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Prevalence and Infection Load Dynamics of Rickettsia felis in Actively Feeding Cat Fleas
research-article
Author(s):
Kathryn E. Reif
1 ,
Rhett W. Stout
1 ,
Gretchen C. Henry
1 ,
Lane D. Foil
2 ,
Kevin R. Macaluso
1
,
*
Publication date (Electronic):
30 July 2008
Journal:
PLoS ONE
Publisher:
Public Library of Science
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Abstract
Background
Rickettsia felis is a flea-associated rickettsial pathogen recurrently identified in both colonized and wild-caught cat fleas, Ctenocephalides felis. We hypothesized that within colonized fleas, the intimate relationship between R. felis and C. felis allows for the coordination of rickettsial replication and metabolically active periods during flea bloodmeal acquisition and oogenesis.
Methodology/Principal Findings
A quantitative real-time PCR assay was developed to quantify R. felis in actively feeding R. felis-infected fleas. In three separate trials, fleas were allowed to feed on cats, and a mean of 3.9×10 6 R. felis 17-kDa gene copies was detected for each flea. A distinct R. felis infection pattern was not observed in fleas during nine consecutive days of bloodfeeding. However, an inverse correlation between the prevalence of R. felis-infection, which ranged from 96% in Trial 1 to 35% in Trial 3, and the R. felis-infection load in individual fleas was identified. Expression of R. felis-infection load as a ratio of R. felis/ C. felis genes confirmed that fleas in Trial 3 had significantly greater rickettsial loads than those in Trial 1.
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Most cited references33
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- Abstract: found
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The Genome Sequence of Rickettsia felis Identifies the First Putative Conjugative Plasmid in an Obligate Intracellular Parasite
Hiroyuki Ogata, Patricia Renesto, Stéphane Audic … (2005)
Introduction Rickettsiae are obligate intracellular small gram-negative bacteria associated with different arthropod hosts. Many Rickettsia species infect human beings and are responsible for mild to severe diseases. Rickettsia felis, the agent of the flea-borne spotted fever rickettsiosis, exhibits several specificities among the currently recognized Rickettsia species. After being identified in fleas in 1990 [1], R. felis has been found worldwide in flea species such as Ctenocephalides felis, parasitizing cats and dogs, and Pulex irritans. R. felis is transovarially transmitted in these insects [2]. Several cases of human infection caused by R. felis have been reported [3,4]. Rickettsia species are phylogenetically classified into two groups: the typhus group and the spotted-fever group (SFG). R. felis belongs to the SFG, together with tick-associated Rickettsia species such as R. conorii, R. sibirica, and R. rickettsii. However its lifestyle resembles that of R. typhi (typhus group), which is also hosted and transovarially transmitted by fleas. Furthermore, R. felis is known to coinfect fleas with Bartonella henselae, B. quintana, and Wolbachia pipientis [5]. The culture conditions of R. felis were established in 2001 using Xenopus laevis tissue culture (XTC) cells at relatively low temperatures (optimally at 28 °C) [3]. Besides these features, little is known about this pathogen. To date, six Rickettsia genome sequences are available. These are from two typhus group species (R. prowazekii [6] and R. typhi [7]) and four SFG species (R. conorii [8], R. sibirica [9], R. rickettsii, and R. akari). To further identify the specificities of R. felis, we determined its genome sequence. Results General Genome Features The genome of R. felis comprises three replicons: a 1,485,148 bp circular chromosome and two circular plasmids identified for the first time in the genus Rickettsia (Figure 1). The predicted total complement of 1,512 protein-coding genes (open reading frames [ORFs]) is the largest among currently sequenced Rickettsia genomes (Table 1). Of these, 1,402 (92.7%) exhibited homologs in the nonredundant database and 1,080 (71.4%) were assigned putative functions. The R. felis chromosome exhibits a long-range (24−277 kbp) colinearity relative to other Rickettsia genomes, although it is more frequently interrupted by inversions/translocations than is observed between other Rickettsia genomes (Figure 2A). This colinearity allowed the precise assessment of orthologous relationships between ORFs of five Rickettsia species (R. felis, R. conorii, R. sibirica, R. prowazekii, and R. typhi). On this basis, we identified 530 R. felis–specific ORFs, that were either absent or degraded (split or fragmented) in the other four Rickettsia genomes (Tables 2 and 3). Consistently, the R. felis genome exhibited a much higher number of gene families than other Rickettsia species (see Table 1). The R. felis–specific ORFs included a remarkably high number of paralogs for transposases, surface cell antigens (sca), global metabolism regulators (spoT), and proteins containing protein–protein interaction motifs such as ankyrin repeats and tetratricopeptide repeats (TPRs). Furthermore, we identified many other ORFs putatively associated with the adaptations of R. felis to its host environment or with its pathogenesis. Plasmids The two R. felis plasmids, named pRF and pRFδ, are 62,829 bp and 39,263 bp long, respectively. Their topologies and sizes were confirmed experimentally (Figures S1 and S2). The pRF plasmid contains 68 ORFs, of which 53 (77.9%) exhibited homologs in public databases and 44 (64.7%) were associated with functional attributes. The nucleotide sequences of pRFδ and pRF are identical, except for an additional 23,566-bp segment that contains 24 ORFs (pRF15–pRF38) in pRF (see Table 3). These plasmids are likely to be R. felis specific since all attempts to detect specific plasmid sequences by polymerase chain reaction (PCR) from DNA of available reference rickettsial species were unsuccessful. In contrast, the same assays against 30 fleas naturally infected by R. felis resulted in amplification of the plasmid sequences in all cases. Plasmids are referred to as conjugative or nonconjugative. The former are disseminated by conjugation from cell to cell, while the latter are only vertically transmitted. The pRF plasmid encodes several homologs of proteins involved in the different conjugative steps (see Table 3; Figure S3). First, it exhibits a split gene (pRF38/pRF39) homologous to the traA Ti of the Agrobacterium tumefaciens tumor-inducing plasmid [10]. TraATi is thought to be a DNA-processing machinery with nickase and helicase activities to generate the transfer strand from the origin of transfer (oriT) [10]. Second, the pRF encodes another split gene (pRF43/pRF44) homologous to the traD F in the Escherichia coli F plasmid. TraDF is a “coupling protein” that connects the DNA-processing machinery (and transfer strand) to the mating pair formation (Mpf) apparatus, a type IV secretion system (T4SS) [11]. Finally, pRF exhibits an ORF (pRF47) similar to TraGF, a protein involved in the F-pilus assembly and aggregate stabilization [12]. Despite the presence of these ORFs linked to the initiation of plasmid transfer, the pRF sequence lacks clear homologs for the proteins involved in the Mpf apparatus found in other bacteria. Nevertheless, the R. felis chromosome (as well as other Rickettsia genomes) encodes most of the components of T4SS, which are highly similar to the vir genes of A. tumefaciens. Since the R. felis T4SS components (virB2 [RF1075], virB3 [RF0087], virB4 [RF0088], virB6 [RF0089, RF0090, RF0091, RF0092, and RF0093], virB8 [RF0463 and RF0465], virB9 [RF0462 and RF0466], virB10 [RF0467], virB11 [RF0468], and virD4 [RF0469]) are conserved in all sequenced Rickettsia genomes that lack plasmids, the primary suspected role of the R. felis T4SS is to translocate virulence factors to hosts. However, the T4SS of A. tumefaciens (vir) and Legionella pneumophila (dot/icm) have been shown to function both as DNA-transfer machineries and as effector translocators [13]. Thus, the R. felis T4SS may also promote the transfer of DNA as in A. tumefaciens. We also noticed that the R. felis chromosome exhibits a DNA primase gene (RF0786) similar to TraC found in the E. coli IncP plasmid. TraC initiates the replication of transferred DNA strands in the recipient cells. Finally, the R. felis chromosome encodes a protein (RF0020) similar to competence protein ComE3, a protein (RF0964) similar to the F-pilin acetylation protein TraX, and a split gene (RF0705/RF0706) homologous to the P-pilus assembly protein FimD. In conclusion, the presence of those putative conjugative transfer genes suggests that the R. felis plasmids have been acquired by conjugation and that R. felis may still retain the capacity of transferring plasmids. Genome Plasticity We identified 