Isolation and molecular characterization of a

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Original article

Isolation and molecular characterization of a relapsing fever Borrelia recovered from Ornithodoros rudis in Brazil ⁎

Sebastián Muñoz-Leala, , Álvaro A. Faccini-Martínezb,c, Francisco B. Costad, Arlei Marcilia,e, Eric T.K.C. Mesquitaf, Edwaldo P. Marques Jr.f, Marcelo B. Labrunaa a Departamento de Medicina Veterinária Preventiva e Saúde Animal, Faculdade de Medicina Veterinária e Zootecnia, Universidade de São Paulo, Av. Prof. Orlando Marques de Paiva, 87, Cidade Universitária, São Paulo, SP, 05508-270, Brazil b Programa de Pós-graduação em Doenças Infecciosas, Centro de Ciências da Saúde, Universidade Federal do Espírito Santo, Vitória, ES, Brazil c Comité de Medicina Tropical, Zoonosis y Medicina del Viajero, Asociación Colombiana de Infectología, Bogotá, Colombia d Departamento de Patologia, Universidade Estadual do Maranhão (UEMA), Cidade Universitária Paulo VI, s/n, São Luís/MA, 65055-970, São Luís, MA, Brazil e Mestrado em medicina e bem estar animal, Universidade Santo Amaro, Av. Prof. Eneas de Siqueira Neto, 340, São Paulo, 04529-300, Brazil f Agência Estadual de Defesa Agropecuária do Maranhão, São Luís, Maranhão, MA, Brazil

A R T I C LE I N FO

A B S T R A C T

Keywords: Ornithodoros rudis Borrelia venezuelensis Tick-borne relapsing fever Brazil South America

In South America, early reports from more than 50 years ago incriminated Ornithodoros rudis as vector of Borrelia venezuelensis, an agent of tick-borne relapsing fever (TBRF). Herein we report the rediscovery of O. rudis by means of morphological, biological and molecular analyses, which also comprise the first report of this tick species in Brazil. Phylogenetic analysis using partial fragments of mitochondrial 16S rRNA gene suggested that O. rudis forms a monophyletic group with Ornithodoros erraticus. By using laboratory rodents as hosts, we isolated a relapsing fever Borrelia from an infected O. rudis female. Phylogenetic analysis inferred from the rrs, flaB, and glpQ genes of Borrelia spp. placed the spirochete harbored by O. rudis closely related to Borrelia turicatae. Until further genetic evidence is not obtained we are referring to this O. rudis spirochete as B. venezuelensis. This is the first in vitro isolation of a TBRF Borrelia from South America. The presence of O. rudis in Brazil should not be overlooked, since this tick has been historically implicated in human cases of TBRF in Colombia, Panama, and Venezuela. This study provides new reports of O. rudis and B. venezuelensis after decades of scientific silence on these agents.

1. Introduction Interest on the study of tick-borne relapsing fever (TBRF) spirochetes in South America started with medical research on severe febrile cases reported from workers at Muzo mines, Colombia, during 1906 and 1907 (Franco et al., 1911). In seek for confirming their diagnoses on spirochete-like extracellular parasites observed in the blood of patients during febrile relapses, R. Franco and his colleagues sent blood smears to M. R. Blanchard from the Paris Academy of Medicine, who determined that the implicated microorganisms indeed were spirochetes, and proposed a priori that Ornithodoros turicata Dugès, 1876 were acting as the main, if not the only, vector to humans (Franco et al., 1911). Some years later, Brumpt (1922) identified Treponema venezuelense from Ornithodoros ticks collected in Venezuela, and stated that the tick involved in the transmission of this agent, previously recognized in this country (Pino-Pou, 1920) and in Colombia (Franco et al., 1911), was its newly described species, Ornithodoros venezuelensis



