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.

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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

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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 ticks) were opportunistically

collected from across the State, with an emphasis on native and feral animals 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,

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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 tick

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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.

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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 Taxonomy ……………………………………………………...……………...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

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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 ECTOPARA SITES

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

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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

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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

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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

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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

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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

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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 animal 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

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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

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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

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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

xvii

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.

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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 3rd century and it is suspected that epidemic typhus was

the cause of a plague described in Athens in the 5th 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

2

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

3

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

4

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).

5

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).

6

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

7

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 arthropod are often less host-specific than

adults, this must be taken into account when considering incidence and host-

specificity of the infecting rickettsiae (Reháček, 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-

8

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

(Reháček 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

9

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.

10

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

11

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).

12

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

13

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.

14

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 sca4 (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

15

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

16

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

17

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, ompA, 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).

18

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).

19

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

20

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).

21

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

22

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

23

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

24

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

25

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

26

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

27

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

28

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

29

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).

30

• 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 Hyalomma 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).

31

• 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

32

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).

33

• 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

34

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).

35

• 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).

36

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

37

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).

38

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).

39

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 Bartonella spp.

Members of the genus Bartonella are fastidious, intracellular, Gram-negative bacilli

that are capable of causing vector-transmitted zoonotic diseases. In the original

40

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 16th 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

41

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).

42

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.

43

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.

44

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.

45

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

46

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.

47

Figure 2.1 Native animal trapping on Barrow Island

Figure 2.2 Tick on the ear of a burrowing bettong

48

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

49

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

50

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

51

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 (fig.s 2.1 and 2.2)

Ectoparasite species No. of animals or 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 Antechinus

Dwellingup Ixodes antechini (fig.2.8) 15

Southern brown Bandicoot

Dwellingup Ixodes australiensis 1

Golden bandicoot Barrow Island Haemaphysalis ratti (fig.2.6) 34

Golden bandicoot Barrow Island Amblyomma limbatum 1

Bandicoot Kimberleys Haemaphysalis spp. 6 Common brushtail Possum

Barrow Island Amblyomma limbatum 15

Common brushtail Possum

Barrow Island Haemaphysalis ratti 4

Burrowing bettong Barrow Island Haemaphysalis ratti 18

Burrowing bettong Barrow Island Amblyomma limbatum 24

Western brush wallaby

Fitzgerald River National Park

Ixodes sp. 1

Western grey kangaroo

Perth Amblyomma triguttatum (fig.2.5)

4

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

National Park Ixodes myremecobii 1

Humans Waychinicup National Park

Ixodes myremecobii 1

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

52

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.

53

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.

54

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

55

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

56

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.

57

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

(Dasyurus 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.

58

Throughout the world, in tropical, temperate and even tundra regions, ticks are serious

veterinary and medical pests. As vectors, ticks surpass all other arthropods 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.

59

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.

60

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

61

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).

62

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

300µL 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

63

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 MgCl2 (Fisher

64

Biotech) concentration of 2mM was found to be optimal. The PCR premix was made

up to 17µL and 3µL 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, 1µL 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 20µL 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).

65

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 20µL 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 5µL of DNA template, 4µL 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 10µL 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.

66

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 MgCl2 concentration of 2mM was found to be

optimal. The PCR premix was made up to 49µL and 1µL 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).

67

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.

68

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

albolimbatum

2/17

Bobtail lizard Lake Magenta Amblyomma

albolimbatum

0/1

Olive python Kimberley Amblyomma spp. 0/1 Yellow-footed Antechinus

Dwellingup Ixodes antechini 8/15

Southern brown Bandicoot

Dwellingup Ixodes australiensis 0/1

Golden bandicoot Barrow Island Haemaphysalis ratti 2/34

Golden bandicoot Barrow Island Amblyomma limbatum 0/1

Bandicoot Kimberleys Haemaphysalis spp. 2/6 Common brushtail Possum

Barrow Island Amblyomma limbatum 7/15

Common brushtail Possum

Barrow Island Haemaphysalis ratti 0/4

Burrowing bettong Barrow Island Haemaphysalis ratti 3/14

Burrowing bettong Barrow Island Amblyomma limbatum 4/4

Western brush wallaby Fitzgerald River National Park

Ixodes spp. 0/1

Western Grey Kangaroo

Perth Amblyomma triguttatum 4/4

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

National Park Ixodes myremecobii 0/1

Humans Waychinicup National Park

Ixodes myremecobii 0/1

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

69

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.

