DETECTION AND CHARACTERIZATION OF

RICKETTSIAE IN WESTERN AUSTRALIA

Helen Clare OWEN, BVMS

This thesis is presented for the degree of Doctor of Philosophy of Murdoch

University, 2007.

I declare that this thesis is my own account of my research and contains as its main content work which has not previously been submitted for a degree at any tertiary

education institution

…………………………………………

Helen Clare OWEN

ii ABSTRACT

The aim of this study was to address the shortfall in current, in-depth knowledge of

Western Australian rickettsiae. Historically, murine typhus had been extensively reported and, more recently, serological studies and a small number of diagnosed cases indicated that spotted fever group rickettsiae were also present in the State, however no attempts had been made to isolate or characterize these rickettsiae.

To facilitate investigation, ectoparasites (principally ) were opportunistically collected from across the State, with an emphasis on native and feral and people. All ectoparasites were screened for rickettsial infection using a polymerase chain reaction incorporating Rickettsia -specific citrate synthase gene ( gltA) primers.

Preliminary sequencing was performed on representative PCR-positive samples from each geographical location, vertebrate host and ectoparasite in order to identify and characterize the infecting rickettsia. Isolation in cell culture and further genotypic characterization was then performed. Finally, a serosurvey and questionnaire were implemented in one of the study areas to determine whether people were being infected with a Rickettsia spp. and whether infection was associated with clinical signs.

Ectoparasite collection produced three genera of ticks ( Ixodes , Amblyomma and

Haemaphysalis ) from native animals, feral pigs and people, primarily from the southwest of Western Australia and Barrow Island in the Pilbara region. Ticks from a number of sources were shown to be infected with rickettsiae by the PCR, including feral pigs, people, bobtail lizards, kangaroos, bandicoots, burrowing bettongs,

iii common brushtail possums and yellow-footed antechinus. Genotypic characterization of positive amplicons from ticks revealed the presence of two novel spotted fever group rickettsiae. Rickettsia gravesii sp. nov ., named in honour of Dr Stephen Graves, was identified extensively throughout the southwest of the State and on Barrow Island in Ixodes , Amblyomma and Haemaphysalis spp. ticks from multiple hosts. Candidatus

“Rickettsia antechini” was detected in Ixodes spp. only from yellow-footed antechinus in Dwellingup. In addition, a novel Bartonella spp. ( Bartonella sp . strain Mu1) was also detected from Acanthopsylla jordani fleas collected from yellow-footed antechinus in Dwellingup.

Rickettsia gravesii sp. nov . is most closely related to the Rickettsia massiliae subgroup of the spotted fever group and to R. rhipicephali in particular. Sequence similarities between this novel species and the subgroup were 99.7%, 98.4%, 95.8% and 97.4% based on its 16S rRNA, gltA , ompA and ompB genes respectively.

Candidatus “Rickettsia antechini” also demonstrated a close relationship to the R. massiliae subgroup (99.4%, 94.8% and 97.1% sequence similarity based on its gltA, ompA and ompB genes respectively). The two novel Western Australian species demonstrated 98.4%, 96.3% and 96.7% sequence similarity to each other based on gltA, ompA and ompB genes respectively indicating separate species. The novel

Bartonella spp. ( Bartonella sp . strain Mu1) detected in fleas collected from yellow- footed antechinus in Dwellingup demonstrated greatest gltA gene sequence similarity to Bartonella strain 40 at 86.1%.

Results from the serosurvey and questionnaire-based investigation into the zoonotic importance of R. gravesii sp. nov . on Barrow Island supported the results of the

iv study and suggested that a tick-borne rickettsia(e) was infecting people on the island.

However, a significant association between seroconversion and a history of symptoms consistent with a rickettsiosis was not found, and it is possible therefore, that R. gravesii sp. nov . produces only asymptomatic infections.

Future work on rickettsiae in Western Australia will involve phenotypic characterization of the novel species, further investigation of their epidemiology and pathogenicity and an ongoing search for additional undiscovered species.

v TABLE OF CONTENTS

Title page ……………………………………………………………………………i

Declaration …………………………………………………………………………..ii

Abstract ……………………………………………………………………………...iii

Table of Contents ……………………………………………………………………vi

List of Tables ……………………………………………………………………….xii

List of Figures ………………………………………………………………………xiii

Acknowledgements …………………………………………………………………xvi

CHAPTER 1. GENERAL INTRODUCTION

1.1 ORDER RICKETTSIALES ………………………………………...... 1

1.1.1 Introduction ……………………………………………………...…………... 1

1.1.2 ……………………………………………………...……………...3

1.1.3 Epidemiology ………………………………………………...……...... 4

1.1.3.1 The association between ectoparasites and rickettsiae... …..………………….6

1.1.3.2 Vertebrate hosts and the effects of rickettsial infection …..……………….....7

1.1.3.3 Public health significance of rickettsial infections ………..…………………..9

1.2 DIAGNOSIS OF RICKETTSIAL INFECTION …………………………....11

1.2.1 Serology ……………………………………………………...……………...11

1.2.2 Culture of rickettsiae ………………………………………….……………...13

1.2.3 PCR detection of rickettsial DNA ……………………………… ………..…14

1.3 CHARACTERIZATION OF NOVEL RICKETTSIAE ……...……………..15

1.3.1 Antigenic and serological characterization ………………………………….15

1.3.2 Molecular-biological assays for identification …………………………….…16

vi 1.4 REVIEW OF RICKETTSIOSES ……………………………………….…...19

1.4.1 Typhus group ……………………………………………………………..…19

1.4.1.1 Epidemic typhus …………………………………………………...... …19

1.4.1.2 Murine/endemic typhus ………………………………………….………...…19

1.4.2 Spotted fever group ……………………………………………...………..…21

1.4.2.1 Rocky Mountain spotted fever ………………………………..…………..…22

1.4.2.2 Mediterranean spotted fever ……………………………………..………….23

1.4.2.3 Queensland tick typhus ……………………………………………..……….23

1.4.2.4 Flinders Island spotted fever ……………………………………..……….…24

1.4.2.5 Rickettsia felis ……………………………………………………………….26

1.4.2.6 “Australian spotted fever” ………………………………………..…………27

1.4.2.7 Unidentified spotted fever group rickettsiae in Australia …………..………28

1.4.2.8 Other pathogenic spotted fever group rickettsiae …………………..………29

1.4.2.9 Rickettsiae of unknown pathogenicity ……………………………..………35

1.4.2.10 Non-pathogenic spotted fever group rickettsiae …………………..………36

1.4.3 Scrub typhus ………………………………………………………………….36

1.4.4 Bartonella spp …………………………………………………………...... 39

1.5 OBJECTIVES …………………………………………………………………42

CHAPTER 2. COLLECTION AND IDENTIFICATION OF ECTOPARASITES

2.1 INTRODUCTION …………………………………………………………...45

2.2 MATERIALS AND METHODS ……………………………………...... 45

2.2.1 Sample collection …………………………………………………….……45

2.2.2 Tick identification …………………………………………………………49

2.3 RESULTS ……………………………………………………………………50

2.4 DISCUSSION ……………………………………………………………...... 54

vii CHAPTER 3. DETECTION OF RICKETTSIAE

3.1 INTRODUCTION ………………………………………………………….….60

3.2 MATERIALS AND METHODS ………………………………………..…..…61

3.2.1 Study design ……………………………………………………………...... 61

3.2.2 Ectoparasite identification …………………………………………………....62

3.2.3 Detection of rickettsial DNA using PCR ………………………………...... 62

3.2.3.1 Extraction of DNA …………………………………………………………62

3.2.3.2 Screening PCR ………………………………………………………...... 62

3.2.3.3 PCR product detection methods …………………………………………..…64

3.2.4 Sequencing ………………………………………………………………….…65

3.2.4.1 Sequencing of gltA and ompA genes ………………………………………...65

3.2.4.2 Sequence analysis ………………………………………………………….…67

3.3 RESULTS ……………………………………………………………………….67

3.3.1 Screening of ectoparasites ……………………………………………….…….67

3.3.2 Prevalence ……………………………………………………………………..69

3.3.3 Sequence similarities ……………………………………………………….…69

3.4 DISCUSSION …………………………………………………………………..72

3.4.1 Aetiology ……………………………………………………………………...72

3.4.2 Prevalence …………………………………………………………………..…73

3.4.3 Life cycle: vectors and vertebrate hosts ………………………………...... 75

3.4.4 Geographical distribution ……………………………………………………...78

3.4.5 Zoonotic potential ……………………………………………………………..80

CHAPTER 4. CHARACTERIZATION OF RICKETTSIA spp . STRAIN BWI-1

4.1 INTRODUCTION …………………………………………………………….…82

4.2 MATERIALS AND METHODS ……………………………………………….84

viii 4.2.1 Sample collection ……………………………………………………………..84

4.2.2 Cell culture ………………………………………………………………...... 84

4.2.3 Immunofluorescence assay ………………………………………………...... 85

4.2.4 Sequencing ………………………………………………………………...... 86

4.2.5 Sequence analysis …………………………………………………………….88

4.3 RESULTS ………………………………………………………………………89

4.3.1 Sequence analysis …………………………………………………………….89

4.4 DISCUSSION ………………………………………………………………….97

CHAPTER 5. DETECTION AND CHARACTERIZATION OF NOVEL

RICKETTSIAE FROM YELLOW-FOOTED ANTECHINUS.

5.1 INTRODUCTION ………………………………………………………...... 102

5.2 MATERIALS AND METHODS …………………………………………...... 103

5.2.1 Sample collection ……………………………………………………...……..103

5.2.2 Ectoparasite identification ……………………………………………..…….103

5.2.3 Cell culture ……………………………………………………………...……103

5.2.4 DNA extraction …………………………………………………..……...... …103

5.2.5 PCR screening ………………………………………………………………..104

5.2.6 Sequencing …………………………………………………………………...104

5.2.7 Sequence analysis …………………………………………………………….105

5.3 RESULTS ………………………………………………………………………105

5.3.1 Prevalence ……………………………………………………………………105

5.3.2 Sequence similarities ……………………………………………………...….107

5.4 DISCUSSION ……………………………………………………………….....115

5.4.1 Aetiology ………………………………………………………………...... 115

5.4.2 Ecology ………………………………………………………………………..117

ix CHAPTER 6. INVESTIGATING THE PATHOGENIC POTENTIAL OF

RICKETTSIA GRAVESII AND THE RISK FACTORS FOR ZOONOTIC

INFECTION.

6.1 INTRODUCTION ………………………………………………………………..120

6.2 MATERIALS AND METHODS ………………………………………………...121

6.2.1 Study design ……………………………………………………………………121

6.2.2 Questionnaire design …………………………………………………………..122

6.2.3 Questionnaire administration …………………………………………………..122

6.2.4 Blood collection ………………………………………………………………..122

6.2.5 Analysis ………………………………………………………………………..123

6.3 RESULTS ……………………………………………………………………….124

6.3.1 Descriptive epidemiology ……………………………………………………..124

6.3.2 Analysis of questionnaire results …………………………………….………..127

6.4 DISCUSSION ………………………………………………………………….. 129

6.4.1 Review of major findings …………………………………………. ………….129

6.4.2 Limitations of the study …………………………………………….………… 131

6.4.3 Public health implications of the study ………………………….…………….133

6.4.4 Recommendations for further research ………………………………………..134

CHAPTER 7. GENERAL DISCUSSION

7.1 INTRODUCTION ……………………………………………………...……… 136

7.2 FINDINGS OF THE STUDY IN RELATION TO AIMS …….. ……………...... 136

7.3 FUTURE DIRECTIONS ………………………………………………………...141

REFERENCES ...... ……………………………………………...145

APPENDICES ………………………………………………………………………190

APPENDIX 1. ABREVIATIONS ………………………………………………….. 190

x APPENDIX 2. SAMPLE COLLECTION INSTRUCTIONS ……………………..192

APPENDIX 3. COMMON AND SCIENTIFIC NAMES …………………………193

APPENDIX 4. GENBANK ACCESSION NUMBERS …………………………...194

APPENDIX 5. GENBANK ACCESSION NUMBERS AND SEQUENCES OF THE

NOVEL STRAINS ……………………………………………………………….. 196

APPENDIX 6. MINIMUM EVOLUTION AND MAXIMUM PARSIMONY TREES

………………………………………………………………………………………199

APPENDIX 7. QUESTIONNAIRE ………………………………………………. 217

APPENDIX 8. RESULTS OF THE SEROSURVEY ……………………...………235

xi LIST OF TABLES

Table 2.1. Hosts from which tick species were collected in Western Australia, the collection sites and the numbers of animals on which ticks were found….. ……...…51

Table 2.2 Tick species reported in Western Australia …………………………….....55

Table 3.1. Tick samples collected from different hosts and locations in Western

Australia and their screening PCR status …………………..…………..…………….68

Table 3.2. Percentage gltA sequence similarity between the two strains identified in this study and those of previously characterised Rickettsia spp ……………………. 70

Table 3.3. Percentage ompA sequence similarity between the two strains identified in this study and those of previously characterised Rickettsia spp. ……………………71

Table 4.1. . Primers used for PCR and sequencing of DNA from Ricketssia spp. strain

BWI-1. ……………………………………………………………………………….87

Table 4.2 Conditions and the premix used for the PCR reactions amplifying Rickettsia spp. BWI-1 DNA ,,,,,,…………………………………………………………..……88

Table 4.3. PCR cycling conditions used in all reactions …………………………….88

Table 4.5 The percentage similarity between Rickettsia sp. strain BWI-1 and the previously characterized rickettsial species …………………………………………91

Table 5.1 Results of PCR screening of ticks and fleas collected from yellow-footed antechinuses in Dwellingup, WA…. ………………………………………………..106

Table 5.2. Comparison of Rickettsia sp. strain D-1 and previously characterized species of Rickettsia based on 3 genes ……………………………………….….....109

Table 5.3. Comparison of Bartonella sp. strain Mu1 and previously characterized species of Bartonella based on gltA gene sequences …………………………...….110

xii Table 6.1 Description of the population of Barrow Island workers that responded to the questionnaire ……………………….……………………………………………126

Table 6.2 Summary of odds ratios for exposure variables versus rickettsial serology

………………………………………………………………………………………128

Table A.1 Rickettsial Genbank accession numbers ………………………………..194

Table A.2 Bartonella sp. GenBank accession numbers …………………………...195

Table A.3 Titration results for the serology performed on respondents to the Barrow Island questionnaire…………………………………………………………………235

LIST OF FIGURES

Figure 1.1 Composite diagram of the life cycle of Rocky Mountain spotted fever,

Rickettsialpox, and murine typhus ………………………………………………...... 5

Figure 2.1. Native trapping on Barrow Island ……………………………….47

Figure 2.2. Tick on the ear of a burrowing bettong …………………………….……47

Figure 2.3 Map of Western Australia showing sample collection sites ..…………….48

Figure 2.4 Map of southwest WA showing sample collection sites .………………..49

Figure 2.5. Dorsal view of Amblyomma triguttatum collected from a person on

Barrow Island ……………………………………………………………………...... 52

Figure 2.6. Ventral view of Haemophysalis ratti collected from a golden bandicoot on

Barrow Island ………………………………………………………………………..53

Figure 2.7. Dorsal view of Amblyomma limbatum collected from a common brushtail possum on Barrow Island ……………………………………………………………53

xiii Figure 2.8. Dorsal view of Ixodes antechini collected from a yellow-footed antechinus in Dwellingup ………………………………………………………………………..54

Figure 2.9 A map of Western Australia demonstrating previously recorded distribution of ticks collected in this study ………………………………..…….…..56

Figure 3.1 Distribution of the rickettsial isolates across WA ………………….…....79

Figure 4.1. Unrooted neighbour-joining tree based on 16S rRNA sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterised rickettsiae. ….. ………………………………………………………..92 Figure 4.2. Neighbour-joining tree based on gltA gene sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterised rickettsiae.. 93

Figure 4.3. Neighbour-joining tree based on ompA gene sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterised rickettsiae ..94

Figure 4.4. Neighbour-joining tree based on ompB gene sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterised rickettsiae ..95

Figure 4.5. Neighbour-joining tree based on sca4 sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterised rickettsiae ..96

Figure 5.1. Acanthopsylla jordani, lateral view ……………………………………107

Figure 5.2. Neighbour-joining tree based on gltA sequences demonstrating the relationship between Rickettsia sp. D-1 and previously characterised rickettsiae ….111

Figure 5.3. Neighbour-joining tree based on ompA sequences demonstrating the relationship between Rickettsia sp. D-1 and previously characterised rickettsiae …112

Figure 5.4. Neighbour-joining tree based on ompB sequences demonstrating the relationship between Rickettsia sp. D-1 and previously characterised rickettsiae ….113

Figure 5.5. Neighbour-joining tree of Bartonella spp. based on gltA sequences demonstrating the relationship between Bartonella sp. strain Mu1 and previously characterised Bartonella spp …………………………………………………... …..114

xiv Figure A.1. Minimum-evolution tree based on 16S rRNA sequences ( R. gravesii )

………………………………………………………………………………………199

Figure A.2. Maximum-parsimony tree based on 16S rRNA sequences ( R. gravesii )

…………………………………………………………………………………..…..200

Figure A.3. Minimum-evolution tree based on gltA sequences ( R. gravesii ) ……...201

Figure A.4. Maximum-parsimony tree based on gltA sequences ( R. gravesii ) ……202

Figure A.5. Minimum-evolution tree based on ompA sequences ( R. gravesii ) ……203

Figure A.6. Maximum-parsimony tree based on ompA sequences ( R. gravesii ) …..204

Figure A.7. Minimum-evolution tree based on ompB sequences ( R. gravesii ) …….205

Figure A.8. Maximum-parsimony tree based on ompB sequences ( R. gravesii )...... 206

Figure A.9. Minimum-evolution tree based on sca4 sequences ( R. gravesii ) …….. .207

Figure A.10 Maximum-parsimony tree based on sca4 sequences ( R. gravesii ) …...208

Figure A.11. Minimum-evolution tree based on gltA sequences ( R. antechini ) …...209

Figure A.12. Maximum-parsimony tree based on gltA sequences ( R. antechini ) ….210

Figure A.13. Minimum-evolution tree based on ompA sequences ( R. antechini ) ….211

Figure A.14 Maximum-parsimony tree based on ompA sequences ( R. antechini )…212

Figure A.15 Minimum-evolution tree based on ompB sequences ( R. antechini )...... 213

Figure A.16. Maximum-parsimony tree based on ompB sequences ( R. antechini )

……………………………………………………………………………………...214

Figure A.17 Minimum-evolution tree based on gltA sequences ( Bartonella sp. strain

Mu1 )………………………………………………………………………..………215

Figure A.18 Maximum-parsimony tree based on gltA sequences ( Bartonella sp. strain

Mu1) …………………………………………………………………. ……………216

xv I had a lot of help. Thank you to my principal supervisor, Associate Professor Stan Fenwick for obtaining funding, recruiting collaborators and for his infectious enthusiasm for the project. Thanks also to my Murdoch co-supervisors, Associate Professor Ian

Robertson and Associate Professor Phillip Clark. Ian agreed to help despite being a self-confessed molecular-biology sceptic, his presence on my supervisory panel was a constant source of reassurance and thanks to Phil especially for his perceptive and often amusing editing of my thesis. Thanks also to Dr Angus Cook, School of

Population Health, UWA who guided me closely through the analysis of the questionnaire and Yazid Abdad, my fellow PhD candidate who kindly allowed me to use his serology results in my analysis.

I am also eternally thankful to everyone at The Australian Rickettsial Reference

Laboratory. Thanks to my co-supervisor Dr John Stenos, colleague Dr Nathan

Unsworth and Chelsea Nyugen for their extensive knowledge of all things rickettsial.

John did a fantastic job of interstate supervising and Nathan performed much of the initial culturing of the Western Australia rickettsiae while trying to pass on some of his formidable skills to me. Thank you also to Dr Stephen Graves, the laboratory’s director, who is such a knowledgeable and willing collaborator and who provided such a catchy name for our novel rickettsia. Thank you to Gorgon, in particular Russell Langdon for their interest in the project and for contributing a great deal towards its funding.

Thank you to my wonderful friend Patchara Phuektes who introduced me to the foreign world of the laboratory with unlimited patience. Thank you to Russ Hobbs for helping me identify my ectoparasite samples and also to Dr Andrew Mikosa, Dr Peter

Spencer and Dr Ryan Jeffries who kindly helped me with the phylogenetic component

xvi of my project. Dr Peter Adams also provided a lot of helpful advice. Thank you to Dr

Liam O’Connor, Dr Pierre-Edouard Fournier and Professor Didier Raoult for allowing me to spend time in their laboratories.

Scientists from the Department of Environmental Conservation (formerly CALM) especially Keith Morris and Tony Start, Kanyana Wildlife Rehabilitation Centre,

Damien Cancilla, Roberta Bencini, Harriet Mills and John Drummond were kind enough to provide me with samples and thank you to Rod Armistead, Maggie Lilith and Felicity Donaldson for letting me come on some very entertaining fieldtrips as well.

I’m fairly certain that I couldn’t have gotten through this without all the moral support and welcome distractions provided by my lovely friends Peter Wai’in and Maggie

Lilith. Thank you also for the companionship provided by my office mates: Kim,

Danka, Celia, Jordan and Nico, who usually let me win the war over the air conditioner controls.

Thank you to my extra-curricular friends Karen, Emma, Leigh, Emma and Russell,

Megs and everyone in the UQ veterinary pathology department for not having the first idea about what rickettsiae are and thank you to Judy and Jan for looking after me while I was working in Victoria.

The largest chunk of my gratitude must go to my family for their unfaltering support, especially my Mum who can always lessen the high drama that seems to accompany all my life’s endeavours.

xvii CHAPTER 1

LITERATURE REVIEW

1.1 ORDER RICKETTSIALES

1.1.1 Introduction

Rickettsioses represent some of the oldest recorded infectious diseases. Scrub typhus was described in China in the 3 rd century and it is suspected that epidemic typhus was the cause of a plague described in Athens in the 5 th century (Groves and Harrington,

1994; Raoult and Roux, 1997). More recently, the importance of rickettsiae as emerging pathogens has also been highlighted (Raoult and Roux, 1997).

The genus Rickettsia is named after Howard Ricketts, a medical practitioner, trained in pathology and microbiology, who was pivotal in the early investigation of Rocky

Mountain spotted fever ( Rickettsia rickettsii ) in 1906 and 1907. Ricketts was the first person to identify what would later be classified as rickettsiae, describing them as obligatory intracellular bacteria, a novel category of pathogen at the time. He also developed laboratory methods for detecting the presence of rickettsiae in ecologic and clinical samples and demonstrated the importance of ticks and the role of mammalian hosts in the maintenance of Rickettsia rickettsii in nature (Walker, 2004).

The order Rickettsiales comprises short, rod-shaped, coccobacillary or pleomorphic micro-organisms which are structurally and biochemically similar to Gram-negative bacteria. They are obligate intracellular parasites requiring a host cell cytoplasm to stabilise their unusually permeable cell membranes and to supply co-factors required for

1 metabolism. They are found not within a phagosome or phagolysosome but lie in free contact with the host cell cytoplasm (Winkler, 1990; Munderloh and Kurtti, 1995).

Rickettsiae range in size from 0.8 to 2.0 micrometres in length and 0.3 to 0.5 micrometres in diameter (La Scola and Raoult, 1997). The rickettsial envelope consists of an inner cytoplasmic membrane, a trilaminar cell wall and a microcapsular layer present on the outside of the outer leaflet of the cell wall (Silverman and Wisseman,

1978; Yano et al ., 2004). Internally, rickettsiae have ribosomes and a small genome consisting of a single circular chromosome, but lack a discrete nucleus (Raoult and

Roux, 1997; Yano et al ., 2004). Rickettsiae are able to function with a small amount of genetic material due to their obligate intracellular nature and their reliance on the host cell for many compounds rather than having to synthesise them. This small genome is thought to be the result of a reductive evolutionary process from an ancestral genome of a larger size (Andersson and Andersson, 1999; Harrison and Gerstein, 2002; Renesto et al ., 2005). Rickettsiae replicate by binary fission in the host cell cytoplasm (Yano et al .,

2004).

The precise origin of rickettsiae remains uncertain. Rickettsiae and the eukaryotic cell mitochondria may have evolved from a common ancestor because many DNA sequences are shared (Weisburg et al ., 1985; Emelyanov, 2003, 2003a). An alternative theory suggests that rickettsiae may have evolved from free-living bacteria (Roux et al .,

1997). The “original” rickettsiae are thought to have parasitised invertebrate host cells and when vertebrates appeared on Earth both the invertebrates and rickettsiae subsequently adapted to become parasites on the vertebrate newcomers. This highly successful adaptation allowed cycling of the organisms between invertebrates and

2 vertebrates and accounts for the widespread dispersal of rickettsiae into many ecological systems today (Graves, 1998).

1.1.2 Taxonomy

The original taxonomy of rickettsiae described in Bergey’s Manual of Systematic

Bacteriology (1939) divided the order Rickettsiales into three families: Rickettsiaceae,

Bartonellaceae and Anaplasmataceae. The family Rickettsiaceae was further divided

into the tribes Rickettsieae, Ehrlicheae and Wolbachieae, with the tribe Rickettsieae

being subdivided into three genera: Coxiella, Rickettsia and Rochalimea (La Scola

and Raoult, 1997) .

Molecular taxonomic methods have demonstrated that this classification is not the

most appropriate one. Substantial revision has subsequently occurred and members of

the order are now placed into subgroups of Proteobacteria based on their 16S rRNA

gene sequences. The genus Rickettsia has been placed into the α-1 subgroup and the

genus Rochalimaea has been combined with the genus Bartonella and placed into the

α-2 subgroup (La Scola and Raoult, 1997).

Initially, species within the genus Rickettsia were divided into two biogroups, the

typhus group (TG) and the spotted fever group (SFG). This distinction was made

based on the particular species’ intracellular position and optimal growth temperature

and the cross reaction of sera from an infected patient with somatic antigens of three

strains of Proteus (Raoult and Roux, 1997). More recently however, it has been

suggested that two species, Rickettsia bellii and Rickettsia canadensis, represent a

phylogenetic line that predates the typhus-spotted fever group split. They have

3 subsequently been placed into a third “ancestral group” (Stothard et al ., 1994). At present the typhus group contains the species Rickettsia typhi and Rickettsia prowazekii and the spotted fever group contains R. rickettsii, R. conorii, R. africae, R. sibirica, R. slovaca, R. honei, R. japonica, R. australis, R. akari, R. felis, R. aeschlimannii, R. helvetica, R. massiliae, R. rhipicephali, R. montanensis, R. parkeri and R. peacockii . In addition to these species, there are many isolates that have not yet been fully characterized which may represent new species (Fournier et al ., 2003).

Rickettsia tsutsugamushi has been recognised as being distinct from other members of the genus Rickettsia warranting its own genus (also in the α-1 subgroup of

Proteobacteria ) designated Orientia to reflect the geographic distribution of scrub typhus throughout the Orient (Raoult and Roux, 1997; Graves, 1998) .

Coxiella burnetii has also been shown to be distinct from other rickettsiae. It has been demonstrated that its 16S rRNA sequence shows greater similarity to members of the

γ-subgroup of Proteobacteria, within which it is now placed (La Scola and Raoult,

1997). It is no longer considered a rickettsia by many authorities (Graves, 1998).

Despite this taxonomy, the classification of the order Rickettsiales continues to be reassessed as new data becomes available (Parola et al ., 2005).

1.1.3 Epidemiology

Rickettsial life cycles are complex and often involve a wide range of vectors, vertebrate hosts and modes of transmission (figure 1.1).

4

Figure 1.1. Composite diagram of the life cycle of Rocky Mountain spotted fever, rickettsialpox, and murine typhus. A. life cycle of Rickettsia rickettsii in its tick and mammalian hosts; B. life cycle of Rickettsia akari ; and C. life cycle of Rickettsia typhi (Azad and Beard, 1998).

5 1.1.3.1 The association between ectoparasites and rickettsiae

Ectoparasites play an integral role in rickettsial life-cycles and much of the rickettsiae’s ability to survive and propagate can be attributed to their association with ectoparasites. Rickettsial diseases can be maintained in silent ectoparasite transovarial and transstadial cycles for many years and re-emerge as epidemics when conditions are favourable (Walker and Fishbein, 1991; Azad and Beard, 1998). The ectoparasite acts as a vector, reservoir and/or amplifier and, with a few exceptions (eg.

Dermacentor andersoni infected with R. rickettsii and Pediculus hominis corporis infected with R. prowazekii ), does not appear to be adversely affected by the infection

(Raoult and Roux, 1997; Azad and Beard, 1998; Parola et al ., 2005).

The ecology of the invertebrate host determines many of the epidemiological features of the associated rickettsial infections. Influential ecological factors include favoured environment, host specificity and feeding behaviour of the ectoparasite, and these in turn can affect geographical and seasonal distribution, occurrence of zoonotic disease and the number of vertebrate hosts one ectoparasite can infect. It has been speculated that any rickettsial species found in an ectoparasite capable of biting humans should be considered a potential human pathogen (Raoult and Roux, 1997; Azad and Beard,

1998).

Rickettsiae infect ectoparasites through one of two routes: 1. When the ectoparasite feeds on a rickettsaemic host. 2. Through transovarial transmission from an infected female ectoparasite to her offspring. Rickettsiae initially infect the ectoparasite’s gut, predominantly in the midgut region, before entering the haemolymph of the ectoparasite’s circulatory system and spreading systemically. Rickettsiae infect and

6 multiply in almost all organs of their invertebrate hosts following infection. If the ovaries of the ectoparasite become infected, rickettsiae can be transmitted to the ectoparasite’s offspring. A small proportion of the rickettsiae may persist in the tick’s gut when it moults to form subsequent life stages, described as transstadial transmission. As immature stages of an are often less host-specific than adults, this must be taken into account when considering incidence and host- specificity of the infecting rickettsiae (Rehá ček, 1989; Munderloh and Kurtti, 1995;

Raoult and Roux, 1997).

Transmission to a vertebrate host can occur through inhalation or wound inoculation with infected ectoparasite faeces; however, when rickettsiae infect the ectoparasite’s salivary gland then transmission can also occur via the ectoparasite’s bite. This is the most common route of infection for SFG rickettsiae (Raoult and Roux, 1997; Azad and Beard, 1998).

Ixodid (“hard-bodied ticks”) plus other species of ticks are the arthropod vectors associated with spotted fever group rickettsiae (except the mite-borne R. akari ) and

Coxiella burnetii . Fleas are the vectors of R. typhi , R. felis and Bartonella henselae .

Mites are the vectors of R. akari and Orientia tsutsugamushi and lice are the vectors of R. prowazekii (Raoult and Roux, 1997; Azad and Beard, 1998).

1.1.3.2 Vertebrate hosts and the effects of rickettsial infection

The range of vertebrate hosts that may be infected by a specific rickettsia depends largely on the host-specificity of the ectoparasite vector. For example, the human body louse ( Pediculus hominis corporis ), the vector of R. prowazekii, is very host-

7 specific whereas Amblyomma species, which transmit R. africae, will bite almost any mammal (Kelly et al ., 1994; Raoult and Roux, 1997).

Most vertebrate hosts have a clinically inapparent infection and do not remain rickettsaemic for long periods of time, however even this brief period is usually long enough for them to serve as significant reservoirs of infection for ectoparasites (Kelly et al ., 1992a). Rickettsia prowazekii, the aetiological agent of epidemic typhus, is an example of a rickettsia that causes a clinically apparent infection in its human vertebrate host. Epidemic typhus is the only rickettsiosis in which humans are the natural host, infection causes disease and then the rickettsiae sequester in the person.

The latent infection can be re-activated (Brill-Zinsser disease) and the subsequent rickettsaemia will provide a source of infection for lice (Azad and Beard, 1998).

