Critical Aspects for Detection of Coxiella burnetii

Abstract

Coxiella burnetii is a globally distributed zoonotic γ-proteobacterium with an obligatory intracellular lifestyle. It is the causative agent of Q fever in humans and of coxiellosis among ruminants, although the agent is also detected in ticks, birds, and various other mammalian species. Requirements for intracellular multiplication together with the necessity for biosafety level 3 facilities restrict the cultivation of C. burnetii to specialized laboratories. Development of a novel medium formulation enabling axenic growth of C. burnetii has facilitated fundamental genetic studies. This review provides critical insights into direct diagnostic methods currently available for C. burnetii. It encompasses molecular detection methods, isolation, and propagation of the bacteria and its genetic characterization. Differentiation of C. burnetii from Coxiella-like organisms is an essential diagnostic prerequisite, particularly when handling and analyzing ticks.

Introduction

C oxiella burnetii is a globally distributed zoonotic γ-proteobacterium whose economic and health importance has recently been underscored following the largest ever reported outbreak, which has occurred in the Netherlands (Roest et al. 2011). C. burnetii possesses several remarkable features, including the ability for proliferation within phagolysosome-like vacuoles of mononuclear phagocytes, a biphasic developmental life cycle, and a lipopolysaccharide phase variation (van Schaik et al. 2013). Infections can either be asymptomatic or result in clinical disease. In humans, the disease is known as Q fever and varies from uncomplicated and self-limited febrile illness (acute Q fever) to long-lasting usually focal disease (chronic Q fever), which may result in fatality (Maurin and Raoult 1999, Million and Raoult 2015). Known as coxiellosis in animals, the disease predominantly manifests as reproductive disorder (Agerholm 2013). Sporadic or clustered cases and large outbreaks have been described worldwide in both humans and animals (Smith 1989, Gilroy et al. 2001, Amitai et al. 2010, Roest et al. 2011, Georgiev et al. 2013).

C. burnetii can infect ticks, birds, and mammals. Ticks are regarded as important vectors for agent transmission between wild animals and for amplification of enzootic cycles to the domestic environment (Cutler et al. 2007, Boarbi et al. 2015). Aerogenic transmission following environmental contamination has been demonstrated between flocks/herds and has resulted in human outbreaks (Hawker et al. 1998); however, direct contact between and with infected animals additionally facilitates spread (Kruszewska and Tylewska-Wierzbanowska 1997, Alsaleh et al. 2011). C. burnetii is excreted in vast numbers during normal parturition as well as abortion. Once aerosolized, the bacteria can be transmitted over long distances by the wind. During the biphasic developmental life cycle, C. burnetii develops highly resistant spore-like structures known as small cell variants (SCVs) providing long-lasting environmental stability. Other body fluids and secretions are also infectious and may facilitate both vertical and sexual transmission (Kruszewska and Tylewska-Wierzbanowska 1997, Maurin and Raoult 1999, Milazzo et al. 2001, Miceli et al. 2010, Agerholm 2013). Small domestic ruminants are the most frequently infected species and are considered as the primary source of human infections.

Eight decades after the first description of Q fever cases, diagnosis remains challenging. Case confirmation in humans and appropriate surveillance of animals depend mostly on the interest of the involved clinician/veterinarian and their diagnostic capabilities, mostly relying upon serology. In this review, direct laboratory detection tests for C. burnetii will be reviewed, especially molecular diagnostic methods and recent improvements in pathogen isolation methods.

Real-time PCR

DNA amplification is most frequently used for direct detection of C. burnetii. This enables investigation of all sample types from vertebrates to ticks and environmental samples such as dust, soil, and water. For acute human cases, whole-blood or buffy coat aliquots collected in EDTA or citrate at onset of symptoms and before antibiotic treatment are most useful (Anderson et al. 2013). Serum, urine, and throat swabs have also proven to be valuable for C. burnetii screening (Klaassen et al. 2009). In more protracted infections, tissue samples from focal regions of infection should be investigated, that is, valvular material from endocarditis, aneurism, or vessel fragments in vascular infections, and bone biopsies in osteomyelitis. For livestock, aborted material (placental material and fetal organs), milk, vaginal swabs, feces, and more rarely semen have proven to be valuable. On a cautionary note, if the herd has been recently vaccinated (first month following vaccination), PCR will not discriminate between the vaccine and wild-type strains (Hermans et al. 2011). As C. burnetii is shed intermittently, consecutive samples are preferred to single collections. Bulk tank milk is recommended for herd monitoring rather than individual samples because of its ease of collection, cost-effectiveness, reduced contamination, and sensitivity for evaluation of the pathogen at the herd level. However, a single collection is not sufficient for detection of C. burnetii in flocks with low numbers of infected animals. Therefore, two to three samples (collected two to three months apart) are more informative (Boarbi et al. 2014). For wildlife screening, blood, urine, feces, vaginal, cloacae, and anal swaps can be useful (Bittar et al. 2014, Tozer et al. 2014, González-Barrio et al. 2015b). In case of dead animals (hunted, road-killed, euthanized, etc.), other samples such as spleen, lung, and liver should also be considered. As for domestic animals, short bacteremia and intermittent shedding can also occur, thus the collection of different sample types obtained during longer sampling periods serves to overcome seasonal fluctuations of C. burnetii in wildlife (González-Barrio et al. 2015a).

For DNA extraction, fresh or frozen samples are preferable, although paraffin-embedded tissues have also been used successfully for the identification of chronic Q fever patients (Costa et al. 2015). DNA extraction protocols vary from column to magnetic particle-based methods. In either case, PCR general guidelines should be rigorously followed to limit sample cross-contamination that might occur when high C. burnetii loads are present. Bacterial numbers are highly variable, with massive C. burnetii burdens in persistently infected tissue samples (as placental/fetal and valvular/vascular material) to very low agent loads in environmental samples, milk samples, and usually in blood samples. For DNA amplification, several real-time PCR protocols targeting different genes are described in the literature as reviewed in Table 1. These have superseded previously used conventional and nested PCRs that are prone to cross-contamination. The multicopy IS1111 repetitive element is often used for agent's detection as this provides increased sensitivity when compared with other targets, but since the exact copy number is unknown for most of the strains, except for C. burnetii Nine Mile I with 20 copies per genome, it cannot be used for quantification (Klee et al. 2006, Tilburg et al. 2010). When results are equivocal (Ct values 35 or greater), additional confirmation using another target or a different region within the same gene should be considered. Furthermore, when investigating arthropod vectors, it must be remembered that the specificity of the IS1111 real-time PCR might be compromised through detection of Coxiella-like variants (Elsa et al. 2015). Confirmation of findings can be verified when necessary by sequencing.