333 repeated DNA sequences (50 to 2,645 bp long) in the R. felis genome, accounting for 4.3% of the sequence, a proportion markedly higher than in other sequenced Rickettsia genomes (see Table 1; Figure 2B). The major source of those repeats is the proliferation of transposase genes, for which we identified 82 copies (or inactivated derivatives). Among other obligate intracellular bacteria, only W. pipientis wMel [14] and Parachlamydia sp. UWE25 [15] exhibit such a high number of large mobile genetic elements. The occurrence of highly similar transposase sequences appears to play a major role in the plasticity of the R. felis genome (see Figure 2A). Transposase ORFs were identified at most extremities of the R. felis genomic segments colinear with the R. conorii genome, suggesting that the R. felis chromosome has been rearranged many times through recombination mediated by these mobile sequences. With the use of the GRAPPA software inferring the most parsimonious genome-rearrangement scenario, we estimated at least 11 inversion events between R. felis and R. conorii. In contrast, only four inversions are required to associate more distantly related R. conorii and R. prowazekii genomes. In addition to transposases, we identified eight phage-related ORFs (see Table 2). The R. felis genome thus appears to have been invaded more frequently by such foreign DNAs than other Rickettsia species. Besides long repeats, Rickettsia genomes are known to contain a number of small palindromic repeats (Rickettsia palindromic elements [RPEs]) capable of invading both coding and noncoding regions [16]. We identified 728 RPEs in the R. felis genome. Of these RPEs, 85 were found within ORFs and three were found in RNA-coding genes. The R. felis chromosome and plasmids share several homologs, suggesting gene exchanges between these replicons. Of 68 ORFs in pRF, 11 have a close homolog (>50% amino acid sequence identity) in the chromosome; these are seven transposases, patatin-like phospholipase (pRF11), thymidylate kinase (pRF13), and two small heat-shock proteins (pRF51 and pRF52). Among these, patatin-like proteins exhibit the most intriguing phylogeny (Figure S4). The genomes of five Rickettsia species (R. prowazekii, R. typhi, R. conorii, R. sibirica, and R. felis) exhibit chromosomal patatin-like phospholipase gene (pat1). Gene organization around pat1 is similar between these Rickettsia. Interestingly, a phylogenetic analysis for these Pat1 and the plasmid-encoded Pat2 indicates a close relationship between Pat1 (RF0360) and Pat2 of R. felis, together being an outgroup of Pat1 sequences of other Rickettsia, suggesting a gene replacement of the chromosomally encoded pat1 by the plasmid-encoded pat2 in the lineage leading to R. felis. Most R. felis genes with orthologs in other Rickettsia have probably been inherited vertically from a common ancestor. On the other hand, genes without orthologs in other Rickettsia may have been acquired by lateral gene transfer. To test this hypothesis, we analyzed the taxonomic distribution of BLASTP best hits of R. felis ORFs against the nonredundant database (excluding rickettsial sequences) (Figure S5). R. felis ORFs with orthologs in other Rickettsia matched preferentially (64%) with sequences from the same taxonomic group as R. felis (i.e., α-proteobacteria). In contrast, the BLAST best hits for the chromosomal ORFs lacking orthologs in other Rickettsia were found preferentially in γ-proteobacteria (31%; 58 ORFs) and cyanobacteria (18%; 33 ORFs). The taxonomic distributions of the best matches for these two ORF sets were significantly different (p < 0.001; χ2 test). This result suggests that many R. felis-specific genes may originate from distantly related organisms by lateral transfer. However, methods based on nucleotide composition bias failed to identify unambiguous candidates for lateral gene acquisition in R. felis. Surface Antigens The sca family is one of the largest paralogous gene families in Rickettsia [8]. Five sca members have been identified in the previously published Rickettsia genomes. Several Sca proteins are known to account for major antigenic differences between Rickettsia species [17] and may play important roles in adhesion to host cells [18]. Sca proteins are characterized by highly variable N-terminal sequences and a conserved C-terminal autotransporter β-domain, which translocates the N-terminal part outside the outer membrane. The R. felis genome exhibits the highest number of sca genes among currently available Rickettsia genomes. We identified nine intact sca paralogs (sca1, sca2, sca3, sca4, sca5/ompB, sca8, sca9, sca12, and sca13) as well as four fragmented or split paralogs (sca0/ompA, sca7, sca10, and sca11). Reverse transcriptase–polymerase chain reaction (RT-PCR) experiments demonstrated that, under mild log growth phase, all R. felis sca paralogs were transcribed, including split ones. Phylogenetic analyses suggest that ancient duplication events gave rise to these paralogs before the divergence of Rickettsia species. We noticed that sca genes exhibit highly different patterns of presence/absence across different Rickettsia species (Table S1). Only ompB and sca4 are conserved in all available Rickettsia genomes [19], remaining members being degraded or absent in one or more species. Together with the accelerated amino acid changes, differential gene degradation of sca paralogs probably contributes to the intra-species variation of those cell-surface proteins and might be linked with their adaptation to different host environments. R. felis is genetically and serologically classified into the SFG of Rickettsia [20]. However, cross-reactivities caused by both proteins and lipopolysaccharides have been found with R. typhi using mouse sera [2] and human sera (Figure S6). R. conorii rarely cross-reacts with R. typhi. We therefore suspected that genes found in both R. felis and R. typhi, but missing in R. conorii, might be responsible for the cross-reactivities of R. felis and R. typhi. A list of such genes includes a sca family gene (sca3), encoding a protein with a predicted molecular weight of 319 kDa, and rfaJ for the lipopolysaccharide 1,2-glucosyltransferase (Table 4). Adaptation to Environment Transcriptional regulation may be of critical importance in R. felis, as the numbers of spoT, the gene regulating “alarmone,” and chromosomal toxin–antitoxin modules are higher in the R. felis genome than in any other sequenced bacterial genome. SpoT and RelA are two hallmark enzymes regulating global cellular metabolism of E. coli in response to starvation [21]. These enzymes control the concentration of alarmone, (p)ppGpp (guanosine tetra- and pentaphosphates), which in turn acts as an effector of transcription. Remarkably, R. felis exhibits 14 spoT, (spoT1–13 and 15) paralogs (Figure S7). Using RT-PCR, we examined the transcription status of 14 R. felis spoT genes. All the spoT ORFs were transcribed. We classified these ORFs into two groups, based on their alignment against the sequence of the Streptococcus dysgalactiae Rel seq that possesses both (p)ppGpp hydrolase and synthetase activities [22]. The first group (SpoT1–10, 14, and 15) was aligned with the sequence of the hydrolase domain, and the second group (SpoT11, 12, and 13) with the sequence of the synthetase domain. Being consistent with the previous observation [23], our phylogenetic analyses suggest that each paralogous gene group originated in early duplication events before the divergence of Rickettsia species. Notably, every sequenced Rickettsia genome encodes at least one ORF exhibiting hydrolase catalytic residues and one ORF exhibiting synthetase catalytic residues, suggesting that both hydrolase and synthetase functions are required for Rickettsia. We also found that seven spoT (spoT1–4 and 7–9) genes were located in the R. felis chromosome next to a gene encoding a transporter of the major facilitator superfamily (MFS) including proline/betaine transporters. MFS is also a large paralogous gene family composed of at least 23 ORF members in R. felis. Toxin–antitoxin systems are composed of tightly linked toxin and antitoxin gene pairs and ensure stable plasmid inheritance when they are encoded in plasmids. In these systems, the toxic effect of a long-lived toxin is continuously inhibited by a short-lived antitoxin only when whole systems are maintained. The toxin–antitoxin modules have also been found on the chromosomes of many free-living prokaryotes, but have rarely been found in obligate intracellular bacteria [24,25]. In the R. felis chromosome, we identified 16 toxin genes (RF0016, RF0095, RF0271, RF0456, RF0490, RF0602, RF0701, RF0732, RF0787, RF0792, RF0898, RF0911, RF0956, RF1272, RF1286, and RF1368) and 14 antitoxin genes (RF0015, RF0094, RF0272, RF0457, RF0489, RF0601, RF0702, RF0731, RF0779, RF0788, RF0899, RF0910, RF0957, and RF1369), comprising at least 13 