- Email: [email protected]
Candidatus Rickettsia hoogstraalii in Ethiopian Argas persicus ticks
Ticks and Tick-borne Diseases 3 (2012) 337–344
Contents lists available at SciVerse ScienceDirect
Ticks and Tick-borne Diseases journal homepage: www.elsevier.com/locate/ttbdis
Original article
Candidatus Rickettsia hoogstraalii in Ethiopian Argas persicus ticks Vera Pader a , Joanna Nikitorowicz Buniak a , Alemseged Abdissa b , Haileeysus Adamu b , Tadele Tolosa c , Abebaw Gashaw c , Ronald R. Cutler d , Sally J. Cutler a,∗ a
School of Health, Sports & Bioscience, University of East London, London, UK College of Public Health and Medical Sciences, Jimma University, Jimma, Ethiopia College of Agriculture and Veterinary Medicine, Jimma University, Jimma, Ethiopia d School of Biological & Chemical Sciences, Queen Mary University of London, London, UK b c
a r t i c l e
i n f o
Keywords: Argas persicus Ornithodoros moubata Amblyomma variegatum Rickettsia Candidatus Rickettsia hoogstraalii Rickettsia felis
a b s t r a c t Ethiopian soft ticks Argas persicus, hard ticks including both Amblyomma variegatum and Rhipicephalus (Boophilus) spp., and fleas were collected from livestock, traditional human dwellings, and cracks and crevices of trees. They were assessed in pools for the presence of Rickettsia using PCR-based methods. The extracted tick DNA was subjected to molecular screening for Rickettsia, which revealed 50.5% of the pooled samples to be positive for Rickettsia spp. These were then subjected to multi-gene analysis using both outer surface proteins and housekeeping genes with proven discriminatory potential. Sequencing of the citrate synthase and outer membrane genes clearly led to the identification of three distinct rickettsial species, Candidatus Rickettsia hoogstraalii in Argas persicus ticks; R. africae in hard tick pools, and R. felis in fleas. Furthermore, we demonstrated the presence of the plasmid-borne small heat-shock protein gene hsp2 in DNA from A. persicus ticks suggesting that Candidatus R. hoogstraalii carried by these ticks possess a plasmid. Unlike chromosomal gene sequences, the hsp2 gene failed to cluster with Candidatus R. hoogstraalii, instead falling into an isolated separate clade, suggesting a different origin for the plasmid. © 2012 Elsevier GmbH. All rights reserved.
Introduction Rickettsiae are transmitted by the bite of various haematophagous arthropods such as ticks, fleas, lice, and mites. With the recent application of molecular methods to arthropods from various locations, many new rickettsial species have been recognised, including several newly recognised pathogenic species. This prompted us to revisit the Rickettsia present in Ethiopian ticks. The presence of R. africae is already established among Amblyomma variegatum and other species of ticks in Ethiopia, for example, R. africae strain ESF-5 originated from the Shulu Province of Ethiopia (Roux et al., 1997) and is found extensively throughout other African countries (Fournier et al., 1998; Mediannikov et al., 2010b; Morita et al., 2004; Mura et al., 2008; Portillo et al., 2007). Similarly, several recent reports have documented the presence of R. felis among fleas from various African nations (Parola, 2011). Other rickettsial species from Africa include R. aeschlimannii, R. conorii, R.
∗ Corresponding author at: School of Health and Bioscience, University of East London, Water Lane, Stratford, London E15 4LZ, UK. Tel.: +44 0208 223 6386; fax: +44 0208 223 4959. E-mail address: [email protected] (S.J. Cutler). 1877-959X/$ – see front matter © 2012 Elsevier GmbH. All rights reserved. http://dx.doi.org/10.1016/j.ttbdis.2012.10.021
massiliae, and R. sibirica mongolitimonae of the spotted fever group and R. prowazekii and R. typhi of the typhus group (Letaief, 2006; Morita et al., 2004; Parola, 2006; Pretorius and Birtles, 2002). Interestingly, it has also been hypothesised that a tick reservoir for R. prowazekii might also exist in Ethiopia (Burgdorfer et al., 1973). Despite this great rickettsial diversity, cases are seldom reported among the indigenous population. Instead, these are more often observed among the ever-increasing numbers of tourists that now visit these regions (Jensenius et al., 2009; Pretorius and Birtles, 2002). Sporadic infections and even clusters of infection have been reported, often diagnosed once these travellers have returned home with fever (Stephany et al., 2009). The burden of rickettsial infection among African populations is only just starting to be recognised, with limited studies investigating the role of Rickettsia in febrile patients attending clinics and hospitals (Mediannikov et al., 2010a; Parola, 2011; Socolovschi et al., 2010). A limiting factor is the lack of molecular diagnostics available within healthcare settings and the relatively poor diagnostic performance of serology in highly endemic situations. Given the greater precision offered by molecular approaches for identification of rickettsiae, we undertook an investigation of various tick species from two regions in Ethiopia to explore the diversity of rickettsiae present.
338
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
Materials and methods
Confirmatory PCR
Ticks collected from Ethiopia
Arthropod pools found to be positive upon screening were further tested using conventional PCR using a MyCycler thermocycler (BioRad) for multi-gene targets previously described including gltA, ompA, dnaK, dnaA, atpA, recA, and the internal rrl-rrf ribosomal spacer (ITS) (Vitorino et al., 2007). Additionally, primers described for amplification of fragments of outer surface proteins A and B were used as previously described ompA (designed using NCBI primer design software, this study) and for partial sequence amplification of the 5 end of ompB (Roux and Raoult, 2000). Larger amplicons for sequencing citrate synthase used the primers and methods described by Roux et al. (1997). To explore the possibility of plasmids among any Rickettsia detected, we used the primers described to characterise plasmids recently described among R. felis (Fournier et al., 2008). Additionally, a small heat shock protein Hsp2 has been reported to reside on rickettsial plasmids, and primers against this target were also used to characterise positive samples according to published protocols (Baldridge et al., 2008). These primers are detailed in Table 1. PCR products were observed using the SybrSafe (Invitrogen) DNA stain in 2% (w/v) agarose gels. The amplicon sizes were measured relative to a 100-bp molecular mass standard (Fermentas).
A total of 98 pools of up to 20 ticks was collected from two different geographical regions of Ethiopia during July to August 2010. The collection areas were Jimma to the southwest and Dire Dawa to the east of Addis Ababa, respectively. Ticks were pooled when collected from the same host, thus some contained multiple species, or when the numbers exceeded 20 from the same environmental location, these were separated into different pools. No pool contained more than 20 ticks. Hard ticks were removed from livestock, whilst soft ticks were collected from cracks and crevices of human dwellings and from under the bark of trees in the vicinity of livestock areas. Limited collections of fleas (Ctenocephalides spp.) were made in addition to the ticks (3 pools: 2 containing 3 fleas; and a single flea in the third), with these being collected from clothing or bedding in traditional human dwellings. All collected arthropod pools were immersed in approximately 0.5 ml of 8 M guanidine hydrochloride and imported to the UK for analysis under licence. DNA extraction Tick and flea pools were transferred into a fresh tube together with glass beads and a volume of sterile PBS equivalent to the size of the arthropods within each pool. These were homogenised using a bead beater prior to incubation with tissue lysis buffer and proteinase K at 56 ◦ C for 1–4 h until liquefied. Debris was pulsed down with a 1-min spin in a microfuge and the supernatant removed for DNA extraction using DNeasy kits (Qiagen) on a QiaCube robot (Qiagen) using the rodent tail protocol. PCR screening All samples were screened using a real-time generic rickettsial PCR (Stenos et al., 2005). Flea samples were additionally screened using real-time assays specific for R. felis biotin synthase (Socolovschi et al., 2010). “Blank” samples containing sterile PBS that were processed alongside the tick pools, and sterile DNA-free water served as negative controls. All real-time assays were performed as originally described, using Stratagene Mx3000p cycler (Agilent). All primers and probes are listed in Table 1.