70

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 Rickettsia sp. strain BWI-1

% Similarity to 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%

71

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

Rickettsia sp. strain BWI-1

% Similarity to

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

72

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

73

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.

74

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

75

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

76

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

77

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

78

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.

79

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

80

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

81

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.

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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

83

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).

84

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

85

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 25cm2 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.

86

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.

87

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

3’

(Stenos et al., 1998) 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

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

16S

rRNA

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

16S

rRNA

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

16S

rRNA

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

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

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

88

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 conditions (see table 4.1)

Primer concentration (µµµµM)

MgCl2

Concentration (mM)

Taq (u/tube)

Premix volume (µµµµL)

DNA volume (µµµµL)

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.

89

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

90

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).

91

Table 4.5. Percentage similarity between Rickettsia sp. strain BWI-1 and Rickettsia

species with validly published names.

Name % Similarity 16S rRNA

GltA ompA ompB sca4

R. prowazekii 98.8% 92.4% NS 76.7% 76.5%

R. typhi 98.7% 92.2% NS 81.5% 79.7%

R. africae 99.5% 97.6% 93.8% 92.7% 95.3%

R. conorii 99.5% 98.1% 92.7% 91.9% 95.3%

R. sibirica 99.5% 98.2% 93.4% 93.2% 95.6%

R. honei 99.5% 98.0% 93.2% 92.3% 96.0%

R. slovaca 99.5% 98.1% 94.0% 95.6% 96.0%

R. parkeri 99.4% 98.2% 93.2% 92.5% 95.2%

R. rickettsii 99.5% 97.8% 92.9% 94.5% 95.9%

R. massiliae 99.7% 98.0% 95.8% 97.1% 95.8%

R. rhipicephali 99.7% 97.9% 95.6% 97.4% 95.7%

R. aeschlimannii 99.5% 98.4% 95.1% 96.9% 96.5%

R. montanensis 99.3% 98.3% 94.5% 96.0% 95.3%

R. japonica 99.5% 97.6% 93.4% 93.8% 96.0%

R. heilongjiangii 99.4% 98.0% 94.3% 93.8% 96.3%

R. helvetica 99.2% 95.8% NS 51.8% 89.2%

R. felis 99.5% 93.4% 53.7% NS 88.6%

R. akari 98.6% 93.4% NS 85.7% 84.7%

R. australis 99.4% 93.7% 85.2% 87.7% 86.0%

R. bellii 99.2% 85.7% NS NS NS

NS – not sequenced.

92

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.

R. parkeri R. sibirica

R. conorii

R. africae R. rickettsii R. honei

R. slovaca R. japonica

R. heilongjiangii R. montanensis

Rickettsia sp. BWI-1 R. aeschlimannii

R. massiliae R. rhipicephali

R. prowazekii R. typhi

R. helvetica R. felis

R. bellii R. akari

R. australis

100

93

0.2%

93

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.

.

R. rickettsii R. conorii R. honei

R. africae R. sibirica R. parkeri R. slovaca

R. japonica R. heilongjiangii

R. aeschlimannii R. massiliae R. rhipicephali

Rickettsia sp. BWI-1 R. montanensis

R. helvetica R. felis

R. akari R. australis R. prowazekii R. typhi

R. bellii 100

99 98

87 94

97

98

99 78

2%

94

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.

.

R. africae R. parkeri

R. sibirica R. conorii

R. slovaca R. honei

R. rickettsii R. japonica

R. heilongjiangii R. aeschlimannii

R. massiliae R. rhipicephali

Rickettsia sp. BWI-1 R. montanensis

R. australis

93 100

100

98 99

71

2%

95

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.

.