modules in operon structures. It is suggested that toxin–antitoxin systems, when encoded on the bacterial chromosome, might be involved in selective killing (a primitive form of bacterial apoptosis) or reversible stasis of bacterial subpopulations during periods of starvation or other stress [26,27]. It is also tempting to speculate that the toxin–antitoxin system could be targeted to the eukaryotic host cells. In this case, this system may help to maintain the presence of bacteria in the host. Notably, in the chromosomally encoded mazEF system of E. coli, the toxin action is regulated by (p)ppGpp. The large number of toxin–antitoxin modules in R. felis, as well as a number of spoT paralogs, might thus be linked to the synchronization of its multiplication within eukaryotic hosts. It is probable that five R. felis–specific ORFs are related to its capacity of antibiotic resistance. We identified a streptomycin resistance protein homolog (RF0774), a class C β-lactamase, AmpC (RF1367), a class D β-lactamase (RF1275), a penicillin acylase homolog with conserved catalytic residues (RF1137), and an ABC-type multidrug transport-system protein, MdlB (RF0981). AmpC β-lactamase is known to be induced by AmpG of the MFS, which was also identified in the R. felis genome (RF0265, RF0608, RF0834, and RF1247). In vivo β-lactamase activity of R. felis was measured using high-performance liquid chromatography (see below). Adaptation to Eukaryotic Hosts R. felis may have developed a specific mechanism to cross-talk with its eukaryotic hosts. It exhibits 22 ankyrin-repeat-containing proteins and 11 TPR-containing proteins. These two protein motifs are frequently found in eukaryotic proteins, but their distributions are rather limited in viruses and bacteria, in both of which they appear to be linked with pathogenicity. The ankyrin repeat is a protein–protein interaction motif, involved in transcription initiation, cell cycle regulation, cytoskeletal integrity, and cell-to-cell signaling [28]. Anaplasma phagocytophilum, a closely related intracellular α-proteobacterium, exhibits a protein containing ankyrin repeats (AnkA), which was detected in the cytoplasm and the nucleus of infected eukaryotic cells (human leukemia-60) [29]. According to the Superfamily database [30], only 15 bacterial species possess more than three ankyrin-repeat-containing proteins, and two species exhibiting the highest number of ankyrin repeats are obligate intracellular bacteria, W. pipientis (21 proteins) and Coxiella burnetii (20 proteins), although Wu et al. [14] reported slightly different numbers of ankyrin-repeat-containing proteins for these species. A recent genome analysis of a facultative intracellular bacterium, L. pneumophila, revealed 20 proteins with ankyrin repeats [31]. Ankyrin repeats were also found in more than 30 ORFs of the giant virus Acanthamoeba polyphaga Mimivirus [32]. TPR, composed of a motif of 34 amino acids organized in tandem, is also recruited by different proteins and facilitates protein–protein interactions [33]. Its role in the adaptation of parasites to their hosts has been suggested. The R. felis genome exhibits 11 TPR-containing ORFs (seven in the chromosome and four in the pRF plasmid). Only Leptospira interrogans (the agent of leptospirosis), Treponema species (including the agent of syphilis), and L. pneumophila [31] exhibit a high number of both TPR and ankyrin repeats. These organisms are eukaryotic parasites. The cryptococcal crooked neck 1 gene of Cryptococcus neoformans (a yeast), containing 16 copies of TPR, appears associated with its virulence [34]. Host Invasion/Pathogenesis Plasmids often carry out functions that benefit bacteria in their survival or expression of virulence. pRF exhibits two ORFs that are possibly associated with the pathogenesis of R. felis: a hyaluronidase and a patatin-like protein. The hyaluronidase homolog (pRF56) exhibits a significant homology to hyaluronidase NagI (1,297 aa) of Clostridium perfringens. Hyaluronidases, which depolymerize hyaluronic acid—an unbranched polysaccharide ubiquitously present in the extracellular matrix of animal tissues—are known as “spreading factors” [35]. Another ORF (pat2) exhibits a significant homology to patatin-like phospholipases. Its paralog (pat1) was also identified in the chromosome, as already mentioned. Patatin is the major storage glycoprotein found in potato tubers, but also exhibits phospholipase A2 activity for protection from infection. Proteins containing patatin-like domains are more frequently found in pathogenic than in nonpathogenic bacteria. McLeod et al. [7] suggested that patatin-like proteins might be responsible for the phospholipase A2 activity identified some years ago in rickettsiae [36]. Potential host-invasion capacity is also provided by R. felis–specific ORFs found on the chromosome, for instance, a chitinase homolog (RF0413) and a chitin-binding protein homolog (RF0710). Chitin is a homopolymer of N-acetylglucosamine and a major component of the exoskeleton of arthropods and of the peritrophic envelope of insects, a lining layer of the midgut. These genes may facilitate the access of bacteria to the insects' gut epithelial cells. R. felis may also use chitin as a nutrient source, as does Vibrio cholerae [37]. We identified a homolog (RF0268) for ecotin, an E. coli periplasmic protein inhibiting activities of a variety of proteases. Two R. felis–specific ORFs (RF0449 and RF0855) exhibit the complete NACHT NTPase domain. In eukaryotes, this NTPase domain has been found in proteins implicated in apoptosis as well as in immune/inflammatory responses [38]. The presence of this domain in other bacterial ORFs is limited to several lineages, such as cyanobacteria and Streptomyces, and their functions are unknown. Higher eukaryotes and prokaryotes nucleotide-binding domain (HEPN) is a recently identified domain detected in a few prokaryotes. We found four genes (two were split) exhibiting HEPN at the C-terminus, and a nucleotidyl transferase domain at the N-terminus. Among other bacteria, only A. tumefasciens, Thermotoga maritima, and Sinorhizobium melitoti were found to exhibit HEPN-containing genes [39]. The nucleotidyl transferase domain has been associated with several classes of bacterial enzymes responsible for resistance to aminoglycosides. HEPN was also found in the human sacsin protein, a chaperonin implicated in a neurodegenerative disease. Finally, R. felis exhibits an ortholog (RF0371) for R. conorii RickA, which induces its actin-based motility [40]. Phenotypic Post-Genomics Analysis The obligate intracellular nature of R. felis hindered progress in the detailed characterization of its phenotypic diversity. Here, we envisaged post-genomics as a way of associating in vivo phenotypes of these bacteria to genomic features. The presence of pili-associated genes prompted us to investigate, by electron microscopy, the presence of such appendages on the cell surface. This approach led to the first characterization of pili on the surface of a Rickettsia; we observed two forms of pili at the surfaces of R. felis (Figure 3). One form of pili establishes direct contact between bacteria, providing a very typical figure of Mpf apparatus; these pili are probably specialized in conjugation. The other form of pili forms small hair-like projections emerging out from the cell surface; these pili are probably involved in the attachment of the bacteria to other cells. Without pili, many disease-causing bacteria lose their invasion capability. The latter type of pili might be considered as virulence factors, as described for Francisella tularensis [41,42]. As previously mentioned, we also found a RickA homolog in the R. felis genome [40]. Based on this finding, we performed immunofluorescence assays. The orientations of actin filaments beside bacteria are distinct from the stress fibers of the host. This further suggests that R. felis is probably capable of using the actin cytoskeleton to disseminate through eukaryotic cells, a method exploited by other SFG rickettsiae [40] (Figure S8). Another R. felis phenotypic character suggested from genomic analyses (three ORFs for patatin-like proteins) was its hemolytic capacity. We confirmed experimentally that R. felis lyses erythrocytes, this effect being inhibited by dithiothreitol. Another genome-guided discovery was β-lactam inhibition, which reached 57% and 53% of the concentration and the minimal inhibitory concentration, respectively, following 2 h incubation of R. felis with amoxicillin. Despite being preliminary results, these findings illustrate the fact that whole-genome sequencing offers opportunities to rapidly gain a better understanding of the phenotypic characters of a fastidious microorganism. Discussion R. felis is the first obligate intracellular bacterium exhibiting a possible conjugative plasmid. Of the nine previously published studies of members