Rickettsia rickettsii has been reported to produce a disease in its canine host characterized by fever, depression and anorexia, neurological signs, scrotal erythema and rash, ecchymoses, epistaxis and death (Kelly et al ., 1992a; Paddock et al ., 2002).

It has been suggested that R. conorii is also capable of causing disease in dogs, however, artificial infection failed to produce conclusive clinical signs (Kelly et al .,

1992a).

Native and feral animals are of major importance in the epidemiology of many zoonotic diseases and their role in rickettsial life cycles should be taken into account when considering the zoonotic risk, control and prevention of specific diseases

(Rehá ček and Tarasevich, 1991; Webster and Macdonald, 1995; Richards et al ., 1997;

Ibrahim et al ., 1999; Parola et al ., 2003; Kruse et al ., 2004). Rickettsiae are

8 effectively maintained in silent sylvatic cycles, and, as reservoir hosts, wild animals are important in the transmission of infection to domestic animals and humans when the opportunity arises. There is increasing interaction between the domestic and sylvatic life cycles as the human population expands and encroaches on previously undisturbed habitats, making vector-borne zoonotic diseases with wildlife reservoirs increasingly significant as emerging infectious diseases (Kruse et al ., 2004).

Wildlife have been shown to be significant in many rickettsial life cycles. In the

U.S.A. for example, R. felis and R. typhi have been found to be present in

Ctenocephalides felis fleas infecting opossums. The infection in this sylvatic cycle can enter the domestic cat-flea cycle when the opossums begin to scavenge around houses in suburban environments (Sorvillo et al ., 1993; Boostrom et al ., 2002). There is also evidence in the U.S.A. that flying squirrels are able to act as non-human reservoirs of Rickettsia prowazekii, and small mammals such as chipmunks, voles, ground squirrels and rabbits have been identified as reservoirs of Rickettsia rickettsii

(Lane et al ., 1981; Raoult and Roux, 1997; Niebylski et al ., 1999). In Australia, the bandicoot has been identified as a reservoir host in the life cycle of Rickettsia australis and Orientia tsutagamushi (Campbell and Domrow, 1974; Raoult and Roux,

1997).

1.1.3.3 Public health significance of rickettsial infections

Humans are accidental hosts for all rickettsiae other than Rickettsia prowazekii . While infection in the natural vertebrate host is usually inapparent, human infection can result in clinical disease ranging from mild to severe.

9 Members of the family Rickettsiaceae have an affinity for endothelial cells. After an arthropod bite, rickettsiae initially infect an undetermined cell type before spreading via the lymphatic vessels to regional lymph nodes and then to the blood stream. A short period of rickettsaemia follows before they infect and then multiply in the endothelial lining of small arteries, veins and capillaries. Lysis of the host cell membrane follows infection. The resulting vasculitis leads to increased vascular permeability, oedema, hypovolaemia and hypotension (Raoult and Roux, 1997; Díaz-

Montero et al ., 2001; Wilson et al ., 2002).

The clinical consequences of rickettsioses range from asymptomatic infections to severe and fatal disease syndromes. The main clinical signs of acute rickettsial infection are non-specific and influenza-like. Fever, headache and malaise usually occur, with or without a rash. The gastrointestinal system can also be involved, resulting in nausea, vomiting and diarrhoea (La Scola and Raoult, 1997; Parola et al .,

2005). In many instances there is an eschar at the site of the ectoparasite bite which starts as a raised, red papule that becomes a vesicle before it ruptures, the centre then becomes dark purple and then black (Andrew et al ., 1946). The course of the disease is usually 2 to 3 weeks (Raoult and Roux, 1997). Occasionally complications such as respiratory, cardiac, neurological, hepatic and renal abnormalities can result (Chi et al ., 1997; Bernabeu-Wittel et al ., 1998; Araki et al ., 2002; Parola et al ., 2005).

Rickettsioses are often not recognised due to their similar clinical manifestation to many other diseases, including measles, chicken pox, unspecified viral exanthema, dengue fever and leptospirosis. Due to the non-specific clinical signs, the often mild course of the diseases, and their transient nature, rickettsioses often go unrecognised

10 and unreported (Sexton et al ., 1990; Walker and Fishbein, 1991; Azad and Beard,

1998; Graves 1998). It is estimated that undiagnosed cases of murine typhus exceed

reported cases by a factor of four to one (Azad, 1990).

1.2 DIAGNOSIS OF RICKETTSIAL INFECTIONS

There are many serological, molecular and culture-based techniques available for the diagnosis of rickettsial infections. When deciding which particular diagnostic test to use, factors such as specimen type, access to equipment and expertise, sensitivity and specificity required and whether the results will be used for clinical or research purposes must be taken into account.

1.2.1 Serology

Serology is widely used to diagnose rickettsial illness however it has the limitation of

only being useful diagnostically when Rickettsia -specific antibodies can be

demonstrated. In most cases these antibodies are not produced for a week or more

following infection by which time the patient has often recovered or died. There have

also been cases where rickettsioses, definitively diagnosed by other means have

produced no increase in antibody titre during or even after the illness (Fournier et al .,

2005). Another limitation of some serological tests is their lack of specificity due to

the presence of cross-reacting antibodies which occur between biogroups (SFG and

TG), between species and with Legionella and Proteus species. Most of the

serological tests currently used for diagnosis however, are Rickettsia -specific.

Serology is a very useful tool in epidemiological studies however, as antibodies,

particularly IgG, will persist in vertebrate hosts for a long time after infection has

passed (Walker and Dumler, 1995; La Scola and Raoult, 1997).

11

The Weil-Felix serological test was one of the first assays developed for the diagnosis of rickettsial infections. It was used to detect antibodies to various Proteus species which contain antigens that cross-react with members of the genus Rickettsia (except

R. akari ). This test is poorly sensitive and not Rickettsia -specific and it is only useful for a short period of time following infection as it detects principally IgM antibodies.

It is rarely used now as it has been superseded by more sophisticated techniques with the advent of improved serological tests and methods of culturing the micro- organisms (Hechemy et al ., 1979; Raoult and Dasch, 1989; La Scola and Raoult,

1997).

The more recently developed serological tests rely on anti-rickettsial antibody/rickettsial antigen interactions for the identification of current or historical disease. These tests include the complement fixation (CF), indirect haemagglutination and latex agglutination tests, the enzyme linked immunosorbent assay (ELISA), indirect immunofluorescence test (IF) and western blotting. The test of choice for the serodiagnosis of rickettsial infection however is the IF test (La Scola and Raoult,

1997; Graves, 1998). The sensitivity of the IF is 94-100% and the specificity can be up to 100% depending on the cut-off titre selected (Walker and Dumler, 1995; Parola et al ., 2005).

Species-specific monoclonal antibodies and western blot immunoassays have also been used to identify the aetiology of SFG rickettsiae infections in epidemiological surveys and are particularly valuable when more than one pathogenic rickettsiae is

12 known to occur in a region (Raoult and Dasch, 1989; Raoult et al ., 1994; Xu et al .,

1997).

1.2.2 Culture of rickettsiae

Culture is used to isolate rickettsiae from clinical samples. Tick, amphibian and mammalian cell lines such as L-929, Vero, BGM, HeLa, XTC, human embryonic lung and primary chick embryo cells may be used for growing rickettsiae. As rickettsiae cannot grow in the presence of most antibiotics, strict attention must be paid to aseptic technique to avoid contamination with bacteria that will overgrow rickettsiae. Rickettsiae have relatively slow growth rates, with an average doubling time of 10 hours. Species of rickettsiae differ in their growth characteristics, such as optimal growth temperature, time until plaque formation and favoured cell-line, and exhibit a range of cytopathic effects in cell cultures (Baird et al ., 1992; Walker and

Dumler, 1995; Raoult and Roux, 1997; Simser et al ., 2001).

Culturing of most Rickettsia spp. presents a degree of risk to laboratory personnel as infection can be acquired through accidental parenteral inoculation or through exposure to infectious aerosols. In 1976 there were 63 reported cases of laboratory aquired infection with Rickettsia ricketsii , 11 of these were fatal. To minimise the risk of infection, it is generally recommended that culture of Rickettsia spp. is undertaken in physical containment 3 (PC3) facilities (http://www.phac-aspc.gc.ca/msds- ftss/msds129e.html). The development of non-culture techniques is therefore desirable due to these requirements and risk posed to personnel.

13 1.2.3 Polymerase chain reaction (PCR) detection of rickettsial DNA

The use of PCR and its application in the diagnosis of rickettsioses has alleviated the need for fresh or frozen specimens and for difficult and hazardous culture procedures

(Tzianabos et al ., 1989; Azad et al ., 1990; Carl et al ., 1990; Sexton et al ., 1994).

Unlike serology, no reference sera are required (Webb et al ., 1990; Raoult and Roux,

1997); as many rickettsial genes are highly conserved, DNA for positive controls can be extracted from generic continuous cell cultures. PCR, in conjunction with DNA sequencing, is more specific than most other diagnostic tests, however traditional

PCR has low sensitivity when used to detect organisms in commonly collected specimens such as sera and whole blood as the rickettsaemia is transient and there are often low concentrations of circulating organisms. A highly sensitive and specific real-time PCR assay has recently been developed by Stenos et al. (2005). This assay is capable of detecting 1 SFG rickettsia per PCR reaction. Traditional PCR is more sensitive when used for clinical specimens such as cerebrospinal fluid, lymphocytes and skin biopsies (Reynolds et al ., 2003).

PCR assays using primers to detect a number of rickettsial genes have been described. These include the 16S rRNA gene (Wilson et al ., 1990), the 17 kilodalton antigen gene (Anderson and Tzianabos, 1989), the citrate synthase gene gltA (Roux et al ., 1997), genes for outer membrane proteins rOmpB (Sekeyová et al ., 2001) and rOmpA for spotted fever group rickettsiae (Roux et al ., 1996; Bouyer et al ., 2001), the groEL gene which encodes for a 60 kilodalton heat shock protein (Lee et al ., 2003) and sca 4 (formerly ‘gene D’) which encodes for the PS120 protein (Sekeyova et al .,

2001). None of these PCR reactions are species-specific however and PCR products

14 must be analysed further by sequencing of amplicons in order that a particular species can be identified (Raoult and Roux, 1997).

1.3 CHARACTERIZATION OF NOVEL RICKETTSIAE

Due to their obligatory intracellular nature rickettsiae express few of the phenotypic and biochemical characteristics traditionally used for bacterial taxonomy, therefore other methods of characterization must be utilized (Raoult and Roux, 1997). In the past, rickettsial isolates were classified into the SFG, TG or scrub typhus group based on the cross reactions of patient’s sera with somatic antigens of strains of Proteus

(Raoult et al ., 2005). Phenotypic criteria such as arthropod vector, pathogenicity, geographical distribution of the strain, optimal culture temperature, plaque formation, growth in embryonic chicken eggs and haemolytic activity have also been used to characterize rickettsial strains (Raoult et al., 2005).

1.3.1 Antigenic and serological characterization

Until the mid-1980s, micro-immunofluorescence using polyclonal mouse sera and monoclonal antibody serotyping were the reference methods for identifying new rickettsiae (Philip et al., 1978; Raoult et al ., 2005). Comparison of proteins by sodium dodecyl sulphate-polyacrylamide gel electrophoresis (SDS-PAGE) and western immunoblotting were also performed so that a novel isolate’s antigenic profile and phylogenetic position could be examined in relation to previously characterized rickettsiae. These techniques primarily targeted the proteins rOmpA and rOmpB , which contain species-specific epitopes. The assays were often difficult to perform and time-consuming and had the drawback of only being possible in a specialised reference laboratory as physical containment level 3 (PC3) clearance is required for

15 the cultivation of rickettsiae; laboratory animal housing, a large panel of specific antisera and appropriately trained staff are also pre-requisites (Bouyer et al ., 2001;

Parola and Raoult, 2001a). Although the recent advent of molecular taxonomic methods has provided an alternative to serological assays, they can still be used to provide valuable information about rickettsial relationships and novelty (Philip et al .,

1978; Beati et al ., 1992; Walker et al ., 1995; Xu et al ., 1997; Xu and Raoult, 1998).

These methods have also become more accessible recently with the development of new culture techniques providing a reliable source of antigens for the assays

(Eremeeva et al ., 1995; Walker et al ., 1995; La Scola and Raoult, 1997; Fang and

Raoult, 2003).

1.3.2 Molecular-biological assays for identification

Genotypic approaches for identifying rickettsiae are being increasingly employed, characteristically providing more objective and definitive information than serotyping

(Parola et al ., 2005), however criteria for the taxonomic definition of novel rickettsial isolates are still under consideration (Raoult and Roux, 1997).

It has been proposed that rickettsiae experience less evolutionary pressures than most convential bacterial species due to their small genome size and strict intracellular nature; these factors result in a low rate of mutation and the 70% DNA-DNA relatedness cut off used to describe conventional bacterial species does not therefore adequately differentiate between rickettsiae. Using this criterion, all the members of the SFG would be classed as a single species, even though they have different

16 ecologies and produce distinct diseases (Stenos et al ., 1998; Fournier et al ., 2003;

Renesto et al ., 2005).

In 2003, Fournier et al. described a set of objective guidelines for the classification of new rickettsial isolates at various taxonomic levels, including genus, group and species, based on sequences from selected genes. While rickettsial taxonomy remains an evolving and controversial field, their criteria are now widely used.

The panbacterial 16S rRNA gene has been used extensively to differentiate bacterial species however, it demonstrates minimal variation across the range of rickettsial species, so significant inferences about intra-genus phylogeny are not possible

(Weisburg et al ., 1991; Roux and Raoult, 1995). The citrate synthase encoding gene

(gltA ) has been shown to mutate more rapidly than other rickettsial genes and therefore provides a more sensitive tool for analysing phylogenetic relationships

(Roux et al ., 1997). One of the outer membrane protein encoding genes, o mpA, is specific for spotted fever group rickettsiae and is relatively divergent between members of the group (Fournier et al ., 1998). An additional outer membrane protein

(ompB ) (Roux and Raoult, 2000) and sca4 (which codes for the PS-120 protein)

(Sekeyová et al ., 2001) are common to both typhus and spotted fever group rickettsiae and are also used in the taxonomic definition of a novel isolate (La Scola and Raoult,

1997; Fournier et al ., 2003). Analysis of these genes concurs with recommendations made by the Ad Hoc Committee for the Re-evaluation of the Species Definition of

Bacteriology that a minimum of five genes should be sequenced when characterising a new isolate (Fournier et al ., 2003).

17 Fournier et al. (2003) recommend that in order to be classified as a member of the genus Rickettsia an isolate should be 98.1% similar to 16S rRNA genes and 86.5% similar to citrate synthase genes from any of the 20 Rickettsia species studied previously. A member of the typhus group should fulfil at least two of the following four criteria: nucleotide sequence similarities with 16S rRNA, citrate synthase, ompB and sca4 genes of either Rickettsia typhi or Rickettsia prowazekii of ≥ 99.4, ≥ 96.6, ≥

92.4, ≥ 91.6% respectively. A member of the spotted fever group should either possess the ompA gene or fulfil at least two of the following criteria: nucleotide sequence similarities with 16S rRNA, citrate synthase, ompB and sca4 genes of any other member of this group of ≥ 98.8, ≥ 92.7, ≥ 85.8 and 82.2% respectively.

To be classified as a new Rickettsia species, an isolate should not exhibit more than one of the following degrees of nucleotide similarity with the most homologous validated species: ≥ 99.8 and 99.9% for the 16S rRNA and the citrate synthase genes respectively and when amplifiable, ≥ 98.8, 99.2 and 99.3% for the ompA, ompB and sca4 genes respectively. The suitability of this classification scheme was confirmed by the study of isolates in vitro (Fournier et al ., 2003).

In addition, it has recently been proposed that bacterial isolates that do not qualify for separate species status based on this criteria can be considered distinct subspecies if they have differing serotypic and epidemio-clinical characteristics. This proposal applies to members of the R. conorii species: “ R. conorii subsp. conorii subsp. nov.”,

“R. conorii subsp. indica subsp. nov.”, “ R. conorii subsp. caspia subsp. nov” and “ R. conorii subsp. israelensis subsp. nov” (Philip et al ., 1978; Parola et al ., 2005; Zhu et al . 2005).

18

1.4. REVIEW OF RICKETTSIOSES

1.4.1 Typhus group

1.4.1.1 Epidemic typhus

Epidemic typhus ( R. prowazekii ) is most frequently seen when conditions are favourable for the proliferation of its vector, the human body louse. Poverty, lack of hygiene and cold weather stimulates the close contact between people that supports the transmission of epidemic typhus. Currently the disease is limited to North and

South America, Asia and Africa (Raoult and Roux, 1997; Raoult et al ., 1997; Roux and Raoult, 1999).

Epidemic typhus is the only rickettsiosis in which humans are the primary vertebrate host. Once a person has contracted epidemic typhus they may retain a latent infection for life. This can re-activate as Brill-Zinsser disease, a milder form of typhus, under stressful conditions. The onset of typhus is usually acute. Patients present with high fever, headaches and severe myalgia. A rash and pneumonia are also common. It is fatal in 10 to 30% of patients depending on underlying diseases and nutritional status

(Raoult and Roux, 1997). There is evidence that flying squirrels are able to act as non- human reservoirs of epidemic typhus as close contact with these animals or exposure to their nests, dander or ectoparasites has been linked to most cases of sporadic epidemic typhus occurring in the United States (Reynolds et al ., 2003).

1.4.1.2 Murine/endemic typhus

The agent of murine typhus, Rickettsia typhi, is closely related to R. prowazekii but the epidemiology, including mode of transmission and the clinical diseases produced

19 are quite different (Wheatland, 1926; Hone, 1927; Fournier et al ., 2003). In contrast to epidemic typhus, murine typhus is more prevalent in warmer countries and it is generally a milder disease (Raout and Roux, 1997). The natural hosts of R. typhi are rats and mice, although other rodents and even other mammals such as shrews, skunks, opossums and cats have been shown to be capable of acting as vertebrate hosts (Azad, 1990). The natural invertebrate vectors are rodent fleas ( Xenopsylla cheopis, Nosopsylla fasciatus and Leptopsylla segnis ). Rickettsia typhi has also been isolated from the cat flea Ctenocephalides felis (Azad and Beard, 1998) but the significance of this vector in the transmission cycle remains unclear. The rickettsia infects the gut and is excreted in the faeces of the flea, with the most common mode of transmission believed to be by inhalation of infected flea faeces. Infection via wound contamination with flea faeces or through flea bites can also occur (Azad,

1990; Graves, 1998). Murine typhus is most prevalent in warmer countries, with most reports occurring in the USA, Africa, Europe and Asia (Raoult and Roux, 1997).

Dr Frank Hone, a general practitioner and the Chief Quarantine Officer for South

Australia recognised an outbreak of acute “typhus-like illness” in waterside workers in 1922 (Hone, 1922; 1923; 1927). He was the first person to describe murine typhus and the first person to suspect a non-ectoparasite mode of transmission for a typhus- like disease. The causative micro-organism, named Rickettsia mooseri at the time, was later identified by scientists working at the United States Public Health Service

(Burnet, 1942; Stewart, 1991). By the 1930s the association of the disease with rodent contact was established and in 1942 inhalation of dust containing infected flea faeces was suggested as the mode of transmission by FM Burnet (Burnet, 1942; O'Connor et al., 1996).

20

After the initial outbreak in Adelaide described by Dr Hone, Wheatland described a

“typhus-like fever” in farm workers in the Toowoomba district of Queensland where a rodent plague had preceded the outbreak in the human hosts (Wheatland, 1926).

Murine typhus also occurred in Western Australia with 1332 cases notified between

1927 and 1952, resulting in 50 deaths, mainly amongst the elderly. The main foci of infection were Perth and Fremantle although cases occurred as far north as Geraldton and as far south as Albany (Saint et al ., 1954). Another outbreak in the Toowoomba area in Queensland was described between 1948 and 1959, when 44 cases occurred

(Derrick and Pope, 1960; O'Connor et al ., 1996). The geographical range of murine typhus in Australia was recently expanded after a human case was reported from

Geelong, Victoria (Jones et al ., 2004).

Murine typhus is probably endemic Australia-wide and cases are documented occasionally; such as the cluster of six cases occurring in Albany, Western Australia between 1995 and 1996 (Forbes et al ., 1991; Graves et al ., 1992; Kelly et al , 1995;

O'Connor et al ., 1996; Graves, 1998; Beaman and Marinovitch, 1999). However, what was once a common disease is now quite rare, perhaps because of effective rat and mouse control programmes (Saint et al ., 1954; Dwyer et al ., 1991). Failure to recognise and report cases may result in under-recognition of murine typhus as a human disease (Forbes et al ., 1991).

1.4.2 Spotted fever group

The use of new culture techniques and molecular biological tools has led to an increase in the identification of novel SFG rickettsiae over the past 20 years. Spotted

21 fever group rickettsiae are widely distributed, occurring on every continent other than

Antarctica (Parola et al ., 2005).

1.4.2.1 Rocky Mountain spotted fever

Rocky Mountain spotted fever (RMSF) was first reported in 1899 (Parola et al ., 2005) and first described in 1904 in an article published by Wilson and Chowning in the first volume of “The Journal of Infectious Diseases”. The authors were investigating the high incidence of cases of an often fatal “spotted fever” occurring on the western side of the Bitterroot River, Western Montana. In their initial investigation they identified an increased incidence of cases in the spring and a lack of communicability between people. They identified outdoor activities as being a risk factor for infection and hypothesised that transmission might have occurred following tick bites and that

Colombian ground squirrels might have played a role in the life cycle of the infectious agent (Walker, 2004).

Rocky Mountain spotted fever ( R. rickettsii) is believed to be the most clinically severe of the SFG rickettsioses. Without treatment, infection in people can lead to severe systemic manifestations, including pneumonitis, myocarditis, hepatitis, renal failure, encephalitis, gangrene and a case-fatality rate as high as 30%. Even with treatment, hospitalisation rates of 72% and case-fatality rates of 4% have been reported (Levy et al ., 2004). Infections can also be lethal in young and previously healthy patients. Rickettsia rickettsii has been recovered from various anthropophilic

(attracted to humans) tick genera including Amblyomma, Dermacentor and

Rhipicephalus spp., which have a wide range of domestic and wild animal vertebrate hosts. Rickettsia rickettsii is also atypical in that it can produce a clinically apparent

22 infection, characterized by fever, neurological signs, scrotal erythema and rash, in its canine vertebrate host (Paddock et al ., 2002). It occurs in areas of Canada, and North,

Central and South America, where it is also the causative agent of Brazilian spotted fever (Fuentes, 1986; Raoult and Roux, 1997; Galvão et al ., 2003; Horta et al ., 2004;

Schoeler et al ., 2005).

1.4.2.2 Mediterranean Spotted Fever

Mediterranean spotted fever (MSF), caused by R. conorii , is also known as

“boutonneuse” fever and Kenyan tick typhus. This disease occurs all around the

Mediterranean coast, in sub-Saharan Africa, India, around the Black Sea and in the eastern part of Russia. It is more common in summer. Typically, MSF has a similar but milder presentation than Rocky Mountain spotted fever with mortality rates of approximately 2.5%. The disease is most often characterized by a single eschar and a generalized macropapular rash that may involve the palms and the soles.

Rhipicephalus spp. act as the vector and dogs, which are the preferred hosts of this tick species, may also be utilised by the rickettsiae as reservoirs (Tinelli et al ., 1989;

Raoult and Roux, 1997; Antón et al ., 2003; Rutherford et al ., 2004).

1.4.2.3 Queensland tick typhus

Queensland tick typhus (QTT), caused by R. australis , was initially recognised in

1946, with many of the first cases observed among Australian troops training in the bush of northern Queensland (Mathew, 1938; Andrew et al ., 1946; Brody, 1946; Plotz et al ., 1946; Raoult and Roux, 1997). Antibodies were detected in bandicoots and rodents in serological studies of vertebrate reservoirs (Fenner, 1946; Campbell and

23 Domrow, 1974; Raoult and Roux, 1997) and Rickettsia australis was isolated from

Ixodes holocyclus and I. tasmani in 1974 (Campbell and Domrow, 1974).

After the initial identification of the disease in north-eastern Queensland, QTT was found to occur in southern Queensland and New South Wales and subsequently down the eastern coast of Australia to Gippsland in Victoria (Streeten et al ., 1948; Neilson,

1955; Pope, 1955; Knyvett and Sanders, 1964; Campbell et al ., 1979; Dwyer et al .,

1991; Sexton et al ., 1991; 1991a; Graves et al ., 1993). Cases all occur on the eastern side of the Great Dividing Range where rainfall is conducive to the survival of Ixodes spp. ticks (Sexton et al ., 1991; Hudson et al ., 1993; Hudson et al ., 1994; Ash and

Smithurst, 1995; Graves, 1998; Roberts et al ., 2000). It has been estimated that 20 cases a year on average are seen by medical practitioners working in the northern beaches region of Sydney (Russell, 2001) and there have been two fatal cases reported to date (Sexton et al ., 1990; Parola et al ., 2005).

While many spotted fever group rickettsiae are genetically very similar, R. australis is relatively distinct (Regnery et al ., 1991). It has been demonstrated that R. australis is the most evolutionarily distant rickettsia in the SFG, this divergence being especially evident in the amino acid sequence of domain II of the ompA gene (Stenos and

Walker, 2000).

1.4.2.4 Flinders Island spotted fever

Flinders Island spotted fever (FISF), caused by R. honei , was first described in 1991 by Dr Robert Stewart, a general practitioner on the island. He had seen 26 cases of this typical rickettsial disease, which occurred most often in the summer months and

24 which usually presented with a macropapular rash most obvious on the trunk and limbs and more prominent centrally than on the periphery (Stewart, 1991). Initially it was suspected that R. australis was the causative agent but the disease was slightly different in presentation and epidemiology when compared to Queensland tick typhus

(Graves et al ., 1991; Sexton et al ., 1991a; Graves, 1998). The aetiological agent of

Flinders Island spotted fever was confirmed as a new species in 1992 when an isolate was characterised, and it was named Rickettsia honei to honour Frank Sandland Hone an early Australian rickettsiologist (Baird et al ., 1996; Raoult and Roux, 1997; Stenos et al ., 1997).

In 2003, it was proposed that R. honei was the same aetiological agent as the rickettsia that causes Thai Tick Typhus (TTT-118) (Graves and Stenos, 2003). This organism had first been isolated from a pool of Ixodes and Rhipicephalus spp . larval ticks in

Thailand in 1962 and was identified as a human pathogen in 2005 (Kollars et al .,

2001; Jiang et al ., 2005). In 1996, R. honei was also identified in Amblyomma cajennense ticks parasitising cattle in Texas, USA making it the SFG rickettsia with the widest recognized distribution to date (Graves and Stenos, 2003).

The tick Bothriocroton hydrosauri (formerly Aponomma hydrosauri ), which has been associated with reptiles such as blue tongue lizards, copperhead and tiger snakes has been identified as an important vector of R. honei on Flinders Island. This was confirmed after demonstration of the rickettsia in the ticks’ salivary glands, malpighian tubules, midgut epithelial cells and oocytes (Stenos et al ., 2003;

Whitworth et al ., 2003). The range of R. honei was recently extended in Australia, when cases of human disease were described from South Australia, and from

25 Schouten Island, south of the Freycinet Peninsula in Tasmania. Recently a R. honei - like agent was detected in a Haemaphysalis tick collected from the Cape York

Peninsula in far north Queensland (Dyer et al ., 2005; Lane et al ., 2005; Unsworth et al ., 2005).

1.4.2.5 Rickettsia felis

In 1990, when cat fleas ( Ctenocephalides felis ) were being examined for the presence of R. typhi using an electron microscope, a Rickettsia -like organism was observed in the flea’s midgut (Adams et al., 1990). The agent was provisionally named the ELB agent after the EL Laboratory (Soquel, California). After initial detection, the ELB agent was identified in blood samples from opossums and their fleas in an area in

California where a rickettsia-like disease of humans had occurred (Williams et al .,

1992). In 1996, the ELB agent was renamed Rickettsia felis and it has subsequently been identified as the cause of human disease in a number of locations including:

North and South America, Europe, Africa, New Zealand and along the Thai-Myanmar border (Schriefer et al ., 1994; Higgins et al ., 1996; Zavala-Velazquez et al ., 2000;

Bouyer et al ., 2001; Raoult et al ., 2001; La Scola et al ., 2002a; Richter et al ., 2002;

Parola et al ., 2003; Kelly et al ., 2004).

Rickettsia felis was initially placed within the typhus group of Rickettsia as it reacts with antibodies of R. typhi and due to the historical association of members of the TG with insects and SFG with acarines. However, data from more recent genetic and antigenic studies, notably the presence of an ompA gene, place R. felis in the SFG

(Bouyer et al ., 2001; Fang and Raoult, 2003). Further analysis of the 16S rRNA gene placed R. felis in a clade with R. akari and R. australis. Assessment of the 17 kDa

26 protein demonstrated that R. felis is 11% divergent from the TG which corresponds with the 10-12% divergence recognised previously between members of the SFG and the TG (Bouyer et al ., 2001). Rickettsia felis has been shown to be transmitted horizontally and vertically in fleas (Fang and Raoult, 2003). In 2003, R. felis was also detected in the ticks Haemaphysalis flava, H. kitasatoe and Ixodes ovatus in Japan.

However, the significance of ticks in the life cycle of R. felis is unknown at this stage

(Ishikura et al ., 2003). The disease produced by R. felis is usually mild and similar to murine typhus (Schoeler et al ., 2005).

Following the outbreak of murine typhus cases in Albany, Western Australia in the

1990s, a survey was conducted using human serum from the south-west of Western

Australia and fleas from rats in Albany and from cats in Perth. No R. typhi DNA was detected from the fleas but 10 out of 37 pools of fleas from the cats in Perth were

PCR/RFLP positive for R. felis infection, however, sequencing for confirmation of the identification was not performed (Kilminster T. An Investigation of Typhus in

Western Australia. Honours Thesis: Department of Microbiology, University of

Western Australia; 1997). In 2004, R. felis was sequenced from flea samples collected from various locations across Western Australia, providing definitive evidence for its existence here (Schloderer et al., 2006). Rickettsia felis has not yet been recorded from a human patient in Australia.

1.4.2.6 “Australian Spotted Fever”

Recently, seven cases of a SFG rickettsiosis caused by infection with the newly- recognised rickettsia “ R. marmionii ” have been diagnosed from Queensland,

Tasmania and South Australia. Genotypic studies have demonstrated a close

27 relationship between R. marmionii and R. honei . Potential vectors include

Haemaphysalis novaeguineae (Graves et al., 2006).

1.4.2.7 Unidentified spotted fever group rickettsiae in Australia

Further evidence is available suggesting that there are other, as yet unidentified SFG rickettsiae present in Australia. For example, a mild SFG rickettsiosis of people has been recognised in northern Tasmania. This has been diagnosed serologically but no isolation and characterization has occurred to date (Chin and Jennens, 1995; Graves,

1998). In addition, a survey performed in 1997 on serum samples collected from the south west of Western Australia demonstrated that 42 out of 866 samples were positive for SFG antibodies (Kilminster T. An Investigation of Typhus in Western

Australia. Honours Thesis: Department of Microbiology, University of Western

Australia; 1997). In 1999, a serosurvey performed on 920 non-randomly collected serum samples from the Kimberley region of Western Australia (WA) demonstrated that 15 out of 920 samples tested were positive for spotted fever group antibodies

(Graves et al., 1999).