Brumpt, 1921 (published as “Ornithodorus venezuelensis”). Brumpt (1921) described this species as morphologically similar to Ornithodoros talaje (Guérin-Méneville, 1849); however, strikingly different in its biology, because larval stages exhibited a rapid feeding behavior (several minutes to few hours), and because first nymphal instar the needed a blood meal to molt to the next developmental stage (Brumpt, 1922). Based on these biological characters, the occurrence of this vector in Panama was confirmed by Dunn (1927a), who not only associated this tick with human relapsing fever cases, but also recognized that it was often misidentified as O. talaje (Dunn, 1927b). Meanwhile, before 1930 eight species of Ornithodoros were known to occur in South America, and following Neumann’s and Nuttall’s works on tick taxonomy (Neumann, 1901; Nuttall et al., 1908), scientists assumed that Ornithodoros rudis Karsch 1880 was a synonym of O. talaje. It was not until early-collected O. venezuelensis from Colombia were sent for examination to Germany, that professor P. Schulze stated that these ticks were morphologically indistinguishable from O. rudis (Osorno-Mesa, 1940).

Corresponding author. E-mail address: [email protected] (S. Muñoz-Leal).

https://doi.org/10.1016/j.ttbdis.2018.03.008 Received 14 December 2017; Received in revised form 28 February 2018; Accepted 6 March 2018 1877-959X/ © 2018 Elsevier GmbH. All rights reserved.

Please cite this article as: Muñoz-Leal, S., Ticks and Tick-borne Diseases (2018), https://doi.org/10.1016/j.ttbdis.2018.03.008

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attached to the skin. Detaching time was then annotated for each larva, thereafter all specimens were individually placed back into the dark incubator (25 °C, 80% RH), and observed for molting to first nymphal instar. Daily checks extended till four weeks in order to verify a possible molting to second nymphal instar without a blood meal. After this period, nymphs were fed on new mice and monitored until molting.

This phenotypic similarity was subsequently corroborated for Brumpt’s and Dunn’s material collected in Colombia, Panama, Paraguay and Venezuela (Cooley and Kohls, 1944). Although the systematic problem concerning the implicated vector was solved, the identity of its transmitted spirochete was still outstanding. After the inquiries of Davis (1955), it has been accepted that O. rudis is the vector of Borrelia venezuelensis, and that this tick was involved in early cases of TBRF in Colombia, Panama, and Venezuela (Dunn, 1927a, 1927b; Osorno-Mesa, 1940). While seminal work on the epidemiology of Ornithodoros ticks and their associated spirochetes performed during the first half of 20th century demonstrated that agents of TBRF did occur in South America (Franco et al., 1911; Pino-Pou, 1920; Brumpt, 1922; Dunn, 1927b), current interest to continue medical research on this topic remains in a stand by status. Discounting the sole in vivo isolation of a Borrelia sp. (i.e., Borrelia brasiliensis) from a nymph of Ornithodoros brasiliensis from Brazil (Davis, 1952), the finding of an unidentified Borrelia sp. massively infecting peripheral blood of a bat in Colombia (Marinkelle and Grose, 1968), the seropositivity to Borrelia parkeri and Borrelia turicatae in autochthonous communities in Bolivia (Ciceroni et al., 1994), and the molecular detection of TBRF Borrelia DNA in an undetermined Ornithodoros species in that same country (Parola et al., 2011), South American scientists have deflected their attention to another range of tick-borne diseases, mainly to those associated with hard ticks (Ixodidae). In an attempt to retake the research on Ornithodoros ticks with historically medical importance in South America, in this study we describe the occurrence of O. rudis in Brazil, and the isolation and molecular characterization of an associated TBRF Borrelia from collected specimens in the northeastern region of the country.