of the Order Rickettsiales (six in Rickettsiaceae, three in Anaplasmateceae), none exhibited a plasmid. Several other obligate intracellular bacteria, such as Chlamydia muridarum, Chlamydophila caviae, C. burnetii, Wigglesworthia glossinidia, and Buchnera aphidicola, are known to possess plasmids. Recently, the reannotation of the genome of Parachlamydia, an obligate intracellular bacteria living in amoeba, predicted an F-like DNA conjugative system encoded in a genomic island [43]. However, no conjugation has yet been observed for those plasmids and genomic island. Transformation of obligate intracellular bacteria remains an elusive goal, although preliminary work on several obligate intracellular bacteria has been reported with limited results [44]. The possible conjugative plasmid identified in R. felis may provide a molecular basis for the future development of new genetic transformation tools in rickettsiae. R. felis is hosted by fleas, as are R. typhi, B. henselae, W. pipientis, and Yersinia pestis. There are surprisingly few common genomic features between R. typhi and R. felis. R. typhi genetically resembles R. prowazekii despite having a lifestyle similar to that of R. felis (Table S2). The comparison with W. pipientis is interesting. This intracellular bacterium also multiplies in arthropods (including fleas) and is transmitted transovarially. The most relevant finding in its genome was the detection of repetitive mobile DNA elements. Many ankyrin repeats and several TPRs were also found. It appears that R. felis and W. pipientis share common genomic features, possibly because of their similar niches (we found two Ct. felis fleas in France coinfected with W. pipientis and R. felis). They both differ significantly from their immediate neighbors [45,46]. Moreover, the phylogenetic relationship and hosts of R. felis and R. prowazekii (transmitted by lice) are comparable with those for B. quintana (transmitted by lice) and B. henselae (transmitted by fleas) [47]. B. henselae exhibits a larger genome with more repeats and integrases than B. quintana. Y. pestis, transmitted by fleas, also exhibits many more insertion sequences than its close relative, Y. pseudotuberculosis [48]. Altogether, flea-infecting bacteria appear to exhibit a specific evolution (i.e., more repeats, transposases, and/or integrases) compared with their non-flea-infecting neighbors. For obligate intracellular bacteria such as rickettsiae, few phenotypic characters have been observed. To date, four intracellular bacterial genomes have been entirely sequenced, the procedure being completed in 7 y or less after their first identification or culture, including R. felis [14,15,49,50]. In the present study, the genome sequencing of R. felis provided evidence of the presence of conjugative plasmids, two types of pili, hemolytic activity, β-lactamase activity, and intracellular motility. We believe that for such recently identified/cultured fastidious organisms, complete genome sequencing is a very potent and timesaving strategy to identify unrecognized phenotypic properties. Materials and Methods Bacterial purification and DNA extraction R. felis (strain California 2) was cultivated on XTC cells growing on RPMI with 5% fetal bovine serum, supplemented with 5 mM L-glutamine. The purification of the bacteria was performed by different steps. First, the bacteria were treated in the presence of 1% trypsine in K36 buffer for 1 h at 37 °C, then centrifuged and digested by DNAseI for 1 h at 37 °C to reduce the eukaryotic DNA contamination. The sample was loaded on a renograffin gradient and the bands of the purified bacteria were washed in K36, treated again by DNAseI. After inactivation with EDTA (50 mM), the bacteria were resuspended in TE, dispatched in 150-μl tubes and stored at −80 °C. Depending on this initial concentration, one or two tubes were diluted in 1 ml of TNE (10 mM Tris [pH 7.5], 150 mM NaCl, 2 mM EDTA) and incubated for 5 h at 37 °C in the presence of lysozyme (2 mg/ml). Lysis was performed for 2 h at 37 °C by adding 1% SDS and RNAseI (25 μg/ml). Overnight treatment with 1 mg/ml of proteinase K followed at 37 °C. After three phenol–chloroform extractions and alcoholic precipitation, the DNA was resuspended in 30 μl of TE and its concentration was estimated by agarose gel electrophoresis. Pulsed-field agarose gel electrophoresis The concentrated bacterial suspension was included in 1% (vol/vol) Incert agarose gel blocks (BMA, Rockland, Maryland, United States). The agarose blocks were digested by Proteinase K (1 mg/ml) (Eurobio Laboratories, Paris, France) in 1% lauroylsarcosine and 0.5 M EDTA (pH 8) (Sigma-Aldrich, St. Louis, Missouri, United States) for 24 h at 50 °C. Fresh Proteinase K was then added and the incubation was continued for 24 h. The blocks were then washed twice in TE (pH 7.6) for 30 min at room temperature. Proteinase K inactivation was performed through incubation in a 4% phenylmethylsulfonyl fluoride (MBI Fermentas, Burlington, Canada) solution for 1 h at 50 °C. This inactivation step was carried out twice. The blocks were then washed two to three times in TE and stored in 0.5 M EDTA (pH 8) at 4 °C. Before restriction enzyme digestion, the agarose blocks were equilibrated twice with TE for 15 min. Digestion was carried out for 4 h, then fresh enzyme was added and the incubation was continued overnight. The digested agarose blocks and molecular-weight markers (Low Range PFG Marker, Lambda Ladder PFG Marker [New England Biolabs, Beverly, Massachusetts, United States]) were equilibrated in 0.5× TBE (50 mM Tris, 50 mM boric acid, 1 mM EDTA). Each agarose block was laid in a 1% PFEG agarose (Sigma-Aldrich) solution in 0.5× TBE. Pulsed-field gel electrophoresis was carried out on a CHEF-DR II device (Bio-Rad, Hercules, California, United States) under different electrophoresis conditions. The 1% agarose gel was run at 200 V using ramped pulse times from 1 to 5 s for 10 h to observe the pattern of small DNA fragments (2–48 kb). The migration was taking place under the following two consecutive conditions: (i) a ramping time from 3 to 10 s at 200 V for 12 h, with the pattern representative for 48- to 242-kb fragments, then (ii) a ramping time from 20 to 40 s at 180 V for 15 h, with the pattern representative for 145- to 610-kb fragments. Shotgun of R. felis genome and sequencing strategy Three shotgun genomic libraries were constructed by mechanical shearing of the genomic DNA using a Hydroshear device (GeneMachine, http://genome.nhgri.nih.gov/genemachine/). DNA fragments were blunt-ended using T4 DNA polymerase (New England Biolabs) and ligated to the BstXI adapter. Fragments of 3, 4.5, and 7 kb were separated on a preparative agarose gel (FMC BioProducts, Rockland, Maryland, United States), extracted with Qiaquick kit (Qiagen, Valencia, California, United States), and ligated into pCDNA2.1 (Invitrogen, Carlsbad, California, United States) for the two smaller inserts and into pCNS (a low copy number vector; C. R., unpublished data) for the largest one. DNA cloning was performed using electrocompetent E. coli DH10B Electromax cells (Invitrogen). Plasmid DNAs were purified and pools of 96 clones were analyzed by gel electrophoresis to validate the libraries. DNA sequencing of insert ends was carried out using Big Dye 3.1 terminator chemistry on an automated capillary ABI3700 sequencer (Applied Biosystems, Foster City, California, United States). Sequences were analyzed and assembled into contigs using Phred, Phrap, and Consed software [51] taking all sequences into account. Sequences were considered valid when at least 75% of the nucleotides had a Phred score of more than 20. The finishing of the genome sequencing included only additional directed reactions that were performed on an ABI3100 sequencer. Two circular plasmid molecules of 63 and 38 kbp, respectively, were identified from the assembled sequences. On the chromosome, three small regions of 41, 155, and 64 bp failed by dropping of sequence. A number of parameters (DMSO, glycerol, hybridization, and elongation temperature) were tested one by one or were combined to sequence over these gaps. We finally succeeded with the association of another type of chemistry, D-rhodamine with 2 M betaine. We designed and used 420 primers (i) to close the sequencing gaps by walking either on shotgun subclones or on the chromosome and (ii) to improve sequence regions of low quality. The integrity of the assembly was validated by comparing the restriction patterns obtained by pulsed-field gel electrophoresis with those deduced from the electronic consensus sequence. The selection of restriction enzymes was based on rare sites. We analyzed single digests of R. felis DNA. The main restriction enzymes used for these studies were ApaI, AfeI, FspI, and SbfI. This comparative study confirmed the predicted length of the R. felis DNA fragments. The structures