Sequence analysis Where amplicons of suitable quality were produced, these were sent to the Genome Centre Queen Mary University of London for sequencing, using an Applied Biosystems ABI 3730xl. The chromatograms were analysed and the sequence data was edited, aligned and trimmed using ClustalW and Mega5. The latter was used for phylogenetic tree construction using a bootstrap value of 1000 replicates (Tamura et al., 2011). Results Arthropod collections Small numbers of hard ticks were also collected by removal from domestic livestock and companion animals. A total of 21 pooled DNA extracts from hard ticks [Amblyomma variegatum and Rhipicephalus (Boophilus) spp.] were collected from animals in both Dire
Table 1 Primers and probes used during investigation. Gene target
Forward primer 5 –3
Reverse primer 5 –3
Probe (where applicable)
Reference
Citrate synthase (gltA)
TCGCAAATGTTCACGGTACTTT
CACAATGGAAAGAAATGCACGA
Stenos et al. (2005)
R. felis biotin synthase
ATGTTCGGGCTTCCGGTATG
CCGATTCAGCAGGTTCTTCAA
FAM-TGCAATAGCAAGAACCGTAGGCTGGATG-BHQ1 6-FAMGCTGCGGCGGTATTTTAGGAATGGG-TAMRA
Citrate synthase Rp877-CS1273 Outer membrane protein (ompA) Outer membrane protein (ompB) F1 Outer membrane protein (ompB) F2 ATP synthase F1 subunit alpha atpA Chromosomal replication initiation protein dnaA Heat shock protein 70 dnaK R. felis plasmid pRFa-b R. felis plasmid pRFc-d hsp2 Hsp2F3-Hsp2R3
GGGGACCTGCTCACGGCGG CGGTGTTGTAGGGACTGCGGC CCGCAGGGTTGGTAACTGC
CATAACGAGTGTAAAGCTG CGGTCCGCCCGGTTTGAAGT CCTTTTAGATTACCGCCTAA
Roux et al. (1997) This study Roux and Raoult (2000)
AATATCGGTGACGGTCAAGG
CCTATATCGCCGGTAATT
Roux and Raoult (2000)
AGTACAGACATATCGAGATGA CTTTACAATCATTACGGTG
CGACTTACCGAAATACCGAC GCAACTAAGCCCCATCC
Vitorino et al. (2007) Vitorino et al. (2007)
GCATTCTAGTCATACCGCC CAAGCTTTTGTACTGCCTCTAT ACATTCCGTAAAGAATATGAGC CTTAGCCTTACTTTGTTCTTTTTTAGG
Vitorino et al. (2007) Fournier et al. (2008) Fournier et al. (2008) Baldridge et al. (2008)
hsp2 Rhoogs qHsp2F1- Rhoogs qHsp2R hsp2 Hsp2F3-Hsp2R
GATGGTAAACTAATGGATCGC
CAAAAAATGAAAGAAACTGCTGA AGTGCATATAGCTACCACACTATCT GCTTATGTTCGCCTTTAGTATTTA CTTTTAATTTATCACGTATTAGAAATAAATCAG TGAGGGCAAAAATGAACAATC
CTTAGCCTTACTTTGTTCTTTTTTAGG
AGGCAAAGGCGAGAGAAATACC
Baldridge et al. (2008)
Socolovschi et al. (2010)
Baldridge et al. (2008)
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
339
Fig. 1. Plate A shows an Amblyomma variegatum tick forming part of a hard tick pool. Plate B shows a representative Argas persicus tick forming part of a soft tick pool. Plate C illustrates the tick density under the bark of trees; whilst Plate D shows ticks within a wooden beam removed from a traditional style Ethiopian dwelling.
Dawa and Jimma regions (see Fig. 1a). Soft ticks identified as Argas persicus (pers. communication Alan Walker, University of Edinburgh, UK) were collected from a village (Asselisso) on the outskirts of Dire Dawa (see Fig. 1b). This village maintained a huge tick burden with ticks not only surrounding livestock areas predominantly under the bark of trees (see Fig. 1c), but also residing within human traditionally built dwellings (see Fig. 1d). We were able to collect 77 pools containing 5–20 ticks from this village. No soft ticks were collected from other sites. Three pools of fleas were collected from human clothing/bedding. Screening of samples Screening of pooled samples revealed 5 of the 21 (24%) hard tick pools and 44 of the 77 (57%) soft tick pools to be positive, whilst the R. felis-specific biotin synthase assay yielded positive results from 2 of the 3 flea pools only (both containing 3 fleas). Identification of Rickettsia detected Conventional PCR to produce fragments for sequence analysis from DNA-extracted tick pools failed to yield full overlapping contigs, thus precluding concatenation for complete gene analysis. Partial gene sequences were generated for citrate synthase (gltA) and outer membrane proteins OmpA and OmpB that have been extensively used for rickettsial identification. Figs. 2–4 show the phylogenetic comparison of our sequences with homologous sequences from validated species and their nearest neighbours
available from GenBank. These phylogenetic trees revealed the presence of R. africae in a hard tick pool (containing Am. variegatum ticks) and R. felis in the positive flea pools (R. felis gltA partial gene sequence deposited under GenBank accession number JN366415; ompB partial gene sequence under accession number JN366420). Rickettsial DNA detected in A. persicus had greatest homology with Candidatus R. hoogstraalii previously reported from other genera of soft ticks collected from Africa, Japan, and the USA (Cutler et al., 2006; Kawabata et al., 2006; Mattila et al., 2007) and among European Haemaphysalis sulcata ticks from Croatia (Duh et al., 2010). This showed 99.5–100% sequence similarity between Candidatus R. hoogstraalii and the amplicons from Ar. persicus over the 428 bp analysed. Representative partial gene sequences for gltA, ompA, and ompB are deposited under GenBank accession numbers JN366413–JN366414 (gltA), JN366416 (ompA), and JN366418–JN366419 (ompB). For comparative purposes, an earlier sample from Tanzanian Ornithodoros moubata soft ticks was submitted for partial ompA and ompB gene sequencing (GenBank accession numbers JN366417 and JN366421, respectively) (Cutler et al., 2006). Sequence results from other “housekeeping” gene targets including dnaA (deposited under GenBank accession JN366410), dnaK (deposited under GenBank accession numbers JN366411 and JN366412), and atpA (deposited under GenBank accession number JN366409) showed poor discriminatory power among rickettsial species (data not shown). The remaining 4 gene targets described for multi-gene identification failed to produce amplicons from positive tick DNA extracts (Vitorino et al., 2007).
340
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
Fig. 2. Neighbour-joining tree citrate synthase (gltA) (bootstrap 1000 replicates, values > 60 shown) over 428 bp. All samples with an Et prefix are samples collected during this study. Sample Et161b from hard ticks clusters with Rickettsia africae whilst sample Et90 from a flea pool clusters with R. felis. Samples from Argas persicus ticks cluster together showing greatest homology with sequences from Candidatus Rickettsia hoogstraalii.
Possession of plasmids Of the primers described for the R. felis plasmids (Fournier et al., 2008), only primer set pRFa-pRFb yielded a positive amplicon of the expected size 159 bp for an Ethiopian flea sample. This is indicative of possession of the pRF plasmid or R. felis. All Rickettsia-positive samples from Ar. persicus ticks were uniformly negative with these primers. Given that primers designed to amplify R. felis plasmids were negative, we targeted a small heat-shock protein, hsp2, believed to reside only on plasmids (Baldridge et al., 2010). Various primer combinations have been designed to target the plasmid-borne hsp2 of Rickettsia and are detailed in Table 1 (Baldridge et al., 2008).
All failed to yield amplicons, with the exception of primer set Hsp2F3-Hsp2R3 designed against the hsp2 gene of R. helvetica that yielded amplicons among 80% of the positive samples from Ar. persicus ticks. All sequences generated were homologous, and a representative sample was deposited under GenBank accession number JN366407. For comparative purposes, the R. felis hsp2 sample was also sequenced and deposited under accession number JN366408. Phylogenetic comparison of these sequences revealed closer homology between sequences derived from R. felis with those of R. helvetica, R. monacensis, and Candidatus R. hoogstraalii, whilst the plasmids from the rickettsial species in Ar. persicus ticks clustered separately (see Fig. 5) only showing 86% homology over 402 bp with that sequence deposited for Candidatus R. hoogstraalii.
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
341
Fig. 3. Neighbour-joining tree for outer membrane protein A (ompA) (bootstrap 1000 replicates, values > 60 shown) over 562 bp. All samples with an Et prefix are samples collected during this study, and these sequences cluster with those from Candidatus R. hoogstraalii. The sequence obtained from Ornithodoros moubata ticks appears more distantly related.
