An Epidemiological and Serological Study of Rickettsia in Western Australia
Presented By
Mohammad Yazid Abdad
Submitted in total fulfilment of the requirement of the degree of
Doctor of Philosophy
School of Veterinary and Biomedical Sciences, Division of Health Sciences, Murdoch University
November 2011
Declaration
I declare this thesis is my own account of my research and contains, as its main content, work which has not been previously submitted for a degree at any tertiary institution.
This thesis is less than 100,000 words in length, inclusive of tables, figures, bibliography and appendices, and it complies with the stipulations set out for the degree of Doctor of Philosophy by Murdoch University.
Mohammad Yazid Abdad School of Veterinary and Biomedical Sciences Division of Health Sciences Murdoch University
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Abstract
The study was aimed at investigating Western Australian rickettsiae, delving deeper into the epidemiology of a recently described rickettsia, Rickettsia gravesii, and any other rickettsiae lurking in the Western Australian bush. Prior to the discovery of R. gravesii, only Rickettsia typhi was known to be endemic to Western Australia. With the addition of R. gravesii and another novel rickettsia named candidatus “Rickettsia antechini”, understanding and investigation of the potential effects these organisms may have on human and animal health becomes paramount. This research adds to the limited information available in the literature pertaining to Australian rickettsial organisms.
Serotyping of R. gravesii in mice demonstrated distinct specificity differences between
R. gravesii, Rickettsia massiliae, Rickettsia australis and Rickettsia honei. R. massiliae was chosen due to R. gravesii’s very close genotypic similarities to the R. massiliae sub-group. R. australis and R. honei were chosen for the possibility of them sharing a similar geographical distribution with R. gravesii.
Collection of ectoparasites, in particular ticks, from humans and larger mammals in south-west Western Australian bush was performed to investigate the prevalence of R. gravesii in tick populations and also to investigate any other rickettsial organisms that may be present. No other rickettsiae were uncovered except for R. gravesii among ixodid ticks collected. Four tick species collected during the course of the study harboured R. gravesii with prevalences of up to 15% in Amblyomma albolimbatum
(n=45), 75% in Amblyomma triguttatum (n=187), 51% in Ixodes australiensis (n=63) and 25% in Ixodes fecialis (n=8). No rickettsial DNA was detected in Haemaphysalis ii longicornis, though only a single specimen was found. Transovarial transmission of R. gravesii in A. triguttatum ticks from infected females to viable larvae was observed by
PCR. Detection of R. gravesii in the salivary glands, ovaries and gut suggests the potential of horizontal, transovarial and transtadial transmission.
Feral pigs (n=148) trapped within the water-catchment areas for Perth were tested for
SFG rickettsiae and a prevalence of 49% was observed by microimmunofluorescence at titres of 1:128 and higher. A baseline control group made up of domestic farmed pigs
(n=67) was used to determine the cut-off titre (1:128) where no non-specific antigen- antibody detectable by microscopy was observed.
Sero-prevalence in humans was also investigated. Volunteers that are exposed to ticks as a result of occupational and recreational activities were recruited over a 3 year period. Sera were collected and a questionnaire asking about participants’ bush activities was filled out. Two occupational groups were involved in the study, workers from Barrow Island (n=67) and Whiteman Park (n=12). The recreational group was made up of members of the Western Australian Rogaining Association (n=104). The control group used in the study was made up of volunteers among the staff and students of Murdoch University (n=60). Rickettsial prevalence was observed at higher levels in the occupational groups than recreational group participants; 44.8% in Barrow Island workers, 50% in Whiteman Park staff, 23.4% in rogainers and 1.7% in the control group.
Analysis of the questionnaires filled out by participants from the human sero- prevalence study demonstrated a significant association between bush activity and exposure to rickettsial organisms and their vectors. A follow-up of volunteers 10-14
iii months after the first serum collection from Whiteman Park staff (n=8) and among the rogainers (n=62) showed no significant rates of seroconversion or seroreversion.
Odds ratio analysis (adjusted for age and gender) based on questionnaire results and sero-prevalence of rickettsial antibodies in the different interest groups and control cohort showed that the recreational group had an odds ratio of 13.70 (95% CI=1.73-
108.49). Odds ratio for the occupational group is 38.81 (95% CI=5.02-300.34). This shows a significant risk for people performing activities in the bush for extended periods of time, with occupational activities posing the highest risk, potentially due to the longer hours compared to recreational activities in general.
The presence of R. gravesii in Western Australia and the data reported in this study underscores the lack of information available to public health authorities with regards to rickettsial organisms and their hosts. Even though no official reports of debilitating spotted fever infections have been reported in WA in the literature thus far, the potential of it happening cannot be disregarded. Increased awareness of rickettsial organisms and infection need to be instilled among public health officials, doctors and veterinarians. Other animal populations and parts of Western Australia not covered in this study need to be further investigated.
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Acknowledgements
First and foremost, I would like to acknowledge and express my deep appreciation towards my three supervisors. Professor Stan Fenwick, whose ideas and enthusiasm had inspired me from the beginning and sticking by me from start to finish. Professor
Cassandra Berry, who had sparked my interest in immunology and all things infectious.
Professor Ian Robertson, who spent countless hours at the start introducing epidemiological research to me.
I am eternally grateful to Professor Stephen Graves, A/Professor John Stenos and Ms
Chelsea Nguyen of the Australian Rickettsial Reference Laboratory, VIC, who in no small part had given this study the needed resource support in the form of laboratory space, culturing facilities, rickettsial strains and validation of my results. They have stuck by me from start to finish.
My sincerest gratitude to Professor Angus Cook of the University of Western Australia and Dr John Dyer of Fremantle Hospital had given me numerous crucial advice and assistance on the direction of my research into the clinical and epidemiological aspect of Rickettsiology.
I am extremely grateful to Professor Pierre-Edouard Fournier of the Unite des
Rickettsies, Marseille, for the time and effort he had put in to introduce me to the field of Rickettsiology from the early stages of my research.
Dr Stuart Blacksell of the Wellcome Trust were kind enough to lend their expertise and host a significant portion of my experiments in their BSL3 laboratory and animal v holding facilities at Mahidol University in Bangkok, Thailand, for which I am most appreciative.
My sincerest gratitude to Dr Peter Adams and Andrew Li for the collection of feral pig and tick samples from the bush and allowing me to use them. Also the rest of the
Trailer Trash, an international group of “rejects” thrown out into a miserable corner of campus, which fortuitously is a couple of steps from the best and cheapest food on campus. Thank you, for all the light banter, international food, friendship and support you shared with me.
I am extremely grateful to the many study volunteers who helped by contributing their blood and time to allow the completion of a significant portion of my thesis. My sincerest thanks to Dr Helen Owen for allowing me the use of her questionnaire data from Barrow Island employees who volunteered for her research.
To Dr Bong Sze How, his friendship and support kept me sane throughout the duration of my candidature. Thank you for being an awesome friend and the brother I never had.
Last but not least, I would like to express my gratitude to my mother (Rosiah Mardi), and sisters (Fawziah Abdad and Sa’adiah Abdad), for their undying support and sacrifice. My sincerest apologies and regret for missing out on so many celebrative occasions and not being on site during moments of tragedy.
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In Memoriam
Embarak Moksin Abdad
1952-2009
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Table of Contents
Declaration ...... i Abstract ...... ii Acknowledgements ...... v Table of Contents ...... viii List of Abbreviations ...... xiii List of Figures and Tables ...... xvi 1 - Introduction ...... 1 1.1 Thesis Introduction ...... 1 1.2 Study Aims and Objectives ...... 3 1.3 Rickettsia ...... 4 1.3.1 Taxonomy ...... 6 1.3.2 Genome ...... 8 1.3.3 Spotted Fever Group (SFG) Rickettsiae ...... 10 1.3.4 Typhus Group (TG) Rickettsiae ...... 13 1.3.5 Ancestral Group (AG) Rickettsiae ...... 13 1.3.6 Transitional Group (TRG) Rickettsiae ...... 13 1.4 Rickettsial Vectors ...... 14 1.4.1 Ticks ...... 14 1.4.2 Fleas ...... 16 1.4.3 Lice ...... 17 1.4.4 Mites ...... 18 1.5 Rickettsia Pathogenesis ...... 19 1.5.1 Transmission ...... 19 1.5.2 Rickettsiae-Host Interactions ...... 19 1.5.3 Pathology ...... 22 1.6 Diagnosis and Treatment ...... 24 1.6.1 Serological Methods ...... 26 1.6.2 Cell Culture ...... 27 1.6.3 Polymerase Chain Reaction (PCR) ...... 28 1.6.4 Treatment ...... 30 1.7 Tick-Borne Rickettsioses ...... 31 viii
1.7.1 Rickettsia rickettsii ...... 31 1.7.2 Rickettsia conorii ...... 32 1.7.3 Rickettsia africae ...... 34 1.7.4 Rickettsia sibirica ...... 35 1.7.5 Rickettsia heilongjiangensis ...... 36 1.7.6 Rickettsia japonica ...... 36 1.7.7 Rickettsia slovaca ...... 37 1.7.8 Rickettsia aeschlimanii ...... 37 1.7.9 Rickettsia massiliae ...... 38 1.7.10 Rickettsia honei ...... 38 1.7.11 Rickettsia parkeri ...... 39 1.8 Non Tick-Borne Rickettsiosis ...... 40 1.8.1 Rickettsia typhi ...... 40 1.8.2 Rickettsia prowazekii ...... 41 1.8.3 Rickettsia akari ...... 42 1.8.4 Rickettsia felis ...... 42 1.8.5 Orientia tsutsugamushi ...... 45 1.9 The Australian Situation ...... 46 2 - General Materials and Methods ...... 52 2.1 Laboratory Conditions Used for Studies...... 52 2.2 Identification of Tick Species ...... 52 2.3 DNA Extraction ...... 55 2.4 Rickettsial Culture ...... 55 2.4.1 Cell Monolayer Cultures ...... 55 2.4.2 Rickettsia Isolation ...... 56 2.4.3 Rickettsia Culture Maintenance ...... 57 2.5 Serology ...... 57 2.5.1 Rickettsial Antigen Preparation ...... 57 2.5.2 Immunofluorescence Assay ...... 57 2.6 Rickettsial PCR ...... 58 2.6.1 Conventional PCR ...... 58 2.6.2 Sequence Analysis of PCR Product ...... 59 3 - Serological Typing of a Novel Spotted Fever Group Rickettsia: Rickettsia gravesii ...... 61 3.1 Introduction ...... 61 ix
3.2 Materials and Methods ...... 63 3.2.1 Rickettsial Suspension Preparation ...... 63 3.2.2 Plaque Assay Culture ...... 63 3.2.3 Measurement of Protein (Rickettsial Antigen) Concentration of Rickettsial Preparation ...... 64 3.2.4 Inoculation of Rickettsiae into Mice and Harvesting Anti-Sera ...... 65 3.2.5 Microimmunofluorescence ...... 66 3.2.6 ELISA ...... 66 3.2.7 Statistical Methods ...... 68 3.3 Results ...... 69 3.3.1 Plaque Assay ...... 69 3.3.2 Protein Concentrations ...... 69 3.3.3 Microimmunofluorescence ...... 70 3.3.4 ELISA ...... 72 3.4 Discussion ...... 73 4 - Amblyomma triguttatum as the Main Arthropod Reservoir of Rickettsia gravesii sp. nov. in Western Australia ...... 77 4.1 Introduction ...... 77 4.2 Materials and Methods ...... 78 4.2.1 Ectoparasite Sampling ...... 78 4.2.2 Ectoparasite Identification ...... 78 4.2.3 Ectoparasite Dissection...... 78 4.2.4 Transfer of Rickettsiae from Female Parent to Larvae...... 82 4.2.5 Rickettsial Isolation by Cell Culture ...... 82 4.2.6 DNA Extraction ...... 82 4.2.7 Rickettsial PCR Screening ...... 83 4.2.8 DNA Sequencing ...... 83 4.3 Results ...... 84 4.3.1 Rickettsia gravesii prevalence in ticks...... 84 4.3.2 Prevalence of R. gravesii in organs of A. triguttatum ...... 86 4.3.3 Transovarial transmission of R. gravesii in A. triguttatum ticks ...... 86 4.4 Discussion ...... 87 5 - Serological Study of Spotted Fever Group Rickettsial Exposure in Feral Pigs (Sus scrofa) of South West WA ...... 93 5.1 Introduction ...... 93
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5.2 Materials and Methods ...... 94 5.2.1 Sample Collection ...... 94 5.2.2 DNA Extraction ...... 95 5.2.3 Rickettsial PCR Screening ...... 95 5.2.4 Microimmunofluorescence ...... 95 5.3 Results ...... 98 5.4 Discussion ...... 101 6 - Rickettsial Exposure in High Risk Occupational and Recreational Groups Western Australia ...... 104 6.1 Introduction ...... 104 6.2 Materials and Methods ...... 106 6.2.1 Study Design ...... 106 6.2.2 Blood Collection ...... 107 6.2.3 Microimmunofluorescence ...... 108 6.2.4 Questionnaire Design ...... 108 6.2.5 Questionnaire Implementation ...... 109 6.2.6 Statistical Analysis ...... 109 6.3 Results ...... 110 6.3.1 Serology (IFA) Results...... 110 6.3.2 Descriptive Epidemiology ...... 113 6.3.3 Odds Ratio (OR) Analysis ...... 116 6.3.4 Spotted Fever Group Rickettsiosis in a WA Patient ...... 118 6.4 Discussion ...... 120 6.4.1 Sero-positivity and its Link to Bush Activity ...... 120 6.4.2 Potential Causes of Bias and Confounding Factors in the Study ...... 122 6.4.3 Public Health Implications of the Study ...... 125 7 - Final Discussion ...... 127 7.1 Fulfilment of Study Aims ...... 127 7.2 Rickettsia gravesii sp. nov...... 133 7.3 Future Directions ...... 136 8 - Bibliography ...... 140 9 - Appendices ...... 168 9.1 Ethics Approval Obtained ...... 168 9.2 GenBank Accession Numbers ...... 169 9.3 Rogainer Study Consent Form ...... 170 xi
9.4 Rogainers’ Questionnaire...... 177 9.5 Rogainers’ Follow-up Questionnaire ...... 184 9.6 Whiteman Park Workers’ Questionnaire ...... 194 9.7 Whiteman Park Workers’ Follow-up Questionnaire ...... 209 9.8 Control/Baseline Group Questionnaire ...... 220
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List of Abbreviations
AF Astrakhan Fever
ARC Australian Research Council
ARRL Australian Rickettsial Reference Laboratory
CBD Central Business District
CD4 Cluster of Differentiation 4
CD8 Cluster of Differentiation 8
CI Confidence Interval
CPE Cytopathic Effect
DEBONEL Dermacentor Borne Necrosis Erythema Lymphadenopathy
DNA Deoxyribonucleic Acid
EDTA Ethylenediaminetetra-Acetic Acid
ELISA Enzyme Linked Immunosorbent Assay
FBSF Flea-Borne Spotted Fever/Cat Flea Typhus
GTPase Guanosine Triphosphatase
HVE Healthy Volunteer Effect
ID50 Infectious Dose 50%
IFA Immunofluorescent Assay
IgA Immunoglobulin A
IgG Immunoglobulin G
IgM Immunoglobulin M
IL-1 Interleukin-1
IL-8 Interleukin-8
IL-12 Interleukin-12
INF Interferon xiii
INF-α Interferon Alpha
INF-β Interferon Beta
INF-γ Interferon Gamma
ISF Israeli Spotted Fever
ITT Indian Tick Typhus
LAR Lymphangitis-Associated Rickettsiosis
LD50 Lethal Dose 50%
MCP-1 Monocyte Chemotactic Protein-1
MIF Microimmunofluorescence
MSF Mediterranean Spotted Fever
NHMRC National Health and Medical Research Council
NK Natural Killer
OD Optical Density
OmpA Outer Membrane Protein A
OmpB Outer Membrane Protein B
OR Odds Ratio
PBS Phosphate Buffered Saline
PCR Polymerase Chain Reaction
RMSF Rocky Mountain Spotted Fever
RNA Ribonucleic Acid rRNA Ribosomal Ribonucleic Acid
SD50 Seroconversion Dose 50%
SFG Spotted Fever Group
SPD Specificity Difference
STG Scrub Typhus Group
STT Siberian Tick Typhus
TCID50 Tissue Culture Infectious Dose 50%
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TG Typhus Group
TIBOLA Tick-Borne Lymphadenopathy
TNF-α Tumour Necrosis Factor Alpha
TRG Transitional Group
TMB Tetramethylbenzidine
WA Western Australia
WARA Western Australia Rogaining Association
WASP Wiskott-Aldrich Syndrome Protein
WHO World Health Organisation
WP Whiteman Park
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List of Figures and Tables
Figures Description Page
1.1 Tree illustrating genetic relationships between the different 7 rickettsial/rickettsia-like organisms found in three different Proteobacteria divisions.
1.2 Illustrations showing location of SFG and TG rickettsiae in the 10 host cell.
1.3 Figure illustrating the methods used in the detection and diagnosis 25 of rickettsiosis in human/animal/arthropod samples. Adapted from Fenollar et al.
1.4 Figure showing the amplification and sequence similarities 29 requirement of 5 rickettsial genes to determine group/ species/novelty of a Rickettsia isolate.
1.5 Geographic distribution of known rickettsial organisms in 48 Australia.
2.1 Morphological characteristics of an Ixodidae tick. 53
2.2 Dichotomous key for the identification of Australian Ixodidae 54 genera.
3.1 Unrooted neighbour-joining tree based on ompA sequences from 62 Rickettsia species with validly published names.
3.2 Illustration of plaque assay plate at dilution 10-1. Similar plates 64 were prepared for each ten-fold dilution up to 10-6.
3.3 Figure showing serum sample dilutions and control locations. 67 Samples were tested in duplicate with two-fold serum dilutions starting from 1/8 to1/4096.
3.4 Formula for calculation of Specificity Difference (SPD). 68
3.5 Relationship between bovine serum albumin (BSA) concentration 70 and absorbance at 750nm.
4.1 Pictures of ticks on various mammals. 79
4.2 Figure showing locations of tick sampling sites. 80
4.3 Figure showing location and direction of cut using a scalpel blade 81 along the circumference of the tick scutum and body.
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Page
4.4 Figure showing the location of the three main organs removed and 81 analysed for rickettsial DNA.
5.1 Three main trapping sites named after the general region they 96 encompass (i) Mundaring, (ii) Serpentine and (iii) Dwellingup.
5.2 Feral pig live trapping using wire cages with apples as bait (A). 97 Multiple pigs were often trapped at the same time (B). Post mortem and ectoparasite sampling performed on site after euthanasia of feral pigs at trap sites (C).
6.1 Eschar and swelling on patient’s leg as a result of a tick bite. 119
7.1 The transmission cycle of Rickettsia gravesii in Western 134 Australia.
7.2 Maps of Western Australia showing (A) comparative size of WA 137 to other nations or states of the world and (B) population distribution based on data from the Australian Bureau of Statistics.
Tables Description Page
1.1 Table summarising rickettsial organisms of the genus Rickettsia 9 whose genomes have been fully sequenced.
1.2 Known pathogenic tick borne spotted fever group rickettsiae. 11
1.3 Invertebrate hosts of Rickettsia felis, some of which carry other 44 potential bacterial pathogens.
2.1 Summary of PCR conditions for Rickettsial DNA amplification. 60
3.1 Mouse groups and rickettsial inoculum size (original and booster) 65 given via the intra-peritoneal route.
3.2 Table of conversion from geometric to arithmetic antibody titres 68 for rickettsial antigen-antibody reactions.
3.3 Successful plaque formations in two of five attempts and the 69 concentration in pfu/mL.
3.4 Summary of absorbance values of rickettsial inocula used and 70 their respective protein antigen concentrations.
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Page
3.5 Microimmunofluorescence antibody titres against rickettsial 71 antigens with arithmetic numerical values in parentheses.
3.6 Table of SPD values for homologous and heterologous 71 antibody/antigen interactions showing level of antigenic difference between various rickettsiae obtained by MIF (SPD ≥ 3 indicates distinct serotypical difference).
3.7 ELISA antibody titres against rickettsial antigens with arithmetic 72 numerical values in parentheses.
3.8 Table of SPD values for homologous and heterologous 72 antibody/antigen interactions showing level of antigenic difference between various rickettsiae obtained by ELISA (SPD ≥ 3 indicates distinct serotype).
4.1 Tick species collected by animal host and their respective habitat. 85
4.2 Prevalence of R. gravesii in ticks by area/region. 86
4.3 Western Australian recorded tick species in the literature and 92 those with potential as vectors of R. gravesii.
5.1 Four animal species considered for study and their fulfilment of 94 five criteria.
5.2 Rickettsial sero-prevalence in feral and domestic pigs against SFG 98 and TG antigens.
5.3 Titration of feral pig sera against Rickettsia antigens. 99
5.4 Seroprevalence in feral pigs by age and location. 99
5.5 Summary of four feral pigs positive by rickettsial citrate synthase 100 gltA gene PCR.
6.1 Seroprevalence for SFG rickettsia from primary and secondary 110 blood sampling of humans residing in WA with MIF titres to SFG of 1:128.
6.2 Rogainers’ seroconversion and seroreversion results over a 10-14 112 month period.
6.3 Whiteman park employees’ seroconversion and seroreversion 112 results over a 10-14 month period.
6.4 Descriptive categories of populations based on questionnaire data. 114
6.5 Risk estimates for seropositivity to SFG Rickettsia sp. 117
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1 - Introduction
1.1 Thesis Introduction
It has been barely 130 years since arthropods were shown to transmit human diseases and knowledge of zoonotic diseases now encompasses hundreds of viruses, bacteria, protozoa and helminthes. Vector borne diseases were responsible for more human disease and deaths in the 17th to early 20th centuries than all other causes combined.
Only after effective preventive and control measures against known disease vectors were developed did the industrial development of the tropics during the 19th and 20th centuries take place.
With the start of the 21st century, there has been ongoing discovery of new species and strains of potentially pathogenic microbes. Most have turned out to be harmless but there are those that have caused severe mortality and morbidity on human and animal populations, such as the Nipah virus outbreaks in Malaysia, Bangladesh and India between 2001 and 2008 and the SARS epidemic of 2003. The description of any new taxa requires extensive efforts on the part of the investigator(s) to fully differentiate a
“new” organism from its most closely related phenotypic and genotypic relatives. Not only is there a need to catalogue any differences or unique features, but also the similarities that the new taxa share with other similar organisms. The wealth of accumulated information and knowledge of new aetiologic agents is essential to assist medical, veterinary and public health professionals in future encounters.
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The rate of new rickettsiae being “discovered” has increased exponentially over the years especially with the advent of molecular methods towards the end of the 20th century. The discovery of a new species of Rickettsia by Helen Owen in 2005, named
Rickettsia gravesii (strain BWI-1), left many questions unanswered (206-208). The main aim of this thesis will be to reduce the gaps in our knowledge of Western
Australian rickettsial organisms and in particular their role as potential animal and/or human pathogens in Western Australia. An extensive review of Rickettsia sp. is crucial to provide a basic understanding of rickettsial organisms. Factors that make up their genotypic and phenotypic characteristics will be included, along with host-rickettsiae interactions, pathogenicity, disease diagnosis and distribution down to the species level.
The design of the study aims and objectives was heavily influenced by the information documented by Helen Owen in her thesis (208). Continuity was the direction decided upon and of the two novel rickettsiae described by Owen (R. gravesii and R. antechini);
R. gravesii was chosen as the focus of this work due to its hypothesized “abundance” in south-west WA native fauna and arthropod populations and suspected higher pathogenicity potential. No “Candidatus R. antechini” was observed during the course of the current study.
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1.2 Study Aims and Objectives
The aim of the study was to address the following objectives derived from the literature on Western Australian rickettsiae.
I. To characterise phenotypic traits of R. gravesii that are distinct from other
rickettsiae.
II. To examine the arthropod host species known to harbour R. gravesii.
III. To determine the sero-prevalence of R. gravesii in a wild animal population.
IV. To identify the regions within south-west WA endemic for R. gravesii.
V. To investigate the relationship between sero-prevalence to spotted fever
rickettsia in humans and their level of bush activity.
VI. To report and characterise any illness linked to Rickettsia sp. infection during
the period of the study.
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1.3 Rickettsia
Zoonotic rickettsial infections occur in both urban and rural communities worldwide.
Transmitted via vectors such as ticks, fleas and lice, rickettsial infections are often viewed as mild or are misdiagnosed as common ailments such as the common cold and flu (322). Through lack of understanding of the actual cause of disease, complications leading to organ failure and possibly even death can occur.
The genus Rickettsia was first described by Howard Ricketts when he investigated an outbreak of Rocky Mountain spotted fever in 1906 and 1907 (257). His work was instrumental in paving the way for modern rickettsiology (258). Most significant were the roles that ectoparasites were found to play as vectors of rickettsial infection and reservoirs of rickettsial organisms in an endemic area. Today we know that the rickettsial life cycle involves both vertebrate and invertebrate hosts. Not only do haematophagous arthropods play an important role as vectors, they are also a primary reservoir and amplifying host, together with some small mammals such as rats and opossums (16).
At present, three groups of organisms are commonly classified as causing rickettsial disease, (i) spotted fever group (SFG), (ii) typhus group (TG), and (iii) scrub typhus group (STG) (69, 324). The first two groups belong to the genus Rickettsia, while the third group belongs to the genus Orientia. A number of major changes to the taxonomy of rickettsia have been made in the past decade, as a result of the advent of molecular taxonomic methods such as 16S rRNA analysis (334). These changes have resulted in the exclusion of a myriad of other organisms from being classified as rickettsiae, although they continue to be described as rickettsia-like, such as Coxiella burnetii and
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Bartonella. These new techniques also enabled scientists to study intracellular bacteria that express few of the phenotypic characteristics traditionally used in taxonomy.
Recently Fournier et al. proposed genetic guidelines for the classification of rickettsial isolates at the genus, group, and species levels by using sequences of the 16S rRNA
(rrs) gene and four protein-coding genes; gltA (citrate synthase), ompA, ompB and sca4
(gene D) (88).
The Rickettsia genus currently includes more than 24 recognized species along with a substantial number of as-yet uncharacterized strains and amplicons. Of the known rickettsioses, 16 of these species have been implicated in human syndromes. However it is hypothesised that any rickettsiae found in arthropods capable of biting humans should be considered potent to be potential human pathogens (93).
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1.3.1 Taxonomy
The genus Rickettsia is split into two major groups, Spotted Fever Group (SFG) and
Typhus Group (TG) (221). Rickettsial organisms within the two groups are small (ca.
0.4 x 0.9 µm), Gram-negative, coccobacillary, aerobic α-Proteobacteria (268). They are classified as either SFG or TG rickettsiae based on their intracellular positions, optimal growth temperatures, percent G+C DNA contents, clinical features, epidemiological aspects and antigenic characteristics (absence of the outer membrane protein in TG) (244). Two other groups have recently been proposed; Ancestral Group
(AG) and Transitional Group (TRG) (109, 295).
Before the use of 16S rRNA sequence to determine the phylogenetic relationship between prokaryotes in the order Rickettsiales, a rigorous set of criteria for classical taxonomy was used. The evolutionary tree that the Rickettsia spp. belonged to was widely accepted, as described by Bergey’s Manual of Systematic Bacteriology (1939), showing the genus Rickettsia in the same tribe, Rickettsieae, as Coxiella and
Rochalimea. Rickettsieae was classified within the family Rickettsiaceae, together with
Wolbachieae and Ehrlichieae. The families Bartonellaceae and Anaplasmataceae were also grouped within the order Rickettsiales. With the advent of new molecular taxonomic methods, the classical taxonomy used was revised and the rickettsiae are now placed into subgroups of Proteobacteria. The genus Rickettsia has been placed in the α-1 subgroup, while Bartonella is in the α-2 subgroup (Figure 1.1).
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Orientia tsutsugamushi (Scrub Typhus)
Rickettsia sp (TG)
Rickettsia sp (SFG) rickettsii
1 Neorickettsia sennetsu
Neorickettsia helminthica
Wolbachia pipientis
Anaplasma phagocytophilum
Anaplasma marginale
Ehrlichia chaffeensis Ehrlichia canis Ehrlichia ruminantium
Bartonella quintana
Bartonella bacilliformis
2 Brucella melitensis
Coxiella burnetii
Francisella persica
Figure 1.1 – Tree illustrating genetic relationships between the different rickettsial/rickettsia-like organisms found in three different Proteobacteria divisions (69).
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The original classification of species within the genus Rickettsia into the SFG and TG groups depended on a variety of characteristics including: intracellular localization, optimal growth temperature and the cross reaction of sera from an infected patient with somatic antigens of three strains of Proteus (Weil-Felix test) (123). The advent of modern molecular and serological techniques both reinforced and disputed original classification of members of the genus and as such the last couple of decades have seen the restructuring of Rickettsia sp. Genomic analyses have now proposed the creation of
2 new groups within the genus Rickettsia; Ancestral Group (AG) consisting of R. bellii and R. canadensis (295) and Transitional Group (TRG) consisting of R. akari and R. felis (109).
Other notable rickettsiae that have been reclassified are Rickettsia tsutsugamushi; which has been placed in its own genus, Orientia (Orientia tsutsugamushi), and is in the same α-1 subgroup as the genus Rickettsia (Figure 1.1) (296). Coxiella burnetii
(originally Rickettsia burnetii) has also been reclassified due to the findings that its 16S
RNA sequence is more similar to members of the γ subgroup and it is no longer viewed as a true rickettsia (185).
1.3.2 Genome
Rickettsial genome sizes are comparatively much smaller than those of other bacteria.
The circular genome ranges in size from 1.1 to 1.6 million base pairs (Table 1.1).
Coding regions range from 75.2% (R. canadensis) to 85.2% (R. belii). Genomic analysis of the eukaryotic mitochondrial genome suggests a close relationship between rickettsiae and mitochondria (338). Based on phylogenetic analysis, mitochondria have been shown to have evolved from the α-proteobacteria and more specifically from
Rickettsiaceae (6, 283).
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R. felis was the first obligate intracellular bacteria reported to have potential conjugative plasmids as part of its genome. The two R. felis plasmids identified were named pRF and pRFδ, and are 62,829 bp and 39,263 bp in size respectively. The size of the main circular chromosome is 1,485,148 bp (197).