R. africae R. parkeri

R. sibirica R. mongolotimonae

R. conorii R. slovaca R. rickettsii

R. honei R. japonica R. heilongjiangii

Rickettsia sp. BWI-1 R. montanensis

R. aeschlimannii R. massiliae R. rhipicephali

R. akari R. australis

R. prowazekii R. typhi 100

98

100

97

96 88

76 99

99

71

5%

96

Error!

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.

.

R. conorii R. parkeri R. sibirica R. africae

R. slovaca R rickettsii R. honei R. japonica

R. heilongjiangii R. montanensis R. aeschlimannii

R. massiliae R. rhipicephali

Rickettsia sp. BWI-1 R. felis

R. akari R. australis

R. helvetica R. prowazekii

R. typhi 100

100 99

100

97 81

74

84

5%

97

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).

98

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

99

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.,

100

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.

101

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.

102

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

103

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 µL of elution buffer rather than 200µL as

104

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).

105

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).

106

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

I.a. – Ixodes antechini; A.j. – Acanthopsylla jordani

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.)

107

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

108

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.

109

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

R. gravesii 98.4% 96.3% 96.7%

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%

R. rhipicephali 99.0% 95.4% 97.1%

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

110

Table 5.3. Comparison of Bartonella sp. strain Mu1 and previously characterized

species of Bartonella based on gltA gene sequences.

Name % Similarity

B. quintana 84.4%

B. henselae 85.7%

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%

111

Error!

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.

R. rickettsii R. conorii R. honei R. africae

R. sibirica R. parkeri R. slovaca

R. japonica R. heilongjiangii

R. aeschlimannii R. massiliae

R. rhipicephali R. gravesii

Rickettsia sp. D-1 R. montanensis

R. helvetica R. felis

R. akari R. australis R. prowazekii R. typhi

R. bellii 100

99 98

82

93

99

95 100

83

2%

112

Error!

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.

R. africae R. parkeri

R. sibirica R. conorii

R. slovaca R. rickettsii

R. honei R. japonica

R. heilongjiangii Rickettsia sp. D-1

R. aeschlimannii R. massiliae R. rhipicephali

R. gravesii

R. montanensis R. australis

94 100

100

84

93 93

75

70 99

2%

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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.

R. africae R. parkeri

R. sibirica R. mongolotimonae

R. conorii R. slovaca R. rickettsii

R. honei R. japonica R. heilongjiangii

R. gravesii R. montanensis

Rickettsia sp. D-1 R. aeschlimannii

R. massiliae R. rhipicephali

R. akari R. australis

R. prowazekii R. typhi 100

98

100

96 94

91 77

99

100

5%

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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.

B. bacilliformis

B. doshiae

B. clarridgeiae

Bartonella sp. Mu1

B. quintana

B. henselae

B. koehlerae

Bartonella sp. Strain40

B. taylorii

B. vinsonii

Bartonella sp. StrainC1phy

Bartonella sp. StrainR1phy

Bartonella sp. StrainC4phy

Bartonella sp. StrainR2phy

B. tribocorum

B. grahamii

Bartonella sp. StrainC7rat

B. elizabethae

B. peromysci

90

99

70

76

20

115

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

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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

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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

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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

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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.

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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

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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.

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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

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participant and allowed to clot at room temperature before serum was removed,

aliquoted and stored at 4oC 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

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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

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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.

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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,

tiredness, scabbing, shortness of breath 10.2

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,

tiredness, scabbing, shortness of breath 8.5

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

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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).

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Table 6.2 Summary of odds ratios (with 95% confidence intervals) for exposure variables versus rickettsial serology

Variable Categories Unadjusted odds ratios

95% CI

Adjusted odds ratios **

95% CI

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 bite

No 1.00 1.00

Yes 0.52 0.09 3.11 0.44 0.07 2.66

Any history of unexplained systemic symptoms; no bite history

No 1.00 1.00

Yes 0.27 0.03 2.57 0.24 0.03 2.34

Total bites (years on island x intensity of tick bites)

Low 1.00 1.00

High 0.91 0.32 2.65 0.68 0.22 2.11

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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

130

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

131

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)

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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

133

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.