Discussion The finding of Ar. persicus in the Dire Dawa region of Ethiopia has been previously reported (Mekonnen et al., 2007). However, we were surprised by the overwhelming abundance of Ar. persicus infestation observed. In the village where these were collected, ticks were primarily located around poultry areas, their natural preferred host; however, non-specific biting of other livestock and humans was a regular occurrence. The density of ticks in this location is indicated in Fig. 1c whereby vast numbers of ticks could be found under the bark of trees. Indeed, blood of human origin was detected in 2 of the tick pools collected (Cutler et al., 2011). The ability to feed on non-preferred host species for Ar. persicus has been reported elsewhere (Hoogstraal, 1956). Although several diverse pathogens have been reported associated with this tick species including Borrelia, Mycoplasma, Mycobacteria, and West Nile virus, to our knowledge, Rickettsia species have not been previously reported among this species in Ethiopia. There is, however, an
isolated report of spotted fever group Rickettsia species previously published from this vector from Armenia, which might be homologous with that reported herein, however, these authors likened this species to R. slovaca, but this could reflect a lack of available strains and inability to undertake sequence analysis for comparative purposes at this point in time (Rehácek et al., 1977). Phylogenetic analysis of gene sequences from Ar. persicus ticks clearly demonstrated a close resemblance to Candidatus R. hoogstraalii. This was evident from analysis of gltA, ompA, and ompB sequences (see Figs. 2–4). The gltA sequences show tight phylogenetic clustering of all Ar. persicus sequenced amplicons. Interestingly, these showed slight divergence to those sequences produced by Candidatus R. hoogstraalii of O. moubata ticks from Tanzania, possibly reflecting co-evolution with their vectors. Surprisingly, gltA sequencing revealed that the rickettsial sequences derived from O. moubata ticks clustered more closely with 2 sequence types described from fleas, one erroneously reported as R. felis, HM582437 (Bermudez et al., 2011) and the other as
342
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
Fig. 4. Neighbour-joining phylogeny for outer membrane protein B (ompB) gene using 1633 bp (bootstrap 1000 replicates, values > 60 shown). All samples with an Et prefix are samples collected during this study, and these sequences cluster with those from Candidatus Rickettsia hoogstraalii. The sequence obtained from Ornithodoros moubata ticks appears more distantly related. The sequence from fleas collected during this study cluster with those for R. felis.
Fig. 5. Neighbour-joining tree for the plasmid-borne small heat-shock protein 2 (hsp2) (bootstrap 1000 replicates, values > 60 shown) over 402 bp. All samples with an Et prefix are samples collected during this study, and these sequences cluster away from those from Candidatus R. hoogstraalii with the sequence from fleas collected during this study clustering more closely with this group together with sequences deposited for R. felis in GenBank.
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
a Rickettsia species AF516331. Both of these sequences cluster alongside the Candidatus R. hoogstraalii and away from the other sequences for R. felis. Whether fleas can also transmit Candidatus R. hoogstraalii remains to be addressed. Support for this notion comes from the report of a sequence type closely related to Candidatus R. hoogstraalii, but detected in a tsetse fly from Senegal (strain SGL01; GenBank GQ255903). Two amplicons generated with citrate synthase primers were highly divergent from other sequences generated (data not shown). A blast search disclosed these to belong to Acinetobacter baumannii, an organism frequently found in ticks that has been previously reported to amplify with these primers (Cutler et al., 2006). Phylogenetic analysis for outer membrane proteins A and B (Figs. 3 and 4) corroborate those for citrate synthase, showing homology of the Rickettsia species in Ar. persicus with Candidatus R. hoogstraalii. Interestingly, the phylogenetic tree for the ompA and ompB genes again show the separate clustering of the Rickettsia present in O. moubata ticks (Cutler et al., 2006). Most studies have focused upon hard ticks, however, fleas have also been recognised as vectors for rickettsial species since the early 20th century. More recently, soft ticks have been implicated as vectors for Rickettsia species such as the primarily human-feeding O. moubata ticks in Tanzania (Cutler et al., 2006) or the avianfeeding Carios capensis ticks in the USA (Mattila et al., 2007) or indeed C. capensis and C. sawaii in Japan (Kawabata et al., 2006). The rickettsial species detected in Ar. persicus appeared to cluster with members of the spotted fever group including R. australis, R. akari, and R. felis. At the same time, a similar rickettsial species was being reported among the hard tick species Haemaphysalis sulcata in Europe (Duh et al., 2010). The name R. hoogstraalii was proposed for these latter strains. More recently, a similar isolate has been obtained from O. erraticus ticks from Portugal (Milhano et al., poster 251, 6th International meeting on Rickettsia and Rickettsial Diseases, 2011). Intriguingly, these appear not just to be restricted to ticks, but have additionally been reported from mosquitoes (Gracner et al., 2009). As yet, little is known regarding the pathogenic potential of Candidatus R. hoogstraalii. Isolates have been obtained from both H. sulcata and C. capensis, and now also from O. erraticus ticks. Growth has been achieved in Vero, CCE3, ISE6, and the mosquito cell line C6/36. In all but the C6/36 cells, cytopathic effect has been noted (Duh et al., 2010). Whether or not, this species exists as an endosymbiont within its tick vector is currently unclear. Investigations of the Haemaphysalis sulcata isolate failed to demonstrate this Rickettsia in tick ovary tissue, but did find that it was abundant in the salivary glands and to a lesser degree in tick midgut tissues (Duh et al., 2010). Whereas phylogenetic analysis of the rickettsial species present in Ar. persicus ticks showed its close relationship with Candidatus R. hoogstraalii, in contrast, the hsp2 gene comparison revealed this gene sequence to be divergent from that described for Candidatus R. hoogstraalii isolated from C. capensis ticks (Baldridge et al., 2008). Multiple origins for rickettsial plasmids have been suggested, however, hsp2 to date has only been described to reside on plasmids (Baldridge et al., 2010). Paradoxically, the lack of hsp2 similarity found between Candidatus R. hoogstraalii and the gene sequence found among Ar. persicus ticks could be explained by a different origin of the plasmid (or hsp2 gene) present in the Ar. persicus Rickettsia. The sequencing of further plasmid gene targets should further substantiate this observation. Further support for this plasmid being divergent from that initially reported in R. felis comes from the failure of R. felis plasmid-specific primers to produce amplicons from these ticks. The finding of R. africae among hard ticks confirms the reports of others who have described this species to be present in several species of tick vector (Fournier et al., 1998; Macaluso et al.,
343
2003). Various Amblyomma species have been associated with R. africae and human cases reported following bites from Am. variegatum ticks in Ethiopia (Stephany et al., 2009). Rhipicephalus species have also been found to carry R. africae, but usually to a lesser degree (Macaluso et al., 2003). As both these tick species often co-feed on Ethiopian cattle and in consequence, our hard tick pools often contained both species, thus we were not able to categorically state the contribution of each tick species to the positive finding. However, as a result of co-feeding, ticks on the same host would potentially both contribute to the positive finding. This rickettsial species is being increasingly recognised as a common cause of febrile infection among travellers returning from Africa (Jensenius et al., 2009, 2004; Mediannikov et al., 2010b). Exposure appears to be significant, in part as a result of the particularly aggressive behaviour of Amblyomma tick vectors evidenced by the frequent detection of multiple inoculation eschars in patients (Raoult et al., 2001). The extent of infection resulting febrile disease among the indigenous population has to date not been explored in depth, but appears to be significant, especially during childhood (Mediannikov et al., 2010a). Similarly, R. felis is becoming increasingly recognised to be prevalent in many African nations. Although fleas were not the main focus of this study, where encountered, they were collected and subsequently tested. Our finding of 2 of the 3 flea pools to be positive is in keeping with the high levels of carriage reported by others and the significant, yet currently largely unrecognised role of R. felis as a cause of febrile infection in sub-Saharan Africa (Parola, 2011; Socolovschi et al., 2010). Drawing conclusions regarding incidence based upon such small numbers as reported herein would be unsound, but we can conclude that R. felis appears abundant among Ethiopian fleas. In conclusion, we document the presence of R. africae and R. felis among hard ticks and fleas from Ethiopia, respectively. We also describe carriage of a Rickettsia-resembling Candidatus R. hoogstraalii present among Ar. persicus ticks in Ethiopia. Although this species is not yet associated with clinical consequences either for humans or their preferred poultry hosts, investigative studies of this species are still in their infancy. Acknowledgements We are indebted to the Presidents Fund from the Society for General Microbiology for making this study possible. We also express our gratitude to Alan R. Walker, Royal (Dick) School of Veterinary Studies, University of Edinburgh, UK, for expert advice regarding the identification of the Argas ticks. References Baldridge, G.D., Burkhardt, N.Y., Felsheim, R.F., Kurtti, T.J., Munderloh, U.G., 2008. Plasmids of the pRM/pRF family occur in diverse Rickettsia species. Appl. Environ. Microbiol. 74, 645–652. Baldridge, G.D., Burkhardt, N.Y., Labruna, M.B., Pacheco, R.C., Paddock, C.D., Williamson, P.C., Billingsley, P.M., Felsheim, R.F., Kurtti, T.J., Munderloh, U.G., 2010. Wide dispersal and possible multiple origins of low-copy-number plasmids in Rickettsia species associated with blood-feeding arthropods. Appl. Environ. Microbiol. 76, 1718–1731. Bermudez, C.S.E., ZaldÌvar, A.Y., Spolidorio, M.G., Moraes-Filho, J., Miranda, R.J., Caballero, C.M., Mendoza, Y., Labruna, M.B., 2011. Rickettsial infection in domestic mammals and their ectoparasites in El Valle de Anton, Cocle, Panama. Vet. Parasitol. 177, 134–138. Burgdorfer, W., Ormsbee, R.A., Schmidt, M.L., Hoogstraal, H., 1973. A search for the epidemic typhus agent in Ethiopian ticks. Bull. World Health Organ. 48, 563–569. Cutler, S.J., Abdissa, A., Adamu, H., Tolosa, T., Gashaw, A., 2011. Borrelia in Ethiopian ticks. Ticks Tick-Borne Dis. 3, 14–17. Cutler, S.J., Browning, P., Scott, J.C., 2006. Ornithodoros moubata, a soft tick vector for Rickettsia in East Africa? Ann. N.Y. Acad. Sci. 1078, 373–377. Duh, D., Punda-Polic, V., Avˇsiˇc-Zˇ upanc, T., Bouyer, D., Walker, D.H., Popov, V.L., Jelovsek, M., Gracner, M., Trilar, T., Bradaric, N., Kurtti, T.J., Strus, J., 2010. Rickettsia hoogstraalii sp. nov., isolated from hard- and soft-bodied ticks. Int. J. Syst. Evol. Microbiol. 60, 977–984.