Table 1.1 – Table summarising rickettsial organisms of the genus Rickettsia whose genomes have been fully sequenced.
Genome No. of % % Species Host Group Size (bp) Genes G+C Coding
R. belii Tick 1,522,076 1,429 31.7 85.2 Ancestral R. canadensis Tick 1,159,772 1,032 29.0 75.2 Group
R. prowazekii Louse/Flea 1,111,523 872 28.9 76.2 Typhus Group R. typhi Flea 1,111,496 877 29.0 76.3
R. akari Mite 1,231,204 1,337 32.3 77.4 Transitional R. felis Flea 1,587,240* 1,512 32.5 83.6 Group
R. rickettsii Tick 1,257,710 1,346 32.5 78.5 Spotted R. conorii Tick 1,268,755 1,412 32.4 81.5 Fever Group R. sibirica Tick 1,250,021 1,234 32.5 77.8 * Includes plasmid(s).
Since the first description of plasmids in R. felis, other members of the groups TRG,
SFG and AG have been published on with harbouring plasmids (19, 20). With the discovery of rickettsial plasmids, it is now better understood how conjugative genes, widespread in the genus, are horizontally transferred (327). The presence of widespread and potentially mobile plasmids in Rickettsia has epidemiologic and evolutionary implications.
9
1.3.3 Spotted Fever Group (SFG) Rickettsiae
The spotted fever group of the genus Rickettsia is of major clinical significance to humans. There are currently 20 recognized species/strains (excluding R. felis and R. akari) of which 12 are recognized as the cause of rickettsioses (Table 1.2). With the recent proposal for a new group for R. felis (flea-borne) and R. akari (mite-borne), the spotted fever group would be exclusively tick-borne (109).
The members of SFG exist in free form in the cytoplasm of their mammalian and arthropod host cells (Figure 1.2). Unlike TG rickettsiae, SFG rickettsiae can pass through the nuclear membrane of their host cell. Rickettsiae have a trilaminar cell wall with a periplasmic space separating the cell wall from the cytoplasm (121). Species- specific antigens are located on a microcapsular layer on the outer leaflet of the cell wall.
TG Rickettsia SFG Rickettsia
Figure 1.2 – Illustrations showing location of SFG and TG rickettsiae in the host cell.
10
Table 1.2 – Known pathogenic tick borne spotted fever group rickettsiae.
Year of first Disease [Year of Known or potential Rickettsia spp identification first clinical Ref vectors in vectors description]
Confirmed Pathogens
Rickettsia rickettsii Dermacentor andersoni 1906 Rocky Mountain (257) Dermacentor variabilis spotted fever Rhipecephalus sanguineus [1899] Amblyomma cajennense Amblyomma aureolatum
Rickettsia conorii Rhipicephalus sanguineus 1932 Mediterranean (171) subsp. conorii spotted fever [1910]
Rickettsia sibirica Dermacentor nuttali Unknown Siberian tick (266) subsp. sibirica Dermacentor marginatus typhus [1934] Dermacentor silvarum Haemaphysalis concinna
Rickettsia parkeri Amblyomma maculatum 1939 Unnamed [2004] (212) Amblyomma americanum Amblyomma triste
Rickettsia conorii Rhipicephalus sanguineus 1950 Indian tick typhus (271) subsp. indica
Rickettsia slovaca Dermacentor marginatus 1968 Tick-borne (280) Dermacentor reticulatus lymphadenopathy [1997] Dermacentor- borne necrosis and lymphadenopathy [1997]
Rickettsia conorii Rhipicephalus sanguineus 1974 Israeli spotted (110, subsp. israelensis fever [1940] 310)
Rickettsia australis Ixodes holocyclus 1974 Queensland tick (45) Ixodes tasmani typhus [1946]
Rickettsia sibirica Dermacentor sinicus 1974 North Asian tick (318) subsp. mongolotimonae typhus [1960]
Hyalomma asiaticum 1991 Lymphangitis (339) Hyalomma truncatum associated rickettsiosis [1996]
Rickettsia heilongjiangensis Dermacentor silvarum 1982 Far Eastern (88) spotted fever [1992]
11
Table 1.2 – Continued.
Year of first Disease [Year of Known or potential Rickettsia spp identification first clinical Ref vectors in ticks description]
Rickettsia africae Amblyomma hebraeum 1990 African tick bite (144, Amblyomma variegatum fever [1934] 156)
Rickettsia conorii Rhipicephalus sanguineus 1992 Astrakhan fever (74, subsp. caspia Rhipicephalus pumilio [1970s] 297)
Rickettsia massiliae Rhipicephalus sanguineus 1992 Unnamed [2005] (23, Rhipicephalus turanicus 315) Rhipicephalus muhsamae Rhipicephalus lunulatus Rhipicephalus sulcatus
Rickettsia honei Bothriocroton (Aponomma) 1993 Flinders Island (115, hydrosauri spotted fever 264) Amblyomma cajennense [1991] Ixodes granulatus
Rickettsia japonica Ixodes ovatus 1996 Oriental or (302) Dermacentor taiwanensis Japanese spotted Haemaphysalis longicornis fever [1984] Haemaphysalis flava
Rickettsia aeschlimannii Hyalomma marginatum 1997 Unnamed [2002] (25) marginatum Hyalomma marginatum rufipes Rhipicephalus appendiculatus
Rickettsia honei Haemaphysalis novaeguineae 2005 Australian (306) strain marmionii spotted fever [2005]
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1.3.4 Typhus Group (TG) Rickettsiae
The causes of epidemic typhus and murine typhus are Rickettsia prowazekii and
Rickettsia typhi respectively. Only these two organisms have been classified within the typhus group thus far (188). Occurrences of the disease are worldwide and transmissions of the pathogens are by the human body louse (Pediculus humanus humanus) for R. prowazekii (80) and rat flea (Xenopsylla cheopis) for R. typhi (13).
Recent publications have shown that the vector for R. typhi may not be limited to just the rat flea but may include other species of fleas such as the cat flea Ctenocephalides felis (194).
1.3.5 Ancestral Group (AG) Rickettsiae
Rickettsia bellii was demonstrated to share common characteristics with both SFG and
TG rickettsiae by Stothard et al. (1994) but also to exhibit unique characteristics of its own. Comparison of the 23S rRNA sequences between R. bellii, R. rickettsii and R. prowazekii indicates that R. bellii predates the divergence that created the SFG and TG genera (295).
1.3.6 Transitional Group (TRG) Rickettsiae
The transitional group was recently proposed by Gillespie et al. (2007) and is made up of R. akari and R. felis. Both rickettsial species are described as being genotypically and phenotypically divergent enough from SFG and TG rickettsiae, to warrant their own grouping (109). Genotypic analysis supports this hypothesis, showing that genes found in members of TRG were possibly acquired through conjugational exchanges with AG rickettsiae (109). R. australis was recently described phylogenetically as a member of TRG (316), however its LPS is antigenically similar to SFG.
13
1.4 Rickettsial Vectors
Arthropods are the only known vectors of rickettsia. They are also reservoirs and amplifiers for rickettsial spread throughout invertebrate and vertebrate populations.
Maintenance of rickettsiae in an arthropod population occurs through transtadial and transovarial means (16). Vertical transmission plays an important survival role when the vector population is low (190).
1.4.1 Ticks
Ticks are obligate haematophagous arthropods that parasitise all classes of vertebrates and have a worldwide distribution. Recognized as human parasites for thousands of years, ticks have been described by ancient writers such as Aristotle and Homer.
However, it was not known that ticks were capable of disease transmission until the end of the 19th century. Ticks are currently considered the second most common vector of human infectious disease after the mosquito. The first demonstration of the potential of ticks as a disease vector (Texas cattle fever) was performed in 1893 (11) through the transmission of the protozoan Babesia bigemina by the tick Boophilus annulatus. It was not until the start of the 20th century that ticks were associated with the transmission of bacterial diseases. Dutton and Todd demonstrated in 1905 (72) that tick relapsing fever was caused by Borrelia duttonii and transmitted by Ornithodoros moubata, and
Ricketts followed closely in 1909 showing that the wood tick Dermacentor andersoni, was involved in the transmission of Rickettsia rickettsii, the agent of Rocky Mountain
Spotted Fever (RMSF) (257). Other common tick-borne bacterial diseases include rickettsioses, ehrlichioses, anaplasmosis, Lyme disease, some relapsing fever borrelioses (others are louse-borne), tularaemia and Q fever (78).
14
There are 2 major tick families; Ixodidae and Argasidae. Ixodidae, commonly known as hard ticks is the most abundant and medically important of the two tick families.
Ixodid ticks have a sclerotized dorsal plate. Argasidae or soft ticks are named because of their flexible cuticle. A third much less common group known as Nuttalliellidae comprises a single species that has been collected in Africa, albeit rarely (154). The main tick vectors for rickettsiae are ixodid ticks. Argasid ticks have previously only been implicated with the non-pathogenic rickettsia R. bellii. Recent work had uncovered the presence of an SFG rickettsiae in soft ticks in Japan (153) and Rickettsia hoogstraalii in the soft tick Cario carpensis in the US (66).
Since the first description of rickettsia by Howard Ricketts, a considerable amount of information on tick-borne rickettsiae has been uncovered via extensive studies of the ticks Dermacentor andersoni and Dermacentor variabilis, the vectors for R. rickettsii
(Rocky Mountain spotted fever) (257, 288, 335). Ticks not only serve as vectors for R. rickettsii, they also act as reservoirs. Maintenance of R. rickettsii in a tick population occurs by both transtadial, transovarial and horizontal transmission (219, 280).
Transtadial transmission within a tick refers to the maintenance of viable rickettsiae from one stage of a tick lifecycle to the next without affecting the viability of the tick or rickettsial organisms. Rickettsia rickettsii can be acquired by non-infected ticks from rickettsaemic animal hosts. The animal hosts that can attain a sufficient level of rickettsaemia and further transmit the bacteria are limited to certain species (37).
Sexual transmission of rickettsiae by ticks during mating have been observed however females infected by sexual transmission were unable to transmit the rickettsiae to their progeny (122).
15
Findings from the early studies of R. rickettsii transmission were also observed in other rickettsiae, such as R. slovaca (65). However variations do exist with some other tick- borne rickettsiae. For example, the inability of R. rhipicephali to be transmitted transovarially (177), which was not observed beyond the first generation.
1.4.2 Fleas
Fleas have a worldwide distribution, are found principally on mammals, and act as important vectors for zoonotic diseases including Yersinia pestis the causative agent of plague, Bartonella quintana the causative agent of bacillary angiomatosis and
Bartonella henselae the causative agent of cat scratch disease (15). Only two members of the genus Rickettsia are known to be transmitted by fleas. Rickettsia typhi of TG and
R. felis of TRG. The transmission of R. typhi by the flea species Xenopsylla cheopis (rat flea) was first described by Maxcy (183). Since then, R. typhi has been recovered worldwide in areas with rat infestation and on other flea species such as the cat flea
Ctenocephalides felis and the mouse flea Leptopsyllia segnis. Transmission of R. felis to humans has been attributed via exposure to flea saliva, differing from R. typhi which has been shown to be transmitted to humans via entry of flea faeces into the flea-bite site or skin abrasions, inhalation or the contamination of the conjunctiva (15).
16
1.4.3 Lice
Lice are well recorded as parasites and are host specific, for example - the human body lice is specific only to humans. Infestations of humans are known as pediculosis. The human body louse (Pediculus humanus humanus and Pediculus humanus corporis) is also recognised as a vector of at least three diseases; epidemic typhus (Rickettsia prowazekii), louse-borne relapsing fever (Borrelia recurrentis) and trench fever
(Bartonella quintana) (245). Various outbreaks and epidemics of the three diseases carried by the human body louse are testament to its close co-evolution with humans.
Some of the most famous outbreaks attributed to human body lice as a vector for R. prowazekii include the Napoleonic Wars where 20% of Napoleon’s troops died of typhus and the 12 million infected Russians between 1917 and 1925 of whom 3 million died (222, 251).
The human body louse does not act as a reservoir for R. prowazekii and becomes infected as a result of feeding on a bacteraemic host. Infected lice usually die within 7 days as a result of damaged epithelial cells caused by R. prowazekii (133). This lethal characteristic of R. prowazekii, for its vector, makes it the only member of the genus
Rickettsia not transmitted transovarially in its vector population. In spite of this limitation, epidemic typhus continues to be reported at the start of the 21st century, especially in the homeless and low-hygiene populations of developed countries. Some factors contributing to the re-emerging threat of epidemic typhus are as a result of increasing incidences of human body louse infestations due to declining social structures and hygiene practices as a result of wars, economic instability and natural catastrophes (41).
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1.4.4 Mites
Only one member of the genus Rickettsia has been described as being transmitted by mites; Rickettsia akari by the mouse mite Liponyssoides sanguineus (211). However R. akari has also been isolated from other rodents in the U.S.A, Eurasia and Africa including the brown rat, gray dwarf hamster, golden spiny mouse, large-eared dormouse and bushy-tailed jird (213). Suggestions of L. sanguineus having a larger host range, including larger vertebrate hosts have been supported with the discovery of antibodies against R. akari in dogs and cats in the U.S.A (54).
Rickettsia akari can be maintained within L. sanguineus populations by transovarial transmission. L. sanguineus mites have five life stages; egg, larva, protonymph, deutonymph and adult (97). Each stage requires a blood meal for the mite to moult and progress to the next stage. With the multiple blood meals within its life cycle, the mite has the opportunity to pass on R. akari to infected vertebrate hosts several times.
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1.5 Rickettsia Pathogenesis
Rickettsial disease or rickettsioses are caused by known pathogenic rickettsiae from the
SFG, TG and TRG. Although all differ in transmission, rickettsiae-host interactions and pathology, much of the pathogenesis of the three groups is similar.
1.5.1 Transmission
All pathogenic rickettsiae require arthropods for transmission; however how the rickettsia is transmitted from the arthropod to humans can differ. Tick and mite borne rickettsiae rely on the transfer of rickettsiae during feeding. Rickettsiae are transferred to the host via the tick or mite saliva at the bite site. Cases of transmission of rickettsiae via the conjunctiva have occurred with exposure of infected tick haemolymph on fingers after crushing an infected tick. Flea and louse borne rickettsiae are transmitted via faeces that enter the host via bite site or cuts on the skin. Inoculation via inhalation is possible and has happened in the laboratory (323). Although target cells at the site of inoculation are currently unknown, rickettsiae possibly infect dermal cells such as vascular endothelium (main cell infected), fibroblasts, lymphatic endothelium and macrophages (321).
1.5.2 Rickettsiae-Host Interactions
Recently with the advent of new culture techniques and imagery technology, a better understanding of the interactions between the rickettsiae and their host has been gained.
Severity of rickettsial disease can be attributed to patient age as the main consistent factor in almost all cases. Initial hypothesis for rickettsioses having a higher fatality in males (esp. for Rocky Mountain spotted fever) has been disproved by recent studies
(320). Other factors affecting rickettsiae-host interaction would be underlying patient 19 diseases and enhanced oxidative stress increasing the severity of illness. Mediterranean spotted fever shows greater virulence in patients with diabetes mellitus (323).
1.5.2.1 Routes of Spread
Rickettsiae appear to target any nucleated cell at the site of inoculation. Cells possibly targeted would include vascular and lymphatic endothelium, dermal dendritic cells, fibroblasts and macrophages (323). Once rickettsiae have penetrated and initiated infection, spread throughout the host is facilitated via the lymphatic and vascular systems. The presence of lymphadenopathy in the same region as the site of inoculation supports the hypothesis of antigenic spread via the lymphatic system. The spread of rickettsiae into susceptible cells is believed to occur mainly by vascular location though the action of rickettsial adhesions acting on specific receptors on host cells do play a significant role (181, 323).
Entry of rickettsiae into a vertebrate or arthropod host cell is via the binding of rickettsial ligands (outer membrane proteins A and B) with host cell protein Ku70 (173,
181, 304). Adhesions of outer membrane protein B (OmpB) with Ku70 result in a cascade reaction which involves the migration of more Ku70 and ubiquitin ligase to the binding site, resulting in the focal-induced endocytosis of the adherent rickettsia into the host cell. Sca1 and Sca2 of the SFG had recently been reported to promote adhesion to mammalian cells (46, 260). The protein complex Arp2/3 has been associated with changes induced in the cytoskeletal actin at the entry site. Proteins involved in the activation of the Arp2/3 complex include GTPase, Cdc 42, protein tyrosine kinase, phosphoinositide 3-kinase and Src-family kinase (180). Once inside the cell, rickettsiae exit the phagosome into the cytosol by digesting the phagosomal membrane via the action of the enzymes phospholipase D and hemolysin C (TlyC) (256, 331). 20
Movement of some rickettsiae from one host cell to another is facilitated by actin polymerization, shown to be present in all spotted fever group rickettsiae except R. peacockii. Activation of the actin polymerization occurs at one pole of the bacterium, and the continuous conversion of globular actin to filamentous actin propels the rickettsia through the cytosol eventually hitting the host cell membrane (124, 298).
Upon collision with the cell membrane there are two possible outcomes; (i) bounce off the inner surface of the cell membrane, or (ii) deformation of the membrane and formation of a filopodium. Should the rickettsia exit the cell in a filopodium, a couple of different possibilities could occur depending on what is on the other side of the cell membrane; rickettsiae enter adjacent cells, or exit as a luminal filopodium entering the blood stream (125). The actin-based mobility seen in SFG rickettsiae has been shown to be initiated by the rickettsial protein RickA and mediated by Sca2 (120, 161). RickA has been described as a WASP-family (Wiskott-Aldrich Syndrome Protein) like protein with similarities such as carboxy terminal domains and activation of the Arp2/3 complex (111, 142). The gene rickA is however absent in both R. prowazekii and R. typhi. This is consistent with the lack of actin polymerization in TG rickettsiae and the burst release model. Erratic actin based mobility has been observed in R. typhi suggesting that members of the genus use multiple redundant mechanisms for actin polymerization which has yet to be clarified (142).
1.5.2.2 Immune Response and Clearance
Innate and adaptive immune responses to rickettsial infection have been a focus of research recently to better understand the rickettsia-host interactions during infection and clearance from the host. In spite of the wealth of information reported, there is still much that is not fully understood. In vitro studies have identified a few critical 21 components making up the host response; gamma interferon (IFN-γ), tumor necrosis factor alpha (TNF-α) and CD8 T cells (81, 82).
Host innate immune response at the onset of rickettsial infection has been observed with the activation of local natural killer (NK) cells. Production of IFN-γ has been associated with the NK cell activation. Type I interferon production has also been found to be stimulated, however its exact role in response to infection is believed to be indirect (30). Effects of type I interferon (IFN-α and IFN-β) production include NK cell activation, maturation of dendritic cells and production of interleukin-12 (IL-12). In vitro infection of human endothelial cells with R. conorii resulted in the production of
IL-1, IL-6, IL-8 and monocyte chemotactic protein-1 (MCP-1) while infected macrophages were shown to secrete TNF-α (150). General vasculitis as a result of R. conorii infection (MSF) could be attributed to the secretion of the above cytokines, which are pro-inflammatory (150, 176, 255).
1.5.3 Pathology
Increased vascular permeability as a result of Rickettsia infection have been associated with bacterial load and TNF-α causing disrupted endothelial cell junctions (309, 336).
Perivascular T lymphocytes and macrophages are heavily involved in the clearance of rickettsial infections. As most rickettsial infection is localized and not widespread, the result of vasculitis is not usually severe except in the most serious of cases.
Perivascular lymphohistiocytic infiltrate has been observed in the organs affected by rickettsial infection including the brain, lung, heart, kidney, skin, gastrointestinal tract, pancreas, gall bladder, skeletal muscle and testes (319).
22
Destruction of endothelial cells as a result of multifocal rickettsial infection usually results in the impairment of normal anticoagulation functions. The occurrence of fibrin- platelet thrombi in both living and dead patients is rare (58). Haemostatic plugs are only observed in the foci of severe rickettsial inflicted vascular injury. In spite of the general hypothesis that thrombosis is the cause of organ infarction and tissue damage among patients, pathological and experimental evidence proves otherwise. Although infarction of organs seldom occurs, it has been observed in the white matter of the brain. Tissue damage to kidneys is usually the result of poor perfusion. Gangrene and cutaneous necrosis have also been observed in some cases of severe rickettsioses resulting in extensive damage to the vascular microcirculation of the affected area
(317).
23
1.6 Diagnosis and Treatment
Diagnosis of rickettsial infections can be hard without appropriate diagnostic assays.
Prior to the advent of modern diagnostic methods, identifying rickettsial infections and prompt treatment relied heavily on the physician’s knowledge of endemic rickettsial diseases in their locality. A classic indicator is the presence of an attached arthropod or inflamed arthropod bite on the patient. But the development of symptoms usually occurs about a week following infection, which may result in the unspecific symptoms being misdiagnosed as another ailment. The two classic symptoms that usually differentiate rickettsioses from other ailments would be the presence of an eschar at the arthropod bite site and a rash. Other major symptoms such as myalgia, fever, headaches, and local lymphadenopathy could be attributed to a myriad of other diseases
(40). There have been reports of “spotless” spotted fever and “escharless” rickettsioses, major symptoms are similar for most rickettsial diseases, variations that do occur can vary greatly from classic symptoms. Thus there is a great need for prompt and accurate laboratory tests to assist physicians with diagnosis of rickettsial disease.
Confirmation of a rickettsial disease is achieved via one of three methods. Eschars and blood samples are screened by serology and PCR or culture respectively (Figure 1.3).
Although seldom performed, the biting arthropod can be screened directly by PCR and/or culture for evidence of rickettsiae.
24
Patient
Blood Sampling Skin biopsy Arthropod
Filter paper Serum Anticoagulant
EDTA Citrate
Serology
Cell Culture
IFA or Gimenez Staining
Polymerase Chain Reaction (PCR)
Figure 1.3 – Figure illustrating the methods used in the detection and diagnosis of rickettsiosis in human/animal/arthropod samples. Adapted from Fenollar et al. (83).
25
1.6.1 Serological Methods
One of the earliest tests developed to serologically diagnose rickettsial infection was the Weil-Felix test (326). The Weil-Felix test was originally designed to detect antibodies to Proteus antigens based on the principle that they cross-reacts with rickettsial antigens. Whole cells of Proteus vulgaris OX-2 react strongly with SFG positive sera except for sera from RMSF patients (R. rickettsii). Typhus Group and
RMSF positive sera react strongly with whole cell P. vulgaris OX-19. Whole cell preparations of Proteus mirabilis OX-K were later found to be reactive with antibodies from scrub typhus patients. The Weil-Felix test works on the principle of agglutination to detect antibodies in sera 5-10 days after the onset of symptoms. It is however not applicable to patients suffering from Brill-Zinsser disease or infected with R. akari due to the inability of the antibodies to agglutinate with the Proteus whole cell preparation used. The antibody type detected in the test is mainly IgM (3). The Weil-Felix test is now considered to be obsolete with the development of rickettsial IF techniques. The poor specificity and sensitivity of the Weil-Felix test is well documented and it would only be used now in rudimentary clinical laboratories (44, 123, 201, 240). The Weil-
Felix method has also been shown to have cross-reactivity when Rickettsia positive sera are tested against Legionella sp. and Proteus sp. (242).
Immunofluorescence Assay
Philip, 1978, suggested the use of immunofluorescence assays (IFA) for the detection of rickettsiae using serology as a replacement to the Weil-Felix method (227). IFA serology has now become the most widely accepted method of detection and is used frequently in providing rapid diagnosis to suspected rickettsial infections (165).
26
IFA is highly sensitive but it faces the problem of the lack of intra-rickettsial group specificity. Diagnosis of rickettsiosis by IFA is fast and accurate but differentiation between species of rickettsiae as the cause of infection is often impossible due to cross reacting antibodies. This is further complicated by the variability in antibody titre determination due to operator subjectivity. A method that has been suggested by the
Australian Rickettsial Reference Laboratory is to increase the cut-off value of antibody titre to 1:128 thus reducing the incidence of false positives (307). Current cut off values used by the Unité des Rickettsies are 1:64 for IgG and 1:32 for IgM (87). In spite of its short-comings, IFA is usually used as the only tool of diagnosis for rickettsial disease.
1.6.2 Cell Culture
The ability to isolate a microbial pathogen in culture is definitive in disease diagnosis and crucial in characterizing the microbes’ genotypic and phenotypic features. In fact, being able to culture a disease causing microorganism forms the second and third criteria of Koch’s postulates (135). With the advent of modern molecular biological techniques in the last couple of decades, the presence of a pathogen’s genetic material in a host’s tissue sample would be enough for a diagnosis. However, molecular techniques alone are currently unable to provide crucial information obtainable only via culture such as pathogen-host cell interactions.
The shell vial technique is one of the most effective methods of isolating rickettsial organisms from extracts of blood, animal and human tissue (313). It was originally adapted from the a similarly named technique used in isolating viruses and is now one of the most effective methods available to researchers for isolating Rickettsia sp. and other rickettsia-like organisms (238, 241). Even so, conventional cell culture using a confluent monolayer of either mammalian (L929, Vero and HEL), amphibian (XTC-2)
27 or insect (DAE100, Aa23 and Sua5B) cell lines acting as host to the rickettsial organism is still popular and well accepted (189, 276).
1.6.3 Polymerase Chain Reaction (PCR)
The application of PCR has revolutionised the detection and diagnosis of rickettsial infections (5). With access to a proper laboratory equipped to perform PCRs on patient samples, accurate diagnosis can be obtained before seroconversion of patient and prior to growth in culture. Samples used for PCR are preferably fresh but tissues that have been preserved in paraffin or fixed to a slide can also be used (290). Various assays have been developed allowing for rapid detection of Rickettsia sp. with most targeting well-described genes such as citrate synthase (gltA) (269), OmpA (ompA) (85), OmpB
(ompB) (270), 16S rRNA (268) and geneD (sca4) (279). The implementation of real- time PCR (qPCR) in rickettsial detection further increased the sensitivity of PCR assays and allows for the quantitation of rickettsiae in a sample (216). Speciation of rickettsiae can now be performed using criteria proposed by Fournier et al. (88) (Figure 1.4) and which have since been adopted worldwide as the standard for identifying rickettsial species by PCR.
28
New bacterial isolate
rss > 98.1% (97.7 – 98.1) AND With at least one gltA > 86.5% (84.4 – 88.6) Rickettsia species
No Yes
Other genus Member of the Rickettsia genus
Presence of the OmpA gene
No Yes
Presence of the OmpB gene Member of the spotted and gene D fever group No Yes Yes
> 2 of these 4 criteria This isolate does rss > 98.8% (98.7 – 98.9) gltA > 92.7% (91.9 – 93.4) With at least one member not belong to any ompB > 85.8% (84.1 – 87.7) of the spotted fever group of the two groups, Gene D > 82.2% (80.9 – 83.5) like R. canadensis and R. bellii No
> 2 of these 4 criteria rss > 98.8% (98.7 – 98.9) gltA > 92.7% (91.9 – 93.4) With either R. typhi and/or Yes ompB > 85.8% (84.1 – 87.7) R. prowazekii Gene D > 82.2% (80.9 – 83.5)
Member of the typhus group
Figure 1.4 – Figure showing the amplification and sequence similarities requirement of 5 rickettsial genes to determine group/ species/novelty of a Rickettsia isolate. Adapted from Fournier et al. (88).
29
1.6.4 Treatment
A rickettsiosis is in some cases a self limiting disease that can be of minor clinical significance or asymptomatic and not require treatment. However, there have been multiple cases of severe rickettsiosis and treatment with antibiotics such as tetracycline or chloramphenicol is essential to avoid morbidity and mortality (87, 106). These drugs however do cause significant side effects in children such as staining of teeth by tetracyclines and bone marrow toxicity by chloramphenicol (8). Symptoms relating to rickettsial infection usually resolve within 48 to 72 hours of treatment. Treatment regimens last from between 5 to 14 days depending on the severity of the disease.
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1.7 Tick-Borne Rickettsioses
Despite the close similarities between the members of the SFG, there are differences that make them distinct enough to warrant separate species classification. The diseases caused by the various members of tick-borne Rickettsia vary in severity though they do share many common symptoms. All tick-borne Rickettsia are currently classified as belonging to the spotted fever group and considered potentially pathogenic to human.
Eleven forms of SFG rickettsiosis as a result of infection by an SFG Rickettsia have been documented in literature outside of Australia since the beginning of the 20th century. Australian rickettsioses will be covered in section 1.9.
1.7.1 Rickettsia rickettsii
Records of spotted fever illness in western Montana, USA, date back to 1873 and were first described by Wilson and Chowning in 1904 (202). Misconception of the aetiologic agent, initially thought to be Pyroplasma hominis (an intraerythrocytic parasite), made by Wilson and Chowning was later corrected by Howard T. Ricketts who further expanded the global medical scientific community’s understanding of rickettsioses.
Ricketts demonstrated that the aetiologic agent for Rocky Mountain spotted fever
(RMSF) was a rickettsia that was transmitted by the tick Dermacentor andersoni. Since
Ricketts’ initial publications, a huge interest in rickettsial infections among physicians and scientists has resulted in a more accurate understanding of the aetiologic agent, R. rickettsii and the disease it causes.
Several tick species including Dermacentor, Rhipicephalus and Amblyomma act as the reservoirs for R. rickettsii (62, 119). Vertebrate amplifying hosts of R. rickettsii include domestic dogs and small mammals such as cotton rats (Sigmodon hispidus), ground squirrels (Spermophilus lateralis), opossums (Didelphis virginiana) and capybaras
31
(Hydrochoerus hydrochaeris) (184, 287). Cases of RMSF and isolation of R. rickettsii from their arthropod vectors have been reported in North, Central and South America
(51). The clinical symptoms of RMSF appear approximately 3-12 days after rickettsial inoculation from a tick bite. RMSF begins with fever followed by a maculopapular rash that develops on the second to fourth day after the onset of fever. Other accompanying symptoms include nausea, vomiting, anorexia, headaches and generalized myalgia (49,
175). In more severe cases of RMSF, pathophysiological manifestations of the disease include vascular permeability leading to pulmonary or cerebral oedema, renal failure, myocarditis and gangrene (51). Rickettsia rickettsii is the most virulent of all known spotted fever rickettsiae with a 70% mortality rate among the earliest recorded cases.