2.1.3. Molecular tools Taxonomic identification of the ticks was complemented by a molecular analysis. For this purpose, DNA was individually extracted from one larva and two nymphs by the Guanidine Isothiocyanate and phenol/chloroform technique (Sangioni et al., 2005). For this purpose, ticks were three-times rinsed with ethanol and ultrapure water, and smashed inside 1.5-mL microtubes. The pelleted DNA was eluted in 10–40 mL of TE buffer. A PCR targeting a fragment of ≈460 bp of the tick mitochondrial 16S rRNA gene was performed following Mangold et al. (1998). Amplicons of expected size were prepared for sequencing using Big Dye Terminator Cycle Sequencing kit (Applied Biosystems, Foster City, CA, USA), and sequenced in an ABI automated sequencer (Applied Biosystems/Thermo Fisher Scientific, model ABI 3500 Genetic Analyzer, Foster City, CA, USA) with the same primers used for PCR. Obtained sequences were assembled, and primer-trimmed with Geneious R9 (Kearse et al., 2012), and submitted to a BLAST analysis (www.ncbi.nlm.nih.gov/blast) to infer closest identities with congeneric ticks (Altschul et al., 1990). 2.2. Spirochetes 2.2.1. Isolation Attempts to isolate spirochetes were performed using 25 field-collected ticks (15 females, and 10 males). Ticks were individually placed on the shaved venter of naïve C. callosus chemically restrained with a Ketamine/Xylazine solution, and were allowed to fully engorge. Once dropped, engorged specimens were placed back at the incubator. Two mL of blood were daily obtained from all 25 rodents by ventral tail vein-puncture, and observed through dark-field microscopy to detect the presence of spirochetes during the following 12 days. Rodents not presenting spirochetemia at the final of this period were considered negative. From spirochetemic animals, 1 mL of blood was collected by intracardiac puncture the second day of spirochetemia, and intraperitoneally injected into naive mice, confirming the presence of spirochetes by the above stated procedure. Samples exhibiting ≈5 × 105 spirochetes were then inoculated into 1 mL of BSK-H medium with 6% rabbit serum (Sigma-Aldrich, USA), kept in a dark incubator at 34 °C and examined every four days by dark-field microscopy at 200X. Blood cultures were performed in duplicates to maximize the probability of successful spirochete growth. When reaching 5–10 organisms per observed field, 1 mL of the culture was subsequently passed into 1 mL of fresh BSK medium. After the first passage reached at least 40 spirochetes per field, the complete medium was stored at −80 °C. Tick feeding in laboratory mice was approved by the Ethical Committee in Animal Research of the Faculty of Veterinary Medicine of the University of São Paulo (projects number 2263/2011 and 9602030214).

2. Material and methods 2.1. Ticks 2.1.1. Collection and identification Ticks of the genus Ornithodoros were collected in between nestdebris inside a hollow palm-tree in July 2017, at Riachão Municipality, Maranhão State, Cerrado Biome, Brazil (07°14¢28¢’S; 46°40¢51¢’W, 586 m). Five nymphs, 15 females and 10 males were brought alive to the laboratory. Females were individually placed in tubes and accommodated inside an incubator at 25 °C, 80% relative humidity (RH), and absolute darkness. Four engorged females were daily checked for oviposition. The third day after arrival, two females started laying eggs that culminated in hatched larvae 22 days after. Ten days after hatching, 10 unfed larvae per female were killed in hot water, clarified with 25% KOH, and mounted in slides using Hoyer’s medium in order to observe discrete morphological characters through optical microscopy (Olympus BX40 optical microscope, Olympus Optical Co. Ltd., Japan). The morphotype of slide-mounted larvae and adults was compared with descriptions of Neotropical Ornithodorinae (Karsch, 1880; Brumpt, 1922; Cooley and Kohls, 1944; Kohls et al., 1965; Jones and Clifford, 1972). 2.1.2. Biological traits Considering that Ornithodoros species historically implicated as vectors of TBRF Borrelia in South America have marked differences in biological characters, especially those concerning feeding period of larvae and molting to successive nymphal instars (Brumpt, 1922; Dunn, 1927a), four cohorts of larvae, each one obtained from four laboratory engorged females were analyzed. Each offspring was individually placed on the shaved back of a chemically restrained tick-naive vesper mouse (Calomys callosus) from an animal room of the Faculty of Veterinary Medicine of the University of São Paulo, São Paulo, Brazil, and visually monitored under a stereomicroscope (Zeiss Stemi SV 11, Zeiss, Münich, Germany). Once on the host, the beginning of the feeding period for each cohort was measured considering the first larva that

