Detection of Coxiella burnetii DNA in Wildlife and Ticks in Northern Queensland,Australia

Abstract

Wild animals and the tick species that feed on them form the natural transmission cycle and reservoir of Coxiella burnetii. The objective of this study was to determine whether C. burnetii was present in the blood of host animals and their ticks in northern Queensland, Australia. Three genomic targets were detected using real-time PCR assays—the Coxiella-specific outer membrane protein coding gene (Com1), the multicopy insertion element (IS1111), and the isocitrate dehydrogenase gene (Icd). Quantification of the single-copy targets identified a range of 1.48×10¹ to 4.10×10³ C. burnetii genome equivalents per microliter in the ticks tested. The detection of Coxiella based on the presence of the genomic targets indicated the occurrence of C. burnetii in both the ticks and whole blood of a variety of native Australian marsupials and confirms these animals are capable of acting as reservoirs of Q fever in northern Queensland.

Introduction

W ild animals and the tick species that feed on them form the natural transmission cycle and reservoir of Coxiella burnetii (Babudieri 1959). The bacterium has been isolated from various tick species that feed on a wide variety of vertebrate hosts. The most frequently reported association is with Ixodes ricinus ticks in a number of European studies (Rehácek et al. 1994, Reye et al. 2010). C. burnetii is vertically transmitted from adult to nymph during egg production in some tick species, resulting in a self-perpetuating reservoir for C. burnetii (Pandurov and Zaprianov 1975, Weyer 1975, Daiter 1977). The host promiscuity of many species of ticks that feed on wild animals may result in the transmission of C. burnetii to domestic animals in endemic areas. C. burnetii has been detected in a variety of tick species in Australia. Early investigations by Smith and Derrick (1940) demonstrated the presence of C. burnetii in Haemaphysalis humerosa (1939) and Ixodes holocyclus (Smith 1942) collected from bandicoots (Isoodon macrourus) in southeastern Queensland. The transmission of C. burnetii by these tick species was also demonstrated in bandicoots (Derrick and Smith 1940, Smith 1942). H. humerosa is primarily a bandicoot tick that is rarely associated with other host species, whereas I. holocyclus is more promiscuous. This tick represents a potential vector for the transmission of C. burnetii from natural hosts to domestic animals, livestock, and humans. Another tick species of importance as a reservoir for C. burnetii is Amblyomma triguttatum. This tick is primarily found on macropodids, but is also promiscuous in host species and has a wide distribution across Australia (Roberts 1970, McDiarmid et al. 2000). In more recent studies, C. burnetii has been detected in another Australian tick species not previously investigated, Bothriocroton auruginans (Vilcins et al. 2009). However, this tick species is unlikely to be an important vector for C. burnetii due to its limited host range in wombats.

C. burnetii has been found to be present in the gut lumen and epithelial lining of these Australian ticks (Smith 1942). Moreover, transovarial passage of C. burnetii has not been observed in Australian tick species investigated. Although tick species known to be potential vectors of C. burnetii are capable of feeding on humans, this route of infection is a rare source of Q fever cases (Lang 1990). Because C. burnetii replicates primarily in the digestive tract of ticks, it is expelled in the feces during feeding, which can result in heavy contamination of the skin of host animals with Coxieallae (Lang 1990). This mechanism of transmission has been proposed as a source of infection in Q fever epidemics (Derrick et al. 1959, Pope et al. 1960). Dried feces from infected ticks have been found to harbor large quantities of phase I C. burnetii and remain infective for approximately 2 years (Stoker and Marmion 1955). In addition, it has been demonstrated that ticks infected with C. burnetii can remain infected for several years and, in some cases, for the life span of the tick (Stoker and Marmion 1955).

Investigations of tick reservoirs of C. burnetii have been performed in Australia previously; however, no investigations have been performed since the 1960s and none have been performed in tropical northern Queensland. Both the potential host and the tick species associated with C. burnetii transmission in past investigations are present in northern Queensland. Current data are required in determining their potential role in Q fever epidemiology in the region, thus this study aimed to determine whether C. burnetii was present in ticks and the blood of host animals in northern Queensland.

Materials and Methods

Collection of blood from Australian native marsupials

Blood samples were collected during trapping of small and noncommercially harvested species and postmortems from commercially harvested species. Trapped animals were captured according to procedures used by the Queensland Parks and Wildlife Service using treadle traps lined with foam and shade cloth. All care was taken to reduce stress on the animals. Blood samples (equivalent to less than 0.5% of the body weight to a maximum 2 mL) collected from each identified animal were taken from the lateral coccygeal vein of the tail with the animal manually restrained in a hessian (burlap) sack. Following blood collection, animals were released at the site at which they were captured. A 200-μL aliquot was removed from each sample prior to centrifugation and stored at −20°C for subsequent DNA extraction. Blood samples from kangaroos were collected via cardiac puncture from animals shot by licensed shooters during routine culling expeditions. Harvesters held a commercial wildlife-harvesting license. A 200-μL aliquot of blood was removed and stored as described for trapped animal samples. Blood samples were collected in 6-mL clot-activated tubes, allowed to clot, and then centrifuged at 1400×g to collect serum. Serum was stored at −20°C for subsequent serological testing.