Table 1.

Most Cited Real-Time Assays (PubMed) for Coxiella Burnetii or Q Fever Over the Last 3 Years

			Fragment amplified (bp)
Target gene	Method	Application	<100	100–200	>200	Reference
Multiplex
IS1111, com1	TaqMan	Detection, quantification	✓			De Bruin et al. (2013)
IS1111, com 1, icd	TaqMan	Detection, quantification		✓		De Bruin et al. (2011)
Singleplex
IS1111	TaqMan	Detection	✓			Harris et al. (2000),Loftis et al. (2006),Schneeberger et al. (2010),Tilburg et al. (2010),Niemczuk et al. (2011),Schets et al. (2013),Di Domenico et al. (2014),Niemczuk et al. (2014)
	TaqMan	Detection		✓		Brouqui et al. (2005),Kim et al. (2005),Fournier et al. (2010),Tilburg et al. (2010)
	TaqMan	Detection			✓	Klee et al. (2006),Barkallah et al. (2014)
	Hyprobe	Detection	✓			Panning et al. (2008)
	Hyprobe	Detection			✓	Stemmler and Meyer (2002)
IS30a	TaqMan	Detection, quantification		✓		Brouqui et al. (2005),Mediannikov et al. (2010)
icd	TaqMan	Detection, quantification	✓			Klee et al. (2006)
com1	TaqMan	Detection, quantification	✓			Marmion et al. (2005),Kersh et al. (2010a) and (2010b),Lockhar et al. (2011),Schets et al. (2013)
	SYBR green	Detection, quantification	✓			Cooper et al. (2007)
	Hyprobe	Detection, quantification		✓		El-Mahallawy et al. (2016)
16S rRNA	TaqMan	Detection, quantification	✓			Marmion et al. (2005)
	SYBR green	Detection, quantification	✓			Millán et al. (2016)
groES and groEL	TaqMan	Detection, quantification	✓			Elsa et al. (2015)
	TaqMan	Detection, quantification		✓		Bond et al. (2016)
ompA	TaqMan	Detection	✓			Jaton et al. (2013)
CBU_678	SYBR green	Detection				Kersh et al. (2010a)
CBU_686	SYBR green	Detection				Kersh et al. (2010a)
dotA	TaqMan	Quantification	✓			Coleman et al. (2004)
rpoS	TaqMan	Quantification	✓			Coleman et al. (2004)

Most of the real-time PCR diagnostic tests described are in-house assays, specifically adapted for detection of C. burnetii DNA in various sample types. Commercial PCR diagnostic tests are also available, but not included in this table as information on components (PCR Master Mix reagents, primer and probe sequences, PCR product lengths) is often omitted due to patent constraints.

Genome and Genetic Characterization

The first whole genome sequence of C. burnetii, from the Nine Mile RSA 493 reference strain, isolated in 1935 from an infected group of ticks (Dermacentor andersoni) was released in 2003. The sequence spans 1.995.275 base pairs and was obtained using the random shotgun method (Seshadri et al. 2003). Four years later, a second genome was published, strain Henzerling RSA 331, isolated from blood of an infected patient in Italy in 1945 (“J. Craig Venter Institute”-CVI, 2007). Later, three additional strains—« K » and « G » derived from human endocarditis and the « Dugway » rodent strain—were published (Beare et al. 2009). Comparative analysis of these genomes highlighted their diversity regarding pseudogene content and number of insertion sequence elements, possibly explaining their biological differences (Beare et al. 2009). Recently, along with the development of powerful sequencing platforms, the numbers of sequenced genomes have blossomed to more than 40, 26 being publically available (D'Amato et al. 2014, 2015, Karlsson et al. 2014, Sidi-Boumedine et al. 2014, Walter et al. 2014, Hammerl et al. 2015). Despite the large number of genome records for C. burnetii since 2003, only nine genomes are fully sequenced and annotated as closed circular genomes, the remainder are available as fragmented scaffolds, contigs, or whole genome shotgun sequences in various genome databases (www.patricbrc.org/portal/portal/patric/Home, www.ncbi.nlm.nih.gov/genome/genomes/543).

Obtaining high quality and host cell-free DNA from an intracellular organism for deep sequencing analyses is a challenging task, but has benefitted more recently from the use of axenic cultivation. When using in vitro cell cultures or embryonated hen eggs, particular care should be taken for complete removal of host DNA. Classical DNA isolation methods are suitable (as cited for real-time PCR). However, bioinformatic filters are required to subtract the host genome sequence. Depending on the degree of host DNA contamination (sometimes in excess of 60%), additional sequencing may be required to obtain a complete genomic coverage for C. burnetii (median genome length 2 Mb). Whole genome sequencing is becoming more affordable, but data analyses remain time-consuming and require specific knowledge and extra funding. Although still not used in routine diagnostics, access and use of whole genome sequence data are steadily increasing and tools for outbreak investigations and trace-back studies applicable in routine diagnostic laboratories will become available. Till then, traditional genotyping approaches are the best choice. Genotyping methods for C. burnetii were fully revised elsewhere (Massung et al. 2012) and therefore will be only briefly described in this review.

The choice of the most appropriate typing option may depend on the research objectives. The simplest and direct tests (lacking further sequencing), with good discriminatory power and lowest DNA demands, are mostly used for rapid tracking of outbreaks. Examples include the multiple-locus variable-number tandem repeat analysis (MLVA), particularly applicable when adapted to capillary electrophoresis for estimation of the number of repeats (Klaassen et al. 2009, Tilburg et al. 2012a), and single-nucleotide-polymorphism (SNP) genotyping (Hornstra et al. 2011, Huijsmans et al. 2011). Both typing approaches were used for the Dutch outbreak investigation (Klaassen et al. 2009, Huijsmans et al. 2011, Tilburg et al. 2012a). Presently, these methods are reviewed toward harmonization and standardized nomenclature (http://mlva.u-psud.fr/mlvav4/genotyping/view.php, Hornstra et al. 2011, Huijsmans et al. 2011).