Rates of mortality declined with the discovery of antibiotics however rates are affected by the promptness of appropriate treatment and correct diagnosis of the disease. This was evident with the high mortality rate of 40% in Brazil in the 1980s (99, 160). The mortality rate for RMSF in the USA has dropped significantly in the later part of the
20th century, now hovering between 2-6% (210).
1.7.2 Rickettsia conorii
Rickettsia conorii, the causative agent of Mediterranean spotted fever (MSF), was first described in Tunisia in 1910 by Conor and Brush (56, 171). Since then, R. conorii has been reported from locations around the Mediterranean, central Europe, India, the
Middle East and central and southern Africa. The main vector for R. conorii is the brown dog tick Rhipicephalus sanguineus, which has a worldwide distribution as a result of the global spread of domesticated dogs along with human populations. There are several known sub-species of R. conorii transmitted by the tick Rh. sanguineus.
These include; R. conorii conorii (agent for MSF), R. conorii israelensis (agent for
Israeli spotted fever, ISF), R. conorii caspia (agent for Astrakhan fever, AF) and R.
32 conorii indica (agent for Indian tick typhus, ITT). The clinical symptoms for all four diseases are varied in severity and presentation. Patients infected with MSF have been reported to present with an eschar, high fever (>39°C) and flu-like symptoms (10).
Mortality as a result of disease is extremely rare, about 2.5% at its highest. The severity of disease is attributed to the susceptibility of the host and the virulence of the infecting strain (4).
ISF was first described in 1946 and its vector and agent later isolated in 1971 (110,
310). Isolates proved to be closely related to R. conorii but slightly different, which led to the proposal by Zhu et al. (2005) to classify it as a sub species of R. conorii; R. conorii israelensis. The distribution of R. conorii israelensis has been reported as far as
Portugal and Sicily (59, 104). Ninety percent of ISF cases do not have an eschar present, the site of inoculation appearing as a small pinkish papule (117).
AF is a more recent spotted fever that was reported first in 1972. Patients were initially reported from the rural areas of Astrakhan, Russia, adjacent to the Caspian Sea (297).
An isolate obtained from a patient in 1991 showed identical genomic patterns with that of R. conorii israelensis (74). The causative agent for AF is now classified as a sub species of R. conorii, R. conorii caspia (341). Other than Rhipicephalus sanguineus, R. conorii caspia has been found in Rh. pumilio and its area of distribution may include
Kosovo and Chad (75, 89, 90). Symptoms for AF are similar to that of MSF, with most patients presenting with maculopapular rash, myalgia and headaches however the presence of an eschar is infrequent with most cases being without an eschar. Other symptoms such as conjunctivitis and hepatomegaly are relatively common (297).
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The agent for ITT was first isolated in 1950 (209, 271). Since then it has been shown to differ antigenically from R. conorii, but there was no doubt that the agent is of rickettsial spotted fever origin (227). Zhu et al. (2005) recently suggested that the ITT agent be classified as a sub species of R. conorii as R. conorii indica (341). There are few differences between ITT and MSF, with the main differences being that rash in ITT is purpuric and the presence of an eschar is rarely observed (141). ITT has been reported outside of India, with cases appearing in Thailand and Laos (220, 228).
1.7.3 Rickettsia africae
Rickettsia africae is the causative agent of African tick bite fever (ATBF) and is endemic in rural sub-Saharan Africa and the Caribbean (144, 156). The first recorded cases date back to the early 20th century in Mozambique and South Africa (144). Ticks belonging to the genus Amblyomma have been reported as the main vector. R. africae transmission by A. hebraeum in southern Africa and A. variegatum in West, Central and East Africa are well documented (24). Amblyomma variegatum is also the tick vector for R. africae in the Caribbean (157).
Infection with R. africae is believed to be asymptomatic in 50% of cases with most cases being asymptomatic or clinically mild and self-limited (143). In cases where symptoms are present, they usually include fever, nausea, headache and myalgia, similar to other spotted fever illness. An eschar is usually seen and multiple eschars are common (247). Complications with ATBF are rare but cases with long lasting fever, subacute neuropathy, chronic fatigue and myocarditis have been reported (27, 145).
There has been no reported mortality as a result of infection or disease.
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1.7.4 Rickettsia sibirica
The species R. sibirica is made up of two sub species; R. sibirica sibirica and R. sibirica mongolotimonae. Both are recognized aetiologic agents for spotted fever rickettsioses with R. sibirica sibirica being the agent for Siberian tick typhus (STT) or
North Asian tick typhus, and R. sibirica mongolotimonae being the agent for lymphangitis-associated rickettsiosis (LAR) (91, 187). There are few similarities between the two sub species, and the main reason for putting them within the same species is the genotypic similarities they share (88).
Siberian tick typhus (STT) was first recorded in Russia in the 1930s. Since then it has been found in various parts of north Asia with recorded cases in Kazakhstan,
Kyrgyzstan, Mongolia and China (266). Tick vectors for the aetiologic agent include
Demacentor nuttali, Dermacentor marginatus, Haemaphysalis concinna and
Dermacentor silvarum (18). Presence of R. sibirica sibirica in small mammals has also been demonstrated, giving rise to the possibility of them acting as a reservoir (50).
Common STT symptoms include eschar, maculopapular rash, high fever, arthralgia and myalgia (244).
Rickettsia sibirica mongolotimonae was first reported in 1991 with an isolate from the tick species Hyalomma asiaticum from Inner Mongolia in China (339). Its next recorded isolation was from Marseille, France, in 1996 from a human patient (243).
Since these two cases were reported, R. sibirica mongolotimonae has been associated with the occurrence of lymphangitis-associated rickettsiosis (LAR) in Europe, Africa and China. Other than Hyalomma asiaticum, it has been isolated from other tick species; H. truncatum, H. anatolicum excavatum and Rh. pusillus (61, 218, 236).
Common symptoms for LAR include rash, multiple eschars, fever and lymphangitis
35 between the inoculation eschar and the draining lymph node thus giving its name lymphangitis-associated rickettsiosis (91). However it was reported that less than 50% of cases had lymphangitis, which is not uncommon among other rickettsiosis (91).
1.7.5 Rickettsia heilongjiangensis
Rickettsia heilongjiangensis is the aetiologic agent for Far-Eastern tick typhus. It was initially mistaken for STT and was only recently described as a new species of spotted fever group rickettsia (88). R. heilongjiangensis is grouped within the same cluster as
R. japonica phylogenetically. Several tick species have been identified as vectors of the rickettsia, including D. sylvarum and H. concinna (186). The clinical symptoms of Far-
Eastern tick typhus include eschar, fever, headache, rash, regional lymphadenopathy and conjunctivitis. The disease is relatively mild in patients and most cases observed occur in the elderly, with only one case under the age of 45 (244).
1.7.6 Rickettsia japonica
The first observed cases of R. japonica infection (Japanese/Oriental spotted fever) were in 1984. The disease closely resembles scrub typhus (ST) however disease occurrence was in a geographical area non-endemic for ST and the high affinity of antibodies for
SFG rickettsial antigens gave reason for it to be described as a new SFG rickettsiosis
(302). The new species of spotted fever group rickettsia, R. japonica, was later isolated from patients in 1985 (303). Several ticks have been found to carry R. japonica, including Haemaphysalis longicornis, Haemaphysalis flava, D. taiwanensis, I. persulcatus, I. monospinosus and I. ovatus (86). Clinical symptoms include fever, chills, macular rash, headache and eschar. Severe cases are not uncommon with fatalities occurring due to encephalitis, multi organ failure and acute respiratory distress syndrome (162). Recently, several genotypic variants of R. japonica have been found in
36
Korea, leading the hypothesis that there are several regional geno-variants of this rickettsia in east Asia (53, 169).
1.7.7 Rickettsia slovaca
Rickettsia slovaca was first isolated in 1968 in Slovakia (then Czechoslovakia) and has since been classified as a novel member of spotted fever Rickettsia (280). It has been isolated from the ticks Dermacentor marginatus and D. reticulatus. The first recorded case of rickettsiosis as a result of R. slovaca infection was not until 1997, almost 30 years after the discovery of the aetiologic agent and it was given the name tick-borne lymphadenopathy (TIBOLA) (167). Signs associated with TIBOLA include eschar and enlarged cervical lymph nodes with sequelae involving alopecia and asthenia. Classic spotted fever symptoms of fever and rash were uncommon among patients (249).
Another disease associated with R. slovaca infection is Dermacentor-borne-necrosis- erythema-lymphadenopathy (DEBONEL). Some of the signs and symptoms recorded from patients diagnosed with DEBONEL include eschar, painful lymphadenopathy and a slight fever (205). Most eschars observed on patients with TIBOLA and DEBONEL were found on the scalp. TIBOLA and DEBONEL, and their aetiologic agent have been reported from Europe only so far.
1.7.8 Rickettsia aeschlimanii
Rickettsia aeschlimanii was first isolated in 1992, but it wasn’t until 1997 that it was deemed to be a novel species of SFG rickettsia (25). First isolated from Hyalomma marginatum marginatum ticks, it has been found in other ticks species, including
Hyalomma marginatum rufipes and Rhipicephalus appendiculatus (218, 235). The geographic distribution of R. aeschlimanii coincides with the tick species that harbours it, specifically southern Europe, north Africa and south Africa. The first human
37 infection was reported in 2002 and the patient presented with multiple eschars and a maculopapular rash (248).
1.7.9 Rickettsia massiliae
Rickettsia massiliae is one of the many spotted fever rickettsiae whose clinical significance is relatively unclear. It was first isolated from Rh. sanguineus ticks near
Marseille, France, in 1992 (23). Since then, it has been found in 4 other tick species which act as reservoirs; Rh. sanguineus, Rh. muhsamae, Rh. lunulatus, Rh. sulcatus and
Rh. turanicus. The geographical distribution of R. massiliae has been determined to be central and southern Europe, central Africa, North and South America (17, 70, 76,
101). First recorded human infection by R. massiliae was in 1985 from Palermo, Italy, but the identification of R. massiliae as the aetiologic agent was only obtained in 2005.
The patient presented with fever, eschar, maculopapular rash and hepatomegaly (315).
1.7.10 Rickettsia honei
Although this species was first characterised in Australia in 1991 (115, 294) (section
1.9), it was recognised but not identified in other locations around the world many years beforehand. The first isolation of R. honei was in 1962 from a pool of Ixodes sp. and Rhipicehalus sp. ticks collected off common rats in Thailand and was designated as
TT-118 (264). The novel rickettsia was coined Thai tick typhus by its discoverers. This find was later reproduced when R. honei DNA was isolated from Ixodes granulatus off
R. rattus in 1974 from Thailand (163). The first recorded case of human Thai tick typhus was in a febrile patient in 2005 (146), from whom R. honei DNA was extracted from their serum. R. honei has also been detected in the USA from Amblyomma cajennense ticks off cattle, although no reports of human rickettsiosis has been reported in the USA to date (29).
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1.7.11 Rickettsia parkeri
Rickettsia parkeri was not recognised as a cause of rickettsiosis until 65 years after it was first described. For 65 years R. parkeri was thought to be non-pathogenic until recent work demonstrated otherwise (212). The “newly” discovered pathogenic nature of R. parkeri emphasises the lack of current knowledge on rickettsial virulence for humans. Even though a large wealth of information had been uncovered in the last century, our understanding on the pathogenicity of rickettsiae worldwide is still limited.
The geographic distribution of R. parkeri has since been determined to stretch from the
U.S.A and all the way down to central South America (215, 299). Rickettsiosis as a result of R. parkeri infection has been observed to be milder than RMSF. Patients present with fever, an eschar and a maculopapular rash (212, 214, 330).
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1.8 Non Tick-Borne Rickettsiosis
There are four known non-tick borne rickettsioses and they split into two different groups. Typhus group rickettsia consists of R. typhi and R. prowazekii and the recently proposed Transitional group rickettsia consists of R. akari and R. felis. Two other known rickettsiae that are not tick-borne, R. canadensis and R. bellii (members of AG) have so far been considered as non-pathogenic to humans; however the current perception is due to limited evidence to prove them as human pathogens and not due to any in depth studies to prove non-pathogenicity (38, 198). The rickettsiosis caused by members of TG and TRG vary in severity, though they do share some clinical similarities to the rickettsiosis of tick-borne/SFG rickettsia
.
1.8.1 Rickettsia typhi
Rickettsia typhi is the causative agent of murine (or endemic) typhus. It is one of the two members of the Typhus group along with R. prowazekii. The vector of R. typhi has been established as the rat flea, Xenopsylla cheopis and its distribution is worldwide and its incidence is higher in the developing parts of the world where hygiene and rat control is inadequate (13, 71, 105). Other vectors that have been shown to transmit or carry R. typhi include the fleas, most notably the cat flea (Ctenocephalides felis), and also lice, mites and ticks (14, 52, 77, 289). The primary reservoir for R. typhi are rats belonging to the genus Rattus, mainly Rattus rattus and R. norvegicus. Other rodents and small mammals have also been implicated as reservoirs for R. typhi, including rats, mice, cats and opossums (48, 285, 333).
The first description of the disease typhus was by Fracastoro in 1546 and the first documentation of typhus in literature was published in 1917 (192). Presentations of clinical symptoms usually occur approximately 6 to 14 days after inoculation. Clinical
40 manifestations as a result of R. typhi infection include; fever, rash, chills, nausea and headaches (105). The rash reported are usually macular in appearance, but there has been instances where the rash appearance were maculopapular, papular, petechial or morbilliform (68). More severe forms of murine typhus are not uncommon and usually resulting in meningoencephalitis and interstitial pneumonia with other signs including hepatomegaly, splenomegaly, pulmonary pleural effusion, respiratory failure, acute renal failure and cardiac dysfunction (68, 282). The rate of mortality is very low and has been recorded to be less than 5% at worst (in the pre-antibiotic era).
1.8.2 Rickettsia prowazekii
The rickettsiosis that results from infection with R. prowazekii is known by various names some of which include “Jail fever”, “epidemic typhus” or just simply “typhus”.
Epidemic typhus has been attributed to be the cause of uncountable deaths in human recorded history, going as far back as the Napoleonic wars and most notable during the
First World War where 3 million Russians died as a result of infection between 1917 and 1925 (222).
The natural vector for R. prowazekii is the human body louse (Pediculus humanus humanus or P. h. corporis) (118). The reservoir for R. prowazekii in North America is the flying squirrel. Another purported reservoir for R. prowazekii is recrudescence in humans (116). The geographical distribution of the human body louse is global with major infestations coinciding with over-crowded and unhygienic human populations
(134). Clinical manifestations of epidemic typhus appear from 10-14 days after infection and include malaise, anorexia, chills, headache, myalgias, arthralgias, rash and fever (225). Rash appears as either macular, maculopapular or petechial. Severe cases of epidemic typhus have been reported with gangrene, seizures, coma, hearing
41 loss, myocarditis, jaundice, pulmonary complications and hepatitis. Rate of mortality in untreated individual can be as high as 30% (64, 95).
1.8.3 Rickettsia akari
Rickettsia akari is the only member of the genus Rickettsia to have mites exclusively as a vector. First observed in 1946 in New York, USA, almost 100 more cases followed closely in the same area within the same year (213). The established vector for R. akari is the mouse mite Liponyssoides sanguineus. Lice have been shown to harbour R. akari although they are unable to transmit the rickettsial agent to humans (328). The natural amplifying host for R. akari is the mouse, Mus musculus. Other rodents and small mammals have been shown to have the potential to act as hosts including the rat Rattus norvegicus and the Korean reed vole, Microtus fortis pelliceus (139). The known geographic distribution of R. akari is in north America, Europe and Africa (41, 103,
274).
Clinical manifestations of rickettsialpox appear 10-15 days after initial infection (213).
Lesions are commonly found at the bite site about 2 days after inoculation before clinical manifestations such as headache, myalgia, high fever and papulovesicular rash
(151). Severe cases of rickettsialpox are rare and no deaths as a result of infection by R. akari have been reported (211).
1.8.4 Rickettsia felis
Rickettsia felis was first discovered in C. felis in 1918 (284), which has since been established as the major carrier of R. felis worldwide. Other flea species associated with
R. felis are Ctenocephalides canis, Pulex irritans, Anomiopsyllus nudata, Xenopsylla cheopis and Archaeopsylla erinacei (15, 31, 147, 220). A recent study into the
42 prevalence of R. felis in Western Australia has added two more flea species to the list;
Echidnophaga gallinacean and Spilopsyllus cuniculi (277). The current known reservoirs of R. felis included more than twenty species of fleas, ticks, lice and mites
(Table 1.3). The large number of reservoirs and potential vectors has made R. felis a growing cause for concern as an emerging health threat worldwide (224, 254). The first report of human infections in Australia was by Williams et al. (332). Clinical manifestations of cat flea typhus include fever, fatigue, headache, maculopapular rash and a lesion at the site of infection (bite) (246). Cat flea typhus is not known to have any serious complications and represents a milder form of rickettsioses. No mortality has ever been reported (342).
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Table 1.3 – Invertebrate hosts of Rickettsia felis, some of which carry other potential bacterial pathogens. Invertebrate Host Vertebrate Host Location (Identified) Other Potential Pathogens* Disease Reference Fleas Ctenocephalides felis Dog, Cat, Rodents, Argentina, Australia, Brazil, Canada, Chile, Cyprus, France, Rickettsia typhi Murine typhus (22, 31, 32, 108, 149, 158, Monkey, Opossums Gabon, Germany, Israel, Mexico, New Zealand, Peru, Spain, Bartonella sp. Cat-scratch disease 159, 166, 179, 191, 200, Taiwan, Thailand, UK, USA, Uruguay 220, 237, 265, 267, 277, 300, 312, 333, 340) Ctenocephalides canis Dog, Cat Algeria, Brazil, France, Spain , Thailand, Uruguay Bartonella sp. Cat-scratch disease (31, 34, 107, 130, 220, 312) Pulex Irritans Human and mammals DR Congo, USA Rickettsia typhi Murine typhus (15, 273) Yersinia pestis Plague Xenopsylla cheopis Rodents, Shrew Indonesia, USA Rickettsia typhi Murine typhus (77, 147) Yersinia pestis Plague Anomiopsyllus nudata Rodents USA - - (293) Archaeopsylla erinacei Hedgehog, Dog, Cat Algeria, France, Germany - - (31, 107, 108) Echidnophaga gallinacean Poultry, Dog, Cat Australia, DR Congo Rickettsia sp. Spotted fever (273, 277) Spilopsyllus cuniculi Cat, Rabbit Australia - - (277) Ctenophthalmus sp. Rodent Portugal Rickettsia typhi Murine typhus (60) Bartonella sp. Cat-scratch disease Xenopsylla brasiliensis Rodent Brazil Yersinia pestis Plague (273) Tunga penetrans Human, Dog, Cat, Pig Brazil - - (273) Polygenis atopus Dog, Cat, Opossum Brazil - - (131) Ticks Haemaphysalis flava Cat Japan Rickettsia japonica Japanese spotted (136) fever Haemaphysalis kitaokai Cattle, Deer Japan - - (136) Rhipicephalus sanguineus Dog, Horse Brazil, Japan Rickettsia sp. Spotted fever (136, 199) Anaplasma sp. Anaplasmosis Amblyomma cajennense Dog, Horse Japan Rickettsia sp. Spotted fever (136) Ixodes granulatus Shrew Taiwan Rickettsia sp. Spotted fever (301) Ehrlichia sp. Ehrlichiosis Ixodes ovatus Cat Japan Rickettsia sp. Spotted fever (136) Carios capensis Seabird USA Coxiella sp. Unknown (67, 252) Rickettsia hoogstraalii Spotted fever Haemaphysalis sulcata Sheep, Goat Croatia Rickettsia hoogstraalii Unknown spotted (67) fever Mites Trombiculid Wild Rodents South Korea Orientia tsutsugamushi Scrub typhus (52) Leptotrombidium deliense Rat Taiwan Orientia tsutsugamushi Scrub typhus (301) Mesostigmata Rat Taiwan - - (301) Lice Liposcelis bostrychophila - Canada - - (26) * Multiple bacterial pathogens present in the same host are simplified with genus only. Fields with no bacterial pathogens listed denotes no record in literature. 44
1.8.5 Orientia tsutsugamushi
Orientia tsutsugamushi (originally Rickettsia tsutsugamushi) though no longer classified as a member of the genus Rickettsia is still considered to be a rickettsial organism with significant public health implications (296). Its main reservoir and vector are mites belonging to the trombiculid family, more specifically, the species
Leptotrombidium deliense. The geographical range of O. tsutsugamushi appears to be restricted only to Asia, Oceania and Australia. In Asia, scrub typhus has been described as far north as Japan (296), and covering the whole of South-East Asia (228) and spreading west-wards to Afghanistan. Scrub typhus has so far been reported from the northern part of Australia, in the states of Western Australia, Northern Territories and
Queensland (112, 114).
The recent discovery of a novel isolate of an Orientia-like organism has seen the addition of a second member to the genus (137). The organism’s name has been proposed as Orientia chuto, isolated from an Australian tourist returning from the
United Arab Emirates, where she was believed to have been infected. This places O. chuto outside the traditional geographic area of O. tsutsugamushi.
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1.9 The Australian Situation
Recorded rickettsial infections in Australia go as far back as the early 20th century approximately 15 years after Rickett’s discovery of Rickettsia rickettsii was published
(128). There are four members of the Rickettsia genus in Australia that has been described thus far, two of them believed to be native to the continent (R. australis and
R. honei) and another two (R. typhi and R felis) believed to have been introduced to
Australia by migration of humans and their accompanying animals and possibly rodents that followed. There have been a few reports of distinctly different Rickettsia-like organisms that are currently being investigated to determine their relation to other members of their suspected genus. Other rickettsial/rickettsia-like organisms reported from the Australian continent but not investigated in this thesis include Bartonella henselae, Bartonella quintana, Bartonella australis, Anaplasma marginale, Anaplasma platys, Coxiella burnetii and O. tsutsugamushi (43, 84, 92, 182, 337).
Rickettsia australis
The first spotted fever rickettsiosis reported in Australia was described as Queensland tick typhus from 13 cases around Cairns in the state of Queensland (7, 39) (Figure 1.5).
It was later established that the agent was related to R. typhi and other members of the genus Rickettsia but distinct enough to be described as a new species and thus named R. australis (98, 226, 234). The vectors for QTT were not established until the mid 1970s, the three tick species that has been identified as vectors of R. australis are Ixodes tasmani, Ixodes holocyclus and Ixodes cornuatus (45, 115). The distribution of the ticks that have been found to harbour R. australis ranges from the Torres Strait islands, northern Queensland, Sydney, New South Wales and south-eastern Victoria (231).
Symptoms attributed to QTT include headaches, chills, malaise, fever, maculopapular
46 rash and an eschar at the site of the tick bite. Of the 40 cases reported in the literature since the first QTT incident, there have been 2 cases of documented mortality (231).
Rickettsia honei
Rickettsiosis as a result of R. honei infection was first reported in Australia in 1991 on
Flinders Island. It was originally thought to be QTT (caused by R. australis), even though there were initial clinical and epidemiological differences. Some of the clinical illness presented includes fever, myalgia, headache, cough and rash (294). The rickettsiosis was named Flinders Island spotted fever (FISF) and the rickettsia after
Frank Sandland Hone, the first Australian rickettsiologist. The vector for R. honei had been determined to be the reptile tick Bothriocroton hydrosauri (previously of the genus
Aponomma) which can be found in South Australia (SA), Victoria, Tasmania and New
South Wales (Figure 1.5). However, the known geographical distribution in Australia for R. honei is limited to South Australia and Tasmania (113, 263).
Unsworth et al. (306) reported the isolation of a R. honei-like rickettsia that causes rickettsiosis in humans in 2007. The isolate was closely related to R. honei and was thus designated R. honei strain “marmionii”. Since then the rickettsiosis has been named
Australian spotted fever (Figure 1.5). R. honei strain “marmionii” has also been isolated from the blood of two patients diagnosed with chronic fatigue syndrome (CFS) (305) giving support to the hypothesis that some cases of CFS may be as a result of chronic rickettsiosis. The name “marmionii” was given to honour the Australian rickettsiologist
Barrie Marmion, who developed the Q fever vaccine.
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A B
C D
E Figure 1.5 – Geographic distribution of known rickettsial organisms in Australia; (A) R. australis, (B) R. honei, (C) R. honei strain “marmionii”, (D) R. typhi, and (E) O. tsutsugamushi. Courtesy of the Australian Rickettsial Reference Laboratory (ARRL).
48
Rickettsia felis
The presence of R. felis was undocumented in Australia until 2006, when a study by
Schloderer et al. (277) demonstrated the presence of R. felis DNA in fleas. There were two flea species isolated from dogs and cats which were identified as novel carriers of the rickettsia; Echidnophaga gallinacean and Spilopsyllus cuniculi. This discovery further added to the growing list of rickettsiae in Australia, especially in Western
Australia where the knowledge of rickettsial epidemiology is least advanced. No other information pertaining to R. felis has been reported since from WA.
The presence of R. felis has recently been reported in several locations in the Eastern states of Australia in fleas and ticks collected off animals (21, 314). A cluster of cases involving 6 patients in Lara, Victoria were the first human reported cases of rickettsiosis attributed to R. felis in Australia (332). All cases had evidence for C. felis as the vector of R. felis. These recent findings and clinical cases highlight the need for rickettsiosis
(cat flea typhus) to be seriously considered as differential diagnoses for cases presenting with fever and rash.
Rickettsia typhi
The first R. typhi infections in Australia were first described as a “typhus-like illness” during an outbreak among South Australian waterside workers in 1922 (127-129). Soon after the report on the initial outbreak of endemic typhus in Adelaide, a similar outbreak was reported from Toowoomba district in Queensland (329). A second outbreak involving 44 cases in Toowoomba was reported between 1948 and 1959 (63).
The first recorded case of murine/endemic typhus in Western Australia (WA) was reported in 1927. A total of 1332 cases were reported since then until 1952 with 50
49 deaths, mostly among the elderly. The geographic distribution of disease occurrence was centred on Perth and Fremantle, but also included other centres of high human habitation in the south west of the state, Geraldton and Albany (275). The most recent
WA cases of recorded murine typhus in humans were a cluster of six cases in Albany between 1995 and 1996 (155, 195). A serosurvey conducted in the Kimberley (northern
WA) showed evidence of seroprevalence for typhus group antibodies in humans (112).
The latest human case of endemic/murine typhus in Australia was reported from
Geelong, Victoria in 2004 (148). R. typhi is thought to be endemic throughout the whole of Australia (Figure 1.5).
Rickettsia gravesii sp. nov.
Rickettsia gravesii was first reported from WA by Owen et al. (206, 207) from ticks collected off various marsupials. The rickettsia was named after Dr Stephen Graves by
Owen et al. in recognition of Graves’ contributions to Australian rickettsiology. Several tick species have been implicated but the tick species believed to be the main reservoir of this new rickettsia was Amblyomma triguttatum (206). A. triguttatum has been recorded to have a geographic distribution as far east as Victoria and far north in WA
(263). The tick species however is made up of 4 different sub-species (262) and the prevalence of R. gravesii within A. triguttatum and other tick species that bite humans in WA has yet to be fully established.
Orientia tsutsugamushi
Even though O. tsutsugamushi was not investigated during the course of this study, its inclusion here is due to its very close relationship to members of the genus Rickettsia.
O. tsutsugamushi is found only in the northern part of Australia covering three states, northern Western Australia, Northern Territories and northern Queensland (Figure 1.5).
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The first reports of scrub typhus (O. tsutsugamushi) in Australia goes back as far as
1935 (168, 308). Isolation of the organism itself was reported in 1955 by mouse inoculation (47) thus confirming presence of O. tsutsugamushi which till then had only been reported by the Weil-Felix test. Since then scrub typhus had been reported in WA
(239), Torres Strait islands of northern QLD (79) and NT (196). The latter was a strain
(Litchfield) characterised as distinct genotypically and serologically from the three main strains of Karp, Kato and Gilliam.
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2 - General Materials and Methods
2.1 Laboratory Conditions Used for Studies
All experiments were performed in laboratories adhering to PC2 guidelines unless stated otherwise.
2.2 Identification of Tick Species
Identification of tick species was performed by light microscopy using a Wild M3 stereomicroscope (Leica, Germany). Standard morphological criteria and dichotomous keys (Figures 2.1 and 2.2) as described by Roberts (262) were used for identification of tick genus and species.
52
A B
Palpi
Coxa Basis capituli Spurs
Eyes Adanal Scutum shield
Festoons Anus
Tarsi Anal Groove
Figure 2.1 – Morphological characteristics of an Ixodidae tick. (A) Dorsal and (B) ventral anatomy of a male Ixodidae tick used in determining genera and species.
53
Anal groove distinct, embracing anus Anal groove obsolete or embracing anus Eyes present? anteriorly; festoons and eyes absent; posteriorly; festoons and eyes maybe present; inornate; male with sclerotized plates ornate/inornate; male with sclerotized plates on covering almost entire ventral surface ventral surface when present, only circumanal
Anal grooves Yes No Ixodes obsolete; festoons absent; palpi short and transversely ridged
Palpi short, conicle, article Palpi elongate, article 2 not Boophilus Anal grooves 2 salient basolaterally; salient basolaterally; distinct; festoons scutum inornate; tarsi of scutum frequently ornate; present; palpi not male not conspicuously tarsi of male conspicuously transversely ridged humped and usually humped and armed with unarmed ventral spurs
Palpi short; basis capituli Palpi short; basis capituli hexagonal dorsally frequently hexagonal dorsally frequently with strong, lateral points; with strong, lateral points; scutum inornate; male with scutum inornate; male with adanal and accessory plates adanal and accessory plates Haemaphysalis Aponomma
Rhipicephalus Amblyomma
Figure 2.2 – Dichotomous key for the identification of Australian Ixodidae genera. Adapted from Roberts (262). 54
2.3 DNA Extraction
DNA extraction from arthropods and cell culture was performed using the QIAamp®
DNA Mini Kit (Qiagen, Germany) according to the manufacturer’s instructions.