Collection of ticks from Australian native marsupials

Up to 10 ticks in either or both nymphal and adult stages were collected from each animal using forceps. Ticks were placed in absolute ethanol in 5-mL sample tubes. The species and area where the animals were sampled was recorded. Ticks were then identified to the species level using a phenotypic key (Roberts 1970).

Extraction of DNA from whole blood of Australian native marsupials

Frozen whole blood samples were thawed and DNA was extracted using a HighPure™ PCR Template Preparation Kit (Roche Diagnostics, Germany) DNA extraction kit according to the manufacturer's instructions for extraction of DNA from tissue. DNA was eluted with molecular biology–grade water (Sigma, Australia) preheated to 70°C. Purified DNA was stored at −20°C for subsequent analysis.

Extraction of DNA from ticks collected from Australian native marsupials

Ticks were treated individually due to size or engorgement, or pooled according to host animal. Ticks were washed with 70% ethanol, air dried for 10 min on sterile paper, and then finely diced with a sterile scalpel blade on a sterile glass slide. DNA was extracted using a HighPure™ PCR Template Preparation Kit (Roche Diagnostics, Germany) DNA extraction kit according to the manufacturer's instructions. DNA was eluted with molecular biology–grade water (Sigma, Australia) preheated to 70°C. Purified DNA was then stored at −20°C prior to further analysis.

Real-time PCR analysis

Primer sets targeting the Com1, IS1111 (Marmion et al. 2005), and Icd (Klee et al. 2006) genes in C. burnetii were obtained from the literature and validated using GeneDoc software. In our laboratory, the detection limits of these assays were determined to be approximately 8 genome equivalents for the Com1 gene, 1 for IS1111, and 20 organisms for the Icd gene. Reactions consisted of 1× Immomix (Bioline, Austalia), 10 μM SYTO-9 (Invitrogen, Australia), and 300 nM primers. PCR reactions with a volume of 10 μL were prepared for a 72-well rotor in a RotorGene 6000 (Corbett Research, Australia). Each sample was tested in duplicate with 1-μL sample per PCR reaction. Standards (genomic standard NMII/C4 C. burnetii) were included, as were positive (tick lysates spiked with NMII/C4) and no-template controls. PCR inhibition in the tick extracts was tested by spiking the extracts with known quantities of the PCR standards and examining results for reduced output. Cycling conditions for qPCR consisted of an initial denaturation for 10 min at 94°C, followed by 40 cycles of denaturation for 10 sec at 94°C, annealing of primers for 10 sec at 62°C, and extension for 20 sec at 72°C. A melt curve analysis was then performed with an increase in temperature from 72°C to 95°C in 1°C increments.

Sequencing of PCR products

To sequence the Com1 PCR product, reactions with a volume of 20 μL were prepared for a 36-well rotor in a RotorGene 6000 (Corbett Research, Australia) as described previously. Representative samples for each melt curve profile were amplified in 10 tubes each. PCR products were quantified using a NanoPhotometer™ (Implen, Germany) and diluted to 100 ng μL⁻¹ in 40-μL aliquots in O-ring–sealed microcentrifuge tubes. Sequencing of PCR products was performed by Macrogen Inc. (Korea) using Com1 primers provided.

Serological analyses

Formalin-inactivated whole-cell C. burnetii antigen of both antigenic phases was prepared as previously described (Cooper et al. 2011a) and used to coat plates for serological testing of marsupial serum. Both phase I and phase II C. burnetii antigens were produced using an Australian C. burnetii isolate (Cumberland strain). This isolate was obtained from the Australian Rickettsial Reference Laboratory (Geelong, Victoria), and was isolated from a patient who contracted Q fever through contact with beef cattle.