2.2.2. DNA extraction, PCR amplification and sequencing Frozen BSK cultures were brought to liquid state at 37 °C, and then centrifuged at 12,000 rpm for 20 min. Pelleted spirochetes were then eluted into 200 mL of PBS buffer. DNA extraction was performed using the DNeasy Blood and Tissue Kit (Qiagen, Valencia, CA, USA). Amplification of 16S ribosomal RNA (rrs), flagellin (flaB) and glycerophosphodiester phosphodiesterase (glpQ) encoding genes were performed by conventional PCR using primers stated elsewhere (Schwan et al., 2005). Initial denaturation began with heating at 94 °C for 3 min, followed with 35 cycles of denaturation at 94 °C for 30 s, annealing at 56 °C for 30 s, and extension at 72 °C for 3 min. An additional 7-min 2

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Fig. 1. Micrographs of unfed slide-mounted larva, and alive field-collected female of Ornithodoros rudis. Larva: (A) dorsal view, (B) dorsal plate, (C) capitulum, and (D) posterior ventral view. Female: (E) dorsal view, and (F) ventral view. Setae of larva are abbreviated as follows: Al, anterolateral; C, central; Pl, posterolateral; Ca, circumanal; Pm, posteromedian. Note that the dotted circle highlights the absence of the postcoxal pair.

was performed using MrBayes v3.1.2 (Huelsenbeck and Ronquist, 2001) with four independent Markov chain runs for 1,000.000 metropolis-coupled MCMC generations, and sampling a tree every 100th generations. The first 25% of the trees represented burn-in, and the remaining trees were used to calculate the Bayesian posterior probability. Sequences of Ixodes holocyclus Neumann, 1899 and Ixodes uriae White 1852 were used as out-groups. GenBank accession numbers of all sequences are shown in the phylogenetic tree.

extension after the 35th cycle was performed at 72 °C. DNA of Borrelia anserina strain PB (Ataliba et al., 2007) was used as positive control in all reactions. Obtained amplicons were visualized with UV light through 1.5% agarose gels stained with SYBR Safe (Thermo Fisher Scientific, Waltham, MA, USA). Products containing a single expectedsize fragment were purified with ExoSAP-IT (USB Corporation, Cleveland, OH, USA) and prepared for sequencing as above stated. Obtained sequences were assembled, trimmed and translated to amino acid (if applicable) with Geneious R9 (Kearse et al., 2012). Sequences not codifying a clean succession of amino acids after translation were discarded. Final contigs obtained for all three genes were submitted to BLAST analysis (www.ncbi.nlm.nih.gov/blast) to infer closest similarities with other spirochetes (Altschul et al., 1990).

2.3.2. Borrelia spp. Individual alignments for each gene were initially constructed with the CLUSTAL W algorithm (Thompson et al., 1994) including sequences obtained in the current study, partial sequences from Borrelia duttonii strains CR2A, 1120K3 (Toledo et al., 2010), and La (Cutler et al., 1999); Borrelia hispanica strains ORIX, and Sp2 (Toledo et al., 2010); B. parkeri strains CA216, CA218, CA219, CA220 and CA221; B. turicatae strains 95PE-570, 99PE-1807, TCB-1, TCB-2, and PE1-926 (Schwan et al., 2005), and Borrelia recurrentis strains A8, A11 and A16 (Cutler et al., 1999). Sequences recovered from complete genome records of other TBRF borreliae were also included as follows: Borrelia anserina strain BA2 (unpublished); Borrelia coriaceae strain Co53 (unpublished), Borrelia crocidurae strains Achema (Elbir et al., 2012) and DOU (unpublished); B. duttonii strain Ly (Lescot et al., 2008); Borrelia hermsii strains DAH, MTW and YBT (unpublished); Borrelia miyamotoi strain LB2001 (Hue et al., 2013); B. parkeri strains HR1 (Barbour and Campeau Miller, 2014) and SLO (unpublished); B. recurrentis strain A1 (Lescot