Live arthropod samples were immersed in 70% ethanol for 2-3 minutes and allowed to air dry on a glass slide. Preserved arthropod samples were dried in the same manner.
Samples were cut to small pieces with a sterile scalpel blade on a glass slide prior to resuspension in 200µL of sterile phosphate buffered saline (PBS) in an eppendorf tube.
Extracted DNA was eluted in elution buffer (70µL) and stored at -20ºC for later use.
DNA was extracted from infected tissue cell culture supernatant (200µL) and eluted in
100µL of elution buffer and stored at -20ºC for later use.
2.4 Rickettsial Culture
Three different cells lines were used for rickettsial culture; L929 (mouse fibroblasts),
Vero (green monkey kidney cells) and XTC-2 (African clawed toad epithelial cells) cells.
2.4.1 Cell Monolayer Cultures
L929 and Vero cells were grown in 25cm2 vented tissue culture flasks at 35ºC with 5%
CO2. Growth media (9ml) used was RPMI 1640 (Gibco, Australia) containing 5% foetal bovine serum (Gibco, Australia), 2mM L-glutamine (Gibco, Australia) and 25mM
HEPES buffered media (Gibco, Australia). XTC-2 cells were grown in 25cm2 non- vented tissue flasks at 28ºC. Cells were grown in 11mL of Leibovitz L-15 media
55
(Gibco, Australia) containing 5% foetal bovine serum (Gibco, Australia), 2mM L-
Glutamine (Gibco, Australia) and 1% tryptose phosphate broth (Oxoid, England).
Flasks with 80% confluency were used for rickettsial infection (sections 2.4.2 and
2.4.3). Cells were passaged 1/10 using trypsin. Cell monolayers with 100% cell confluency were gently washed with 5mL of sterile PBS and then 1ml of 0.5% Trypsin solution with 5.3mM EDTA4Na (Gibco, Australia) was added and incubated at room temperature for 1 minute, followed by a couple of hard knocks on the flask to remove any adherent cells. Growth media (10mL) was added to the cells which were then incubated at appropriate temperature and CO2 content.
2.4.2 Rickettsia Isolation
Rickettsia isolation was mainly performed from arthropods in this study. Arthropods were identified as described in section 2.2. Removal of potential contaminants was done by immersion in 95% sodium hypochlorite solution for 10min, followed by 70% ethanol and air dried in a level 2 biohazard hood. Arthropods were then halved with a sterile scalpel blade and placed in microcentrifuge tubes with PBS for PCR analysis
(500µL PBS) and rickettsia isolation (1mL PBS). Rickettsia isolation involved crushing the arthropod with the blunt end of a sterile plastic inoculation loop (Sarstedt, Germany) and thus releasing the rickettsiae harbored within the arthropod as a suspension in the
PBS. Tick homogenate was then passed through a 0.45µm filter (Millex, Ireland) and inoculated into monolayers of Vero, L929 and/or XTC-2 cells with 80-90% confluency in 25cm2 flasks. Flasks were then centrifuged at 600g for 20min to induce entry of rickettsia into the cell monolayer. Cultures were incubated for up to two months at 35C in 5% CO2 (L929/Vero) or 28˚C (XTC-2), with fortnightly media changes. Cultures were examined microscopically weekly and by microimmunofluorescence monthly. If 56 cell cultures were thought to be positive, by virtue of a CPE, DNA was extracted and the presence of rickettsia confirmed by PCR.
2.4.3 Rickettsia Culture Maintenance
Isolated rickettsial cultures were maintained for DNA extraction and antigen preparation for genetic and phenotypic characterisation. When an existing rickettsial culture had destroyed the cell monolayer, 0.5mL to 2mL of rickettsial suspension was inoculated into a new flask containing a similar cell monolayer with 80-90% confluency centrifuged at 600g for 20min and incubated as appropriate. Residual rickettsial suspensions were used for DNA extraction (section 2.3) and PCR (section 2.6) or antigen prepared for serology (section 2.5).
2.5 Serology
2.5.1 Rickettsial Antigen Preparation
Cell suspensions from rickettsial cultures were heat inactivated at 56ºC for 30 minutes prior to use as antigens for IFA. Antigen concentrations were titrated in order to optimise fluorescence intensity.
2.5.2 Immunofluorescence Assay
Paired human sera (when applicable) were analyzed using the indirect IFA. Rickettsial whole cell antigen preparations (R. australis strain JC, R. akari strain Kaplan, R. conorii strain Malish, R. honei strain RB, R. rickettsii strain Bitteroot, R. sibirica strain 246, R. typhi strain 18 and R. prowazekii strain Breinl) were acetone fixed to 40 well slides.
Sera were diluted 1:128 with 2% casein buffer (PBS with 2% skim milk powder) and
57 incubated with antigen slides in a humid environment at 35˚C for 30min. Slides were washed 3 times with 10% PBS solution and air-dried. Goat fluorescein isothiocyanate
(KPL, USA) labelled antibodies specific for human antibodies (IgM + IgA + IgG)
(diluted 1:100 in 2% casein buffer) were applied and incubated for 30min in a humid environment at 35˚C. The slides were washed as described previously and mounted with a coverslip (Biolab Scientific, Australia) using fluorescent mounting fluid (Dako, USA).
Antibody titres were read using an illuminator-equipped microscope (Leica, Germany).
Positive values were reported for endpoint titres ≥1:128. This cut off value was recommended by the Australian Rickettsial Reference Laboratory (ARRL) to reduce influence of non-specific binding and cross-reacting antibodies (112).
2.6 Rickettsial PCR
2.6.1 Conventional PCR
Conventional PCR analysis was performed on DNA extracts from materials suspected of harbouring rickettsiae including arthropods, rickettsial cultures and infected tissues.
Products were subsequently sequenced to identify the rickettsia detected within the samples. Gene targets used in the PCR assays were gltA, ompA, ompB, sca4, orf17 and tsa56.
Each conventional rickettsial PCR was performed using the same reagent mix and cycling conditions with the exception of primer types, MgCl2 concentrations and annealing temperatures (Table 2.1). Reactions contained 1M of each primer
(GeneWorks, Australia), 200M of each deoxyribonucleotide triphosphate (dNTP), 10 reaction buffer, 2U Taq polymerase (Invitrogen, Australia), 1.5L of DNA extract and water made up to a 25μL final reaction volume. The amplification were performed in a
58
GeneAmp PCR thermal cycler (Perkin Elmer, USA) with an initial denaturation at 95C for 3min, followed by 40 cycles of denaturation at 95C for 30s, 30s at the appropriate annealing temperature (Table 2.1) and extension at 72˚C for 1min followed by a final extension at 72˚C for 10min.
PCR products were electrophoresed in a 1% Tris acetate EDTA agarose gel (Sigma-
Aldrich, USA), stained with ethidium bromide (Sigma-Aldrich, USA) for 30min at
100V and visualized using an ultraviolet transilluminator for comparative analysis with known positive and negative controls.
2.6.2 Sequence Analysis of PCR Product
PCR amplicons positive for rickettsial DNA were excised from gels and purified using the QIAquick DNA clean up kit (Qiagen, Germany) and sequenced using facilities at the State Agricultural Biotechnology Centre (SABC) (Perth, Western Australia).
Sequencing reactions were performed using an ABI Prism Dye Terminator Cycle
Sequencing Core Kit (Applied Biosystems, California) according to the manufacturer’s instructions.
All sequenced products were analysed using the computer program Chromas Lite and were compared to sequence data available from GenBank, using the BLAST 2.2.14 program. Phylogenetic alignment of DNA sequences utilised the programme ClustalX.
Determination of the genetic difference between the Rickettsia spp. by phylogenetic analysis was conducted using Mega2 software (using Tamura and Nei and neighbour- joining algorithms).
59
Table 2.1 – Summary of PCR conditions for Rickettsial DNA amplification.
MgCl2 Annealing Primer Nucleotide sequence Gene concentration (mM) temperature (C) Reference
RpCS877p GGGGGCCTGCTCACGGCGG gltA 3 51 (253)
RpCS1258n ATTGCAAAAAGTACAGTGAACA gltA 3 51 (253)
Rr190.70p ATGGCGAATATTTCTCCAAAA ompA 2 48 (253)
Rr190.602n AGTGCAGCATTCGCTCCCCCT ompA 2 48 (253)
120M59 CCGCAGGGTTGGTAACTGC ompB 2 51 (270)
120807 CCTTTTAGATTACCGCCTAA ompB 2 51 (270)
MTO-1 GCTCTTGCAACTCTATGTT orf17 3 51 (325)
MTO-2 CATTGTTCGTCAGGTTGGCG orf17 3 51 (325)
D1f ATGAGTAAAGACGGTAACCT sca4 2 51 (279)
D928r AAGCTATTGCGTCATCTCCG sca4 2 51 (279)
60
3 - Serological Typing of a Novel Spotted Fever Group Rickettsia: Rickettsia gravesii
3.1 Introduction
Serological typing or serotyping is a classification method based on the bacterial cell antigen reactivity with antisera. It is used to classify viruses, bacteria and other microorganisms. A single species can have multiple serological types (serovars) as demonstrated in Salmonella enterica with more than 2,500 serovars (272). By using serotyping of a Rickettsia against closely related members within the genus, it is possible to classify the organism as either a separate species or serovar. Philip et al.
(227) introduced a method by which rickettsiae could be serotyped and thus phenotypically differentiated. This serotyping method has been accepted as the method of phenotypically describing a new species of Rickettsia, based on antigenic differences
(250).
This study was undertaken to demonstrate R. gravesii differs antigenically from R. australis and R. honei as they are all found in Australia. R. massiliae was chosen as a representative of the R. massiliae sub-group of SFG rickettsia as it is phylogenetically the most closest related to R. gravesii (206) (Figure 3.1).
61
R slovaca 43 31 R africae 29 R sibirica 31 R honei 62 R parkeri 100 R conorii 67 R rickettsii R heilongjiangensis 97 100 R japonica R montanensis
58 R raoultii 78 R gravesii strain BWI-1 96 92 R aeschlimannii 79 R rhipicephali 94 R massiliae R tamurae R australis R felis
Figure 3.1 – Unrooted neighbour-joining tree based on ompA sequences from Rickettsia species with validly published names. Bootstrap values are indicated at the nodes. Genbank accession numbers used listed in Appendix 9.2.
The main bulk of the work described in this chapter was performed at the Faculty of
Tropical Medicine, Mahidol University, Bangkok, with the help of Stuart Blacksell and his team, who kindly provided the BSL3 facilities for rickettsial culture and animal holding facilities. Animal ethics approvals for the work performed were obtained from
Murdoch University (NS2049/07) and Mahidol University (FTM-ACUC 007/2007)
(Appendix 9.1).
62
3.2 Materials and Methods
3.2.1 Rickettsial Suspension Preparation
Rickettsial strains used in this chapter were R. gravesii strain BWI-1, R. australis strain
JC, R. honei strain RB and R. massiliae strain Mtu1. They were grown as previously described in 2.4.3. Rickettsial cultures were removed from their flasks using a scraper and transferred to a sterile centrifuge tube and centrifuged at 1500g for 10min to pellet
Vero cell debris. The supernatant was transferred to a fresh tube and centrifuged again at 8500g for 30 minutes to pellet rickettsiae in the suspension. The supernatant was then discarded and the pellet resuspended in 5mL of sterile PBS and snap-frozed in cryotubes (Nunc, USA) at -80ºC in 1mL aliquots. Aliquots were then used for the plaque assay, mouse inoculations, serology and qPCR.
3.2.2 Plaque Assay Culture
Vero cells were used for the plaque assay and were grown as described in section 2.4.1 before passage into 12 well plates. Monolayers were allowed to develop overnight with
4 an initial seed of 1.5mL of 4.0 x 10 cells/mL and incubated at 32ºC with CO2. When confluent, the medium was removed and wells were seeded with 200μL rickettsial suspension in dilutions ranging from 10-1 to 10-6 with one ten-fold dilution per plate
(Figure 3.1). An undiluted aliquot of each rickettsial suspension was also seeded into a
25cm2 flask with a Vero cell monolayer (Section 2.4.1) as a positive control to ensure viability of rickettsial organisms. Plates were then placed on a mechanical rocking platform at room temperature for 1 hour to allow rickettsial entry into the cell monolayer and the rickettsial suspension was then removed. 1.5mL of 1% agarose overlay with RPMI 1640 and 10% FCS was applied to the monolayers and the plates 63 were then incubated at 32ºC with 5% CO2 for 6 days. A second overlay medium (1% agarose, RPMI 1640, 10% FCS and 0.01% neutral red dye) was added to the wells on day 6. Plates were wrapped in aluminium foil and incubated at 32ºC with 5% CO2 for 4 days and observed for visible plaques every 24 hours.
C 10-1 10-1
R. australis
R. gravesii
R. honei
R. massiliae
Samples in duplicate
Figure 3.2 – Illustration of plaque assay plate at dilution 10-1. Similar plates were prepared for each ten-fold dilution up to 10-6.
3.2.3 Measurement of Protein (Rickettsial Antigen) Concentration of
Rickettsial Preparation
Protein concentration was used as a surrogate measure of approximate antigen concentration used when inoculating rickettsiae into mice (Section 3.2.4). The Biorad
DC Protein Assay kit (based on the Lowry assay) was used with a slight variation to the manufacturer’s protocol. Four dilutions of a BSA protein standard were prepared with concentrations of 0.5mg/mL, 1.0mg/mL, 1.5mg/mL and 2.0mg/mL to produce a
64 standard curve. Standards and samples were prepared using carbonate-bicarbonate buffer of pH 9.6. 50uL of standards and samples were placed in 5mL plastic test tubes and 250uL of Reagent A added to each tube. Tubes were vortexed and 2mL of Reagent
B was added to each. Tubes were allowed to stand for 15 minutes at room temperature and their absorbance read at 750nm. Protein concentration of samples was extrapolated from the standard curve.
3.2.4 Inoculation of Rickettsiae into Mice and Harvesting Anti-Sera
Four week old female BALB/c mice were split into 5 groups of 5-8 mice each as shown in Table 3.1. Rickettsiae were prepared as detailed in section 2.4.1 and quantified as described in section 3.2.2. Rickettsial inocula were administered into mice via the intra- peritoneal route.
Table 3.1 – Mouse groups and rickettsial inoculum size (original and booster) given via the intra-peritoneal route.
Anti-serum Mouse Inoculum Number of Mice Group (Volume = 0.1mL) Control 5 Sterile PBS Anti-R. australis 7 R. australis Anti-R. gravesii 8 R. gravesii Anti-R. honei 7 R. honei Anti-R. massiliae 8 R. massiliae
Inoculated mice were kept in metal shoe box sized containers in an isolated animal holding facility after the first inoculation on day 0. A booster shot with the same antigen preparation as the original inoculum was given to each mouse on day 7. On day 10, mice were anaesthetized using a glass beaker (air tight lid included) with isofluorane, in
65 a fume hood, before blood was collected via the orbital sinus. Blood were collected in
1.5mL eppendorf tubes and allowed to clot at room temperature.
3.2.5 Microimmunofluorescence
As described in section 2.5.2.
3.2.6 ELISA
Enzyme linked immunosorbent assays (ELISA) were performed on the samples to compare the results with those obtained from the IFA. The ELISA method was adapted from the protocol developed by Tanganuchitcharnchai, Mahidol University (pers. comm.). Flat bottomed ELISA plates were coated with 50μL of antigen suspension, comprising 10μg/mL of semi-purified antigen (see section 3.2.1 for antigen preparation and section 3.2.3 for measurement of protein concentration), in carbonate-bicarbonate buffer (coating buffer) per well at 4ºC overnight in a moist chamber. Plates were washed three times the next day with 0.05% PBS-Tween20 and the wells blocked with
3% skim milk for 1 hour at room temperature. Wells were washed again three times and
50μL of serum sample or control were added to appropriate wells as detailed in Figure
3.2. Plates were incubated for 1 hour at 32ºC in a moist chamber followed by another four washes with 0.05% PBS-Tween20. Anti-mouse antibodies conjugated with horse radish peroxidase were added to each well and the plates incubated at room temperature in a moist chamber for 1 hour. Plates were then washed another 4 times with 0.05%
PBS-Tween20 before 50μL of tetramethylbenzidine (TMB) substrate was added to each well and placed in a dark chamber at room temperature for 30min. The reaction was stopped using 50μl of 1M HCl in each well and the optical density (O.D.) read at 450nm in a spectrophotometer.
66
Anti-serum dilutions
Rickettsial Controls
32 64 128 256 512 1024 2048 4096
Antigens 16 + -
1/ 1/ 1/ 1/ 1/ 1/ 1/ 1/ 1/ 1/8
R. australis
R. gravesii
R. honei
R. massiliae
Figure 3.3 – Figure showing serum sample dilutions and control locations. Samples were tested in duplicate with two-fold serum dilutions starting from 1/8 to1/4096. Wells in each plate were coated with rickettsial antigen as shown
67
3.2.7 Statistical Methods
The serotypic differences between the 4 rickettsial strains tested in this chapter were compared by determining the differences between homologous and heterologous antibody titres. The difference between the titres were presented as specificity difference (SPD) as described by Fraser (94). This method was applied to rickettsial serotyping by Philip (227).
SPD = (Aa + Bb) – (Ab + Ba)
Figure 3.4 – Formula for calculation of Specificity Difference (SPD). Aa is the antibody titre of serum A with the homologous antigen a. Bb is the antibody titre of serum B with the homologous antigen b. Ab is the antibody titre of serum A with the heterologous antigen b. Ba is the antibody titre of serum B with the heterologous antigen a.
Working units used in determining the SPD were arithmetic instead of geometric and converted as shown in Table 3.2. Any sera that were negative for antibody titre at titre
1/8 were given a numerical value of 0 and considered to lack any antibody against the tested antigen.
Table 3.2 – Table of conversion from geometric to arithmetic antibody titres for rickettsial antigen-antibody reactions.
Geometric 1/8 1/16 1/32 1/64 1/128 1/256 1/512 1/1024 1/2048 value Arithmetic 1 2 3 4 5 6 7 8 9 value
The SPD of serotypically similar strains would have a value of 0 to 2 when the sum of the heterologous interactions was subtracted from the sum of homologous interactions.
An SPD value of 3 or higher was defined as denoting a significant antigenic difference between the two rickettsial strains/species i.e. they were different serotypes. 68
3.3 Results
3.3.1 Plaque Assay
Using the method described in section 3.2.2, attempts at plaque assay by counting plaque forming units (pfu) were generally unsuccessful in measuring the concentration of rickettsiae. However plaque formation was observed in two of five attempts (Table
3.3). Of the three unsuccessful attempts, one resulted in a generalized CPE rather than plaques and the other two yielded neither plaques nor CPE.
Table 3.3 – Successful plaque formations in two of five attempts and the concentration in pfu/mL.
Organism Concentration of rickettsiae obtained in pfu/mL (a) (b) R. australis 70 20 R. gravesii ND 40 R. honei 70 40 R. massiliae 390 0 ND – Not done.
3.3.2 Protein Concentrations
Absorbance values plotted against BSA protein standards yielded a linear graph (Figure
3.4). From this graph, rickettsial protein/antigen concentrations were determined to fall within between 0.70mg/mL and 0.85mg/mL (Table 3.4). These were later diluted to
50ug/mL for the coating of ELISA plates used in sections 3.2.6 and 3.3.4.
69
OD 750nm
0.5 1.0 1.5 2.0 BSA Concentration mg/mL
Figure 3.5 – Relationship between bovine serum albumin (BSA) concentration and absorbance at 750nm.
Table 3.4 – Summary of absorbance values of rickettsial inocula used and their respective protein antigen concentrations.
Organism O.D. (750nm) Concentration R. australis 0.111 0.70mg/mL R. gravesii 0.136 0.85mg/mL R. honei 0.129 0.80mg/mL R. massiliae 0.122 0.78mg/mL
3.3.3 Microimmunofluorescence
Serology results are shown in Table 3.5. Antibody titre values were converted to a numerical value based on the conversion recommended by Philip et al. (227) (Table
3.2).
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Table 3.5 – Microimmunofluorescence antibody titres against rickettsial antigens with arithmetic numerical values in parentheses.
Antigen Sera R. australis R. gravesii R. honei R. massiliae R. australis 1:256 (6) 1:32 (3) 1:256 (6) 1:16 (2) R. gravesii 1:128 (5) 1:512 (7) 1:256 (6) 1:32 (3) R. honei 1:64 (4) 1:32 (3) 1:256 (6) 1:16 (2) R. massiliae 1:8 (1) 1:8 (1) 1:8 (1) 1:64 (4)
Calculated SPD values are summarised Table 3.6. Surprisingly, R. honei and R. australis were shown to be not antigenically distinct (SPD=2*). SPD of R. gravesii against the other rickettsiae were shown to be at 4 or higher. As expected, R. massiliae has a high SPD of 7 when compared to the other rickettsiae tested.
Table 3.6 – Table of SPD values for homologous and heterologous antibody/antigen interactions showing level of antigenic difference between various rickettsiae obtained by MIF (SPD ≥ 3 indicates distinct serotypical difference).
Antigen Sera R. australis R. gravesii R. honei R. massiliae R. australis 0 5 2* 7 R. gravesii 5 0 4 7 R. honei 2* 4 0 7 R. massiliae 7 7 7 0 *R. australis and R. honei not antigenically distinct by this method.
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3.3.4 ELISA
ELISA serology conducted using the same principles showed higher antibody titres and higher numerical values for SPD calculation (Table 3.7).
Table 3.7 – ELISA antibody titres against rickettsial antigens with arithmetic numerical values in parentheses.
Antigen Sera R. australis R. gravesii R. honei R. massiliae R. australis 1:2048 (9) 1:256 (6) 1:1024 (8) 1:256 (6) R. gravesii 1:512 (7) 1:4096 (10) 1:512 (7) 1:256 (6) R. honei 1:256 (6) 1:64 (4) 1:512 (7) 1:32 (3) R. massiliae 1:256 (6) 1:512 (7) 1:256 (6) 1:256 (6)
SPD values derived from the ELISA results (Table 3.8) showed a similar trend to the
MIF results. R. australis and R. honei were again demonstrated to be not antigenically distinct (SPD=2*). R. gravesii was significantly distinct from the 3 other rickettsiae tested. R. massiliae was antigenically distinct from other 3 rickettsiae tested but with a much lower SPD value as compared to those derived from MIF.
Table 3.8 – Table of SPD values for homologous and heterologous antibody/antigen interactions showing level of antigenic difference between various rickettsiae obtained by ELISA (SPD ≥ 3 indicates distinct serotype).
Antigen Sera R. australis R. gravesii R. honei R. massiliae R. australis 0 6 2* 3 R. gravesii 6 0 6 3 R. honei 2* 6 0 4 R. massiliae 3 3 4 0 *R. australis and R. honei not antigenically distinct by this method. 72
3.4 Discussion
Because Rickettsia are obligate intracellular organisms, plaque assay is the best method of quantifying them. Developed initially for quantifying viruses, the main factor required for plaque assays to work is that the target organism causes cell lysis or CPE.
Other methods that allow for the quantification of rickettsial organisms, such as electron-microscopy, haemagglutination, ELISA and real-time qPCR, are not capable of determining the number of viable infectious particles (36, 311).
Due to time constraints and limited availability of facilities to conduct the study only five attempts at plaque assays were made, three with disappointing results. This highlighted the need for a rickettsial quantitative tool that was reliable and easily performed with average technical competency. We were able to observe some plaque formation in two assays. However the concentration of plaques was much lower and more variable than expected. This could not simply be attributed to a low rickettsial count as one attempt with the same rickettsial culture preparation showed a generalised
CPE even at the highest dilution. All non-infected controls had an intact monolayer without CPE.
An alternate approach to quantifying the concentration of rickettsial protein (surrogate for antigens) used to inoculate mice was the Lowry assay (Section 3.2.3). Purification prior to quantification was performed using differential centrifugation. The main purpose of the Lowry assay was to ensure that the crude rickettsial suspensions inoculated into the mice were all of a similar protein concentration and hence could be assumed to be antigenically equivalent. To ensure viability of rickettsiae after purification, an aliquot of each rickettsial species tested was inoculated into cell culture
73 to test for viability. Viability was confirmed within 7 days post-inoculation via CPE and
IFA.
A more appropriate method, had time been available, would have been the quantification of rickettsial organisms using the TCID50 assay (291). The TCID50 assay has been used to quantify R. australis and with centrifugation was found to be the most sensitive method for obtaining counts of viable rickettsiae when compared to other assay systems tested, e.g. LD50 (lethal dose 50%) and SD50 (seroconversion dose 50%).
Stenos et al. (291) used a different host cell system and a different surface area of the monolayer for plaque formation to this current study.
Even though antibody titres of mice infected with R. massiliae and R. gravesii were low, when titrated against R. gravesii and R. massiliae antigens respectively, the antibody response was considered to be type-specific rather than group-specific. and induced antibodies did not cross-react with heterologous species of rickettsiae in the
SFG (229).
Analysis of the mouse sera to determine the specificity difference between the 4 different tested species of Rickettsia showed a significant difference in serotype (SPD
≥3) between R. massiliae and all 3 Australian isolates (R. gravesii strain BWI-1, R. honei and R. australis). R. gravesii sp. BWI-1 was demonstrated to be serologically distinct from the other Australian isolates (R. honei and R. australis). R. honei however demonstrated a close antigenic relationship with R. australis. This information was surprising considering the distinct genotypic differences between the two species.
74
Based on the geographic distribution of the Australian strains of R. australis and R. honei (Figure 1.6), we observed overlaps with regards to vector distribution and known clinical cases that have been described in the literature. Of particular interest was the presence of R. honei strain “marmionii” (not used in this study) which shares geographic distributions with R. honei and R. australis. All three organisms have different known vectors, thus the unexpected serotypic similarity between R. australis and R. honei cannot be explained by the experiments conducted in this study. Further work needs to be done to explain this phenomenon. There is evidence in the literature of the sharing of genomic material between rickettsial species (33). However no evidence of R. australis and R. honei dual infections in humans or animals have been observed in the literature, potentiating physical transfer of nucleic acids that might explain the antigenic similarities between the two species observed in this chapter.
Similar specificity differences were generated when tested with MIF and ELISA.
However antibody levels were much higher with ELISA suggesting that it may be more sensitive than MIF. In spite of the higher titres obtained with ELISA (Table 3.8), the
SPD values generated were similar to those obtained via IFA (Table 3.6). There were some differences, i.e. the SPD between R. massiliae and all Australian isolates were lower. However, the SPD value obtained was still enough to classify them as different serotypes. The debate as to methodology is ongoing (73, 164). However IFA has become accepted as the gold standard worldwide for rickettsial serology (88).
The findings of this study clearly demonstrated that R. gravesii strain BWI-1 is serotypically distinct from its closest genotypic relative sub-group, represented by R. massiliae, and from the other recognised Australian species, R australis and R. honei.
This indicates that incorporating R. gravesii strain BWI-1 antigens into future routine
75
IFAs for rickettsial diagnostics in Australia may be warranted especially if R. gravesii strain BWI-1 is shown to be pathogenic for humans.
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4 - Amblyomma triguttatum as the Main Arthropod Reservoir of Rickettsia gravesii sp. nov. in Western Australia
4.1 Introduction
The role of ticks acting as potential reservoirs and vectors for spotted fever group rickettsiae in WA was clearly demonstrated by Owen et al. (206-208). Of particular interest was the reporting of two novel rickettsial strains, tentatively named Rickettsia gravesii and Rickettsia antechini, in ticks found in southwest Western Australia (WA) and Barrow Island (208). A larger sample size of ticks biting humans and animals was needed to ascertain the prevalence of these rickettsiae in ectoparasites from the WA bush. Of particular interest was the prevalence within ticks known to bite larger mammals including humans. The tick species of interest were Amblyomma triguttatum and Ixodes australiensis as they are common and widely distributed in WA (262). There are a total of twenty-three recorded tick species in WA (262). The natural hosts of the ticks are varied, ranging from native mammals and marsupials to feral and introduced animals in the bush. Further investigation into other tick species in WA was required to paint a clearer picture of prevalence of infection in ticks that could potentially act as a vector or reservoir of R. gravesii and R. antechini.
The study reported in this chapter aims to address the question of the prevalence of rickettsiae in those tick species with an affinity for larger mammals in southwest WA and Barrow Island.
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4.2 Materials and Methods
4.2.1 Ectoparasite Sampling
Collection of ectoparasites was performed opportunistically as part of other studies conducted on large mammals (Figure 4.1A) in WA by staff of the Department of
Environment and Conservation (DEC) and the School of Veterinary and Biomedical
Sciences, Murdoch University. Samples were also obtained from wildlife rehabilitation centres on the outskirts of Perth and from various human (Figure 4.1B) recreational and occupational groups located throughout WA. A map showing areas from which ticks were sampled is shown in figure 4.2. Ectoparasites were stored in either a humidified container or in 70% (v/v) ethanol and sent to Murdoch University for identification and analysis.
4.2.2 Ectoparasite Identification
Ectoparasite identification was performed using the method outlined in section 2.2.
4.2.3 Ectoparasite Dissection
Ectoparasite dissection was performed by attaching the ectoparasite to Blu-Tack under a dissecting microscope. A sterile scalpel blade was then inserted into one side of the tick and used to cut along the circumference of the tick’s body/scutum as shown in Figure
4.3. Using a pair of sterile metal forceps, the top part of the tick’s body/scutum was pulled forward or removed to expose the internal organs as shown in Figure 4.4.
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A Ixodes australiensis
Amblyomma triguttatum
B
Amblyomma triguttatum
Figure 4.1 – Pictures of ticks on various mammals. (A) Two different tick species (I. australiensis and A. triguttatum) feeding at the same time on the same feral pig. (B) A. triguttatum prior to attachment on a human.
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Figure 4.2 – Figure showing locations of tick sampling sites. 1 - Barrow Island, 2 – Perth metropolitan areas (including Whiteman Park and Kanyana Wildlife Rescue Centre), 3 – Water catchment areas of Dwellingup, Serpentine and Mundaring, 4 – Southwest WA including Bibulman track and roganing areas, 5 – Two Peoples Bay, Albany.
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2 Scalpel blade
1
Direction of cut
Figure 4.3 – Figure showing location and direction of cut using a scalpel blade along the circumference of the tick scutum and body. 1 – Point of insertion of scalpel blade. 2 – End point of cut on opposite side of start site.
