INTRODUCTION
Coxiella burnetii infection causes “Query (Q) fever” in humans and animals. Q fever is ranked as one of the top 13 priority zoonoses globally and has been designated as one of the most contagious diseases [1]. C. burnetii infects a wide variety of mammals (particularly rodents), reptiles, and birds [2]. Cattle, sheep, and goats are the primary reservoir hosts and are responsible for the majority of human infections. The primary mode of transmission in humans involves inhalation of aerosolized bacteria spread from infected reservoirs, while the primary mode of transmission in animals involves inhalation of infectious organisms and ingestion of contaminated feed and bedding [3]. In addition to aerosol transmission, ticks have been shown to exhibit vector competence in transferring pathogen to its hosts. Additionally, transstadial and transovarian transmission of C. burnetii in ticks has been reported. Moreover, ticks aid in the transmission of the pathogen to wild and domestic animals [4]. Natural infections have been reported in > 40 species of Ixodidae and Argasidae ticks [5]. The brown dog tick, Rhipicephalus sanguineus, has tested positive for C. burnetii [6]. Transmission of the Q fever agent by tick bites in humans has not been established; however, there are reports of humans acquiring Q fever infection by crushing the tick between the fingers [7] and via dried faeces containing spore-like forms of C. burnetii [3].
Rodents have been reported as reservoirs for Q fever; however, rodent contribution to pathogen maintenance, transmission, and geographic spread remains to be elucidated. Small rodents serve as an important intermediate linking the sylvatic and domestic cycles, thereby contributing to C. burnetii transmission from rodents-to-livestock and incidentally to humans [8]. In this study we have clarified the role of synanthropic rodents/shrews and their ectoparasites in the epidemiology of Q fever under natural settings in Puducherry, India. Such data are essential to assess the mode of maintenance and spread of C. burnetii and risk of human infection in the future.
MATERIALS AND METHODS
This study was conducted in 39 villages within the Union Territory of Puducherry (Fig 1). The Institutional Animal Ethics Committee (IAEC-2018/ ICMR-VCRC/P-2) approved the study. Rodents and shrews were trapped in randomly selected villages using Sherman traps.
The traps were set by 5:00 pm and retrieved by 6:30 am the next morning. As the trapped rodents and shrews were potential sources of other zoonotic infections, the trapped animals were immobilized by exposure to chloroform to avoid accidental handling injuries. The anesthetized animals were euthanized by injecting an overdose of pentobarbital sodium (250 mg/kg) via the intraperitoneal route. The euthanized rodents were identified after recording their morphologic features [9]. Blood samples (0.5-1.0 ml) were collected from rodents and shrews via direct cardiac puncture using sterile syringes. The ears, snout, axillary regions, and limbs of individual rodents/shrews were examined under a stereo microscope. Ectoparasites were retrieved using thin tweezers and preserved in labelled vials containing 70% ethanol. The ectoparasites were identified based on the morphologic characteristics using standardard taxonomic keys [10–15]. The following formula was used to calculate the ectoparasite index: ectoparasite index = total number of ectoparasites collected/total number of animals examined.
A commercially available DNA extraction kit (GenElute Blood Genomic DNA kit; Sigma-Aldrich, St. Louis, MO, USA) was used to extract DNA from the ectoparasites and rodent blood samples following the manufacturerˈs protocols. Preliminary screening for C. burnetii in rodent/shrew and ectoparasite DNA was carried out using real-time PCR by targeting the 70-bp fragment of the IS 1111 gene [16]. The samples that tested negative by real-time PCR were re-screened for C. burnetii according to the published protocol of Dhaka et al. [17] and De Bruin et al. [18] and Zhang et al. [19] to amplify the IS1111 and com 1 genes, respectively. A C. burnetii-positive DNA sample (kindly provided by Dr. Stephen Selvaraj, Professor of Microbiology at MGAMRI, Puducherry) was used as a PCR-positive control and standardization of all PCR assays.
RESULTS
Details of the trapped rodents/shrews and ectoparasites
In this study a total of 724 traps were placed and 140 animals were trapped (a trap-positive rate of 19.34%). Of the 140 animals trapped, 33 were identified as Rattus rattus and 107 were identified as Suncus murinus. Tick infestation was noticed in 15 animals, of which 11 were S. murinus and 4 were R. rattus. Mite infestation was detected in 89 trapped animals, of which 79 were S. murinus and 10 were R. rattus. Flea infestation was detected in one R. rattus. In total, 57 ticks, 3290 mites, and 6 fleas were collected. The tick-, mite-, and flea-positive rates are shown in Table 1. The retrieved ticks were identified as Rhipicephalus sanguineus, the mites were identified as Leptotrombidium deliense and Schoengastiella spp., and the fleas were identified as Xenopsylla cheopis.
