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Appl Environ Microbiol. Mar 2010; 76(6): 1718–1731.
Published online Jan 22, 2010. doi:  10.1128/AEM.02988-09
PMCID: PMC2838022
Wide Dispersal and Possible Multiple Origins of Low-Copy-Number Plasmids in Rickettsia Species Associated with Blood-Feeding Arthropods[down-pointing small open triangle]
Gerald D. Baldridge,4* Nicole Y. Burkhardt,4 Marcelo B. Labruna,1 Richard C. Pacheco,1 Christopher D. Paddock,2 Philip C. Williamson,3 Peggy M. Billingsley,3 Roderick F. Felsheim,4 Timothy J. Kurtti,4 and Ulrike G. Munderloh4
Department of Entomology, University of Minnesota, St. Paul, Minnesota 55108,4 Departamento de Medicina Veterinaria Preventiva e Saude Animal, Faculdade de Medicina Veterinaria e Zootecnia, Universidade de Sao Paulo, Sao Paulo, Brazil,1 Infectious Diseases Pathology Branch, Division of Viral and Rickettsial Diseases, Centers for Disease Control and Prevention, Atlanta, Georgia,2 Department of Forensic and Investigative Genetics, University of North Texas Health Science Center, Fort Worth, Texas3
*Corresponding author. Mailing address: Department of Entomology, University of Minnesota, 1980 Folwell Ave., St. Paul, MN 55108. Phone: (612) 624-3688. Fax: (612) 625-5299. E-mail: baldr001/at/umn.edu
Received December 10, 2009; Accepted January 14, 2010.
Plasmids are mobile genetic elements of bacteria that can impart important adaptive traits, such as increased virulence or antibiotic resistance. We report the existence of plasmids in Rickettsia (Rickettsiales; Rickettsiaceae) species, including Rickettsia akari, “Candidatus Rickettsia amblyommii,” R. bellii, R. rhipicephali, and REIS, the rickettsial endosymbiont of Ixodes scapularis. All of the rickettsiae were isolated from humans or North and South American ticks. R. parkeri isolates from both continents did not possess plasmids. We have now demonstrated plasmids in nearly all Rickettsia species that we have surveyed from three continents, which represent three of the four major proposed phylogenetic groups associated with blood-feeding arthropods. Gel-based evidence consistent with the existence of multiple plasmids in some species was confirmed by cloning plasmids with very different sequences from each of two “Ca. Rickettsia amblyommii” isolates. Phylogenetic analysis of rickettsial ParA plasmid partitioning proteins indicated multiple parA gene origins and plasmid incompatibility groups, consistent with possible multiple plasmid origins. Phylogenetic analysis of potentially host-adaptive rickettsial small heat shock proteins showed that hsp2 genes were plasmid specific and that hsp1 genes, found only on plasmids of “Ca. Rickettsia amblyommii,” R. felis, R. monacensis, and R. peacockii, were probably acquired independently of the hsp2 genes. Plasmid copy numbers in seven Rickettsia species ranged from 2.4 to 9.2 per chromosomal equivalent, as determined by real-time quantitative PCR. Plasmids may be of significance in rickettsial evolution and epidemiology by conferring genetic plasticity and host-adaptive traits via horizontal gene transfer that counteracts the reductive genome evolution typical of obligate intracellular bacteria.
The alphaproteobacteria of the genus Rickettsia (Rickettsiales; Rickettsiaceae) have undergone the reductive genome evolution typical of obligate intracellular bacteria, resulting in A/T-rich genomes (1.1 × 106 to 1.5 × 106 bp) with a high content of pseudogenes undergoing elimination (3, 10, 20, 26). Initial sequencing of rickettsial genomes focused on the important arthropod-borne pathogens Rickettsia prowazekii, Rickettsia conorii, and Rickettsia typhi and appeared to confirm the prevailing belief that plasmids were absent and transposons were rare among Rickettsia spp. (2, 28, 39, 44). As mobile genetic elements in bacteria, plasmids and transposons drive horizontal gene transfer (HGT) and the acquisition of virulence determinants and environmental adaptive traits (30, 43, 60, 70). Subsequent sequencing of the Rickettsia felis genome revealed the surprising presence of abundant transposase paralogs and the 63-kbp pRF plasmid, with 68 open reading frames (ORFs) encoding predicted proteins, as well as a 39-kbp deletion form, pRFδ (45). Although pRF was suggested to be conjugative, it was initially thought to be unique among the rickettsiae, a reasonable inference given that plasmids are uncommon among the reduced genomes of obligate intracellular bacteria and were previously unknown in the Rickettsiales (3, 4, 13). However, a phylogenetic analysis implied an origin for pRF in ancestral rickettsiae and the possible existence of other rickettsial plasmids (28), which was soon confirmed by the cloning of the 23.5-kbp pRM plasmid from Rickettsia monacensis (6). Some of the 23 ORFs on pRM had close pRF homologs, and both plasmids carried transposon genes and the molecular footprints of transposition events associated with HGT from other bacterial taxa.
The discoveries of pRF and pRM made obsolete the long-held dogma that plasmids were not present in members of the genus Rickettsia and implied a source of unexpected genetic diversity in the reduced rickettsial genomes, particularly if potentially conjugative plasmids carrying transposon genes proved to be common among members of the genus. That hypothesis gained credence when pulsed-field gel electrophoresis (PFGE) and Southern blot surveys (7) using plasmid gene-specific probes demonstrated plasmids in Rickettsia helvetica, “Candidatus Rickettsia hoogstraalii” (38), and Rickettsia massiliae and possible multiple plasmids in “Candidatus Rickettsia amblyommii” (71) isolates. The same study demonstrated the loss of a plasmid in the nonpathogenic species Rickettsia peacockii during long-term serial passage in cultured cells and the absence of a plasmid in Rickettsia montanensis M5/6, an isolate with a long laboratory passage history. Genome sequencing of R. massiliae and Rickettsia africae revealed the 15.3-kbp pRMA and 12.4-kbp pRAF sequences, with 12 and 11 ORFs, respectively, that were more similar to those of pRF than to those of pRM (11, 24).