A more robust and conservative typing system preferably supported by large databases and broadly accepted/used would provide the best overall option for eco-epidemiological investigations and data integration, at both local and global scales. Multispacer sequence typing (MST) is a good example of this case (Glazunova et al. 2005, Tilburg et al. 2012b). It has the advantage of using standardized nomenclature and genotypes can be identified using a web-based MST database (http://ifr48.timone.univ-mrs.fr/mst/coxiella_burnetii/), enabling comparison of results between laboratories.

Cultivation

Although cultivation is not usually required for a definitive diagnosis, it is valuable when new clinical presentations or atypical epidemiological situations in association with a C. burnetii infection occur. Isolation and propagation from clinical samples enable phenotypic and genotypic characterization using molecular typing methods or deeper genetic analyses such as whole genome sequencing. Cultivation is also of paramount importance to build strain collections to aid further research. It is laborious, time-consuming, and success largely depends upon sample quality, freshness, and pathogen load. Furthermore, technical expertise and availability of suitable laboratory biosafety level 3 facilities are essential. Handling and processing of samples or cultures with a high bacterial load bear the risk of generating contaminated aerosols and sets that involved personnel at risk as demonstrated by several laboratory-acquired infections (Johnson and Kadull 1966, Curet and Paust 1972, Hall et al. 1982, Graham et al. 1989, Wurtz et al. 2016). Despite this, increasing numbers of isolates are now available.

Isolation from Clinical Samples

In vitro isolation

Several in vitro cell lines support C. burnetii replication, including those from macrophage (P388D1, J774, DH82), fibroblast (L929, HEL), and epithelial lineages (Vero E6) (Maurin and Raoult 1999, Mediannikov et al. 2010, Santos et al. 2012). The human embryonic lung fibroblast cell line—HEL—is one of the most widely used as it is easy to maintain, preserves monolayer integrity during prolonged incubations, and is highly susceptible to infection (Gouriet et al. 2005, Lagier et al. 2015). The canine malignant histiocytic macrophage cell line—DH82 (ATCC CRL-10389)—traditionally used for culturing other mononuclear leucocytes targeting bacteria, such as Ehrlichia canis and E. chaffeensis, has been increasingly adopted as an in vitro system for C. burnetii (Mediannikov et al. 2010, Lockhart et al. 2012, Santos et al. 2012, Cumbassa et al. 2015). In vitro isolation is usually performed using the shell vial technique (Gouriet et al. 2005, Santos et al. 2012). Cultures are incubated at 37°C and 5% CO₂ atmosphere for 2 months possibly extending up to 4–5 months, with periodical evaluation of microbial growth using either light or fluorescence microscopy. During this period, supplementation by partial replacement of culture medium is required with a frequency adapted according to the cell line in use. Fetal bovine serum (FBS) concentration can be reduced to 5% (v/v) in culture medium to decrease cell proliferation and maintain monolayer longevity. Appearance of parasitophorous vacuoles can be checked directly using an inverted microscope (magnification 20–40× ). Monthly assessment of culture aliquots should also be undertaken with initial cytoconcentration, stained by Gimenez, and examined by microscopy (by immersion at 1000× ) for the characteristic tightly packed C. burnetii vacuoles (Gimenez 1964). Positive findings should be confirmed by PCR (see above Real-time PCR section).

Various fresh or frozen samples (≤−80°C) can be used with the shell vial technique, including anticoagulated whole-blood, buffy coat, other biological fluids, tissue biopsies or necropsies, and ticks, etc. Fluids are directly inoculated, while tissue samples should be macerated with a pestle or disrupted with a scalpel in culture medium before being inoculated into the shell vial. An important prerequisite is the absence of microbial contaminants, which is challenging when working with postmortem or aborted tissues, ticks, and environmental samples. Ticks can be surface decontaminated by serial passages in bleach 10% and/or alcohol 70% and rinsed in sterile water before further manipulations. For placenta, fetal, and other samples that are associated with high C. burnetii loads (Ct values <25), a tissue homogenate filtration step can increase recovery. Briefly, samples are homogenized in FBS-free medium and exposed to frozen–thaw cycles and low-speed centrifugation, with the resulting supernatant subjected to sequential filtration, using 1 and 0.45-μm syringe filters, and directly inoculated into shell vials. During the initial days of cultivation, a broad-spectrum antibiotic–antifungal cocktail containing 10,000 units/mL of penicillin, 10,000 μg/mL of streptomycin, and 25 μg/mL of Fungizone^® (amphotericin B) can be added to culture medium to limit unwanted microbial growth.

In vivo isolation

In vivo isolation using rodent models, mice, or guinea pigs has proven particularly well suited for contaminated samples, such as environmental (such as ticks, etc.) or veterinary field samples, including milk or products of conception. Inoculation of the sample into a vertebrate host provides a buffer against unwanted microbial contamination. Furthermore, in vivo models are essential for maintenance of the native virulent form (phase I) of C. burnetii. The mouse strain, OF1, is the genetic lineage frequently used for isolation because of its relative sensitivity compared with either BALB/c or C57/BL6 mice (authors' experience). Milk samples should be decreamed first by simple decantation. Inoculum being aspirated from just under the fat layer can be directly injected intraperitoneally into adult >50-day mice, with volumes complying with ethical requirements. Successive injections (up to three) 5–7 days apart can be used where material permits and low microbial load is suspected (Ct values >32). For abortive material, tissues should be macerated and diluted at least twice in physiological water or PBS before injection. Following inoculation, the host should be monitored for clinical signs and by indirect serology (Mori et al. 2013), or postmortem evaluation, at 3–5 weeks postinfection. The spleen, liver, and lungs are preferred organs for C. burnetii monitoring by either microscopy or real-time PCR. Infection is typically accompanied by measurable splenomegaly caused by massive C. burnetii propagation.