Rickettsial infections have also been reported in both people travelling into Australia and Australians travelling abroad. In 1988, two cases of Rickettsia conorii infection were diagnosed in Perth, Western Australia. Both patients had recently returned from

Africa (Graves, 1998). A Japanese man who spent a short time on the Gold Coast,

Queensland in 2000 later developed symptoms consistent with a rickettsiosis, these symptoms were thought to be associated with an Ixodes holocyclus bite. Western blot analysis demonstrated reactivity of the serum to proteins specific for Rickettsia helvetica however no further evidence for the presence of R. helvetica in Australia has

28 been reported (Inokuma et al ., 2003). Until the current study, no native SFG rickettsiae had been isolated in WA, however, cases have been defined serologically

(Dyer, pers. comm., 2006).

1.4.2.8 Other pathogenic spotted fever group rickettsiae

• Israeli spotted fever ( Rickettsia conorii israelensis )

The agent of Israeli spotted fever (ISF) belongs to the R. conorii complex. It was first isolated from ticks and humans in 1974, and it has been reported from Israel, Portugal and Italy. The brown dog tick, Rhipicephalus sanguineus, has been identified as the most common vector of the organism. The Israeli spotted fever rickettsia is capable of producing a life-threatening illness especially as treatment is often delayed as there is usually no eschar present to assist with the diagnosis (Aharonowitz et al ., 1999;

Bacellar et al ., 1999; Giammanco et al ., 2003).

• Astrakhan fever ( Rickettsia conorii caspia )

The agent of Astrakhan fever (AF) was first described in 1991 in Astrakhan on the

Caspian Sea. It has since been detected in Africa and Kosovo. Astrakhan fever shows a similar clinical presentation and seasonal distribution to MSF. Its clinical presentation differs in that AF produces a milder disease, is yet to cause a fatality and has a low rate of eschar formation. The suspected vectors are the dog ticks

Rhipicephalus pumilio and R. sanguineus (Tarasevich et al ., 1991; Eremeeva et al .,

1994; Eremeeva et al ., 1995; Rydkina et al ., 1999; Parola and Raoult, 2001a; Parola et al ., 2005).

29

• African tick bite fever ( R. africae )

African tick bite fever was first described in Mozambique and South Africa in 1911.

Although these and subsequent cases were generally thought to be MSF, there was some debate as to whether tick bite fever could be caused by a distinct species based on a different epidemiology and clinical presentation. This debate was not resolved until 1990 when Kelly et al . isolated and characterized R. africae from the tick species

Amblyomma hebraeum (Parola et al ., 2005) .

Rickettsia africae is transmitted by Amblyomma hebraeum and A. variegatum and has been reported from South and Central Africa and the West Indies. It usually produces a mild, self-limiting disease with fever, multiple eschars, regional lymphadenopathy and, rarely, a skin rash (Kelly et al., 1992; 1992a; Kelly et al ., 1996; Parola et al .,

2001; Parola and Raoult, 2001a; Jensenius et al . 2003; Toutous-Trellu, 2003;

Rutherford et al., 2004; Kelly, 2006).

• Rickettsia sibirica mongolotimonae

This rickettsial species was isolated from asiaticum ticks in Mongolia and was initially thought to be non-pathogenic. Later, R. sibirica mongolotimonae was isolated from patients in southern France with fever, headache, an eschar, lymphangitis and lymphadenopathy. This presentation led to the proposed naming of the disease as “lymphangitis-associated rickettsiosis”. It has subsequently been detected in Hyalomma truncatum ticks from cattle in sub-Saharan Africa (Parola et al ., 2001; Parola and Raoult, 2001a; Fournier et al ., 2005).

30

• North Asian tick typhus ( R. sibirica sibirica )

This rickettsiosis was first described in 1936 and has now been shown to occur widely in Asiatic Russia. Disease occurs most often in summer and ranges from asymptomatic to serious infections with complications such as pneumonia and sepsis.

The tick vectors of R. sibirica are Dermacentor spp. and Haemaphysalis spp. (Yu et al ., 1993; Xu et al ., 1997; Chen et al ., 1998; Rydkina et al ., 1999; Lewin et al ., 2003;

Parola et al ., 2005).

• “Far Eastern Spotted Fever” (R. heilongjiangensis )

Rickettsia heilongjiangensis was first isolated from Dermacentor sylvarum ticks collected in the Heilongjian Province of China in 1982. Subsequent serological studies have confirmed that R. heilongjiangensis is the agent of a mild human rickettsiosis that occurs in China and the Russian far east. Infections occur most frequently at the end of June and July, in contrast to infections caused by R. sibirica sibirica in the same geographical area, which occur more commonly at the end of

April and May (Zhang et al ., 2000; Mediannikov et al ., 2004). Haemaphysalis spp. ticks are believed to be the vectors of this rickettsia (Parola et al., 2005).

• Rickettsialpox ( R. akari )

Rickettsialpox, an asymptomatic to mild disease characterized by low-grade fever, headache and a vesicular eruption over the trunk and extremities (similar in appearance to chicken pox) was first described in New York City in 1946. It has also recently been detected in the Ukraine, Slovenia, Croatia, Turkey and Korea

(Radulovic et al ., 1996; Raoult and Roux, 1997; Ozturk et al ., 2003). It is transmitted

31 by Liponyssoides sanguineus (formerly Allodermanyssus sanguineus ), a mite that parasitises the domestic mouse, and high risk groups include intravenous drug users and people living within densely populated areas (Raoult and Roux, 1997; Parola et al ., 1998; Comer et al ., 1999; Comer et al ., 2001; Choi et al ., 2005; Schoeler et al .,2005).

• Rickettsia slovaca

Rickettsia slovaca is an example of a SFG rickettsia that does not usually produce a rash in affected patients. It was first isolated from Dermacentor marginatus ticks in

Slovakia in 1968 however, its pathogenic potential was not confirmed until 1980. A rickettsiosis caused by R. slovaca was first described in 1997 in a patient who presented with a single lesion on the scalp and enlarged cervical lymph nodes after receiving a bite from a Dermacentor spp. tick. These clinical signs led to the infection being termed “tick-borne lymphadenopathy” (TIBOLA). The clinical syndrome produced by R. slovaca has also recently been referred to as “ Dermacentor -borne necrosis-erythema-lymphadenopathy” (DEBONEL) in Spain (Oteo et al ., 2004;

Parola et al ., 2005). Most reported cases have been mild and afebrile however the full spectrum of disease is still unknown as R. slovaca infection has been suspected in some patients displaying neurological signs. Rickettsia slovaca is thought to be widely distributed across Europe and can be differentiated from MSF based on its geographical distribution, the absence of a rash that is common in cases of R. conorii infection and differences in seasonal occurrence (Beati et al ., 1994; Raoult et al .,

1997a; Sekeyová et al ., 1998; Parola and Raoult, 2001a; Raoult et al ., 2002a).

32 • Rickettsia helvetica

Rickettsia helvetica was originally designated the ‘Swiss agent’ after being isolated from Ixodes ricinus ticks in Switzerland in 1979 (Beati et al ., 1993). More recently, this rickettsia has been implicated as the cause of two cases of fatal cases of peri- myocarditis in young patients in Sweden and an unexplained febrile illness in a

French patient. Serological evidence of R. helvetica infections has been demonstrated in eight patients from Italy, France and Thailand who presented with fever, headache and myalgia, but no cutaneous rash (Fournier et al ., 1998; Nilsson et al ., 1999;

Fournier et al ., 2004). Rickettsia helvetica has also been reported from Slovenia,

Japan and along the central Thai-Myanmar border. The vector is the tick Ixodes ricinus (Parola et al ., 1998; Nilsson et al ., 1999; Fournier et al ., 2000; Parola and

Raoult, 2001; Fournier et al ., 2002; Satoh et al ., 2002; Giammanco et al ., 2003;

Ishikura et al ., 2003; Parola et al ., 2005).

• Japanese spotted fever (JSF), Oriental spotted fever ( R. japonica )

The first case of JSF was reported in a patient from Tokushima Prefecture, Japan in

1984 and the causative agent was isolated in 1986 (Takao et al ., 2000). Since then it has been shown to be endemic in areas of southern Japan, where it occurs most frequently between April and November (Chiya et al ., 2000; Takao et al ., 2000; Satoh et al ., 2001; Ishikura et al ., 2003; Choi et al ., 2005). Japanese spotted fever presents as a typical rickettsiosis with skin rashes that develop initially on the extremities and then spread to the entire body. Central nervous system involvement and a single mortality have also been reported (Takao et al ., 2000; Araki et al ., 2002). Initial studies identified wild mice as the main mammalian reservoir for R. japonica.

However, recent studies have suggested that dogs and wild deer may also play an

33 important role in the life cycle (Takao et al ., 2000; Satoh et al ., 2001). The vector ticks identified are Dermacentor taiwanensis, Haemaphysalis flava and Ixodes spp. in

Japan (Takao et al ., 2000; Fournier et al ., 2002; Ishikura et al ., 2003) and recently,

Haemaphysalis longicornis in Korea (Choi et al ., 2005).

• Rickettsia parkeri

Rickettsia parkeri was first detected in Amblyomma maculatum ticks in the southern

United States in 1939. It was thought to be non-pathogenic until a case of human disease was described in 2004. Rickettsia parkeri produces a mild disease characterized by multiple eschars and a maculopapular rash. Its true prevalence is unknown as it is likely that R. parkeri has been mistaken previously for RMSF or rickettsialpox as it occurs in the same geographical areas (Paddock et al ., 2004;

Raoult, 2004). Recently, R. parkeri DNA has also been detected from A. maculatum ticks in South America (Parola et al ., 2005).

• Rickettsia massiliae

Rickettsia massiliae was identified in Rhipicephalus turanicus ticks near Marseilles,

France in 1990. Since then it has also been detected in Rhipicephalus spp. ticks in other areas of France, Greece, Spain, Portugal and Central Africa (Beati and Raoult,

1993; Parola and Raoult, 2001a). This rickettsia was classified as a “rickettsia of unknown pathogenicity” until R. massiliae DNA was identified in the blood of an

Italian man who had presented with an eschar and macropapular rash 20 years previously. A rickettsia was cultured from a blood sample collected at the time of presentation in 1985 and then stored, making this retroactive study possible (Vitale et al ., 2005).

34

• R. aeschlimannii

Rickettsia aeschlimannii was first detected in Hyalomma marginatum rufipes ticks collected from Zimbabwe and H. marginatum marginatum ticks collected from

Portugal, however it wasn’t characterized until 1997 following its isolation from H. marginatum marginatum ticks from Morocco. Rickettsia aeschlimannii has also been detected in H. marginatum rufipes ticks from Niger and Mali (Pretorius and Birtles,

2002). In 2002, the pathogenicity of R. aeschlimannii was established when an infection was reported in a patient returning from Morocco to France, and an infection was subsequently reported in a patient in South Africa (Pretorius and Birtles, 2002).

The rickettsiosis caused by R. aeschlimannii is characterized by an eschar, fever and a generalized maculopapular skin rash. The widespread nature of R. aeschlimannii has recently been demonstrated following its isolation in Croatia, Spain and Corsica and the detection of a rickettsia closely related to R. aeschlimannii from Haemaphysalis punctata ticks in Kazakhstan and Hyalomma dromedarii and H. truncatum ticks from the Sudan (Raoult et al ., 2002; Fernández-Soto et al ., 2003; Punda-Polic et al ., 2003;

Matsumoto et al ., 2004; Morita et al ., 2004; Shpynov et al ., 2004).

1.4.2.9 Rickettsiae of unknown pathogenicity

Many rickettsiae whose pathogenicities are still not determined have been identified worldwide, including R. bellii , R. canadensis , R. montanensis , R. rhipicephali , BJ-90,

R. hulinensis, Rickettsia tamurae sp. nov. and Rickettsia spp. Strain S (Raoult and

Roux, 1997; Zhang et al ., 2000; Mediannikov et al., 2004; Fournier et al ., 2006).

35 1.4.2.10 Non-pathogenic spotted fever group rickettsiae

Rickettsia peacockii is the only non-pathogenic rickettsia characterized to date. It is believed to be a tick-symbiotic rickettsia that interferes with the maintenance and transmission of R. rickettsii in Dermacentor andersoni tick populations . It has been suggested that the high prevalence of R. peacockii in tick ovaries on the east side of the Bitterroot Valley in Montana, and in areas of Colorado, interferes with the ability of R. rickettsii to be transmitted transovarially in these ticks, however it can still be acquired orally and transmitted transstadially or horizontally. Rickettsia peacockii is closely related to R. rickettsii however it differs in its limited ability to translate ompA. Consequently, adhesion to host cells is impaired as is its ability to undergo actin-based motility, which may explain its restriction to tick vectors and its non- pathogenic nature (Niebylski et al ., 1997; Munderloh and Kurtti, 1995; Azad and

Beard, 1998; Simser et al ., 2001; Baldridge et al ., 2004).

Non-pathogenic SFG rickettsiae are speculated to have arisen from pathogenic rickettsiae that over time lost the ability to cause disease in both mammalian and tick hosts. It is possible that this process could be reversed under certain conditions. Study of rickettsiae such as R. peacockii may help in defining mechanisms that would allow non-pathogenic organisms to re-emerge as pathogens and may provide information on how non-pathogenic endosymbionts could be manipulated for control of the ticks and the pathogens they harbour (Simser et al ., 2001).

1.4.3 Scrub typhus ( Orientia tsutsugamushi )

Scrub typhus is an ancient disease. It was described in China in the third century and it appeared in Western medical literature as early as 1879. Historically, disease had

36 been associated with farmers who entered lowlands to plant crops following spring floods. Investigators believed that the small red mites that were always plentiful during disease outbreaks probably played a role in transmission. In the early 1920s, investigators in Malaysia noticed that there were two distinct forms of typhus fever occurring. One, probably murine typhus, was seen most often in urban areas, while the other appeared to be a rural disease and was associated mainly with exposure to areas of “scrub”, cleared forest lands with secondary vegetation. In 1930 the rickettsial agent was identified by Nagayo and named Rickettsia orientalis. “ Scrub typhus” was the term used to describe the disease and it is still used commonly around the world, except in Japan where it is referred to as tsutsugamushi disease (Groves and Harrington, 1994).

The organism has undergone a number of taxonomic changes and currently belongs to the genus Orientia . Orientia tsutsugamushi (“tsutsugamushi” means “noxious mite” in

Japanese) is made up of a collection of antigenically and genetically diverse serotypes

(Walker and Fishbein, 1991). The three classic serotypes are: Karp (from Papua New

Guinea), Kato (from Japan) and Gilliam (from Burma) although there are many other serotypes present in east and south-east Asia and northern Australia (Graves, 1998).

The devastating effect of scrub typhus was particularly noticeable during World War II, when an estimated 40 000 soldiers were affected by the disease in southern Asia. It was also thought to be the major cause of “fever of unknown origin” in US troops in

Vietnam (Groves and Harrington, 1994).

37 Scrub typhus has a long history of occurrence in Australia. “Coastal fevers” were first described in northern Queensland in the agricultural population of farm labourers, wood and cane cutters in 1877. Scrub typhus was subsequently found to account for a significant proportion of these cases (Langan and Mathew, 1935) after all attempts to equate the fevers with known infections such as malaria, typhoid and plague failed

(Derrick, 1957; Graves et al ., 1999). Six possible cases of scrub typhus were recorded in the tropical region of the Northern Territory from 1937 to 1939 (Odorico et al .,

1998). Scrub typhus was again identified as the cause of infections in northern

Queensland during jungle and amphibious warfare training of troops on the Atherton

Tableland during World War II (McCulloch, 1944; Southcott, 1947). In 1941, Gordon

Heaslip demonstrated that the rickettsia was present in rats and bandicoots (Derrick,

1957) and the importance of bandicoots as reservoirs of infection in the scrub typhus life cycle was emphasised by Campbell and Domrow in 1974.

Since 1990, ten cases (one fatal) of scrub typhus have been reported in Litchfield

Park, Northern Territory and it is suspected that other foci of infection will be identified in similar rainforest pockets with increased tourism (Currie et al ., 1993;

Ralph et al ., 2004). The causative agent of this disease has been identified as a novel strain and called O. tsutsugamushi “Litchfield” (Odorico et al ., 1998; Graves et al .,

1999). In 1996 and 1997 there was another outbreak of scrub typhus in army personnel in north Queensland and in 2000 and 2001, a cluster of scrub typhus cases were diagnosed in the Torres Strait islands of northern Australia (McBride et al .,

1999; Faa et al ., 2003).

38 In 1999, a survey was performed on 920 non-randomly collected serum samples from the Kimberley region of Western Australia. Twenty four of the 920 samples tested positive for scrub typhus antibodies. This result, together with a confirmed case from the area, is highly suggestive of the presence of infection in the north of Western

Australia (Quinlan et al ., 1993; Odorico et al ., 1998; Graves et al ., 1999).

Larvae of the trombiculid mites of the genus Leptotrombidium are the only known vectors of O. tsutsugamushi. Mites acquire infection at a low rate by feeding on subdermal lymph and tissue fluids from an infected vertebrate. Infection is maintained in the mite population through extremely efficient transovarial transmission (Groves and Harrington, 1994). The trombiculid mites are very susceptible to drying so disease occurs only in areas of high rainfall and within these areas scrub typhus occurs in focal areas as there are sharply delineated islands of infected mites (Cook, 1944; Derrick,

1957; 1961; Groves and Harrington, 1994).

Infection of vertebrates occurs after a mite bite. Small rodents of the genus Rattus are the natural hosts, although over 55 species of mammal have been shown to be naturally infected with O. tsutsugamushi (Groves and Harrington, 1994). Scrub typhus is one of the more serious rickettsioses, with mortality prior to the discovery of effective antibiotic treatment sometimes reaching 60% (Devine, 2003).

1.4.4 B artonella spp.

Members of the genus Bartonella are fastidious, intracellular, Gram-negative bacilli that are capable of causing vector-transmitted zoonotic diseases. In the original

39 organisation of bacterial taxonomy, Bartonella was placed in the order Rickettsiales.

However, subsequent revision removed the genus Bartonella (which does not contain any obligate intracellular pathogens) from the order. The genus Bartonella has since been combined with the genus Rochalimaea and the new genus Bartonella has been placed into the α-2 subgroup of Proteobacteria based on 16S rRNA sequence analysis

(La Scola and Raoult, 1997).

The pathogenesis of Bartonella spp. infections differs from that of other rickettsiae in that they target erythrocytes in the host, however they can also cause proliferation of vascular endothelial cells and the formation of new blood vessels, a characteristic unique to the genus (Anderson and Neuman, 1997; Windsor, 2001; La Scola et al .,

2002). Bartonelloses in people can result in a highly variable pattern of disease, with presentations including haemolytic anaemia, septicaemia, endocarditis, osteolysis, bacillary angiomatosis, myositis, retinitis, encephalopathy and lymphadenopathy

(Kordick et al ., 1999) To date, 10 of the 19 recognised species of Bartonella (B. bacilliformis , B. quintana , B. henselae , B. elizabethae, B. vinsonii subsp. berkhoffii , B. vinsonii subsp. arupensis, . B. koehlerae, B. grahamii, B. washoensis and B. clarridgeiae ) have been implicated in human disease (Zeaiter et al ., 2002; Jardine et al ., 2006).

“Carrion’s disease”, which is caused by Bartonella bacilliformis, was perhaps the first recognised bartonellosis, with descriptions dating back to the 16 th century (Ihler,

1996). It is transmitted by sandflies and infection can result in two forms of disease.

The acute form manifests as severe, life threatening anaemia, while the chronic form results in vascular proliferative lesions of the skin termed “verruga peruana”. It is

40 limited to certain regions of the Andes mountains and although outbreaks still occur occasionally it is principally viewed as a medical curiosity (Ihler, 1996; Anderson and

Neuman, 1997).

Other Bartonella spp. have had a more devastating impact. “Trench fever”, caused by

Bartonella quintana and transmitted by the human body louse ( Pediculus humanus ), was estimated to have affected over 1 million troops during World War I (Anderson and Neuman, 1997). Cat scratch disease (CSD) which is caused by Bartonella henselae is also the cause of significant morbidity in many areas of the world. The cat flea ( Ctenocephalides felis ) has been shown to be responsible for transmission of B. henselae between cats, human infection results when the organism is inoculated via contaminated cat teeth or claws (Windsor, 2001). Cat scratch disease manifests as lymphoid hyperplasia, granuloma formation, microabscess development, and in some cases suppuration, and while it is usually a benign and self-limiting disease, complications and mortalities can occur especially in immuno-compromised individuals.

Bartonella henselae is the only Bartonella spp. to be extensively recorded in Australia however there has also been a single case of Bartonella quintana infection reported in an immuno-compromised patient (Rathbone et al ., 1996; Branley, 1997).

41 1.5 OBJECTIVES

The overall objective of this study was to address the shortfall of in-depth knowledge on Western Australian rickettsiae, paying particular attention to the role of native and feral animals and their ectoparasites as vertebrate hosts and vectors. More specifically, the aims of this study were:

1. To gain a better understanding of whether species of Rickettsia are present in

WA, with the main emphasis on the tick-borne spotted fever group.

Our understanding of the occurrence of SFG rickettsiae in particular in the State is lacking. We have serological data and unpublished case information indicating the presence of members of the biogroup, but so far none have been isolated or characterized. The first aim of this study therefore is to screen ectoparasite samples

(concentrating on ticks) for rickettsial infection, to confirm their presence and to identify them either as a previously characterized or a novel species of rickettsia.

2. To identify geographical areas, vectors and vertebrate hosts of interest in local

rickettsial life cycles, in particular, the role of native and feral animals in

rickettsial ecology.

Samples will be opportunistically collected from different geographical locations and screened to determine whether there is any difference in infection prevalence across the areas sampled. The potential role of different vertebrate host species will also investigated, with particular emphasis on the role of WA’s unique native fauna as reservoir hosts.

42 3. To perform a preliminary investigation into the pathogenic potential of any

novel species identified during this study.

It is important to investigate the zoonotic potential of any novel isolate recovered as this will largely dictate the course of any future studies on the organism and will determine whether the investment of resources into control and prevention strategies is warranted. As yet there is no laboratory-based method for distinguishing pathogenic from non-pathogenic Rickettsia species and a species can only be established as pathogenic after it has been isolated from a person demonstrating symptoms of a rickettsiosis. A questionnaire will be issued and a serosurvey performed on a population at risk to investigate whether infection is occurring and whether this infection is associated with symptoms.

4. To contribute to the body of knowledge on rickettsial ecology worldwide in

order to aid in developing a better understanding of the genus Rickettsia .

Due to their obligate intracellular nature, small amount of genetic information and slow rate of mutation it is very difficult to develop tools for genetic manipulation to gain a better understanding of rickettsial gene function. Rickettsiologists therefore have relied almost entirely on the comparison of sequences and phenotypes from different Rickettsia species to gain an understanding of gene function, rickettsial ecology, evolution and dissemination. In order to gain the maximum amount of benefit from this comparison it is imperative therefore to have as much sequence information as possible from around the world. At the moment, rickettsial research is very active in Europe and the USA however shortfalls in rickettsial research in other parts of the world mean that there are still pieces of the rickettsial puzzle missing.

Sequence information from WA Rickettsia species will aid in filling this deficit.

43 Identifying and understanding methods of rickettsial dissemination using molecular epidemiological techniques is also critical for the development of effective control strategies.

5. To contribute, through opportunistic sampling, to the current knowledge of tick

presence in Western Australia, including the species of tick present, their

distribution and vertebrate host preference.

Knowledge of the tick species present in an area can aid in predicting which

vector-borne diseases are endemic and will also aid in prompt recognition of

exotic vectors, before they become established in an area.

44 CHAPTER 2

COLLECTION AND IDENTIFICATION OF

ECTOPARASITES

2.1 INTRODUCTION

In 1970, F. H. S. Roberts published a comprehensive survey of Australian ticks performed over a period of approximately 30 years. Since then there has been little published on the subject (Halliday, 2001). The aim of the initial stage of this project was to opportunistically collect ticks from different locations throughout the state, in particular from native animals, and to use the information on vector distribution, in addition to that published by Roberts, to determine the likelihood of tick-borne diseases being present in a particular geographical or ecological area of the state.

2.2 MATERIALS AND METHODS

2.2.1 Sample collection

Ectoparasites (ticks primarily, although some fleas were also collected) were opportunistically collected from wildlife trapped by researchers from the Departments of Biological Sciences and Veterinary and Biomedical Sciences, Murdoch University; the Department of Animal Science, University of Western Australia and the

Department of Conservation and Land Management (CALM). These ectoparasites were collected using guidelines developed by the author (Appendix 2). Additionally, ectoparasites were collected from animals at a wildlife rehabilitation centre in the outer suburbs of the Perth metropolitan area. Ticks were also collected from individuals whose work (researchers trapping animals, workers on Barrow Island) or

45 recreational activities (rogainers, bush walkers) necessitated spending time in heavily vegetated areas which exposed them to ticks. Maps showing the locations of trap sites and other collection sites are shown in Figures 2.3 and 2.4. All ticks collected were placed in Eppendorf vials containing 70% ethanol and catalogued prior to identification and analysis for the presence of rickettsiae. A list of all ticks collected and their hosts is shown in Table 2.1.

Trapping of animals on Barrow Island (BWI) was performed in established grids as part of an annual wildlife census with CALM and strategically around burrowing bettong warrens (which are also utilised by common brushtail possums and golden bandicoots) as part of a concurrent PhD project on bettong populations (figure 2.1).

Trapping of yellow-footed antechinuses in Dwellingup was performed in established grids, again as part of a concurrent PhD project. Trapping by CALM staff used methods established by the agency over many years.

Briefly, animals were trapped in cage or small Elliot traps placed strategically in study areas and baited with a mixture of peanut butter, rolled oats and sardines. The traps were opened in the evening and checked for animals in the early morning. Animals were handled in bags made from soft material. All trapped animals were examined for ectoparasites, with particular attention being paid to areas around the animals’ face, ears, elbows and scrotum or pouch (figure 2.2) (Roberts, 1970). This work was approved by Murdoch University’s animal ethics committee.

46

Figure 2.1 Native animal trapping on Barrow Island

Figure 2.2 Tick on the ear of a burrowing bettong

47

Figure 2.3. Map of Western Australia showing sample collection sites.

48

Figure 2.4. Map of the southwest of Western Australia showing sample collection sites.

2.2.2 Tick identification

Ticks were identified based on their morphology by light microscopy, using a dissecting microscope and the taxonomic key outlined in Roberts, 1970. Where possible, the species was identified. However, in some cases, mouthparts or other areas of the body used for differentiation were damaged during removal from the host

49 and it was only possible to identify them to the genus level. Fleas were also identified using light microscopy and a standard key (Dunnet and Mardon, 1974).

2.3 RESULTS

The following is a description and discussion of the results from the tick collection.

Results pertaining to the fleas will be discussed in chapter 5. Table 2.1 shows the collection site, tick species, the tick’s vertebrate hosts and the number of animals infected. The scientific names for the animals are listed in Appendix 3.

Tick samples belonging to three different genera ( Ixodes , Amblyomma and

Haemaphysalis ) were collected in 208 instances from animals, humans or the environment. Three species of Ixodes were identified, namely I. australiensis, I. antechini and I. myremecobii. Three different species of Amblyomma were also collected ( A. triguttatum, A. albolimbatum and A. limbatum ). One species of

Haemaphysalis, H. ratti was identified. The tick species most commonly collected are pictured in figures 2.5 – 2.8.

Many of the vertebrate hosts were only infected with one species of tick. For example, Ixodes antechini was the only tick species collected from yellow-footed antechinuses, and bobtail lizards were exclusively infected with Amblyomma albolimbatum, even though the reptiles sampled were from widely separated locations

(Perth and Lake Magenta). Two different species of tick were collected from humans:

Ixodes myremecobii from Lake Magenta and the Fitzgerald River National Park, the most south-eastern sites from which samples were collected, and Amblyomma triguttatum from the south-west and from Barrow Island. Of the samples collected

50 from the mainland, Amblyomma triguttatum was the only tick species collected from more than one vertebrate host species (kangaroos and people).

Table 2.1. Hosts from which tick species were collected in Western Australia, the collection sites and the numbers of animals on which ticks were found.

Host species Collection site Ectoparasite species No. of animals or (fig.s 2.1 and 2.2) humans with ticks or number of ticks collected from the environment Bobtail lizard Perth Amblyomma albolimbatum 17

Bobtail lizard Lake Magenta Amblyomma albolimbatum 1

Olive python Kimberleys Amblyomma spp. 1 Yellow-footed Dwellingup Ixodes antechini (fig.2.8) 15 Antechinus Southern brown Dwellingup Ixodes australiensis 1 Bandicoot Golden bandicoot Barrow Island Haemaphysalis ratti (fig.2.6) 34

Golden bandicoot Barrow Island Amblyomma limbatum 1

Bandicoot Kimberleys Haemaphysalis spp. 6 Common brushtail Barrow Island Amblyomma limbatum 15 Possum Common brushtail Barrow Island Haemaphysalis ratti 4 Possum Burrowing bettong Barrow Island Haemaphysalis ratti 18

Burrowing bettong Barrow Island Amblyomma limbatum 24

Western brush Fitzgerald River Ixodes sp. 1 wallaby National Park

Western grey Perth Amblyomma triguttatum 4 kangaroo (fig.2.5) Kimberley rock rat Kimberley Haemaphysalis sp. 1 Rat Kimberley Haemaphysalis sp. 1 Feral pig Serpentine Ixodes sp. 1 Feral pig Nannup Ixodes sp. 1 Humans Fitzgerald River Ixodes myremecobii 1 National Park Humans Waychinicup National Ixodes myremecobii 1 Park Humans Lake Magenta Ixodes myremecobii 1 Humans Perth Amblyomma triguttatum 1 Humans Williams Amblyomma triguttatum 14 Humans Barrow Island Amblyomma triguttatum 32

Environment Boddington Amblyomma triguttatum 12

51

Two species of ticks ( Haemaphysalis ratti and Amblyomma limbatum ) were found on all three marsupial species trapped on Barrow Island. Golden bandicoots were predominantly infected with Haemaphysalis ratti ticks (34/35 animals) however a small number of Amblyomma limbatum ticks (1/35 animals) were also collected from trapped animals. Conversely, common brushtail possums were predominantly parasitised by Amblyomma limbatum, however one animal had a Haemaphysalis ratti tick . Burrowing bettongs had similar numbers of each species of tick collected from them: Amblyomma limbatum on 24/39 animals , Haemaphysalis ratti on 18/39 animals). Three bettongs were co-infected with both species of ticks at the time of capture.

Figure 2.5. Dorsal view of Amblyomma triguttatum collected from a person on Barrow Island.

52

Figure 2.6. Ventral view of Haemaphysalis ratti collected from a golden bandicoot on Barrow Island.

Figure 2.7. Dorsal view of Amblyomma limbatum collected from a common brushtail possum on Barrow Island.

53

Figure 2.8. Dorsal view of Ixodes antechini collected from a yellow-footed antechinus in Dwellingup.

2.4 DISCUSSION

Most of the results obtained in this chapter concur with those reported by Roberts in his survey (table 2.2 and figure 2.9), the novel findings of this study are discussed below.

The occurrence of Ixodes antechini in Western Australia has not been previously reported. Records of this tick species in Australia have been from Antechinus species

(Antechinus flavipes, A. minimus minimus, A. swainsonii swainsonii, A. s. mimetes, A. m. maritimus, A. stuartii ), eastern quolls ( Dasyurus viverrinus) and dunnarts

(Smithopsis spp.) in New South Wales, Victoria, Tasmania and South Australia

(Roberts, 1970). A single I. antechini has also been recorded from a bird, the fairy prion ( Pachyptila turtur ) in Tasmania (Roberts, 1970). Based on its population density at the time of trapping, I. antechini seems well established on yellow-footed antechinuses in the Dwellingup area. It is likely therefore that it has always been present but was not detected in the survey performed by Roberts. It is not surprising to

54 find that Roberts’ data is incomplete as his survey was a very ambitious undertaking and could not have hoped to be exhaustive.