The absence of plasmids in R. montanensis and important Rickettsia pathogens maintained as laboratory isolates has left unresolved the question of the true extent of plasmid distribution among Rickettsia spp. Until recently, the genus was thought to consist of closely related species, known chiefly as typhus and spotted fever pathogens transmitted by lice, fleas, mites, and ticks (31). It is now apparent that many, and possibly most, Rickettsia spp. inhabit a diverse range of arthropods that do not feed on blood, as well as leeches, helminths, crustaceans, and protozoans, suggesting an ancient and complex evolutionary history (54). A multigene phylogenetic analysis of the Rickettsiales resulted in a “molecular clock” which indicated that the order arose from a presumably free-living ancestor and then adapted to intracellular growth during the appearance of metazoan phyla in the Cambrian explosion (76). A transition to a primary association with arthropods followed during the Ordovician and Silurian periods. The genus Rickettsia arose approximately 150 million years ago and evolved into several clades, including the early-diverging hydra and torix lineages associated with leeches and protozoans. A rapid radiation occurred about 50 million years ago in the arthropod-associated lineages (76).
Whole-genome sequencing has led to a revision of phylogenetic relationships among Rickettsia spp. associated with blood-feeding arthropods (10, 26, 28). A newly defined ancestral group (AG) contains the earliest-diverging species, Rickettsia bellii and Rickettsia canadensis, while R. prowazekii and R. typhi, transmitted by lice and fleas, respectively, constitute the typhus group (TG). A proposed transitional group (TRG), consisting of the mite-borne Rickettsia akari, the flea-borne R. felis, and the tick-borne Rickettsia australis, bridges the genotypic and phenotypic differences between the TG and the much larger spotted fever group (SFG), consisting of tick-borne rickettsiae (28). However, some presumptive SFG rickettsiae remain poorly characterized and are of uncertain phylogenetic status, while the accumulation of genomic data from rickettsiae found in a diverse range of invertebrate hosts may have profound impacts on the currently understood phylogeny of rickettsiae associated with blood-feeding arthropods. For example, it appears that the above AG and TRG species have many close relatives in insects (76). Despite the recent phylogenomic advances, the genetic and host-adaptive mechanisms underlying the evolution of arthropod-transmitted pathogens of vertebrates from ancestral Rickettsia spp., including any possible role of plasmids, remain poorly understood.
In this report, we have taken advantage of recent isolations of rickettsiae from North and South America to conclusively demonstrate that low-copy-number plasmids are indeed common in low-passage isolates of AG, TRG, and SFG rickettsiae. The only exceptions were multiple isolates of R. parkeri, obtained from ticks and human eschar biopsy specimens and newly recognized as a mildly pathogenic SFG rickettsia (49, 50, 52, 79), and the previously characterized species R. montanensis (7). We confirmed that some Rickettsia isolates harbor more than one plasmid by cloning and sequencing multiple plasmids from “Ca. Rickettsia amblyommii” isolates AaR/SC and Ac/Pa, and we obtained PCR- and gel-based evidence that supported genome sequence evidence for the existence of multiple plasmids in REIS, the rickettsial endosymbiont of Ixodes scapularis. Phylogenetic analysis provided strong evidence for multiple plasmid incompatibility groups and possible multiple origins of plasmid-carried parA genes in the genus Rickettsia. Other than genes encoding plasmid replication initiation and partitioning proteins, the newly sequenced “Ca. Rickettsia amblyommii” plasmids resembled the previously sequenced rickettsial plasmids in sharing limited similarities in coding capacity (6, 7, 22). However, we have previously drawn attention to the presence of hsp genes, encoding α-crystalline small heat shock proteins, as a conserved feature of most rickettsial plasmids that may play a role in host adaptation (7). Phylogenetic analysis indicated that the hsp2 genes were plasmid specific, while the hsp1 genes found on four rickettsial plasmids may have been acquired by a chromosome-to-plasmid transfer event in a TRG-like species.
Rickettsiae.
All rickettsiae were originally isolated in Vero (primate) cells, unless stated otherwise (Table (Table1).1). Whenever possible, low-passage isolates were used to eliminate the possibility that isolates testing negative for the presence of plasmids had done so due to the loss of plasmids during serial passage in the laboratory (7). The identity of each isolate was confirmed before analysis for the presence of plasmids by PCR amplification, DNA sequencing, and comparison to GenBank reference sequences of the gltA, ompA, and 17-kilodalton antigen genes, commonly used to genotype Rickettsia spp. (1, 23). Rickettsiae were cultivated in Vero E6 or tick ISE6 cells as described previously (41), with the exception of REIS, the rickettsial endosymbiont of Ixodes scapularis (42), which was cultivated in IRE11 cells. Rickettsiae were released from host cells, separated from cellular debris by filtration, and concentrated by centrifugation (7).
TABLE 1.
TABLE 1.
Origins and passage histories of Rickettsia species evaluated for plasmids and plasmid copy number
PFGE.
Purified rickettsiae were resuspended and embedded in low-melting-point agarose, digested with proteinase K in the presence of sodium lauryl sarcosine and 0.5 M EDTA, and subjected to PFGE as described previously (6), except that R. bellii isolates were run in 0.9% agarose gels to enhance the separation of large plasmid isomers from chromosomal DNA.
Southern blot analyses.
Rickettsial DNA was prepared and electrophoresed in PFGE gels, depurinated, and transferred to a Zeta Probe GT genomic membrane (Bio-Rad, Hercules, CA) as described previously (5). The blots were hybridized with digoxigenin-labeled probes prepared by PCR amplification of R. monacensis pRM plasmid genes, washed, and exposed to Kodak X-Omat AR film (6, 7).
Determination of plasmid copy numbers.