Propagation of Bacterial Isolates

Embryonated egg inoculation

Propagation of highly concentrated C. burnetii cultures is achieved through the use of yolk sac infection. This method was historically used for direct isolation, but it is no longer recommended in favor of in vitro or in vivo protocols (see above). Nonetheless, it remains useful for massive propagation in specific settings (vaccine production, fundamental studies) and therefore the protocol will be briefly reviewed. Surface disinfected, 7-day-old, specific, pathogen-free chicken eggs are candled to locate the yolk sac. Once identified, the edge of the air sac should also be localized and marked on the eggshell. Inoculation with a suspension containing C. burnetii-infected material is injected through a hole drilled few mm above the marked air sac. Inoculation material might arise from in vitro or in vivo isolation procedures (see paragraphs above), including cell culture suspensions or macerated mouse organs. The latter might require a 1:2 to 1:10 dilution in physiological water or PBS prior injection. The eggshell holes are sealed with scotch tape or solvent-free glue and the eggs are incubated at 35–37°C until day 21. Bacterial growth may result in death of the embryo, but only eggs dying after day 5 postinjection are collected. Once opened, the yolk sac should be harvested by detachment, washed several times in physiological water or PBS, and then macerated and processed for further use.

Axenic media

Over the last decades, our understanding has evolved regarding the physiological and structural characteristics of the destructive phagolysosomal-like compartment with its acidic pH (∼4.5) and antimicrobial factors, such as hydrolytic and proteolytic enzymes, yet it is this same environment that provides the required intracellular niche of C. burnetii. Early studies demonstrated the necessity of an acidic pH for metabolic activation (transport of nutrients, glucose and glutamate, and intracellular replication) (Hackstadt and Williams 1981). Understanding this acid activation and the ability to decipher the metabolic pathways of C. burnetii by genome analyses led to the development an axenic medium, namely Complex Coxiella Medium, which supports metabolic activity of C. burnetii (Omsland et al. 2008, Omsland and Heinzen 2011). This axenic medium has subsequently been refined to its third-generation formulation, the defined Acidified Citrate Cysteine Medium (ACCM-D), which contains amino acids, glutamine as carbon source, and methyl-β-cyclodextrin to sequester inhibitory metabolites (Omsland et al. 2011). It has a low pH of 4.75 and cultivation requires specific microaerophilic atmosphere conditions of 5% CO₂ and 2.5% O₂ achieved by the use of a dual-gas incubator or alternatively using an anaerobic pouch in case of a monogas incubator (Omsland et al. 2009, 2011). ACCM-D supports the biphasic transition from the SCV to the replicative large cell variant of C. burnetii (Sandoz et al. 2016). Typically, there is an initial lag phase of 2 days, followed by an exponential phase until day 8 and transition into stationary phase. The second-generation formula, ACCM-2, has occasionally been used for direct isolation of C. burnetii from in vivo experimental or clinical samples (Omsland et al. 2011, Boden et al. 2015). ACCM-2 or ACCM-D may not support growth of all C. burnetii strains and therefore axenic cultivation is more frequently used for amplification of bacteria from cell culture or inoculation of macerated organs into mice. The sensitivity of axenic cultivation has been estimated to fall between 10 and 100 GE/mL (genome equivalents), depending on the quality of the sample (authors' experience). The impact of repeated axenic propagation on virulence remains to be fully elucidated (Kersh et al. 2011, Kuley et al. 2015).

Coxiella-like Organisms

Initially, the Coxiella genus was thought to comprise solely C. burnetii species, but is now recognized to contain other members, namely Coxiella cheraxi and novel Coxiella-like organism identified in birds and in nonvertebrate species. C. cheraxi was first isolated in 2000 from connective and hepatopancreatic tissues of a dead crayfish, displaying inclusion bodies with Rickettsia-like gram-negative bacteria (Tan and Owens 2000). The partial 16S rDNA, sodB, and com1 sequences of C. cheraxi (strain TO-98) shared highest homology with C. burnetii sequences, achieving similarity of 96%, 96%, and 100%, respectively (Tan and Owens 2000, Cooper et al. 2007). Birds are commonly infected with C. burnetii without apparent clinical signs, but in contrast, show pathology when infected with Candidatus Coxiella avium, a pleomorphic Coxiella-like organism multiplying in macrophage vacuoles and leading to inflammation of liver, lung, and spleen or systemic infection and death of the host (Shivaprasad et al. 2008, Vapniarsky et al. 2012). Further diversity among the genus has been described with reports of Coxiella-like organisms as endosymbionts among several species of ticks (Duron et al. 2015), with extremely high (close to 100%) infection frequency. Indeed, it has been postulated that these might represent ancestral species of C. burnetii (Duron et al. 2015). The genetic classification of these organisms within the Coxiella genus is complex, with common patterns of codivergence within tick species (tick species-specific clades) and horizontal gene transfer events complicating the phylogenetic separation (Duron et al. 2015). The genome is further reduced in comparison with that of C. burnetii (Smith et al. 2015) and traditional cultivation methods for C. burnetii have been unsuccessful to date (Duron et al. 2015). Importantly, several IS1111 sequence haplotypes are present in Coxiella-like tick endosymbionts (Duron 2015), consequently caution is needed to avoid misidentification between Coxiella-like bacteria and C. burnetii, as previously mentioned in the above Real-time PCR section. Table 2 summarizes PCR assays used to screen samples for Coxiella-like bacteria.

Table 2.

PCR ^* Assays (PubMed) Used for Detection of Coxiella-Like Bacteria

		Fragment amplified (bp)
Target gene	Method	<100	100–500	>500	Reference
sodB	Conventional PCR + sequencing		✓		Stein and Raoult (1992),Papa et al. (2016)
				✓	Tan and Owens (2000)
16S rRNA	Real-time PCR + sequencing	✓			Liu et al. (2013)
	Conventional PCR + sequencing		✓		Almeida et al. (2012)
	Conventional PCR + sequencing			✓	Bernasconi et al. (2002),Mediannikov et al. (2003),Duron et al. (2014),Wilkinson et al. (2014),Duron et al. (2015),Al-Deeb et al. (2016),Machado-Ferreira et al. (2016)
23S rRNA	Conventional PCR + sequencing			✓	Noda et al. (1997),Duron et al. (2015)
groEL	Conventional PCR + sequencing			✓	Duron et al. (2014),Duron et al. (2015)
icd	LAMP PCR + sequencing			✓	Al-Deeb et al. (2016)
com1	Conventional PCR + sequencing		✓		Raele et al. (2015)
	Conventional PCR + sequencing			✓	Tan and Owens (2000)
fusA	Conventional PCR + sequencing		✓		Zhong (2012)
rpoB	Conventional PCR + sequencing			✓	Duron et al. (2015)
dnaK	Conventional PCR + sequencing			✓	Duron et al. (2015)

Amplified fragments were either directly sequenced or cloned into plasmid vectors before sequencing.