344
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
Fournier, P.-E., Roux, V., Caumes, E., Donzel, M., Raoult, D., 1998. Outbreak of Rickettsia africae infections in participants of an adventure race in South Africa. Clin. Infect. Dis. 27, 316–323. Fournier, P.E., Belghazi, L., Robert, C., Elkarkouri, K., Richards, A.L., Greub, G., Collyn, F., Ogawa, M., Portillo, A., Oteo, J.A., Psaroulaki, A., Bitam, I., Raoult, D., 2008. Variations of plasmid content in Rickettsia felis. PLoS One 3, e2289, doi:2210.1371/journal.pone.0002289. Gracner, M., Avˇsiˇc-Zˇ upanc, T., Punda-Polic, V., Dolinsek, J., Bouyer, D., Walker, D.H., Zavala-Castro, J.E., Bradaric, N., Crocquet-Valdes, P.A., Duh, D., 2009. Comparative ompA gene sequence analysis of Rickettsia felis-like bacteria detected in Haemaphysalis sulcata ticks and isolated in the mosquito C6/36 cell line. Clinical Microbiology and Infection 15, 265–266. Hoogstraal, H., 1956. Ticks of the Sudan. In: African Ixodoidea. U.S. Government Printing Office, Washington, pp. 1–1101. Jensenius, M., Davis, X., von Sonnenburg, F., Schwartz, E., Keystone, J.S., Leder, K., Lopéz-Véléz, R., Caumes, E., Cramer, J.P., Chen, L., Parola, P., 2009. Multicenter GeoSentinel analysis of rickettsial diseases in international travelers, 1996–2008. Emerg. Infect. Dis. 15, 1791–1798. Jensenius, M., Fournier, P.E., Raoult, D., 2004. Rickettsioses and the international traveler. Clin. Infect. Dis. 39, 1493–1499. Kawabata, H., Ando, S., Kishimoto, T., Kurane, I., Takano, A., Nogami, S., Fujita, H., Tsurumi, M., Nakamura, N., Sato, F., Takahashi, M., Ushijima, Y., Fukunaga, M., Watanabe, H., 2006. First detection of Rickettsia in soft-bodied ticks associated with seabirds, Japan. Microbiol. Immunol. 50, 403–406. Letaief, A., 2006. Epidemiology of rickettsioses in North Africa. Ann. N.Y. Acad. Sci. 1078, 34–41. Macaluso, K.R., Davis, J.O.N., Alam, U., Korman, A.M.Y., Rutherford, J.S., Rosenberg, R., Azad, A.F., 2003. Spotted fever group rickettsiae in ticks from the Masai Mara region of Kenya. Am. J. Trop. Med. Hyg. 68, 551–553. Mattila, J., Burkhardt, N., Hutcheson, H., Munderloh, U., Kurtti, T., 2007. Isolation of cell lines and a rickettsial endosymbiont from the soft tick Carios capensis (Acari: Argasidae: Ornithodorinae). J. Med. Entomol. 44, 1091–1101. Mediannikov, O., Diatta, G., Fenollar, F., Sokhna, C., Trape, J.-F., Raoult, D., 2010a. Tickborne rickettsioses, neglected emerging diseases in rural Senegal. PLoS Negl. Trop. Dis. 4, e821. Mediannikov, O., Trape, J.-F., Diatta, G., Parola, P., Fournier, P.-E., Raoult, D., 2010b. Rickettsia africae, Western Africa. Emerg. Infect. Dis. 16, 571–573. Mekonnen, S., Pegram, R.G., Gebre, S., Mekonnen, A., Jobre, Y., Zewde, S., 2007. A synthesis review of ixodid (Acari:Ixodidae) and argasid (Acari: Argasidae) ticks in Ethiopia and their possible roles in disease transmission. Ethiopian Vet. J. 11, 1–24. Morita, C., El Hussein, A.R., Matsuda, E., Abdel Gabbar, K.M., Muramatsu, Y., Abdel Rahman, M.B., Eleragi, A.M., Hassan, S.M., Chitambo, A.M., Ueno, H., 2004.
Spotted fever group rickettsiae from ticks captured in Sudan. Jpn. J. Infect. Dis. 57, 107–109. Mura, A., Socolovschi, C., Ginesta, J., Lafrance, B., Magnan, S., Rolain, J.M., Davoust, B., Raoult, D., Parola, P., 2008. Molecular detection of spotted fever group rickettsiae in ticks from Ethiopia and Chad. Trans. R. Soc. Trop. Med. Hyg. 102, 945–949. Parola, P., 2006. Rickettsioses in sub-Saharan Africa. Ann. N.Y. Acad. Sci. 1078, 42–47. Parola, P., 2011. Rickettsia felis from a rare disease in the USA to a common cause of fever in sub-Saharan Africa. Clin. Microbiol. Infect. 17, 996–1000. Portillo, A., Perez-Martinez, L., Santibanez, S., Blanco, J.R., Ibarra, V., Oteo, J.A., 2007. Detection of Rickettsia africae in Rhipicephalus (Boophilus) decoloratus ticks from the Republic of Botswana, South Africa. Am. J. Trop. Med. Hyg. 77, 376–377. Pretorius, A.-M., Birtles, R.J., 2002. Rickettsia aeschlimannii: a new pathogenic spotted fever group Rickettsia, South Africa. Emerg. Infect. Dis. 8, 874. Raoult, D., Fournier, P.E., Fenollar, F., Jensenius, M., Prioe, T., de Pina, J.J., Caruso, G., Jones, N., Laferl, H., Rosenblatt, J.E., Marrie, T.J., 2001. Rickettsia africae, a tick-borne pathogen in travelers to sub-Saharan Africa. N. Engl. J. Med. 344, 1504–1510. Rehácek, J., Urvölgyi, J., Kovácová, E., 1977. Massive occurrence of rickettsiae of the spotted fever group in fowl tampan, Argas persicus, in the Armenian S.S.R. Acta Virol. 21, 431–438. Roux, V., Raoult, D., 2000. Phylogenetic analysis of members of the genus Rickettsia using the gene encoding the outer-membrane protein rOmpB (ompB). Int. J. Syst. Evol. Microbiol. 50, 1449–1455. Roux, V., Rydkina, E., Eremeeva, M., Raoult, D., 1997. Citrate synthase gene comparison, a new tool for phylogenetic analysis, and its application for the rickettsiae. Int. J. Syst. Bacteriol. 47, 252–261. Socolovschi, C., Mediannikov, O., Sokhna, C., Tall, A., Diatta, G., Bassene, H., Trape, J.F., Raoult, D., 2010. Rickettsia felis-associated uneruptive fever. Senegal. Emerg Infect Dis 16, 1140–1142. Stenos, J., Graves, S.R., Unsworth, N.B., 2005. A highly sensitive and specific real-time PCR assay for the detection of spotted fever and typhus group Rickettsiae. Am. J. Trop. Med. Hyg. 73, 1083–1085. Stephany, D., Buffet, P., Rolain, J.M., Raoult, D., Consigny, P.H., 2009. Rickettsia africae infection in man after travel to Ethiopia. Emerg. Infect. Dis. 15, 1867–1869. Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., Kumar, S., 2011. MEGA5: Molecular Evolutionary Genetics Analysis using Maximum Likelihood, Evolutionary Distance, and Maximum Parsimony Methods. Mol. Biol. Evol., http://dx.doi.org/10.1093/molbev/msr121. Vitorino, L., Chelo, I.M., Bacellar, F., Ze-Ze, L., 2007. Rickettsiae phylogeny: a multigenic approach. Microbiology 153, 160–168.