Plasmid copy numbers were determined as ratios of the single-copy plasmid hsp2 and chromosomal gltA genes, using real-time quantitative PCR (QPCR) and the relative quantification method (36, 37). To construct plasmids for generation of species-specific standard curves, gltA sequences were PCR amplified from each Rickettsia sp., using a 0.25 μM concentration (each) of the primers CS877F and CS1273R (63) and PfuTurbo Hotstart DNA polymerase (Stratagene, La Jolla, CA). Cycle parameters on a Stratagene Robocycler instrument were as follows: 1 cycle at 95°C for 2 min; 40 cycles at 95°C for 30 s, 42°C for 30 s, and 68°C for 1 min; and a final 10-min cycle at 68°C. The PCR products were purified on spin columns (Qiagen, Valencia, CA), treated with Taq enzyme (Promega, Madison, WI) to create 3′-A overhangs, and cloned into the pCR4 vector (see below). The hsp2 sequences were amplified using the Hsp2F3-Hsp2R3 primer pair (Table (Table2)2) (all primers were synthesized by Integrated DNA Technologies, Coralville, IA) and GoTaq DNA polymerase (Promega) with the following cycle parameters: 1 cycle at 95°C for 2 min; 40 cycles at 95°C for 30 s, 46°C for 30 s, and 72°C for 1 min; and a final 7-min period at 72°C. Spin column-purified products were ligated into the pCR4 vector with a Topo TA cloning kit and transformed into Oneshot Top10 competent cells according to the manufacturer's protocol (Invitrogen, Carlsbad, CA). Plasmid DNAs from kanamycin-resistant clones were prepared using a High Pure plasmid isolation kit (Roche, Indianapolis, IN) and sequenced to verify the identities of the cloned PCR products (ABI 377 automated sequencer at Advanced Genetic Analysis Center, University of Minnesota).
TABLE 2.
TABLE 2.
Species-specific and general PCR amplification primers for hsp2, gltA, and parA genesb
For QPCR, serial dilutions of plasmid and rickettsial genomic DNAs were adjusted to 10 ng DNA per sample with salmon sperm DNA (Promega) and then transferred to 96-well plates for PCR amplification. The hsp2 sequences were amplified with species-specific primers and the gltA sequences with the CS-F-CS-R primer pair (69) or species-specific primers (Table (Table2),2), using Brilliant or Brilliant II SYBR green QPCR master mix, a 240 nM concentration of each primer, and an Mx3005p qPCR cycler according to the manufacturer's protocol (Stratagene). Cycle parameters were 1 cycle at 95°C for 10 min, followed by 40 cycles at 95°C for 30 s, 54°C for 1 min, and 72°C for 30 s. Amplification specificity was confirmed by melting-curve analysis. Data acquisition and analysis were carried out with the MxPro software package, version 4. Plasmid copy numbers per chromosomal equivalent were determined by referencing values obtained for genomic samples to the standard curves and were expressed as the mean ratios of hsp2/gltA amplification products from three separate plates, with all samples run in triplicate on each plate.
Cloning and sequencing of “Ca. Rickettsia amblyommii” plasmids.
Genomic DNAs from “Ca. Rickettsia amblyommii” isolates AaR/SC and Ac/Pa were partially digested with HpaI and SwaI (New England Biolabs, Ipswich, MA), whose recognition sites occur at low frequencies in rickettsial DNA. The digestion products were ligated into the linear plasmid vector pJAZZ (Lucigen Corp., Madison, WI), which is optimized for cloning of large, A/T-rich inserts. Colony lifts of the libraries were hybridized with probes derived from the R. monacensis pRM6, -8, and -16 plasmid genes (6). Positive clones that contained inserts of approximately 6.5 to 25 kbp were Sanger sequenced by a primer walking strategy, using custom primers (Invitrogen, Carlsbad, CA) and a BigDye Terminator v.3.1 cycle sequencing kit (Applied Biosystems, Inc., Foster City, CA) with an ABI Prism 310 or 3130xl genetic analyzer (Applied Biosystems). Sequence analysis, assembly, and editing were performed using Sequencher v4.7 (Gene Codes Corporation, Ann Arbor, MI). The edited sequence was prepared for submission to GenBank by use of the Sequin application, version 9.5 (NCBI), and was annotated using the NCBI Prokaryotic Genomes Automatic Annotation Pipeline (PGAAP).
Phylogenetic analyses of predicted ParA and Hsp proteins.
Maximum parsimony (67) and neighbor-joining (61) phylogenetic analyses were conducted using ParA, Hsp1, and Hsp2 proteins deduced from sequenced PCR products (see below) or plasmids and/or chromosomal sequences of Rickettsia spp. and members of other bacterial taxa available at GenBank. Sequences were manually aligned with Clustal X, edited to remove gaps, and imported into PAUP* 4.0b (72) to construct phylogenetic trees. The node stability of dendrograms was estimated with 1,000,000 random bootstrap replications (21). Additional Hsp2 sequences from R. bellii and “Ca. Rickettsia hoogstraalii” were derived from sequenced GoTaq PCR amplification products obtained using the Hsp2F3-Hsp2R primer pair and from R. helvetica products obtained using the Hsp2F3-Hsp2R3 and Hsp2F-Hsp2R primer pairs (Table (Table2).2). Cycle parameters were as follows: 1 cycle at 95°C for 2 min; 40 cycles at 95°C for 30 s, 46°C for 30 s, and 72°C for 1 min; and a final 7-min step at 72°C, except that R. helvetica-specific Hsp2F-Hsp2R-containing reaction mixtures were annealed at 50°C. The ParA sequences of REIS were derived from sequenced Accu Taq (Sigma, St. Louis, MO) amplification products obtained using pREIS1, -2, -3, and -4 primer pairs (Table (Table2).2). Cycle parameters were as follows: 1 cycle at 94°C for 3 min; 35 cycles at 94°C for 30 s, 50°C for 30 s, and 72°C for 1 min; and a final 5-min step at 72°C.
Nucleotide sequence accession numbers.
The “Ca. Rickettsia amblyommii” plasmid and PCR-amplified rickettsial hsp2 sequences reported here have been deposited in GenBank (http://www.ncbi.nlm.nih.gov/GenBank) under the following accession numbers: pRAM18, GU322808; pRAM23, GU322807; R. bellii hsp2, GU180086; R. helvetica hsp2, GU180087; and “Ca. Rickettsia hoogstraalii” hsp2, GU180088.