Conclusion

Direct detection of C. burnetii, although challenging, fulfills a much needed diagnostic gap. Recovery of isolates is essential to address our evolving understanding of this pathogen and to decipher our understanding of the intricate interactions between this microbe and its vertebrate host. This will pave the way for better-targeted intervention and control strategies. Furthermore, direct detection is essential to provide categorical association of emerging clinical sequelae with C. burnetii infection. Finally, the discriminatory methods reviewed above furnish us with tools to detect hitherto undescribed species, expanding our understanding of the Coxiella genus and highlighting potential limitations of our current diagnostic tools.

Footnotes

Acknowledgment

This work was done under the frame of COST action TD1303.

Disclosure Statement

No competing financial interests exist.

References

Agerholm

. Coxiella burnetii associated reproductive disorders in domestic animals-a critical review. Acta Vet Scand, 2013; 55:13.

Al-Deeb

, Frangoulidis

, Walter

, Kömpf

, et al. Coxiella-like endosymbiont in argasid ticks (Ornithodoros muesebecki) from a Socotra Cormorant colony in Umm Al Quwain, United Arab Emirates. Ticks Tick Borne Dis, 2016; 7:166–171.

Almeida

, Marcili

, Leite

, Nieri-Bastos

, et al. Coxiella symbiont in the tick Ornithodoros rostratus (Acari: Argasidae). Ticks Tick Borne Dis, 2012; 3:203–206.

Alsaleh

, Pellerin

, Rodolakis

, Larrat

, et al. Detection of Coxiella burnetii, the agent of Q fever, in oviducts and uterine flushing media and in genital tract tissues of the non pregnant goat. Comp Immunol Microbiol Infect Dis, 2011; 34:355–360.

Amitai

, Bromberg

, Bernstein

, Raveh

, et al. A large Q fever outbreak in an urban school in central Israel. Clin Infect Dis, 2010; 50:1433–1438.

Anderson

, Bijlmer

, Fournier

, Graves

, et al. Diagnosis and management of Q fever-United States, 2013: Recommendations from CDC and the Q Fever Working Group. MMWR Recomm Rep, 2013; 62:1–30.

Barkallah

, Gharbi

, Hassena

, Slima

, et al. Survey of infectious etiologies of bovine abortion during mid- to late gestation in dairy herds. PLoS One, 2014; 9:e91549.

Beare

, Unsworth

, Andoh

, Voth

, et al. Comparative genomics reveal extensive transposon-mediated genomic plasticity and diversity among potential effector proteins within the genus Coxiella . Infect Immun, 2009; 77:642–656.

Bernasconi

, Casati

, Péter

, Piffaretti

. Rhipicephalus ticks infected with Rickettsia and Coxiella in Southern Switzerland (Canton Ticino). Infect Genet Evol, 2002; 2:111–120.

10.

Bittar

, Keita

, Lagier

, Peeters

, et al. Gorilla gorilla gorilla gut: A potential reservoir of pathogenic bacteria as revealed using culturomics and molecular tools. Sci Rep, 2014; 24:7174.

11.

Boarbi

, Fretin

, Mori

. Coxiella burnetii, agent de la fièvre Q. Can J Microbiol, 2015:1–21.

12.

Boarbi

, Mori

, Rousset

, Sidi-Boumedine

, et al. Prevalence and molecular typing of Coxiella burnetii in bulk tank milk in Belgian dairy goats, 2009–2013. Vet Microbiol, 2014; 170:117–124.

13.

Boden

, Wolf

, Hermann

, Frangoulidis

. First isolation of Coxiella burnetii from clinical material by cell-free medium (ACCM2). Eur J Clin Microbiol Infect Dis, 2015; 34:1017–1022.

14.

Bond

, Vincent

, Wilks

, Franklin

, et al. One Health approach to controlling a Q fever outbreak on an Australian goat farm. Epidemiol Infect, 2016; 144:1129–1141.

15.

Brouqui

, Rolain

, Foucault

, Raoult

. Short report: Q fever and Plasmodium falciparum malaria co-infection in a patient returning from the Comoros archipelago. Am J Trop Med Hyg, 2005; 73:1028–1030.

16.

Coleman

, Fischer

, Howe

, Mead

, et al. Temporal analysis of Coxiella burnetii morphological differentiation. J Bacteriol, 2004; 186:7344–7352.

17.

Cooper

, Layton

, Owens

, Ketheesan

, et al. Evidence for the classification of a crayfish pathogen as a member of the genus Coxiella . Lett Appl Microbiol, 2007; 45:558–563.

18.

Costa

, Morais

, Santos

, Tavares

, et al. Q fever chronic osteomyelitis in two children. Pediatr Infect Dis J, 2015; 34:1269–1271.

19.

Cumbassa

, Barahona

, Cunha

, Azorin

, et al. Coxiella burnetii DNA detected in domestic ruminants and wildlife from Portugal. Vet Microbiol, 2015; 180:136–141.

20.

Curet

, Paust

. Transmission of Q fever from experimental sheep to laboratory personnel. Am J Obstet Gynecol, 1972; 114:566–568.

21.

Cutler

, Bouzid

, Cutler

. Q fever. J Infect, 2007; 54:313–318.

22.

D'Amato

, Eldin

, Georgiades

, Edouard

, et al. Loss of TSS1 in hypervirulent Coxiella burnetii 175, the causative agent of Q fever in French Guiana. Comp Immunol Microbiol Infect Dis, 2015; 41:35–41.

23.