Table 2.2 Tick species reported in Western Australia (Roberts, 1970; Stenos et al .,

2003). *: indicates species collected in this study.

Genus Ixodes eudyptidis

tasmani hydromyidis australiensis* fecialis vestitus myremecobii*

Genus Haemaphysalis humerosa ratti* lagostrophi

Genus Boophilus microplus

Genus Rhipicephalus sanguineus

Genus Amblyomma calabyi postoculatum limbatum* australiense triguttatum*

Genus Aponomma fimbriatum

Genus Bothriocroton hydrosauri (formerly Aponomma hydrosauri )

Genus Ornithodoros gurneyi

55

Figure 2.9 A map of Western Australia demonstrating previously recorded distribution of ticks collected in this study (Roberts, 1970; Pearce and Grove, 1987; M. Harvey, pers. comm., 2003). IA – Ixodes australiensis; IM - Ixodes myremecobii; AT – Amblyomma triguttatum ; HR – Haemaphysalis ratti; AL – Amblyomma limbatum.

56 If I. antechini is a recent introduction to the state however, it is possible that movement of the fat-tailed dunnart ( Sminthopsis crassicaudata ) or the western quoll

(D asyurus geoffroyi) within their habitat range was responsible for this introduction.

The fat-tailed dunnart’s range extends from the eastern states across to WA. The distribution of the western quoll is now confined to the southwest of WA although previously its distribution extended across to the eastern states. It is less likely that the migration of other marsupial species could have facilitated the introduction as although Antechinus flavipes occurs in both WA and the eastern states, these populations have been shown to be separated by a large geographical barrier

(Menkhorst and Knight, 2001). Migration of fairy prions could have been responsible for the introduction of this tick species as they have been recorded in Tasmania and

WA. This would be less likely however, as fairy prions are uncommon in WA, they are also seabirds and Dwellingup is located a considerable distance inland (Roberts,

1970; Simpson and Day, 1986). These observations however are merely speculative and would need to be proved by comparing populations of ticks across Australia using genetic methods.

Another interesting finding of this study is the identification of the reptile-associated tick Amblyomma limbatum (Roberts, 1970) from mammals, namely the common brushtail possum and the burrowing bettong on Barrow Island. Although, Amblyomma limbatum ticks are relatively non-specific about their choice of reptile host, finding such a large number on mammalian hosts is unusual (Belan and Bull, 1995). This could mean that the range of this tick species is not dependent on the presence of reptiles.

57 Throughout the world, in tropical, temperate and even tundra regions, ticks are serious veterinary and medical pests. As vectors, ticks surpass all other in the variety of microparasites that they transmit to humans as well as to livestock. These include fungi, viruses, bacteria, rickettsia and protozoan parasites (Randolph, 2000).

For this reason it is important to determine the range of tick species present in the

State, and their geographical distribution and ecology. This information can then be used to assess the risk of tick-borne diseases, which are often associated with specific tick genera or species (eg. the association of Queensland tick typhus with Ixodes spp. in the eastern states) and to prioritise the areas where control and prevention measures are implemented. Baseline knowledge of the tick species present in an area will also aid in the prompt recognition of exotic species which can be introduced by migratory birds and people travelling into the country. For example, Dermacentor variabilis ticks, which are vectors of Rocky Mountain spotted fever, have been removed from travellers returning to the east coast of Australia, and an eco-climatic analysis indicated that they would be capable of establishing and propagating in the area if left to do so unchecked (Russell, 2001).

There are no previous reports incriminating Ixodes antechini as a vector for agents of zoonotic or animal disease. At this stage therefore, the likelihood of tick-borne diseases being present in WA is not increased by finding this previously unreported tick species here. Of the ticks collected in this study and already reported in the state,

Amblyomma triguttatum and the Ixodes spp. are of potential zoonotic importance.

Amblyomma triguttatum is a recognised vector of Coxiella burnetii and is likely to be involved in its life cycle in Western Australia (McDiarmid, et al., 2000) and Ixodes spp. are known vectors of many zoonotic pathogens including Rickettsia australis, R.

58 helvetica and Borrelia spp. (Estrada-Pen~ and Jongejan, 1999), it is possible therefore that these pathogens are present or could become established in the state if introduced.

Although this study is by no means comprehensive, due to the logistical problems of sampling across a large State, the findings have contributed to our knowledge of tick ecology in Australia and have set a firm basis for further studies of sub-populations in the future.

59 CHAPTER 3

DETECTION OF RICKETTSIAE IN ECTOPARASITES COLLECTED IN WA

3.1 INTRODUCTION

The significance of rickettsioses to human and animal health is becoming increasingly recognised worldwide (Raoult and Roux, 1997). In contrast to the eastern states, there is a limited amount of information on the presence of rickettsiae in Western Australia, a State which is effectively isolated by the Nullarbor Plain in the south and by the

Great Sandy, Gibson and Great Victoria Deserts further north. Murine typhus was first reported in the state in 1927 and cases have since been described from Albany in the south to Exmouth in the north (Saint et al ., 1954; Forbes et al ., 1991; Kelly et al .,

1995; O'Connor et al ., 1996; Beaman and Marinovitch, 1999). The presence of scrub typhus was also confirmed in the north of the state when a case was diagnosed in the

Kimberleys in 1998 (Odorico et al ., 1998). In addition, there is strong circumstantial evidence that spotted fever group rickettsiae are present in WA as a few cases of spotted fever have been diagnosed clinically and serologically in people who have not been outside the State in the months preceding their illness, including one case in a veterinarian associated with Murdoch University (J. Stenos, pers. comm., 2005).

Preliminary epidemiological studies have detected spotted fever antibodies in human sera from the north and southwest of the state and scrub typhus antibodies in the north, however no rickettsiae have been isolated thus far (Burnet, 1942; Stewart,

1991; Kilminster T. An Investigation of Typhus in Western Australia. Honours

Thesis: Department of Microbiology, University of Western Australia; 1997; Graves et al ., 1999). The aim of the current study was to investigate the presence of

60 rickettsiae in WA and to determine some epidemiological aspects of the infections, including distribution, vertebrate hosts and tick vectors.

3.2 MATERIALS AND METHODS

3.2.1 Study design

A comprehensive survey of the whole State was not possible therefore the aim of this study was to use the opportunistically collected ectoparasite samples described in chapter 2 for analysis. Barrow Island (BWI), which is located about 1300km north of

Perth and approximately 60 km off the coast, was particularly targeted for sampling as there had been anecdotal evidence for the occurrence of a disease of unknown but possibly rickettsial origin (J. Drummond, pers. comm., 2003) and workers on the island suffered a high incidence of tick bites.

Ectoparasite samples were collected for analysis as, in addition to being a relatively non-invasive, easily collected sample, they are also a vital part of the rickettsial life cycle, making them a sensitive indicator of the presence of rickettsiae in an area. Once infected, ectoparasites remain so for life and rickettsial infection is easily detected using molecular techniques, while in vertebrate hosts molecular-based screening relies entirely on the detection of the transient rickettsaemic phase (Kelly et al ., 1992a;

Raoult and Roux, 1997).

61 3.2.2 Ectoparasite identification

The methods and results for ectoparasite collection and identification are described in chapter 2.

3.2.3 Detection of rickettsial DNA Using PCR

3.2.3.1 Extraction of DNA

DNA was extracted from the ethanol preserved ticks using Chelex-100 resin (Bio-rad

Laboratories, Hercules, CA, USA) and a protocol based on that previously described by Guttman et al. (1996). If the tick was large (approximately 4mm or greater in length) it was bisected longitudinally and the remaining half was stored in 70% ethanol. If the ticks were smaller, up to four ticks from a single animal were used for extraction.

Ticks were removed from the 70% ethanol and washed with distilled water.

Dissection was performed using a sterile scalpel blade and a new disposable glass microscope slide. Each dissected tick was placed into an Eppendorf tube containing

300 µL of 10% Chelex and the tube was incubated at 56 °C overnight. After vortexing for 15 s the samples were heated at 95 °C for 15 min and then vortexed for a further 15 s. The samples were finally centrifuged at 14 000 rpm for 5 min before the supernatant containing the DNA was removed for use in the PCR reactions. The DNA was stored at 4 °C between uses (12-48 hours) and at -20 °C for longer-term storage.

3.2.3.2 Screening PCR

The preparation of the PCR pre-mix was performed in a “DNA-free” room. DNA extraction, the loading of DNA into the PCR premix, the running of the PCR assays

62 and the loading of PCR products into gels were also performed in separate areas to avoid cross-contamination.

DNA from Rickettsia typhi , R. honei and R. australis was used as positive control material (kindly provided by Dr Stephen Graves from the Australian Rickettsial

Reference Laboratory (ARRL)) and DNA from Coxiella burnetii , Bartonella henselae and Rickettsia felis was supplied by the World Health Organisation (WHO) Rickettsia

Reference Center, Centers for Disease Control and Prevention, Atlanta, U.S.A.

(kindly supplied by Dr. Greg Dasch). Positive and negative controls (distilled water) were used with every PCR reaction, the reaction was repeated if the expected result was not obtained for either of the controls. All primers were purchased from

Invitrogen Life Technologies (Mount Waverley, Victoria, Australia).

Primers that amplify a 381 base pair segment of the citrate synthase gene ( gltA ) of

Rickettsia prowazekii (RpCS.877p GGG GGC CTG CTC ACG GCG G and

Rp.CS.1258n ATT GCA AAA AGT ACA GTG AAC A) were chosen from published studies for the initial screening of samples (Regnery et al ., 1991). These primers detect DNA from members of the genus Rickettsia, the genus Bartonella and Coxiella burnetii (Regnery et al ., 1991; Birtles and Raoult, 1996). They do not amplify DNA from Orientia tsutsugamushi .

PCR was performed using a buffer concentration of 1x (Fisher Biotech, West Perth,

Western Australia), dideoxynucleotide triphosphate concentrations of 0.2 mM (dNTP,

Fisher Biotech), primer concentrations of 0.2 µM and 0.5 units of Taq DNA polymerase (Fisher Biotech) per tube. During validation of the PCR, a MgCl 2 (Fisher

63 Biotech) concentration of 2mM was found to be optimal. The PCR premix was made up to 17 L and 3 L of DNA template was added for each reaction.

PCR cycling conditions described by Regnery et al . (1991) were used with an increased annealing temperature found to be more optimal for the laboratory’s PCR thermocyclers. A hot start of 95 °C for 5 minutes was followed by 35 cycles of denaturation (20s at 95 °C), annealing (30s at 53 °C) and extension (2min at 60 °C). A final extension (7 min at 72 °C) was then performed and reactions were held at 14 °C.

Amplification was carried out using a Perkin Elmer 2400 thermocycler machine

(Perkin Elmer, Wellesley, MA, USA).

3.2.3.3 PCR product detection methods

PCR products were visualised using ultraviolet light following electrophoresis on 1% agarose gels. One gram of agarose (Bio-rad Laboratories) was added to 100 ml of tris-acetate-EDTA (TAE) buffer (40 mM Tris-HCL; 20 mM acetate; 2 mM EDTA; pH 7.9) and dissolved in the microwave. After the agarose had cooled slightly, 1 µL of ethidium bromide was added. The agarose solution was then poured into a gel tray with a 20 well comb that had been taped with autoclave tape. Once the gel had set,

4µL of loading buffer (Sigma, Saint Louis, Missouri, USA) was added to 20 µL of

PCR product and the mixture was loaded into the wells. The DNA was electrophoresed on horizontal gels in electrophoretic cells (Bio-rad Laboratories) at 80 volts for approximately 40 minutes and visualised under ultraviolet light. The image was then saved digitally for analysis with the Molecular Analyst program (Bio-rad

Laboratories).

64 3.2.4 Sequencing

3.2.4.1 Sequencing of the gltA and ompA genes

At least one PCR positive sample from each location (and within these locations from each vector and vertebrate host), was sequenced using the gltA gene primers previously described. Eighty microlitres of PCR product were electrophoresed on a

1% agarose gel (4 x 20 µL of PCR product were run in adjacent wells), visualized using ultraviolet light and cut out of the gel using a sterile scalpel blade. The PCR product was purified from unincorporated primers and nucleotides using the

QIAquick Gel Extraction kit (Qiagen, Hilden, Germany) and the protocol described by the manufacturer, with the exception that the final DNA was stored in 30 µL of elution buffer.

Sequencing was performed using 5 µL of DNA template, 4 µL of Big Dye version 3.1 terminator mix (Applied Biosystems, Foster City, CA, USA) and 3.2pM of primer.

The reaction mix was made up to 10 µL using PCR grade water. The sequencing PCR amplification conditions were as follows: an initial step of 95 °C for 2 min followed by

25 cycles of denaturing (96 °C for 10s), annealing (50 °C for 5s) and extension (60 °C for 4 min) before being held at 14 °C indefinitely.

The dye terminators were removed using Centrisep columns (Princeton Separations,

Inc., Adelphia, NJ, USA) and the protocol described by the manufacturer. The purified PCR product was sequenced using an Applied Biosystems 377 XL automatic sequencer and sequences were analysed using the Chromas version 1.45 program

(Technelysium Pty. Ltd., Tewantin, QLD, Australia). Each gene fragment was sequenced in the forward and reverse directions.

65 The initial gltA sequences of several of the PCR-positive samples were compared to previously characterized rickettsiae using the Basic Local Alignment Search Tool

(BLAST, http://www.ncbi.nlm.nih.gov/BLAST ) program. As some of these isolates were identified as members of the SFG it was thought to be prudent to do further sequencing of these samples using primers for the SFG-specific ompA gene. The ompA gene mutates more rapidly than the gltA gene and therefore is a more sensitive indicator of any divergence between the closely related SFG rickettsial species

(Regnery et al ., 1991). The primers used were Rp.190.70 (ATG GCG AAT ATT TCT

CCA AAA) and Rp.190.701 (GTT CCG TTA ATG GCA GCA TCT) (Fournier et al .,

1998). These primers amplify an approximately 630 base pair region of the ompA gene.

PCR was performed using a buffer concentration of 1x, dNTP concentrations of 0.2 mM, primer concentrations of 0.1 µM and 1.25 units of Taq DNA polymerase per tube. During validation of the PCR, a MgCl 2 concentration of 2mM was found to be optimal. The PCR premix was made up to 49 L and 1 L of DNA template was added for each reaction.

PCR cycling conditions involved a “hot start” of 95 °C for 3 minutes followed by 35 cycles of denaturation (20s at 95 °C), annealing (30s at 46 °C) and extension (1min at

65 °C). A final extension (7 min at 72 °C) was then performed before holding at 14 °C

(Fournier et al ., 1998).

66 3.2.4.2 Sequence analysis

After an initial alignment the sequences were cut so that they were all the same length. In order to determine how many species of rickettsia had been detected in this study, fragments of the gltA and ompA genes from a selection of samples from different geographical areas, vectors and vertebrate hosts were sequenced, aligned and compared using the multisequence alignment program CLUSTAL W in the

BISANCE software package. The novel strains were then compared to Rickettsia species with validly published names (Genbank accession numbers are listed in appendix 4). Sequence similarities were calculated using the INFOALIGN program in the EMBOSS software package (Williams, 2001).

3.3 RESULTS

3.3.1 Screening of ectoparasites

The ectoparasite samples collected and the results of their PCR screening are displayed in table 3.1.

67

Table 3.1. Tick samples collected from different hosts and locations in Western

Australia and their screening PCR status

Host species Collection site Tick species GltA screening (+ve/no. screened)

Bobtail lizard Perth Amblyomma 2/17

albolimbatum

Bobtail lizard Lake Magenta Amblyomma 0/1

albolimbatum

Olive python Kimberley Amblyomma spp. 0/1 Yellow-footed Dwellingup Ixodes antechini 8/15 Antechinus Southern brown Dwellingup Ixodes australiensis 0/1 Bandicoot Golden bandicoot Barrow Island Haemaphysalis ratti 2/34

Golden bandicoot Barrow Island Amblyomma limbatum 0/1

Bandicoot Kimberleys Haemaphysalis spp. 2/6 Common brushtail Barrow Island Amblyomma limbatum 7/15 Possum Common brushtail Barrow Island Haemaphysalis ratti 0/4 Possum Burrowing bettong Barrow Island Haemaphysalis ratti 3/14

Burrowing bettong Barrow Island Amblyomma limbatum 4/4

Western brush wallaby Fitzgerald River Ixodes spp. 0/1 National Park

Western Grey Perth Amblyomma triguttatum 4/4 Kangaroo Kimberley Rock Rat Kimberley Haemaphysalis spp. 0/1 Rat Kimberley Haemaphysalis spp. 0/1 Feral pig Serpentine Ixodes spp. 1/1 Feral pig Nannup Ixodes spp. 0/1 Humans Fitzgerald River Ixodes myremecobii 0/1 National Park Humans Waychinicup National Ixodes myremecobii 0/1 Park Humans Lake Magenta Ixodes myremecobii 0/1 Humans Perth Amblyomma triguttatum 1/1 Humans Williams Amblyomma triguttatum 4/14 Humans BWI Amblyomma triguttatum 15/32

Humans/environment Boddington Amblyomma triguttatum 0/12

68

3.3.2 Prevalence

Of the ticks collected on the mainland 8/15 (54%) Ixodes antechini ticks collected from yellow-footed antechinuses were PCR positive for infection using gltA gene primers. Four out of fourteen (29%) of the Amblyomma triguttatum ticks from

Williams were also positive as were 2/17 (12%) of the Amblyomma albolimbatum ticks collected from bobtail lizards from Perth.

Of the ticks collected from Barrow Island 15/32 (47%) Amblyomma triguttatum ticks collected from people, 2/34 (6%) Haemaphysalis ratti ticks collected from bandicoots and 7/15 (47%) Amblyomma limbatum ticks collected from possums were PCR positive. Although other ticks were positive for rickettsiae, the sample sizes from the other sites and hosts were too small to use for any meaningful indications of prevalence.

3.3.3 Sequence Similarities

The percentage nucleotide similarity comparing 328 bp of the gltA gene and 542 bp of the ompA gene from the two strains identified in this study and previously characterized Rickettsia species are displayed in tables 3.2 and 3.3.

69 Table 3.2. Percentage gltA sequence similarity between the two strains identified in this study and those of previously characterised Rickettsia spp.

Name % Similarity to % Similarity to Rickettsia sp. strain BWI-1 Rickettsia sp. strain D-1 Rickettsia sp. strain BWI- 1 100% 98.8%

Rickettsia sp. strain D-1 98.8% 100%

R. prowazekii 93.3% 93.3%

R. typhi 93.0% 93.0%

R. africae 98.2% 98.8%

R. conorii 98.2% 98.8%

R. sibirica 98.5% 99.1%

R. honei 98.5% 99.1%

R. slovaca 98.2% 98.8%

R. parkeri 98.5% 98.8%

R. rickettsii 97.5% 98.2%

R. massiliae 97.9% 98.5%

R. rhipicephali 98.2% 98.8%

R. aeschlimannii 98.8% 99.4%

R. montanensis 98.5% 99.1%

R. japonica 98.5% 99.1%

R. heilongjiangii 98.8% 99.4%

R. Helvetica 94.8% 94.8%

R. felis 95.1% 95.7%

R. akari 94.8% 95.4%

R. australis 95.7% 96.3%

R. bellii 86.0% 86.3%

70 Table 3.3. Percentage ompA sequence similarity between the two strains identified in this study and those of previously characterised Rickettsia spp.

Name % Similarity to % Similarity to

Rickettsia sp. strain BWI-1 Rickettsia sp. strain D-1

Rickettsia sp. strain BWI-1 100% 96.7%

Rickettsia sp. strain D-1 96.7% 100%

R. africae 93.5% 94.6%

R. conorii 92.3% 93.2%

R. sibirica 93.2% 94.6%

R. honei 93.0% 94.3%

R. slovaca 93.7% 95.0%

R. parkeri 93.0% 93.7%

R. rickettsii 92.6% 93.2%

R. massiliae 95.4% 94.8%

R. rhipicephali 95.4% 94.8%

R. aeschlimannii 94.8% 94.1%

R. montanensis 94.4% 93.7%

R. japonica 93.2% 93.7%

R. heilongjiangii 94.1% 94.6%

R. felis 53.1% 53.9%

R. australis 85.9% 85.5%

Based on the alignment of the gltA gene segment the rickettsial strain present in the samples from the feral pig from Serpentine, the people from Williams, the bobtail lizards, kangaroos and people from Perth and the people, bandicoots, burrowing bettongs and possums from BWI are identical. This was designated Rickettsia spp. strain BWI-1. The other rickettsial strain was only present in the ticks collected from

71 the yellow-footed antechinuses, this strain was present in all the PCR positive amplicons sequenced from these ticks. It is 98.8% similar to Rickettsia sp. strain

BWI-1 and was named Rickettsia spp. strain D-1. The ompA gene sequences confirmed the gltA gene sequence results. Rickettsia sp. strain BWI-1 is 96.7% similar to Rickettsia sp. strain D-1 based on the ompA gene sequences.

Sequencing of amplicons from the PCR-positive tick samples showed the possible presence of two novel rickettsia strains. The gltA gene segment Rickettsia sp. strain

BWI-1 is most similar to R. heilongjiangii , R. aeschlimannii and Rickettsia sp. strain

D-1 at 98.8% sequence similarity. The ompA gene segment of Rickettsia sp. strain

BWI-1 is 95.6% similar to R. massiliae and R. rhipicephali , these are its closest relative from the previously characterized rickettsial species however it shows a higher degree of sequence similarity to Rickettsia sp. strain D-1 at 96.7%. Based on the segment of the gltA gene sequenced, Rickettsia sp. strain D-1 shares 99.4% nucleotide similarity with both R. heilongjiangii and R. aeschlimannii . The sequenced ompA segment of Rickettsia sp. strain D-1 demonstrates 95.0% similarity with R. slovaca.

3.4 DISCUSSION

3.4.1 Aetiology

In order to be classified as a novel species Fournier et al ., 2003 recommend that an isolate should be less than 99.9% similar with the most related species of previously characterized rickettsia based on its gltA gene and less than 98.8% based on its ompA gene. Based on these criteria, the percentage similarity shown by the two strains to previously characterized rickettsiae and to each other provides strong evidence that

72 both Rickettsia sp. strain BWI-1 and Rickettsia sp. strain D-1 are novel and distinct species.

3.4.2 Prevalence

Rickettsia sp. strain BWI-1 showed a wide range of prevalences in different tick species, spanning from 6 to 47%. It is likely however that there were PCR inhibitors present in the ticks that may not have been completely removed in the DNA extraction process (Schwartz et al ., 1997). These and other factors would have contributed to the sensitivity of the PCR assay being less than 100%, so true prevalence is probably higher than these figures suggest.

An infection rate of 47% is relatively high, as although tick surveys performed in

Africa have demonstrated a high prevalence of infection (up to 72%), studies conducted in other parts of the world generally report lower rates of infection (Dupont et al , 1994; Beati et al, 1995).

The range in prevalences may be due to different tick species being exposed to rickettsiae at different rates or inherent differences in the vector capabilities of different tick species, or it could be due to differences between the island and mainland ecosystems where Rickettsia sp. strain BWI-1 occurs. It is also possible that the vertebrate hosts identified in this study have varying levels of host competency for an infecting rickettsia (i.e. susceptibility to infection, ability to support effective proliferation of the rickettsiae and duration of rickettsaemia). Tick species that favour a host with a relatively poor host competency are less likely to be exposed to the rickettsia and therefore can be expected to have a lower prevalence of infection.

73 Conversely, if a tick species favours an efficient host, they are likely to have a higher prevalence of infection.

It has previously been demonstrated that tick species can transmit rickettsial species transovarially with different levels of efficiency. Dermacentor variabilis ticks, for example, can become infected with either Rickettsia montanensis or R. rhipicephali.

A study found that while these ticks still had high levels of R. montanensis infection after two generations they were unable to sustain infection with R. rhipicephali for more than one generation. It is possible therefore, that even though infected ticks are found in nature, Dermacentor ticks are not optimal hosts for R. rhipicephali , a species originally isolated from the brown dog tick Rhipicephalus sanguineus (Macaluso et al ., 2002). Extrapolating this to the present study, it is possible that Haemaphysalis ratti ticks are not as efficient vectors of Rickettsia sp. strain BWI-1 as Amblyomma triguttatum.

Differences in prevalence rates could also be due to the ecosystems. As Barrow Island is a closed ecosystem, a rickettsia that is transovarially transmitted efficiently and that infects ticks permanently could reach a very high prevalence within a few generations.

In contrast, a mainland ecosystem would theoretically, have new, uninfected ticks constantly moving in from surrounding areas which would mean that a lower prevalence of infection could be expected.

The differences in prevalence of infection between the tick species could also be, in part due to laboratory and statistical artifact. The unengorged Haemaphysalis ratti are significantly smaller than the other species of ticks and it is possible that methods

74 used, particularly the extraction techniques may not detect infection as well in this species. In addition, the number of these ticks screened in this study was relatively small and may not have given a true indication of the prevalence of infection.

3.4.3 Life cycle: vectors and reservoir hosts

Rickettsiae often have complex life cycles involving a number of different ectoparasite vectors, domestic animal and wildlife reservoir hosts. This ability to utilise a range of vectors and hosts maximises the likelihood that the rickettsial infection will be maintained and propagated. Sylvatic life cycles are particularly advantageous being, in many cases, a reservoir of infection that is not reliant on or contained by human activity. Ectoparasites and vertebrates can interact to maintain infection in these sylvatic life cycles and consequently infection can be readily transmitted from sylvatic to domestic life cycles where they can infect people.

The importance of sylvatic life cycles in the epidemiology of rickettsioses has been widely demonstrated. For example, Ctenocephalides felis on opossums have been shown to transmit R. felis and R. typhi infections to the domestic cat-flea cycle when the opossums begin to scavenge around suburban houses (Sorvillo et al ., 1993;

Boostrom et al ., 2002). In Australia, R. australis utilizes the tick vectors Ixodes tasmani , I. holocyclus and I. cornuatus in its life cycle. All these tick species can infect both native and domestic animals and could therefore link the sylvatic and domestic life cycles (Roberts, 1970; Pinn and Sowden, 1998).

The life cycle of Rickettsia sp. strain BWI-1 appears to involve several different tick species and potentially a wide range of vertebrate hosts, including feral pigs, golden

75 bandicoots, kangaroos, burrowing bettongs, common brushtail possums and bobtail lizards. It has not yet been established whether these vertebrates become infected and therefore act as reservoirs of disease. This is especially true of the lizards, as despite

R. honei being found in a reptile-associated tick, the ability of reptiles to harbour rickettsial infection is unknown (Stenos et al ., 2003).

It is certainly possible that other species of vertebrate parasitised by these vectors could also be involved in the ecology of the novel rickettsiae. It was not possible to determine the species of the Ixodes tick on the pig however there are several species of Ixodes known to occur in the southwest of the state that are non-specific in their choice of vertebrate host. Ixodes australiensis were the only Ixodes spp. ticks collected in a recent study on the tick species parasitizing feral pigs in the southwest of WA, therefore it is highly likely that these ticks were also I. australiensis (Lee, pers. comm., 2006). Haemaphysalis ratti has been shown by other researchers to parasitise western-barred bandicoots ( Perameles bougainville ) on Dorre Island, off the mid-west coast of WA. Amblyomma albolimbatum is known to infest a wide variety of reptiles as is Amblyomma limbatum, which prior to this study has predominantly been recorded from reptiles (Roberts, 1970). In this study it was found on common brushtail possums, burrowing bettongs and golden bandicoots. If

Amblyomma limbatum is indeed prevalent on mammals it is possible that it could transmit Rickettsia sp. strain BWI-1 to a wide range of hosts. Amblyomma triguttatum ticks were only collected from people on Barrow Island; however, of the animal species present on the island, the black-footed rock wallaby ( Petrogale lateralis ), spectacled hare wallaby ( Lagorchestes conspicillatus ) and the euro

(Macropus robustus ) are potential vertebrate hosts (Roberts, 1970; Menkhorst and

76 Knight, 2001). The western grey kangaroo ( Macropus fuliginosus ), a common host for Amblyomma triguttatum ticks, is widespread throughout the state, although it is not found on Barrow Island.

Another interesting finding is that, of the three marsupial species trapped on BWI, golden bandicoots were predominantly infected with Haemaphysalis ratti, however, one animal was also infected with an Amblyomma limbatum tick. In contrast, common brushtail possums were most frequently infected with Amblyomma limbatum ticks but a small number of Haemaphysalis ratti were also collected from them. Burrowing bettongs were infested with relatively even numbers of each species of tick. It is noteworthy that some bettongs and one possum were co-infected with both tick species at the time of capture. This may explain the presence of Rickettsia sp. strain

BWI-1 in more than one species of tick as a rickettsaemic animal infected with both

Haemaphysalis ratti and Amblyomma limbatum ticks may be able to transmit the same species of rickettsia to both tick types. There is a slight possibility however, that two species of tick feeding in close proximity could transmit rickettsiae horizontally without the host animal becoming infected (Raoult and Roux, 1997). If this is in fact the case, it would alter how we must view the epidemiology of the novel strain, as a reservoir host might not be necessary for the maintenance of the rickettsia (Richter et al ., 2002). Based on these results, it appears that Rickettsia sp. strain BWI-1 has a complex life cycle that would make implementing effective control and prevention measures difficult.

In contrast to Rickettsia sp. strain BWI-1, Rickettsia sp. strain D-1 has only been identified in one tick species to date, Ixodes antechini, and on one vertebrate host, the

77 yellow-footed antechinus. Ixodes antechini , the tick vector of Rickettsia sp. strain D-1 has not been previously recorded in WA. In the eastern states it has been known to parasitise Antechinus species ( Antechinus flavipes, A. minimus minimus, A. swainsonii swainsonii, A. s. mimetes, A. m. maritimus, A. stuartii) , eastern quolls ( Dasyurus viverrinus), dunnarts ( Smithopsis spp.) and fairy prions ( Pachyptila turtur ). It would be prudent to collect and analyse other tick species in the area to try and establish whether this is a rickettsia that utilises a simple and specific life cycle or whether these tick and vertebrate species are part of a more complex life cycle. If the life cycle is in fact simple and specific then there is a decreased likelihood that people will become infected and it is unlikely to be of great zoonotic significance.

3.4.4 Geographical distribution

Rickettsia sp. strain BWI-1 appears to be widely distributed across the sites sampled in the southwest of the mainland and also on BWI. Rickettsia sp. strain D-1 has thus far only been identified in one location. The locations where the strains were identified are depicted in Figure 3.1.

78 Figure 3.1 Distribution of the rickettsial strains across WA . * - Rickettsia sp. strain BWI-1; ^ - Rickettsia sp. strain D-1.

As rickettsiae can only be maintained in areas that are conducive to the survival of their ectoparasite vectors an idea of their potential geographical distribution can be obtained by examining the distribution of these vectors.

Most of the tick species utilised by Rickettsia sp. strain BWI-1 have a relatively limited geographical distribution. Amblyomma triguttatum ticks occur in the southwest of the state and on BWI. Ixodes spp. only occur in the state’s southwest where there is sufficient rainfall for their survival. Haemaphysalis ratti , in addition to

79 occurring on BWI is present on Dorre Island (Roberts, 1970; M. Harvey, pers.comm,

2003). Amblyomma limbatum has only been reported from Abrolhos, Barrow and

Bernier Islands and Mount Wynne (Roberts, 1970). Amblyomma albolimbatum in contrast, is known to occur extensively throughout the state. Consequently Rickettsia sp. strain BWI-1 may have a wider geographical distribution than demonstrated by this study if Amblyomma albolimbatum is a significant vector.

Ixodes antechini has not previously been recorded in Western Australia. Other species of Australian Ixodes are distributed only where there is sufficient rainfall to provide the ambient moisture they require, therefore it is unlikely that the range of Ixodes antechini extends far into the drier areas surrounding the State’s southwest. Therefore, unless Rickettsia sp. strain D-1 is able to utilise other tick species as vectors, it is unlikely that it has a geographical range extending beyond the State’s southwest.