Ectoparasite positivity rate and index in the trapped animals.
| Species of rodents/shrews trapped (No. of animals trapped) | Number of animals positive for ticks | Tick positivity rate | Number of ticks collected | Tick index | Number of animals positive for mites | Mite positivity rate | Number of mites collected | Mite index | Number of animals positive for fleas | Flea positivity rate | Number of fleas collected | Flea index |
|---|---|---|---|---|---|---|---|---|---|---|---|---|
|
Rattus rattus
(n=33) | 4 | 12.12% | 7 | 0.21 | 10 | 30.3% | 7 | 0.21 | 1 | 3.0% | 6 | 0.18 |
|
Suncus murinus
(n=107) | 11 | 10.28% | 50 | 0.47 | 79 | 73.83% | 3283 | 30.68 | 0 | 0.0% | 0 | 0.0 |
Molecular detection of C. burnetii in rodent/shrew and ectoparasite DNA samples
The DNA samples from 140 rodents and shrews, 45 ticks (9 pools), 1375 mites (55 pools), and 6 fleas (1 pool) tested C. burnetii negative using real-time and conventional PCR methods (Fig 2A–D).
(A) Real-time PCR screening of C. burnetii in rodents/shrews and their ectoparasites targeting the IS 1111 gene. (B) Results of screening C. burnetii in rodents and their ectoparasites by PCR targeting the IS1111 gene. (C) Results of screening C. burnetii in rodents and their ectoparasites by trans- PCR targeting the IS1111 gene. (D) Results of screening C. burnetii in a rodent by PCR targeting the com1 gene.
DISCUSSION
The lack of C. burnetii in the present study is in agreement with the results obtained by Sahu et al. [20], who reported that of 38 rodents collected from paddy fields adjoining the goat farms in Chattishgarh and Odisha, none were positive for C. burnetii [20]. Pluta et al. [21] did not detect C. burnetii among 119 rodents trapped from 3 Q fever endemic areas in southern Germany. Similarly, Minichova et al. [22] reported zero prevalence of C. burnetii in rodents in Slovakia. However, Reusken et al. [23] reported the presence of C. burnetii in black and brown rats trapped from animal farms located close to bulk milk-positive goat farms associated with a Q fever outbreak in The Netherlands. Gonzalez et al. [24] reported the presence of C. burnetii in micromammals, such as Apodemus spp., Crocidura spp. and Rattus rattus in Spain. Alotaibi et al. [25] demonstrated C. burnetii DNA in 17.5% of rodents trapped in Saudi Arabia. The absence of C. burnetii in rodents and shrews in the current study could be due to a lack of pathogen exposure from infected livestock or neutralization and clearance of the pathogen by the antibodies developed in exposed rodents/shrews.
In addition to the molecular evidence, exposure to C. burnetii in rodents was also confirmed based on serologic results. Meredith et al. [26] reported an overall seroprevalence of 17.3% among rodents in the UK (range, 15.6%–19.1%) based on species. Hence, a sero-surveillance in rodents/shrews in the current study would have helped confirm exposure to C. burnetii; however, this remains a major limitation.
None of the ticks (n=45), mites (n=1375), and fleas (n=5) infesting rodents and shrews tested positive for C. burnetii. Our findings are consistent with earlier reports of C. burnetii absence among 8593 tick samples in Slovakia [22]. Similarly, Kamani et al. [27] did not detect C. burnetii in rodent ticks (Rhipicephalus sanguineus), mites (Haemolaelaps spp. and Hemimerus talpoides), and fleas (Xenopsylla cheopis and Ctenophthalmus spp.) in Nigeria. Our findings suggest that the natural foci of C. burnetii are limited, which accounted for our negative results in the ectoparasites.
The IS1111 gene is a transposase-like insertion sequence with a wide range of copy numbers (7–100 copies per genome), offering higher sensitivity of C. burnetii detection by PCR. It has been reported that ticks also harbor Coxiella-like endosymbionts (CLEs), which may also test positive for the IS1111 gene by PCR, leading to false-positive reports of C. burnetii [28]. Therefore, we also performed screening with PCR targeting the com1 gene, which encodes outer membrane protein 1 with a single copy in the C. burnetii genome [4]. To rule out non-specific amplifications, a standard positive control was also used. We believe that the absence of C. burnetii in rodents and their ectorparasites might be due to the absence of the pathogen or very low-copy numbers of pathogen DNA.
Given that serologic and molecular evidence in Puducherry indicated exposure to C. burnetii in sheep, goats [29], and a buffalo [30], future longitudinal studies with serologic and molecular markers are warranted in rodents. Such investigations will help to delineate the factors facilitating the sustenance and spread of C. burnetii in Puducherry.
Overall, our findings indicate that the enzootic maintenance of C. burnetii and its transmission via ticks in rodents/shrews has a minor role in the tansmission of Q fever to animals and humans in Puducherry compared to the major route of transmission by aerosol from the infected livestock. Further longitudinal studies are warranted to delineate the influence of seasonal variations and the role of rodents and their ectoparasites in Q fever disease dynamics.