Competitive enzyme-linked immunosorbent assays (ELISAs) were optimized, validated, and performed as previously described (Cooper et al. 2011b). Briefly, NUNC™ 96-well Maxisorp plates were coated with 50 μL of phase II or phase I antigen at 25 μg mL⁻¹ in carbonate/bicarbonate coating buffer (pH 9.0) and incubated overnight at 37°C. Plates were coated with 50 μL of postcoating buffer (Tropbio, Australia), incubated at room temperature for 2 h, and dried. Test sera were applied at a dilution of 1:100 in 50-μL aliquots in duplicate and incubated at 37°C for 1 h. The wells were washed three times with phosphate-buffered saline + Tween 20 (PBS-T), after which 50 μL horseradish peroxidase (HRP)-conjugated rabbit anti-bovine immunoglobulin (Ig; Serotec, UK) at 1:1000 was applied and incubated at 37°C for 1 h. The wells were washed again, after which 50 μL ABTS buffer was applied and incubated at 37°C for 30 min. Optical density readings were obtained using a Multiskan Ascent plate reader at 414/494 nm. A reduction in optimal density of >70% from that of the indicator serum alone was considered to be a positive result. Results of duplicates for each sample were averaged to produce a mean result for each animal.

Statistical analyses

Chi-squared tests were performed to determine whether there was any association between the presence of DNA in ticks, presence of DNA in blood samples from native marsupials, and seropositivity in the same animals.

Results

Whole blood was collected from 35 common northern bandicoots (I. macrourus), 17 eastern grey kangaroos (Macropus giganteus), 5 agile wallabies (Macropus agilis), 4 red kangaroos (Macropus rufus), 3 common wallaroos (Macropus robustus), 2 brushtail possums (Trichosurus vulpecula), 1 black striped wallaby (Macropus dorsalis), and 1 rufous bettong (Aepyprymnus rufescens).

A total of 280 ticks were collected from 34 common northern bandicoots (I. macrourus). Of these, 250 were identified as H. humerosa and 30 as I. holocyclus. These were divided according to host animal into 38 pools of H. humerosa and three pools of I. holocyclus. A total of 43 A. triguttatum specimens were collected from 9 eastern grey kangaroos (M. giganteus), 4 agile wallabies (M. agilis), and 1 rufous bettong (A. rufescens).

All A. triguttatum samples were tested separately due to the size of the specimens. A summary of all samples collected is detailed in Table 1. Tick species containing C. burnetii DNA included I. holocyclus collected from the common northern bandicoot (10/30) and A. triguttatum collected from the eastern grey kangaroo (12/31). Although H. humerosa collected from the common northern bandicoot made up the large majority of total ticks collected (250/323), all were negative for the presence of C. burnetii DNA.

Table 1.

Summary of Amplicon Types Detected in Ticks and Whole Blood Collected from Australian Native Marsupials

Host animal			Ticks collected from host animal
Species	Number	PCR positive (blood)	Species	Number	PCR positive
Common northern bandicoot (Isoodon macrourus)	35	6	Haemaphysalis humerosa	250	0
			Ixodes holocyclus	30	10
Eastern grey kangaroo (Macropus giganteus)	17	6	Amblyomma triguttatum	31	12
Agile wallaby (Macropus agilis)	5	1	Amblyomma triguttatum	6	0
Red kangaroo (Macropus rufus)	4	1	No ticks collected	0	NA
Common wallaroo (Macropus robustus)	3	1	Amblyomma triguttatum	4	0
Brushtail possum (Trichosurus vulpecula)	2	1	No ticks collected	0	NA
Rufous bettong (Aepyprymnus rufescens)	1	0	Amblyomma triguttatum	2	0
Black-striped wallaby (Macropus dorsalis)	1	1	No ticks collected	0	NA

NA, Not applicable.

Quantification of the single-copy targets (Com1 and Icd) identified a range of 1.48×10¹ to 4.10×10³ genome equivalents per microliter of tick extract. All tick samples that were positive for the Com1 gene amplicon also contained the Icd amplicon. However, the IS1111 gene target was only observed in 60% of these same samples. To test for PCR inhibitors in the tick preparations, they were spiked with the C. burnetii used to prepare the standard curves using concentrations from 10¹ to 10⁶ genome equivalents. No inhibition of the PCR was observed in the tick extracts at any of these concentrations.

No association was found between the detection of C. burnetii DNA in the tick samples collected from a host animal and seropositivity in serum of the host animal, or detection of C. burnetii DNA in the tick sample and detection of C. burnetii DNA in the whole blood sample for each animal. There was also no association found between the detection of DNA in the whole blood sample and seropositivity in the serum or the detection of DNA in the whole blood samples and detection of DNA in ticks. A summary of seropositivity to phase II and I antigens is included in Table 2. Sequencing results indicated that the Com1 amplicon matched the sequence for Com1 from Nine Mile I C. burnetii (AE016828.2), with homology of 98%.

Table 2.