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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

135

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.

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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

137

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.

138

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

139

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

140

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

141

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.

142

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

143

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

144

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.

145

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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)

191

SFG (spotted fever group) TAE (tris-acetate-EDTA) TG (typhus group) TIBOLA (“tick-borne lymphadenopathy”) WA (Western Australia) WHO (World Health Organisation)

192

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.

193

APPENDIX 3. COMMON AND SCIENTIFIC NAMES

Black-footed rock wallaby (Petrogale lateralis)

Bobtail lizard (Trachysaurus 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 (Dasyurus geoffroyi)

Yellow-footed antechinus (Antechinus flavipes)

194

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

195

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

196

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

197

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

198

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

199

APPENDIX 6. MINIMUM EVOLUTION AND MAXIMUM PARSIMONY TREES.

Figure A4.1 Minimum evolution tree based on 16S rRNA sequences.

R. parkeri R. sibirica

R. conorii R. africae R. honei

R. slovaca R. rickettsii

R. japonica R. heilongjiangii

R. montanensis R. aeschlimannii

R. massiliae R. rhipicephali

Rickettsia sp.BWI-1 R. helvetica

R. felis R. prowazekii

R. typhi R. bellii

R. akari R. australis

100

93

0.2%

200

Figure A4.2 Maximum parsimony tree based on 16S rRNA sequences.

R. conorii R. sibirica R. africae R. parkeri R. honei R. rickettsii R. slovaca R. japonica R. heilongjiangii R. aeschlimannii Rickettsia sp.BWI-1 R. massiliae R. rhipicephali R. montanensis R. helvetica R. felis R. akari R. prowazekii R. typhi R. bellii R. australis

94

99

201

Figure A4.3. Minimum evolution tree based on gltA sequences.

R. sibirica R. parkeri

R. africae R. slovaca R. honei

R. rickettsii R. conorii

R. japonica R. heilongjiangii

R. aeschlimannii R. massiliae R. rhipicephali

Rickettsia sp. BWI-1 R. montanensis

R. helvetica R. felis R. akari

R. australis R. prowazekii R. typhi

R. bellii 100

99

98

92 81

94

99

100 76

72

2%

202

Figure A4.4. Maximum parsimony tree based on gltA sequences.

R. rickettsii R. conorii R. honei R. slovaca R. africae R. sibirica R. parkeri R. japonica R. heilongjiangii Rickettsia sp.BWI11

R. aeschlimannii R. massiliae R. rhipicephali R. montanensis R. helvetica R. felis R. akari R. australis R. prowazekii R. typhi R. bellii

97

96 70

86

100

99 96

83

203

Figure A4.5 Minimum evolution tree based on ompA sequences.

R. africae R. parkeri

R. sibirica R. conorii R. rickettsii

R. slovaca R. honei

R. japonica R. heilongjiangii

R. aeschlimannii R. massiliae R. rhipicephali

Rickettsia sp. BWI-1

R. montanensis R. australis

93 100

100 84

97

98

2%

204

Figure A4.6 Maximum parsimony tree based on ompA sequences.

R. africae R. sibirica R. parkeri R. slovaca R. honei R. rickettsii R. conorii R. japonica

R. heilongjiangii Rickettsia sp. BWI-1

R. aeschlimannii R. massiliae R. rhipicephali R. montanensis R. australis

78 99

97

99

94

77

205

Figure A4.7 Minimum evolution tree based on ompB sequences.

R .africae R .parkeri

R. sibirica R. mongolotimonae

R. conorii R. slovaca R. rickettsii

R. honei R. japonica R. heilongjiangii

R. aeschlimannii Rickettsia sp. BWI-1

R. montanensis R. massiliae

R. rhipicephali R. akari

R. australis R. prowazekii

R. typhi 100

99

100

76

94 92

85 80

99

71

99

77

5%

206

Figure A4.8. Maximum parsimony tree based on ompB sequences.