for pRF and pRFδ plasmids were controlled by specific primer amplifications (see Figure S1). Three PCRs were performed and the amplification results were in agreement with the expected hypothesis. These PCR results validate the two distinct plasmid forms (62.8 and 39 kbp, respectively). Meanwhile, a Southern blot was performed through a pulsed-field electrophoresis gel. Uncut genomic R. felis DNA and R. felis DNA digested by the restriction enzyme PvuI (corresponding to a unique site in the pRF-specific region) were analyzed. These blocks of DNA were loaded twice onto the gel with the molecular-weight markers: Lambda Marker (Bio-Rad) and Low Range PFG Marker (New England Biolabs) as described above, with a pulse time from 1 to 5 s for 12 h at 180 V. The gel was treated and transferred onto Hybond N+ (Amersham Biosciences, Little Chalfont, United Kingdom) with a vacuum blot. The DNA was fixed by heating for 2 h at 80 °C, and the membrane was cut into two pieces. Two probes were derived from two PCR products. The first, pRFh–pRFi (726 bp), was designed within the pRF-specific insert, and the second, pRFa–pRFg (251 bp), was designed to encompass the deletion site of the pRFδ. These two probes were labeled with dCTP32 and hybridized at 65 °C for 17 h on each membrane. Membranes were washed three times in 1× SSC and 0.1% SDS at 65 °C. The exposure time ranged from 6 h to overnight at −80 °C on ECL film. The hybridizations were clearly established on R. felis digested by PvuI and led to one signal with the pRFh–pRFi probe and two signals for the two plasmid structures with the pRFa–pRFg probe at a predicted molecular weight compatible with our prediction (see Figure S2). We tested 30 samples of fleas naturally infected by R. felis obtained from different geographic areas (Algeria [11 fleas], France [15 fleas], and New Zealand [four fleas]) with three pairs of primers: (i) primers designed in the traD gene (pRF37F1/R1), (ii) primers in the pRF plasmid (pRFe–pRFf), and (iii) primers in the pRFδ plasmid (pRFa–pRFg). We confirmed positive PCR products of (i) 196 bp, (ii) 208 bp, and (iii) 251 bp for all the 30 cases. Annotation We predicted protein-coding genes (ORFs) using SelfID [52] as previously described [8]. tRNA genes were identified using tRNAscan-SE [53]. Database searches were performed using BLAST programs [54] against Swiss-Prot/TrEMBL [55], the NCBI CDD database [56], and SMART [57]. The number of transposases, ankyrin/TPR-containing genes, autotransporter domains, and integrases were computed using PSI-BLAST with NCBI/CDD entries related to those domains with an E-value threshold of 10−5. Repeated DNA sequences were identified with the use of RepeatFinder [58], by ignoring the sequence similarity between pRF and pRFδ. To identify Rickettsia palindromic elements, we used hidden Markov models [59] based on the previously identified RPE sequences [60]. By taking advantage of genome colinearity, we identified orthologous relationships of genes in R. felis, R. conorii, R. sibirica, R. prowazekii, and R. typhi with the use of Genomeview (S. Audic, unpublished software). Based on the gene orthology, we defined R. felis–specific ORFs, which were of one of the following three classes: Class I ORFs exhibiting no homologous ORFs in the other four Rickettsia genomes; Class II ORFs exhibiting homologous ORFs but no orthologous ORFs in the other four Rickettsia genomes; and Class III ORFs exhibiting orthologous ORFs in some or all of the other four Rickettsia, all of which exhibit degraded (split or fragmented) genes relative to the R. felis ORF. Plasmid-encoded ORFs were by definition classified into Class I or II. A gene composed of more than one ORF was defined as “split gene.” A gene composed of a single ORF whose length is shorter than 50% of the longest ortholog was defined as a “fragmented” ORF. We used T-Coffee [61] and MEGA [62] for multiple sequence alignment and phylogenetic tree analyses, respectively. The analyses of horizontal gene transfer were performed by BLAST search against the Swiss-Prot/TrEMBL nonredundant database, excluding rickettsial sequences, as well as by methods based on nucleotide composition bias [63,64]. We obtained the minimum number of inversions to associate a pair of Rickettsia genomes using GRAPPA release 2.0 [65]. Ultrastructural characterization of pili by electronic microscopy R. felis cells were carefully collected from the supernatant of XTC cells infected for 5 d and grown at 28 °C. Following centrifugation (400 g, 10 min), bacteria were fixed for 1 h at 4 °C in glutaraldehyde (2.5% in phosphate-buffered saline [PBS]). Cells were then washed in PBS and placed on a carbon–formvar-coated 400-mesh copper grid (Electron Microscopy Sciences, Hatfield, Pennsylvania, United States) for 15 min then negatively stained with 2% phosphotungstic acid for 10 s, before analysis by electron microscopy (Philips Morgagni 268D, Philips Electronics, Eindhoven, the Netherlands). Estimation of β-lactamase activity To evaluate the level of β-lactamase activity, 104 R. felis cells grown on XTC cells and then sonicated were mixed with amoxicillin to a final concentration of 20 μg/ml, and incubated for 2 h at 28 °C. The concentration of amoxicillin was measured in the R. felis + amoxicillin suspension as well as in a suspension of XTC cells without bacteria + amoxicillin, before and after incubation, using high-performance liquid chromatography. In addition, the minimum inhibitory concentrations of these four suspensions were estimated by growth inhibition of a Micrococcus luteus strain. RNA extraction and RT-PCR Approximately 6.5 × 105 bacteria were used to infect one 25-cm3 flask of confluent XTC cells maintained at 28 °C. Infected cells were harvested 48 h later, centrifuged (12,000 g, 10 min), and pellets were immediately frozen in liquid nitrogen before being stored at −80 °C. Total RNA was isolated by using the RNeasy Mini Kit (Qiagen) according to the manufacturer's instructions. At the end of the extraction procedure, all samples were treated with RNase-Free DNase Set (Qiagen) for 30 min. The concentration and quality of isolated RNA were determined with the Agilent 2100 bioanalyzer (Agilent Technologies, Englewood, New Jersey, United States). Aliquots of the DNase-treated total RNA samples were stored at −80 °C until use. RT-PCR was performed from 2 μl of RNA (25 μl final reaction volume) with the Superscript One-Step RT-PCR with Platinum Taq (Invitrogen). Possible DNA contamination was assessed with the Expand high-fidelity polymerase (Roche, Basel, Switzerland). Cycling conditions were 30 min at 50 °C, 5 min at 95 °C, and 40 cycles at 30 s at 95 °C, 30s at 50 °C, and 1 min at 72 °C, followed by a final extension cycle of 7 min at 72 °C. The RT-PCRs were conducted on the PTC-100 thermocycler (Bio-Rad). Amplification products were run on 2% (wt/vol) agarose gels, and the DNA was stained with ethidium bromide. The size of the PCR product was determined by comparison with DNA molecular-weight marker VI (Boehringer Ingelheim, Ingelheim, Germany). Detection of F-actin and immunofluorescence staining Vero cells grown to semiconfluence on glass coverslips were infected with R. felis for 24–48 h at 28 °C in a humidified CO2 incubator (5% CO2). Infected cells were then fixed for 1 h at 4 °C with formaldehyde (3% wt/vol in PBS supplemented with 1 mM MgCl2 and 1 mM CaCl2), washed three times in PBS, and then made permeable with 0.2% Triton X-100 in PBS for 1 min. After three washings in PBS, the coverslips were incubated for 1 h with a monoclonal anti–R. felis antibody. Bacteria were visualized by staining with anti-mouse-Alexa 594 antibody (1:300) and F-actin with FITC-phalloidin (1:250). The coverslips were mounted using Fluoprep (BioMérieux, Marcy-l'Etoile, France) and were examined with a confocal laser scanning microscope using a 100× oil immersion objective lens. Hemolysis experiments Human blood (10 ml) was centrifuged (1,500 g, 10 min), and after three PBS washings, erythrocytes were resuspended in 20 ml of PBS. This suspension (100 μl) was mixed with 800 μl of PBS and 100 μl of rickettsial suspension (106, 105, and 104 bacteria, respectively). In some experiments, rickettsiae were incubated for 1 h at 35 °C in the presence of 2 mM DTT. Complete hemolysis was determined by adding 900 μl of H2O to erythrocytes, and spontaneous hemolysis corresponded to control without bacteria. Following 3 h of incubation at 35 °C, the samples were fixed using paraformaldehyde (0.3% final concentration) and centrifuged. Hemoglobin release was estimated by measurement of the optical density of the supernatant at 545 nm. This experiment was performed in duplicate. Primers The sequences of the primers for PCR and RT-PCR are provided in Table S3. Supporting Information Figure S1 Confirmation of Plasmid Topologies for pRF and pRFδ by PCR (A) The locations of the three primer sets (pRFa–pRFb, pRFc–pRFd, and