Plasmids occur in R. rhipicephali, REIS, and R. akari.
In this study, we expanded the range of arthropod-associated Rickettsia spp. analyzed for the presence of plasmids, using our previously developed PFGE and Southern blot methods (7). In a PFGE gel lane loaded with an extract of the South American R. rhipicephali HJ#5 isolate cultivated in tick ISE6 cells, plasmid isomers were visible as SYBR green-stained DNA bands that migrated at approximately 20, 45, and 55 kbp relative to the linear DNA markers (Fig. (Fig.1A).1A). The relative migration of the plasmid isomer and DNA markers does not allow precise estimation of plasmid size because the plasmid isomers represent a mixture of supercoiled and open circular forms as well as presumptive single-stranded replication intermediates (6). Host cell mitochondrial DNA bands (7) comigrated with those for an uninfected ISE6 cell extract, at approximately 25 and 35 kbp. In a lane loaded with REIS cultivated in IRE11 cells, plasmid isomer bands migrated at approximately 40, 45, and 55 kbp, but only the smaller mitochondrial band was visible. The gel was Southern blotted and hybridized (Fig. (Fig.1B)1B) with a probe derived from the R. monacensis pRM16 gene, encoding a DnaA-like replication initiator protein. In the lane with R. monacensis IrR/Munich, the characteristic hybridization pattern of linear, open circular, and supercoiled pRM isomers was present, with the major bands migrating at approximately 25 and 45 kbp (6, 7). In the R. rhipicephali and REIS lanes, four- and three-band hybridization patterns, respectively, were present at positions of the plasmid bands in Fig. Fig.1A1A consistent with their identity as plasmid isomers. A replicate gel was hybridized (Fig. (Fig.1C)1C) with a probe derived from the pRM6 gene, hsp2, which produced the expected three-band hybridization pattern in the R. monacensis lane. In the R. rhipicephali lane, the pRM6 probe hybridized in a two-band pattern versus the four-band pattern of the pRM16 probe, suggesting the presence of plasmid isomers with different gene complements. In the REIS lane, the pRM6 probe replicated the hybridization pattern of the pRM16 probe, but a fourth, higher-molecular-weight band was also present. A second replicate gel was hybridized (Fig. (Fig.1D)1D) with a probe derived from the pRM8 gene, encoding a RecD/TraA helicase protein. It hybridized as expected in the R. monacensis lane but to only two plasmid isomer bands in the REIS lane and did not produce a signal in the R. rhipicephali lane. Analysis of R. akari Bronx revealed a plasmid that migrated as a predominant isomer at approximately 47 kbp (Fig. (Fig.2A).2A). It hybridized to the pRM16 probe (Fig. (Fig.2B)2B) but not to the pRM6 and pRM8 probes (Fig. 2C and D).
FIG. 1.
FIG. 1.
Presence of plasmids in R. rhipicephali from South America and REIS from North America. (A) PFGE of DNAs from REIS isolate Camp Ripley, R. rhipicephali isolate HJ#5, and R. monacensis isolate IrR/Munich. Relative migration positions of rickettsial chromosomal (more ...)
FIG. 2.
FIG. 2.
Presence of plasmids in R. akari and “Ca. Rickettsia amblyommii” isolates and absence of plasmids in R. parkeri isolates from North America. (A) PFGE of DNAs from R. akari isolate Bronx, R. parkeri isolates Ft. Story, Portsmouth, and Tates (more ...)
In summary, the results demonstrated that the mite-associated TRG member R. akari and three SFG Rickettsia spp. isolated from ticks collected on three continents carry plasmids. R. rhipicephali and REIS appear to have more than one plasmid. Four putative plasmid sequences from REIS were obtained as a consequence of the I. scapularis genome sequencing project and were deposited in GenBank as the circular pREIS1 and pREIS2 plasmids, the incompletely circularized pREIS3 plasmid, and the plasmid scaffold (accession no. NZ_GG688316) (referred to here as pREIS4). We confirmed the REIS identity of the parA genes on those plasmids by PCR amplification of DNA extracts of our REIS isolate, using primers flanking the parA genes in the GenBank-deposited pREIS sequences. In all four cases, PCR products of the predicted lengths (815 to 1,075 bp) were obtained, and their sequences were identical to those of the cognate pREIS1, -2, -3, and -4 sequences (data not shown), consistent with a REIS origin of the plasmids, rather than other bacteria that may have been present in tissues used to prepare I. scapularis DNA.
Plasmids occur in North and South American “Ca. Rickettsia amblyommii” but not in R. parkeri.
Analysis of the North American SFG member “Ca. Rickettsia amblyommii” Darkwater revealed plasmid isomers that migrated at approximately 47 and 62 kbp (Fig. (Fig.2A).2A). In contrast, plasmids were absent in lanes containing R. parkeri Ft. Story, Portsmouth, and Tates Hell (Fig. (Fig.2A)2A) and for three additional North American isolates, High Bluff, Grand Bay, and Oktibbeha (data not shown). The pRM16 probe hybridized to the “Ca. Rickettsia amblyommii” plasmids but did not hybridize to R. parkeri DNA (Fig. (Fig.2B).2B). A replicate gel hybridized with the pRM6 probe (Fig. (Fig.2C)2C) reproduced the “Ca. Rickettsia amblyommii” two-band hybridization pattern of the pRM16 probe and identified a third band that probably corresponded to a plasmid that was poorly resolved from the 25-kbp mitochondrial DNA band shown in Fig. Fig.1A.1A. The pRM6 probe did not hybridize to DNA of R. parkeri. A second replicate gel hybridized with the pRM8 probe (Fig. (Fig.2D)2D) reproduced the “Ca. Rickettsia amblyommii” pRM6 hybridization pattern but did not yield a signal in the R. parkeri lanes.
