomal large 15 44 83 subunit pseudouridine synthase F BJD94_036 1246305.4.peg.77 Up 2.2883 0.0005204 Hypothetical protein 00 1 06 BJD94_052 1246305.4.peg.11 Up 2.2935 6.15573E- Trimethylamine-N- 70 13 10 oxide reductase BJD94_113 1246305.4.peg.23 Up 2.2995 0.0104787 ATP-dependent DNA 45 23 2 helicase Rep BJD94_209 1246305.4.peg.42 Up 2.3910 0.0314180 Acetate kinase 25 30 56 BJD94_163 1246305.4.peg.32 Up 2.4207 0.0016353 Hypothetical protein 15 69 38 BJD94_158 1246305.4.peg.31 Up 2.4581 4.17521E- Hypothetical protein 40 71 06 BJD94_054 1246305.4.peg.11 Up 2.5368 2.54142E- Hypothetical protein 50 51 07 BJD94_079 1246305.4.peg.16 Up 2.6072 0.0013800 S-formylglutathione 45 42 35 hydrolase fghA BJD94_096 1246305.4.peg.19 Up 2.6623 6.42961E- Hypothetical protein 25 81 08 BJD94_054 1246305.4.peg.11 Up 2.6986 5.16867E- Cyclopropane-fatty- 45 50 15 acyl-phospholipid synthase cdfA BJD94_193 1246305.4.peg.38 Up 2.7082 0.0026223 Catalase 10 90 77 BJD94_054 1246305.4.peg.11 Up 2.7183 1.68364E- Amine oxidase 55 52 14 BJD94_021 1246305.4.peg.44 Up 2.7681 1.8523E- Response regulator 55 1 05 containing a CheY- like receiver domain and an HD-GYP domain BJD94_054 1246305.4.peg.11 Up 2.7943 2.71693E- Putative 65 54 18 transcriptional regulator

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BJD94_170 1246305.4.peg.34 Up 2.9186 0.0045961 Membrane fusion 70 24 65 protein of RND family multidrug efflux pump BJD94_052 1246305.4.peg.11 Up 2.9192 0.0022438 Cytochrome c-type 75 14 43 protein NapC BJD94_068 1246305.4.peg.14 Up 2.9350 4.14846E- Hypothetical protein 90 37 11 BJD94_094 1246305.4.peg.19 Up 2.9439 7.16913E- TRAP-type 70 50 05 uncharacterized transport system2C fused permease component BJD94_198 1246305.4.peg.39 Up 2.9512 0.0401803 Permease of the 25 93 53 drug/metabolite transporter (DMT) superfamily BJD94_157 1246305.4.peg.31 Up 3.1029 0.0400210 AzlC family protein 90 59 34 :branched-chain amino acid ABC transporter permease BJD94_005 1246305.4.peg.10 Up 3.1711 0.0025817 Hypothetical protein 10 8 83 BJD94_087 1246305.4.peg.17 Up 3.2151 0.0072678 Hypothetical protein 00 89 63 :Hemerythrin-like; BJD94_154 1246305.4.peg.30 Up 3.3385 6.21123E- Alkyl hydroperoxide 60 92 36 reductase protein C ahpC BJD94_150 1246305.4.peg.30 Up 3.5370 0.0454705 Anaerobic C4- 50 10 19 dicarboxylate transporter DcuC BJD94_054 1246305.4.peg.11 Up 3.5701 4.39006E- short-chain 60 53 08 dehydrogenase/reduct ase family BJD94_032 1246305.4.peg.69 Up 3.6379 0.0314559 Hypothetical protein 80 7 99 BJD94_086 1246305.4.peg.17 Up 3.6787 0.0039640 Hypothetical protein 30 74 77 purA BJD94_163 1246305.4.peg.32 Up 3.7405 0.0005204 Adenylosuccinate 25 71 06 synthetase BJD94_087 1246305.4.peg.17 Up 3.8163 3.6532E- Pyruvate formate- 15 92 108 lyase tdcE or grcA BJD94_079 1246305.4.peg.16 Up 4.2150 5.30061E- S-(hydroxymethyl) 40 41 33 glutathione dehydrogenase frmA BJD94_012 1246305.4.peg.25 Up 4.2489 0 Pyruvate formate- 15 1 lyase BJD94_116 1246305.4.peg.23 Up 4.7018 0.0070724 Hypothetical protein : 80 86 53 BJD94_054 1246305.4.peg.11 Up 4.7289 0.0006470 Biotin synthase bioB 90 60 49 BJD94_054 1246305.4.peg.11 Up 4.7604 0.0059386 Biotin synthesis 80 58 99 protein BioC bioC

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BJD94_005 1246305.4.peg.11 Up 5.2078 0.0012895 Predicted dye- 30 2 31 decolorizing peroxidase (DyP) subgroup tyrA BJD94_156 1246305.4.peg.31 Up 5.2368 0.0010441 Tripeptide 15 22 14 aminopeptidase BJD94_196 1246305.4.peg.39 Up 6.3833 2.54054E- periplasmic 55 60 05 phosphate-binding protein PstS BJD94_085 1246305.4.peg.17 Up 6.5970 1.0274E- Sodium-dependent 80 64 05 phosphate transporter nptA BJD94_054 1246305.4.peg.11 Up 6.6448 7.66124E- Dethiobiotin 75 57 06 synthetase bioD BJD94_185 1246305.4.peg.37 Up 6.6752 1.52348E- Beta-glucosidase 80 45 06 BJD94_051 1246305.4.peg.10 Up 7.4182 5.59611E- Hydroxylamine 20 82 11 reductase hcP BJD94_164 1246305.4.peg.35 Up 7.8656 6.17339E- Hypothetical protein 65 0 09 * Up or down- regulation refers to iron-replete samples vs iron-deplete controls.

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