D'Amato

, Rouli

, Edouard

, Tyczka

, et al. The genome of Coxiella burnetii Z3055, a clone linked to the Netherlands Q fever outbreaks, provides evidence for the role of drift in the emergence of epidemic clones. Comp Immunol Microbiol Infect Dis, 2014; 37:281–288.

24.

De Bruin

, de Groot

, de Heer

, Bok

, et al. Detection of Coxiella burnetii in complex matrices by using multiplex quantitative PCR during a major Q fever outbreak in the Netherlands. Appl Environ Microb, 2011; 77:6516–6523.

25.

De Bruin

, Janse

, Koning

, de Heer

, et al. Detection of Coxiella burnetii DNA in the environment during and after a large Q fever epidemic in the Netherlands. J Appl Microbiol, 2013; 114:1395–1404.

26.

Di Domenico

, Curini

, De Massis

, Di Provvido

, et al. Coxiella burnetii in Central Italy: Novel genotypes are circulating in cattle and goats. Vector Borne Zoonotic Dis, 2014; 14:710–715.

27.

Duron

. The IS1111 insertion sequence used for detection of Coxiella burnetii is widespread in Coxiella-like endosymbionts of ticks. FEMS Microbiol Lett, 2015; 362:fnv132.

28.

Duron

, Jourdain

, McCoy

. Diversity and global distribution of the Coxiella intracellular bacterium in seabird ticks. Ticks Tick Borne Dis, 2014; 5:557–563.

29.

Duron

, Noel

, McCoy

, Bonazzi

, et al. The recent evolution of a maternally-inherited endosymbiont of ticks led to the emergence of the Q Fever pathogen, Coxiella burnetii . PLoS Pathog, 2015; 11:e1004892.

30.

Elsa

, Duron

, Severine

, Gonzalez-Acuna

, et al. Molecular methods routinely used to detect Coxiella burnetii in ticks cross-react with Coxiella-like bacteria. Infect Ecol Epidemiol, 2015; 5:29230.

31.

El-Mahallawy

, Kelly

, Zhang

, Yang

, et al. Serological and molecular evidence of Coxiella burnetii in samples from humans and animals in China. Ann Agric Environ Med, 2016; 23:87–91.

32.

Fournier

, Thuny

, Richet

, Lepidi

, et al. Comprehensive diagnostic strategy for blood culture-negative endocarditis: A prospective study of 819 new cases. Clin Infect Dis, 2010; 51:131–140.

33.

Georgiev

, Afonso

, Neubauer

, Needham

, et al. Q fever in humans and farm animals in four European countries, 1982 to 2010. Euro Surveill, 2013; 18:pii: 20407.

34.

Gilroy

, Formica

, Beers

, Egan

, et al. Abattoir-associated Q fever: A Q fever outbreak during a Q fever vaccination program. Aust N Z J Public Health, 2001; 25:362–367.

35.

Gimenez

. Staining rickettsiae in yolk-sac cultures. Stain Technol, 1964; 39:135–140.

36.

Glazunova

, Roux

, Freylikman

, Sekeyova

, et al. Coxiella burnetii genotyping. Emerg Infect Dis, 2005; 11:1211–1217.

37.

González-Barrio

, Maio

, Vieira-Pinto

, Ruiz-Fons

. European rabbits as reservoir for Coxiella burnetii . Emerg Infect Dis, 2015a; 21:1055–1058.

38.

González-Barrio

, Martín-Hernando

, Ruiz-Fons

. Shedding patterns of endemic Eurasian wild boar (Sus scrofa) pathogens. Res Vet Sci, 2015b; 102:206–211.

39.

Gouriet

, Fenollar

, Patrice

, Drancourt

, et al. Use of shell-vial cell culture assay for isolation of bacteria from clinical specimens: 13 years of experience. J Clin Microbiol, 2005; 43:4993–5002.

40.

Graham

, Yamauchi

, Rountree

. Q fever in animal laboratory workers: An outbreak and its investigation. Am J Infect Control, 1989; 17:345–348.

41.

Hackstadt

, Williams

. Biochemical stratagem for obligate parasitism of eukaryotic cells by Coxiella burnetii . Proc Natl Acad Sci U S A, 1981; 78:3240–3244.

42.

Hall

, Richmond

, Caul

, Pearce

, et al. Laboratory outbreak of Q fever acquired from sheep. Lancet, 1982; 1:1004–1006.

43.

Hammerl

, Mertens

, Sprague

, Hackert

, et al. First draft genome sequence of a human Coxiella burnetii isolate, originating from the largest Q fever outbreak ever reported, the Netherlands, 2007 to 2010. Genome Announc, 2015; 3: pii: e00445–15.

44.

Harris

, Storm

, Lloyd

, Arens

, et al. Long-term persistence of Coxiella burnetii in the host after primary Q fever. Epidemiol Infect, 2000; 124:543–549.

45.

Hawker

, Ayres

, Blair

, Evans

, et al. A large outbreak of Q fever in the West Midlands: Windborne spread into a metropolitan area?. Commun Dis Public Health, 1998; 1:180–187.

46.

Hermans

, Huijsmans

, Schellekens

, Savelkoul

, et al. Coxiella burnetii DNA in goat milk after vaccination with Coxevac(®). Vaccine, 2011; 29:2653–2656.

47.

Hornstra

, Priestley

, Georgia

, Kachur

, et al. Rapid typing of Coxiella burnetii . PLoS One, 2011; 6:e26201.

48.

Huijsmans

, Schellekens

, Wever

, Toman

, et al. Single-nucleotide-polymorphism genotyping of Coxiella burnetii during Q fever outbreak in The Netherlands. Appl Environ Microbiol, 2011; 77:2051–2057.

49.

Jaton

, Peter

, Raoult

, Tissot

, et al. Development of a high throughput PCR to detect Coxiella burnetii and its application in a diagnostic laboratory over a 7-year period. New Microbes New Infect, 2013; 1:6–12.

50.

Johnson

, Kadull

. Laboratory-acquired Q fever. A report of fifty cases. Am J Med, 1966; 41:391–403.

51.

Karlsson

, Macellaro

, Bystrom

, Forsman

, et al. Eight new genomes and synthetic controls increase the accessibility of rapid melt-MAMA SNP typing of Coxiella burnetii . PLoS One, 2014; 9:e85417.