2.3. Phylogenetic analyses 2.3.1. Tick mitochondrial 16S rRNA gene An alignment of the obtained 16S rRNA mitochondrial DNA partial sequences with 59 homologous sequences from other Argasinae and Ornithodorinae available in GenBank was constructed using CLUSTAL X (Thompson et al., 1997), and manually adjusted with GenDoc (Nicholas et al., 1997). Phylogenetic analyses were inferred by Maximum Parsimony (MP) and Bayesian methods. The MP analysis was performed using PAUP 4.0b10 (Swofford, 2002) with 500 bootstrap replicates, random stepwise addition starting trees (with random addition of sequences), and TBR branch swapping. The Bayesian analysis 3

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3.2. Spirochetes

et al., 2008), and B. turicatae strains 91E135 (unpublished) and BTE5EL (Kingry et al., 2016). Subsequently, ambiguous alignments of rrs, flaB and glpQ genes were manually concatenated, and submitted to a phylogenetic inference. A Bayesian analysis was performed with the same software and specifications as above. Sequences of Borrelia chilensis strain VA1 (Huang et al., 2015) were selected as out-group. GenBank accession numbers of all sequences used to perform the phylogenetic analysis are specified in the resulted tree.

Overall, only one female tick was positive to borrelial infection. Spirochetes were visualized in the blood of the infected rodent at the fourth day post tick exposure, and successfully passed to a second host at the third day of spirochetemia (Video 1). Replicated spirochetes were then successfully recovered into two BSK-H media, cultured until the second passage, and frozen at 80 °C. The content of one of the frozen media was submitted to DNA extraction. Multilocus sequencing approach yielded trimmed fragments of 1450 bp, 1002 bp, and 1005 bp for the rrs, flaB, and glpQ genes, respectively. By a BLAST search, obtained sequences shared > 99% nucleotide identity with homologous sequences of B. turicatae BTE5EL (CP015629) as follows: rrs, 1449/ 1450 bp; flaB, 999/1002 bp; glpQ, 1003/1005 bp. The concatenated alignment of 2949 bp submitted to a Bayesian phylogenetic analysis confirmed this trend, since high posterior probability supported that the Borrelia sp. harbored by O. rudis (from now onwards, B. venezuelensis) forms a clade with all seven B. turicatae strains included in the analysis (Fig. 3). Although closely related, B. venezuelensis did not cluster between any of the B. turicatae strains, and rather appeared at a basal position forming a separated taxon within the clade. The characterized strain of B. venezuelensis has been named as RMA01, and its DNA sequences were deposited in GenBank under the following accession numbers: MG651649 (rrs), MG651650 (flaB), and MG651651 (glpQ).