It is also possible that an unidentified rickettsia species is present in the Kimberley region of WA. Spotted fever group and TG antibodies were detected in the region in a survey conducted on human sera in 1999 (Graves et al ., 1999) and the unsequenced

PCR-positive samples obtained from Haemaphysalis spp . collected from bandicoots in this study provide further evidence for the existence of other species.

3.4.5 Zoonotic Potential

The pathogenic potential of Rickettsia sp. strain BWI-1 and Rickettsia sp. strain D-1 is unknown. However, all SFG rickettsiae found in ticks that bite people should be considered potential pathogens (Raoult, 2004) and in the current study Rickettsia sp. strain BWI-1 has been found in Amblyomma triguttatum ticks most of which were

80 removed from people. It was not possible to speciate the Ixodes spp. tick infected with

Rickettsia sp. strain BWI-1 due to damage to the mouthparts, however, of the Ixodes spp. present in WA, I. tasmani is the only species conclusively known to bite humans

(Roberts, 1970), and it is also a known vector of R. australis (Campbell and Domrow,

1974). The other vectors for Rickettsia sp. strain BWI-1 ( Haemaphysalis ratti ,

Amblyomma albolimbatum and Amblyomma limbatum ) are not known to bite people.

Ixodes antechini is not known to bite people, however this does not definitely discount Rickettsia sp. strain D-1 as a human pathogen as other anthropomophilic vectors may be involved in the life cycle.

Subsequent stages of this project will investigate further the epidemiological and genotypic characterization of the novel strains and their zoonotic potential.

81 CHAPTER 4

CHARACTERIZATION OF RICKETTSIA sp. STRAIN BWI-1

4.1 INTRODUCTION

Extensive sequencing of novel rickettsial isolates gives us important insights not only into the novel isolate itself but also into the Rickettsia genus as a whole, providing valuable information on rickettsial pathogenicity, obligate intracellular parasitism and bacterial evolution (Wood and Azad, 2000).

Rickettsial taxonomy is a controversial, continuously evolving field. Rickettsiologists have demonstrated that the 70% DNA-DNA relatedness used to describe bacterial species does not adequately differentiate rickettsiae. It has been proposed that rickettsiae have been subjected to different evolutionary pressures due to their small genome size and strict intracellular nature (Stenos et al ., 1998; Fournier et al ., 2003).

In 2003, Fournier et al. described a set of objective guidelines for the classification of new rickettsial isolates based on sequences from several genes, namely the 16S rRNA, citrate synthase ( gltA ), ompA and ompB and sca4 genes. Multiple genes with different evolutionary pressures provide the maximum amount of information about a new isolate.

Extensive characterization of the genotypic and phenotypic properties of novel isolates and their comparison to previously characterized species of rickettsiae also provides valuable information about the Rickettsia genus. The obligate intracellular

82 nature of rickettsiae causes technical difficulties which have long prohibited detailed study of their gene function through traditional molecular-based methods. These difficulties include having to ensure any genetic manipulation will not interfere with the rickettsia’s ability to infect host cells or else the manipulation will automatically fail. The successful isolation of the first “knock-out” of R. prowazekii was not obtained until early 2005 (Renesto et al ., 2005). The identification and isolation of rickettsial transformants has also been difficult due to a lack of selectable markers.

In the absence of easily utilised genetic tools, rickettsiologists have relied on information gained from comparing rickettsial genome sequences in order to identify proteins to be targeted in specific diagnostic assays, novel treatments and vaccine development. The recent inclusion of rickettsiae in the list of potential bio-terrorism agents has made the development of tools to prevent, diagnose and treat these infections a new priority (Renesto et al ., 2005).

Extensive sequence information can also be utilised for purposes of molecular epidemiology. For some infectious microbiological agents molecular epidemiology can provide very specific information on the nature of transmission events, points of outbreak and origin of an epidemic and whether a disease is caused by an endemic or an introduced agent. Molecular epidemiology is not as sensitive for the study of rickettsiae due to their low rate of genetic exchange, however it is still a valuable technique that can be used to identify possible modes of dissemination and evolution

(Chae et al ., 2003; Tarasevich et al ., 2003; Renesto et al ., 2005).

83 In Chapter 3, a rickettsia of unknown pathogenicity was detected from Barrow Island and the southwest of Western Australia in Amblyomma triguttatum ticks collected collected from people and kangaroos, Haemaphysalis ratti ticks collected from golden bandicoots and burrowing bettongs, Amblyomma limbatum ticks collected from burrowing bettongs and common brushtail possums, Amblyomma albolimbatum ticks collected from bobtail lizards and an Ixodes spp. tick collected from a feral pig. In this chapter, further characterization was performed on Rickettsia sp. strain BWI-1 in order to gain a better understanding of the evolutionary mechanisms of Australian rickettsiae in particular. This genetic information can also be used in conjunction with any ecological and phenotypic properties of the isolate that may be identified in the future in order to gain a better understanding of the Rickettsia genus as a whole.

4.2 MATERIALS AND METHODS

4.2.1 Sample collection

Amblyomma triguttatum ticks, a vector of Rickettsia sp. strain BWI-1 identified in

Chapter 3, were collected from people working on BWI. These ticks were kept alive in sterile, humid containers so that culture of the infecting rickettsiae could be performed in the laboratory.

4.2.2 Cell culture

Culture of Rickettsia spp. from these ticks was then undertaken at the Australian

Rickettsia Reference Laboratory in Geelong, Victoria. In order to remove as much of the surface bacteria as possible the live ticks were placed into a petri-dish and immersed in 95% sodium hypochlorite solution for 10 minutes. The ticks were dried, sprayed with 70% ethanol and then crushed in 1.5 ml of sterile phosphate-buffered

84 saline (PBS) using the end of a disposable bacterial loop. One millilitre of the tick solution was passed through a filter (0.45uM filter, Millex, Carrigtwohill, Ireland) and the filtrate was inoculated into a 25cm 2 flask (Nunc, Rochester, NY, USA) containing confluent XTC-2 cells. The XTC-2 cells were incubated at 28°C in flasks containing

Leibovitz L-15 media (Invitrogen) supplemented with 5% heat inactivated foetal bovine serum, 0.4% tryptose phosphate (Oxoid, Hampshire, England) and 200mM L- glutamine (Invitrogen). Cultures were maintained and observed for evidence of cytopathic effect by light microscopy weekly and by immunofluorescence assay (IFA) monthly by scientists working at the laboratory.

4.2.3 Immunofluorescence assay

For the IFA, cells from the cultures were acetone-fixed to a slide before being exposed to human serum that contained antibodies to spotted fever group, typhus group and scrub typhus group rickettsiae (titre ≥ 1024, supplied by the Australian

Rickettsia Reference Laboratory). Slides were then stained with fluorescein isothiocyanate (FITC) labelled goat anti-human (IgM, IgG and IgA) antibody

(Biomediq, Doncaster, Vic., Australia). Each conjugate was diluted 1:100 in casein buffer (2% skim milk powder in PBS, heat inactivated at 56°C for 30 min), incubated at 35°C for 30 minutes and then washed in 1:10 PBS. A continuous rickettsial culture and an uninfected XTC-2 cell culture were used as positive and negative controls respectively. DNA was extracted from IFA positive cultures using Gentra DNA extraction kits (Gentra, Minneapolis, MN, USA) and the protocol described by the manufacturer, their IFA-positive status was then confirmed by PCR. Cultures that were IFA and PCR positive were then passaged into newly confluent XTC-2 flasks.

85 4.2.4 Sequencing

Once it had been established that the cell cultures were infected, gltA and ompA gene segments from the infecting rickettsial isolate were amplified, sequenced and compared to the sequences obtained from the tick isolate (Rickettsia sp. strain BWI-

1). DNA was extracted from the cultures using Gentra DNA extraction kits and the protocol described by the manufacturer. The primers used were RpCS877p and

RpCS1258n for the gltA gene and Rr190.70p and Rr190.701 for the ompA gene.

These primers and their PCR cycling conditions were previously described in chapter

3.

Further PCR amplification and sequencing was performed on the isolate in culture using other sections of the gltA gene along with segments of the ompB, sca4 and 16S rRNA encoding genes. A 1098 bp gltA gene fragment, a 482 bp ompB encoding gene fragment, a 763 bp sca4 fragment and a 1335 bp 16S rRNA fragment were all sequenced using the primers listed in table 4.1, these were optimised with positive control DNA before they were used on the samples. PCR conditions are listed in tables 4.2 and 4.3. All primers were sourced from the literature where possible or designed using the computer program Amplify 1.2 (Engels, 1993). Sequencing has been previously described in section 3.2.4.

86 Table 4.1. Primers used for PCR and sequencing of DNA from Ricketssia spp. strain

BWI-1.

Gene primer Primer sequence Reference PCR conditions (table

4.2)

GltA RpCS877p 5’ GGGGGCCTGCTCACGGCGG 3’ (Regnery et al ., 1991) 1

GltA RpCS1258n 5’ATTGCAAAAAGTACAGTGAACA 3’ (Regnery et al ., 1991) 1

OmpA Rr190.70p 5’ATGGCGAATATTTCTCCAAAA 3’ (Regnery et al., 1991) 2

OmpA Rr190.701 5’ GTTCCGTTAATGGCAGCATCT 3’ (Fournier et al., 1998) 2

GltA CS162F 5’GCAAGTATCGGTGAGGATGTAATC (Stenos et al ., 1998) 3

3’

GltA CS398SF 5’ATTATGCTTGCGGCTGTCGG 3’ (Stenos et al ., 1998) 3

GltA CS764SF 5’ATTGCCTCACTTTGGGGACC3’ (Stenos et al ., 1998) 3

GltA CS731SR 5’AAGCAAAAGGGTTAGCTCC 3’ (Stenos et al ., 1998) 3

GltA CS411SR 5’AATGCCGCAAGAGAACCGACAG3’ (Stenos et al ., 1998) 3

GltA CS877R 5’CCGCCGTGAGCAGGCCCCC Designed 3

Omp B ompB120M59 5’CCGCAGGGTTGGTAACTGC 3’ (Roux and Raoult, 2000) 4 ompB ompBr 5’GAGGAGCTTTTTGAGTTGTAG 3’ Designed 4

16s rRNA 1 5’GACTCGGTCCTAGTTTGAGA 3’ (Rogall et al ., 1990) 5 rRNA

16S rRNA3 5’TTTGAGTTTCCTTAACTCCC3’ (Rogall et al ., 1990) 5 rRNA

16S rRNA2 5’CAGCCGACCTAGTGGAGGAA 3’ (Rogall et al ., 1990) 5 rRNA

16S rRNA 4 5’CATAATGGCGCCGACGAC 3’ (Rogall et al ., 1990) 5 rRNA

Sca4 geneD1f 5’ATGAGTAAAGACGGTAACCT 3’ (Sekeyova et al ., 2001) 4

Sca4 Gene D928 5’ AAGCTATTGCGTCATCTCCG (Sekeyova et al ., 2001) 4

87

Table 4.2. Conditions and the premix used for the PCR reactions amplifying Rickettsia spp. BWI-1 DNA - all reactions used a buffer concentration of 1x and a dNTP concentration of 0.2 mM.

PCR Primer MgCl 2 Taq Premix DNA conditions concentration ( µµµM) Concentration (u/tube) volume volume (see table (mM) (µµµL) (µµµL) 4.1) 1 and 3 0.2 2 0.5 17 3 2 0.1 2 1.25 49 1 4 0.25 2.5 1 45 5 5 1 2 2 21 4

Table 4.3. PCR cycling conditions used in all reactions

PCR conditions (see table 4.1) Number of cycles Denaturation Annealing extension

1 35 20 s at 95 °C 30 s at 53 °C 2 min at 60 °C 2 35 20 s at 95 °C 30 s at 46 °C 1 min at 65 °C 3 35 20 s at 95 °C 30 s at 50 °C 2 min at 70 °C 4 40 20 s at 95 °C 30 s at 50 °C 1.5 min at 68 °C 5 40 30 s at 95 °C 30 s at 51 °C 1 min at 72 °C

4.2.5 Sequence analysis.

The sequences were initially subjected to BLAST analysis. The sequences of the gltA , ompA , ompB , sca4 and 16S rRNA genes for Rickettsia species with validly published names were then accessed from GenBank (accession numbers in appendix 4) before the sequences were aligned using the multi-sequence alignment program CLUSTAL

X in the BISANCE software package. All the rickettsial sequences for a gene were cut to the length of the shortest sequence.

88

The percentage of similarity between the species was determined using the

INFOALIGN program in the EMBOSS software package (Williams, 2001). Sequence similarities were calculated based on pairwise transitions and transversions between sequences. Phylogenetic and molecular evolutionary analyses were conducted using

MEGA version 2.1 (Kumar et al ., 2001). Evolutionary distance matrices were determined using Kimura 2-parameter methods (Kimura, 1980) and based on these matrices dendrograms were inferred using the neighbour-joining (NJ), minimum evolution (ME) and maximum parsimony (MP) analyses. The stability of the predicted branches was determined using bootstrap analysis. Bootstrap values were obtained for a consensus tree based on 1000 randomly generated trees using the same program.

4.3 RESULTS

4.3.1 Sequence analysis .

The rickettsial isolate from the tick culture was 100% homologous with Rickettsia sp. strain BWI-1, previously characterized in Chapter 3 based on citrate synthase and ompA gene segments. It was therefore assumed that Rickettsia sp. strain BWI-1 had been isolated successfully. Segments of the 16S rRNA (1293 bp), gltA (1098 bp), ompA (548 bp), ompB (451 bp) and sca4 (758 bp) genes were then sequenced and compared to previously characterized rickettsial species. These sequences have been submitted to GenBank (Appendix 5).

The percentage similarity between Rickettsia sp. strain BWI-1 and the previously characterized rickettsial species is displayed in Table 4.5. Rickettsia sp. strain BWI-1 consistently shows greatest sequence similarity to the R. massiliae subgroup of the

89 spotted fever group ( R. massiliae , R. massiliae strain Bar 29, R. rhipicephali , R. aeschlimanni and R. montanensis (Roux et al ., 1997; Fournier et al ., 1998)). Based on the 16S rRNA gene sequences, Rickettsia sp. strain BWI-1 is most closely related to

R. massiliae and R. rhipicephali at 99.7% sequence similarity. Based on the gltA gene sequences it is most closely related to R. aeschlimannii at 98.4% sequence similarity.

OmpA gene sequences place Rickettsia sp. strain BWI-1 closest to R. massiliae at

95.8% similarity. Rickettsia sp. strain BWI-1 is 97.4% similar to R. rhipicephali based on its ompB gene sequence and 96.5% similar to R. aeschlimannii based on its sca4 gene sequence.

For most of the genes, the three methods of dendrogram inference (NJ, ME and MP) produced trees with similar arrangements (Appendix 6), the main difference being the level of bootstrap support for the branches. Rickettsia sp. strain BWI-1 appears most commonly on a branch of its own lying just outside the main clustering of the R. massiliae subgroup (Figures 4.1- 4.5).

90

Table 4.5. Percentage similarity between Rickettsia sp. strain BWI-1 and Rickettsia species with validly published names.

Name % Similarity GltA ompA ompB sca4 16S rRNA 98.8% 92.4% NS 76.7% 76.5% R. prowazekii 98.7% 92.2% NS 81.5% 79.7% R. typhi 99.5% 97.6% 93.8% 92.7% 95.3% R. africae 99.5% 98.1% 92.7% 91.9% 95.3% R. conorii 99.5% 98.2% 93.4% 93.2% 95.6% R. sibirica 99.5% 98.0% 93.2% 92.3% 96.0% R. honei 99.5% 98.1% 94.0% 95.6% 96.0% R. slovaca 99.4% 98.2% 93.2% 92.5% 95.2% R. parkeri 99.5% 97.8% 92.9% 94.5% 95.9% R. rickettsii 99.7% 98.0% 95.8% 97.1% 95.8% R. massiliae 99.7% 97.9% 95.6% 97.4% 95.7% R. rhipicephali 99.5% 98.4% 95.1% 96.9% 96.5% R. aeschlimannii 99.3% 98.3% 94.5% 96.0% 95.3% R. montanensis 99.5% 97.6% 93.4% 93.8% 96.0% R. japonica 99.4% 98.0% 94.3% 93.8% 96.3% R. heilongjiangii 99.2% 95.8% NS 51.8% 89.2% R. helvetica 99.5% 93.4% 53.7% NS 88.6% R. felis 98.6% 93.4% NS 85.7% 84.7% R. akari 99.4% 93.7% 85.2% 87.7% 86.0% R. australis 99.2% 85.7% NS NS NS R. bellii

NS – not sequenced.

91

R. conorii R. parkeri

R. sibirica R. africae R. rickettsii

R. honei R. slovaca R. japonica

93 R. heilongjiangii R. montanensis Rickettsia sp. BWI-1 R. aeschlimannii R. massiliae R. rhipicephali R. prowazekii 100 R. typhi R. helvetica R. felis R. bellii

R. akari R. australis

0.2%

Figure 4.1. Unrooted neighbour-joining tree based on 16S rRNA sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterized rickettsiae. The percentage of similarity between the species was determined using the INFOALIGN program in the EMBOSS software package (Williams, 2001). Sequence similarities were calculated based on pairwise transitions and transversions between sequences. Phylogenetic and molecular evolutionary analyses were conducted using MEGA version 2.1 (Kumar et al ., 2001). Evolutionary distance matrices were determined using Kimura 2-parameter methods (Kimura, 1980). Bootstrap values were obtained for a consensus tree based on 1000 randomly generated trees using the same program.

Scale indicates percentage divergence. Bootstrap values >70% are indicated at the nodes. The strain of interest in this study is highlighted in red, the other species of rickettsiae recorded in Australia are highlighted in blue.

92

R. rickettsii R. conorii

R. honei R. africae R. sibirica

R. parkeri R. slovaca R. japonica

98 R. heilongjiangii R. aeschlimannii R. massiliae 78 97 R. rhipicephali 99 Rickettsia sp. BWI-1 94 R. montanensis 87 R. helvetica R. felis R. akari 98 99 R. australis R. prowazekii

R. typhi 100 R. bellii

2%

Figure 4.2. Neighbour-joining tree based on gltA gene sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterised rickettsiae. Scale indicates percentage divergence. Bootstrap values >70% are indicated at the nodes. The strain of interest in this study is highlighted in red, the other species of rickettsiae recorded in Australia are highlighted in blue.

.

93

71 R. africae R. parkeri R. sibirica R. conorii R. slovaca 99 R. honei 98 R. rickettsii R. japonica 100 R. heilongjiangii R. aeschlimannii 100 R. massiliae 93 R. rhipicephali Rickettsia sp. BWI-1 R. montanensis R. australis

2%

Figure 4.3. Neighbour-joining tree based on ompA gene sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterised rickettsiae. Scale indicates percentage divergence. Bootstrap values >70% are indicated at the nodes. The strain of interest in this study is highlighted in red, the other species of rickettsiae recorded in Australia are highlighted in blue.

.

94 R. africae R. parkeri 96 R. sibirica 88 97 R. mongolotimonae 76 R. conorii 99 R. slovaca R. rickettsii R. honei R. japonica 99 100 R. heilongjiangii Rickettsia sp. BWI-1 R. montanensis R. aeschlimannii R. massiliae 71 R. rhipicephali R. akari 98 R. australis R. prowazekii 100 R. typhi

5%

Figure 4.4. Neighbour-joining tree based on ompB gene sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterised rickettsiae. Scale indicates percentage divergence. Bootstrap values >70% are indicated at the nodes. The strain of interest in this study is highlighted in red, the other species of rickettsiae recorded in Australia are highlighted in blue.

.

95

R. conorii R. parkeri 74 R. sibirica

R. africae Error! 84 R. slovaca R rickettsii 81 R. honei R. japonica 97 R. heilongjiangii R. montanensis 100 R. aeschlimannii R. massiliae R. rhipicephali Rickettsia sp . BWI-1 R. felis R. akari 99 100 R. australis

R. helvetica R. prowazekii 100 R. typhi

5%

Figure 4.5. Neighbour-joining tree based on sca4 sequences demonstrating the relationship between Rickettsia sp. BWI-1 and previously characterised rickettsiae. Scale indicates percentage divergence. Bootstrap values >70% are indicated at the nodes. The strain of interest in this study is highlighted in red, the other species of rickettsiae recorded in Australia are highlighted in blue.

.

96 4.4 DISCUSSION

These results demonstrate that Rickettsia sp. strain BWI-1 is sufficiently divergent to be classified as a novel species and they also provide some information on the possible origin of the species and the evolutionary route that Rickettsia sp. strain

BWI-1 may have taken from its ancestors.

Although bootstrap support for the phylogenetic position of Rickettsia sp. strain BWI-

1 is not strong (>70) for most of the gene sequences examined, its placement is fairly consistent when examining different genes with different evolutionary origins, giving weight to the results.

Taking all the gene sequences into account, Rickettsia sp. strain BWI-1 is more closely related to members of the R. massiliae subgroup of the SFG than to the

Australian SFG species. This is not totally unexpected however as previous studies have found that the other species of Australian SFG rickettsiae are also quite distinct from one another. Rickettsia australis is a comparatively divergent member of the

SFG while R. honei is closely related to species in the core SFG branch in phylogenetic analyses. It is estimated that that the divergence of R. honei and R. australis occurred 65 million years ago (Stenos et al ., 1998). It seems unlikely therefore that rickettsiae in the eastern states of Australia evolved from a recent common ancestor as might be expected given their close proximity, instead it has been suggested that the occurrence of R. honei in southeast Australia could be the result of migrating birds carrying infected ticks (Stenos et al ., 1998).

97 Similarly, it also seems likely that Rickettsia sp. strain BWI-1 does not share a recent common ancestor, and has not evolved in parallel, with either of the recognised

Australian species of SFG. An alternative explanation for its occurrence must therefore be hypothesised. Of interest is that Rickettsia sp. strain BWI-1 occurs both on the mainland of WA and on Barrow Island, which has been separated from the mainland since the last ice age approximately 8000 years ago. It therefore represents a rickettsia that has not evolved significantly during this time. Alternatively, a breach in

BWI quarantine may have occurred, with ticks moving off BWI onto the mainland or vice versa. Due to the wide range of this rickettsial species however, the most likely scenario is that the species was present in WA prior to the island being formed.

A closer examination of the SFG subgroup with which Rickettsia sp. strain BWI-1 clusters provides some information on its characteristics and possible evolutionary route. The R. massiliae subgroup contains rickettsiae of unknown ( R. rhipicephali and

R. montanensis ), or recently recognised, pathogenicity ( R. massiliae, R. massiliae strain Bar 29 and R. aeschlimannii ). Rickettsia massiliae DNA was recently identified in the blood of an Italian man who had presented with an eschar and macropapular rash 20 years previously. A rickettsia had been cultured and stored from a blood sample collected at the time of presentation, making this retroactive study possible

(Vitale et al ., 2005). Rickettsia massiliae strain Bar 29 has been detected in tick saliva, and a human serosurvey performed in Spain identified R. massiliae strain Bar

29 antibodies, both of these factors imply a possible pathogenic role although no human cases have been reported to date (Parola et al ., 2005). The pathogenicity of R. aeschlimannii was not established until 2002 when an infection was reported in a patient returning from Morocco to France. An infection was later reported from a

98 patient in South Africa. Rickettsia aeschlimannii produces a disease characterized by an eschar, fever and a generalized maculopapular skin rash (Roux et al ., 1997;

Cardenosa et al ., 2003; Raoult, 2004).

Members of the R. massiliae subgroup of SFG have been identified in Europe, Africa,

North and South America, the Astrakhan region, Siberia, Kazakhstan, Thailand and

Japan (Babalis et al ., 1994; Beati et al ., 1996; Azad and Beard, 1998; Rydkina et al .,

1999; Bernasconi et al ., 2002; Pertorius and Birtles, 2002; Satoh et al ., 2002;

Cardenosa et al ., 2003; Parola et al ., 2003; Ammerman et al ., 2004; Morita et al .,

2004; Labruna et al ., 2005; Eremeeva et al., 2006). These last three locations are the closest geographically to Western Australia and although it has been suggested that rickettsiae readily switch to different tick hosts over evolutionary time (Weller et al .,

1998), the rickettsiae in Kazakhstan were detected in Haemaphysalis spp. tick vectors and those in Japan were detected in Amblyomma spp. which are both genera of vectors from which Rickettsia sp. strain BWI-1 has been amplified in this study. The

Rickettsia spp. identified in Thailand were detected in Dermacentor spp. (Parola et al ., 2003). It is possible that the presence of closely related species of rickettsiae in

Kazakhstan, Thailand, Japan and Australia may be the result of dissemination of tick vectors (not necessarily this particular tick species) through the movement of migratory birds as suggested in other instances (Stenos et al ., 1998; Kelly, 2006;

Santos-Silva et al ., 2006).

The pathogenic potential of Rickettsia sp. strain BWI-1 is, at this stage unknown.

Although several people working on Barrow Island have presented with influenza-like clinical signs and rash characteristic of a rickettsiosis (J. Drummond, pers. comm.,

99 2003) diagnostic confirmation was not performed at the time. The R. massiliae subgroup contains rickettsiae of both known and unknown pathogenicity however, the pathogenic members were initially classified as rickettsiae of unknown pathogenicity before subsequent reclassification. Another factor that increases the likelihood that

Rickettsia sp. strain BWI-1 is a human pathogen is its presence in an anthropophilic tick species, Amblyomma triguttatum (Roberts, 1970; Pearce and Grove, 1987; Storer et al ., 2005) . The statement has been made that all rickettsiae utilising tick vectors that feed on people are capable of infecting people (Raoult, 2004).

It has been speculated previously that a mechanism exists by which non-pathogenic symbiotic rickettsiae that possess an ompA gene may evolve into pathogenic strains through changes in their outer membrane proteins (Weller et al ., 1998), so even if evidence to support the pathogenicity of Rickettsia sp. strain BWI-1 is lacking in this study this is by no means conclusive, and does not discount the need for further research on the rickettsia and its possible disease syndromes.

Description of a new species:

Given the results of experiments detailed in this chapter, we therefore propose the creation of a new species name for Rickettsia sp. strain BWI-1: Rickettsia gravesii

(graves.i.i. L. gen. n. gravesii ) after Stephen Graves, an Australian pathologist and microbiologist who is the founder and director of The Australian Rickettsial

Reference Laboratory and who has a long history of promoting and contributing to

Australian rickettsial research.

100 Rickettsia gravesii sp. nov. grows on XTC-2 cells at 28 °C in Leibovitz L-15 media supplemented with 5% foetal calf serum, 0.4% tryptose phosphate and 200mM L- glutamine.

The type strain is strain BWI-1. This was isolated from an Amblyomma triguttatum tick from Barrow Island, Western Australia in 2004 and the isolate has been deposited in the Australian Rickettsial Reference Laboratory collection. Preparations are underway to also deposit it in the Collection de souches de L’Unité des Rickettsies

(CSUR), World Health Organization Collaborative Center for Rickettsioses,

Borrelioses and Tick-borne Infections, Marseilles, France.

101 CHAPTER 5

DETECTION AND CHARACTERIZATION OF NOVEL RICKETTSIAE FROM YELLOW-FOOTED ANTECHINUSES

5.1 INTRODUCTION

The role of wildlife as reservoirs of rickettsioses and bartonelloses has been widely documented from many countries, however it is still poorly understood in Australia

(Niebylski et al., 1999; Birtles et al., 2001; Boostrom et al ., 2002; Jacomo et al .,

2002; Holmberg et al. , 2003; Bown et al. , 2004; Kim et al. , 2006). Yellow-footed antechinuses (mardos, Antechinus flavipes) are small, mainly insectivorous marsupials that are native to Australia. There are several subspecies and these are found in a variety of habitats across Queensland, New South Wales, Victoria, South Australia and the southwest of WA. All the males of the species die after two weeks of intensive mating activity in August (Menkhorst and Knight, 2001). The male mortality is precipitated by a failure of the animals’ glucocorticoid feedback mechanism. The resulting excess of corticosteroids produces immunosuppression and other stress-related pathology including gastric bleeding, negative nitrogen balance and hypoglycaemia (McAllan et al ., 1998).

During a concurrent study to examine the ecology of yellow-footed antechinuses, ectoparasites were collected and a novel rickettsia ( Rickettsia sp. strain D-1) was identified (described in Chapter 3). The aim of this section of the study was to further

102 characterize Rickettsia sp. strain D-1 and to determine whether yellow-footed antechinuses and their ectoparasites harbour other potentially zoonotic rickettsiae.

5.2 MATERIALS AND METHODS

5.2.1 Sample collection

Additional trapping was performed in dry sclerophyll forests surrounding the town of

Dwellingup in the southwest of WA, an area of interest identified in chapter 2 and 3.

The methods used for trapping and ectoparasite removal have been described previously in Chapter 2.

5.2.2 Ectoparasite identification

Ticks were identified by light microscopy according to a standard key (Roberts,

1970). Similarly, fleas were identified using light microscopy and a standard taxonomic key (Dunnet and Mardon, 1974).

5.2.3 Cell culture

Culture from the tick samples was attempted. The methods for cell culture are described in section 4.2.2.

5.2.4 DNA extraction

The protocol for extraction of DNA from ticks has been described in Chapter 3.

DNA was extracted from the fleas using Qiagen minikis. Up to five fleas from an individual animal were pooled to increase the efficiency of the screening process.

DNA was extracted from fleas following the manufacturer’s tissue protocol, with the exception that the DNA was stored in 70 L of elution buffer rather than 200 L as

103 recommended by the manufacturer. This modification was made in order to increase the final concentration of DNA.

5.2.5 PCR screening

The methods used for PCR screening have been described previously in Chapter 3.

5.2.6 Sequencing

The methods used for sequencing have been described previously in Chapter 4. The sequences generated were initially compared to those in Genbank using the BLAST program. Samples that showed sequences similar to those of previously characterized

SFG rickettsiae were further sequenced using gltA , ompA and ompB gene primers. If the BLAST analysis indicated that the positive sample was a Bartonella spp., for which there is no amplifiable ompA or ompB gene, only gltA sequencing was performed.

Repeated attempts to culture the Rickettsia spp. identified ( Rickettsia sp. strain D-1) from the ticks were unsuccessful therefore no 16S rRNA sequencing was undertaken.

As the 16S rRNA gene is highly conserved across the eubacteria, attempts to sequence it directly from tick samples could result in the amplification of endosymbionts that are often present in ticks, leading to confusing results (Edwards et al ., 1989; Noda et al ., 1997; Stenos et al ., 2005).

104 5.2.7 Sequence Analysis

The methods for sequence analysis have been described previously in Chapter 4.

5.3 RESULTS.

5.3.1 Prevalence

Ectoparasites were collected in 48 instances of trap success. This figure includes the animals described in chapter 2 and 3 and animals trapped on subsequent field trips.

Thirty six individual animals were captured and this number includes initial trapping of an animal and any subsequent re-trappings (table 5.1). Two species of flea (38/48 of the animals with ectoparasites carried Acanthopsylla jordani ; Echidnophagia perilis was collected from one animal ) and one species of tick (27/48 of the animals with ectoparasites had Ixodes antechini ) were collected from the yellow-footed antechinuses. In 17 instances, an animal was co-infected with A. jordani and I. antechini .

Following screening with gltA gene primers, fourteen of the twenty seven animals infested with Ixodes antechini (51.9%) had PCR positive ticks and 44.7% (17/38) of animals with Acanthopsylla jordani had PCR positive fleas. The single E. perilis specimen collected was PCR negative. Of the animals that were co-infected with two species of ectoparasite, there were 6 cases of PCR positive A. jordani and PCR negative I. antechini, 7 cases of positive I. antechini and negative A. jordani, 2 instances in which both ectoparasites were positive and 2 where they were both negative (table 5.1).