52.

Kersh

, Lambourn

, Self

, Akmajian

, et al. Coxiella burnetii infection of a Steller sea lion (Eumetopias jubatus) found in Washington State. J Clin Microbiol, 2010a; 48:3428–3431.

53.

Kersh

, Oliver

, Self

, Fitzpatrick

, et al. Virulence of pathogenic Coxiella burnetii strains after growth in the absence of host cells. Vector Borne Zoonotic Dis, 2011; 11:1433–1438.

54.

Kersh

, Wolfe

, Fitzpatrick

, Candee

, et al. Presence of Coxiella burnetii DNA in the environment of the United States, 2006 to 2008. Appl Environ Microbiol, 2010b; 76:4469–4475.

55.

Kim

, Kim

, Lafferty

, Dubovi

. Coxiella burnetii in bulk tank milk samples, United States. Emerg Infect Dis, 2005; 11:619–621.

56.

Klaassen

, Nabuurs-Franssen

, Tilburg

, Hamans

, et al. Multigenotype Q fever outbreak, the Netherlands. Emerg Infect Dis, 2009; 15:613–614.

57.

Klee

, Tyczka

, Ellerbrok

, Franz

, et al. Highly sensitive real-time PCR for specific detection and quantification of Coxiella burnetii . BMC Microbiol, 2006; 6:2.

58.

Kruszewska

, Tylewska-Wierzbanowska

. Isolation of Coxiella burnetii from bull semen. Res Vet Sci, 1997; 62:299–300.

59.

Kuley

, Smith

, Frangoulidis

, Smits

, et al. Cell-free propagation of Coxiella burnetii does not affect its relative virulence. PLoS One, 2015; 10:e0121661.

60.

Lagier

, Edouard

, Pagnier

, Mediannikov

, et al. Current and past strategies for bacterial culture in clinical microbiology. Clin Microbiol Rev, 2015; 28:208–236.

61.

Liu

, Li

, Liu

, Hu

, et al. Coinfection of Dermacentor silvarum olenev (acari: ixodidae) by Coxiella-Like, Arsenophonus-like, and Rickettsia-like symbionts. Appl Environ Microbiol, 2013; 79:2450–2454.

62.

Lockhart

, Graves

, Banazis

, Fenwick

, et al. A comparison of methods for extracting DNA from Coxiella burnetii as measured by a duplex qPCR. Lett Appl Microbiol, 2011; 52:514–520.

63.

Lockhart

, Islam

, Fenwick

, Graves

, et al. Comparative sensitivity of four different cell lines for the isolation of Coxiella burnetii . FEMS Microbiol Lett, 2012; 334:75–78.

64.

Loftis

, Reeves

, Szumlas

, Abbassy

, et al. Rickettsial agents in Egyptian ticks collected from domestic animals. Exp Appl Acarol, 2006; 40:67–81.

65.

Machado-Ferreira

, Vizzoni

, Balsemão-Pires

, Moerbeck

, et al. Coxiella symbionts are widespread into hard ticks. Parasitol Res, 2016; 115:4691–4699.

66.

Marmion

, Storm

, Ayres

, Semendric

, et al. Long-term persistence of Coxiella burnetii after acute primary Q fever. QJM, 2005; 98:7–20.

67.

Massung

, Cutler

, Frangoulidis

. Molecular Typing of Coxiella burnetii (Q Fever). Adv Exp Med Biol, 2012; 984:381–396.

68.

Maurin

, Raoult

. Q fever. Clin Microbiol Rev, 1999; 12:518–553.

69.

Mediannikov

, Fenollar

, Socolovschi

, Diatta

, et al. Coxiella burnetii in humans and ticks in rural Senegal. PLoS Negl Trop Dis, 2010; 4:e654.

70.

Mediannikov

, Ivanov

, Nishikawa

, Saito

, et al. Molecular evidence of Coxiella-like microorganism harbored by Haemaphysalis concinnae ticks in the Russian Far East. Ann NY Acad Sci, 2003; 990:226–228.

71.

Miceli

, Veryser

, Anderson

, Hofinger

, et al. A case of person-to-person transmission of Q fever from an active duty serviceman to his spouse. Vector Borne Zoonotic Dis, 2010; 10:539–541.

72.

Milazzo

, Hall

, Storm

, Harris

, et al. Sexually transmitted Q fever. Clin Infect Dis, 2001; 33:399–402.

73.

Millán

, Proboste

, Fernández de Mera

, Chirife

, et al. Molecular detection of vector-borne pathogens in wild and domestic carnivores and their ticks at the human-wildlife interface. Ticks Tick Borne Dis, 2016; 7:284–290.

74.

Million

, Raoult

. Recent advances in the study of Q fever epidemiology, diagnosis and management. J Infect, 2015; 71 Suppl 1:S2–S9.

75.

Mori

, Boarbi

, Michel

, Bakinahe

, et al. In vitro and in vivo infectious potential of Coxiella burnetii: A study on Belgian livestock isolates. PLoS One, 2013; 8:e67622.

76.

Niemczuk

, Szymańska-Czerwińska

, Śmietanka

, Bocian

. Comparison of diagnostic potential of serological, molecular and cell culture methods for detection of Q fever in ruminants. Vet Microbiol, 2014; 171:147–152.

77.

Niemczuk

, Szymanska-Czerwinska

, Zarzecka

, Konarska

. Q fever in a cattle herd and humans in the southeastern Poland. Laboratory diagnosis of the diseases using serological and molecular methods. Bull Vet Inst Pu1awy, 2011; 55:593–598.

78.

Noda

, Munderloh

, Kurtti

. Endosymbionts of ticks and their relationship to Wolbachia spp. and tick-borne pathogens of humans and animals. Appl Environ Microbiol, 1997; 63:3926–3932.

79.

Omsland

, Beare

, Hill

, Cockrell

, et al. Isolation from animal tissue and genetic transformation of Coxiella burnetii are facilitated by an improved axenic growth medium. Appl Environ Microbiol, 2011; 77:3720–3725.

80.

Omsland

, Cockrell

, Fischer

, Heinzen

. Sustained axenic metabolic activity by the obligate intracellular bacterium Coxiella burnetii . J Bacteriol, 2008; 190:3203–3212.