Salivary glands
Mid-gut
Ovaries
Figure 4.4 – Figure showing the location of the three main organs removed and analysed for rickettsial DNA. Organs were removed in the order of salivary glands, mid-gut and ovaries to prevent cross-contamination.
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The tick organs of interest were removed using a clean pair of metal forceps, the salivary glands first, followed by the mid-gut and finally the ovaries. Extra care was taken not to break the tissues while handling them to avoid cross-contamination from one organ to the next. Changing of metal forceps and cleaning with sterile 70% ethanol in between organs was performed to reduce the likelihood of carryover of rickettsial organisms. Removed organs were placed in sterile PBS and kept at 4˚C prior to DNA extraction in separate eppendorf tubes.
4.2.4 Transfer of Rickettsiae from Female Parent to Larvae
To demonstrate the ability of rickettsiae to transfer from an adult female to tick larvae via vertical (ovarial) transmission, engorged live adult female ticks were maintained in containers in humid conditions at room temperature (~22-26°C) until eggs were laid.
Adult females were then removed, their DNA was extracted and PCR detection for rickettsiae was conducted. Eggs were maintained in the same containers with similar conditions and allowed to hatch into tick larvae. Live larvae were collected and their
DNA was extracted and PCR detection for rickettsiae performed.
4.2.5 Rickettsial Isolation by Cell Culture
Attempts at cell culture were made to isolate rickettsial organisms from ectoparasite samples using the methods detailed in section 2.4.2.
4.2.6 DNA Extraction
The protocol for DNA extraction of sampled ectoparasites is described in section 2.3.
DNA was extracted from whole ticks, tick organs and eggs.
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4.2.7 Rickettsial PCR Screening
PCR screening for rickettsial DNA from ectoparasite samples were conducted as described in section 2.6.1. Targets used for identification of rickettsiae in tick samples were the gltA and ompA genes.
4.2.8 DNA Sequencing
PCR products from 4.2.7 were sequenced as described in section 2.6.2.
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4.3 Results
4.3.1 Rickettsia gravesii prevalence in ticks
Five tick species were identified during the course of this study, of which one of the species, Haemaphysalis longicornis, is “newly” introduced into WA (28). The other four species collected were Amblyomma albolimbatum (a reptile tick common in WA),
Amblyomma triguttatum (kangaroo tick), Ixodes australiensis and Ixodes fecialis (Table
4.1). A total of 304 ticks were characterised in this study, of which A. triguttatum made up the majority with 187 isolates (Table 4.1). Of interest was the high number of A. triguttatum ticks collected from humans of which 76% (25/33) were found to be positive for SFG rickettsiae. No SFG rickettsiae were found in I. australiensis infesting humans, but observed in feral pigs and the Gilbert’s Pottoroo. Attempts at isolating R. gravesii in culture from ticks during the study were unsuccessful.
A. triguttatum had the highest prevalence for R. gravesii in WA with a prevalence of
75.3% and 56.8% in southwest WA and Barrow Island respectively (Table 4.2). The only H. longicornis tick examined was negative for rickettsiae.
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Table 4.1 – Tick species collected by animal host and their respective habitat.
Vertebrate W.A. SFG Tick Species Host Region Prevalence* Amblyomma Carpet Python Dwellingup 2/13 15.3% albolimbatum (Morelia spilota)
Bobtail lizard Kanyana 4/32 12.5% (Tiliqua rugosa) 6/45 13.3% Amblyomma Human Southwest WA 25/33 75.8% triguttatum (Homo sapiens) Barrow Island 12/22 54.5%
Feral Pig Dwellingup 6/7 85.7% (Sus scrofa) Serpentine 42/54 77.8% Mundaring 21/27 77.8%
Kangaroo Perth and surrounds 19/29 65.5% (Macropus rufus and Barrow Island 9/15 60.0% Macropus fuliginosus) 134/187 71.7% Haemaphysalis Feral Pig Dwellingup 0/1 0% longicornis (Sus scrofa) Ixodes Human Southwest WA 0/5 0 australiensis (Homo sapiens) Barrow Island 0/9 0
Feral Pig Dwellingup 9/15 60.0% (Sus scrofa) Serpentine 8/13 61.5%
Gilbert’s Pottoroo Two People’s Bay 4/21 19.0% (Potorous gilbertii) 21/63 33.3% Ixodes fecialis Gilbert’s Pottoroo Two People’s Bay 2/8 25% (Potorous gilbertii) *Amplicons detected for PCR targets ompA and gltA. All samples were positive for both ompA and gltA.
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Table 4.2 – Prevalence of R. gravesii in ticks by area/region.
Tick Species W.A. Region R. gravesii Prev. % Prevalence Amblyomma albolimbatum Dwellingup 2/13 15.4 Kanyana 4/32 12.5 Amblyomma triguttatum Southwest WA* 113/150 75.3 Barrow Island 21/37 56.8 Haemaphysalis longicornis Southwest WA* 0/1 0.0 Ixodes australiensis Barrow Island 0/9 0.0 Southwest WA* 17/33 51.5 Two People’s Bay 4/21 19.0 Ixodes fecialis Two People’s Bay 2/8 25.0 *Southwest WA include regions of Perth suburbs, Dwellingup, Mundaring, Serpentine, Kanyana and the “wheat-belt” between Perth and Albany.
4.3.2 Prevalence of R. gravesii in organs of A. triguttatum
Four individual A. triguttatum ticks’ organs (salivary glands, ovaries and gut) were screened for SFG gene targets (gltA and ompA). All the ticks were female and unengorged. All organs examined from three of the four ticks (salivary glands, ovaries and gut) had rickettsial DNA present. Sequencing of the PCR amplicons confirmed it to be R. gravesii.
4.3.3 Transovarial transmission of R. gravesii in A. triguttatum ticks
Four of the six engorged female Amblyomma triguttatum ticks laid eggs. The two ticks that did not lay any eggs died prematurely and they were both discarded and not analysed for R. gravesii as the focus of this experiment was to observe transovarial transmission. All four egg clusters successfully hatched and produced viable larvae. Out of the four larval cohorts, three were positive for SFG rickettsiae, which were determined to be R. gravesii.
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4.4 Discussion
The ectoparasites obtained for this study were made up mostly of ticks that commonly infest humans and large mammals such as kangaroos. Thus a priority was to collect ticks from the southwest region of WA where the majority of humans live and human activities occurred. Reptile ticks were also collected (A. albolimbatum), and although different from the ticks (B. hydrosauri) harbouring R. honei (292), their potential to harbour rickettsiae was demonstrated by Owen (206-208) and hence they were included in this study.
The predominant tick found on humans and other large mammals such as kangaroos and feral pigs was A. triguttatum. This tick demonstrated an affinity for larger mammals as host. As A. triguttatum had the highest prevalence of R. gravesii, the transmission of R. gravesii by A. triguttatum is possible. The apparent absence of locally acquired spotted fever illness in WA raises the question as to whether effective transmission occurs or whether infections are asymptomatic or mild and therefore undiagnosed. The hypothesis that rickettsial exposure of large mammals and humans occurs in the bush is tested in the next two chapters via a sero-prevalence study.
Rickettsia gravesii was observed in two additional tick species (I. australiensis and I. fecialis) neither described by Owen in her thesis, thus expanding the range of potential host vectors and reservoirs for novel Rickettsia spp. Owen only received a single individual I. australiensis and this would explain why the significant prevalence of R. gravesii in the tick species was not reported in her work (208).
The high prevalence of R. gravesii in A. triguttatum (75.3%) and I. australiensis
(51.5%) in southwest WA and the high affinity these ticks have for biting humans
87 compared to other ticks make them the most likely vectors for R. gravesii. However, spotted fever infections in humans in WA are rarely reported and the disease is not legally notifiable. This could be attributed to a low efficacy of transmission of the organism or the self-limiting nature of the illness, typical of spotted fevers, resulting in under-reporting or misdiagnosis.
R. gravesii was not observed in I. australiensis ticks obtained from humans. However on feral pigs, R. gravesii was found in 60.7% (17/28) of that tick species. This prevalence was similar to the prevalence of R. gravesii in A. triguttatum from humans
(67.3%; 37/55) and feral pigs (78.4%; 69/88). The high prevalence found in A. triguttatum from humans and feral pigs gives credence to the hypothesis that A. triguttatum is the main arthropod reservoir of R. gravesii. The absence of R. gravesii in
I. australiensis from humans but similarly high prevalence from feral pig to those of A. triguttatum points towards the possibility of I. australiensis being a vector although possibly not a reservoir. The R. gravesii observed in I. australiensis ticks could possibly have been acquired via horizontal transmission. The close proximity of feeding sites as observed on trapped feral pigs (Figure 4.1A) would allow for transfer of rickettsial organisms from an infected tick to a naive tick of another species. Recent studies have demonstrated the ability of animal hosts to act as amplifiers of Rickettsia spp. and to infect naive ticks without themselves becoming rickettsaemic (132, 233, 287).
The decision to use A. triguttatum to investigate the presence of R. gravesii in the three major organs (salivary glands, gut and ovaries) was mainly due to (i) the adult tick’s relative size compared to other tick species and (ii) the abundance of live A. triguttatum collected. The other tick species of comparative size to A. triguttatum was A. albolimbatum, however, most of the A. albolimbatum ticks were engorged or preserved
88 in 70% ethanol (v/v). Only 12 A. triguttatum ticks were deemed suitable for dissection in terms of size, viability and unfed state. Out of the twelve, four were dissected and
PCR of extracted DNA was performed. The results of the four “successful” dissections demonstrated that R. gravesii was present in all three organs of A. triguttatum analysed.
Its presence in the salivary glands implies transmission while feeding; in ovaries it would suggest transovarial and possibly sexual transmission while its presence in the gut points at maintenance of R. gravesii within the tick itself via transstadial transmission.
Questions may arise concerning cross-contamination between tick organs during removal and between different tick samples. Considerable care was taken to reduce the risk of contamination and the presence of a tick that was entirely negative for R. gravesii in all its organs suggests that cross contamination was absent. All ticks were dissected in one session using the same set of tools. The tools were cleaned between organs within the same tick and between tick samples. However the risk of contamination as a result of breakage of organs during removal was possible. To reduce this risk as much as possible, organs were removed in the sequence in which they were most accessible; salivary glands, mid gut and ovaries. Removal of organs in any other sequence would have caused the organ to be forced through other more superficial organs and thus increase the risk of contamination.
Positive results for SFG rickettsiae and subsequent sequencing of the product that demonstrated R. gravesii in both adult female and larval A. triguttatum indicates transovarial transmission of R. gravesii to viable offspring. For transovarial transmission to occur successfully the rickettsiae must remain viable through all stages of development. To prevent cross contamination from parent to larvae, parents were
89 removed after successful laying of the eggs and the eggs were allowed to hatch undisturbed.
The question still remains as to whether R. gravesii can be transmitted transstadially. Its presence in the mid-gut of A. triguttatum suggests that this is likely. However we were not able to attempt long term maintenance of ticks across their complete life cycle stages and subsequent generations due to laboratory restrictions.
The presence of Haemaphysalis longicornis on a feral pig in Western Australia suggests that H. longicornis infests animals in the WA bush. H. longicornis is of significant veterinary concern in QLD and NSW, and has been demonstrated to be a vector for
Theileria mutans (259, 262). First reported in WA by Besier and Wroth (28), the introduction of H. longicornis was purported to be from a domestic pet from the eastern states of Australia. Its potential to be a vector of rickettsiae is unknown, though work performed by Smith et al (286) has demonstrated its capacity to be an experimental vector for Coxiella burnetii. Even though our study showed a negative PCR result for
Rickettsia in a single H. longicornis, its potential as a vector for rickettsiae must not be overlooked due to the extremely small sample size.
Despite the current knowledge of rickettsiae in Australia, H. longicornis has yet to be attributed as a vector of rickettsiae in the eastern states. With its “recent” introduction into WA, the potential of H. longicornis to be infected with R. gravesii is worth investigating. No reports of R. gravesii have been made in tick populations in the eastern states. R. gravesii is abundant in WA, involving most tick populations so far investigated by Owen (208) and in this study. A future finding of R. gravesii in H.
90 longicornis would support the hypothesis that it may be transmitted to other tick species from a main invertebrate reservoir i.e. A. triguttatum.
With the evidence presented in this chapter, the tentative conclusion is that A. triguttatum is a strong candidate as the reservoir and main vector for R. gravesii.
Further investigation is warranted. We have established (i) the viability of A. triguttatum eggs and larvae with R. gravesii present, (ii) the high prevalence of R. gravesii in A. triguttatum populations from various parts of southwest WA and Barrow
Island especially in those ticks biting humans, and (iii) the presence of R. gravesii in the three main organs of A. triguttatum. The list of potential vectors of R. gravesii has also been expanded from four (208) to six (Table 4.3). Of potential concern is the very high prevalence of R. gravesii in A. triguttatum ticks found on humans. To investigate this further, a sero-prevalence and risk study in humans was designed and the results are presented in Chapter 6.
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Table 4.3 – Western Australian recorded tick species in the literature and those with potential as vectors of R. gravesii.
Genus Species R. gravesii Recorded in WA Reference
Ixodes eudyptidis ● (262) tasmani ● (262) hydromyidis ● (262) australiensis ☼ ● (262) fecialis ☼ ● (262) vestitus ● (262) myremecobii ● (262) antechini ▼ (208)
Haemaphysalis ratti ▼ ● (262) humerosa ● (262) lagostrophi ● (262) longicornis ● (28)
Boophilus microplus ● (262)
Rhipicephalus sanguineus ● (262)
Amblyomma calabyi ● (262) postoculatum ● (262) limbatum ▼ ● (262) australiense ● (262) triguttatum ☼ ▼ ● (262) albolimbatum ☼ ▼ ● (262)
Aponomma fimbriatum ● (262)
Bothriocroton hydrosauri ● (262)
Ornithodoros gurneyi ● (262)
● Published in the literature ▼ Described in Owen (2007) thesis ☼ Observed in this study
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5 - Serological Study of Spotted Fever Group Rickettsial Exposure in Feral Pigs (Sus scrofa) of South West WA
5.1 Introduction
There is evidence of spotted fever group rickettsiae in the Western Australian bush
(206, 208), however, as yet there is no published evidence of animal or human exposure to rickettsial organisms. This study was designed to investigate the prevalence of SFG rickettsial antibodies in a large animal species highly exposed to the main tick vector of a newly-recognised spotted fever group rickettsia in WA. Several criteria were formulated to select animal species suitable for this study. These included;
i) High exposure rate to the tick species that feed on large mammals in the
bush and also humans i.e. Amblyomma triguttatum and Ixodes
australiensis.
ii) Similar physiological constitution to humans and same approximate size
of organs.
iii) Animal’s limited ability to groom itself. This allows for maximum
duration of the ectoparasite attachment to the animal.
iv) Target population is sufficiently abundant for a statistically viable
sample size.
v) Part of a culling or research programme to allow for opportunistic
collection of samples thus reducing logistical and funding constraints.
vi) Conjugated species specific antibody readily available.
The only mammal that fulfilled all of the criteria was the feral pig (Sus scrofa) (see
Table 5.1). Samples were collected from feral pigs trapped within the Perth water catchment area as part of a Department of Environment and Conservation (DEC) 93 trapping and culling program. Animal ethics approval was obtained from the Murdoch
University Animal Ethics Committee, approval number R1155/05 (Appendix 9.1).
Table 5.1 – Four animal species considered for study and their fulfilment of five criteria.
Criteria Kangaroo Feral Pigs Feral Goats Feral Cats
I ● ● ●
II ●
III ● ●
IV ● ● ●
V ● ● ● ●
VI ● ● ●
5.2 Materials and Methods
5.2.1 Sample Collection
Collection of samples was conducted collaboratively in south-west Western Australia with the DEC. Traps were set by the DEC in water catchment areas in Serpentine,
Dwellingup and Mundaring (Figure 5.1) where feral pigs were known to inhabit in large numbers.
Ten to fifteen live traps were set up in each trapping area using apples as the bait
(Figure 5.2). Traps were checked and reset on a daily basis as part of the ethics requirement so as not to cause unnecessary distress to captured animals, and to release animals other than feral pigs. Feral pigs were euthanized/culled via a shotgun shot to the temple. Trapping and collection of samples was conducted between November 2006 94 and April 2007. Domestic pig samples were obtained from an abattoir in Gingin, approximately 20km north of Perth. Ectoparasites were removed from carcases as part of a post mortem protocol, and these were stored in alcohol until analysed at Murdoch
University (Chapter 4).
5.2.2 DNA Extraction
DNA was extracted from the liver and spleen of feral pigs. Organs were homogenized using sterile scalpel blades prior to start of extraction. Extraction was performed according to the manufacturer’s protocol (QIAamp® DNA Mini Kit) as outlined in section 2.3.
5.2.3 Rickettsial PCR Screening
PCR screening for rickettsial DNA from samples is described in section 2.6.1.
5.2.4 Microimmunofluorescence
Serological analysis of collected serum samples for SFG rickettsial antibodies using microimmunofluorescence is described in section 2.5.2.
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Trapping Areas
Water Catchment Areas
Figure 5.1 – Three main trapping sites named after the general region they encompass (i) Mundaring, (ii) Serpentine and (iii) Dwellingup. Water catchment areas appear as coloured regions and actual trapping areas are highlighted in transparent grey.
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A
B C
Figure 5.2 – Feral pig live trapping using wire cages with apples as bait (A). Multiple pigs were often trapped at the same time (B). Post mortem and ectoparasite sampling performed on site after euthanasia of feral pigs at trap sites (C).
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5.3 Results
Domestic pig sera (non-feral) used as controls were positive up to a titre of 1:64 (Table
5.2). Thus the cut off-point for positive sera among feral pig samples was determined to be at a titre of 1:128 for both SFG and TG antigens. Of the 149 feral pig samples tested,
70 (46.9%) had titres greater than 1:128 for SFG antigen. Of these, 48 (32.2%) were positive at 1:128 titre; 15 (10.1%) at 1:256 titre and 7 (4.7%) at 1:512 titre. Sixty-two
(41.6%) samples were positive at 1:128 (only) against TG antigen. All TG-positive sera were also SFG-positive.
Table 5.2 – Rickettsial sero-prevalence in feral and domestic pigs against SFG and TG antigens.
Antibody End Point Titre Total Type Grouping (%) +ve 1/32* 1/64* 1/128 1/256 1/512 1/1024
Dom. SFG 9 12 0 0 0 0 0 (0%) Pig (n=56) TG 7 9 0 0 0 0 0 (0%)
Feral SFG NT NT 48 15 7 0 70 (47%) Pig (n=149) TG NT NT 62 0 0 0 62 (42%)
* Positive results obtained at these dilutions were treated as non-specific antigen- antibody binding and regarded as false-positives, not indicative of SFG rickettsia exposure. NT – Not Tested
Titration of positive feral pig sera against 4 different SFG rickettsia antigens was performed to determine if any had a higher titre against a particular SFG rickettsial species (Table 5.3). The four rickettsial species chosen were R. australis, R. honei, R. gravesii and R. rickettsii.
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Table 5.3 – Titration of feral pig sera against Rickettsia antigens.
Antigen 1/128 1/256 1/512 1/1024 Total
R. australis 62 3 0 0 65
R. gravesii sp. nov. 48 15 7 0 70
R. honei 60 4 0 0 64
R. rickettsii 59 1 0 0 60
Prevalence was higher in adult feral pigs (≥12 months) with an average of 66.7% compared to the young (<12 months), with a prevalence of 33.7% (p<0.02) (Table 5.4).
The higher prevalence in older feral pigs was observed in feral pigs from Dwellingup,
Serpentine (approximately 1.5 times higher) and Mundaring (approximately 3.5 times higher) (Table 5.4).
Table 5.4 – Sero-prevalence in feral pigs by age and location.
Age Group Mundaring Dwellingup Serpentine Total
Adult 6/7 18/24 16/29 40/60* (>12 mths) 86% 75% 55% 67%
Young 4/16 9/26 17/47 30/89* (<12 mths) 25% 35% 36% 34%
10/23 27/50 33/76 70/149 Total 44% 54% 43% 47%
*p value < 0.02
PCR conducted on DNA extracts from 129 liver and spleen samples revealed 4 positives (3.1% prevalence) for rickettsial DNA (gltA) in the liver only. A summary of the information available for each sample that was PCR positive is shown in Table 5.5.
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Table 5.5 – Summary of four feral pigs positive by rickettsial citrate synthase gltA gene PCR.
Sample No. Age Gender Area Trapped SFG Serology
1 Adult (~3 years) Male Mundaring Negative
2 Young (4-5 months) Female Dwellingup Negative
3 Young (6-8 months) Male Serpentine Positive*
4 Adult (>3 years) Male Serpentine Negative
*Serology result was equivocal with titres at 1/128 to R. gravesii and R. honei and negative to R. australis and R. rickettsii antigens.
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5.4 Discussion
The high rickettsial seroprevalence among feral pigs in and around the water catchment areas of Perth demonstrates a high rate of exposure to rickettsial organisms. This may be due to ticks in these regions harbouring a high prevalence of the novel rickettsia,
Rickettsia gravesii sp. nov. (172). A high cut-off titre (≥1:128) was based on the control group (domestic pigs never exposed to ticks in the bush) which exhibited no reactivity at >1:128 dilution. This is similar to the cut-off used for human diagnostics. In spite of this relatively high cut-off, a sero-prevalence of 47% at a cut-off titre of 1/128 was observed. Cross-reactivity between species of the SFG is well established in the literature and hence makes it difficult to differentiate the infecting rickettsia from other members of the SFG. As no other SFG rickettsiae had been found in the water catchment areas and higher titres were observed against R. gravesii antigens, it is most likely that exposure to R. gravesii, was responsible for the high seroprevalence in the feral pigs.
Typhus group antibodies were observed in 41.6% of the test group however the titre was not above 1/128. All TG-positive sera were SFG-positive. Also, no typhus group rickettsia or R. felis have been reported from ectoparasites in the WA bush. Thus it was concluded that the typhus group antibodies observed were a result of cross-reactivity between the SFG and TG.
Adult feral pigs in all three locations had rickettsial antibodies prevalence of at least 1.5 times higher than the young. Mundaring adult feral pigs had a prevalence of 85%, compared to only 25% among the young. The large difference in prevalence of rickettsial antibodies observed between adult and young feral pigs suggests that as the feral pig population ages, more pigs are exposed to rickettsial organisms probably by
101 increasing risk of exposure to infected ticks. No other studies have reported this previously, as they have tended to focus on general prevalence of rickettsial antibodies to determine exposure levels. A similar trend was observed in the other two trapping sites. However, the numbers in each area are too low to provide a significant result on their own. Anti-SFG antibodies when compared between young and adult feral pigs as a whole showed a doubling in prevalence as a feral pig ages. Statistical analysis of the total prevalence between the age groups using the student t-test demonstrated a significant result with p < 0.02 (Table 5.4).
Various organs have been reported to harbour rickettsiae such as the brain (2), kidney
(55), lymph nodes (281), liver (178) and spleen (278). The organs sampled (liver and spleen) and tested for rickettsial DNA was chosen due to ease of access at post-mortem, a major role played in the circulatory system and as such is the most likely location of infections. The low prevalence of 3.1% (4/129) observed in the liver indicates that feral pigs probably act as an incidental host rather than a reservoir for rickettsiae in WA. No rickettsiae were observed in the spleen indicating possible rapid degradation of foreign
DNA in the pigs’ spleen. The presence of SFG rickettsia in the liver (but not the spleen) is difficult to explain as both organs are major components of the reticulo-endothelial system, which readily clears microbes from the blood system.
Only one of the feral pigs positive for rickettsial DNA was also positive for anti- rickettsial antibodies. However, it should be noted that the antibody levels were only
1/128 for two of the rickettsial organisms tested against (R. gravesii and R. honei). This fits with the hypothesis that antibodies raised as a result of natural infection are only observed weeks after the initial exposure as the immune system requires time to produce antibodies to a detectable level. As the levels of antibodies rise, the amount of
102 rickettsial organisms in the host is expected to drop. The level of antibodies in the animal/human host is expected to stay at detectable levels for weeks if not months after the organism has been cleared from the host. Although we were unable to perform a longitudinal study on a controlled population, the evidence from this study suggests that a high number of feral pigs are exposed to rickettsial organisms but a very low number have the organism present in their organs at any point in time. Thus the timeframe for detecting rickettsia in these animals may be very small.
Recent reports in the literature have demonstrated the transfer of rickettsial organisms from one tick to another via horizontal transmission through an animal host without the animal becoming rickettsaemic (132, 233, 287). With the low prevalence of rickettsiae observed in feral pig organs, compared to the high sero-prevalence levels, it is possible that feral pigs do not act as a reservoir for rickettsia but can potentially be an amplifier for rickettsial organisms in the WA bush without harbouring the organism itself. The low prevalence of rickettsial DNA in feral pigs, in spite of high exposure demonstrated by the high sero-prevalence, suggests that the maintenance of rickettsial organisms in the feral pig is transitional. The feral pig thus fills the role as an amplifier for rickettsiae by exposing uninfected ticks to rickettsiae it is harbouring, thus potentially increasing the prevalence of the rickettsiae in tick populations infesting the pigs. The ticks affected may or may not be effective vectors and/or reservoir of rickettsial organisms.
This study represents the first investigation of rickettsial sero-prevalence in a wild animal population in Australia. Other sero-prevalence studies investigating Rickettsia exposure in Australia has only looked at dogs and cats as companion animals (21, 138).
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6 - Rickettsial Exposure in High Risk Occupational and Recreational Groups Western Australia
6.1 Introduction
The discovery of two novel spotted fever group rickettsiae (R. antechini and R. gravesii) by Owen et al. (206, 207) provides a new focus on endemic rickettsial organisms and their public health importance in Western Australia. Based on the known epidemiology of rickettsial infections, there is the potential for both organisms to affect the health of both humans and animals. In view of the potential risk posed by such organisms, an investigation to determine their geographical range and other ecological information was undertaken.
Earlier chapters have investigated the potential for feral pigs to act as rickettsial reservoirs, and have discussed the diversity of vectors that may harbour and transmit rickettsial organisms in the bush. With various industries located in these natural environments, including those related to mining and agriculture, there are concerns over the risk of exposure to rickettsial organisms in an occupational setting. Additionally, the increasing number of WA residents pursuing recreational activities in the bush increases the risk of tick bites and hence possibly exposure to vector-borne rickettsioses. In order to the address the potential risks through occupational and recreational activities, this study was designed to expand on Owen’s (208) tick exposure study by including the following groups:
(i) Employees of various organisations and companies working on Barrow
Island
(ii) Employees of Whiteman Park wildlife sanctuary in Perth
(iii) Members of a rogaining organisation – an outdoor recreational sport.
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Barrow Island lies 50km off the northwest coast of the Pilbara region of WA. The closest major town is Onslow. The island is classified as a class A nature reserve, and as a result, has very strict quarantine measures. Oil and gas extraction is however allowed under strictly managed conditions. The human population on the island, at the time of this study, was approximately 200, most of them involved in drilling and mining work.
Visits to the island are restricted to employees of the oil company and Department of
Environment and Conservation (DEC) staff.
Whiteman Park (wildlife sanctuary) is located within the northern Perth suburbs and has a large wildlife population within its boundaries. The research interest in Whiteman park was generated by reports of illnesses related to tick bites among their employees.
They reported a high incidence of tick bites (mainly A. triguttatum) on site, probably related to the large number of free roaming macropods.
Rogaining is an outdoor recreational sport that was first developed in 1947 by students of Melbourne University. The sport itself mainly resembles orienteering, with the main difference being a larger cross-country element. Rogaining events may last between 6 to
24 hours with participants travelling through the bush around a designated course. It is not uncommon for groups to stay out in the bush for the entire duration of the event, including sleeping under trees or in bushes with minimal protection from arthropods.
Rogaining events occur within a 200km range of the outskirts of Perth. All rogaining volunteers who took part in this study were members of the Western Australia
Rogaining Association (WARA).
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For each of these three distinctive exposure groups, sera were examined for SFG rickettsia. A control (urban) group was recruited from Murdoch University for comparison.
6.2 Materials and Methods
6.2.1 Study Design
The study was designed as a (i) cross-sectional analysis/serosurvey with an external comparator group; in conjunction with (ii) a prospective component for some subjects to assess rates of seroconversion and seroreversion. The serological survey involved the collection of up to two blood samples from each volunteer: one at the time of recruitment, and (for a subset) a second sample 10-14 months later.
Collection of blood samples coincided with the completion of a questionnaire enquiring about the participant’s outdoor activities, symptoms associated with rickettsioses, other risk factors and demographic details.
Samples were collected from volunteers from three main groups: (i) Occupational
Group (OG), (ii) Recreational Group (RG) and (iii) Control Group (CG). Volunteers from two organisations contributed samples as part of the OG group cohort: staff working on Barrow Island and at Whiteman Park (WP) in Perth. The RG cohort was made up of members of the Western Australian Rogaining Association (WARA). Staff and students of Murdoch University were recruited as a control group (CG).
There were no strict recruitment criteria for volunteers from the OG and RG cohorts have no strict criteria other than being employed in the areas under study or as a
106 member of their respective organisations and over the age of 18 years at the time of sampling. More stringent criteria were set for the CG baseline control group to ensure that they were of comparable age and had been WA residents for a reasonable period of time. Criteria for their admission into the study were:
1) Minimum 18 years old at time of recruitment.
2) Residing within the Perth metropolitan area.
3) Not having resided outside of WA for more than 6 months cumulatively within
the last 5 years prior to sample collection.
Follow-up sample collections were performed 10-14 months after the first collection, although only a limited number of volunteers from WP and WARA were available.
Human ethics approval was obtained from the Fremantle Hospital and Murdoch
University Human Ethics Committees (see Appendix 9.1).
6.2.2 Blood Collection
Eight to ten millilitre blood samples were collected from the study participants between
2006 and 2009. After clotting, blood samples were centrifuged and between two-three millilitres of serum were recovered from each sample. Where possible, blood was collected on site together with questionnaire completion, with the exception of the first group of rogainers, who had their serum collected by PathWest and questionnaires completed by telephone. Subsequently, the follow-up collection for rogainers involved on-site collection and questionnaire completion similar to all study groups. Barrow
Island workers’ blood samples were collected by a registered nurse at the onsite medical centre. Whiteman Park employees, rogainers and Murdoch staff and students had their blood collected onsite by a qualified phlebotomist.