Analysis of the South American “Ca. Rickettsia amblyommii” An13 and Ac37 isolates revealed plasmid isomers that migrated at approximately 17 and 50 kbp and approximately 17, 33, and 55 kbp, respectively (Fig. (Fig.3A).3A). The South American R. parkeri isolates At#24 and At#5 did not have plasmid DNA bands. The pRM16 probe hybridized to plasmid isomers of “Ca. Rickettsia amblyommii” An13 in a three-band pattern that included an additional 33-kbp band not revealed by SYBR green staining but recognized only the 17- and 33-kbp plasmid isomers of “Ca. Rickettsia amblyommii” Ac37 (Fig. (Fig.3B).3B). The probe did not hybridize to R. parkeri DNA. Replicate gels hybridized with the pRM6 probe (Fig. (Fig.3C)3C) and the pRM8 probe (Fig. (Fig.3D)3D) produced similar hybridization patterns, except that the 17-kbp plasmid isomer of “Ca. Rickettsia amblyommii” An13 was not recognized. The results were consistent with the presence of pRM6, -8, and -16 homologs in the “Ca. Rickettsia amblyommii” isolates and their absence in the R. parkeri isolates.
FIG. 3.
FIG. 3.
Presence and absence of plasmids in South American isolates of “Ca. Rickettsia amblyommii” and R. parkeri, respectively. (A) PFGE of DNAs from “Ca. Rickettsia amblyommii” isolates An13 and Ac37 and R. parkeri isolates At#24 (more ...)
In summary, the results demonstrated the presence of plasmids in North and South American “Ca. Rickettsia amblyommii” isolates, which carried pRM6, -8, and -16 homologs. In contrast, eight R. parkeri isolates from both continents did not carry plasmids, and none of the probes hybridized to their sheared chromosomal or low-molecular-weight DNA, consistent with the absence of plasmid gene homologs on the rickettsial chromosomes. The R. parkeri isolates were the only low-passage rickettsia isolates that we analyzed that did not possess a plasmid. Previously analyzed high-passage R. montanensis also lacked a plasmid, while low-passage R. peacockii possessed a plasmid that was lost during serial passage (7).
Plasmids occur in low-passage R. bellii but not in a high-passage isolate.
Analysis of South American isolates of R. bellii, a member of the AG rickettsiae, revealed prominent plasmids that migrated at approximately 55 to 65 kbp in lanes containing isolates ovale, Ad25, An4, and Mogi and at 75 and 90 kbp in the lane containing isolate HJ#7 (Fig. (Fig.4A).4A). The upper mitochondrial DNA band migrated as a doublet in the 0.9% agarose gels. Plasmid bands were absent in the lane loaded with R. bellii 369-C, which was originally isolated in 1966 from Dermacentor variabilis ticks collected in Arkansas. Replicate gels hybridized with the pRM16 probe (Fig. (Fig.4B)4B) and the pRM6 probe (Fig. (Fig.4C)4C) revealed plasmids in the two- and three-band patterns seen in Fig. Fig.4A,4A, with less-abundant smaller isomers. However, none of the patterns were identical, and the plasmid of the ovale isolate did not hybridize with the pRM6 probe. The pRM8 probe hybridized weakly or not all (isolate An4) to the plasmid bands but strongly to sheared chromosomal DNA, including that of isolate 369-C (Fig. (Fig.4D).4D). Because R. bellii occupies a basal position in rickettsial phylogenetic trees, we hybridized a replicate gel blot with a probe derived from the pRM12 gene, encoding a proline/betaine transporter. It was previously shown to recognize the plasmid of “Ca. Rickettsia hoogstraalii” isolated from a North American argasid tick but not the plasmids of Rickettsia spp. from other, ixodid ticks or R. felis from cat fleas (7). The pRM12 probe hybridized to plasmids of three of the five R. bellii isolates and to chromosomal DNA of all five (Fig. (Fig.4E).4E). In summary, the results demonstrated the absence of a plasmid in the high-passage 369-C isolate, in contrast to the universal presence of relatively large plasmids in the low-passage R. bellii isolates. The plasmids varied in size, and those of the An4 and ovale isolates were less conserved than the others relative to pRM of R. monacensis.
FIG. 4.
FIG. 4.
Presence of plasmids in South American isolates of R. bellii. (A) PFGE of DNAs from R. bellii isolates ovale, Ad25, An4, Mogi, HJ#7 (from South America), and 369-C (from North America). (B) Southern blot of the same gel hybridized with the pRM16 gene (more ...)
To further assess the conservation of the rickettsial plasmid gene complement, we hybridized PFGE Southern blots for rickettsiae reported in this study with a probe derived from the R. monacensis pRM23 gene, encoding a transposon resolvase similar to that of Burkholderia thailandensis and probably derived by HGT (6). The probe hybridized to plasmid isomers of R. akari Bronx, R. rhipicephali HJ#5, and REIS (Fig. (Fig.5B)5B) but not to plasmids of “Ca. Rickettsia amblyommii” or R. bellii (data not shown). The results of the Southern blot analyses shown in Fig. Fig.11 to to55 are summarized in Table Table33.
FIG. 5.
FIG. 5.
A probe derived from the R. monacensis pRM23 gene, encoding a homolog of a Burkholderia thailandensis transposon resolvase, hybridizes to plasmids of three Rickettsia spp. (A) PFGE of DNAs from R. monacensis IrR/Munich, R. akari Bronx, REIS, and R. rhipicephali (more ...)
TABLE 3.
TABLE 3.
Southern blot detection of pRM gene homologs on plasmids of New World Rickettsia isolates
Plasmid copy numbers.
Rickettsial plasmid copy numbers were estimated by QPCR as the relative ratios of plasmid-carried hsp2 gene homologs and the single-copy gltA genes found on rickettsial chromosomes. Plasmid copy numbers among seven Rickettsia spp. fell within a fourfold range, from 2.4 in R. helvetica to 9.2 in R. peacockii (Table (Table4).4). Means and standard deviations of standard curve DNA and rickettsial sample DNA hsp2 reaction efficiencies were 96.8 ± 2.6 and 94.1 ± 2.9, respectively, while the cognate values for gltA reactions were 99.5 ± 1.5 and 95.9 ± 2.9, respectively. In all cases, reaction products had well-defined single-peak melting curves.