pRFa–pRFd) used to validate the presence of the two distinct plasmid forms are indicated. (B) The result of the PCR assay with these primers. Two pairs of primers (pRFa–pRFg and pRfh–pRFi) used to obtain the probes for the Southern blot (see Figure S2), as well as another pair of primers (pRF37F1/R1) used in plasmid detection in fleas infected by R. felis, are also indicated in (A). (1.5 MB TIF). Click here for additional data file. Figure S2 Characterization of R. felis Plasmids by Southern Blot The two membranes (A and B) loaded with R. felis genomic DNA (#1/4) and R. felis DNA digested by PvuI (#2/3) were hybridized either by the probe pRFa–pRFg or by pRFh–pRFi. (39 KB PDF). Click here for additional data file. Figure S3 A Model for the Conjugative Plasmid Transfer of R. felis This model is based on gram-negative bacterial conjugation systems involving T4SS. Homologs responsible for different steps of conjugation were identified in the R. felis genome. DNA-processing machinery, plasmid-encoded pRF38/39 (TraATi); the coupling protein, plasmid-encoded pRF43/44 (TraDF) or chromosomally encoded virD4 (RF0469); Mpf apparatus, chromosomal genes for virB2 (RF1075), virB3 (RF0087), virB4 (RF0088), virB6 (RF0089, RF0090, RF0091, RF0092, and RF0093), virB8 (RF0463 and RF0465), virB9 (RF0462 and RF0466), virB10 (RF0467), and virB11 (RF0468); priming for DNA replication in the recipient cell, chromosomally encoded RF0786 (TraC). (57 KB PDF). Click here for additional data file. Figure S4 Phylogenetic Trees for the Patatin-Like Proteins, Thymidylate Kinases, and Small Heat-Shock Proteins Phylogenetic trees were constructed using the neighbor-joining method with Jones-Taylor-Thornton model. (16 KB PDF). Click here for additional data file. Figure S5 Taxonomic Distribution of BLAST Best Hits of R. felis ORFs R. felis ORFs were searched against the nonredundant database (excluding rickettsial sequences). The distribution difference between ORFs with rickettsial orthologs and ORFs lacking rickettsial orthologs remained significant even after the removal of transposase ORFs. (16 KB PDF). Click here for additional data file. Figure S6 Cross-Reactivity of R. felis Western immunoblot showing the preferential cross-reactivity of antibodies with R. felis and R. typhi in a patient with murine typhus (lanes a–c), and with R. felis and R. conorii in a patient with Mediterranean spotted fever (lanes d–f). Lanes a and d, R. conorii antigen; lanes b and e, R. typhi antigen; lanes c and f, R. felis antigen; MM, molecular mass. (30 KB PDF). Click here for additional data file. Figure S7 Domain Structures and the Presence/Absence Patterns of spoT Genes in Different Rickettsia With reference to the S. dysgalactiae Relseq, four (H: 53H, 77H, 78D, and 144D) and five (241R, 243K, 251K, 264D, and 323E) catalytic residues were examined for the (p)ppGpp hydrolase and synthetase domains, respectively. ORF sizes were those for R. felis genes, except SpoT14, for which the R. prowazekii ORF size is indicated. a, absent; Ch, conserved hydrolase catalytic residues; Cs, conserved synthetase catalytic residues; s, split or fragmented genes. (17 KB PDF). Click here for additional data file. Figure S8 Confocal Laser Analysis of R. felis–Infected Vero Cells Bacteria were stained by indirect immunofluorescence using a monoclonal anti–R. felis antibody followed by an anti-mouse-Alexa 594 antibody (red). F-actin was stained with FITC-phalloidin (green). Arrows indicate R. felis with actin tail. (29 KB PDF). Click here for additional data file. Table S1 Distribution of sca Genes among Rickettsia Genomes (30 KB DOC). Click here for additional data file. Table S2 Comparison of Different Features of Bacteria Infecting Fleas with Their Close Relatives (32 KB DOC). Click here for additional data file. Table S3 Nucleotide Sequences of the Primers Used in the Present Study (28 KB DOC). Click here for additional data file. Accession Numbers The genome sequence of R. felis is accessible via GenBank (http://www.ncbi.nlm.nih.gov/Genbank) under the accession numbers: CP000053, CP000054, and CP000055. The EMBL Nucleotide Sequence Database (http://www.ebi.ac.uk/embl) accession number for Cl. perfringens NagI is Q8XM09. The Pfam (http://www.sanger.ac.uk/Software/Pfam/) accession number for ComE3 is PF03772. The Protein Data Bank (http://www.rcsb.org/pdb/) accession number for S. dysgalactiae Relseq is 1VJ7.
- Record: found
- Abstract: found
- Article: found
Molecular Detection of Bartonella quintana, B. koehlerae, B. henselae, B. clarridgeiae, Rickettsia felis, and Wolbachia pipientis in Cat Fleas, France
Jean Marc Rolain, Michel Franc, Bernard Davoust … (2003)
The prevalences of Bartonella, Rickettsia, and Wolbachia were investigated in 309 cat fleas from France by polymerase chain reaction (PCR) assay and sequencing with primers derived from the gltA gene for Rickettsia, the its and pap31 genes for Bartonella, and the 16S rRNA gene for Anaplasmataceae. Positive PCR results were confirmed by using the Lightcycler and specific primers for the rOmpB of Rickettsia and gltA of Bartonella. R. felis was detected in 25 fleas (8.1%), W. pipientis, an insect symbiont, in 55 (17.8%), and Bartonella in 81 (26.2%), including B. henselae (9/81; 11.1%), B. clarridgeiae (55/81; 67.9%), B. quintana (14/81; 17.3%), and B. koehlerae (3/81; 3.7%). This is the first report of the amplification of B. quintana from fleas and the first description of B. koehlerae in fleas from an area outside the United States. Cat fleas may be more important vectors of human diseases than previously reported. Fleas can be found worldwide and are vectors of several important zoonoses, including plague caused by Yersinia pestis ( 1 ). The classic cycle of Rickettsia typhi, the agent of murine typhus, involves rats and the rat flea, Xenopsylla cheopis, the main vector ( 2 ). The disease is transmitted by flea bites or contact with flea feces. Recently, murine typhus has been shown to exist in some endemic foci where neither rats or their fleas are found. Subsequently, in the United States, R. typhi was found to be maintained in the cat flea, Ctenocephalides felis, collected from opossums ( 2 ). R. felis is the recently recognized agent of flea-borne spotted fever, which has been reported in various countries, including the United States, Mexico, Brazil, Germany, and France ( 3 – 6 ). C. felis is apparently the main vector of this new rickettsial disease, and R. felis has been found in this flea in several countries, including the United States ( 2 ), Brazil ( 7 ), Spain ( 8 ), and Ethiopia ( 6 ). A reservoir of flea-borne spotted fever in the United States may be the opossum. Of major importance to the epidemiology of the above rickettsioses is the maintenance of R. typhi or R. felis in their hosts by transovarial transmission ( 9 ) and the fact that neither organism is lethal for fleas. Bartonellae are gram-negative bacteria that cause various human diseases and have various arthropods, such as lice, ticks, and fleas, as vectors ( 10 ). Transmission to humans may also occur by scratches or bites from reservoir hosts, especially cats. Among the genus Bartonella, four species have been isolated from the blood of cats. Two of these species occur worldwide, B. henselae, the agent of cat-scratch disease, and B. clarridgeiae, which might be another agent of cat-scratch disease ( 11 ), and two have been reported only from the United States, B. koehlerae and B. bovis Bermond (B. weissii) ( 12 – 14 ). The main vector of B. henselae infections in cats is most likely the cat flea ( 2 ), whereas the vectors of B. koehlerae and B. weissii are unknown. Detection of B. clarridgeiae in cat fleas by polymerase chain reaction (PCR) amplification has indicated the possible role of fleas as vectors of the organism ( 15 ). Cats are the reservoirs of the bacteria, and the prevalence of cats with B. henselae and B. clarridgeiae bacteremia ranges from 4% to 70%, according to the geographic location ( 15 ). We describe experiments in which we used PCR amplification and DNA sequencing to detect Rickettsia, Bartonella, and Ehrlichia species in C. felis collected from various sites in France. Materials and Methods Source and Identification of Cat Fleas Cat fleas (C. felis), identified according to current taxonomic keys ( 16 ), were obtained from various departments of the Veterinary School of Toulouse, located throughout France. Cat fleas were sent to our laboratory in sealed, preservative-free, plastic tubes at room temperature. To prevent contamination problems, as positive controls we used DNA from R. montanensis (ATCC VR-611) and Bartonella elizabethae F9251 (ATCC 49927), which react with the primer pairs we used in PCR but give sequences distinct from the species we were investigating. For negative controls, we used sterile water and human body lice reared in our laboratory; both negative controls