105 Table 5.1 Results of PCR screening of ticks and fleas collected from yellow-footed antechinuses in Dwellingup, WA.

Individual Number of times sampled Result for PCR screening 1 1 Negative (I.a.) 2 1 Negative (I.a) 3 4 Positive (A.j) Negative (A.j.) Positive (A.j), negative (I.a) Negative (A.j.), positive (I.a.) 4 1 Negative (A.j.) 5 1 Negative (I.a.) 6 1 Negative (I.a.) 7 1 Negative (I.a.) 8 1 Positive (A.j.) 9 3 Positive (A.j.), negative (I.a.) Negative (A.j.), positive (I.a.) Negative (A.j.) 10 1 Positive (A.j.), negative (I.a.) 11 1 Positive (A.j.) 12 1 Negative (A.j.) 13 2 Positive (A.j.) Positive (A.j), negative (I.a.) 14 3 Positive (A.j.) Negative (A.j.), positive (I.a) Negative (A.j.), positive (I.a) 15 1 Positive (A.j.), positive (I.a.) 16 2 Negative (A.j.) Positive (A.j.), negative (I.a) 17 1 Negative (A.j.) 18 1 Negative (A.j.) 19 1 Negative (A.j.), negative (I.a) 20 1 Negative (A.j.), Positive (I.a) 21 1 Positive (A.j.) 22 1 Negative (A.j.), Positive (I.a) 23 2 Positive (I.a.) Positive (A.j.), positive (I.a.) 24 1 Positive (I.a.), negative (A.j) 25 1 Negative (A.j.) 26 2 Negative (A.j.), negative (I.a.) Positive (I.a.) 27 1 Positive (A.j.) 28 1 Positive (I.a.) 29 1 Positive (I.a.) 30 1 Positive (A.j.), positive (I.a.) 31 1 Negative (A.j.) 32 2 Positive (A.j.), negative (I.a.) Negative (A.j.) 33 1 Negative (A.j.) 34 1 Positive (A.j.) 35 1 Negative (E. perilis), negative (A.j.) 36 1 Negative (A.j.)

I.a. – Ixodes antechini ; A.j. – Acanthopsylla jordani

106

Figure 5.1. Acanthopsylla jordani : lateral view.

5.3.2 Sequence similarities

The sequences from the novel strains have been submitted to GenBank (Appendix 5).

Sequencing of multiple PCR positive tick samples revealed identical sequences. The percentage nucleotide similarity between the novel strains identified in this study and previously characterized Rickettsia and Bartonella species are shown in tables 5.2 and

5.3.

Sequencing of the gltA gene segment from multiple PCR positive samples obtained from A. jordani revealed that the fleas were infected with a Bartonella spp. which is

107 most closely related to Bartonella sp. strain 40 at 86.1% sequence similarity. Based on its gltA gene, Rickettsia sp. strain D-1 is most closely related to R. aeschlimannii, demonstrating 99.4% sequence similarity. Comparison of ompA gene sequences demonstrates a 95.6% sequence similarity to R. massiliae and R. slovaca and it is

97.1% similar to R. rhipicephali based on its ompB gene. The phylogenetic position of the Rickettsia sp. strain D-1 based on analysis of these genes is demonstrated in figures 5.2 – 5.5 (and ME and MP trees in appendix 6). The phylogenetic position of the novel Bartonella species ( Bartonella sp. strain Mu1) based on gltA sequences is demonstrated in figure 5.6.

108 Table 5.2. Comparison of Rickettsia sp. strain D-1 and Rickettsia species with validly published names based on 3 genes.

Name GltA ompA ompB

98.4% 96.3% 96.7% R. gravesii R. prowazekii 92.9% NA 76.0%

R. typhi 92.6% NA 80.6%

R. africae 98.8% 95.2% 92.3%

R. conorii 99.1% 93.9% 90.5%

R. sibirica 99.3% 95.2% 92.3%

R. honei 99.1% 94.8% 93.2%

R. slovaca 99.2% 95.6% 94.1% R. parkeri 99.2% 94.3% 91.9%

R. rickettsii 98.7% 93.7% 93.8%

R. massiliae 99.1% 95.6% 96.9% 99.0% 95.4% 97.1% R. rhipicephali

R. aeschlimannii 99.4% 94.6% 96.3% R. montanensis 99.2% 93.9% 95.6% R. japonica 98.7% 94.1% 93.8%

R. heilongjiangii 99.0% 95.0% 93.3%

R. helvetica 96.6% NA 52.1%

R. felis 94.5% 53.9% NA

R. akari 93.9% NA 85.0%

R. australis 94.8% 85.2% 87.7%

R. bellii 86.8% NA NA

109 Table 5.3. Comparison of Bartonella sp. strain Mu1 and previously characterized species of Bartonella based on gltA gene sequences.

Name % Similarity 84.4% B. quintana

85.7% B. henselae

B. koehlerae 85.2%

B. bacilliformis 80.2%

B. doshiae 83.5%

B. elizabethae 81.9%

B. grahamii 83.1%

B. tribocorum 82.7%

B. vinsonii 84.8% B. clarridgeiae 84.4%

B. taylorii 83.1%

B. peromysci 35.4%

Bartonella sp . Strain 1phy 84.8% Bartonella sp . Strain 4phy 83.1%

Bartonella sp. Strain R phy1 85.2%

Bartonella sp . Strain phy2 83.1%

Bartonella sp. Strain C4phy 83.1%

Bartonella sp. Strain C7rat 81.9%

Bartonella sp . Strain 40 86.1%

110

R. rickettsii R. conorii

R. honei R. africae R. sibirica

R. parkeri Error! R. slovaca R. japonica

99 R. heilongjiangii R. aeschlimannii 83 R. massiliae 95 R. rhipicephali 100 R. gravesii 93 Rickettsia sp. D-1 R. montanensis 82 R. helvetica R. felis R. akari 98 R. australis 99 R. prowazekii 100 R. typhi R. bellii 2%

Figure 5.2. Neighbour-joining tree based on gltA sequences comparing the novel strain and previously characterised rickettsiae.

Scale indicates percentage divergence. Bootstrap values >70% are indicated at the nodes. The strain of interest in this study is highlighted in red, the other species of rickettsiae recorded in Australia are highlighted in blue.

111

75 R. africae

R. parkeri 70 R. sibirica 99 R. conorii

R. slovaca Error! 93 R. rickettsii R. honei 93 R. japonica 100 R. heilongjiangii Rickettsia sp. D-1 R. aeschlimannii 84 100 R. massiliae 94 R. rhipicephali R. gravesii R. montanensis

R. australis

2%

Figure 5.3. Neighbour-joining tree based on ompA sequences comparing the novel strain and previously characterised rickettsiae.

Scale indicates percentage divergence. Bootstrap values >70% are indicated at the nodes. The strain of interest in this study is highlighted in red, the other species of rickettsiae recorded in Australia are highlighted in blue.

112

R. africae 94 R. parkeri R. sibirica 91 96 R. mongolotimonae 77 R. conorii 99 R. slovaca

R. rickettsii R. honei R. japonica 100 100 R. heilongjiangii R. gravesii R. montanensis Rickettsia sp. D-1 R. aeschlimannii R. massiliae R. rhipicephali R. akari

98 R. australis R. prowazekii 100 R. typhi

5%

Figure 5.4. Neighbour-joining tree based on ompB sequences comparing the novel strain and previously characterised rickettsiae.

Scale indicates percentage divergence. Bootstrap values >70% are indicated at the nodes. The strain of interest in this study is highlighted in red, the other species of rickettsiae recorded in Australia are highlighted in blue.

113

B. bacilliformis

B. doshiae B. clarridgeiae Bartonella sp. Mu1 B. quintana B. henselae

B. koehlerae Bartonella sp. Strain40 B. taylorii

76 B. vinsonii 99 Bartonella sp. StrainC1phy Bartonella sp. StrainR1phy

70 Bartonella sp. StrainC4phy 90 Bartonella sp. StrainR2phy B. tribocorum

B. grahamii Bartonella sp. StrainC7rat B. elizabethae B. peromysci

20

Figure 5.5. Neighbour-joining tree of Bartonella spp. based on gltA sequences comparing the novel strain and previously characterised Bartonella spp.

Scale indicates percentage divergence. Bootstrap values >70% are indicated at the nodes. The strain of interest in this study is highlighted in red, the other species of rickettsiae recorded in Australia are highlighted in blue.

114 5.4 DISCUSSION

5.4.1 Aetiology

Attempts to cultivate Rickettsia sp. strain D-1 from ticks in XTC and Vero cell lines were unsuccessful. This could be due to laboratory factors however it is also possible that an inability to grow in these cell lines is a phenotypic property of the strain. This is the case for R. peacockii, which has only been successfully propagated in tick cell lines (Simser et al ., 2001). Rickettsia sp. strain D-1 has been assigned the name

Candidatus “Rickettsia antechini” following instructions given for the naming of bacteria that are yet to be cultured (Murray and Stackenbrandt, 1995; Raoult et al.,

2005). The culture of Bartonella sp. strain Mu1 was deemed to be outside the scope of this project, it will be performed as part of future work.

The identification of novel Rickettsia and Bartonella species and the examination of their evolutionary lineages increases our understanding of the natural history of the genera, their ecology and their mechanisms of pathogenicity. As previously described, in order to be classified as a novel species of rickettsia, Fournier et al (2003) recommended that an isolate should not exhibit more than one of the following degrees of nucleotide similarity with the most homologous validated species: ≥ 99.8,

≥99.9%, ≥98.8%, ≥99.2% and ≥99.3% for the 16S rRNA, gltA , ompA, ompB and sca4 genes respectively. Based on the results from this study therefore, it is likely that

Candidatus “Rickettsia antechini” constitutes a novel species, however insufficient genotypic information has been gathered to propose it as such at this stage.

Candidatus “Rickettsia antechini” falls within the R. massiliae subgroup of the SFG, and is closely related to R. gravesii, the other novel species identified during this

115 study. Rickettsia gravesii has also been detected in geographical locations close to

Dwellingup, the yellow-footed antechinus study site. It seems possible therefore that

R. gravesii and Candidatus “R. antechini” could have evolved from a recent common ancestor.

The classification of the genus Bartonella is less well defined (Birtles and Raoult,

1996). It has been determined that rpoB and gltA gene sequences provide the most sensitive indication of the phylogenetic position of a Bartonella species (La Scola et al . 2003). The value of other genes is limited by the high degree of sequence similarity in the gene fragments between species and the lack of sequence data across the range of Bartonella species (Houpikian and Raoult, 2001; La Scola et al ., 2003).

Phylogenetic studies based on gltA gene sequences have demonstrated a range of similarities between previously characterized Bartonella species (from 83.7% to

93.2%) and it has been suggested that a newly detected isolate should be considered a novel species if it demonstrates <96.0% similarity to the corresponding 327 bp gltA fragment of previously validated species (Birtles and Raoult, 1996; La Scola et al.,

2003). Based on this criterion, Bartonella sp. strain Mu1 falls within the genus and is sufficiently divergent to be classified as a novel species. Further characterisation based on sequencing of the genes mentioned above (in particular rpoB ) was not performed due to difficulties in collecting follow-up samples, this should be performed in the future in order to more accurately classify this novel Bartonella species.

Bartonella henselae has been widely reported from Australian cats (Branley et al.,

1996; Rathbone et al ., 1996) and a single case of Bartonella quintana infection has

116 been reported in an immunocompromised patient in 1996 (Branley et al., 1996;

Rathbone et al ., 1996). From the sequences generated it seems unlikely that

Bartonella sp. strain Mu1 shares a recent common ancestor with other Australian members of the genus and the Bartonella species occurring in Australia are probably the result of multiple independent introductions.

The zoonotic potential of the two novel species identified are presently unknown. A previous serological survey of people in the southwest of the state detected the presence of SFG antibodies so it is likely that at least one infectious Rickettsia species is present in the area (Kilminster T. An Investigation of Typhus in Western Australia.

Honours Thesis: Department of Microbiology, University of Western Australia;

1997). Further investigations need to be performed to determine whether the rickettsia detected in this study is responsible for the sero-conversion or whether there are other, yet to be isolated, species of rickettsia present.

5.4.2 Ecology

Sylvatic life cycles for infectious agents are particularly advantageous, involving, in many cases, reservoirs of infection that are not reliant on or contained by human activity. Both ectoparasites and vertebrates are utilised in order to maintain infection in these sylvatic life cycles and to allow the transition into domestic life cycles.

The potential vector of Candidatus “R. antechini”, Ixodes antechini, has not been recorded from WA previously (Roberts, 1970). In addition, there is a limited amount of information available on the flea species Acanthopsylla jordani, which has previously only been recorded from dunnarts ( Smithopsis murina ) in the Dryandra

117 Forestry Reserve (Dunnet and Mardon, 1974). Therefore we cannot speculate as to whether Candidatus “R. antechini” and Bartonella sp. strain Mu1 are likely to occur elsewhere in WA.

The role of the yellow-footed antechinus in the life cycles of Candidatus “R. antechini” and Bartonella sp. strain Mu1 life cycles is unclear, as it is yet to be determined if they are competent hosts for either of these micro-organisms (i.e. susceptible to infection, able to support effective proliferation of the micro-organisms and infective to the vectors for a significant period of time (Richter et al ., 2000)). No intra-erythrocytic Bartonella spp. were detected microscopically in blood smears obtained from the ears of yellow-footed antechinus following collection of tissue samples for DNA analysis (P. Clark, pers. comm.., 2003); however, it has been previously noted that conventional staining and microscopy is a relatively insensitive technique for Bartonella spp. detection and even high levels of bacteraemia can go undetected (Kordick et al ., 1999). Inherent limitations involved with working on small native animals in the field prevented the collection of larger volumes of blood for serology, molecular based screening and blood culture at this stage.

It has been hypothesized that co-evolution may have occurred between bacteria and their mammalian reservoir hosts and information on novel Bartonella spp. will go some way to proving or disproving this hypothesis (Houpikian and Raoult, 2001;

Jacomo et al ., 2002). Based on the limited sequence information obtained in this study, the Bartonella sp. strain Mu1 is most closely related to Bartonella spp. strain

40 which was detected in wood mice ( Apodemus sylvaticus ) in the United Kingdom.

This result would therefore support the hypothesis of co-evolution of bacteria and

118 host. Wood mice and yellow-footed antechinuses share many common features, as opposed to other vertebrate hosts used by Bartonella spp. such as dogs and coyotes

(B. visonii subsp. berkhoffii ), cattle ( B. bovis ), humans ( B. quintana and B. bacilliformis ) and cats ( B. koehlerae, B. clarridgeiae, B. henselae and B. bovis )

(Breitschwerdt and Kordick, 2000; Jacomo et al ., 2002).

It would also be interesting to examine the role of rickettsiae and Bartonella spp. in the annual male yellow-footed antechinus die off. It has been demonstrated that some

Bartonella and Rickettsia spp. can be pathogenic to their vertebrate hosts (Kelly et al .,

1992a; O'Reilly et al ., 1999; Paddock et al ., 2002), and it is therefore possible that infection with these agents could contribute to the demise of severely immunosuppressed male yellow-footed antechinuses.

The zoonotic potential of Candidatus “R. antechini” and Bartonella sp. strain Mu1 is unknown. As stated previously, a previous serological survey in the southwest of the state detected the presence of SFG antibodies in human sera so it is likely that an infectious rickettsia is present in the area (Kilminster T. An Investigation of Typhus in

Western Australia. Thesis: Department of Microbiology, University of Western

Australia; 1997). Nevertheless, further investigation needs to be performed to determine whether the rickettsia detected in this study is responsible for this seroconversion or whether there are other, yet to be isolated species of rickettsia in the area.

119 CHAPTER 6

INVESTIGATION OF THE PATHOGENIC POTENTIAL OF RICKETTSIA GRAVESII AND THE RISK FACTORS

FOR ZOONOTIC INFECTION

6.1 INTRODUCTION

Determining the pathogenicity of a novel Rickettsia spp. is an important step in understanding its role in infections and will largely influence the nature of any future work on the organism. The emphasis of research on pathogenic rickettsiae is more likely to focus on their zoonotic significance, epidemiology, control and prevention, whereas future studies relating to non-pathogenic Rickettsia spp. are aimed at defining mechanisms that allow non-pathogenic organisms to emerge as pathogens and may provide information on how non-pathogenic endosymbionts could be manipulated for control of ticks and the pathogens they harbour (Simser et al ., 2001).

The many technical difficulties involved in the detailed study of rickettsial gene functions have prevented the definitive identification of their virulence factors thus far

(Renesto et al. , 2005). It is not yet possible therefore to classify a novel isolate as pathogenic or non-pathogenic based on genotypic or proteomic methods: the pathogenicity of a Rickettsia spp. can only be definitively established after its isolation from a patient presenting with the clinical signs of a rickettsiosis (Sarov et al. , 1992; Paddock et al ., 2004).

The main objective of this preliminary study was to explore the pathogenic potential of Rickettsia gravesii sp. nov . by investigating its links to any clinical signs reported

120 in people living or working in an area of endemnicity (as established by the work described in previous chapters).

6.2 MATERIALS AND METHODS

6.2.1 Study design

The study design used was a combined serosurvey and retrospective cohort study using an internal control group. The serological survey was based on the collection of one blood sample from each participant. Exposure and symptom data for the retrospective cohort analysis were collected utilising a questionnaire.

Barrow Island was chosen as the site of the study as it provided a well-defined geographical area with an established R. gravesii presence. The island, a class A nature reserve, contains an abundance of native animals and therefore a large tick population. The island’s human population consists entirely of employees of a mining company and scientists monitoring the impact of the mining activities on the environment, thus providing an easily contactable and organised population to study.

People who work on Barrow Island also represent a high-risk population because much of their work is performed in the bush adjacent to the wildlife. These people are not permanent residents on the island, instead spending a discrete period of time on the island, usually 2 weeks, alternating with 2 weeks on the mainland. The only criterion for participation in the study was having recently spent time on the island.

This work was approved by Murdoch University’s Human Ethics committee.

121 6.2.1 Questionnaire design

The main objective of the questionnaire was to establish whether people on the island were being exposed to a tick-borne rickettsia and whether infection was associated with any characteristic symptoms. The symptoms being investigated were chosen based on descriptions of clinical signs commonly displayed by people with established rickettsioses.

A second objective of the questionnaire was to obtain demographic information and to identify other factors that could increase the respondent’s exposure to tick-borne illnesses both on the island (job description, time spent on the island, time spent in the bush and exposure to potential vertebrate hosts of ticks), on the Western Australian mainland, interstate or overseas (Appendix 7).

6.2.3 Questionnaire administration

Two sessions were arranged with a two week interval to administer the questionnaires

(22.6.06 and 6.7.06) in order to give both shifts of workers the opportunity to participate in the survey. A presentation explaining the project was provided for the group, and then interested volunteers filled out the self-administered questionnaire over approximately 20-30 minutes. Informed consent forms were signed by all study participants (Appendix 7).

6.2.4 Blood collection

In addition to filling out the questionnaire, participants were asked to submit a blood sample at each session, collected at the island’s medical centre by a registered nurse hired for the purpose. Approximately 8 mls of blood was collected from each

122 participant and allowed to clot at room temperature before serum was removed, aliquoted and stored at 4 oC prior to testing. The serology was performed at the

Australian Rickettsial Reference Laboratory, Geelong, by Yazid Abdad, PhD candidate, School of Veterinary and Biomedical Sciences, Murdoch University as part of his PhD thesis. He kindly permitted the inclusion of his preliminary results for analysis alongside the questionnaire. A standard micro-immunofluorescence assay

(IFA) was utilised to test the reactivity of the serum to R. australis , R. honei , R. conorii , R. sibirica , R. rickettsii , R. akari , R. typhi and R. prowazekii antigens (Graves et al , 1999). Although a titre of 1:64 is usually accepted as the cut-off for positive sera, a cut-off of 1:128 was used in this study so that the influence of cross-reacting antibodies would be minimised (Walker and Dumler, 1995; Graves et al , 1999).

6.2.5 Analysis

Microsoft Excel, SPSS 14 and Stata 9.0 were used for data management, descriptive analysis and the calculation of logistic regression and odds ratios. Three categories of symptoms were proposed as being consistent with a rickettsiosis and respondents who fell into any of these categories were considered to have had a syndrome “consistent with a rickettsiosis” and were grouped together for the analysis. The proposed definition of “consistent with a rickettsiosis” was kept reasonably non-specific in order to avoid false-negative results. Thus, based on a review of the literature describing clinical signs associated with rickettsioses (section 1.1.3.3) respondents who reported at least two symptoms from rash, fever, headache, aching sensations, tiredness, scabbing around a tick bite or shortness of breath were considered positive for the purposes of analysis. Respondents reporting a generalised rash (that is, a skin reaction not in immediate proximity to the tick bite) in the

123 absence of other symptoms were also examined in a separate analysis. Actitivities and incidents occurring the preceeding 2 years were concentrated on as it was thought that the respondants would remember these most accurately.

The presence of a significant association between the variables “age, gender, length of time the respondent had been working on the island, time spent in the bush on the island, use of insect repellent and whether the respondent was frequently bitten by ticks” and the dependent variable “symptoms consistent with a rickettsiosis” was investigated using stepwise regression followed by binary logistic analysis. A significant association between the same variables (and symptoms) and a positive serology result was also investigated using binary and ordinal logistic regression (titres of 1:128 were placed in one group, 1:256 in another and >1:256 into the final group) and the calculation of odds ratios, both unadjusted and adjusted for age and gender. A limitation of this study was that a comparative serosurvey on a control group from the Western Australian mainland was not able to be performed therefore the respondents who were not exposed to the potential risk factors in any category were used as the reference group for the calculation of odds ratios.

6.3 RESULTS

6.3.1 Descriptive epidemiology

Approximately 30% of people on the island at the time of the questionnaire administration session (30/100 at each visit) consented to participate in the study. The study sample consisted of 59 volunteers and this population is described in table 6.1.

The majority of the population was male (93.0%) and over 40 years old (55.9%).

Most of the respondents had been working on the island for over 3 years (40.8%) and most spent time in the bush everyday (45.8%). The majority of respondents recalled

124 finding ticks on themselves only occasionally after spending time in the bush (55.9%) and 22.1% (percentage of respondents reporting symptoms after a tick bite plus percentage of respondents with symptoms but no tick bite history plus percentage of respondents responding positively to both these categories) of the population reported having had symptoms consistent with a rickettsiosis. The titration results from the serology are listed in appendix 8. Forty six percent (27/59) of the population had an antibody titre greater than or equal to 1:128: 21/27 had a titre of 1:128, 5/27 had a titre of 1:256 and 1/27 had a titre >1:256. On analysis, the majority of serological samples demonstrated a higher affinity for rickettsial serotypes in the spotted fever group compared with the typhus group.

125 Table 6.1 Description of the population of Barrow Island workers that responded to the questionnaire.

Variable Categories % Age <28 years old 20.4 28-40 23.7 >40 55.9

Gender Male 93.0 Female 7.0

Time worked on BWI <1year 25.4 1-3 years 25.4 >3 years 40.8 Infrequent visitor 8.5

Time in bush on BWI Everyday 45.8 Often (>3x/week) 23.7 Occasionally 16.9 Hardly ever 13.9

Time in bush on mainland WA More than once per year 47.5 Once per year or less 50.8 1.7 Time in bush (other Aust. States) More than once per year 6.8 Once per year or less 91.5

Time in bush (overseas) More than once per year 11.9 Once per year or less 86.4

Tick bites Most or every time in bush 1.7 Often (25-75%) 13.6 Occasionally 55.9 Never 22.0 Don’t know 6.8

Symptoms (following bite) None or 1 symptom 84.7 Rash (not just round bite) 5.1 2 symptoms from rash, fever/chills and headache 0 Any 2 symptoms from rash, fever, headache, aching, 10.2 tiredness, scabbing, shortness of breath

Symptoms (no bite history) None or 1 symptom 89.8 Rash (not just round bite) 1.7 2 symptoms from rash, fever/chills and headache 0 Any 2 symptoms from rash, fever, headache, aching, 8.5 tiredness, scabbing, shortness of breath Total symptoms None or 1 symptom 77.9 Symptoms after tick bite 11.9 Symptoms with no bite history 6.8 Both 3.4

Medical conditions None 79.6 Minor 8.5 Major (more systemic eg. Heart, asthma, diabetes) 11.9

126

6.3.2 Analysis of questionnaire results

Logistic regression and the calculation of odds ratios demonstrated that there were (i) no significant associations between any of the variables and symptoms, and (ii) no significant associations between any of the variables (including symptoms) and a positive serology result (table 6.2).

127 Table 6.2 Summary of odds ratios (with 95% confidence intervals) for exposure variables versus rickettsial serology Variable Categories Unadjusted odds 95% Adjusted odds 95% CI ratios CI ratios ** Time worked on BWI ≤1 year (baseline) 1.00 1.00

>1 year 1.43 0.48 4.25 1.27 0.41 3.96

Tick repellent used? No 1.00 1.00 Yes 1.11 0.33 3.79 1.06 0.30 3.67

Time in bush on BWI Never/rarely 1.00 1.00 Everyday/often 2.10 0.66 6.67 1.96 0.55 6.94

Time in bush on the mainland (WA) Never/infrequently 1.00 1.00 Often (>once per year) 1.61 0.57 4.51 1.64 0.58 4.66

Time in bush (other Aust. States) Never/infrequently 1.00 1.00 Often (>once per year) 3.29 0.93 11.61 3.09 0.86 11.12

Time in bush (overseas) Never/infrequently 1.00 1.00 Often (>once per year) 0.86 0.17 4.28 0.91 0.18 4.67

Tick bites Never/Infrequently 1.00 1.00 Often 0.87 0.21 3.67 0.77 0.18 3.29

Any history of systemic symptoms following No 1.00 1.00 bite Yes 0.52 0.09 3.11 0.44 0.07 2.66

Any history of unexplained systemic No 1.00 1.00 symptoms; no bite history Yes 0.27 0.03 2.57 0.24 0.03 2.34

Total bites Low 1.00 1.00 (years on island x intensity of tick bites) 128 High 0.91 0.32 2.65 0.68 0.22 2.11 6.4 DISCUSSION

6.4.1 Review of major findings

A particularly noteworthy finding of this study is the high percentage of the population with evidence of antibodies to rickettsiae. International serosurveys of the general population have demonstrated a prevalence rate of 5% or less, therefore the prevalence rate detected in this study (46%) suggests that there is at least one species of infectious Rickettsia on the island. From a global perspective, a prevalence rate this high is unusual even in Rickettsia -endemic areas, and similar rates have only been reported in regions associated with R. africae transmission in Africa (Parola, 1999).

Thus, the findings in the study population could be the result of a very high and continuous rate of infection, perhaps unsurprising given that most of the study population spend a high proportion of their work day in scrub, surrounded by an abundant, tick-carrying native animal population. The workers would, therefore, be constantly exposed to ticks. The most anthropophilic of the tick species on the island, the ornate kangaroo tick, Amblyomma triguttatum , has been shown to have a high rate of Rickettsia spp. infection (chapter 3). It is also possible that the antibodies resulting from infection persist for a long period of time.

The regression and odds ratio results indicate that there is no increased risk of symptoms or seropositivity associated with a particular age group or sex, and that seropositivity was not associated with symptoms consistent with a rickettsiosis

[OR=0.52, 95% CI=0.09, 3.11 (with a tick bite history) and OR=0.27, 95% CI=0.03,

2.57 (without a tick bite history)] or a history of tick bite (OR=0.87, 95% CI=0.21,

3.67) or with related variables that would increase the respondents exposure to ticks such as the length of time worked on the island (OR=1.43, 95% CI=0.48, 4.25) and

129 the amount of time spent in the bush (OR=2.10, 95% CI=0.66, 6.67). The use of insect repellent did not appear to have a protective effect (OR=1.11, 95% CI=0.33,

3.79).

These results suggest that the people working on Barrow Island are being exposed to a rickettsia (or multiple rickettsiae) that produces an asymptomatic infection. Similar results were obtained in a prospective study of soldiers who had been bitten by

Amblyomma triguttatum ticks while in the southwest of WA, an area also identified as harbouring R. gravesii . Although no specific pathogen was being studied, no symptomatic evidence to incriminate the ticks as vectors of disease was observed

(Pearce and Grove, 1987).

It is unlikely that the lack of a significant association between a history of tick bite and an antibody titre is a true result, as SFG rickettsiae are transmitted, in the vast majority of instances, via a tick bite. Although it was not overtly reflected in this study, informal discussions with people working on the island, and personal experience (!), indicated that a large population of ticks that avidly bite people exists on the island, therefore it is more likely that the bites have occurred and that they have gone unnoticed. Other studies surveying people who were most likely exposed to a known tick-transmitted SFG Rickettsia spp. have also failed to demonstrate a significant association between antibody titre and recollection of tick bites (Hilton et al, 1999).

The lack of any association between age or gender and antibody titre (although there was only a very small population of females in this study) is in agreement with the

130 results of previous studies on other Rickettsia spp .. Where people from a particular demographic were infected more often, it was generally conjectured that this related to participation in activities that exposed them more frequently to vectors of rickettsiae rather than to any factor inherent in a particular age group or sex

(Aharonowitz et al ., 1999; Anton et al., 2003).

6.4.2. Limitations of the study

This study was designed to gain the most informative and robust results within the limitations imposed by the respondents’ employer. Barrow Island is remote and flights for non-essential personal are only sporadically available, employees also work

12 hour shifts so they are not always available to attend sessions such as the ones performed for this study. Results from this retrospective, questionnaire-based study cannot be considered conclusive however, given that it was based on a relatively small sample size (emphasized by the wide confidence intervals in the odds ratio calculations) and was reliant on the accuracy of people’s recollections. It is also possible that the clinical signs nominated as being consistent with a rickettsiosis did not include symptoms that may have been relevant, because such infections can manifest as a diverse range of conditions (Helmick et al ., 1984; Fournier et al ., 2004;

Oteo et al ., 2004; Fournier et al., 2005). There was also potential in the extremely hot, humid environment where the study was based, for relevant symptoms to be dismissed as heat stress, sunburn, sweat rash or the result of hard manual labour. The frequency of tick bites could also have been under-represented as they are often painless and due to the very small, immature stages may have therefore gone unnoticed (Gross et al ., 1983; Helmick et al ., 1984)

131 It is possible that there was a degree of selection bias; people who were more concerned about tick bites (and therefore more sensitive to bites, relevant symptoms etc.) may have been more likely to respond. It was not possible to determine whether selection bias would have influenced the results.

Factors inherent in the serology results used in this study also prohibit the drawing of definitive conclusions. Specificity was maximised through the use of a high cut-off titre (so that antibodies cross reacting with non-rickettsia-specific antigens were eliminated from consideration) (Walker and Dumler, 1995) and by the demonstration of a higher affinity for antigens from the spotted fever group members tested than for antigens from the typhus group members. However, results were not specific for members within the spotted fever biogroup. It was not possible to determine, therefore, whether the antibody titre was due to R. gravesii or to another species encountered on or off the island. In addition, the use of a high cut-off titre increases specificity to the detriment of sensitivity. Although this was considered a worthwhile sacrifice in this preliminary study to minimise false positives, the low sensitivity may have resulted in the prevalence of infection being under-represented.