Contents lists available at SciVerse ScienceDirect
Ticks and Tick-borne Diseases journal homepage: www.elsevier.com/locate/ttbdis
Original article
Candidatus Rickettsia hoogstraalii in Ethiopian Argas persicus ticks Vera Pader a , Joanna Nikitorowicz Buniak a , Alemseged Abdissa b , Haileeysus Adamu b , Tadele Tolosa c , Abebaw Gashaw c , Ronald R. Cutler d , Sally J. Cutler a,∗ a
School of Health, Sports & Bioscience, University of East London, London, UK College of Public Health and Medical Sciences, Jimma University, Jimma, Ethiopia College of Agriculture and Veterinary Medicine, Jimma University, Jimma, Ethiopia d School of Biological & Chemical Sciences, Queen Mary University of London, London, UK b c
a r t i c l e
i n f o
Keywords: Argas persicus Ornithodoros moubata Amblyomma variegatum Rickettsia Candidatus Rickettsia hoogstraalii Rickettsia felis
a b s t r a c t Ethiopian soft ticks Argas persicus, hard ticks including both Amblyomma variegatum and Rhipicephalus (Boophilus) spp., and fleas were collected from livestock, traditional human dwellings, and cracks and crevices of trees. They were assessed in pools for the presence of Rickettsia using PCR-based methods. The extracted tick DNA was subjected to molecular screening for Rickettsia, which revealed 50.5% of the pooled samples to be positive for Rickettsia spp. These were then subjected to multi-gene analysis using both outer surface proteins and housekeeping genes with proven discriminatory potential. Sequencing of the citrate synthase and outer membrane genes clearly led to the identification of three distinct rickettsial species, Candidatus Rickettsia hoogstraalii in Argas persicus ticks; R. africae in hard tick pools, and R. felis in fleas. Furthermore, we demonstrated the presence of the plasmid-borne small heat-shock protein gene hsp2 in DNA from A. persicus ticks suggesting that Candidatus R. hoogstraalii carried by these ticks possess a plasmid. Unlike chromosomal gene sequences, the hsp2 gene failed to cluster with Candidatus R. hoogstraalii, instead falling into an isolated separate clade, suggesting a different origin for the plasmid. © 2012 Elsevier GmbH. All rights reserved.
Introduction Rickettsiae are transmitted by the bite of various haematophagous arthropods such as ticks, fleas, lice, and mites. With the recent application of molecular methods to arthropods from various locations, many new rickettsial species have been recognised, including several newly recognised pathogenic species. This prompted us to revisit the Rickettsia present in Ethiopian ticks. The presence of R. africae is already established among Amblyomma variegatum and other species of ticks in Ethiopia, for example, R. africae strain ESF-5 originated from the Shulu Province of Ethiopia (Roux et al., 1997) and is found extensively throughout other African countries (Fournier et al., 1998; Mediannikov et al., 2010b; Morita et al., 2004; Mura et al., 2008; Portillo et al., 2007). Similarly, several recent reports have documented the presence of R. felis among fleas from various African nations (Parola, 2011). Other rickettsial species from Africa include R. aeschlimannii, R. conorii, R.
∗ Corresponding author at: School of Health and Bioscience, University of East London, Water Lane, Stratford, London E15 4LZ, UK. Tel.: +44 0208 223 6386; fax: +44 0208 223 4959. E-mail address: [email protected] (S.J. Cutler). 1877-959X/$ – see front matter © 2012 Elsevier GmbH. All rights reserved. http://dx.doi.org/10.1016/j.ttbdis.2012.10.021
massiliae, and R. sibirica mongolitimonae of the spotted fever group and R. prowazekii and R. typhi of the typhus group (Letaief, 2006; Morita et al., 2004; Parola, 2006; Pretorius and Birtles, 2002). Interestingly, it has also been hypothesised that a tick reservoir for R. prowazekii might also exist in Ethiopia (Burgdorfer et al., 1973). Despite this great rickettsial diversity, cases are seldom reported among the indigenous population. Instead, these are more often observed among the ever-increasing numbers of tourists that now visit these regions (Jensenius et al., 2009; Pretorius and Birtles, 2002). Sporadic infections and even clusters of infection have been reported, often diagnosed once these travellers have returned home with fever (Stephany et al., 2009). The burden of rickettsial infection among African populations is only just starting to be recognised, with limited studies investigating the role of Rickettsia in febrile patients attending clinics and hospitals (Mediannikov et al., 2010a; Parola, 2011; Socolovschi et al., 2010). A limiting factor is the lack of molecular diagnostics available within healthcare settings and the relatively poor diagnostic performance of serology in highly endemic situations. Given the greater precision offered by molecular approaches for identification of rickettsiae, we undertook an investigation of various tick species from two regions in Ethiopia to explore the diversity of rickettsiae present.
338
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
Materials and methods
Confirmatory PCR
Ticks collected from Ethiopia
Arthropod pools found to be positive upon screening were further tested using conventional PCR using a MyCycler thermocycler (BioRad) for multi-gene targets previously described including gltA, ompA, dnaK, dnaA, atpA, recA, and the internal rrl-rrf ribosomal spacer (ITS) (Vitorino et al., 2007). Additionally, primers described for amplification of fragments of outer surface proteins A and B were used as previously described ompA (designed using NCBI primer design software, this study) and for partial sequence amplification of the 5 end of ompB (Roux and Raoult, 2000). Larger amplicons for sequencing citrate synthase used the primers and methods described by Roux et al. (1997). To explore the possibility of plasmids among any Rickettsia detected, we used the primers described to characterise plasmids recently described among R. felis (Fournier et al., 2008). Additionally, a small heat shock protein Hsp2 has been reported to reside on rickettsial plasmids, and primers against this target were also used to characterise positive samples according to published protocols (Baldridge et al., 2008). These primers are detailed in Table 1. PCR products were observed using the SybrSafe (Invitrogen) DNA stain in 2% (w/v) agarose gels. The amplicon sizes were measured relative to a 100-bp molecular mass standard (Fermentas).
A total of 98 pools of up to 20 ticks was collected from two different geographical regions of Ethiopia during July to August 2010. The collection areas were Jimma to the southwest and Dire Dawa to the east of Addis Ababa, respectively. Ticks were pooled when collected from the same host, thus some contained multiple species, or when the numbers exceeded 20 from the same environmental location, these were separated into different pools. No pool contained more than 20 ticks. Hard ticks were removed from livestock, whilst soft ticks were collected from cracks and crevices of human dwellings and from under the bark of trees in the vicinity of livestock areas. Limited collections of fleas (Ctenocephalides spp.) were made in addition to the ticks (3 pools: 2 containing 3 fleas; and a single flea in the third), with these being collected from clothing or bedding in traditional human dwellings. All collected arthropod pools were immersed in approximately 0.5 ml of 8 M guanidine hydrochloride and imported to the UK for analysis under licence. DNA extraction Tick and flea pools were transferred into a fresh tube together with glass beads and a volume of sterile PBS equivalent to the size of the arthropods within each pool. These were homogenised using a bead beater prior to incubation with tissue lysis buffer and proteinase K at 56 ◦ C for 1–4 h until liquefied. Debris was pulsed down with a 1-min spin in a microfuge and the supernatant removed for DNA extraction using DNeasy kits (Qiagen) on a QiaCube robot (Qiagen) using the rodent tail protocol. PCR screening All samples were screened using a real-time generic rickettsial PCR (Stenos et al., 2005). Flea samples were additionally screened using real-time assays specific for R. felis biotin synthase (Socolovschi et al., 2010). “Blank” samples containing sterile PBS that were processed alongside the tick pools, and sterile DNA-free water served as negative controls. All real-time assays were performed as originally described, using Stratagene Mx3000p cycler (Agilent). All primers and probes are listed in Table 1.