TABLE 4.
TABLE 4.
Plasmid copy numbers in selected Rickettsia spp.
Ca. Rickettsia amblyommii” AaR/SC and Ac/Pa isolates each have multiple distinct plasmids.
In this (Fig. (Fig.1)1) and a previous study (7), PFGE, Southern blot, PCR, and I. scapularis genome sequence data suggested the existence of two or more distinct plasmid species in “Ca. Rickettsia amblyommii,” R. rhipicephali, and REIS. We verified that hypothesis by isolation, cloning, and sequencing of plasmids from genomic libraries of “Ca. Rickettsia amblyommii” AaR/SC and Ac/Pa. Screening of the libraries by hybridization with a cocktail of probes derived from pRM genes yielded positive clones at a frequency of <1%. Eight AaR/SC clones and seven Ac/Pa clones were selected for further study. Southern blot analysis revealed plasmid DNA inserts of 6.5 to 24 kbp (data not shown). Four “Ca. Rickettsia amblyommii” AaR/SC clones were sequenced to confirm the existence of at least two plasmids in that isolate. The entire 18,497-bp pRAM18 plasmid was contained in a single Swa1 clone (verified by PCR amplification using terminal primers), while the 22,781-bp pRAM23 sequence was obtained from a 21.9-kbp HpaI clone and a 0.9-kbp PCR product amplified with primers complementary to end sequences of the HpaI clone. An as yet incompletely circularized sequence represents a provisional third plasmid (pRAM30), estimated to be 30 kbp in length. The completed pRAM18 and pRAM23 sequences encode homologs of the DnaA-like replication initiator and ParA plasmid partitioning proteins encoded by the pRM16 and pRM18 genes. Homologs of the pRM6 and pRM7 genes, encoding Hsp1 and Hsp2 proteins, were present on pRAM23 but not on pRAM18, which was nearly identical to a slightly larger Ac/Pa plasmid contained in a single clone. Incompletely sequenced Ac/Pa clones indicated the presence of at least one more plasmid in that isolate. Other notable aspects of the AaR/SC plasmids included the presence on pRAM23 of a homolog of the R. bellii and Orientia tsutsugamushi phrB genes, which encode DNA UV damage repair enzymes, as well as two sca12 gene homologs encoding outer membrane proteins with potential roles in host cell interactions. A predicted gene on pRAM18 encoded a chimeric protein containing both SpoT and leucine-rich repeat domains, thus combining potential stringent response and protein interaction functions.
The pRAM18 gene complement, but not that of pRAM23, was highly similar to that of pRMA from R. massiliae and included a homolog of the pRMA p05 site-specific recombinase gene. In a PFGE/Southern blot analysis, a probe derived from the pRAM18 p05 homolog hybridized to a subset of the “Ca. Rickettsia amblyommii” AaR/SC plasmid isomers that were not recognized by the pRM6 probe (data not shown), further confirming the existence of multiple plasmid species in a single rickettsia isolate.
Phylogenetic analysis of rickettsial ParA proteins.
Maximum parsimony analysis of 154 amino acids (145 informative amino acids) of the rickettsial ParA proteins showed that those encoded on rickettsial chromosomes clustered tightly within the tree (Fig. (Fig.6,6, area at left center), as a group that diverged from a node on a branch bearing a ParA protein from Ehrlichia chaffeensis, also a member of the Rickettsiaceae. Of all the rickettsial plasmid-encoded type Ib ParA proteins, only that encoded by pREIS4 from the rickettsial endosymbiont of I. scapularis was present on the same branch with the chromosomal ParA proteins, diverging from the most basal node on the branch. The rickettsial plasmid ParA proteins were highly diverse relative to the chromosomal ParA proteins, as illustrated by comparison with the tree positions of plasmid-encoded ParA proteins from other families of bacteria. The majority of the rickettsial plasmid ParA sequences clustered in three groups, with good bootstrap support. The first group (at right) was at a distal position on a branch with a basal node that led off to pRAF-encoded ParA of R. africae and a medial node that led off to plasmid-encoded ParA sequences from Yersinia and Salmonella spp. The distal group consisted of highly similar ParA homologs, encoded by pRAM23 of “Ca. Rickettsia amblyommii” and pRMA of R. massiliae, as well as a less similar homolog encoded on a contig reported from an incomplete genome sequence assembly of the aquatic eukaryote Trichoplax adhaerens (68), the simplest known multicellular animal. We detected many homologs of Rickettsia genes on the Trichoplax contigs and predict that previously observed bacterial endosymbionts in Trichoplax (62) will prove to be rickettsiae. The second rickettsial plasmid ParA group (bottom right) consisted of homologs encoded by pRPR of R. peacockii and pRF of R. felis, which clustered at the distal ends of a branch containing an adjacent node that led to a ParA protein encoded by a plasmid of Pseudomonas syringae. A more basal node on the branch led to ParA proteins from pREIS1 and plasmids of Yersinia and Pseudomonas spp., while the most basal node led to the ParA protein encoded by pRM of R. monacensis. The third group (upper left) consisted of ParA proteins encoded by pRFδ of R. felis, pREIS2, and pREIS3. The ParA proteins encoded by pRAM18 and the provisional pRAM30 plasmid diverged from nodes without bootstrap support on a branch (top center) that also contained a ParA protein encoded by the Borrelia burgdorferi cp32 plasmid. Neighbor-joining analysis supported the maximum parsimony results (not shown).
FIG. 6.
FIG. 6.
Maximum parsimony phylogenetic tree (unrooted) of rickettsial ParA proteins. The scale bar indicates the number of amino acid changes within branch lengths. Bootstrap scores are indicated by numerals. GenBank accession numbers for the sequences used are (more ...)
Phylogenetic analysis of rickettsial small heat shock proteins.