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6.2.3 Microimmunofluorescence
Serological analysis of human serum samples for rickettsial antibodies via microimmunofluorescence was performed as described in section 2.5.2. Titration was performed on serum samples that were positive to SFG and TG antigens at the screening dilution of 1/128. SFG positive samples were titrated against total SFG antigens containing equal concentrations of R. honei, R. australis and R. gravesii antigen. TG positive samples were titrated against R. typhi. Control sera and antigens were provided by the Australian Rickettsial Reference Laboratory, Geelong.
6.2.4 Questionnaire Design
Questionnaires were designed to establish whether patterns of tick exposure in the bush including activities that involved a high-risk of tick exposure, duration of such exposures and the frequency of tick bites during such activities. Information on presentation of any symptoms or signs that may have been attributable to a rickettsiosis, was also obtained. Other information collected in the questionnaire included travel and medical histories and demographic information (Questionnaires are included in
Appendix 9.4, 9.6 and 9.8).
A second questionnaire was also designed for follow-up with the volunteers 10-14 months after the first questionnaire to coincide with the second blood sample. The focus of the second questionnaire was to determine any changes to the participant’s data from the first questionnaire and the development of any rickettsiosis-like symptoms following activities in the bush during the (approximately) one year time interval
(Appendix 9.5 and 9.7).
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6.2.5 Questionnaire Implementation
All potential volunteers were given either a presentation or an informal briefing on the study giving them an overview of rickettsiae and rickettsioses, describing rickettsial organisms in Western Australia and discussion of the potential public health impact of such organisms. Details were provided on the procedures to be undertaken i.e. a blood sample and questionnaire at recruitment, and a follow up blood sample and questionnaire 10-14 months after. Volunteers were then asked to fill out the consent form before their participation in the study proceeded (Appendix 9.3).
6.2.6 Statistical Analysis
Using the data gathered from the serological analysis and questionnaire, a statistical analysis was performed on the relationship between the predictive variables and major outcome variable (seropositive to SFG Rickettsia sp. at 1/128). Descriptive statistics were generated for occupational and recreational groups, other activity-related descriptors and the age and gender of the study population. To determine the risk of infection in the WA bush, final risk estimates were generated using logistic regression.
The outcomes of the survey conducted on participants were converted to binary data
(“Yes” or “No” answers) and odds ratios were calculated from the collated information.
Odds ratios were used to compare various exposure groups with reference to the control
(baseline) group. Other risk factors were also assessed, such as bush activities in other
States and overseas and the presence of major health issues. Both crude and adjusted risk estimates were calculated by statistical modelling using Stata/MP 10 (USA).
Odds ratios were chosen to demonstrate the relative risk of each group of being infected by an SFG rickettsia in the WA bush. Odds ratio analysis was adjusted for age and gender. The choice of odds ratio was made because it allows examining of the effects of
109 other variables on a particular relationship. Presentation of odds ratio is in the form of probability (35). The calculation of odds ratio in this study was by logistic regression with adjustment for age and gender. The control group was used as a baseline (odds ratio 1.0).
6.3 Results
6.3.1 Serology (IFA) Results
A total of 60 sera were collected for the base-line control group, 104 from rogainers, 67 from Barrow Island workers and 12 from Whiteman Park staff (Table 6.1). Positives were regarded as SFG titres of 1:128 as determined by MIF. Whiteman Park employees were observed to have the highest rickettsial antibody prevalence at 50% (6/12) (but the sample size was small), with Barrow Island workers showing a comparative result
(44.8%; 30/67). The seroprevalence for rogainers was 23.1% (24/104), whereas that of the baseline “control” group sero-prevalence was 1.7% (1/60).
Table 6.1 – Seroprevalence for SFG rickettsia from primary and secondary blood sampling of humans residing in WA with IF titres to SFG of 1:128.
First Sample Follow-Up Sample Group Collected Positive Prevalence Collected Positive Prevalence Control Group 60 1 1.7% - - - (Baseline)
Recreational Group 104 24 23.1% 62 18 29% (Rogainers)
Occupational Group 67 30 44.8% - - - (Barrow Island)
Occupational Group 12 6 50% 8 3 37.5% (Whiteman Park)
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A follow-up blood sample was collected from the rogainers (n=62) and Whiteman Park employees only (n=8) (Table 6.1). Follow up samples were not possible for Barrow
Island staff due to the transient nature of employment, the difficulties of getting to the island and the wide range of areas that the fly-in fly-out workers came from. The seroprevalence for rogainers was slightly higher at 10-14 months after the first blood test (29%; 18/62), whereas the Whiteman Park employees’ seroprevalence had fallen to
37.5% (3/8).
Seroconversion and seroreversion rates were calculated using the formulae below;
i) Seroconversion Rate = (t=0 Negative, t=1 Positive)/(t=1 Positive) X 100
ii) Seroreversion Rate = (t=0 Positive, t=1 Negative)/(t=1 Negative) X 100
Seroconversion and seroreversion rates for the rogainers were 15.5% and 35.5% respectively (Table 6.2). Whiteman park employees seroconversion and seroreversion rates were 0% and 50% respectively (Table 6.3).
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Table 6.2 – Rogainers’ seroconversion and seroreversion results over a 10-14 month period (t=0 is serum collected at first collection; t=1 is serum collected at follow-up at 10-14 months; N is number of volunteers).
t=0 t=1 N
- - 38
- + 7*
+ - 6^
+ + 11
Total 62 * Seroconversion ^ Seroreversion
Table 6.3 – Whiteman park employees’ seroconversion and seroreversion results over a 10-14 month period (t=0 is serum collected at first collection; t=1 is serum collected at follow-up at 10-14 months; N is number of volunteers). t=0 t=1 N
- - 2
- + 0*
+ - 3^
+ + 3
Total 8 * Seroconversion ^ Seroreversion
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6.3.2 Descriptive Epidemiology
There were approximately 1500 members in the Western Australian Rogaining
Association. A total of 135 questionnaires were completed, giving a participation rate of
9.0%. Questionnaire completion rates for occupational groups were 29.5% (59/200) for
Barrow Island employees and 32.1% (9/28) for Whiteman Park employees. Follow-up rates of 34.8% (47/135) and 44.4% (4/9) were obtained with the rogainers and
Whiteman park employees respectively. We were unable to follow-up on Barrow Island employees due to logistical constraints documented above. The populations surveyed are summarised in Table 6.4.
From the questionnaire results obtained, the two groups that spent most time in the bush has a large male majority, with 93% for Barrow Island employees and 66.7% for
Whiteman park employees. Gender distribution in participants from the control group and rogainers were similar (Table 6.4). The majority of the participants in the recreational and occupational groups were above the age of 40 (median age of each group was Barrow Island employees=42, rogainers=50 and Whiteman park employees
=51). In the control group, the median age was 24. There were further variations in the median age male and female participants within each group. Median age for the control group and the rogainers were fairly similar between male and female participants.
Wider variations in age were observed in the Barrow Island workers (male=42, female=28) and Whiteman park employees (male=53, female=38).
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Table 6.4 – Descriptive categories of populations based on questionnaire data.
Whiteman Whiteman Control Barrow Island Rogainers Park Park Employees Rogainers Employees Categories Baseline Follow-Up Employees Follow-Up # % # % # % # % # % # % Gender Male 20 42.6 55 93.0 66 48.9 25 53.2 6 66.7 3 75.0 Female 27 57.4 4 7.0 69 51.1 22 46.8 3 33.3 1 25.0 Age groups <25 years old 25 53.2 8 13.6 1 0.7 0 0 1 11.1 1 25.0 25 to 50 years old 21 44.7 37 62.7 75 55.6 24 51.1 3 33.3 2 50.0 >50 years old 1 2.1 14 23.7 59 43.7 23 48.9 5 55.6 1 25.0 Recreational time in bush in WA Once a month or more 18 38.3 29 49.2 135 100 38 100 7 87.8 2 50.0 Rarely (<2 times/year) 29 61.7 30 50.8 0 0 0 0 2 22.2 2 50.0 Occupational time in bush in WA Once a month or more 2 4.3 50 84.7 11 8.1 3 6.4 7 87.8 3 75.0 Rarely (<2 times/year) 45 95.7 9 15.3 124 91.9 44 93.6 2 22.2 1 25.0 Time in bush in other Australian states More than once per year 4 8.5 15 25.4 14 10.4 0 0 1 11.1 0 0 Once per year or less 43 91.5 44 74.6 120 89.6 47 100 8 88.9 4 100 Time in bush overseas More than once per year 2 4.3 7 11.9 4 3.0 0 0 0 0 0 0 Once per year or less 45 95.7 52 86.4 131 97.0 47 100 9 100 4 100
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Table 6.4 – continued.
Whiteman Whiteman Control Barrow Island Rogainers Park Park Employees Rogainers Employees Categories Baseline Follow-Up Employees Follow-Up # % # % # % # % # % # % Tick bite frequency Most or every time in bush 0 0 1 1.7 14 10.4 2 4.3 2 22.2 0 0 Often (25-75%) 0 0 8 13.6 41 30.4 9 19.1 1 11.1 2 50.0 Occasionally 2 4.3 33 55.9 60 44.4 16 34.0 5 55.6 2 50.0 Never 45 95.7 13 22.0 14 10.4 17 36.2 1 11.1 0 0 Don’t know 0 0 4 6.8 6 4.4 3 6.4 0 0 0 0 Symptoms following tick bite None or one symptom 50 84.7 120 88.9 42 89.4 6 66.7 1 25.0 Rash unrelated to bite 3 5.1 3 2.2 2 4.3 1 11.1 1 25.0 Two symptoms of rash, fever/chills or headache NA 0 0 5 3.7 0 0 0 0 0 0 Any two symptoms of rash, fever, headache, aching, 6 10.2 7 5.2 3 6.4 2 22.2 2 50.0 tiredness, scab, shortness of breathe Symptoms with no bite history None or one symptom 42 89.4 53 89.8 124 91.9 45 95.7 6 66.7 3 75.0 Rash unrelated to bite 1 2.1 1 1.7 5 3.7 0 0 0 0 0 0 Two symptoms of rash, fever/chills or headache 2 4.3 0 0 1 0.7 1 2.1 2 22.2 0 0 Any two symptoms of rash, fever, headache, aching, 2 4.3 5 8.5 5 3.7 1 2.1 1 11.1 1 25.0 tiredness, scab, shortness of breathe Medical conditions None 47 79.6 98 72.6 37 78.7 8 88.9 2 50.0 NA Major (e.g. heart, asthma, diabetes) 12 20.4 37 27.4 10 21.3 1 11.1 2 50.0
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All the rogainers who participated in the study reported that they spent time in the bush at least once a month for recreational purposes. This contrasted with the other groups: only 49.2% of Barrow Island workers and 38.3% of the control group reported spending time in the bush recreationally. However, for activities in the bush due to occupational activities, the workers on Barrow Island rated the highest with 84.7% spending time in the bush at least once a month, as compared to 8.1% of rogainers and 95.7% of the control group. Whiteman park employees spent equal time on average in the bush while at work and recreationally (87.8%).
Rogainers (40.7%) and Whiteman park employees (33.3%) reported the highest rates of tick bites (25-100% of their time in the bush) occurring when they were in the bush for both occupational and recreational activities. Only 15.3% of Barrow Island workers reported tick bites while in the bush. Reports of symptoms as a result of tick bites that may be linked to a rickettsiosis were highest among Whiteman Park staff (22.2%), followed by the workers on Barrow Island (10.2%), and lastly the rogainers (8.9%).
However very similar rates were also observed for rickettsiosis-like symptoms not linked to tick bites.
6.3.3 Odds Ratio (OR) Analysis
Logistic regression and calculation of odds ratios demonstrated that there were significant associations between age, gender, work related bush activities and specific group classifications (occupational and recreational) with positive rickettsial serology.
No significant associations were observed between variables such as recreational activity, tick bite frequency and symptoms related and/or unrelated to tick bites with positive serology (Table 6.5).
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Table 6.5 – Risk estimates for seropositivity to SFG Rickettsia sp. (adjusted for age and gender). N Unadjusted 95% CI Adjusted 95% CI (Sero +ve/ Odds Ratio (p-value) Odds Ratio (p-value) Sero -ve) Demographic Variables Age Strata
<25y (baseline) 4/30 1.0 - 1.0 -
0.93-9.00 0.75-7.64 25-50y 27/70 2.89 2.39 (p=0.066) (p=0.14) 1.14-12.62 0.95-11.63 >50y 15/30 3.75 3.33 (p=0.033) (p=0.059) Gender Female 9/56 1.0 - 1.0 - (baseline) 1.39-6.97 1.29-6.57 Male 37/74 3.11 2.91 (p=0.006) (p=0.010) Reported Work in W.A. Bushland
No 18/95 1.0 - 1.0 -
2.08-8.59 1.63-7.77 Yes 28/35 4.22 3.56 (p<0.001) (p=0.001) Specific Occupational and Recreational Classifications Control Group 1/46 1.0 - 1.0 - (baseline) Rogainers 1.73-108.49 1.38-142.07 14/47 13.70 14.02 (recreational) (p=0.013) (p=0.025) Workers on 5.02-300.34 3.48-298.80 27/32 38.81 32.24 Barrow Is (p<0.001) (p=0.002) Wildlife park 3.41-396.72 2.34-1109.27 4/5 36.80 50.89 employees (p=0.003) (p=0.012) Frequency of Recreational Activities in W.A. Bushland <1x per month 14/47 1.0 - 1.0 - (baseline) 0.63-2.67 0.58-2.71 ≥1x per month 32/83 1.29 1.24 (p=0.48) (p=0.57) Experienced Frequent Tick Bites in W.A. Bushland
No 32/99 1.0 - 1.0 -
0.65-3.15 0.45-2.53 Yes 12/26 1.43 1.07 (p=0.38) (p=0.88) Reports of Systemic Symptoms Specifically Following Tick Bites˄
No 37/69 1.0 - 1.0 -
0.41-3.76 0.48-4.78 Yes 6/9 1.24 1.51 (p=0.70) (p=0.48) Reports of Any Systemic Symptoms (Not Necessarily Following Tick Bites)˄
No 41/76 1.0 - 1.0 -
0.26-3.26 0.27-3.46 Yes 4/8 0.93 0.96 (p=0.91) (p=0.95) ˄Only for occupational and recreational groups
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6.3.4 Spotted Fever Group Rickettsiosis in a WA Patient
While two workers on Barrow Island presented to a doctor with possible symptoms of a rickettsiosis, only one definitive case of rickettsiosis was reported during the course of the study. The diagnosis was made by Dr John Dyer at the Fremantle Hospital in
Western Australia in May 2006. The patient presented with clinical symptoms of a rickettsiosis (fever, malaise, myalgia, rash and abnormal liver function) and had positive serology to SFG rickettsial antibodies. PCR analysis on a biopsy of the patient’s eschar was positive for the Rickettsia sp. citrate synthase gene. However, PCR on the patient’s
EDTA blood sample was negative and this was mainly attributed to the early prescription of doxycycline. The patient was hiking the Bibbulmun track, which stretches for more than 500km between Albany and Perth. The patient had spent more than 3 weeks in the bush and had become infested with ticks. Shortly after returning, he fell ill with fever, chills, rash, hepatitis and eschar (Figure 6.1) at a tick bite site. He was admitted to Fremantle Hospital and given a regimen of doxycycline for suspected rickettsial disease. The patient’s clinical symptoms cleared up within 3 days of starting the treatment. A tick collected from the patient, although not attached anywhere close to the eschar, was submitted for analysis as part of this study. The tick was identified as belonging to the genus Ixodes, although speciation was not possible due to damage to the mouth parts (palp articles) during removal from the patient. PCR analysis of the tick was negative for Rickettsia sp.
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Figure 6.1 – Eschar and swelling on patient’s leg as a result of a tick bite (Courtesy of Dr John Dyer, Fremantle Hospital).
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6.4 Discussion
6.4.1 Sero-positivity and its Link to Bush Activity
In this study, it was noted that the unadjusted OR for sero-positivity to SFG rickettsia was significant for participants over the age of 50 (unadjusted OR=3.75, 95% CI=1.14-
12.62). However, after adjustment for gender, the risk estimate became non-significant
(Table 6.5). An age-related effect for this cohort was anticipated, given the association between age and sero-positivity, as observed in many infectious disorders, because of cumulative seroconversion over time. It is also notable that a significant proportion of participants from both the occupational and recreational groups were above the age of
50, so age was a factor in predicting the likelihood of exposure to ticks (Table 6.4).
With respect to gender, males were also observed to be significantly more at risk
(OR=2.91, 95% CI=1.29-6.57). This occurrence can be attributed to the higher number of males in occupational roles that puts them at risk in the bush on Barrow Island (93%) and Whiteman Park (66.7%). As hypothesized, there was a significant risk to participants who reported having an occupational role in the bush (OR=3.56, 95%
CI=1.63-7.77). For specific occupational and recreational groups, the risk was most significant in the Whiteman park staff (OR=50.89, 95% CI=2.34-1109.27). The wide confidence intervals were a result of the small sample size in this occupational category.
Workers on Barrow Island were also significantly at risk with an OR of 32.24 (95%
CI=3.48-298.80), as were rogainers, with an OR of 14.02 (95% CI=1.38-142.07) (Table
6.5).
The high rate of sero-prevalence to SFG Rickettsia sp. was unexpected among the groups studied in WA. It was acknowledged that people performing high risk activities would be exposed to ticks and rickettsiae, but the high sero-prevalence points towards a higher exposure rate/transmission efficiency than that seen elsewhere in high risk
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“healthy” (asymptomatic) populations (57, 232). In Australia, comparably high prevalences were more commonly observed in patients with overt symptoms of rickettsial infection (305). Other studies have demonstrated high seroprevalences to rickettsial exposure including; R. africae in Guadeloupe (French West Indies) (217), R. japonica in Korea (140) and R. helvetica in Denmark (193). The low sero-prevalence observed in the current control group was similar to that observed in other studies of a general population, where prevalences of less than 5% have been reported (112).
The mere presence of an endemic rickettsia in an environment does not necessarily mean that it will be transmitted to humans and hence a high level of seropositivity in humans cannot be assumed. The WA data reflects a high rate of rickettsial transmission and human infection in the WA bush. However, clinical disease from rickettsia exposure appears to be rare. Thus a rickettsiosis caused by SFG infection in WA may be mild or asymptomatic. As no other rickettsiae have been found in ticks that commonly bite humans in WA (A. triguttatum and I. australiensis) it is postulated that a significant proportion of seropositives may be due to exposure to R. gravesii, the most common rickettsia identified in ticks in the state.
A previous WA study investigating A. triguttatum tick infestations of humans failed to identify a clear link between any rickettsiosis-like illness and tick bites (223). Only delayed hypersensitivity reactions to tick bites (24-48 hours) and secondary bacterial infections were reported. Considering that symptoms of rickettsiosis usually present 7-
14 days after infection, any presenting symptoms may not have been attributed to the earlier tick bites. Some of the reported symptoms associated with tick bites may be due to an allergic reaction which is not uncommonly recorded in Australia (42, 102).
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The lack of a significant association between self-reported tick bites and seropositivity is unlikely to be an accurate reflection of the true exposure profile. The majority of SFG rickettsioses worldwide are transmitted by a tick bite, and therefore the exposure event must have occurred at some point. The other known SFG rickettsia endemic to WA, R. felis, presents serologically as a TG rickettsiae and is transmitted by fleas (1, 332). It is more likely that the seroprevalence observed in bush-active groups in south-west WA is as a result of exposure to R. gravesii. However, this hypothesis needs confirmation through the isolation of R. gravesii from a patient diagnosed with SFG rickettsiosis.
Other similar cases to the one reported in Section 6.3.4 have been reported in south- west WA, but with little to no clinical or microbiological follow up to confirm rickettsial aetiology (John Dyer, pers. comm.). The “lack” of records in WA pertaining to rickettsial disease may relate to factors such as: (i) the treatment of patients with antibiotics without undertaking pathology testing; and (ii) the endemic SFG rickettsiosis may be mainly asymptomatic or very mild and thus may not come to medical attention.
6.4.2 Potential Causes of Bias and Confounding Factors in the Study
There were limitations to the study that may have introduced bias to the observed results. In this discussion, these are separated into selection bias and information bias.
Selection Bias
All participants in the study were volunteers and thus may have particular characteristics that could have influenced the outcome of the result. People who volunteer in community-based studies have demographic attributes that may introduce bias to the study and the interpretation of results (100). Although characteristics of volunteers vary between studies, it has often been observed that healthier individuals in
122 the community are more inclined to volunteer. This phenomenon is known as the
Healthy Volunteer Effect (HVE). HVE has been demonstrated in several studies (12,
96, 174) and is acknowledged as an important source of bias in epidemiological research. HVE may explain the lack of any link to adverse symptoms linked to rickettsiosis after a tick bite, given that healthier individuals may be less likely to show severe symptoms of infection.
There were also different gender numbers in the study across the study groups. There were slightly more females in the rogainers (57.4%) and control (51.1%) groups, whereas respondents were overwhelmingly more likely to be male in the Barrow Island
(93%) and Whiteman park (66.7%) groups. The slight bias towards females in the rogainers and control groups was unexpected as both populations were approximately equal, whereas the overwhelming bias towards males in the Barrow Island and
Whiteman park groups was expected due to the male-oriented nature of the work performed at the two locations.
High participation rates were observed with the control, Barrow Island and Whiteman park groups because of “on the spot” recruitment and immediate follow-up blood collection and questionnaire completion. The rogainers underwent a slightly different method of recruitment, in which volunteers were recruited and were followed up with a phone interview and blood collection at a pathology centre close to their residence. As a result, retention rate was low, with volunteers pulling out or unable to provide a blood sample due to problems faced trying to visit or liaise with the pathology laboratory.
Attempts to rectify this problem were made by subsequent collection of blood samples and completion of questionnaires at rogaine sites, although this option was logistically
123 more demanding than collection of samples with full pathology facilities (as occurred for the other groups).
Information Bias
The surveys conducted required participants to recall information going as far back as five years from the time of the questionnaire application. The accuracy of the information may not be reliable and may be represented as an over-reporting (or under- reporting) of information deemed useful (or unrelated) to the study. For example, mild symptoms following tick bites may not have been accurately recalled or may not have been linked to the exposure event.
Confounding Factors
There were several identified possible confounding factors such as age, gender, location of home (bush, suburb, rural), keeping of pets and living in close proximity to wildlife that may have affected the patterns of exposure or clinical outcomes. Of these potential risk factors, age and gender were identified as the most significant confounding factors and were thus adjusted for in the calculation of the odds ratios.
Low Participant Numbers
The relatively modest sample sizes in some groups were related to the relatively small baseline populations in some occupations (Section 6.3.2). The low numbers resulted in wide CI intervals for a number of the risk estimates.
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6.4.3 Public Health Implications of the Study
The information collected from this study demonstrates the potential risk that novel
SFG rickettsia, R. gravesii, may pose to humans active in the WA bush. The high rate of infection/exposure observed among high-risk populations reinforces the hypothesis that there are endemic SFG rickettsiae in WA, and that the most likely agent is R. gravesii.
The tick is the main vector of SFG rickettsial transmission similar to other rickettsial diseases. Prolonged attachment of ticks to their host have been demonstrated to increase the rate of transmission of pathogenic bacteria (152, 230). Although there is currently no data on the transmission of Rickettsia sp. in WA, evidence suggests that attachment for longer than 24 hours would result in a high rate of rickettsial transmission.
Amblyomma triguttatum had been identified previously in this study as the main reservoir and vector of R. gravesii. The affinity of this tick for large vertebrate mammals in the bush is well documented (262).
The single confirmed case of SFG rickettsiosis during the course of the study suggests that cases of rickettsiosis do occur even though most of those arising through activities in the WA bush maybe asymptomatic. However due to the distance between towns and cities and the low population in the state, even in the most populated region of the southwest, cases of rickettsiosis may have gone unnoticed, unreported or misdiagnosed as being caused by another pathogen. This pattern of under-detection has been observed to occur with other Rickettsia sp. elsewhere (126, 212).
In conclusion, the study has led to an increased awareness of arthropod-borne infections by the affected populations. By the end of the study, precautionary measures were being implemented by volunteers and those with whom they come into contact. Simple
125 precautionary measures, such as frequent checking for ticks during prolonged periods in the bush, were encouraged by the participating organisations. In some cases light coloured work clothing was issued to allow spotting of ticks easily. Other methods of prevention included treatment of clothing with insect repellents, long trousers tucked into socks and the application of effective repellent to exposed skin. An extra precaution suggested by this research would be to avoid locations where large macropods
(kangaroos) and feral pigs may wallow or take shelter, thus reducing the likelihood of encountering the tick species associated with these animals.
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7 - Final Discussion
There has been renewed interest in human rickettsial disease in Western Australia although much is still unknown. There are three groups of rickettsial organisms known to be present in WA; spotted fever group, typhus group and scrub typhus. The latter two are well characterised with typhus group rickettsia (Rickettsia typhi) mainly reported from urban centres (155, 195, 275) and scrub typhus (Orientia tsutsugamushi) in the northern, tropical regions of the state (239). So far the only spotted fever group rickettsiae reported have been Rickettsia felis and more recently two novel species,
Rickettsia gravesii and candidatus “Rickettsia antechini” (206, 208). The distribution of
SFG rickettsiae in WA required further investigation and thus became this study’s focus. Due to the large size of the state, the study focus was restricted to the most populated south-west corner.
7.1 Fulfilment of Study Aims
I. To characterise phenotypic traits of R. gravesii that are distinct from other
rickettsiae.
Rickettsia gravesii sp. nov. was demonstrated to be serotypically distinct from other closely related rickettsiae. Serotypical analysis by Specificity Difference (SPD) (227) demonstrated that R. gravesii was distinct from R. massiliae, a close genotypic relative.
Rickettsia gravesii was also compared with two other abundant Australian rickettsiae, R. australis and R. honei, as they may have overlapping regions of endemicity. The three rickettsiae were serotypically distinct. Other characteristics, such as vector species and geographical distribution, also emphasise the uniqueness of R. gravesii (208). This gives support to its acceptance as a novel member of the Rickettsia genus.
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II. To examine the arthropod host species known to harbour R. gravesii.
Expanding the arthropod host species harbouring R. gravesii from four to six was deemed an important step to further understand the ecology of the organism and its role within the West Australian biosphere. Two tick species were added, I. australiensis and
I. fecialis (Table 4.3). Amblyomma triguttatum’s role as an arthropod reservoir was further demonstrated by the high prevalence observed in all populations tested and the evidence of potential transtadial, transovarial and horizontal transmission (Chapter 4).
Other tick species collected had a relatively low prevalence of R. gravesii comparative to A. triguttatum (Table 4.2).
Amblyomma triguttatum’s abundance and affinity for larger vertebrate mammals
(including humans) compared to other tick species suggests that it is the principle vector for R. gravesii in WA (172, 261). Of the 4 subspecies of A. triguttatum, the subspecies
A. triguttatum triguttatum (172, 206-208) was investigated in this study. None of the other three subspecies has ever been collected or observed in southwest WA or Barrow
Island i.e. A. triguttatum queenslandense, A. triguttatum ornatissimum or A. triguttatum rosei. Amblyomma triguttatum triguttatum has also been reported in other regions of
Australia, especially in QLD and NSW, however no studies have been performed to investigate the presence of Rickettsia sp. in these states. Thus, R. gravesii may or may not be present in those tick populations, for even though the ticks are of the same subspecies, the different populations are separated by desert in the middle of Australia.
No records of A. triguttatum triguttatum have been found from the Northern Territory,
South Australia or far western Queensland. One can only theorise on the split and sub- speciation of A. triguttatum triguttatum on the continent to different areas without a visible land connection. Investigating the presence of R. gravesii in tick populations on
128 the eastern part of Australia may shed light on how closely related the populations are to each other and whether R. gravesii is unique to WA.
The serendipitous identification of a species of tick (H. longicornis) recently determined to be new to WA and possibly introduced via the importation of cattle from SA (28), highlights the potential for unintentional spread of rickettsial organisms by humans via their normal activities. The report of H. longicornis on a feral pig points to the possible establishment of the tick species among local fauna following cattle importation. The tick species has yet to be reported from any other vertebrate hosts in WA. As only one individual was obtained, further investigation into the encroachment of H. longicornis among WA fauna is warranted. More relevant to this study, H. longicornis has been implicated as a vector for SFG rickettsiae and Coxiella sp. in Korea (169, 170). Thus it has the potential to acquire local rickettsiae and become a rickettsial vector.
III. To determine the sero-prevalence of R. gravesii in a wild animal population.
With the high prevalence of R. gravesii in A. triguttatum ticks in WA, and the ticks’ affinity for large vertebrate bush mammals (and humans), a bush animal species was needed that would give an insight into the A. triguttatum burden and resulting rickettsial seroprevalence. The feral pig was the ideal animal for this part of the study.
Results from the study of ticks and rickettsiae in feral pigs revealed a high sero- prevalence to SFG rickettsia (47%) and provided strong evidence of transmission of a spotted fever group rickettsia by ticks (Chapter 5). The seroprevalence together with the presence of rickettsial DNA in 3.1% of feral pig livers suggested that a small proportion of the feral pig population may be rickettsaemic at any one time. However, the duration of rickettsaemia in the animal is unknown.
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The feral pig is widely distributed in Australia, with an estimated size of more than 20 million free roaming pigs. Its overlap with Australian native mammal populations is well documented and thus the ectoparasites found on the feral pig have also been found on other mammals (262). The study on the feral pig in south west WA has given an indication of probable high rickettsial exposure in other species of local large vertebrate mammals.
In Spain, the prevalence of the SFG rickettsia R. slovaca in Dermacentor marginatus ticks collected off wild boars, and their anti-rickettsial sero-prevalence was high (203,
204). The sero-prevalence (~50%) was similar to that obtained in the current study however the prevalence of SFG rickettsiae detected in the tick populations was higher in
Australia. Nevertheless, no physical rickettsial presence (genomic or culture isolates) were reported in the Spanish study.