Sequence analysis Where amplicons of suitable quality were produced, these were sent to the Genome Centre Queen Mary University of London for sequencing, using an Applied Biosystems ABI 3730xl. The chromatograms were analysed and the sequence data was edited, aligned and trimmed using ClustalW and Mega5. The latter was used for phylogenetic tree construction using a bootstrap value of 1000 replicates (Tamura et al., 2011). Results Arthropod collections Small numbers of hard ticks were also collected by removal from domestic livestock and companion animals. A total of 21 pooled DNA extracts from hard ticks [Amblyomma variegatum and Rhipicephalus (Boophilus) spp.] were collected from animals in both Dire
Table 1 Primers and probes used during investigation. Gene target
Forward primer 5 –3
Reverse primer 5 –3
Probe (where applicable)
Reference
Citrate synthase (gltA)
TCGCAAATGTTCACGGTACTTT
CACAATGGAAAGAAATGCACGA
Stenos et al. (2005)
R. felis biotin synthase
ATGTTCGGGCTTCCGGTATG
CCGATTCAGCAGGTTCTTCAA
FAM-TGCAATAGCAAGAACCGTAGGCTGGATG-BHQ1 6-FAMGCTGCGGCGGTATTTTAGGAATGGG-TAMRA
Citrate synthase Rp877-CS1273 Outer membrane protein (ompA) Outer membrane protein (ompB) F1 Outer membrane protein (ompB) F2 ATP synthase F1 subunit alpha atpA Chromosomal replication initiation protein dnaA Heat shock protein 70 dnaK R. felis plasmid pRFa-b R. felis plasmid pRFc-d hsp2 Hsp2F3-Hsp2R3
GGGGACCTGCTCACGGCGG CGGTGTTGTAGGGACTGCGGC CCGCAGGGTTGGTAACTGC
CATAACGAGTGTAAAGCTG CGGTCCGCCCGGTTTGAAGT CCTTTTAGATTACCGCCTAA
Roux et al. (1997) This study Roux and Raoult (2000)
AATATCGGTGACGGTCAAGG
CCTATATCGCCGGTAATT
Roux and Raoult (2000)
AGTACAGACATATCGAGATGA CTTTACAATCATTACGGTG
CGACTTACCGAAATACCGAC GCAACTAAGCCCCATCC
Vitorino et al. (2007) Vitorino et al. (2007)
GCATTCTAGTCATACCGCC CAAGCTTTTGTACTGCCTCTAT ACATTCCGTAAAGAATATGAGC CTTAGCCTTACTTTGTTCTTTTTTAGG
Vitorino et al. (2007) Fournier et al. (2008) Fournier et al. (2008) Baldridge et al. (2008)
hsp2 Rhoogs qHsp2F1- Rhoogs qHsp2R hsp2 Hsp2F3-Hsp2R
GATGGTAAACTAATGGATCGC
CAAAAAATGAAAGAAACTGCTGA AGTGCATATAGCTACCACACTATCT GCTTATGTTCGCCTTTAGTATTTA CTTTTAATTTATCACGTATTAGAAATAAATCAG TGAGGGCAAAAATGAACAATC
CTTAGCCTTACTTTGTTCTTTTTTAGG
AGGCAAAGGCGAGAGAAATACC
Baldridge et al. (2008)
Socolovschi et al. (2010)
Baldridge et al. (2008)
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
339
Fig. 1. Plate A shows an Amblyomma variegatum tick forming part of a hard tick pool. Plate B shows a representative Argas persicus tick forming part of a soft tick pool. Plate C illustrates the tick density under the bark of trees; whilst Plate D shows ticks within a wooden beam removed from a traditional style Ethiopian dwelling.
Dawa and Jimma regions (see Fig. 1a). Soft ticks identified as Argas persicus (pers. communication Alan Walker, University of Edinburgh, UK) were collected from a village (Asselisso) on the outskirts of Dire Dawa (see Fig. 1b). This village maintained a huge tick burden with ticks not only surrounding livestock areas predominantly under the bark of trees (see Fig. 1c), but also residing within human traditionally built dwellings (see Fig. 1d). We were able to collect 77 pools containing 5–20 ticks from this village. No soft ticks were collected from other sites. Three pools of fleas were collected from human clothing/bedding. Screening of samples Screening of pooled samples revealed 5 of the 21 (24%) hard tick pools and 44 of the 77 (57%) soft tick pools to be positive, whilst the R. felis-specific biotin synthase assay yielded positive results from 2 of the 3 flea pools only (both containing 3 fleas). Identification of Rickettsia detected Conventional PCR to produce fragments for sequence analysis from DNA-extracted tick pools failed to yield full overlapping contigs, thus precluding concatenation for complete gene analysis. Partial gene sequences were generated for citrate synthase (gltA) and outer membrane proteins OmpA and OmpB that have been extensively used for rickettsial identification. Figs. 2–4 show the phylogenetic comparison of our sequences with homologous sequences from validated species and their nearest neighbours
available from GenBank. These phylogenetic trees revealed the presence of R. africae in a hard tick pool (containing Am. variegatum ticks) and R. felis in the positive flea pools (R. felis gltA partial gene sequence deposited under GenBank accession number JN366415; ompB partial gene sequence under accession number JN366420). Rickettsial DNA detected in A. persicus had greatest homology with Candidatus R. hoogstraalii previously reported from other genera of soft ticks collected from Africa, Japan, and the USA (Cutler et al., 2006; Kawabata et al., 2006; Mattila et al., 2007) and among European Haemaphysalis sulcata ticks from Croatia (Duh et al., 2010). This showed 99.5–100% sequence similarity between Candidatus R. hoogstraalii and the amplicons from Ar. persicus over the 428 bp analysed. Representative partial gene sequences for gltA, ompA, and ompB are deposited under GenBank accession numbers JN366413–JN366414 (gltA), JN366416 (ompA), and JN366418–JN366419 (ompB). For comparative purposes, an earlier sample from Tanzanian Ornithodoros moubata soft ticks was submitted for partial ompA and ompB gene sequencing (GenBank accession numbers JN366417 and JN366421, respectively) (Cutler et al., 2006). Sequence results from other “housekeeping” gene targets including dnaA (deposited under GenBank accession JN366410), dnaK (deposited under GenBank accession numbers JN366411 and JN366412), and atpA (deposited under GenBank accession number JN366409) showed poor discriminatory power among rickettsial species (data not shown). The remaining 4 gene targets described for multi-gene identification failed to produce amplicons from positive tick DNA extracts (Vitorino et al., 2007).
340
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
Fig. 2. Neighbour-joining tree citrate synthase (gltA) (bootstrap 1000 replicates, values > 60 shown) over 428 bp. All samples with an Et prefix are samples collected during this study. Sample Et161b from hard ticks clusters with Rickettsia africae whilst sample Et90 from a flea pool clusters with R. felis. Samples from Argas persicus ticks cluster together showing greatest homology with sequences from Candidatus Rickettsia hoogstraalii.
Possession of plasmids Of the primers described for the R. felis plasmids (Fournier et al., 2008), only primer set pRFa-pRFb yielded a positive amplicon of the expected size 159 bp for an Ethiopian flea sample. This is indicative of possession of the pRF plasmid or R. felis. All Rickettsia-positive samples from Ar. persicus ticks were uniformly negative with these primers. Given that primers designed to amplify R. felis plasmids were negative, we targeted a small heat-shock protein, hsp2, believed to reside only on plasmids (Baldridge et al., 2010). Various primer combinations have been designed to target the plasmid-borne hsp2 of Rickettsia and are detailed in Table 1 (Baldridge et al., 2008).
All failed to yield amplicons, with the exception of primer set Hsp2F3-Hsp2R3 designed against the hsp2 gene of R. helvetica that yielded amplicons among 80% of the positive samples from Ar. persicus ticks. All sequences generated were homologous, and a representative sample was deposited under GenBank accession number JN366407. For comparative purposes, the R. felis hsp2 sample was also sequenced and deposited under accession number JN366408. Phylogenetic comparison of these sequences revealed closer homology between sequences derived from R. felis with those of R. helvetica, R. monacensis, and Candidatus R. hoogstraalii, whilst the plasmids from the rickettsial species in Ar. persicus ticks clustered separately (see Fig. 5) only showing 86% homology over 402 bp with that sequence deposited for Candidatus R. hoogstraalii.
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
341
Fig. 3. Neighbour-joining tree for outer membrane protein A (ompA) (bootstrap 1000 replicates, values > 60 shown) over 562 bp. All samples with an Et prefix are samples collected during this study, and these sequences cluster with those from Candidatus R. hoogstraalii. The sequence obtained from Ornithodoros moubata ticks appears more distantly related.