If R. gravesii is responsible for the antibodies detected in this study it is possible that it produces an asymptomatic infection. However, it is not possible to completely dismiss the pathogenic potential of R. gravesii based on these results because of the limitations of the study described above and the fact that asymptomatic rickettsial infections have been recorded in other serological studies, sometimes in people thought to be exposed to rickettsiae of known pathogenic potential such as R. rickettsii , R. conorii and R. africae (Gross et al ., 1983; Mansueto et al ., 1984; Liu et

132 al ., 1995) . For example , in one serological and questionnaire-based study in a R. rickettsii endemic area, 4% of the subjects were seropositive to SFG rickettsia but none recalled having had relevant symptoms (Hilton et al. , 1999). Additionally, serosurveys performed in areas where R. africae is endemic have demonstrated that despite a prevalence of up to 100%, reports of symptomatic infection in indigenous people are extremely rare (Jensenius et al ., 2003; Kelly, 2006). Explanations offered for such asymptomatic infections in populations thought to be exposed to rickettsiae which are in fact pathogenic include the development of immunity through exposure at a young age, variation in strain virulence, and infection with an antigenically similar, but non-pathogenic, organism (Marx et al ., 1982; Mansueto et al ., 1984;

Sanchez et al., 1992; Sarov et al ., 1992; Hilton et al ., 1999; Jensenius et al ., 2003;

Kelly, 2006). Nevertheless, in the present study, workers on Barrow Island would not have been exposed to infection at a young age (unless they were infected with R. gravesii on the mainland), and therefore it is unlikely that they would have developed immunity to explain the lack of symptoms recorded. It is possible however that variation in strain virulence does occur in R. gravesii , or that some of the antibody titres were in response to infection with another, non-pathogenic rickettsiae.

6.4.3 Public health implications of the study

This work has identified that a large proportion of the workforce on Barrow Island are being exposed to a member of the spotted fever group of Rickettsia . Although the findings of this study also suggest that this rickettsia produces an asymptomatic infection, increasing awareness of the risks amongst workers and detailing measures to limit exposure are warranted until further work is performed on the Rickettsia spp. to confirm its non-pathogenic nature.

133

A significant association between exposure to ticks and an antibody titre was not shown, however, as all SFG rickettsiae are transmitted by ticks it is likely that they are also involved in transmission of the recently identified rickettsia on Barrow

Island, R. gravesii . Tick exposure can be minimised by identifying high risk areas (eg. shady, moist environments frequented by animals) and either minimising time spent in these areas or increasing vigilance when at risk. People entering high risk areas could treat clothing with repellents, wear long trousers tucked into their shoes and apply an effective repellent to exposed skin to minimise risk of tick attachment.

Clothing should be checked regularly while in tick infested areas, this is facilitated by wearing light coloured clothes.

If a tick does attach, removing it promptly is critical, as the risk of transmission of a rickettsia or other pathogen by a tick increases with the duration of attachment and generally requires 24-48 hours (Gammons and Salam, 2002). Ticks are removed most effectively using blunt, rounded forceps to grasp the mouthparts of the tick as close to the skin as possible. Other methods of removing ticks, such as using fingers, or irritants to kill the ticks in situ , should be avoided because they may increase the risk of regurgitation by the tick and consequently, the transmission of infectious agents.

6.4.4 Recommendations for further research

This preliminary study has produced some interesting results however they cannot be considered conclusive due to the reasons discussed above. Information obtained from this study should be used in the future to design more focused and specific projects.

Further blood sampling and serology from these individuals and a Western Australian

134 based control group should be performed to determine whether the infection is current or historical, and, together with a prospective study of new workers on the island, will produce more robust results.

135 CHAPTER 7.

GENERAL DISCUSSION

7.1 INTRODUCTION

Spotted fever group rickettsiae are becoming increasingly recognised as emerging infectious diseases of significance (Parola et al ., 2005). As rickettsiology is a developing discipline, new species of rickettsiae are constantly being detected and the knowledge of established species is often revised. This study attempted to address some of the shortfalls in our knowledge of rickettsial presence in Western Australia, concentrating particularly on the role of native and feral animals as reservoir hosts.

7.2 FINDINGS OF THE STUDY IN RELATION TO AIMS

The overall objective of this study was to make a preliminary assessment of rickettsial presence in Western Australia. More specifically the aims were:

Aim 1. To gain a better understanding of whether species of Rickettsia are present in

WA, with the main emphasis on the tick-borne spotted fever group.

This aim was met as two novel SFG rickettsiae [designated Rickettsia gravesii sp. nov . (Chapter 4) and Candidatus “Rickettsia antechini” (Chapter 5)] and one novel

Bartonella species [ Bartonella sp. strain Mu1 (Chapter 5)] were identified in the study. Rickettsia gravesii sp. nov . is most closely related to the Rickettsia massiliae subgroup of the SFG rickettsiae, demonstrating sequence similarities of 99.7%,

98.4%, 95.8% and 97.4% for 16S rRNA, gltA , ompA and ompB gene fragments respectively. Candidatus “Rickettsia antechini” also demonstrated a close relationship to the R. massiliae subgroup (99.4%, 94.8% and 97.1% sequence similarity based on its gltA, ompA and ompB genes respectively). The two novel Western Australian

136 species demonstrated 98.4%, 96.3% and 96.7% sequence similarity to each other based on gltA, ompA and ompB genes respectively. The third novel species identified,

Bartonella sp. strain Mu1, demonstrated greatest gltA gene sequence similarity to

Bartonella strain 40 at 86.1%. The close relationship between the two species of rickettsiae detected in this study could indicate that they both evolved from a recent common ancestor. The lack of a close genetic relationship with R. australis and R. honei (the two documented pathogenic SFG species in Australia) however, suggests that the species of rickettsiae in WA have evolved independently of those present in the eastern states in recent times and probably had different routes of introduction.

This is not a surprising finding as there is increasing evidence that Rickettsia spp. are capable of wide dissemination, illustrated by the presence of R. honei in Australia,

Thailand and the USA (Graves and Stenos, 2003). No previously characterised species of rickettsiae were detected in the study.

Aim 2. To identify geographical areas, vectors and vertebrate hosts of interest in

local rickettsial life cycles, in particular, the role of native and feral animals

in rickettsial ecology.

The tick species identified as potential vectors of R. gravesii sp. nov. by PCR are

Amblyomma triguttatum, Amblyomma limbatum, Haemaphysalis ratti, Amblyomma albolimbatum and Ixodes spp.. Of these, Amblyomma triguttatum is most likely to have the greatest zoonotic significance as it has a relatively high prevalence of infection, a wide host range and it feeds on humans readily (Chapter 3). Ixodes antechini has been identified as a potential vector of Candidatus “R. antechini”

(Chapter 3), however, I. antechini has not been found to infest humans to date and therefore the zoonotic significance of this Rickettsia spp. is unknown.

137

Rickettsia gravesii sp. nov. was detected in several, widely separated areas in the state while Candidatus “R. antechini” and Bartonella sp. strain Mu1 were only detected in

Dwellingup in the State’s southwest (Chapter 3 and 5). Potential vertebrate hosts of R. gravesii sp. nov. identified in this study (i.e. found in ectoparasites collected from the animals but not from the animals themselves) are golden bandicoots, common brushtail possums, burrowing bettongs, western grey kangaroos, feral pigs and bobtail lizards (Chapter 3). The yellow-footed antechinus was identified as a potential vertebrate host of Candidatus “R. antechini” and Bartonella sp. strain Mu1 (Chapter 3 and 5).

Early Australian rickettsiologists undertook preliminary investigations into the role of native animals, particularly bandicoots, in the life cycle of rickettsiae, however little work has been performed in the area recently (Campbell and Domrow, 1974; Huang et al ., 1990). The potential role of feral pigs in rickettsial life cycles has been identified in north-eastern Spain, where Rickettsia slovaca was identified in

Dermacentor marginatus ticks removed from wild boars killed by hunters (Ortuno et al ., 2006). Strategies are already in place to control the feral pig population in the south west of WA due to their destructive nature, nevertheless the potential role of the population as a reservoir of R. gravesii is a finding that could contribute to the argument for control of the feral pig population.

Although an Australian Rickettsia spp., R. honei, has been shown to be transmitted by a reptile-associated tick, it has not yet been determined whether reptiles can function as a vertebrate host of the organisms as their physiology is vastly different from that

138 of mammals (Stenos et al. , 2003). Therefore, the potential role of reptiles and the reptile-associated ticks Amblyomma limbatum and Amblyomma albolimbatum in the life cycle of R. gravesii is unclear and should be investigated further.

Aim 3. To perform a preliminary investigation into the pathogenic potential of any

novel rickettsia species identified during this study.

The serosurvey performed on Barrow Island workers suggests that there is at least one species of rickettsia infective for people on the island, an area which has been shown to be endemic for R. gravesii sp. nov. Although these results, together with the results from the questionnaire, imply that infection is not significantly associated with symptoms, asymptomatic rickettsial infections have been recorded in serological studies performed in other parts of the world, sometimes in people thought to be exposed to rickettsiae of known pathogenic potential such as R. rickettsii , R. conorii and R. africae (Gross et al. , 1983; Mansueto et al ., 1984; Liu et al ., 1995). Therefore, further investigation into the pathogenic potential of R. gravesii should be performed before it is definitively established as a non-pathogenic species, particularly as it is highly prevalent in Amblyomma triguttatum, a tick species that feeds readily on humans.

Aim 4. To contribute to the body of scientific knowledge on rickettsial ecology

worldwide, in order to aid in developing a better understanding about the

genus Rickettsia .

The sequences for the novel Rickettsia and Bartonella species identified in this study have been submitted to GenBank ( www.ncbi.nlm.nih.gov/Genbank/index.html )

(appendix 5), adding to the rapidly growing global literature on the genera (Chapters 4

139 and 5). Cultures of R. gravesii sp. nov . have been submitted to The Australian

Rickettsial Reference Laboratory in Geelong, and are in the process of being submitted to the Collection de souches de L’Unité des Rickettsies (CSUR), World

Health Organization Collaborative Center for Rickettsioses, Borrelioses and Tick- borne Infections, Marseilles, France. This information will be used by rickettsial researchers throughout the world to compare rickettsial species and to determine lineages, routes of transmission and many other factors of the epidemiology and ecology of these fascinating organisms.

This study has provided important information on the potential routes for dissemination of rickettsiae, both within Australia and in a global context. Results from this study have identified ectoparasite vectors and potential vertebrate hosts of novel rickettsiae, many of which are unique to WA and have not been investigated thus far. The demonstration of a high prevalence of rickettsial infection in a tick population on Barrow Island, an ecosystem where no new animals or ectoparasites have been introduced for thousands of years, is a fascinating finding. Results from this study have highlighted the high level of exposure to rickettsiae that has occurred in people working in this environment, with the potential for the SFG rickettsia to be an emerging infectious disease resulting in the incursion of humans into a new niche.

Aim 5. To contribute, through opportunistic sampling, to the current knowledge of

tick presence in Western Australia, including the species of tick present, their

distribution and vertebrate host preference.

Most of the findings from this section of the study (Chapter 2) confirmed those of previous authors (Roberts 1970; Halliday, 2001). A novel finding however was the

140 detection of Ixodes antechini in WA. This tick species may have always been present here and was just not detected in Roberts’ survey, however it is possible that it may have been more recently introduced through the migration of its’ previously identified vertebrate hosts the fat-tailed dunnart and the western quoll. In the past, migration of tick species across the breadth of the continent would have taken a significant amount of time. Now however, captive breeding and release programs and relocation of native species (particularly those that are endangered) are common practices and the potential of such strategies to also disseminate ectoparasites and any infectious agent they may harbour should be taken into consideration.

Another unusual finding of this study was the collection of the reptile-associated tick

Amblyomma limbatum from mammals (golden bandicoots and common brushtail possums) (Roberts, 1970). As Rickettsia gravesii sp. nov. was also detected in

Amblyomma limbatum ticks in this study (Chapter 3), it is possible that this rickettsia is not dependent solely on a mammalian host population for its maintenance and propagation.

7.3 FUTURE DIRECTIONS

While significant findings have been made in this study, it is highly likely that this work represents the “tip of the iceberg” for rickettsiology in Western Australia. The results have raised many important questions, including the zoonotic nature of rickettsiae in WA, and there are many geographical areas, vectors and vertebrate hosts that are yet to be explored.

141 Future work in this area should involve further attempts to culture Candidatus

“Rickettsia antechini” to determine whether culturing of this organism is possible, or whether it is possibly a non-pathogenic endosymbiont similar to R. peacockii, which has been unamenable to culture thus far . Mouse serotyping and phenotypic characterization of both novel strains to support the proposal of R. gravesii sp. nov . as a new rickettsial species will also be part of future work on Western Australian rickettsiae as these studies were not able to be performed during the current study.

Phenotypic properties, including cell culture growth characteristics (temperature, growth media and cell line preference) and patterns of antibiotic resistance should also be investigated. For example, other members of the R. massiliae subgroup have been found to be the only species belonging to a subgroup of the SFG to be rifampin- resistant (Rolain et al ., 1998). It would be interesting to determine whether the novel strains, which appear to be part of the R. massilae subgroup, also demonstrate this phenotypic property and have the associated genotypic characteristics.

Further investigation into the pathogenic potential of the novel strains is also warranted, as although preliminary results suggest that R. gravesii sp. nov. results in asymptomatic infections, this is by no means conclusive. Several cases of SFG rickettsial infection have been diagnosed serologically in the outer suburbs of Perth in recent years (J. Stenos, pers. comm. 2005; J. Dyer, pers. comm. 2006), therefore R. gravesii (which has been detected in this area) or another, as yet unidentified, rickettsia is capable of producing a symptomatic infection. Future work will hopefully involve ongoing serological testing of the Barrow Island individuals already tested and also people who are about to commence work on the island for the first time, in order to identify people with a rising titre and therefore an active infection. These

142 people can then be questioned more closely about the development of symptoms.

Serosurveys in other areas of the State should also be performed in the future, adding to the knowledge gained in this study. Attempts to isolate and sequence rickettsiae from human blood samples and biopsies from humans diagnosed with spotted fevers in the State should also continue so that the identity of any infecting rickettsia can be determined. Finally, education of the medical fraternity in WA on the potential role of rickettsiae in fevers of unknown origin could also play an important part in investigating their role in the State.

Determining the role of the local tick species and vertebrate hosts in the rickettsial life cycles would also provide important information that could be used to better predict distribution of the novel strains and the risk factors for infection, as shown in previous studies (Rehacek and Tarasevich, 1991; Mills and Childs, 1998; Kitron, 2000;

Randolph, 2000; 2004). For example, the role of the tick species as biological vectors could be examined though the use of transmission studies or electron microscopy of different anatomical structures. Such studies could differentiate true vectors from tick species that have just ingested a blood meal containing the Rickettsia spp. Electron microscopy of the vectors could also provide further supporting information on the potential pathogenicity of the rickettsiae. Salivary gland infection would suggest that the Rickettsia spp. can be transmitted to people during tick feeding, while detecting the Rickettsia spp. only in the ticks’ ovaries is a finding that suggests it is more likely to be a non-pathogenic endosymbiont (Raoult and Roux, 1997). Transmission studies on native animals to determine which species are susceptible to infection, are able to support effective proliferation of the rickettsiae, and are infective to vectors for significant periods of time are not practical, however a serological test has been

143 developed to detect Rickettsia australis antibodies in native fauna and development of a similar test for the novel strains may be possible (Huang et al ., 1990). Continued screening of ticks collected from animals in the State will continue to provide information on the distribution and prevalence of the novel strains and may also detect the presence of other novel species.

In conclusion, this “pioneer” study in Western Australia has just scraped the surface of this fascinating genus and the results suggest that there is tremendous scope for further exciting and enlightening work to be performed in the area of rickettsiology in this geographically isolated region in the future.

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189 APPENDICES

APPENDIX 1. ABREVIATIONS

AF (Astrakhan fever) A.j. ( Acanthopsylla jordani ) ARRL (Australian Rickettsial Reference Laboratory) BLAST (Basic local alignment search tool) Bp (base pairs) BWI (Barrow Island) °C (degrees Celsius) CALM (Department of Conservation and Land Management) CF (complement fixation) CI (confidence interval) CSD (cat scratch disease) DNA (Deoxyribonucleic acid) DEBONEL (“ Dermacentor -borne necrosis-erythema-lymphadenopathy”) Dntp (Deoxyribonucleotide triphosphate) ELISA (enzyme linked immunosorbent assay) FISF (Flinders Island spotted fever) FITC (fluorescein isothiocyanate) gltA (citrate synthase gene) I.a. ( Ixodes antechini ) IF (indirect immunofluorescence test) IFA (immunofluorescence assay) IgA (immunoglobulin A) IgG (immunoglobulin G) IgM (immunoglobulin M) ISF (Israeli spotted fever) JSF (Japanese spotted fever) KDa (kilodalton) ME (minimum evolution analyses) Min (minutes) MM (millimoles) MP (maximum parsimony analyses) MSF (Mediterranean spotted fever) NJ (neighbour-joining) OmpA (outer membrane protein gene A) OmpB (outer membrane protein gene B) OR (odds ratio) PBS (phosphate-buffered saline) PCR (polymerase chain reaction) PCR/RFLP (polymerase chain reaction/ restriction fragment length polymorphism ) QTT (Queensland tick typhus) RMSF (Rocky Mountain spotted fever) Rpm (revolutions per minute) rRNA (ribosomal ribonucleic acid) S (seconds) SDS-PAGE (sodium dodecyl sulphate-polyacrylamide gel electrophoresis)

190 SFG (spotted fever group) TAE (tris-acetate-EDTA) TG (typhus group) TIBOLA (“tick-borne lymphadenopathy”) WA (Western Australia) WHO (World Health Organisation)

191

APPENDIX 2. SAMPLE COLLECTION INSTRUCTIONS

PROTOCOL FOR ECTOPARASITE COLLECTION

Background:

The broad aim of our project is to investigate the epidemiology of rickettsial diseases in Western Australia. Rickettsiae are responsible for several human diseases known to be present in this state which include Q fever, murine and scrub typhus, spotted fevers and cat scratch disease. Ticks, fleas, lice and trombiculid mite larvae are known vectors of these diseases but the ability of other ectoparasites to act as vectors has not been ruled out.

We aim to achieve a better understanding of these diseases and possibly identify new ones by investigating the distribution and prevalence of infection in Western Australian ectoparasites and native and domestic animals. The vertebrate host range is also thought to be wide for many of these diseases so we will be looking at ectoparasites from any mammals/marsupials (although reptile and bird ectoparasites are worth collecting if you see them).

The first step of our project will be to collect ectoparasites, which are an essential part of the rickettsial life cycle, from different regions of the state. These will be examined using a sensitive molecular diagnostic test (PCR) to detect the presence of infection. While we would like to concentrate on collection of ticks and fleas, we would be interested in any ectoparasites that were found as unknown ecological cycles may exist in WA.

Suggested protocol :

1. Collect any ectoparasites with flea comb or tweezers. Hair with lice eggs on it can be cut. Ticks can be removed by grasping the mouthparts against the skin with tweezers and pulling firmly but slowly backwards until the tick is eased out of the skin. Trombiculid mite larvae are very small, red-orange and are often found in animals’ ears.

The vials contain 70% ethanol as a preservative and can probably fit 2-3 ectoparasites from the same animal. Hair samples and fleas can be put in the dry containers if more convenient.

2. Record vial number, animal species and where the animal comes from on the data sheet. If possible, information such as approximate number of ectoparasites on the animal (+, ++, +++) and condition of the animal (poor, average, good) would be helpful.

192

APPENDIX 3. COMMON AND SCIENTIFIC NAMES

Black-footed rock wallaby ( Petrogale lateralis )

Bobtail lizard (T rachysaurus rugosus )

Burrowing bettong ( Bettongia lesuer )

Common brushtailed possum ( Trichosurus vulpecular )

Dunnart ( Smithopsis spp.)

Eastern quoll ( Dasyurus viverrinus)

Euro ( Macropus robustus )

Fairy prion ( Pachyptila turtur )

Fat-tailed dunnart ( Sminthopsis crassicaudata )

Feral Pig (Sus scrofa )

Golden bandicoot ( Isoodon auratus )

Kimberley rock rat ( Zyzomys woodwardi )

Olive python ( Liasis olivaceous )

Southern brown bandicoot ( Isoodon obesulus )

Spectacled hare wallaby ( Lagorchestes conspicillatus )

Western-barred bandicoot ( Perameles bougainville )

Western brush wallaby (Macropus irma)

Western grey kangaroo (Macropus fuliginosus )

Western quoll (D asyurus geoffroyi)

Yellow-footed antechinus ( Antechinus flavipes )

193 APPENDIX 4. GENBANK ACCESSION NUMBERS

Table A.1 Rickettsial genbank accession numbers NA – not available

Species 16S rRNA GltA ompA ompB sca4

R. prowazekii M21789 M17149 NA AF200340 AF200340

R. .typhi L36221 U59714 NA L04661 L04661

R. bellii L36103 U59716 NA NA NA

R. helvetica L36212 U59723 NA AF123725 AF163009

R. rickettsii L36217 U59729 U43804 X16353 AF163000

R. conorii AF541999 U59730 U43806 AF163008 AF163008

R. africae L36098 U59733 U43790 AF123706 AF151724

R. sibirica L36218 U59734 U43807 Af123722 AF155057

R. honei L36220 U59726 U43809 AF123724 AF163004

R. slovaca L36224 U59725 U43808 AF123723 AF155054

R. parkeri L36673 U59732 U43802 AF123717 AF155059

R. japonica L36213 U59724 U43795 AF123713 AF155055

R. akari L36099 U59717 NA AF123707 AF213016

R. australis L36101 U59718 AF149108 AF123709 AF187982

R. felis L28944 AF210692 AF210694 AF210695 AF196973

R. massiliae L36214 U59719 U43799 AF123714 AF163003

R. montanensis L36215 U74756 U43801 AF123716 AF163002

R. rhipicephali L36216 U59721 U43803 AF123719 AF155053

R. aeschlimannii U74757 U59722 U43800 AF123705 AF163006

R. heilongjiangii AF178037 AF178034 AF179362 AY260451 AY331396

194 Table A.2 Bartonella spp. Genbank Accession Numbers.

Species gltA

Bartonella bacilliformis Z70021

B. doshiae Z70017

B. elizabethae Z70009

B. grahamii Z70016

B. quintana Z70014

B. taylorii Z70013

B. vinsonii Z70015

Bartonella sp. Strain C1-phy Z70022

Bartonella sp. Strain C4-phy Z70019

Bartonella sp. Strain C5-rat Z70018

Bartonella sp. Strain C7-rat Z70020

Bartonella sp. Strain R-phy 1 Z70010

Bartonella sp. Strain R-phy2 Z70011

Bartonella sp. Strain N40 Z70012

B.clarridgeiae U84386

B.koehlerae AF176091

B.peromysci AY805110

B. tribocorum AJ005494

195 APPENDIX 5. GENBANK ACCESSION NUMBERS AND SEQUENCES OF THE NOVEL STRAINS

Rickettsia gravesii sp. nov. 16S rRNA gene - accession number: DQ269434 gttagtggca gacgggtgag taacacgtgg gaatctgtcc atcagtacgg aataactttt agaaataaaa gctaataccg tatattctct acggaggaaa gatttatcgc tgatgggtga gcccgcgtca gattaggtag ttggtgaggt aatggctcac caagccgacg atctgtagct ggtctgagag gatgatcagc cacactggga ctgagacacg gcccagactc ctacgggagg cagcagtggg gaatattgga caatgggcga aagcctgatc cagcaatacc gagtgagtga tgaaggcctt agggttgtaa agctctttta gcaaggaaga taatgacgtt acttgcagaa aaagccccgg ctaactccgt gccagcagcc gcggtaagac ggagggggct agcgttgttc ggaattactg ggcgtaaaga gtgcgtaggc ggtttagtaa gttggaagtg aaagcccggg gcttaacctc ggaattgctt tcaaaactac taatctagag tgtagtaggg gatgatggaa ttcctagtgt agaggtgaaa ttcttagata ttaggaggaa caccggtggc gaaggcggtc atctgggcta caactgacgc tgatgcacga aagcgtgggg agcaaacagg attagatacc ctggtagtcc acgccgtaaa cgatgagtgc tagatatcgg aagattctct ttcggtttcg cagctaacgc attaagcact ccgcctgggg agtacggtcg caagattaaa actcaaagga attgacgggg gctcgcacaa gcggtggagc atgcggttta attcgatgtt acgcgaaaaa ccttaccaac ccttgacatg gtggtcgcgg atcgcagaga tgcttttctt cagctcggct ggaccacaca caggtgttgc atggctgtcg tcagctcgtg tcgtgagatg ttgggttaag tcccgcaacg agcgcaaccc tcattcttat ttgccagcgg gtaatgccgg gaactataag aaaactgccg gtgataagcc ggaggaaggt ggggacgacg tcaagtcatc atggccctta cgggttgggc tacacgcgtg ctacaatggt gtttacagag ggaagcaaga cggcgacgtg gagcaaatcc ctaaaagaca tctcagttcg gattgttctc tgcaactcga gagcatgaag ttggaatcgc tagtaatcgc ggatcagcat gccgcggtga atacgttctc gggccttgta cacactgccc gtcacgccat gggagttggt ttt

Rickettsia gravesii sp. nov. gltA gene - accession number: DQ269435 gcaagtatcg gtgaggatgt aatcgatata agtagggtat ctgcggaagc cgattgcttt acttccgacc cgggttttat gtctcctgct tcttgtcagt ctactatcac ctatatagac ggtgataaag gaaacttgcg gcatcgagga tatgatatta aagacttagc tgagaaatgt gattttttag aagtagctta tttactgatt tatggggaac taccaagtgg cgagcagtat aataatttca ctaaacaggt tgctcatcat tctttagtga atgaaagatt acactattta tttcaaacct tttgtagctc ttctcatcct gtggctatta tgcttgcggc tgtcggttct ctttcggcat tttatcctga tttattgaat tttaaggaag cagattacga acttaccgct attagaatga ttgctaagat acctaccatc gccgcaatgt cttataaata ttctatagga caaccgttta tttatcctga taattcgtta gattttaccg aaaattttct gcatatgatg tttgcaacgc cttgtacgaa atataaagta aatccaataa taaaaaatgc tcttaataag atatttatcc tacatgccga tcatgagcag aatgcttcta cttcaacagt ccgaattgct ggttcatccg gagctaaccc ttttgcttgt attagcacgg gtattgcctc actttggggg cctgctcacg gcggggctaa tgaagcggta ataaatatgc ttaaaaaaat tggtagttct gagtatattc ctaaatacat agctaaagct aaggataaaa atgatccatt taggttaatg ggttttggtc atcgtgtata taaaaactat gacccgcgtg ccgcagtact taaaaaaacg tgcaaagaag tattaaagga actcgggcag ctagacaaca atccgctctt acaaatagca atagaacttg aagctatcgc tcttaaagat gaatatttta ttgagagaaa attatatcca aatgttgatt tttattcggg tattatctat aaagctatgg gtataccgtc gcaaatgttc

196 actgtacttt ttgcaata

Rickettsia gravesii sp. nov. ompA gene - accession number: DQ269437 tcttaaagcc gctttattca ccacctcaac cgcagcgata gtgctgagta gtagcggggc actcggtgtt gctgcaggtg ttatttctac taataatgca gcatttagtg atcttgctgt tgccaataat tggaatgaga taacggctga aggggtagct aatggtgctc ctgctgacgg tcctcaagac aattgggcat ttacttacgg tggtgatcat actatcactg cagataaagt cggtcgtatt attacggcta taaatgttgc gggtactact cccgtaggtc taaatattac tcaaaatacc gtcgttggtt cgattatgac gggaggtaac ttgttgcctg ttactattac tgccggcaaa agcttaactt taaatggtac taatgctgtt gctgcaaatc atggttttga tgctcctgcc gataattata caggtttagg aaatataact ttagggggag tgaatgctgc actaattata caatctgcaa ccccggcaaa gataacactt gcaggaaata tatatggag

Rickettsia gravesii sp. nov. ompB gene - accession number: DQ269438 ccatagtagc cagttttgca ggttcagcta tgggtgctgc tatacagcag aatagaacaa caaacggagt tgctacaact gttgatggtg cgggatttga ccaaactgcc gccgttcctc ctgtaaatgt tgcggttgct ctaaatgcag ttattactgc taatgctaat aatggtatta atttaaatac tccagccggt agttttaacg gtttgttttt agatactgca aacaatttag cagtgacagt gagtgcagat actaccttag ggttcatcac taatgttgct aataaaggta acttctttga ccttacgctt gatgccggta aaactcttac tataacaggt caaggtatta ctaatgcaca agctgctgtt acaaaaaatg ctcaaaatgt tgttgcacaa tttaatggtg gtgctgctat tgccaataat gatcttagcg gtgtaggaag

Rickettsia gravesii sp. nov. sca4 gene - accession number: DQ269438 agcaataagg aatatacaga agaacaagag caaaaagaat ttttatctca aactacaacc ccagaactag aagctgacga tggttttatc gttacttctg catcttctgc tcaatttacc ccttcaatta gtgctttatc ggacaatatc tctcctgaca gtcagatatc agacccaata accaaggctg taagggaaac aattatacaa ccgcaaaaag ataatttaat agaacaaata ttaaaaaacc tggcagtcct tacagaccgt gatttagctg aacaaaaaag aaaagaaata gaagaagaaa aagataaagc attaagtact tttttcggta atccggctaa tagagagttt attgataagg ctttagaaaa tcctgagctt aaaaagaaat tagaatcaat agaaatagcc ggctataaaa atattcataa tacatttagt gccgctagtg ggtaccctgg tggatttaaa ccggtacagt gggaaaatca agtaagtgca agcgatctta gagcaacagt agttaaaaat gatgcaggtg atgaactctg taccttaaat gaagcaactg ttaaaactaa gccttttact ttagctaaac aagacggtac tcaggttcag atcagctcat atagggaaat agattttcct ataaaacttg ataaagccga tggatcaatg catttatcga tggtagcatt aaaagctgat ggcacaaagc cctctaaaga taaagccgta tatttcactg tccactac

Candidatus “Rickettsia antechini” gltA gene - accession number: DQ372954 gcaagtatcg gtgaggatgt aatcgatata agtagggtat ctgcggaagc tgattgcttt acttacgacc cgggttttat gtctactgct tcttgtcagt ctactatcac ctatatagac ggtgataaag gaatcttgcg gcatcgagga tatgatatta aagacttagc tgagaaaagt gattttttag aagtggcata tttactgatt tatggggaac taccaagtgg cgagcagtat aataatttca ctaaacaggt tgctcatcat tcattagtga atgaaagatt acactattta tttcaaacct tttgtagctc ttctcatcct atggctatta tgcttgcggc tgtcggttct ctttcggcat tttatcctga tttattgaat tttaaggaag cagattacga acttaccgct

197 attagaatga ttgctaagat acccaccatc gccgcaatgt cttataaata ttctatagga caaccgttta tttatcctga taattcgtta gattttaccg aaaattttct gcatatgatg tttgcaacgc cttgtacgaa atataaagta aatccaataa taaaaaatgc tcttaataag atatttatcc tacatgccga tcatgagcag aatgcttcta cttcaacagt ccgaattgcc ggctcatccg gagctaaccc ttttgcttgt attagcacgg gtattgcctc actttggggg cctgctcacg gcggggctaa tgaagcggta ataaatatgc ttaaagaaat cggtagttct gagtatattc ctaaatatat agctaaagct aaggataaaa atgatccatt taggttaatg ggttttggtc atcgtgtata taaaaactat gacccgcgtg ccgcagtact taaagaaacg tgcaaagaag tattaaagga actcgggcag ctagacaaca atccgctctt acaaatagca atagaacttg aagctatcgc tcttaaagat gaatatttta ttgagagaaa attatatcca aatgttgatt tttattcggg tattagctat aaagctatgg gtataccgtc gcaa

Candidatus “Rickettsia antechini” ompA gene - accession number: DQ372955 tcttaaagcc gctttattca ctacctcaac cgcagcgata atgctgagta gtagcggggc actcggtgtt gctgcaggtg ttattgctac taataatgca gcatttagtg atcttgctgt tgccaataat tggaatgaga taacggctgg aggggtagct aatggtactc ctgctggcgg tcctcaaaac aatggggcat ttacttacgg tggtgatcat actatcactg cagatgcagt tgatcgtatt attacggcta taaatgttgc gggtactact cccgtaggtc taaatattgc tcaaaatacc gtcgttggtt cgattatgac gggaggtaac ttgttgcctg ttactattac tgccggcaaa agcttaactt taaacggtac taatgctgtt gctgcaaatc atggttttga tgctcctgtc gataattata caggtttagg aaatataact ttagggggag cgaatgctgc actaattata caatctgcaa ccccggcaaa gttaacactt gcaggaaata tagatggag

Candidatus “Rickettsia antechini” ompB gene - accession number: DQ372956 ccatagtagc cagttttgca ggttcagcta tgggtgctgc tatacagcag aatagaacaa caaacggagc tgctacaact gttgatggtg cgggatttga ccaaattgcc gctcctgcaa atgttgcggt tgctctaaat gcagttatta ctgctaatgc taataatggt attaatttaa atactccagc cggtagtttt aacggtttgt ttttagatac tgcaaacaat ttagcagtga cagtgagtgc agatactacc ttagggttca tcactaatgc tgctaataac ggtaactcct ttaaccttac gcttggtgct ggtaaaactc ttactataac aggtcaaggt attactaatg cacaagctgc tgttacacac aatgctaaaa atgttgttgt acaatttaat ggtggtgctg ctattgccaa taatgatctt agcggtgtgg gaac

Bartonella spp. strain Mu1 gltA gene – accession number: DQ372957 gaaaaaattc ctcaatttat tgcacgtgcg aaagataaag acgacccttt ccgccttatg ggttttggcc accggatata taaaaattat gatccgcgcg cgaaaattat gcaaaaaact tgccataaag ttttaaaaga attgaatatt aaagatgatc cactccttga tatcgctata aaacttgaaa atatagctct taatgacgaa tattttgttg aaaagaaact ttatcct

198 APPENDIX 6. MINIMUM EVOLUTION AND MAXIMUM PARSIMONY TREES.

R. parkeri R. sibirica

R. conorii R. africae R. honei

R. slovaca

R. rickettsii R. japonica 93 R. heilongjiangii

R. montanensis R. aeschlimannii R. massiliae

R. rhipicephali Rickettsia sp.BWI-1 R. helvetica

R. felis R. prowazekii 100 R. typhi

R. bellii

R. akari R. australis

0.2%

Figure A4.1 Minimum evolution tree based on 16S rRNA sequences.