81.

Omsland

, Cockrell

, Howe

, Fischer

, et al. Host cell-free growth of the Q fever bacterium Coxiella burnetii . Proc Natl Acad Sci U S A, 2009; 106:4430–4434.

82.

Omsland

, Heinzen

. Life on the outside: The rescue of Coxiella burnetii from its host cell. Annu Rev Microbiol, 2011; 65:111–128.

83.

Panning

, Kliwiński

, Greiner-Fischer

, Peters

, et al. High throughput detection of Coxiella burnetii by real-time PCR with internal control system and automated DNA preparation. BMC Microbiol, 2008; 8:77.

84.

Papa

, Tsioka

, Kontana

, Papadopoulos

, et al. Bacterial pathogens and endosymbionts in ticks. Ticks Tick Borne Dis, 2016; pii: S1877-959X(16)30149–2.

85.

Raele

, Galante

, Pugliese

, De Simone

, et al. Coxiella-like endosymbiont associated to the “Anatolian brown tick” Rhipicephalus bursa in Southern Italy. Microbes Infect, 2015; 17:799–805.

86.

Roest

, Tilburg

, van der Hoek

, Vellema

, et al. The Q fever epidemic in The Netherlands: History, onset, response and reflection. Epidemiol Infect, 2011; 139:1–12.

87.

Sandoz

, Beare

, Cockrell

, Heinzen

. Complementation of arginine auxotrophy for genetic transformation of Coxiella burnetii by use of a defined axenic medium. Appl Environ Microbiol, 2016; 82:3042–3051.

88.

Santos

, Tilburg

, Botelho

, Barahona

, et al. Genotypic diversity of clinical Coxiella burnetii isolates from Portugal based on MST and MLVA typing. Int J Med Microbiol, 2012; 302:253–256.

89.

Schets

, de Heer

, de Roda Husman

. Coxiella burnetii in sewage water at sewage water treatment plants in a Q fever epidemic area. Int J Hyg Environ Health, 2013; 216:698–702.

90.

Schneeberger

, Hermans

, van Hannen

, Schellekens

, et al. Real-time PCR on serum samples is indispensable for early diagnosis of acute Q fever. Clin Vaccine Immunol, 2010; 17:286–290.

91.

Seshadri

, Paulsen

, Eisen

, Read

, et al. Complete genome sequence of the Q-fever pathogen Coxiella burnetii . Proc Natl Acad Sci U S A, 2003; 100:5455–5460.

92.

Shivaprasad

, Cadenas

, Diab

, Nordhausen

, et al. Coxiella-like infection in psittacines and a toucan. Avian Dis, 2008; 52:426–432.

93.

Sidi-Boumedine

, Ellis

, Adam

, Prigent

, et al. Draft genome sequences of six ruminant Coxiella burnetii isolates of european origin. Genome Announc, 2014; 2:e00285–14.

94.

Smith

. Q fever outbreak in Birmingham, UK. Lancet, 1989; 2:557.

95.

Smith

, Driscoll

, Gillespie

, Raghavan

. A Coxiella-like endosymbiont is a potential vitamin source for the Lone Star tick. Genome Biol Evol, 2015; 7:831–838.

96.

Stein

, Raoult

. Detection of Coxiella burnetti by DNA amplification using polymerase chain reaction. J Clin Microbiol, 1992; 30:2462–2466.

97.

Stemmler

, Meyer

. Rapid and specific detection of Coxiella burnetii by LightCycler PCR. In: Reischl U, Wittwer C, Cockerill F, eds. Rapid Cycle realtime PCR: methods and applications. Microbiology And Food Analysis . Berlin:Springer Berlin Heidelberg;, 2002: 149–154.

98.

Tan

, Owens

. Infectivity, transmission and 16S rRNA sequencing of a rickettsia, Coxiella cheraxi sp. nov., from the freshwater crayfish Cherax quadricarinatus . Dis Aquat Organ, 2000; 41:115–122.

99.

Tilburg

, Melchers

, Pettersson

, Rossen

, et al. Interlaboratory evaluation of different extraction and real-time PCR methods for detection of Coxiella burnetii DNA in serum. J Clin Microbiol, 2010; 48:3923–3927.

100.

Tilburg

, Roest

, Buffet

, Nabuurs-Franssen

, et al. Epidemic genotype of Coxiella burnetii among goats, sheep and humans in The Netherlands. Emerg Infect Dis, 2012b; 18:887–889.

101.

Tilburg

, Rossen

, van Hannen

, Melchers

, et al. Genotypic diversity of Coxiella burnetii in the 2007–2010 Q fever outbreak episodes in The Netherlands. J Clin Microbiol, 2012a; 50:1076–1078.

102.

Tozer

, Lambert

, Strong

, Field

, et al. Potential animal and environmental sources of Q fever infection for humans in Queensland. Zoonoses Public Health, 2014; 61:105–112.

103.

van Schaik

, Chen

, Mertens

, Weber

, et al. Molecular pathogenesis of the obligate intracellular bacterium Coxiella burnetii . Nat Rev Microbiol, 2013; 11:561–573.

104.

Vapniarsky

, Barr

, Murphy

. Systemic Coxiella-like infection with myocarditis and hepatitis in an eclectus parrot (Eclectus roratus). Vet Pathol, 2012; 49:717–722.

105.

Walter

, Vincent

, Stenos

, Graves

, et al. Genome Sequence of Coxiella burnetii Strain AuQ01 (Arandale) from an Australian Patient with Acute Q Fever. Genome Announc, 2014; 2:pii: e00964–14.

106.

Wilkinson

, Dietrich

, Lebarbenchon

, Jaeger

, et al. Massive infection of seabird ticks with bacterial species related to Coxiella burnetii. Appl Environ Microbiol, 2014; 80:3327–3333.

107.

Wurtz

, Papa

, Hukic

, Di Caro

, et al. Survey of laboratory-acquired infections around the world in biosafety level 3 and 4 laboratories. Eur J Clin Microbiol Infect Dis, 2016; 35:1247–1258.

108.

Zhong

. Coxiella-like endosymbionts. Adv Exp Med Biol, 2012; 984:365–379.