3. Results 3.1. Ticks Morphological analyses of slide-mounted larvae allowed the identification of O. rudis based on the following discrete traits: dorsum provided with a small irregularly shaped plate, and 17–18 pairs of setae (seven anterolateral, six posterolateral and four to five central); venter provided with seven pairs and one posteromedian setae, postcoxal pair absent; hypostome apically rounded, dentition 3/3 in the distal fifth portion, then 2/2 towards the insertion, with 9–11 denticles in the first, 8–10 in the second and 3–4 in the third file (Kohls et al., 1965) (Fig. 1A, B, C, and D). Alive adults were also recognized as O. rudis by exhibiting light-brown mammillae; long-ovate body shape with parallel lateral sides; dorsoventral groove absent; barely defined dorsal discs; legs and coxae short; cheeks small, not capable of completely covering the hypostome and palps, and not reaching the ventral surface of basis capituli (Karsch, 1880; Brumpt, 1922; Cooley and Kohls, 1944) (Fig. 1E, and F). Vouchers of O. rudis were deposited in the following tick collections: “Coleção Nacional de Carrapatos Danilo Gonçalves Saraiva” (CNC, São Paulo, Brazil), accession number CNC-3628 (20 larvae mounted in individual slides); and “Coleção de Ixodidae e Argasidae do Nordeste Brasileiro” (CIANE, Maranhão, Brazil), accession number CIANE-002 (one male, one female, and one nymph). The feeding period of larvae did not exceed 180 min; engorged larvae molted to the first nymphal instar in up to nine days (Table 1). None of the first instar-nymphs molted during the period of four weeks, and avidly fed when exposed to the host after this fasting time. Only after this meal they molted to the second nymphal instar. One haplotype of 427 bp was sequenced from the larva, and two nymphs submitted to PCR targeting the tick mitochondrial 16S rRNA gene. By BLAST comparison, the three closest congeneric homologous sequences sharing highest genetic identity were Ornithodoros kohlsi Guglielmone and Keirans, 2002 (KX130783, 83.3%, 360/432 bp, 11 gaps), Ornithodoros saraivai Muñoz-Leal and Labruna, 2017 (KX812526, 82.8%, 357/431 bp, 13 gaps), and Ornithodoros peropteryx Kohls, Clifford & Jones, 1969 (KC493651, 82.2%, 356/433 bp, 12 gaps). The mitochondrial 16S rDNA sequence of O. rudis was deposited in GenBank database under the accession number MG653563. Phylogenetic analyses confirmed the genetic identity of O. rudis as a separated taxon within the Argasidae family, and pointed to a degree of relatedness with Ornithodoros erraticus. Together with Ornithodoros cavernicolous as a sister clade, O. rudis and O. erraticus formed a separated lineage within the Ornithodorinae subfamily (Fig. 2).

4. Discussion 4.1. Ornithodoros rudis This is the first report of O. rudis after a scientific gap of more than 50 years, since the last work on this species concerned the description of its larval stage in 1965 (Kohls et al., 1965). Our findings confirmed the identity of this tick by a morphological analysis of larvae and adult stages, and observations on its biology. As previously mentioned (Brumpt, 1922), larvae of O. rudis are rapid feeders, as they engorge and detach from host after several minutes to few hours (Table 1). Moreover, the first nymphal instar needs a meal to molt to the next nymphal instar (Brumpt, 1922), a behavioral trait that separates this tick from representatives of Alectorobius subgenus (Clifford et al., 1964). Previous collections of O. rudis were made in human dwellings and associated fowl (Cooley and Kohls, 1944), yet its natural habitat and hosts remained unknown. Here we report the collection of O. rudis in wilderness for the first time, and pose the possible association that this tick might have with wild avian hosts, since we extracted all the specimens in between debris of a bird nest. Still, wild rodents should not be discarded as potential hosts, since in the laboratory immature and mature stages of O. rudis rapidly attached and fed upon C. callosus. From a genetic point of view, the obtained haplotype confirms that the identified tick indeed correspond to a previously uncharacterized taxon, because it exceeds a 17% of genetic divergence with any closest relative. However, the phylogenetic position of O. rudis with O. erraticus in a monophyletic group is still discussable, since both MP and Bayesian supports were not conclusive in the trees (Fig. 2). Interestingly, larval stages of both Ornithodoros species are indeed biologically related, since they share a fast-feeding behavior (Brumpt, 1922; El Shoura, 1987). Moreover, the phenotype of both larvae are compatible with traits of Pavlovskyella subgenus sensu Kohls et al. (1965) (i.e., oddly shaped dorsal plate, blunt hypostome, absence of postcoxal pair of setae). In contrast, when noticing the morphology of adult stages, O. rudis becomes a unique species unclassifiable to subgeneric level (Hoogstraal, 1985), since males and females are provided with cheeks, and lack tarsal humps, two characters that rather match the phenotype of Alectorobius subgenus (Kohls et al., 1965). So far, O. rudis still appears as a conflictive species in terms of its taxonomic position, and this fact might be reflected in the phylogenetic analysis as well, since apart from O. erraticus, which is considered an Ornithodoros (Pavlovskyella)

Table 1 Feeding and molting periods of Ornithodoros rudis larvae. Cohorts

Female Female Female Female

No. larvae

1 2 3 4

15 29 26 22

Feeding (minutes)

Molting (days)

Mean

SD

Min–Max

First–Last

105 80 111 120

± 33.7 ± 27.7 ± 43.0 ± 30.7

54–169 34–127 51–177 72–180

7–9 7–9 7–8 6–8

SD, Standard Deviation; Min, Minimum; Max, Maximum.