R. africae R .parkeri R. sibirica R. mongolotimonae R. conorii R. slovaca R. rickettsii R. honei R. aeschlimannii R. massiliae R. rhipicephali Rickettsia sp. BWI-1 R. montanensis R. japonica R. heilongjiangii R. akari R. australis R. prowazekii R. typhi

80 83

92

89

95

96

99

99

100

207

Figure A4.9. Minimum evolution tree based on sca4 sequences.

R. conorii R. parkeri R. sibirica R. africae

R. slovaca R. rickettsii R. honei R. japonica

R heilongjiangii R. montanensis R. aeschlimannii

R. massiliae R. rhipicephali

Rickettsia sp. BWI-1 R. felis

R. akari R. australis

R. prowazekii

R. typhi 100

100 100

83 91

100

98

5%

208

Figure A4.10. Maximum parsimony tree based on sca4 sequences.

R. conorii R. parkeri R. sibirica R. africae

R. honei R. rickettsii

R. slovaca R. japonica R. heilongjiangii R. montanensis R. aeschlimannii R. massiliae R. rhipicephali Rickettsia sp.BWI1 R. felis R. akari R. australis R. helvetica R. prowazekii R. typhi

100

83

71

99

74

100

90

100

209

Figure A4.11. Minimum evolution tree based on gltA sequences (Rickettsia sp. D-1).

R. sibirica R. parkeri

R. africae R. slovaca R. honei

R. rickettsii R. conorii

R. japonica R. heilongjiangii

R. gravesii Rickettsia sp. D-1

R. aeschlimannii R. massiliae R. rhipicephali

R. montanensis R. helvetica

R. felis R. akari

R. australis R. prowazekii R. typhi

R. bellii 100

99 98

89

83

99

94

100 75

2%

210

Figure A4.12. Maximum parsimony tree based on gltA sequences.

R. rickettsii R. conorii R. honei R. sibirica R. parkeri R. slovaca R. africae R. japonica R. heilongjiangii R. gravesii Rickettsia sp. D-1 R. aeschlimannii R. massiliae

R. rhipicephali R. montanensis R. helvetica R. felis R. akari R. australis R. prowazekii R. typhi R. bellii

98

95 70

99

90

96

84

100

211

Figure A4.13. Minimum evolution tree based on ompA sequences.

R. africae R. parkeri

R. sibirica R. conorii R. rickettsii

R. honei R. slovaca

R. japonica R. heilongjiangii

Rickettsia sp. D-1 R. aeschlimannii

R. massiliae R. rhipicephali

R. gravesii R. montanensis

R. australis

94 99

100

85

91

94

99

2%

212

Figure A4.14. Maximum parsimony tree based on ompA sequences.

R. africae R. parkeri R. sibirica R. conorii R. rickettsii R. honei R. slovaca R. japonica R. heilongjiangii Rickettsia sp. D-1 R. aeschlimannii R. massiliae R. rhipicephali R. gravesii R. montanensis R. australis

99

98

93

76 99

90

85

213

Figure A4.15. Minimum evolution tree based on ompB sequences.

R. africae R. parkeri

R. sibirica R. mongolotimonae

R. conorii R. slovaca R. rickettsii

R. honei R. japonica R. heilongjiangii

R. aeschlimannii R. gravesii

Rickettsia sp. D-1 R. montanensis

R. massiliae R. rhipicephali

R. akari R. australis

R. prowazekii R. typhi 100

99

99

77

94

90 88

82 99

99

5%

214

Figure A4.16. Maximum parsimony tree based on ompB sequences.

R. africae R. parkeri R. sibirica R. mongolotimonae R. conorii R. slovaca R. rickettsii R. honei R. japonica R. heilongjiangii R. aeschlimannii R. gravesii R. massiliae R. rhipicephali Rickettsia sp. D-1 R. montanensis R. akari R. australis R. prowazekii R. typhi

97

80 83

92

90

96

99 99

100

215

Figure A.17. Minimum evolution tree based on gltA sequences (Bartonella sp. strain Mu1).