Discussion The finding of Ar. persicus in the Dire Dawa region of Ethiopia has been previously reported (Mekonnen et al., 2007). However, we were surprised by the overwhelming abundance of Ar. persicus infestation observed. In the village where these were collected, ticks were primarily located around poultry areas, their natural preferred host; however, non-specific biting of other livestock and humans was a regular occurrence. The density of ticks in this location is indicated in Fig. 1c whereby vast numbers of ticks could be found under the bark of trees. Indeed, blood of human origin was detected in 2 of the tick pools collected (Cutler et al., 2011). The ability to feed on non-preferred host species for Ar. persicus has been reported elsewhere (Hoogstraal, 1956). Although several diverse pathogens have been reported associated with this tick species including Borrelia, Mycoplasma, Mycobacteria, and West Nile virus, to our knowledge, Rickettsia species have not been previously reported among this species in Ethiopia. There is, however, an
isolated report of spotted fever group Rickettsia species previously published from this vector from Armenia, which might be homologous with that reported herein, however, these authors likened this species to R. slovaca, but this could reflect a lack of available strains and inability to undertake sequence analysis for comparative purposes at this point in time (Rehácek et al., 1977). Phylogenetic analysis of gene sequences from Ar. persicus ticks clearly demonstrated a close resemblance to Candidatus R. hoogstraalii. This was evident from analysis of gltA, ompA, and ompB sequences (see Figs. 2–4). The gltA sequences show tight phylogenetic clustering of all Ar. persicus sequenced amplicons. Interestingly, these showed slight divergence to those sequences produced by Candidatus R. hoogstraalii of O. moubata ticks from Tanzania, possibly reflecting co-evolution with their vectors. Surprisingly, gltA sequencing revealed that the rickettsial sequences derived from O. moubata ticks clustered more closely with 2 sequence types described from fleas, one erroneously reported as R. felis, HM582437 (Bermudez et al., 2011) and the other as
342
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
Fig. 4. Neighbour-joining phylogeny for outer membrane protein B (ompB) gene using 1633 bp (bootstrap 1000 replicates, values > 60 shown). All samples with an Et prefix are samples collected during this study, and these sequences cluster with those from Candidatus Rickettsia hoogstraalii. The sequence obtained from Ornithodoros moubata ticks appears more distantly related. The sequence from fleas collected during this study cluster with those for R. felis.
Fig. 5. Neighbour-joining tree for the plasmid-borne small heat-shock protein 2 (hsp2) (bootstrap 1000 replicates, values > 60 shown) over 402 bp. All samples with an Et prefix are samples collected during this study, and these sequences cluster away from those from Candidatus R. hoogstraalii with the sequence from fleas collected during this study clustering more closely with this group together with sequences deposited for R. felis in GenBank.
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
a Rickettsia species AF516331. Both of these sequences cluster alongside the Candidatus R. hoogstraalii and away from the other sequences for R. felis. Whether fleas can also transmit Candidatus R. hoogstraalii remains to be addressed. Support for this notion comes from the report of a sequence type closely related to Candidatus R. hoogstraalii, but detected in a tsetse fly from Senegal (strain SGL01; GenBank GQ255903). Two amplicons generated with citrate synthase primers were highly divergent from other sequences generated (data not shown). A blast search disclosed these to belong to Acinetobacter baumannii, an organism frequently found in ticks that has been previously reported to amplify with these primers (Cutler et al., 2006). Phylogenetic analysis for outer membrane proteins A and B (Figs. 3 and 4) corroborate those for citrate synthase, showing homology of the Rickettsia species in Ar. persicus with Candidatus R. hoogstraalii. Interestingly, the phylogenetic tree for the ompA and ompB genes again show the separate clustering of the Rickettsia present in O. moubata ticks (Cutler et al., 2006). Most studies have focused upon hard ticks, however, fleas have also been recognised as vectors for rickettsial species since the early 20th century. More recently, soft ticks have been implicated as vectors for Rickettsia species such as the primarily human-feeding O. moubata ticks in Tanzania (Cutler et al., 2006) or the avianfeeding Carios capensis ticks in the USA (Mattila et al., 2007) or indeed C. capensis and C. sawaii in Japan (Kawabata et al., 2006). The rickettsial species detected in Ar. persicus appeared to cluster with members of the spotted fever group including R. australis, R. akari, and R. felis. At the same time, a similar rickettsial species was being reported among the hard tick species Haemaphysalis sulcata in Europe (Duh et al., 2010). The name R. hoogstraalii was proposed for these latter strains. More recently, a similar isolate has been obtained from O. erraticus ticks from Portugal (Milhano et al., poster 251, 6th International meeting on Rickettsia and Rickettsial Diseases, 2011). Intriguingly, these appear not just to be restricted to ticks, but have additionally been reported from mosquitoes (Gracner et al., 2009). As yet, little is known regarding the pathogenic potential of Candidatus R. hoogstraalii. Isolates have been obtained from both H. sulcata and C. capensis, and now also from O. erraticus ticks. Growth has been achieved in Vero, CCE3, ISE6, and the mosquito cell line C6/36. In all but the C6/36 cells, cytopathic effect has been noted (Duh et al., 2010). Whether or not, this species exists as an endosymbiont within its tick vector is currently unclear. Investigations of the Haemaphysalis sulcata isolate failed to demonstrate this Rickettsia in tick ovary tissue, but did find that it was abundant in the salivary glands and to a lesser degree in tick midgut tissues (Duh et al., 2010). Whereas phylogenetic analysis of the rickettsial species present in Ar. persicus ticks showed its close relationship with Candidatus R. hoogstraalii, in contrast, the hsp2 gene comparison revealed this gene sequence to be divergent from that described for Candidatus R. hoogstraalii isolated from C. capensis ticks (Baldridge et al., 2008). Multiple origins for rickettsial plasmids have been suggested, however, hsp2 to date has only been described to reside on plasmids (Baldridge et al., 2010). Paradoxically, the lack of hsp2 similarity found between Candidatus R. hoogstraalii and the gene sequence found among Ar. persicus ticks could be explained by a different origin of the plasmid (or hsp2 gene) present in the Ar. persicus Rickettsia. The sequencing of further plasmid gene targets should further substantiate this observation. Further support for this plasmid being divergent from that initially reported in R. felis comes from the failure of R. felis plasmid-specific primers to produce amplicons from these ticks. The finding of R. africae among hard ticks confirms the reports of others who have described this species to be present in several species of tick vector (Fournier et al., 1998; Macaluso et al.,
343
2003). Various Amblyomma species have been associated with R. africae and human cases reported following bites from Am. variegatum ticks in Ethiopia (Stephany et al., 2009). Rhipicephalus species have also been found to carry R. africae, but usually to a lesser degree (Macaluso et al., 2003). As both these tick species often co-feed on Ethiopian cattle and in consequence, our hard tick pools often contained both species, thus we were not able to categorically state the contribution of each tick species to the positive finding. However, as a result of co-feeding, ticks on the same host would potentially both contribute to the positive finding. This rickettsial species is being increasingly recognised as a common cause of febrile infection among travellers returning from Africa (Jensenius et al., 2009, 2004; Mediannikov et al., 2010b). Exposure appears to be significant, in part as a result of the particularly aggressive behaviour of Amblyomma tick vectors evidenced by the frequent detection of multiple inoculation eschars in patients (Raoult et al., 2001). The extent of infection resulting febrile disease among the indigenous population has to date not been explored in depth, but appears to be significant, especially during childhood (Mediannikov et al., 2010a). Similarly, R. felis is becoming increasingly recognised to be prevalent in many African nations. Although fleas were not the main focus of this study, where encountered, they were collected and subsequently tested. Our finding of 2 of the 3 flea pools to be positive is in keeping with the high levels of carriage reported by others and the significant, yet currently largely unrecognised role of R. felis as a cause of febrile infection in sub-Saharan Africa (Parola, 2011; Socolovschi et al., 2010). Drawing conclusions regarding incidence based upon such small numbers as reported herein would be unsound, but we can conclude that R. felis appears abundant among Ethiopian fleas. In conclusion, we document the presence of R. africae and R. felis among hard ticks and fleas from Ethiopia, respectively. We also describe carriage of a Rickettsia-resembling Candidatus R. hoogstraalii present among Ar. persicus ticks in Ethiopia. Although this species is not yet associated with clinical consequences either for humans or their preferred poultry hosts, investigative studies of this species are still in their infancy. Acknowledgements We are indebted to the Presidents Fund from the Society for General Microbiology for making this study possible. We also express our gratitude to Alan R. Walker, Royal (Dick) School of Veterinary Studies, University of Edinburgh, UK, for expert advice regarding the identification of the Argas ticks. References Baldridge, G.D., Burkhardt, N.Y., Felsheim, R.F., Kurtti, T.J., Munderloh, U.G., 2008. Plasmids of the pRM/pRF family occur in diverse Rickettsia species. Appl. Environ. Microbiol. 74, 645–652. Baldridge, G.D., Burkhardt, N.Y., Labruna, M.B., Pacheco, R.C., Paddock, C.D., Williamson, P.C., Billingsley, P.M., Felsheim, R.F., Kurtti, T.J., Munderloh, U.G., 2010. Wide dispersal and possible multiple origins of low-copy-number plasmids in Rickettsia species associated with blood-feeding arthropods. Appl. Environ. Microbiol. 76, 1718–1731. Bermudez, C.S.E., ZaldÌvar, A.Y., Spolidorio, M.G., Moraes-Filho, J., Miranda, R.J., Caballero, C.M., Mendoza, Y., Labruna, M.B., 2011. Rickettsial infection in domestic mammals and their ectoparasites in El Valle de Anton, Cocle, Panama. Vet. Parasitol. 177, 134–138. Burgdorfer, W., Ormsbee, R.A., Schmidt, M.L., Hoogstraal, H., 1973. A search for the epidemic typhus agent in Ethiopian ticks. Bull. World Health Organ. 48, 563–569. Cutler, S.J., Abdissa, A., Adamu, H., Tolosa, T., Gashaw, A., 2011. Borrelia in Ethiopian ticks. Ticks Tick-Borne Dis. 3, 14–17. Cutler, S.J., Browning, P., Scott, J.C., 2006. Ornithodoros moubata, a soft tick vector for Rickettsia in East Africa? Ann. N.Y. Acad. Sci. 1078, 373–377. Duh, D., Punda-Polic, V., Avˇsiˇc-Zˇ upanc, T., Bouyer, D., Walker, D.H., Popov, V.L., Jelovsek, M., Gracner, M., Trilar, T., Bradaric, N., Kurtti, T.J., Strus, J., 2010. Rickettsia hoogstraalii sp. nov., isolated from hard- and soft-bodied ticks. Int. J. Syst. Evol. Microbiol. 60, 977–984.