were tested after every seventh cat-flea sample in our PCR. DNA Extraction Fleas were immersed for 5 min in a solution of 70% ethanol/0.2% iodine, washed for 5 min in sterile distilled water and crushed individually in sterile Eppendorf tubes with the tip of a sterile pipette. Their DNA was extracted by using the QIAamp Tissue Kit (QIAGEN, Hilden, Germany) according to the manufacturer’s instructions. This kit was also used to extract DNA from the human body lice reared in our laboratory under standard conditions and used as negative controls. Detection of Bartonella spp., Rickettsia spp., and Anaplasmataceae DNA extracts were amplified in two different runs with different target genes to confirm the results. In the first run we tested all the cat fleas by using genus-specific primers (Table 1) derived from the intergenic spacer region (its gene), the pap 31 gene for Bartonella ( 19 , 20 ), the citrate synthase–encoding gene for Rickettsia ( 17 ), and the 16S RNA gene for Anaplasmataceae ( 21 , 22 ). A total volume of 2.5 μL of the extracted DNA was amplified in a 25-μL reaction mixture containing 12.5 pmol of each primer, 200 μM of dATP, dCTP, dGTP, and dTTP, and 1 U of Elongase in 1X PCR buffer with 0.8 μL of 25 mM MgCl2 (Life Technologies, Cergy Pontoise, France). PCR was carried out in a Peltier Thermal Cycler PTC-200 (MJ Research, Inc., Watertown, MA) with an initial 3-min denaturation step at 95°C, followed by 40 cycles of denaturation at 95°C (30 s), annealing at 50°C (30 s), and extension at 72°C (1 min). Amplification was completed by holding the reaction mixture at 68°C for 3 min to allow complete extension of the PCR products. PCR products were resolved by electrophoresis in 1% agarose gels, and when appropriately sized products were found, they were purified by using Qiagen columns (QIAquick Spin PCR purification kit; QIAGEN) before sequencing. Table 1 Oligonucleotide primers used for polymerase chain reaction amplification and sequencing Primer (reference) Nucleotide sequence Detected organism References CS-877 (gltA gene) GGG GGC CTG CTC ACG GCG G Rickettsia species (17) CS-1273 (gltA gene)
BM59 (rOmpB gene)
B807 (rOmpB gene) ATT GCA AAA AGT ACA GTG AAC A
CCG CAG GGT TGG TAA CTG C
CCT TTT AGA TTA CCG CCT AA Rickettsia species
Rickettsia species
Rickettsia species (17)
(18)
(18) URBarto1 (its gene) CTT CGT TTC TCT TTC TTC A Bartonella species (19) URBarto2 (its gene) CTT CTC TTC ACA ATT TCA AT Bartonella species (19) PAPn1 (pap31 gene) TTC TAG GAG TTG AAA CCG AT Bartonella species (20) PAPn2 (pap31 gene) GAA ACA CCA CCA GCA ACA TA Bartonella species (20) BartogltAForward (gltA gene) TTC CGY CTT ATG GGT TTT GG Bartonella species This report Bartokoehlerae (gltA gene) AAC AAA ATA TTC ATC ATT CAG G B. koehlerae This report Bartoclarridgeiae (gltA gene) AAA GCA ATT TTT TCA AGT TCC B. clarridgeiae This report Bartohenselae (gltA gene) CAT TTC TGT TGG AAA TCC TAG B. henselae This report Bartoquintana (gltA gene)
EHR16SD (16S rRNA gene)
EHR16SD (16S rRNA gene) TTT TAA TGT AAT GCC AGA ATA A
GGT ACC YAC AGA AGA AGT CC
TAG CAC TCA TCG TTT ACA GC B. quintana
Wolbachia
Wolbachia This report
(21)
(21) Any positive sample was tested again by using different primers and real-time PCR technology. For Bartonella, the forward primer of the gltA gene (Table 1) was used for all samples and species-specific primers targeting B. quintana, B. henselae, B. clarridgeiae, and B. koehlerae were designed as reverse primers. For Rickettsia, we used rOmpB-specific primers targeting R. felis (Table 1) ( 18 ). A real-time PCR assay was performed on DNA extracts in a Lightcycler instrument (Roche Biochemicals, Mannheim, Germany). The amplification program began with a denaturation step of 95°C for 120 sec, followed by 40 cycles of denaturation at 95°C for 15 s, annealing at 52°C for 5 sec, and extension at 72°C for 10 s with fluorescence acquisition at 54°C in single mode. Melting curve analysis was done at 45°C to 90°C (temperature transition, 20°C/s) with stepwise fluorescence acquisition by real-time measurement of fluorescence directly in the clear glass capillary tubes. Sequence specific standard curves were generated by using 10-fold serial dilutions (105 to 106 copies) of standard bacterial concentration of Bartonella. The positive PCR products of the two runs for Bartonella, Rickettsia, and Wolbachia were sequenced by using the d-rhodamine terminator cycle-sequencing ready reaction kit (PE Applied Biosystems, Les Ulis, France) according to the manufacturer’s protocol. Sequences obtained were compared with those in the GenBank DNA database by using the program BLAST (version 2.0, National Center for Biotechnology Information; available from URL: http://www.ncbi.nlm.nih.gov). Results Overall, 309 cat fleas from 92 cats from all areas of France (north, west, east, and south) were tested. Almost two thirds (60/92; 65%) of the cats lived both outdoors and indoors, 20% lived predominantly outdoors, and 15% lived exclusively indoors. Our negative controls consistently failed to yield detectable PCR products, whereas our positive controls always gave expected PCR products. We found a total of 89 fleas (28.8%) that were infected: 25 were positive for R. felis (25/309; 8.1%) as determined by citrate synthase–gene sequencing, and 81 were positive for Bartonella species (81/309; 26.2%) as determined either by its gene or pap31 gene sequencing (Table 2). The sequences of the DNA amplicons we obtained were identical to those of R. felis (Genbank accession no. U33922), B. henselae (Genbank accession no. AF369527), B. clarridgeiae (GenBank accession no. AF312497), B. koehlerae (GenBank accession no. AF312490), or B. quintana (GenBank accession no. AF368391). The Bartonella species we identified were B. henselae (9/89; 11.1%), B. clarridgeiae (55/89; 67.9%), B. quintana (14/89; 17.3%), and B. koehlerae (3/89; 3.7%). Our results were confirmed by a second PCR with the Lightcycler and specific primers for the gltA gene for Bartonella and rOmpB for Rickettsia. Because the primers were species-specific, we were able to demonstrate that none of the fleas contained more than one Bartonella species. Seventeen fleas, however, contained R. felis and B. quintana (12 fleas) or B. clarridgeiae (5 fleas) (Table 2). Lastly, in 55 fleas (17.8%) our Anaplasmataceae-specific primers amplified DNA with a sequence identical to that of Wolbachia pipientis (Genbank accession no. U23709). Table 2 Repartition of 89 Bartonella-positive fleas B. henselae B. clarridgeiae B. quintana B. koehlerae No coinfection Total Rickettsia felis 0 5 12 0 8 25 No coinfection 9 50 2 3 17 No. of fleas positive for Bartonella 9 55 14 3 81 Discussion Rickettsia and Bartonella infections occur worldwide and may cause serious diseases in people. Most of these pathogenic bacteria are transmitted to people by arthropod vectors such as ticks, fleas, and lice, which are also involved in the maintenance of the bacteria. The detection of these pathogenic bacteria in their vector arthropods can be used in epidemiologic studies and control strategies ( 2 ). We tested cat fleas from around France for the presence of Rickettsia, Bartonella, and Ehrlichia species. We found DNA of R. felis and various Bartonella species in these fleas by using PCR with primers for different specific genes and sequencing to confirm our results. All our negative controls gave no PCR products, and all fleas that tested positive were also positive with other PCRs with different target genes and different techniques. We report for the first time the presence of R. felis in cat fleas from France. This bacteria has been detected previously in wild cat fleas from various countries, including the United States ( 2 ), Ethiopia ( 6 ), and, very recently, Spain ( 8 ), and Brazil ( 7 ). Since C. felis has a worldwide distribution and infestation with these fleas is very common, some have assumed that R. felis and flea-borne spotted fever should occur worldwide. We found that 8.1% of fleas from domestic cats were infected with R. felis, suggesting that clinical cases in humans may be prevalent in France and probably in Europe. In the United States, the infection rates of fleas, as determined by PCR amplification, have been reported to vary from 43% to 93% ( 2 ). Since bacteria are maintained transovarially, R. felis may be used as a marker to follow changes in the infection rates over time ( 2 ). In people, clinical cases of flea-borne spotted fever have been reported in the United States (Texas) ( 3 ), Mexico ( 4 ), France, and Brazil ( 6 ), and, very recently, in Germany ( 5 ). Preliminary serologic results indicate that flea-borne spotted fever might occur in France ( 6 ), and we have now shown that fleas in France are infected with R. felis. The disease is probably more prevalent than expected, even if the risk of transmission by fleas is unknown. Cross-reactions in serologic testing for R. felis