B. elizabethae

Strain C7rat

B. grahamii

StrainR1phy

Strain C4phy

Strain R2phy

B. taylorii

B. vinsonii

Strain C1phy

B. bacilliformis

B. doshiae

Strain40 Bartonella sp.strain Mu1

B. quintana

B. henselae

100

100

100

99

85

95

79

0.02

216

Figure A.18. Maximum-Parsimony Tree Based on gltA Sequences (Bartonella sp. strain Mu1).

B. elizabethae

Strain C7rat

B. grahamii

Strain R1phy

Strain C4rphy

Strain R2phy

Strain 40 Bartonella sp.strain Mu1

B. quintana

B. henselae

B. bacilliformis

B. doshiae

B. vinsonii

Strain C1phy

B. taylorii

98

100

79

99

95

217

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.

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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.

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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

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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 st udy 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 inform ation 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.

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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

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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

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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.

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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

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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

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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

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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

in the bush or forest in other parts of Western Australia (except for Barrow

Island) or interstate

(please tick the answer that most applies to you)

Bushwalking, hiking, camping or working in the bush or forest overseas (outside

Australia)

(please tick the answer

that most applies to you)

� Not at all in past 2 years

� Not at all in past 2 years

� Less than once a year � Which state(s)/territories?…………

� Less than once a year

� Which country/countries?…………

� About once a year � Which state(s)/territories?…………

� About once a year

� Which country/countries?…………

� More than once a year � Which state(s)/territories?…………

� More than once a year

� Which country/countries?…………

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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 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

- 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 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

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- 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 nodes under the skin, usually away from the bite mark)*

TICK ALL THAT APPLY � in the groin � in the armpits � in the neck � other; where_____________

� When did this occur? (e.g. Feb 2005) ……/……….

- Aching in joints, limbs, back or neck without reason*

� When did this occur? (e.g. Feb 2005) ……/……….

- Tiredness/ Feeling ‘out of energy’ without reason*

� When did this occur? (e.g. Feb 2005) ……/……….

- Shortness of breath or difficulty breathing without reason*

� When did this occur? (e.g. Feb 2005) ……/……….

- Blistering/scabbing around skin area where bitten*

� When did this occur? (e.g. Feb 2005) ……/……….

����If the subject answered “At any time in the 2 weeks after the tick bite” to ANY OF THE SYMPTOMS, please continue. Otherwise, pl ease 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 th e

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tick bites that occurred in the two weeks before th ese 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 th e 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

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� 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

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����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 th e 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

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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 th en please go straight to question 20 )

Nature of reaction

Please tick any

symptoms that you have had

How many times have these

symptoms occurred since you started

working on Barrow?

What time of the year did these

symptoms occur? (eg:

February)

Do you remember being bitten by a tick anytime in the 2 weeks before these symptoms developed?

Skin swelling/ tenderness

� Yes�� � No � Not sure

if YES:

|_|_|_|

� Yes � No � Can’t remember

Prolonged headache (> 3 days)

� Yes�� � No � Not sure

if YES:

|_|_|_|

� Yes � No � Can’t remember

Rash over parts or all of my body

� Yes�� � No � Not sure

if YES:

|_|_|_| 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

� Yes � No � Can’t remember

General aching in joints, limbs, back or neck without reason

� Yes�� � No � Not sure

if YES:

|_|_|_|

� Yes � No � Can’t remember

Prolonged tiredness/ Feeling ‘out of energy’ (> 3 days) without reason

� Yes�� � No � Not sure

if YES:

|_|_|_|

� Yes � No � Can’t remember

Fever or chills � Yes��

� No � Not sure

if YES:

|_|_|_|

� Yes � No � Can’t remember

Shortness of breath or difficulty breathing without reason

� Yes�� � No � Not sure

if YES:

|_|_|_|

� Yes � No � Can’t remember

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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 surv ey.

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APPENDIX 8. RESULTS OF THE SEROSURVEY

Table A3. Titration results for the serology performed on respondents to the Barrow Island questionnaire. Respondant No.

austral honei conorii sibir rickett akari typhi prow

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 - - - - - - - -

Titration results

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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.

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