344
V. Pader et al. / Ticks and Tick-borne Diseases 3 (2012) 337–344
Fournier, P.-E., Roux, V., Caumes, E., Donzel, M., Raoult, D., 1998. Outbreak of Rickettsia africae infections in participants of an adventure race in South Africa. Clin. Infect. Dis. 27, 316–323. Fournier, P.E., Belghazi, L., Robert, C., Elkarkouri, K., Richards, A.L., Greub, G., Collyn, F., Ogawa, M., Portillo, A., Oteo, J.A., Psaroulaki, A., Bitam, I., Raoult, D., 2008. Variations of plasmid content in Rickettsia felis. PLoS One 3, e2289, doi:2210.1371/journal.pone.0002289. Gracner, M., Avˇsiˇc-Zˇ upanc, T., Punda-Polic, V., Dolinsek, J., Bouyer, D., Walker, D.H., Zavala-Castro, J.E., Bradaric, N., Crocquet-Valdes, P.A., Duh, D., 2009. Comparative ompA gene sequence analysis of Rickettsia felis-like bacteria detected in Haemaphysalis sulcata ticks and isolated in the mosquito C6/36 cell line. Clinical Microbiology and Infection 15, 265–266. Hoogstraal, H., 1956. Ticks of the Sudan. In: African Ixodoidea. U.S. Government Printing Office, Washington, pp. 1–1101. Jensenius, M., Davis, X., von Sonnenburg, F., Schwartz, E., Keystone, J.S., Leder, K., Lopéz-Véléz, R., Caumes, E., Cramer, J.P., Chen, L., Parola, P., 2009. Multicenter GeoSentinel analysis of rickettsial diseases in international travelers, 1996–2008. Emerg. Infect. Dis. 15, 1791–1798. Jensenius, M., Fournier, P.E., Raoult, D., 2004. Rickettsioses and the international traveler. Clin. Infect. Dis. 39, 1493–1499. Kawabata, H., Ando, S., Kishimoto, T., Kurane, I., Takano, A., Nogami, S., Fujita, H., Tsurumi, M., Nakamura, N., Sato, F., Takahashi, M., Ushijima, Y., Fukunaga, M., Watanabe, H., 2006. First detection of Rickettsia in soft-bodied ticks associated with seabirds, Japan. Microbiol. Immunol. 50, 403–406. Letaief, A., 2006. Epidemiology of rickettsioses in North Africa. Ann. N.Y. Acad. Sci. 1078, 34–41. Macaluso, K.R., Davis, J.O.N., Alam, U., Korman, A.M.Y., Rutherford, J.S., Rosenberg, R., Azad, A.F., 2003. Spotted fever group rickettsiae in ticks from the Masai Mara region of Kenya. Am. J. Trop. Med. Hyg. 68, 551–553. Mattila, J., Burkhardt, N., Hutcheson, H., Munderloh, U., Kurtti, T., 2007. Isolation of cell lines and a rickettsial endosymbiont from the soft tick Carios capensis (Acari: Argasidae: Ornithodorinae). J. Med. Entomol. 44, 1091–1101. Mediannikov, O., Diatta, G., Fenollar, F., Sokhna, C., Trape, J.-F., Raoult, D., 2010a. Tickborne rickettsioses, neglected emerging diseases in rural Senegal. PLoS Negl. Trop. Dis. 4, e821. Mediannikov, O., Trape, J.-F., Diatta, G., Parola, P., Fournier, P.-E., Raoult, D., 2010b. Rickettsia africae, Western Africa. Emerg. Infect. Dis. 16, 571–573. Mekonnen, S., Pegram, R.G., Gebre, S., Mekonnen, A., Jobre, Y., Zewde, S., 2007. A synthesis review of ixodid (Acari:Ixodidae) and argasid (Acari: Argasidae) ticks in Ethiopia and their possible roles in disease transmission. Ethiopian Vet. J. 11, 1–24. Morita, C., El Hussein, A.R., Matsuda, E., Abdel Gabbar, K.M., Muramatsu, Y., Abdel Rahman, M.B., Eleragi, A.M., Hassan, S.M., Chitambo, A.M., Ueno, H., 2004.
Spotted fever group rickettsiae from ticks captured in Sudan. Jpn. J. Infect. Dis. 57, 107–109. Mura, A., Socolovschi, C., Ginesta, J., Lafrance, B., Magnan, S., Rolain, J.M., Davoust, B., Raoult, D., Parola, P., 2008. Molecular detection of spotted fever group rickettsiae in ticks from Ethiopia and Chad. Trans. R. Soc. Trop. Med. Hyg. 102, 945–949. Parola, P., 2006. Rickettsioses in sub-Saharan Africa. Ann. N.Y. Acad. Sci. 1078, 42–47. Parola, P., 2011. Rickettsia felis from a rare disease in the USA to a common cause of fever in sub-Saharan Africa. Clin. Microbiol. Infect. 17, 996–1000. Portillo, A., Perez-Martinez, L., Santibanez, S., Blanco, J.R., Ibarra, V., Oteo, J.A., 2007. Detection of Rickettsia africae in Rhipicephalus (Boophilus) decoloratus ticks from the Republic of Botswana, South Africa. Am. J. Trop. Med. Hyg. 77, 376–377. Pretorius, A.-M., Birtles, R.J., 2002. Rickettsia aeschlimannii: a new pathogenic spotted fever group Rickettsia, South Africa. Emerg. Infect. Dis. 8, 874. Raoult, D., Fournier, P.E., Fenollar, F., Jensenius, M., Prioe, T., de Pina, J.J., Caruso, G., Jones, N., Laferl, H., Rosenblatt, J.E., Marrie, T.J., 2001. Rickettsia africae, a tick-borne pathogen in travelers to sub-Saharan Africa. N. Engl. J. Med. 344, 1504–1510. Rehácek, J., Urvölgyi, J., Kovácová, E., 1977. Massive occurrence of rickettsiae of the spotted fever group in fowl tampan, Argas persicus, in the Armenian S.S.R. Acta Virol. 21, 431–438. Roux, V., Raoult, D., 2000. Phylogenetic analysis of members of the genus Rickettsia using the gene encoding the outer-membrane protein rOmpB (ompB). Int. J. Syst. Evol. Microbiol. 50, 1449–1455. Roux, V., Rydkina, E., Eremeeva, M., Raoult, D., 1997. Citrate synthase gene comparison, a new tool for phylogenetic analysis, and its application for the rickettsiae. Int. J. Syst. Bacteriol. 47, 252–261. Socolovschi, C., Mediannikov, O., Sokhna, C., Tall, A., Diatta, G., Bassene, H., Trape, J.F., Raoult, D., 2010. Rickettsia felis-associated uneruptive fever. Senegal. Emerg Infect Dis 16, 1140–1142. Stenos, J., Graves, S.R., Unsworth, N.B., 2005. A highly sensitive and specific real-time PCR assay for the detection of spotted fever and typhus group Rickettsiae. Am. J. Trop. Med. Hyg. 73, 1083–1085. Stephany, D., Buffet, P., Rolain, J.M., Raoult, D., Consigny, P.H., 2009. Rickettsia africae infection in man after travel to Ethiopia. Emerg. Infect. Dis. 15, 1867–1869. Tamura, K., Peterson, D., Peterson, N., Stecher, G., Nei, M., Kumar, S., 2011. MEGA5: Molecular Evolutionary Genetics Analysis using Maximum Likelihood, Evolutionary Distance, and Maximum Parsimony Methods. Mol. Biol. Evol., http://dx.doi.org/10.1093/molbev/msr121. Vitorino, L., Chelo, I.M., Bacellar, F., Ze-Ze, L., 2007. Rickettsiae phylogeny: a multigenic approach. Microbiology 153, 160–168.