4

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Fig. 2. Maximum Parsimony (MP) and Bayesian (B) inferred phylogenetic trees. Support values (MP/B) are given for major branches. The position of Ornithodoros rudis is highlighted in bold.

5

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Fig. 3. Bayesian inferred phylogenetic tree of Borrelia spp. using concatenated rrs, flaB, and glpQ genes. The position of Borrelia venezuelensis RMA01 is highlighted in bold.

B. venezuelensis as an independent sister taxon. However, whether this genetic evidence implies conspecificity with B. turicatae should be carefully assessed, and needs further analyses. Based on low divergence exhibited by rrs, flaB, and glpQ genes, the taxonomic status of B. duttonii and B. recurrentis was once questioned, and precluded a clear specific separation (Ras et al., 1996; Cutler et al., 2008;). In this case, the study of additional loci, more strains (Cutler et al., 2008), and the analysis of whole-genome sequences from both Borrelia species (Lescot et al., 2008), confirmed their taxonomic independence. Considering this scenario, and relying on historical and ecological evidence, a conservative criterion should be adopted, and treat the spirochete isolated from O. rudis as B. venezuelensis until extended genetic analyses of this and additional strains are performed. Considering historical records (Cooley and Kohls, 1944; Kohls et al., 1965), it must not be overlooked that vectors of B. turicatae and B. venezuelensis (O. turicata and O. rudis, respectively) might overlap their

representative (Kohls et al., 1965), this species also clustered into a major clade that includes O. cavernicolous, a bat-associated tick classified as an Ornithodoros (Alectorobius) species (Dantas-Torres et al., 2012). In any case, the characterization of additional genes is now needed in order to elucidate the phylogenetic position of O. rudis under higher resolution.

4.2. Borrelia venezuelensis The recovery and in vitro culture of spirochetes from an O. rudis female confirmed early research (Brumpt, 1922; Dunn, 1927b) and constitutes the first in vitro isolation of spirochetae from this tick. Phylogenetic analysis inferred from three loci (rrs, flaB, and glpQ) indicated this spirochete to be closely related to B. turicatae. Although B. venezuelensis formed a monophyletic group with B. turicatae, seven strains of this last species still clustered into a separated branch, leaving 6

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distributional ranges in Central America, and eventually share hosts, which could represent a reasonable explanation for the occurrence of phylogenetically closely related Borrelia adapted to survive in different soft tick species. From a human epidemiological perspective, although TBRF has never been reported in Brazil, clinical symptoms of this disease can be obscured by other arthropod-borne affections such as malaria (Nordstrand et al., 2007), Dengue, and Yellow fever (Dworkin et al., 2008), which are endemic in this country (Gallego et al., 2014). Moreover, persisting relapsing fever borreliosis can mimic chronic Lyme disease symptoms (Lange et al., 1991), and in this case, the use of improper diagnostic procedures could directly preclude a correct elucidation of the etiological agent. This epidemiological scenario becomes particularly important in Brazil, since 15 years of suspicions have been unsuccessful to prove that Borrelia burgdorferi sensu lato (s. l.) is the causative agent of an alleged Lyme-like disease named as the BaggioYoshinari Syndrome (Yoshinari et al., 2010). Finally, considering that B. venezuelensis-infected O. rudis does occur in the country, and that serologic methods used for the diagnosis of infections of B. burgdorferi s. l. can cross-react with borreliae of the relapsing fever group (Rath et al., 1992; Schwan et al., 1996), TBRF should be included as a differential diagnosis in Brazilian patients showing an unspecific febrile syndrome.

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