IV. To identify the regions within south-west WA endemic for R. gravesii.
The current study showed that R. gravesii was prevalent throughout southwest WA. As the A. triguttatum ticks are also known to infest kangaroo populations in the region
(262), it is likely that kangaroos in WA may also be involved in the ecology of the infections.
With R. gravesii now known to be endemic in southwest WA, the question remains as to its distribution elsewhere. A similarly high prevalence was identified on Barrow
Island in the north of the state (Chapter 5) (208) suggesting a possible state-wide distribution. Antibodies to spotted fever group rickettsiae have been observed previously in the Kimberley region (112). Amblyomma triguttatum and kangaroos are
130 abundant in these areas, as are other larger mammals such as feral pigs, feral goats and dogs. It would be valuable to investigate whether the observed seroprevalence was due to exposure to R. gravesii or some other SFG rickettsia.
V. To investigate the relationship between sero-prevalence to spotted fever
rickettsia in humans and their level of bush activity.
The prevalence of R. gravesii in ticks and feral pigs strengthens the hypothesis that humans involved in bush activities are also exposed to the organism. Previous studies in
WA (112) had shown serological evidence of SFG infections in humans and thus a background level was expected. A risk analysis study was designed, using logistic regression to calculate odds ratios. A bush-associated occupational risk was observed to be the highest in the two tested groups. A bush-associated recreational group had lower risk of being seropositive for SFG rickettsiae, but were still significantly higher than the state baseline. The high rates of seroprevalence observed in the high risks groups tested demonstrated a clear risk for those living and working on Barrow Island and for individuals in the southwest region of WA who are active in the bush.
VI. To report and characterise any illness linked to Rickettsia sp. infection
during the period of the study.
With the high rates of seroprevalence to SFG Rickettsia sp. observed, it was expected that human rickettsiosis would occur more frequently. However, only one known case occurred during the duration of this study (2005-2009), suggesting that rickettsioses in
WA are rare or under-reported. It is possible that R. gravesii infection is generally asymptomatic or causes only a mild illness with insignificant symptoms that diagnostic investigations are not undertaken.
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Another possibility is that ticks are not attached to humans for very long before being removed. People who spend time in the bush were observed to be highly aware of ticks and removed them promptly when they attached or were found crawling on their bodies.
SFG Rickettsia sp. transmission does not occur for several hours after tick attachment and hence the infectious dose in many cases may be small and below the threshold required to cause symptomatic illness but enough to stimulate an immune response and seroconversion of the person bitten.
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7.2 Rickettsia gravesii sp. nov.
This study has led to a better understanding of the ecology of SFG rickettsiae in WA.
The predominant SFG rickettsia appears to be R. gravesii sp. nov. that is endemic to
WA. The main arthropod vector and reservoir is Amblyomma triguttatum with maintenance of the organism by transovarial and transtadial transmission (Figure 7.1 I).
Vertical transmission was observed in this study following the identification of rickettsial DNA in viable larvae from infected parents and in all other stages of the tick
(nymph and adult). The transmission of rickettsiae to humans and animals may thus occur during all three stages of the life cycle of A. triguttatum’s where a blood meal is required (Figure 7.1 I and II). Amblyomma triguttatum is a three-host tick although questions remain as to whether the larval and nymphal stages of the tick have a similar affinity for host species as the adult.
A high prevalence of anti-SFG rickettsia antibodies occurred in humans and animals with high tick exposure suggesting successful transmission via tick bites (Figure 7.1
II). No other rickettsial species was observed in this study other than R. gravesii, suggesting that it may be the main cause of anti-SFG rickettsia sero-prevalence in south-west WA. No significant illness was observed in the animals (feral pigs) and humans involved in the study, although sporadic cases of human rickettsiosis have been suspected. The illness caused by R. gravesii in humans may be mild or asymptomatic in most cases. This study investigated prevalence in the feral pig only, but similar prevalences may occur in other vertebrate mammals that share similar ectoparasite burdens within similar geographical boundaries.
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Figure 7.1 – The transmission cycle of Rickettsia gravesii in Western Australia. 134
Amblyomma triguttatum is the predominant tick species collected from animals and from the environment by flagging. Other tick species such as I. australiensis were only observed to harbour R. gravesii when they shared similar hosts with A. triguttatum. This may suggest that the horizontal transmission of rickettsiae from A. triguttatum
(reservoir) to other tick species can occur. (Figure 7.1 II to IV). Observation of tick locations on the host supports this hypothesis, where the close co-proximity during feeding may increase the chance of horizontal transmission. This was however untested in the study and would be worth pursuing to demonstrate whether or not it does occur.
This study has revealed a close relationship between R. gravesii and A. triguttatum ticks from south-west WA. Its relationship to other tick species and their hosts is still unknown and needs investigation. Evidence presented in this thesis gives support for A. triguttatum as the main reservoir for R. gravesii. Other tick species may also be reservoirs, but with the high prevalence of R. gravesii, and its abundance and affinity for larger mammals, including humans, it is the most likely tick species associated with this newly-recognised rickettsial species.
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7.3 Future Directions
Western Australia is approximately a third of the total land mass of Australia, with a land area of 2,529,875 km2 (Figure 4.2). Thus WA is larger than the state of Texas
(USA), Japan and the UK combined (Figure 7.2A). The total population of WA in 2010 was 2,296,411 (9). This equate to less than 1 person per square kilometre of land. The majority of land in WA is virtually uninhabited, with most of the population (75%; ABS
2007 data) living in and around Perth and south-west WA (Figure 7.2B). These figures demonstrate the massive undertaking it would require to fully investigate rickettsial organisms and their relationship with local animal populations. With current knowledge; O. tsutsugamushi occurs in the north (112, 239), R. felis and R. typhi in cities and towns (155, 195, 277) and the novel R. gravesii and R. antechini on Barrow island and the south-west part of the state (206-208).
The focus of this study was to attempt to understand the mechanics of transmission and prevalence of R. gravesii in the bush in south-west WA. Even though the focus was on
Rickettsia sp., only R. gravesii was observed in samples, with no other members of the
Rickettsia genus detected. The apparent absence of R. felis and R. typhi in the WA bush further supports the hypothesis that they are “urban rickettsioses” involving fleas infesting domestic pets and urban rats, respectively. A possible future direction for rickettsial research in WA would be investigating rickettsia-like illnesses in the towns and cities of WA.
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Figure 7.2 – Maps of Western Australia showing (A) comparative size of WA to other nations or states of the world and (B) population distribution based on data from the Australian Bureau of Statistics.
There is still much about the novel rickettsia R. gravesii that is not yet known. Little or no investigations have been performed in the regions between south-west WA and
Barrow Island. The lack of specimens and information can be attributed to the few towns in the area and to conservation efforts that tend to be focused on ecologically rich regions of the south-west and the Kimberley in the north. Efforts to collect samples from both humans and animals in this region should be pursued to confirm the hypothesis that R. gravesii occurs throughout the state.
There are 23 species of ticks in WA and so far only 6 species have been shown to harbour R. gravesii. In addition, only one of the six species (A. triguttatum) has been
137 investigated extensively. The remaining 5 species of ticks harbouring R. gravesii are still to be investigated to determine if they are effective reservoirs of R. gravesii and if they can act as vectors. While there is evidence for R. gravesii being transmitted transovarially and transstadially in A. triguttatum, this may or may not occur in other tick species.
This study focused on the relationship between ticks, rickettsiae and a large vertebrate mammal - the feral pig. However, WA has many other mammals that are much smaller in stature and less numerous. The ectoparasite burdens of these smaller mammals are usually different from those of the feral pig, kangaroos and other large animals that roam the West Australian bushland. Investigations into these smaller mammals would have to be multi-pronged due to their diversity. Animal behavioural factors need consideration as they are highly variable, including nocturnal, arboreal, burrowers, carnivores, omnivores, varied lifespan and geographical range of each marsupial. These factors may influence the tick species found on the various marsupials of WA. Species of ticks not observed in this study may give new insights into the ecology of R. gravesii and candidatus “R. antechini”.
Much of the sample collecting and analysis for rickettsial organisms prior to this study has been opportunistic. The deliberate collection of human, feral pig and ectoparasite samples for rickettsial analysis is the first study of its kind in south-west WA. This has contributed to an increase in awareness among various groups and organisations in WA, either directly or indirectly linked to the study of rickettsial organisms and their transmission. This increased awareness has seen the implementation of strategies to prevent tick bites and to treat adverse reactions relating to tick infestation with more
138 concern (pers. comm. Whiteman Park staff and committee members of Western
Australia Rogaining Association).
Rickettsia gravesii is genotypically closely related to the massiliae sub-group and this close relationship raises the question of how two rickettsiae geographically so far apart can be so closely related genetically. There are several possible explanations for such a close relationship but the most plausible is the introduction of a common ancestor to
WA by migratory birds. Rickettsia gravesii has retained some of the similarities between itself and members of the R. massiliae subgroup. However over time, and due to isolation, it has become distinctively different and is thus considered to be a novel species. The best way to examine the close relationship between R. gravesii and its distant relatives would be by an in-depth investigation into its genome. Currently work is being performed in collaboration with the Unite de Rickettsies in Marseille to sequence the entire genome of R. gravesii.
In conclusion, there is still much to be learned about the status and ecology of rickettsiae in WA. Some questions have been answered in this study with regards to
SFG prevalence and R. gravesii transmission, but many questions remain unanswered; the wealth of information waiting to be discovered makes the discoveries of this study seem miniscule. Australia is indeed the “Eldorado of Rickettsiology” (Pierre Edouard
Fournier, pers. comm.).
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300. Tsai, KH, Lu, HY, Huang, JH, Wang, PJ, Wang, HC, Huang, CG, Wu, WJ and Shu, PY (2009) Rickettsia felis in cat fleas in Taiwan. Vector Borne Zoonotic Diseases, 9; 561-563.
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308. Unwin, ML (1935) "Coastal fever" and endemic tropical typhus in north Queensland: recent investigation, clinical and laboratory findings. Medical Journal of Australia, Sept 7; 303-308.
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313. Vestris, G, Rolain, JM, Fournier, PE, Birg, ML, Enea, M, Patrice, JY and Raoult, D (2003) Seven years' experience of isolation of Rickettsia spp. from clinical specimens using the shell vial cell culture assay. Annals of the New York Academy of Sciences, 990; 371-374.
314. Vilcins, IM, Old, JM and Deane, E (2009) Detection of a Hepatozoon and spotted fever group Rickettsia species in the common marsupial tick (Ixodes tasmani) collected from wild Tasmanian devils (Sarcophilus harrisii), Tasmania. Veterinary Parasitology, 162; 23-31.
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320. Walker, DH and Fishbein, DB (1991) Epidemiology of rickettsial diseases. European Journal of Epidemiology, 7; 237-245.
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324. Watt, G and Parola, P (2003) Scrub typhus and tropical rickettsioses. Current Opinions on Infectious Diseases, 16; 429-436.
325. Webb, L, Carl, M, Malloy, DC, Dasch, GA and Azad, AF (1990) Detection of murine typhus infection in fleas by using the polymerase chain reaction. Journal of Clinical Microbiology, 28; 530-534.
326. Weil, E and Felix, A (1916) Zur serologischen Diagnose des Fleckfiebers. Wien Klin Wochenschr, 29; 33-35.
327. Weinert, LA, Welch, JJ and Jiggins, FM (2009) Conjugation genes are common throughout the genus Rickettsia and are transmitted horizontally. Proceedings of the Royal Society of Biological Sciences, 276; 3619-3627.
328. Weyer, F (1952) The behavior of Rickettsia akari in the body louse after artificial infection. American Journal of Tropical Medicine and Hygiene, 1; 809- 820.
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330. Whitman, TJ, Richards, AL, Paddock, CD, Tamminga, CL, Sniezek, PJ, Jiang, J, Byers, DK and Sanders, JW (2007) Rickettsia parkeri infection after tick bite, Virginia. Emerging Infectious Diseases, 13; 334-336. 166
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333. Williams, SG, Sacci, JBJ, Schriefer, ME, Andersen, EM, Fujioka, KK, Sorvillo, FJ, Barr, AR and Azad, AF (1992) Typhus and typhuslike rickettsiae associated with opossums and their fleas in Los Angeles County, California. Journal of Clinical Microbiology, 30; 1758-1762.
334. Woese, CR (1987) Bacterial Evolution. Microbiological Reviews, 51; 221-271.
335. Wolbach, SB (1919) Studies on Rocky Mountain spotted fever. Journal of Medical Research, 41; 1-197.
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337. Woolley, MW, Gordon, DL and Wetherall, BL (2007) Analysis of the first Australian strains of Bartonella quintana reveals unique genotypes. Journal of Clinical Microbiology, 45; 2040-2043.
338. Yang, D, Oyaizu, Y, Oyaizu, H, Olsen, GJ and Woese, CR (1985) Mitochondrial origins. Proceedings of the National Academy of Sciences, USA, 82; 4443-4447.
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340. Zavala-Velazquez, JE, Zavala-Castro, JE, Vado-Solis, JA, Ruiz-Sosa, JA, Moron, CG, Bouyer, DH and Walker, DH (2002) Identification of Ctenocephalides felis fleas as a host of Rickettsia felis, the agent of spotted fever rickettsiosis in Yucatan, Mexico. Vector Borne Zoonotic Diseases, 2; 69-75.
341. Zhu, Y, Fournier, PE, Eremeeva, M and Raoult, D (2005) Proposal to create subspecies of Rickettsia conorii based on multi-locus sequence typing and an emended description of Rickettsia conorii. BMC Microbiology, 5; 1-11.
342. Znazen, A and Raoult, D (2007) Flea-Borne Spotted Fever. IN Raoult, D and Parola, P (Eds.) Rickettsial Diseases. New York, Informa Healthcare.
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9 - Appendices
9.1 Ethics Approval Obtained
Permit Type Project Title Institution Date of Number Approval
2005/099 Human The epidemiology of Murdoch May 2005 rickettsial diseases in University, WA, Western Australia. Australia
R1155/05 Animal Risks to human health and Murdoch October the ecosystem from feral University, WA, 2005 pigs in Perth metropolitan Australia water catchments.
05/609 Human Rickettsial Spotted Fever in Fremantle November Western Australia: Hospital, WA, 2005 Identifying the infectious Australia disease burden in a high-risk group.
2006/023 Human Rickettsial Spotted Fever in Murdoch March Western Australia: University, WA, 2006 Identifying the infectious Australia disease burden in a high-risk group.
FTM- Animal Raising of polyclonal Mahidol July 2007 ACUC antibodies to enable the University, 007/2007 serologic differentiation of Bangkok, spotted fever group Thailand rickettsia.
NS2049/07 Animal Serologic typing of Murdoch August “Rickettsia gravesii” by University, WA, 2007 Microimmunofluorescence. Australia
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9.2 GenBank Accession Numbers
Species 16S RNA gltA ompA ompB sca4
R. aeschlimanii U74757 U59722 U43800 AF123705 AF163006
R. africae L36098 U59733 U43790 AF123706 AF151724
R. akari L36099 U59717 NA AF123707 AF213016
R. australis L36101 U59718 AF149108 AF123709 AF187982
R. belii L36103 U59716 NA NA NA
R. canadensis U15162 U59713 EF160122 NA NA
R. conorii AF541999 U59730 U43806 AF149110 AF163008
R. felis L28944 AF210692 AF210694 AF210695 AF196973
R. gravesii strain DQ269434 DQ269435 DQ269437 DQ269438 DQ269439 BWI-1
R. heilongjiangii AF178037 AF178034 AF179362 AY260451 AY331396
R. honei L36220 U59726 U43809 AF123724 AF163004
R. japonica L36213 U59724 U43795 AF123713 AF155055
R. massiliae L36214 U59719 U43799 AF123714 AF163003
R. montanensis L36215 U74756 U43801 AF123716 AF163002
R. parkeri L36673 U59732 U43802 AF123717 AF155059
R. prowazekii M21789 M17149 NA AF123718 AF200340
R. raoultii DQ365810 DQ365804 DQ365801 DQ365798 DQ365808
R. rhipicephali L36216 U59721 U43803 AF123719 AF155053
R. rickettsii L36217 U59729 U43804 X16353 AF163000
R. sibirica L36218 U59734 U43807 AF123722 AF155057
R. slovaca L36224 U59725 U43808 AF123723 AF155054
R. tamurae AY049981 AF394896 DQ103259 DQ113910 DQ113911
R. typhi L36221 U59714 NA NC006142 NC006142
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9.3 Rogainer Study Consent Form
Note: The information sheets disseminated to the respective groups were very similar to each other, unlike the questionnaires. The rogainer information sheet is thus used as an example. The most important part of the information sheets were the Consent Form and Revocation of Consent Form, which is the same for all groups.
Identification code: |_|_|_|_|_|
STUDY OF INFECTION RISKS FROM TICKS
PATIENT INFORMATION STATEMENT AND CONSENT FORM
______
Principal Investigator: Dr John Dyer Infectious Diseases Physician SMAHS Infectious Diseases Service Level 2 B Block, Fremantle Hospital Alma St Fremantle WA 6160 Telephone: 08 9431 2149 Facsimile: 08 9431 2035
Contact Person: Dr John Dyer Infectious Diseases Physician SMAHS Infectious Diseases Service Level 2 B Block, Fremantle Hospital Alma St Fremantle WA 6160 Telephone: 08 9431 2149 Facsimile: 08 9431 2035
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Patient Information and Consent Form
You are invited to participate in the research study outlined below.
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 you will undergo, and other important study information. Please read this information carefully and, if you have further questions, please ask the Principal Investigator.
Rickettsial bacteria that are transmitted to humans by tick bite may be an important unrecognised cause of infection in Western Australia. We plan to investigate the occurrence of rickettsial infection in rogainers, who are frequently exposed to ticks in pursuit of their sport. Any rickettsial germs found as a result of these studies will be analysed in detail to determine to what extent they resemble other species known to cause human disease.
2. Purpose to Study
The objective of the study is to identify transmission pathways and disease risks of rickettsial spotted fever in Western Australia. We wish to discover whether individuals engaged in activities such as rogaining, who have a high risk of tick contact, may have increased rates of tick-transmitted rickettsial infection.
The ultimate goals of this research are:
to identify the transmission risks and occurrence of rickettsial infection in Western Australia to reduce the risk of rickettsial transmission in individuals residing, recreating or working in high-risk areas to develop health and safety guidelines for individuals at high risk of tick- borne infection
3. The Study Design
An ethics committee has examined and approved this study before any participant is allowed to enroll in the study. A total of approximately 144 active rogainers recruited by advertisement through the WA Rogaining Association will participate.
This study protocol will consist of: (a) baseline analysis of a blood test obtained at the beginning of the summer rogaining season to determine previous exposure to rickettsial infection, and to correlate this with the likely intensity of tick exposure; (b) a second blood test at least one month after the final rogaining event of the summer season to ascertain any cases of rickettsial infection, whether symptomatic or not, that may have occurred over this period of time. At the time of collection of the initial blood test, participants in the study
171 will be asked to fill out a self-administered questionnaire to measure previous level of tick exposure.
Participants will also be asked to keep a diary of recognised tick exposures during the course of the season, and to report illnesses (especially with fever &/or rash) that may represent rickettsial infection to the study co-ordinator, who will arrange for further medical assessment and investigation as medically indicated. Blood tests obtained from such symptomatic study participants will be sent to a reference laboratory for additional testing (culture and polymerase chain reaction/sequencing analysis) so as to identify the causative microbe. Finally, participants will be asked to collect ticks to which they are exposed during rogaining events and submit them to the study investigators for further analysis for potentially pathogenic microbes at the School of Veterinary and Biomedical Sciences, Murdoch University.
4. Procedures
If, after reading this information sheet and discussing the trial with the study investigators, you decide to take part, you will be asked to sign a form confirming that you agree to participate.
As part of this study you will have blood drawn (approximately 10mL or 1.5 tablespoons) at the time of enrolment and a further 10mL sample will be taken one or month or more after the final event of the season. At the time of your first visit you will be asked to fill in a questionnaire aimed at determining your past level of risk for tick exposure.
You will also be asked to keep a diary noting and describing any tick exposures during the course of the season. These documents will be collected by the study investigators and analysed to determine correlations between blood test results and intensity of tick exposure. You will also be asked to collect any ticks with which you come in direct contact and submit them for further analysis to the study investigators.
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 medical doctor (Dr. John Dyer), 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 needlestick.
6. Benefits of Participating in the Study
We anticipate increased knowledge of the presence and transmission of rickettsial infection in WA to result in direct benefits for participants in sports such as rogaining that involve significant contact with the natural environment.
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7. Study Expenses
The only specific cost in this study will be the travel costs for attending the clinic on 2 occasions for study specific visits. Reimbursement for these costs will be available from the study co-ordinator.
8. Financial Support for the Study
Financial support to cover the costs of this study is being provided by a research grant from the Fremantle Hospital Medical Research Foundation. 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. The account is subject to audit by the Auditors to Fremantle Hospital.
9. Questions about the Study
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. 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.
Your doctor 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.
All information obtained in connection with this study will remain confidential. You will not be identified in any publication or public presentation of data from this study. Only authorised personnel from the SMAHS ethics committee will have access to study documents and results. By signing this form, you give permission that these authorised persons may have access to study documents if necessary.
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 John Dyer
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Contact No: 08 9431 2149
If you have a problem during the study and would like to talk to an independent person you may contact: Fremantle Hospital Ethics Committee Representative Contact No: 08 9431 2929.
You will be given a copy of this form to keep.
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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 John Dyer on 08 9431 2149, who will be happy to answer them.
8. I acknowledge receipt of a copy of this Consent Form and the Subject Information Statement. Complaints may be directed to (Chair of South Metropolitan Area Health Service – Human Research Ethics Committee on 08 9431 2929.
______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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CONSENT FORM 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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9.4 Rogainers’ Questionnaire
Identification code: |_|_|_|_|_|
STUDY OF INFECTION RISKS FROM TICKS QUESTIONNAIRE
Thank you for your interest and co-operation in our research.
The purpose of this questionnaire is to find out whether rogainers are at risk from bacterial infections carried by ticks.
Please circle the appropriate answer, tick the box or enter an answer as appropriate.
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GENERAL QUESTIONS
1. Date today: / / 2006
2. Gender: Male Female
3. Year of birth: |_|_|_|_|
4. What is your current occupation (please include paid or unpaid work):
…………………………………..
5. How long have you been an active rogainer?
|_|_| Months |_|_| Years
6. How long have you been a member of the WARA?
|_|_| Months |_|_| Years
7. On average, how many times a year do you participate in a rogaining event? |_|_| times
8. At what level do you currently compete in rogaining?
…………………………………..
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RECORD OF PAST AND CURRENT ACTIVITIES
9. Apart from rogaining, how often do you spend time in the bush or scrub for work or recreation? (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 apart from rogaining
If you answered “I spend NO time in the bush apart from rogaining ” to question 9, please go straight to question 12 (next page). Otherwise, please proceed to question 10.
10. If you spend time in the bush or scrub APART from rogaining, how long do you usually go for? |_|_| hours at a time
11. In the past 2 years, how often have you been bushwalking, hiking or camping in other states of Australia or overseas? (tick ALL boxes that apply)
Interstate Overseas (outside Australia)
Not at all in past 2 years Not at all in past 2 years
Less than once a year Less than once a year Which Which state(s)/territories?………… country/countries?…………
About once a year About once a year Which Which state(s)/territories?………… country/countries?…………
More than once a year More than once a year Which state(s)/territories?………… Which country/countries?…………
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RECORD OF TICK BITES
12. IN THE PAST 2 YEARS, have you EVER been bitten by a tick?
Yes No (Go to Q13) Not sure (Go to Q13) If YES:
a. In which month and year did this occur? (e.g. Feb 2005)
……/………. b. Where were you when this occurred? Location (eg name of park or district): ……………. State: …………….
c. 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
d. Did you remove the tick yourself? Yes No: removed at a medical clinic No : removed by someone else: who?......
e. Did you suffer a reaction to the tick bite? Yes No (Go to question 12g) Not sure (Go to question 12g)
f. What was the nature of the reaction to the tick bite?
Nature of reaction Please tick any symptoms that you have had Immediate (up to a couple of days after the tick bite) - Swelling of the skin around the bite - Tenderness of the skin around the bite - Discharge, pus or fluid coming out of the skin around the bite In the 7-14 days after tick bite - Headache - Fever or chills - 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 - Aching in joints, limbs, back or neck without reason - Tiredness/ Feeling „out of energy‟ without reason - Shortness of breath or difficulty breathing without reason - Blistering/scabbing around skin area where bitten 180
g. Did you seek medical advice for the tick bite? Yes No Not sure
h. Were you treated with antibiotics after the tick bite? Yes No Not sure
ADDITIONAL DETAILS: PLEASE ANSWER THESE QUESTIONS FOR ANY OTHER TICK BITES THAT OCCURRED IN THE LAST 2 YEARS
a. In which month and year did this occur? (e.g. Feb 2005)
……/………. b. Where were you when this occurred? Location (eg name of park or district): ……………. State: …………….
c. 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
d. Did you remove the tick yourself? Yes No: removed at a medical clinic No : removed by someone else: who?......
e. Did you suffer a reaction to the tick bite? Yes No (Go to question g) Not sure (Go to question g)
f. What was the nature of the reaction to the tick bite?
Nature of reaction Please tick any symptoms that you have had Immediate (up to a couple of days after the tick bite) - Swelling of the skin around the bite - Tenderness of the skin around the bite - Discharge, pus or fluid coming out of the skin around the bite In the 7-14 days after tick bite - Headache - Fever or chills - Rash TICK ALL THAT APPLY around the bite on chest, back or abdomen (stomach region) hands, arms or shoulders
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feet or legs on neck, face or scalp on groin or buttocks - Aching in joints, limbs, back or neck without reason - Tiredness/ Feeling „out of energy‟ without reason - Shortness of breath or difficulty breathing without reason - Blistering/scabbing around skin area where bitten
g. Did you seek medical advice for the tick bite? Yes No Not sure
h. Were you treated with antibiotics after the tick bite? Yes No Not sure
13. Have you EVER experienced any of the following symptoms since you started rogaining?
Nature of Please How many What Do you reaction tick any times have time of remember being sympto these symptoms the year bitten by a tick ms that occurred since did these anytime in the you you began symptom fortnight before have rogaining? s occur? these symptoms had (eg. developed? February) Skin swelling/ Yes if YES: Yes tenderness No No Not sure |_|_|_| Can‟t remember
Prolonged headache Yes if YES: Yes No No Not sure |_|_|_| Can‟t remember
Rash on the arms or Yes if YES: Yes chest No No Not sure |_|_|_| Can‟t remember
General aching in Yes if YES: Yes joints, limbs, back or No No neck without reason Not sure |_|_|_| Can‟t remember
Prolonged tiredness/ Yes if YES: Yes Feeling „out of energy‟ No No without reason Not sure |_|_|_| Can‟t remember
Fever or chills Yes if YES: Yes No No Not sure |_|_|_| Can‟t remember
Shortness of breath or Yes if YES: Yes difficulty breathing No No without reason Not sure |_|_|_| Can‟t remember
182
OTHER MEDICAL HISTORY
14. Do you have any major medical conditions?
Yes No
If YES: Please describe ……………………………….……………………
……………………………………………………
15. Are you routinely on any medications or treatments?
Yes No
If YES: a. What is the name of the medication?
……………………………….……………………
……………………………………………………
b. What condition or health problem do you take this for? ……………………………….……………………
……………………………………………………
Thank you for taking the time to complete this survey.
183
9.5 Rogainers’ Follow-up Questionnaire
Identification code: |_|_|_|_|_|
STUDY OF INFECTION RISKS FROM TICKS QUESTIONNAIRE
Thank you for your interest and co-operation in our research.
The purpose of this questionnaire is to find out whether rogainers are at risk from bacterial infections carried by ticks.
Date today: |_|_| /|_|_| / 2008
184
GENERAL QUESTIONS
1. What is your current occupation (please include paid or unpaid work):
…………………………………..
2. Are you still an active rogainer?
|_|_| Months |_|_| Years
3. Are you still a member of the WARA?
|_|_| Months |_|_| Years
4. On average, how many times in the past year do you participate in a rogaining event? |_|_| times
5. When you are at a rogaining event, in the past year, do you usually complete:
The full course Part of the course Other………………………..
6. On average, in the past year, how long do you participate in an event for?
|_|_| hours |_|_| minutes
185
RECORD OF PAST AND CURRENT ACTIVITIES
7. Apart from rogaining, in the past year how often do you spend time in the bush or scrub for work or recreation? (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 apart from rogaining
If you answered “I spend NO time in the bush apart from rogaining” to question 9, please go straight to question 12 (next page). Otherwise, please proceed to question 10.
8. If you spend time in the bush or scrub APART from rogaining, what activities are you involved in and how long do you usually go for? Activity………………………………………|_|_| hours at a time Activity………………………………………|_|_| hours at a time Activity………………………………………|_|_| hours at a time
9. In the past year, how often have you been bushwalking, hiking or camping in bushland, forests or other wilderness areas in other states of Australia or overseas? (tick ALL boxes that apply)
Interstate Overseas (outside Australia)
Not at all in the past year Not at all in the past year
Once Once Which Which state(s)/territories?………… country/countries?…………
More than once More than once Which state(s)/territories?………… Which country/countries?…………
186
RECORD OF TICK BITES
10. IN THE PAST YEAR, how often are you been bitten by ticks while rogaining or doing any other activity in bushland? This refers to tick bites that may have occurred while 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 Q13) Not sure (Go to Q13)
If the subject answered “Never” or “Not sure” to question 12, please go straight to question 13. Otherwise, please proceed.
a. In which months do the bites usually occur? (e.g. Mar-May) …….
b. In what locations were you when you were bitten by ticks IN THE PAST YEAR? This includes while rogaining or doing some other activity.
Location (eg name of park or district): …………….State: ……………. What was the activity? rogaining other......
Location (eg name of park or district): …………….State: ……………. What was the activity? rogaining other......
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 YEAR?