Maximum parsimony analysis of the rickettsial plasmid-encoded Hsp2 proteins (113 amino acids [100 informative amino acids]) showed that they clustered tightly as a single group (Fig. (Fig.7,7, circled at bottom right). The most basal group member was encoded by the R. felis chromosomal RF1004 gene. Sequence analysis indicated that it was a chimera that may have originally been a plasmid gene that was integrated into the chromosome by a recombination event adjacent to RF1005, which suffered a 5′-end deletion of its ORF (data not shown). The chromosome-encoded Hsp proteins of three facultative intracellular pathogens from other bacterial taxa and of Wolbachia pipientis (Rickettsiales) occupied intermediate positions in the tree. The Rickettsia chromosome-encoded Hsp1 proteins clustered tightly as a group that branched away from the Legionella pneumophila node. The “R. rickettsii complex” in that group included the SFG members R. rickettsii, R. africae, R. conorii, R. massiliae, R. peacockii, and R. sibirica. The plasmid-encoded Hsp1 proteins of R. felis, R. monacensis, R. peacockii, and “Ca. Rickettsia amblyommii” pRAM23 (pRAM18 has no hsp genes) formed a second group (circled at top) that clustered as a side branch to the chromosomal Hsp1 cluster. Neighbor-joining analysis produced the same tree topology (not shown).
FIG. 7.
FIG. 7.
Maximum parsimony phylogenetic tree (unrooted) of rickettsial small heat shock proteins. The scale bar indicates the number of amino acid changes within branch lengths. Bootstrap scores are indicated by numerals. GenBank accession numbers for the sequences (more ...)
With this and previous PFGE/Southern blot analyses (6, 7), we have demonstrated the presence of plasmids in low-passage isolates of AG, TRG, and SFG Rickettsia spp. obtained from arthropods or clinical samples collected from humans in North and South America and Europe. Similar isolates of R. africae from Africa have plasmids (24), and it now seems likely that plasmids occur in rickettsiae associated with blood-feeding arthropods throughout the world. However, plasmids were not present in R. parkeri, newly recognized as a mild SFG pathogen (49, 50, 52), or in the GenBank-deposited genome sequences of major pathogens in the SFG (R. conorii and R. rickettsii) and TG (R. prowazekii and R. typhi). Plasmids were detected in five low-passage isolates of R. bellii but not in R. bellii 369-C (Fig. (Fig.4)4) or R. montanensis M5/6 (7), both of which have undergone serial passage since their isolations in the 1960s (9, 56; E. J. Bell, unpublished data). In this context, it would be reasonable to reevaluate the major rickettsial pathogens (i.e., R. conorii, R. prowazekii, R. rickettsii, and R. typhi) as low-passage isolates for the presence of plasmids by using PFGE/Southern blot assays.
We obtained the first estimates of rickettsial plasmid copy numbers. Single-copy hsp2 genes occur on the sequenced pRF, pRM, pRAM23, and pRPR plasmids. The unsequenced plasmids of R. bellii, R. helvetica, and “Ca. Rickettsia hoogstraalii” have hsp2 homologs (Fig. (Fig.4C)4C) (7), but their existence as single copies was an assumption. Rickettsial chromosomes typically carry single-copy gltA genes, but whether this is true for the unsequenced species “Ca. Rickettsia amblyommii,” R. helvetica, and “Ca. Rickettsia hoogstraalii” awaits confirmation. With the exception of R. felis, hsp2 homologs are not present in known rickettsial chromosome sequences. Given these caveats, plasmid copy numbers in seven Rickettsia spp. representing the AG, TRG, and SFG averaged 4.4 per chromosomal equivalent (Table (Table4).4). The rickettsiae thus have low-copy-number plasmids (i.e., fewer than 10 copies per chromosome), consistent with their possession of par genes, which are essential for the maintenance and stable inheritance of such plasmids (25, 27, 74). Plasmids are absent in members of the other genera of the Rickettsiales and are rare in other obligate intracellular bacteria associated with arthropods. Among such bacteria, the plasmids of the Buchnera endosymbionts of aphids (Hemiptera: Aphididae) are the best known. Ratios of the leuABCD and trpEG genes, carried on separate plasmids, to single-copy chromosomal genes among Buchnera spp. associated with three aphid hosts ranged from 0.6 to 23.5 (35, 58, 73). However, copy number interpretations of those values may be complicated by trpEG gene amplification and fluctuations in Buchnera chromosome ploidy during host lifetimes (73).
Apparent homologs of the R. monacensis pRM16 and pRM6 genes, encoding plasmid maintenance and probable host-adaptive functions, respectively, were well conserved among the plasmids of 21 Rickettsia isolates (Table (Table3)3) (7), but there were interesting divergences in plasmid gene conservation versus host association and phylogeny. Among SFG members, the plasmid gene complement of “Ca. Rickettsia hoogstraalii,” isolated from a North American argasid tick, was much better conserved relative to that of R. monacensis than was that of R. helvetica, both of which were isolated from the same European ixodid tick. Apparent homologs of the pRM23 gene, encoding a transposon resolvase, were present on plasmids of the mite-borne species R. akari of the TRG as well as on those of R. rhipicephali and REIS, SFG members that were isolated from South and North American ticks of different genera (Fig. (Fig.5).5). In contrast, pRM23 homologs were not present on plasmids of other SFG rickettsiae or those of R. felis (TRG) and R. bellii (AG). The plasmid gene complements were not wholly consistent either with rickettsial phylogeny and host associations or with descent from a single ancestral plasmid, suggesting possible multiple origins and/or the influence of HGT.