are unpredictable in our experience, and thus serologic tests for R. felis should be performed in patients suffering from fever of unknown origin. Our findings of R. felis in French fleas indicate, then, that R. felis should be used, along with other spotted fever group rickettsiae, in serologic tests on patients suspected of having a spotted fever group rickettsiosis. To date, B. henselae is the only recognized agent of cat-scratch disease; epidemiologic studies have implicated cats, which remain bacteremic for months to years, as the major reservoirs of B. henselae ( 23 , 24 ). Its DNA has been amplified from fleas found on bacteremic cats, and transmission of B. henselae to cats by C. felis has been demonstrated ( 23 , 25 , 26 ). Flea infestation was found to be more common in bacteremic cats than in nonbacteremic cats. The prevalence of B. henselae in fleas in our study was 3%, whereas B. henselae has been isolated from the blood of 4% to 70% of cats, depending on location, cat population, and flea infestation rate but not depending on the infection rate of fleas on the cats or the seropositivity of the cats ( 15 , 25 ). Our study has confirmed that B. clarridgeiae may be detected in fleas, and we found a 17.8% prevalence in infected fleas, that is, 67.9% of all fleas positive for Bartonella by PCR. Until now, B. henselae was the most common species isolated from cats, and the prevalence of B. clarridgeiae has ranged from 16% to 30% ( 25 ). B. clarridgeiae is more prevalent in European cats than in American cats ( 25 ). The difference of recovery rate of B. clarridgeiae in these studies may be explained by the fact that the number of bacteria in the blood of cats infected with B. clarridgeiae is low (unpub. data) and because B. clarridgeiae grows more slowly in culture ( 27 ). Also, freezing of whole blood improves the recovery rate of B. clarridgeiae since bacteria are probably localized in erythrocytes ( 15 ). We report, for the first time, the presence of B. koehlerae in cat fleas. This recently described species has only been reported in the blood of cats in the United States ( 12 ), and our findings support the idea that this bacteria might be transmitted by cat fleas, like other Bartonella, and that it has a worldwide distribution. The prevalence of this new Bartonella species in cats remains unknown and is probably underestimated because the bacteria are extremely fastidious and only grow on chocolate agar and not on heart infusion agar with rabbit blood, Columbia, or on sheep blood agar, which are currently widely used for the isolation of Bartonella ( 12 ). We also report for the first time that cat fleas can contain B. quintana; this is surprising because, to date, the body louse was the only known vector of this species, and people are the only known natural reservoirs. Previously, however, in two clinical reports of chronic adenopathy attributed to B. quintana infection, the only epidemiologic risk identified was the presence of infected cat fleas ( 28 , 29 ). The role of the cat flea as a potential vector of B. quintana and its related diseases needs to be clarified, and new investigations of patients in contact with cats and their fleas are indicated. Also, we have reported cases of endocarditis due to B. quintana for which no epidemiologic risks (alcoholism or homelessness) were found ( 30 , 31 ). These findings and the fact that B. quintana DNA sequences can be found in ticks ( 32 ) indicate that other arthropod vectors apart from lice may be involved in the epidemiology of B. quintana infections. Coinfection of cats with B. clarridgeiae and B. henselae has been reported, as has infection with the two different genotypes of B. henselae, Marseille and Houston ( 25 , 33 ). Recent reports suggest that interspecific competition between closely related rickettsiae may control rickettsial establishment in arthropods (dual infections in arthropod vectors are rare and have not yet been observed in individual fleas) ( 34 ). Fleas, however, feed intermittently on different hosts and thus may acquire multiple bacterial strains and pass these to their progeny transovarially ( 2 ). Our finding of R. felis with either B. clarridgeiae or B. quintana in fleas suggests that dual infections may occur in humans infected by flea feces. We did not, however, find fleas infected with several species of Bartonella although cats may be bacteremic with two different species. We believe that the techniques we used should have been sensitive enough to detect such coinfections, and that competition between two Bartonella species may result in one species being eliminated and the flea becoming infected with the other Bartonella species. Further studies are indicated to confirm this possibility. Finally, we report, for the first time, the presence of W. pipientis in fleas, which we found while seeking Ehrlichia using Anaplasmataceae-specific primers. This bacteria is known to be an endosymbiont of several arthropods, primarily insects, and it has a role in parthenogenesis ( 35 ). Any effects that it might have on fleas, however, are unknown. In summary, our study has provided evidence that cat fleas are commonly infected by R. felis and Bartonella species in France. Further, we have shown for the first time that B. quintana may infect fleas. The description of B. quintana-related diseases in patients with histories of contact with fleas indicates the possibility that fleas may be vectors of the organism. Finally, we reported for the first time the presence of B. koehlerae outside the United States.
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- Abstract: found
- Article: not found
Rickettsial infection in Dermacentor variabilis (Acari: Ixodidae) inhibits transovarial transmission of a second Rickettsia.
This study examined the ability of ticks to maintain multiple species of spotted fever group rickettsiae via transovarial transmission. Using a capillary feeding method, previously established Ricketisia montana- and Rickettsia rhipicephali-infected cohorts of Dermacentor variabilis (Say) were exposed to R. rhipicephali and R. montana, respectively, in two reciprocal challenge experiments. Eggs collected from individual females, for two successive generations, of each cohort were assessed for rickettsial infection by polymerase chain reaction for each challenge experiment. Assessment of the eggs from challenged ticks identified that both B. montana- and R. rhipicephali-infected ticks were refractory to their respective challenge rickettsiae. The prechallenged infection rate for both F1 and F2 generations (100%) of the B. montana-infected cohort was resistant to transovarial transmission of the second rickettsia species, and only R. montana was detected in the eggs of FL = (50%) and F2 = (74%) challenged females. The R. rhipicephali-infected cohort maintained a lower level of infection (20%) in the population and did not transovarially transmit the challenge species, however, detectable levels of infection were lost after the first generation. Second-generation ticks, no longer infected with B. rhipicephali, became susceptible to infection with R. montana and female ticks (approximately 4%) were able to transmit R. montana to their progeny. The resistance of the ovaries to co-infection and apparent host-specific nature of infection suggests that rickettsial infection of tick ovaries may alter the molecular expression of the oocytes so as to preclude secondary infection with other rickettsiae.
Author and article information
Contributors
: Role: Editor
Journal
Journal ID (nlm-ta): PLoS ONE
Journal ID (publisher-id): plos
Journal ID (pmc): plosone
Title:
PLoS ONE
Publisher:
Public Library of Science
(San Francisco, USA
)
ISSN
(Electronic):
1932-6203
Publication date Collection: 2008
Publication date
(Electronic):
30
July
2008
Volume: 3
Issue: 7
Electronic Location Identifier: e2805
Affiliations
[1
]Department of Pathobiological Sciences, Louisiana State University, Baton Rouge, Louisiana,
United States of America
Author notes
* E-mail:
kmacaluso@
123456vetmed.lsu.edu
Article
Publisher ID: 08-PONE-RA-04166R2DOI: 10.1371/journal.pone.0002805
PMC ID: 2474969
PubMed ID: 18665265
SO-VID: ece6dfff-cd41-4065-9085-6e918b670a3e
Copyright © Reif et al. This is an open-access article distributed under the terms of the Creative
Commons Attribution License, which permits unrestricted use, distribution, and reproduction
in any medium, provided the original author and source are credited.
History
Date
received
: 2
April
2008
Date
accepted
: 20
June
2008
Page count
Pages: 8
Categories
Subject:
Research Article
Subject:
Ecology/Environmental Microbiology
Subject:
Microbiology/Microbial Growth and Development
Subject:
Infectious Diseases/Bacterial Infections
Subject:
Public Health and Epidemiology/Infectious Diseases
ScienceOpen disciplines: Uncategorized
Data availability:
ScienceOpen disciplines: Uncategorized