Nature of reaction Please tick any symptoms that you have had Immediate (up to a couple of days after tick bites) - Swelling of the skin around the bite Most or every 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 every time I get bitten (more than 75% of the time) Often when I get bitten (25-75% of the time) Occasionally when I get bitten (Less than 25% of the time) Never Not sure - Discharge, pus or fluid coming out of Most or every time I get bitten (more than 75% of the time) the skin around the bite Often when I get bitten (25-75% of the time) Occasionally when I get bitten (Less than 25% of the time) Never Not sure - Itchiness around the bite Most or every 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 187
At any time during 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 over any part or all of your body* TICK ALL THAT APPLY around the bite on chest, back or abdomen (stomach region) hands, arms or shoulders feet or legs on neck, face or scalp on groin or buttocks
When did this occur? (e.g. Feb 2005)
……/……….
- Tender or painful „glands‟ (lumps or TICK ALL THAT APPLY nodes under the skin, usually away from in the groin the bite mark)* in the armpits in the neck other; where______
When did this occur? (e.g. Feb 2005)
……/……….
- Aching in joints, limbs, back or neck When did this occur? (e.g. Feb 2005) without reason* ……/……….
- Tiredness/ Feeling „out of energy‟ When did this occur? (e.g. Feb 2005) without reason* ……/……….
- Shortness of breath or difficulty When did this occur? (e.g. Feb 2005) breathing without reason* ……/……….
- Blistering/scabbing around skin area When did this occur? (e.g. Feb 2005) where bitten* ……/……….
If the subject answered “At any time in the 2 weeks after the tick bite” to ANY OF THE SYMPTOMS, please continue. Otherwise, please go straight to question 13.
I would like to ask you in more detail about the [*symptoms/symptoms] which occurred in ……/………. (START WITH THE EARLIEST DATE). Thinking back to the tick bites that occurred in the two weeks before these symptoms: e. In what locations were you when you were bitten by ticks in [date]…/….? Location (eg name of park or district): …………….State: ……………. What was the activity? rogaining other......
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 188
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
NOW PROCEED TO THE NEXT DATE ON THE TABLE ABOVE: I would now like to ask you about the [*symptoms/symptoms] which occurred in ……/………. Thinking back to the tick bites that occurred in the two weeks before these symptoms:
e. In what locations were you when you were bitten by ticks in [date]…/….? Location (eg name of park or district): …………….State: ……………. What was the activity? rogaining other......
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
189
NOW PROCEED TO THE NEXT DATE ON THE TABLE ABOVE: I would now like to ask you about the [*symptoms/symptoms] which occurred in ……/………. Thinking back to the tick bites that occurred in the two weeks before these symptoms:
e. In what locations were you when you were bitten by ticks in [date]…/….? Location (eg name of park or district): …………….State: ……………. What was the activity? rogaining other......
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
190
11. APART FROM ANY SYMPTOMS YOU HAVE MENTIONED ALREADY, have you EVER experienced any of the following symptoms? This may not necessarily be associated with a tick bite that you recall. Nature of Please How many What Do you remember reaction tick any times have time of being bitten by a symptom these symptoms the year tick anytime in the s that you occurred since did these 2 weeks before have had you began symptom these symptoms rogaining? s occur? developed? (eg. February) Skin swelling/ Yes if YES: Yes tenderness No No Not sure |_|_|_| Can‟t remember
Prolonged headache Yes if YES: Yes (> 3 days) No No Not sure |_|_|_| Can‟t remember
Rash over parts or all Yes if YES: Yes of my body No No Not sure TICK ALL THAT Can‟t remember APPLY around the bite on chest, back or abdomen (stomach region) hands, arms or shoulders feet or legs on neck, face or scalp on groin or buttocks
General aching in Yes if YES: Yes joints, limbs, back or No No neck without reason Not sure |_|_|_| Can‟t remember
Prolonged tiredness/ Yes if YES: Yes Feeling „out of energy‟ No No (> 3 days) without Not sure |_|_|_| Can‟t remember reason Fever or chills Yes if YES: Yes No No Not sure |_|_|_| Can‟t remember
Shortness of breath or Yes if YES: Yes difficulty breathing No No without reason Not sure |_|_|_| Can‟t remember
191
12. APART FROM ANY SYMPTOMS YOU HAVE MENTIONED ALREADY IN THE PAST YEAR, have you EVER experienced any other symptoms after spending time in the bush in Western Australia, Australia, or overseas?
What symptoms did Year In what locations were you experience? you when you experienced this?
Location (eg name of park or district): …………….State: …………….
What was the activity? ...... Location (eg name of park or district): …………….State: …………….
What was the activity? ...... Location (eg name of park or district): …………….State: …………….
What was the activity? ......
192
OTHER MEDICAL HISTORY
13. Do you have any major medical conditions?
Yes No
If YES: Please describe ……………………………….……………………
……………………………………………………
14. Are you routinely on any medications or treatments?
Yes No If YES: a. What is the name of the medication? ……………………………….…………………… ……………………………………………………
b. What condition or health problem do you take this for? ……………………………….…………………… ……………………………………………………
Thank you for taking the time to complete this survey.
193
9.6 Whiteman Park Workers’ Questionnaire
Identification code: |_|_|_|_|_|
WWHHIITTEEMMAANN PPAARRKK HHEEAALLTTHH AANNDD SSAAFFEETTYY QQUUEESSTTIIOONNNNAAIIRREE
Thank you for your interest and co-operation in our research. We are a group of doctors and veterinarians from the Division of Health at 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 in Western Australia. 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.
194
GENERAL QUESTIONS
1. Date today: / / 2006
2. Gender: Male Female
3. Year of birth: |_|_|_|_|
4. What is your current job/role in Whiteman Park:
…………………………………..
5. What are the main work tasks you do in this job?
……………………………………………………………………………………
………………..…………………………………………………………………
………………………………………………………………….…………………
6. How long have you worked in Whiteman Park?
|_|_| Months |_|_| Years
7. Approximately how many days or weeks a year do you work in Whiteman Park?
|_|_|_| Days OR |_|_| Weeks
8. Approximately how long do you spend on the park on an average working day?
|_|_| Hours
195
RECORD OF PAST AND CURRENT ACTIVITIES IN WHITEMAN PARK
9. While you are in Whiteman Park, 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 in Whiteman Park, how often do you go into the bush or scrub 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 in Whiteman Park:
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 in Whiteman Park?
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
196
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 in Whiteman Park?
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
197
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 in Whiteman Park.
Bushwalking, hiking, camping or working Bushwalking, hiking, camping or working in the bush or forest in other parts of in the bush or forest overseas (outside Western Australia (except for Australia) Whiteman Park) or interstate
(please tick the answer (please tick the answer that most applies to you) that most applies to you)
Not at all in past 2 years Not at all in past 2 years
Less than once a year Less than once a year Which Which state(s)/territories?………… country/countries?…………
About once a year About once a year Which Which state(s)/territories?………… country/countries?…………
More than once a year More than once a year Which state(s)/territories?………… Which country/countries?…………
198
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 in Whiteman Park, 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 work at Whiteman Park” 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)
in Whiteman Park elsewhere in Australia or overseas Location (eg name of area, park or district): ……………. State: ……………. Country: …………….
c. In which part(s) of your body do the bite(s) usually occur? (TICK ALL THAT APPLY) on chest, back or abdomen (stomach region) hands, arms or shoulders feet or legs on neck, face or scalp on groin or buttocks
d. Have you experienced any of the following after tick bites IN THE PAST 2 YEARS? Nature of reaction Please tick any symptoms that you have had Immediate (up to a couple of days after tick bites)
- Local swelling of the skin around the Most or e very time I get bitten (more than 75% of the time) bite Often when I get bitten (25-75% of the time) Occasionally when I get bitten (Less than 25% of the time) Never Not sure
- Tenderness of the skin around the bite Most or e very time I get bitten (more than 75% of the time) Often when I get bitten (25-75% of the time) Occasionally when I get bitten (Less than 25% of the time) Never Not sure
- Discharge, pus or fluid coming out of Most or e very time I get bitten (more than 75% of the time) the skin around the bite Often when I get bitten (25-75% of the time) Occasionally when I get bitten (Less than 25% of the time) Never Not sure
- 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
199
At any time during the 2 weeks after tick bites - Prolonged headache (> 3 days)* When did this occur? (e.g. Feb 2005)
……/……….
- Fever or chills* When did this occur? (e.g. Feb 2005)
……/……….
- Rash* TICK ALL THAT APPLY around the bite on chest, back or abdomen (stomach region) hands, arms or shoulders feet or legs on neck, face or scalp on groin or buttocks
When did this occur? (e.g. Feb 2005)
……/………. - Tender or painful „glands‟ (lumps or TICK ALL THAT APPLY nodes under the skin, usually away from in the groin the bite mark)* in the armpits in the neck other; where______
When did this occur? (e.g. Feb 2005)
……/……….
- Aching in joints, limbs, back or neck When did this occur? (e.g. Feb 2005) without reason* ……/……….
- Tiredness/ Feeling „out of energy‟ When did this occur? (e.g. Feb 2005) without reason* ……/……….
- Shortness of breath or difficulty When did this occur? (e.g. Feb 2005) breathing without reason* ……/……….
- Blistering/scabbing around skin area When did this occur? (e.g. Feb 2005) where bitten* ……/……….
If the subject answered “At any time in the 2 weeks after the tick bite” to ANY OF THE SYMPTOMS, please continue. Otherwise, please go straight to question 13.
I would like to ask you about the [*symptoms/symptoms] which occurred in ……/………. (START WITH THE EARLIEST DATE). Thinking back to the tick bites that occurred in the two weeks before these symptoms:
e. Where were you when you were bitten? in Whiteman Park 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 200
on neck, face or scalp on groin or buttocks
g. Did you remove the tick(s) yourself? Yes No: removed at a medical clinic No : removed by someone else: who?......
h. Did you seek medical advice for the tick bite(s)? Yes No Not sure
i. Were you treated with antibiotics after the tick bite(s)? Yes No Not sure
j. Did you miss any work because of the tick bite(s)? Yes if yes, How many days in total? ...... No
NOW PROCEED TO THE NEXT DATE ON THE TABLE ABOVE: I would now like to ask you about the [*symptoms/symptoms] which occurred in ……/………. Thinking back to the tick bites that occurred in the two weeks before these symptoms:
e. Where were you when you were bitten? in Whiteman Park 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
201
NOW PROCEED TO THE NEXT DATE ON THE TABLE ABOVE: I would now like to ask you about the [*symptoms/symptoms] which occurred in ……/………. Thinking back to the tick bites that occurred in the two weeks before these symptoms:
e. Where were you when you were bitten? in Whiteman Park 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
202
19. APART FROM ANY SYMPTOMS YOU HAVE MENTIONED ALREADY, have you EVER experienced any of the following since you started working in Whiteman Park?
(Please tick all the symptoms you have had and answer the questions in that row. If you haven‟t had any of the following symptoms then please go straight to question 20)
Nature of Please How many times What time of Do you remember tick any have these the year did being bitten by a tick reaction symptoms symptoms occurred these anytime in the 2 weeks that you since you started symptoms before these symptoms have had working in occur? (eg: developed? Whiteman Park? February)
Skin swelling/ Yes if YES: Yes tenderness No No Not sure |_|_|_| Can‟t remember
Prolonged headache Yes if YES: Yes (> 3 days) No No Not sure |_|_|_| Can‟t remember
Rash over parts or all Yes if YES: Yes of my body No No Not sure |_|_|_| Can‟t remember
TICK ALL THAT APPLY around the bite on chest, back or abdomen (stomach region) hands, arms or shoulders feet or legs on neck, face or scalp on groin or buttocks
General aching in Yes if YES: Yes joints, limbs, back or No No neck without reason Not sure |_|_|_| Can‟t remember
Prolonged tiredness/ Yes if YES: Yes Feeling „out of energy‟ No No (> 3 days) without Not sure |_|_|_| Can‟t remember reason Fever or chills Yes if YES: Yes No No Not sure |_|_|_| Can‟t remember
Shortness of breath or Yes if YES: Yes difficulty breathing No No without reason Not sure |_|_|_| Can‟t remember
203
CHEMICAL EXPOSURE HISTORY
20. Are you routinely in contact with any potentially toxic or dangerous chemicals either in or out of work hours? (e.g. pesticides, herbicides, fungicides, industrial cleaners etc.)
Yes No If YES: Please describe type or names of chemicals
…………………….……………………
Between which years did you use these chemicals? |_|_|_|_| - |_|_|_|_|
During this time, how often were/are you exposed?
Daily Weekly Monthly
Approximately how many minutes or hours are you in contact with these chemicals at a time?
|_|_| Hours OR |_ |_|_| Minutes
21. Have you ever been involved in an accident involving potentially toxic or dangerous chemicals? (e.g. pesticides, herbicides, fungicides)
Yes No If YES: Please describe type or names of chemicals
…………………….………………
In which year did this occur? |_|_|_|_|
Approximately how many minutes or hours are you in contact with these chemicals?
|_|_| Hours OR |_ |_|_| Minutes
Were you given/seek medical attention after the incident?
Yes No If YES: Were you ever diagnosed with any illness related to the
accident, please describe …………………….……………………
204
MEDICAL HISTORY
22. I would now like to ask you about whether you have had any particular kinds of infections in the past few years. In the past 5 years, Have you ever been diagnosed with an attack of gastroenteritis („gastro‟‟/ a stomach infection)?
Yes No
If YES:
Date or Year of diagnosis ………………
Was the cause of the infection or the bug ever identified? If yes: what was the cause/ name of the bug……………………………….……………………
Were you prescribed any medication after this illness?
Yes No
If YES: Please describe ………………………………………
23. In the past 5 years, have you ever been diagnosed with any infections of the urinary tract, including the kidney, bladder or urethra (which is the tube that drains the bladder)?
Yes No
If YES:
Date or Year of diagnosis ………………
Was the cause of the infection or the bug ever identified? If yes: what was the cause/ name of the bug……………………………….……………………
Were you prescribed any medication after this illness?
Yes No
If YES: Please describe ………………………………………
205
24. I am now going to read out a list of medical conditions that you may or may not have had in the past. Do not worry if you have not heard of many of them. Have you ever been diagnosed with any of the following medical conditions? (tick for any that apply)
Unexplained pain, redness or irritation of one or both eyes in the past 5 years yes: Date or Year of diagnosis ……………… Are you taking any prescribed medication for this condition? Yes No If YES: Please describe ………………………………………
ankylosing spondylitis (AS) If yes: Date or Year of diagnosis ……………… Are you taking any prescribed medication for this condition? Yes No If YES: Please describe ………………………………………
rheumatoid arthritis (this is different from so-called 'wear-and-tear' arthritis, or osteoarthritis) If yes: Date or Year of diagnosis ……………… Are you taking any prescribed medication for this condition? Yes No If YES: Please describe ………………………………………
Reiter‟s disease (INTERVIEWER NOTE: may need to spell out) If yes: Date or Year of diagnosis ……………… Are you taking any prescribed medication for this condition? Yes No If YES: Please describe ………………………………………
Crohn‟s disease or ulcerative colitis If yes: Date or Year of diagnosis ……………… Are you taking any prescribed medication for this condition? Yes No If YES: Please describe ………………………………………
Psoriasis If yes: Date or Year of diagnosis ……………… Are you taking any prescribed medication for this condition? Yes No
206
If YES: Please describe ………………………………………
25. Apart from those mentioned, have you ever been diagnosed with arthritis or any long-term joint, back or neck conditions ?
Yes No
If YES: Please describe which joints or parts of your neck or spine are affected ……………………………….……………………
Date or Year of diagnosis ………………
Are you still experiencing this medical condition? Yes No
Are you taking any prescribed medication for this condition? Yes No If YES: Please describe ………………………………………
26. Have any of your close family members and/or relatives been diagnosed with any form of arthritis or any long-term joint, back or neck conditions?
Yes No
If YES:
a. What is his/her relation to you?
……………………………….……………………
b. What is the name of the condition?
……………………………….……………………
……………………………………………………
207
OTHER MEDICAL HISTORY
27. Do you have any major medical conditions not already mentioned?
Yes No
If YES: Please describe ……………………………….……………………
……………………………………………………
28. Are you routinely on any medications or treatments not already mentioned?
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 the time to complete this survey.
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9.7 Whiteman Park Workers’ Follow-up Questionnaire
Identification code: |_|_|_|_|_|
WHITEMAN PARK STUDY OF INFECTION RISKS FROM TICKS QUESTIONNAIRE
Thank you for your interest and co-operation in our research.
The purpose of this questionnaire is to find out whether rogainers are at risk from bacterial infections carried by ticks.
Date today: |_|_| /|_|_| / 2008
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GENERAL QUESTIONS
1. What is your current job/role in Whiteman Park:
…………………………………..
2. What are the main work tasks you do in this job?
……………………………………………………………………………………
………………..…………………………………………………………………
………………………………………………………………….…………………
3. How long have you worked in Whiteman Park?
|_|_| Months |_|_| Years
4. Approximately how many days or weeks a year do you work in Whiteman Park?
|_|_|_| Days OR |_|_| Weeks
5. Approximately how long do you spend on the park on an average working day?
|_|_| Hours
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RECORD OF PAST AND CURRENT ACTIVITIES IN WHITEMAN PARK IN THE PAST YEAR
6. While you are in Whiteman Park in the past year, 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
7. While you are in Whiteman Park in the past year, how often do you go into the bush or scrub 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 6 AND 7, please go straight to QUESTION 14. Otherwise, please proceed to QUESTION 8.
8. 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
9. Is time you spend in the bush in Whiteman Park:
Mainly for work reasons Mainly recreational reasons Equally related to work and recreation
10. How often do you wear short-sleeved shirts and/or shorts during recreational time spent in bush or scrub in Whiteman Park?
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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11. In the past year, 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
12. In the past year, How often do you find ticks on yourself after time spent in the bush in Whiteman Park?
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
13. 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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14. In the past 1 year, 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 in Whiteman Park.
Bushwalking, hiking, camping or working Bushwalking, hiking, camping or working in the bush or forest in other parts of in the bush or forest overseas (outside Western Australia (except for Australia) Whiteman Park) or interstate
(please tick the answer (please tick the answer that most applies to you) that most applies to you)
Not at all in past 1 year Not at all in past 1 year
Less than once a year Less than once a year Which Which state(s)/territories?………… country/countries?…………
About once a year About once a year Which Which state(s)/territories?………… country/countries?…………
More than once a year More than once a year Which state(s)/territories?………… Which country/countries?…………
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RECORD OF TICKBITES
15. IN THE PAST 1 YEAR, how often have you been bitten by ticks? This refers to tick bites that may have occurred while in Whiteman Park, 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 work at Whiteman Park” or “Not sure” to question 15, please go straight to question 16. 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)
in Whiteman Park 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 1 YEARS?
Nature of reaction Please tick any symptoms that you have had Immediate (up to a couple of days after tick bites)
- Local swelling of the skin around the Most or e very time I get bitten (more than 75% of the time) bite Often when I get bitten (25-75% of the time) Occasionally when I get bitten (Less than 25% of the time) Never Not sure
- Tenderness of the skin around the bite Most or e very time I get bitten (more than 75% of the time) Often when I get bitten (25-75% of the time) Occasionally when I get bitten (Less than 25% of the time) Never Not sure
- Discharge, pus or fluid coming out of Most or e very time I get bitten (more than 75% of the time) the skin around the bite Often when I get bitten (25-75% of the time) Occasionally when I get bitten (Less than 25% of the time) Never Not sure
- 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 214
At any time during the 2 weeks after tick bites
- Prolonged headache (> 3 days)* When did this occur? (e.g. Feb 2005)
……/……….
- Fever or chills* When did this occur? (e.g. Feb 2005)
……/……….
- Rash* TICK ALL THAT APPLY around the bite on chest, back or abdomen (stomach region) hands, arms or shoulders feet or legs on neck, face or scalp on groin or buttocks
When did this occur? (e.g. Feb 2005)
……/………. - Tender or painful „glands‟ (lumps or TICK ALL THAT APPLY nodes under the skin, usually away from in the groin the bite mark)* in the armpits in the neck other; where______
When did this occur? (e.g. Feb 2005)
……/……….
- Aching in joints, limbs, back or neck When did this occur? (e.g. Feb 2005) without reason* ……/……….
- Tiredness/ Feeling „out of energy‟ When did this occur? (e.g. Feb 2005) without reason* ……/……….
- Shortness of breath or difficulty When did this occur? (e.g. Feb 2005) breathing without reason* ……/……….
- Blistering/scabbing around skin area When did this occur? (e.g. Feb 2005) where bitten* ……/……….
If the subject answered “At any time in the 2 weeks after the tick bite” to ANY OF THE SYMPTOMS, please continue. Otherwise, please go straight to question 17.
I would like to ask you about the [*symptoms/symptoms] which occurred in ……/………. (START WITH THE EARLIEST DATE). Thinking back to the tick bites that occurred in the two weeks before these symptoms:
e. Where were you when you were bitten? in Whiteman Park elsewhere in Australia or overseas Location (eg name of area, park or district): ……………. State: ……………. Country: …………….
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f. In which part(s) of your body did the bite(s) occur? (TICK ALL THAT APPLY) on chest, back or abdomen (stomach region) hands, arms or shoulders feet or legs on neck, face or scalp on groin or buttocks
g. Did you remove the tick(s) yourself? Yes No: removed at a medical clinic No : removed by someone else: who?......
h. Did you seek medical advice for the tick bite(s)? Yes No Not sure
i. Were you treated with antibiotics after the tick bite(s)? Yes No Not sure
j. Did you miss any work because of the tick bite(s)? Yes if yes, How many days in total? ...... No NOW PROCEED TO THE NEXT DATE ON THE TABLE ABOVE: I would now like to ask you about the [*symptoms/symptoms] which occurred in ……/………. Thinking back to the tick bites that occurred in the two weeks before these symptoms:
e. Where were you when you were bitten? in Whiteman Park 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 216
NOW PROCEED TO THE NEXT DATE ON THE TABLE ABOVE: I would now like to ask you about the [*symptoms/symptoms] which occurred in ……/………. Thinking back to the tick bites that occurred in the two weeks before these symptoms:
e. Where were you when you were bitten? in Whiteman Park 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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16. APART FROM ANY SYMPTOMS YOU HAVE MENTIONED ALREADY, have you EVER experienced any of the following in the past year?
(Please tick all the symptoms you have had and answer the questions in that row. If you haven‟t had any of the following symptoms then please go straight to question 17)
Nature of Please How many times What time Do you remember tick any have these of the year being bitten by a tick reaction symptoms symptoms occurred did these anytime in the 2 that you since you started symptoms weeks before these have had working in occur? (eg: symptoms Whiteman Park? February) developed?
Skin swelling/ Yes if YES: Yes tenderness No No Not sure |_|_|_| Can‟t remember
Prolonged headache Yes if YES: Yes (> 3 days) No No Not sure |_|_|_| Can‟t remember
Rash over parts or all Yes if YES: Yes of my body No No Not sure |_|_|_| Can‟t remember
TICK ALL THAT APPLY around the bite on chest, back or abdomen (stomach region) hands, arms or shoulders feet or legs on neck, face or scalp on groin or buttocks
General aching in Yes if YES: Yes joints, limbs, back or No No neck without reason Not sure |_|_|_| Can‟t remember
Prolonged tiredness/ Yes if YES: Yes Feeling „out of energy‟ No No (> 3 days) without Not sure |_|_|_| Can‟t remember reason Fever or chills Yes if YES: Yes No No Not sure |_|_|_| Can‟t remember
Shortness of breath or Yes if YES: Yes difficulty breathing No No without reason Not sure |_|_|_| Can‟t remember
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OTHER MEDICAL HISTORY
17. Do you have any major medical conditions?
Yes No
If YES: Please describe ……………………………….……………………
……………………………………………………
18. Are you routinely on any medications or treatments?
Yes No
If YES: a. What is the name of the medication?
……………………………….……………………
……………………………………………………
b. What condition or health problem do you take this for? ……………………………….……………………
……………………………………………………
Thank you for taking the time to complete this survey.
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9.8 Control/Baseline Group Questionnaire
Identification code: |_|_|_|_|_|
RISK OF TICK-BORNE INFECTIONS IN WESTERN AUSTRALIA
Thank you for your interest and co-operation in our research.
The purpose of this questionnaire is to assess the risk of the average Western Australian resident at contracting a tick-borne bacterial infection.
Please circle the appropriate answer, tick the box or enter an answer as appropriate. Should you have any clarifications needed while completing the questionnaire, please forward them to the study facilitator present.
SCREENING QUESTION Before completing the interview, please inform the study facilitator with the answer to the question below.
S-1. In the past 5 years, have you mainly been living in Western Australia? This means living in this State without going away for longer than 12 months altogether over the last 5 years. (Single response) 1. Yes – START INTERVIEW 2. No– Thank you very much for your time. That is all we need to ask you today. 998. Unsure/Don‟t know/Can‟t remember -Thank you very much for your time. That is all we need to ask you today. 999. Refused – Thank you very much for your time. That is all we need to ask you today.
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GENERAL QUESTIONS
1. Date today: / / 2008
2. What is your gender?: Male Female
3. What is your year of birth: |_|_|_|_|
4. What is your current occupation:
…………………………………..
5. What are the main work tasks you do in this occupation?
…………………………………………………………………………………………...
………………..………………………………………………………………………......
………………………………………………………………….…………………………
6. Have you ever lived overseas for longer than 6 months? (Single response) (Single response) 1. Yes – Go to Q6.1 2. No– Go to Q7 998. Unsure/Don‟t know/Can‟t remember – Go to Q7 999. Refused – Go to Q7
6.1 Please tell me which countries and which years you lived there? (Single Response) Country: ______|_|_| Year arrived |_|_| Year departed Country: ______|_|_| Year arrived |_|_| Year departed Country: ______|_|_| Year arrived |_|_| Year departed
7. In the past 2 years, have you lived in country areas or in areas on the outskirts of town? (Single response) (Single response) 1. Yes – Go to Q7.1 2. No– Go to Q8 998. Unsure/Don‟t know/Can‟t remember – Go to Q8 999. Refused – Go to Q8
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7.1 How many months did you live in these areas over the past 2 years? (Single Response)
|_|_| months
The next section is about any pets you may have:
8. At any time in the past 2 years, have you kept any cats? (Single response) 1. Yes – Go to Q8.1 2. No– Go to Q9 998. Unsure/Don‟t know/Can‟t remember – Go to Q9 999. Refused – Go to Q9
8.1 Did any of your cats ever go into bushland or other wilderness areas? (Single Response) 1. Yes 2. No 998. Unsure/Don‟t know/Can‟t remember 999. Refused
9. At any time in the past 2 years, have you kept any dogs? (Single Response) 1. Yes – Go to Q9.1 2. No Go to Q10 998. Unsure/Don‟t know/Can‟t remember – Go to Q10 999. Refused – Go to Q10
9.1 Did any of your dogs ever go into bushland or other wilderness areas? (Single Response) 1. Yes 2. No 998. Unsure/Don‟t know/Can‟t remember 999. Refused
10. At any time in the past 2 years, have you regularly kept any other animals on your property that you have not already mentioned? (Single Response) 1. Yes – What type of animals? ______Go to Q10.1 2. No Go to Q11 998. Unsure/Don‟t know/Can‟t remember – Go to Q11 999. Refused – Go to Q11
10.1 Did any of these animals ever go into bushland or other wilderness areas? (Single Response) 1. Yes 2. No 998. Unsure/Don‟t know/Can‟t remember 999. Refused 222
11. In the past 2 years, how often have you been bushwalking, hiking, camping or working in the bush or forest in WA and other parts of Australia or overseas?
Bushwalking, hiking, camping or Bushwalking, hiking, camping or working in the bush or forest in other working in the bush or forest overseas parts of Western Australia or (outside Australia) interstate
(please tick the answer that most (please tick the answer that most applies to you) applies to you)
Not at all in past 2 years go to Not at all in past 2 years go to Q.12 Q.12
Less than once a year Less than once a year Which Which state(s)/territories?………… country/countries?…………
About once a year About once a year Which Which state(s)/territories?………… country/countries?…………
More than once a year More than once a year Which Which state(s)/territories?………… country/countries?…………
11.1. IN THE PAST 2 YEARS, how often have you been bitten by ticks?
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 Not sure
12. Except for anything you have already mentioned, in the past 2 years have you regularly come into contact with animals through your work or hobbies? (Single Response) 1. Yes – What is the work or hobby that brings to into contact with animals? What type of animals? ______Go to Q12.1
2. No Go to Q13 998. Unsure/Don‟t know/Can‟t remember – Go to Q13 223
999. Refused – Go to Q13
12.1 On average, how many hours per month are you in contact with these animals? (Single Response)
|_|_|_| hours per month
13. APART FROM ANY SYMPTOMS YOU HAVE MENTIONED ALREADY, have you EVER experienced any of the following right after a trip into the bush or after contact with animals?
(Please tick all the symptoms you have had and answer the questions in that row. If you haven‟t had any of the following symptoms then please go straight to question 14)
Nature of reaction Please tick How many times What time of Do you remember any have these the year did being bitten by a symptoms symptoms these tick anytime in the that you occurred in the symptoms 2 weeks before have had past 3 years? occur? (e.g.: these symptoms February) developed?
Skin swelling/ Yes if YES: Yes tenderness No No Not sure |_|_|_| Can‟t remember
Prolonged headache Yes if YES: Yes (> 3 days) No No Not sure |_|_|_| Can‟t remember
Rash over parts or all of Yes if YES: Yes my body No No Not sure |_|_|_| Can‟t remember
TICK ALL THAT APPLY around the bite on chest, back or abdomen (stomach region) hands, arms or shoulders feet or legs on neck, face or scalp on groin or buttocks General aching in joints, Yes if YES: Yes limbs, back or neck No No without reason Not sure |_|_|_| Can‟t remember
Prolonged tiredness/ Yes if YES: Yes Feeling „out of energy‟ (> No No 3 days) without reason Not sure |_|_|_| Can‟t remember
Fever or chills Yes if YES: Yes No No Not sure |_|_|_| Can‟t remember
Shortness of breath or Yes if YES: Yes difficulty breathing No No without reason Not sure |_|_|_| Can‟t remember
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14. Have you ever seen a doctor as a result of symptoms linked to a tick bite?
YES
NO
If YES: Month/Year of consultation
………………………………………………………………………………………
Please describe the symptoms or diagnosis and any medication prescribed by the attending physician.
………………..……………………………………………………………………
………………………………………………………………….……………………
Thank you for the time taken to complete this survey.
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