We obtained evidence for multiple origins of the plasmids through phylogenetic analysis. Plasmids carry conserved partitioning genes (par) that are usually organized in an autoregulated operon and are required for plasmid segregation at cell division (25, 74). The encoded ParA proteins are Walker-type ATPases whose ATP-bound forms interact with a nucleoprotein complex consisting of ParB protein dimers bound to sequence repeats at the parS centromere to mediate intracellular location, movement, segregation, and incompatibility of plasmids (14, 25). The ParB protein sequences are highly conserved, but the more variable ParA sequences allow phylogenetic analysis of plasmid lineages (27). Maximum parsimony analysis showed that most rickettsial plasmid-encoded ParA proteins fell into three groups that clustered with ParA proteins encoded on plasmids from other bacterial genera rather than with the highly conserved ParA sequences encoded on rickettsial chromosomes (Fig. (Fig.6).6). In conjunction with the evidence for multiple plasmids in single Rickettsia spp. discussed below, the results provided strong evidence for the presence of plasmids from multiple incompatibility groups in the genus Rickettsia. Similar phylogenetic analyses of plasmid maintenance proteins encoded by repABC operons of the repABC plasmids have demonstrated the presence of multiple plasmid incompatibility groups within several genera of alphaproteobacteria, including as many as nine in the Roseobacter clade that can stably coexist in the same cell, while six occur in each of two Rhizobium spp. (17, 55). Similar to the rickettsial ParA phylogeny, different repABC replicons within the same bacterial strain tend to belong to different phylogenetic clades with lineages that are not congruent with species trees, suggesting that incompatibility groups arise as a consequence of divergent evolution that may be interrupted by HGT events between plasmid lineages (16).
Differential hybridization patterns of the pRM6 and pRM16 probes to plasmid isomers of both R. rhipicephali and REIS (Fig. (Fig.1)1) and to those of several “Ca. Rickettsia amblyommii” isolates (Fig. (Fig.2)2) (7) were reminiscent of the simultaneous presence of the 63-kbp pRF plasmid and the 39-kbp pRFδ deletion form in R. felis (45). Although the pREIS1,- 2, -3, and -4 plasmid sequence scaffolds from the I. scapularis genome sequence project have various degrees of similarity, they possess different parA genes, and none clearly represents a deletion form of another. We obtained PCR evidence for their legitimate identity as REIS plasmids and obtained physical confirmation that multiple plasmids exist in single rickettsia isolates by cloning and sequencing pRAM18 and pRAM23 from “Ca. Rickettsia amblyommii” AaR/SC. Their sequence similarities were confined to genes that encoded DnaA-like replication initiators and Par proteins, and they were therefore distinct plasmids rather than a major plasmid accompanied by a deletion form. We are sequencing a third plasmid (pRAM30) from the AaR/SC isolate and two from the “Ca. Rickettsia amblyommii” Ac/Pa isolate. Because those isolates represent uncloned bacterial populations, it is not yet clear whether individual cells contain only single or multiple plasmid species.
The presence of multiple plasmid species in single “Ca. Rickettsia amblyommii” isolates is intriguing given the biology of their primary hosts. Amblyomma ticks are widely distributed from tropical to temperate climates and are known for their aggressive propensity to feed on a wide range of hosts that are parasitized by other ticks and blood-feeding arthropods (18, 29, 78). Those attributes are well suited to facilitating HGT within the “intracellular arena” of bacterial genetic exchange in potentially coinfected arthropods (12, 13). A surprisingly diverse range of bacteria occur in ticks (40), and coinfections of single ticks with different obligate intracellular microbes have been demonstrated for several genera, including Amblyomma (19); in addition, as many as three Rickettsia spp. have been found in single ticks (15). The wide distribution and host biology of Amblyomma ticks and the presence of multiple plasmids that may be of different lineages in single “Ca. Rickettsia amblyommii” strains are consistent with the possibility that Amblyomma ticks have been an active locus of HGT into and within the genus Rickettsia.
The high content of transposon-related sequences characteristic of all sequenced rickettsial plasmids suggests that they may be HGT “hot spots” within rickettsial genomes, perhaps as a consequence of their exposed positions as cytoplasmic episomes relative to the packaged chromosomes associated with the bacterial cell walls. As mobile genetic elements, plasmids are crucial drivers of HGT that enhance bacterial diversity and often provide the host bacterium with functions, such as drug resistance and environmental adaptive capacity, that play roles in pathogenicity (30, 43, 60, 70, 75). Plasmids may play those roles in rickettsiae by acting as a mechanism for gain of new genes in the otherwise reductive Rickettsia genomes. The rickettsial α-crystalline hsp genes, which are prime candidates for provision of host-adaptive functions (7), provide a likely example. Phylogenetic analysis indicated that the hsp2 genes of the Rickettsia species were plasmid specific, with the exception of a chromosomal homolog in R. felis, and were probably acquired independently of the hsp1 genes (Fig. (Fig.7).7). The plasmid-encoded Hsp1 cluster consisted of proteins from R. felis (TRG) and three SFG spp. (R. monacensis, R. peacockii, and “Ca. Rickettsia amblyommii”) but branched from a deep node within the chromosomal Hsp1 group that lies between R. bellii (AG) and the TRG and TG rickettsiae. That result was consistent with a potential evolutionary origin of the plasmid hsp1 genes in a chromosome-to-plasmid transfer event in a TRG-like Rickettsia species. This possibility is supported by phylogenetic analyses showing that the R. monacensis OmpA and OmpB proteins have much greater similarity to homologs from TRG rickettsiae than to those from SFG rickettsiae (38), consistent with a much closer affinity between R. monacensis and the TRG members than has previously been realized. Additional support derives from the presence of a 12-kbp plasmid-like sequence in the R. typhi chromosome (28, 39) and our observation that the R. felis RF1004 gene may have undergone a plasmid-to-chromosome transfer.
We have now demonstrated that plasmids occur in nearly all arthropod-borne AG, TRG, and SFG rickettsiae that we have surveyed for their presence. The pRF plasmid of R. felis was the first to be discovered in the genus Rickettsia and was suggested to be conjugative on the basis of encoding conjugative transfer gene products and the presence of pili on the surfaces of R. felis cells (45). It is now known that conjugative genes are widespread in the genus and that they are horizontally transmitted (77). The presence of widespread and potentially mobile plasmids in Rickettsia spp. has evolutionary and epidemiologic implications. The true impact of those implications requires further investigation of the full extent of the distribution of rickettsial plasmids within the many members of the genus not found in arthropod vectors and whether the plasmids are currently mobile.
Acknowledgments
This research was supported by NIH grant RO1 AI49424 to U.G.M.
Footnotes
[down-pointing small open triangle]Published ahead of print on 22 January 2010.
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