199

R. conorii R. sibirica R. africae

R. parkeri

R. honei

R. rickettsii R. slovaca R. japonica R. heilongjiangii 94 R. aeschlimannii

Rickettsia sp.BWI -1

R. massiliae R. rhipicephali R. montanensis

R. helvetica R. felis R. akari

R. prowazekii

99 R. typhi R. bellii R. australis

Figure A4.2 Maximum parsimony tree based on 16S rRNA sequences.

200

R. sibirica R. parkeri

R. africae R. slovaca R. honei

72 R. rickettsii R. conorii R. japonica 99 R. heilongjiangii R. aeschlimannii

76 R. massiliae

100 94 R. rhipicephali

Rickettsia sp. BWI-1 81 R. montanensis 92 R. helvetica

R. felis 98 R. akari 99 R. australis

R. prowazekii

100 R. typhi R. bellii

2%

Figure A4.3. Minimum evolution tree based on gltA sequences.

201

R . rickettsii

R. conorii R. honei R. slovaca

R. africae

R. sibirica R. parkeri

R. japonica

97 R. heilongjiangii Rickettsia sp.BWI11 86 R. aeschlimannii

70 R. massiliae 96 R. rhipicephali R. montanensis 83 R. helvetica R. felis 96 R. akari 99 R. australis

R. prowazekii 100 R. typhi R. bellii

Figure A4.4. Maximum parsimony tree based on gltA sequences.

202

R. africae R. parkeri

R. sibirica

98 R. conorii R. rickettsii

R. slovaca 97 R. honei R. japonica 100 R. heilongjiangii

R. aeschlimannii 84 100 R. massiliae 93 R. rhipicephali

Rickettsia sp. BWI-1 R. montanensis R. australis

2%

Figure A4.5 Minimum evolution tree based on ompA sequences.

203

R. africae

R. sibirica

R. parkeri R. slovaca 97 R. honei

94 R. rickettsii R. conorii R. japonica 99 R. heilongjiangii 77 Rickettsia sp. BWI-1 R. aeschlimannii

99 R. massiliae 78 R. rhipicephali R. montanensis

R. australis

Figure A4.6 Maximum parsimony tree based on ompA sequences.

204

R .africae 76 92 R .parkeri R. sibirica 85 94 R. mongolotimonae 80 99 R. conorii R. slovaca R. rickettsii 71 R. honei

R. japonica 99 100 R. heilongjiangii R. aeschlimannii

Rickettsia sp. BWI-1 R. montanensis R. massiliae

77 R. rhipicephali R. akari 99 R. australis

R. prowazekii

100 R. typhi

5%

Figure A4.7 Minimum evolution tree based on ompB sequences.

205

80 R. africae 83 R .parkeri

92 R. sibirica

R. mongolotimonae 89 R. conorii R. slovaca 95 R. rickettsii R. honei

R. aesc hlimannii R. massiliae

99 R. rhipicephali Rickettsia sp. BWI-1 R. montanensis 100 R. japonica 99 R. heilongjiangii R. akari

96 R. australis

R. prowazekii R. typhi

Figure A4.8. Maximum parsimony tree based on ompB sequences.

206

R. conorii R. parkeri R. sibirica

R. africae R. slovaca R. rickettsii

R. honei

R. japonica 98 R heilongjiangii R. montanensis

100 R. aeschlimannii R. massiliae 91 83 R. rhipicephali

Rickettsia sp. BWI-1 R. felis R. akari 100 100 R. australis

R. prowazekii

100 R. typhi

5%

Figure A4.9. Minimum evolution tree based on sca4 sequences.

207

R. conorii R. parkeri

R. sibirica R. africae 74 R. honei

R. rickettsii

R. slovaca R. japonica 83 R. heilongjiangii

R. montanensis 100 R. aeschlimannii R. massiliae

90 71 R. rhipicephali Rickettsia sp.BWI1

100 R. felis

99 R. akari

100 R. australis R. helvetica

R. prowazekii

R. typhi

Figure A4.10. Maximum parsimony tree based on sca4 sequences.

208

R. sibirica

R. parkeri R. africae R. slovaca

R. honei R. rickettsii R. conorii

R. japonica

99 R. heilongjiangii R. gravesii Rickettsia sp. D-1 100 R. aeschlimannii R. massiliae 83 75 94 R. rhipicephali

R. montanensis 89 R. helvetica R. felis

98 R. akari

99 R. australis R. prowazekii

100 R. typhi

R. bellii

2%

Figure A4.11. Minimum evolution tree based on gltA sequences ( Rickettsia sp. D-1).

209

R. rickettsi i R. conorii

R. honei

R. sibirica R. parkeri R. slovaca

R. africae

R. japonica 98 R. heilongjiangii R. gravesii

Rickettsia sp. D -1 90 R. aeschlimannii 70 R. massiliae 95 R. rhipicephali R. montanensis 84 R. helvetica

R. felis

96 R. akar i 99 R. australis

R. prowazekii

100 R. typhi R. bellii

Figure A4.12. Maximum parsimony tree based on gltA sequences.

210

R. africae

R. parkeri R. sibirica

99 R. conorii

R. rickettsii

94 R. honei R. slovaca 91 R. japonica

100 R. heilongjiangii Rickettsia sp. D-1 R. aeschlimannii

85 R. massiliae 99 94 R. rhipicephali R. gravesii

R. montanensis

R. australis

2%

Figure A4.13. Minimum evolution tree based on ompA sequences.

211

R. africae R. parkeri

R . sibirica

98 R. conorii

R. rickettsii 93 R. honei R. slovaca 90 R. japonica 99 R. heilongjiangii Rickettsia sp. D -1

R. aeschlimannii 85 R. massiliae 99 76 R. rhipicephali R. gravesii

R. montanensis R. australis

Figure A4.14. Maximum parsimony tree based on ompA sequences.

212

77 R. africae 90 R. parkeri R. sibirica 88 94 R. mongolotimonae 82 R. conorii 99 R. slovaca

R. rickettsii

R. honei R. japonica 99 99 R. heilongjiangii

R. aeschlimannii R. gravesii Rickettsia sp. D-1

R. montanensis R. massiliae R. rhipicephali

R. akari

99 R. australis R. prowazekii

100 R. typhi

5%

Figure A4.15. Minimum evolution tree based on ompB sequences.

213

R. africae 80 83 R. parkeri

92 R. sibirica R. mongolotimonae 90 R. conorii

96 R. slovaca

R. rickettsii R. honei R. japonica 99 R. heilongjiangii 99

R. aeschlimannii R. gravesii

100 R. massiliae R. rhipic ephali Rickettsia sp. D -1

R. montanensis

R. akari 97 R. australis

R. prowazekii

R. typhi

Figure A4.16. Maximum parsimony tree based on ompB sequences.

214

100 B. elizabethae 99 Strain C7rat

B. grahamii

StrainR1phy

95 Strain C4phy

Strain R2phy 100 B. taylorii

79 B. vinsonii 100 Strain C1phy

B. bacilliformis B. doshiae Strain40

Bartonella sp .strain Mu1

B. quintana

85 B. henselae

0.02

Figure A.17. Minimum evolution tree based on gltA sequences ( Bartonella sp. strain Mu1).

215

99 B. elizabethae 95 Strain C7rat

B. grahamii

Strain R1phy

Strain C4rphy 79 100 Strain R2phy Strain 40

Bartonella sp.strain Mu1

B. quintana

B. henselae B. bacilliformis

B. doshiae

B. vinsonii 98 Strain C1phy

B. taylorii

Figure A.18. Maximum-Parsimony Tree Based on gltA Sequences ( Bartonella sp. strain Mu1).

216 APPENDIX 7. QUESTIONNAIRE

BARROW ISLAND HEALTH AND SAFETY QUESTIONNAIRE

PATIENT INFORMATION STATEMENT AND CONSENT FORM

Principal Investigators :

A/Prof Stan Fenwick Epidemiology & Public Health School of Veterinary & Biomedical Sciences Murdoch University

Dr John Dyer Infectious Diseases Physician SMAHS Infectious Diseases Service Fremantle Hospital

Dr Angus Cook, Director, Ecology and Heath Research Group, School of Population Health, University of Western Australia

Contact Person :

A/Prof Stan Fenwick Epidemiology & Public Health School of Veterinary & Biomedical Sciences Murdoch University

Ph 08 9360 7418

You are invited to participate in the research study outlined below.

217 1. Background to Research Study

Before you decide whether or not to take part in this research study, it is important that you understand why the study is being done, the procedures and assessments involved, and other important study information. Please read this information carefully and, if you have further questions, please ask the Principal Investigator.

Rickettsias are a type of bacteria that are passed to humans by tick bite, and they may be an important unrecognised cause of infection in Western Australia.

We plan to investigate whether this infection affects people working on Barrow Island, who are frequently exposed to ticks during their work or recreational activities. Any germs found as a result of these studies will be analysed to see if they are like other species known to cause human disease.

2. Purpose of the Study

This study will find out if the bacteria (rickettsias) causing spotted fever are present in Western Australia. We want to discover if individuals engaged in activities in the bush, who have a high risk of tick contact, may have increased rates of infection from ticks.

The ultimate goals of this research are:

• to identify the risk of transmission of this type of infection in Western Australia • to reduce the chance of individuals living, working or recreating in high-risk areas getting infections • to develop health and safety guidelines for individuals at risk of infection

3. The Study Design

An ethics committee has approved this study before any participant is allowed to enrol in the study. We are seeking a total of approximately 100-120 workers on Barrow Island.

This study will involve: (a) a blood test obtained at the beginning of the study period season to see if there has been any previous exposure to infection, and if this relates to contact with ticks; (b) a second blood test 6 months later to see if any new cases of infection have occurred over this period of time.

At the time of collection of the initial blood test, those who participate will be asked to fill out a questionnaire to measure previous level of contact with ticks.

4. Procedures

If you decide to take part after reading this information sheet and discussing the trial with the study investigators, you will be asked to sign a form confirming that you agree to participate.

218 As part of this study you will have blood drawn (approximately 10mL or 1.5 tablespoons) at the time of enrolment, and another 10mL sample will be taken about 6 months later. This is a very important stage as it tells us about whether your body has been contact with these bacteria.

At the time of your first visit you will be asked to fill in a questionnaire asking about your past level of risk for tick exposure. These documents will be collected by the study investigators and analysed to determine correlations between blood test results and intensity of tick exposure.

The information you provide is confidential and will only be used for the purposes of this study.

5. Risks of Study Procedures

Risks of Blood Drawing

As part of this study, you will have your blood drawn (approximately 10mL) at each visit. Blood will be drawn by a health professional with extensive experience in blood taking. This procedure is uncomfortable but rarely results in any significant problems. Side effects that have been noted with drawing blood include feeling light-headed or faint, fainting, formation of a blood clot, bruising and/or infection at the site of the sampling.

6. Benefits of Participating in the Study

We expect that this study, which has never been done before in WA, will benefit people involved in a number of jobs that involve a high level of contact with the natural environment. For the first time, we will understand if there are risks of this important type of infection in WA and will be able to make suggestions for reducing the chance of getting infected.

7. Study expenses

There are no financial costs to you by your involvement.

8. Financial Support for the Study

Financial support to cover the costs of this study is being provided by Chevron and Fremantle Hospital. These funds are placed in a nominated account of the Hospital and the expenditure of the funds is in accordance with the approval given by the Human Research Ethics Committee.

9. Questions about the study

219

The Principal Investigator will do all that is possible to answer your questions and concerns as they arise.

10. Statement of Subject Rights

Your participation in this study is entirely voluntary. This means that if you decide not to take part or to withdraw from the study at any time, you will not be penalised nor lose any benefits to which you are otherwise entitled.

The research team will have explained the details of this trial to you and answered any questions you may have. You should be satisfied with the information you have been given and had adequate time to consider whether you want to participate.

If you decide you would like to take part in this study, you will be asked to sign a consent form. If you do not want to take part, or if you choose to withdraw from the trial at any time, you will continue to receive the best medical care offered by your doctor. You do not have to give a reason if you withdraw.

All information obtained in connection with this study will remain confidential. You will never be identified in any publication or public presentation of data from this study. Only authorised personnel at Fremantle Hospital and the School of Population Health, University of Western Australia will have access to the study documents and results. This information is kept in a locked, password protected facility. By signing this form, you give permission that these authorised persons may have access to study documents if necessary.

Your employer has NO access to this personal information at any time and cannot ever request it.

If new information becomes available during the course of the study that is relevant to your willingness to continue taking part in the study you will be informed promptly.

If you would like more information about the study, do not hesitate to ask Dr Stan Fenwick: Contact No: 08 9360 7418

If you have a problem during the study and would like to talk to an independent person you may contact: Murdoch University Human Ethics Committee Representative Contact No: 08 9360 2393.

You will be given a copy of this form to keep.

220

1.1.1 CONSENT FORM

1. I, ...... of ...... , aged ...... years, agree to participate in the experiment described in the patient information statement set out in the attached form.

2. I acknowledge that I have read the patient information statement, which explains why I have been selected, the aims of the experiment and the nature and the possible risks of the investigation, and the statement has been explained to me to my satisfaction.

3. Before signing this consent form, I have been given the opportunity of asking any questions relating to any possible physical and mental harm I might suffer as a result of my participation and I have received satisfactory answers.

4. I understand that I can withdraw from the experiment at any time without prejudice to my relationship with my doctor.

5. I agree that research data gathered from the results of the study may be published, provided that I cannot be identified.

6. I understand that if I have any questions relating to my participation in this research, I may contact Dr Stan Fenwick at Murdoch University on 08 9360 7418 , who will be happy to answer them.

7. I acknowledge receipt of a copy of this Consent Form and the Subject Information Statement. Complaints may be directed to Chair of Murdoch University Human Research Ethics Committee on 08 9360 2393

______Signature of Participant Please PRINT name Date

______Signature of Investigator(s) Please PRINT name Date

______Signature of Witness Please PRINT name Date

______Nature of Witness

221

1.1.2 CONSENT FORM

1.1.3 REVOCATION OF CONSENT

I hereby wish to WITHDRAW my consent to participate in the research proposal described above and understand that such withdrawal WILL NOT jeopardise any treatment or my relationship with my doctor.

______Signature Date

______Please PRINT Name

______Signature of Investigator(s)

______Please PRINT name Date

______Signature of Witness

______Please PRINT name Date

______Nature of Witness

222

Identification code: |_|_|_|_|_|

BBAARRRROOWW IISSLLAANNDD HHEEAALLTTHH AANNDD SSAAFFEETTYY QQUUEESSTTIIOONNNNAAIIRREE

Thank you for your interest and co-operation in our research. We are a group of doctors and veterinarians from Murdoch University, Fremantle Hospital, and the School of Population Health at the University of Western Australia and are looking at whether contact with ticks affects human health.

We have a particular interest in a range of diseases which ticks can carry and transmit to people, such as Q fever, Flinders Island spotted fever and Queensland tick typhus in other parts of Australia.

It is not known whether any of these problems exist on Barrow Island. The information you give us is vital to understanding whether such diseases affect humans and what activities may affect a person’s chances of catching them. The research you are involved in has never been done before in Western Australia.

223

GENERAL QUESTIONS

1. Date today: / / 2006

2. Gender:  Male  Female

3. Year of birth: |_|_|_|_|

4. What is your current job/role on Barrow Island:

…………………………………..

5. What are the main work tasks you do in this job?

………………………………………………………………………………… ……………

………………..………………………………………………………………… ………….

………………………………………………………………….……………… …………..

6. How long have you worked on Barrow Island?

|_|_| Months |_|_| Years

7. Approximately how many times a year do you go to the island?

|_|_| Times

8. Approximately how long do you spend on the Island each time you work there?

|_|_|_| Days OR |_|_|Weeks OR |_|_| Months

224

RECORD OF PAST AND CURRENT ACTIVITIES ON BARROW ISLAND

9. While you are on Barrow Island, how often do you go into the bush or scrub for work ? (please tick the answer that most applies to you)

 Very frequently (5 or more days a week)  Frequently (1-4 days a week)  Often (1-3 times a month)  Occasionally (Less than once a month)  I spend NO time in the bush for work

10. While you are on Barrow Island, how often do you go into the bush or scrub on Barrow Island for recreation (eg bush walking) ? (please tick the answer that most applies to you)

 Very frequently (5 or more days a week)  Frequently (1-4 days a week)  Often (1-3 times a month)  Occasionally (Less than once a month)  I spend NO time in the bush for recreation

If you answered “ I spend NO time in the bush ” to question 9 AND 10, please go straight to QUESTION 17 . Otherwise, please proceed to QUESTION 11.

11. How long do you spend in the bush on an average day ?

 No time  Less than 1 hour  1 to 3 hours  Greater than 3 hours

12. Is time you spend in the bush on Barrow Island:

 Mainly for work reasons  Mainly recreational reasons  Equally related to work and recreation

13. How often do you wear short-sleeved shirts and/or shorts during recreational time spent in bush or scrub on Barrow Island?

 Every time I go out for recreational reasons  Often (50% or more of the time I go out for recreational reasons)  Sometimes (less than 50% of the time I go out for recreational reasons)  Never

225

14. How often do you see animals (such as marsupials) or evidence that they have been there?

 Every time I go out  if so , What animals? …………………………………..  Often (50% or more of the time I go out)  if so , What animals? …………………………………..  Sometimes (less than 50% of the time I go out )  if so , What animals? …………………………………..  Never

15. How often do you find ticks on yourself after time spent in the bush on Barrow Island?

 Every time I go out  Often (50% or more of the time I go out)  Sometimes (less than 50% of the time I go out)  Never

16. How often do you apply tick repellent before spending time in the bush or scrub?

 Every time I go out  Often (50% or more of the time I go out)  Sometimes (less than 50% of the time I go out )  Never

226 17. In the past 2 years, how often have you been bushwalking, hiking, camping or working in the bush or forest in other parts of Australia or overseas ? This does NOT include your time on Barrow Island.

Bushwalking, hiking, camping or working Bushwalking, hiking, camping or working in the bush or forest in other parts of in the bush or forest overseas (outside Western Australia (except for Barrow Australia) Island) or interstate

(please tick the answer (please tick the answer that most applies to you) that most applies to you)

 Not at all in past 2 years  Not at all in past 2 years

 Less than once a year  Less than once a year  Which  Which state(s)/territories?………… country/countries?…………

 About once a year  About once a year  Which  Which state(s)/territories?………… country/countries?…………

 More than once a year  More than once a year  Which state(s)/territories?…………  Which country/countries?…………

227

RECORD OF TICK BITES

18. IN THE PAST 2 YEARS, how often have you been bitten by ticks? This refers to tick bites that may have occurred while on Barrow Island, or while in some other location in Australia or overseas.

 Most or every time I am in the bush (more than 75% of the time)  Often when I am in the bush (25-75% of the time)  Occasionally when I am in the bush (Less than 25% of the time)  Never (Go to Q19)  Not sure (Go to Q19)

 If the subject answered “ Never I spend no time in the bush part from rogaining ” or “Not sure” to question 18, please go straight to question 19. Otherwise, please proceed.

a. In which months do the bites usually occur? (e.g. Mar-May) …….

b. In what locations have you been training when you were bitten by ticks IN THE PAST 2 YEARS? (TICK ALL THAT APPLY)

 on Barrow lsland  elsewhere in Australia or overseas Location (eg name of area, park or district): ……………. State: ……………. Country: …………….

c. In which part(s) of your body do the bite(s) usually occur? (TICK ALL THAT APPLY)  on chest, back or abdomen (stomach region)  hands, arms or shoulders  feet or legs  on neck, face or scalp  on groin or buttocks

d. Have you experienced any of the following after tick bites IN THE PAST 2 YEARS?

Nature of reaction Please tick any symptoms that you have had Immediate (up to a couple of days after tick bites)

- Local swelling of the skin around the  Most or e very time I get bitten (more than 75% of the time) bite  Often when I get bitten (25-75% of the time)  Occasionally when I get bitten (Less than 25% of the time)  Never  Not sure

- Tenderness of the skin around the bite  Most or e very time I get bitten (more than 75% of the time)  Often when I get bitten (25-75% of the time)  Occasionally when I get bitten (Less than 25% of the time)  Never  Not sure

- Discharge, pus or fluid coming out of  Most or e very time I get bitten (more than 75% of the time) the skin around the bite  Often when I get bitten (25-75% of the time)  Occasionally when I get bitten (Less than 25% of the time)  Never  Not sure

228

- Itchiness around the bite  Most or e very time I get bitten (more than 75% of the time)  Often when I get bitten (25-75% of the time)  Occasionally when I get bitten (Less than 25% of the time)  Never  Not sure

At any time in the 2 weeks after tick bites

- Prolonged headache (> 3 days)*  When did this occur? (e.g. Feb 2005)

……/……….

- Fever or chills*  When did this occur? (e.g. Feb 2005)

……/……….

- Rash* TICK ALL THAT APPLY  around the bite  on chest, back or abdomen (stomach region)  hands, arms or shoulders  feet or legs  on neck, face or scalp  on groin or buttocks

 When did this occur? (e.g. Feb 2005)

……/……….  - Tender or painful ‘glands’ (lumps or TICK ALL THAT APPLY nodes under the skin, usually away from  in the groin the bite mark)*  in the armpits  in the neck  other; where______

 When did this occur? (e.g. Feb 2005)

……/……….

- Aching in joints, limbs, back or neck  When did this occur? (e.g. Feb 2005) without reason* ……/……….

- Tiredness/ Feeling ‘out of energy’  When did this occur? (e.g. Feb 2005) without reason* ……/……….

- Shortness of breath or difficulty  When did this occur? (e.g. Feb 2005) breathing without reason* ……/……….

- Blistering/scabbing around skin area  When did this occur? (e.g. Feb 2005) where bitten* ……/……….

If the subject answered “At any time in the 2 weeks after the tick bite” to ANY OF THE SYMPTOMS, please continue. Otherwise, please go straight to question 13.

I would like to ask you about the [*symptoms/symptoms] which occurred in ……/………. (START WITH THE EARLIEST DATE). Thinking back to the

229 tick bites that occurred in the two weeks before these symptoms:

e. Where were you when you were bitten?  on Barrow lsland  elsewhere in Australia or overseas Location (eg name of area, park or district): ……………. State: ……………. Country: …………….

f. In which part(s) of your body did the bite(s) occur? (TICK ALL THAT APPLY)  on chest, back or abdomen (stomach region)  hands, arms or shoulders  feet or legs  on neck, face or scalp  on groin or buttocks

g. Did you remove the tick(s) yourself?  Yes  No: removed at a medical clinic  No : removed by someone else: who?......

h. Did you seek medical advice for the tick bite(s)?  Yes  No  Not sure

i. Were you treated with antibiotics after the tick bite(s)?  Yes  No  Not sure

j. Did you miss any work because of the tick bite(s)?  Yes  if yes , How many days in total? ......  No

NOW PROCEED TO THE NEXT DATE ON THE TABLE ABOVE: I would now like to ask you about the [*symptoms/symptoms] which occurred in ……/………. Thinking back to the tick bites that occurred in the two weeks before these symptoms:

e. Where were you when you were bitten?  on Barrow lsland  elsewhere in Australia or overseas Location (eg name of area, park or district): ……………. State: ……………. Country: …………….

f. In which part(s) of your body did the bite(s) occur? (TICK ALL THAT APPLY)  on chest, back or abdomen (stomach region)  hands, arms or shoulders  feet or legs  on neck, face or scalp  on groin or buttocks

g. Did you remove the tick(s) yourself?  Yes

230  No: removed at a medical clinic  No : removed by someone else: who?...... h. Did you seek medical advice for the tick bite(s)?  Yes  No  Not sure i. Were you treated with antibiotics after the tick bite(s)?  Yes  No  Not sure

j. Did you miss any work because of the tick bite(s)?  Yes  if yes , How many days in total? ......  No

231

NOW PROCEED TO THE NEXT DATE ON THE TABLE ABOVE: I would now like to ask you about the [*symptoms/symptoms] which occurred in ……/………. Thinking back to the tick bites that occurred in the two weeks before these symptoms:

e. Where were you when you were bitten?  on Barrow lsland  elsewhere in Australia or overseas Location (eg name of area, park or district): ……………. State: ……………. Country: …………….

f. In which part(s) of your body did the bite(s) occur? (TICK ALL THAT APPLY)  on chest, back or abdomen (stomach region)  hands, arms or shoulders  feet or legs  on neck, face or scalp  on groin or buttocks

g. Did you remove the tick(s) yourself?  Yes  No: removed at a medical clinic  No : removed by someone else: who?......

h. Did you seek medical advice for the tick bite(s)?  Yes  No  Not sure

i. Were you treated with antibiotics after the tick bite(s)?  Yes  No  Not sure

j. Did you miss any work because of the tick bite(s)?  Yes  if yes , How many days in total? ......  No

232 19. APART FROM ANY SYMPTOMS YOU HAVE MENTIONED ALREADY, have you EVER experienced any of the following since you started working on Barrow Island?

(Please tick all the symptoms you have had and answer the questions in that row. If you haven’t had any of the following symptoms then please go straight to question 20 )

Nature of Please How many times What time Do you remember being tick any have these of the year bitten by a tick anytime in reaction symptoms symptoms occurred did these the 2 weeks before these that you since you started symptoms symptoms developed? have had working on Barrow? occur? (eg: February)

Skin swelling/  Yes  if YES:  Yes tenderness  No  No  Not sure |_|_|_|  Can’t remember

Prolonged headache  Yes  if YES:  Yes (> 3 days)  No  No  Not sure |_|_|_|  Can’t remember

Rash over parts or all  Yes  if YES:  Yes of my body  No  No  Not sure |_|_|_|  Can’t remember

TICK ALL THAT APPLY  around the bite  on chest, back or abdomen (stomach region)  hands, arms or shoulders  feet or legs  on neck, face or scalp  on groin or buttocks

General aching in  Yes  if YES:  Yes joints, limbs, back or  No  No neck without reason  Not sure |_|_|_|  Can’t remember

Prolonged tiredness/  Yes  if YES:  Yes Feeling ‘out of energy’  No  No (> 3 days) without  Not sure |_|_|_|  Can’t remember reason Fever or chills  Yes  if YES:  Yes  No  No  Not sure |_|_|_|  Can’t remember

Shortness of breath or  Yes  if YES:  Yes difficulty breathing  No  No without reason  Not sure |_|_|_|  Can’t remember

233

OTHER MEDICAL HISTORY

20. Do you have any major medical conditions?

 Yes  No

If YES: Please describe ……………………………….……………………

……………………………………………………

21. Are you routinely on any medications or treatments?

 Yes  No

If YES:

a. What is the name of the medication?

……………………………….……………………

……………………………………………………

b. What condition or health problem do you take this for? ……………………………….……………………

……………………………………………………

Thank you for taking the time to complete this survey.

234 APPENDIX 8. RESULTS OF THE SEROSURVEY

Table A3. Titration results for the serology performed on respondents to the Barrow Island questionnaire.

Titration results Respondant austral honei conorii sibir rickett akari typhi prow No. 1 512 512 256 256 128 128 256 256 2 512 512 256 256 128 128 256 256 3 256 256 256 256 256 256 256 256 4 256 128 128 256 128 128 128 128 5 256 512 256 256 128 128 256 256 6 256 256 256 512 256 256 256 256 7 256 256 256 256 256 256 256 256 8 128 1024 128 256 256 128 256 256 9 512 256 256 256 256 256 512 512 10 256 256 256 256 256 128 256 256 11 512 256 256 256 256 512 512 512 12 256 512 256 256 256 256 256 256 13 256 256 256 256 256 256 512 512 14 256 512 256 256 256 128 256 256 15 256 256 256 256 128 256 256 256 16 256 256 256 512 256 256 512 512 17 256 1024 256 512 256 128 256 256 18 256 512 128 512 256 256 512 512 19 256 512 512 256 512 256 256 256 20 256 256 512 256 256 256 256 256 21 256 256 512 256 256 256 256 256 22 256 256 256 256 256 256 128 128 23 ------24 ------25 ------26 ------27 256 128 128 128 256 128 256 256 28 512 256 256 256 256 256 256 256 29 256 256 256 256 512 128 256 256 30 128 256 256 128 256 256 256 256 31 256 256 128 128 256 256 256 256 32 256 256 256 128 128 - 256 256 33 256 256 256 128 256 256 256 256 34 512 256 512 256 256 256 512 512 35 128 128 128 128 256 128 128 128 36 ------37 256 256 256 128 256 128 256 256 38 256 128 128 256 256 128 256 256 39 ------

235 40 128 128 256 128 256 256 256 256 41 256 128 512 128 128 128 256 256 42 256 128 256 128 256 128 256 256 43 256 128 256 128 128 128 256 256 44 ------45 128 256 256 256 128 256 256 256 46 128 256 256 256 128 256 256 256 47 256 256 256 256 256 128 256 256 48 ------49 128 256 128 129 256 128 256 256 50 256 128 128 128 512 256 128 128 51 256 512 256 256 256 256 128 128 52 256 256 256 128 256 128 256 256 53 ------54 256 128 256 256 256 256 256 256 55 ------56 512 128 256 128 128 128 128 128 57 256 256 256 256 256 128 256 256 58 256 256 256 256 256 256 256 256 59 128 128 128 128 256 128 128 128

austral – R. australis ; sibir – R. sibirica ; rickett – R. rickettsii ; prow – R. prowazekii. - no antibody titre.

236

237