Summary of Seropositivity for Australian Native Marsupials

Host animal		Seropositivity (n)
Species	Number	Phase II (%)	Phase I (%)	Phase II and/or Phase I (%)
Common northern bandicoot (Isoodon macrourus)	35	10 (28.5)	9 (25.7)	11 (31.4)
Eastern grey kangaroo (Macropus giganteus)	17	5 (29.4)	7 (41.1)	7 (41.1)
Agile wallaby (Macropus agilis)	5	2 (40.0)	3 (60.0)	3 (60.0)
Red kangaroo (Macropus rufus)	4	0 (0.0)	0 (0.0)	0 (0.0)
Common wallaroo (Macropus robustus)	3	2 (66.6)	2 (66.6)	2 (66.6)
Brushtail possum (Trichosurus vulpecula)	2	0 (0.0)	0 (0.0)	0 (0.0)
Rufous bettong (Aepyprymnus rufescens)	1	1 (100)	0 (0.0)	1 (100)
Black-striped wallaby (Macropus dorsalis)	1	1 (100)	1 (100)	1 (100)
Total number of animals tested	68	21 (30.8)	22 (32.3)	25 (36.7)

Discussion

A quantitative (q) PCR was successfully used for the detection of C. burnetii in ticks and whole blood collected from Australian native marsupials. The detection of the 3 target genes indicated the presence of C. burnetii in either or both the ticks and whole blood of bandicoots and a variety of macropods in northern Queensland. The presence of C. burnetii in both the ticks and whole blood of Australian native marsupials suggests these animals are capable of acting as reservoirs of Q fever in northern Queensland.

The current study confirmed the presence of C. burnetii in the ticks of bandicoots and macropods via the detection of C. burnetii DNA. This study also represents the first known detection of C. burnetii in blood of the agile wallaby, common wallaroo, and rufous bettong. While detection of the Coxiella-specific genes in tick extracts indicates the presence of C. burnetii, detection via PCR is unable to distinguish between localization in the blood meal or tick organs.

Almost one-third of animals tested contained antibodies to phase 1 antigens, suggesting that these animals were previously infected with C. burnetii. However, antibody responses to C. burnetii in the animals tested were highly heterogeneous, a finding that is consistent with human Q fever serology. Such heterogeneity of the antibody response complicates serological investigations and epidemiological studies. Although serological surveys have provided the bulk of epidemiological data on Q fever, no standardized technique has been employed and there is a large degree of variability between the serological tests used. Further work is required in animal serology to determine the pattern of antibody production in response to C. burnetii. Such patterns may differ between species and could indicate relative time since exposure and chronicity of infection in the host. Further work would also be required to establish the identity of the immunoglobulin subsets detected in the current study, as isotype-specific and subclass-specific regents are not currently available for native Australian marsupials.

Although C. burnetii DNA was detected in the blood and ticks of the native Australian marsupials surveyed in this study, there was no association between the presence of C. burnetii DNA in either ticks or blood and seropositivity. There was also no association found between the presence of C. burnetii DNA in ticks and presence of C. burnetii DNA in blood. However, these results were not unexpected, because bacteremia is often transient in animals (Mantovani and Benazzi 1953) and not necessarily coincidental with seroconversion (McQuiston and Childs 2002).

The detection of C. burnetii DNA in both ticks and whole blood of native Australian marsupials indicates transmission of C. burnetii between these species in northern Queensland. This detection also indicates these species may be reservoirs of C. burnetii in the region. Both A. triguttatum and I. holocyclus have promiscuous host ranges, and the detection of C. burnetii DNA in these species indicates they may be capable of transmitting C. burnetii to a wide range of host species, including livestock, domestic animals, and their feral counterparts. Although this study was not comprehensive, the results suggest that these domestic animals are a potential source of human infection. This investigation, which acts as a proof of principle study, has demonstrated the possibility of native Australian animals and their ticks to be hosts and vectors of C. burnetii. On the basis of these results, a more comprehensive survey is warranted.

In conclusion, the detection of C. burnetii in both the ticks and whole blood of Australian native marsupials confirmed that these animals were capable of acting as reservoirs of Q fever in northern Queensland. The confirmation of the presence of C. burnetii in these species indicates they may be the source of human Q fever cases where no contact with typical reservoir species is reported. The detection of C. burnetii in bandicoots and possums is particularly important, due to the adaptation of these species to urban environments. The detection of C. burnetii in macropods in the region may also have public health implications due to these animals often ranging into outlying suburban areas that border bushland. The evidence of C. burnetii in macropod ticks, in addition to the findings of another Australian study indicating macropods shed C. burnetii in their feces (Banazis et al. 2010), suggests that these animals may act as a vector of C. burnetii to domestic animals and the human population.

Footnotes

Acknowledgments

The authors would like to thank Associate Professor Leigh Owens for assistance with statistical analyses. The authors would also like to thank the professional kangaroo harvesters involved in this study for their assistance in the collection of samples.

Author Disclosure Statement

No competing financial interests exist.

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