Volume 16, Number 3—March 2010
Borrelia, Ehrlichia, and Rickettsia spp. in Ticks Removed from Persons, Texas, USA
Highlight and copy the desired format.
|EID||Williamson PC, Billingsley PM, Teltow GJ, Seals JP, Turnbough MA, Atkinson SF, et al. Borrelia, Ehrlichia, and Rickettsia spp. in Ticks Removed from Persons, Texas, USA. Emerg Infect Dis. 2010;16(3):441-446. https://dx.doi.org/10.3201/eid1603.091333|
|AMA||Williamson PC, Billingsley PM, Teltow GJ, et al. Borrelia, Ehrlichia, and Rickettsia spp. in Ticks Removed from Persons, Texas, USA. Emerging Infectious Diseases. 2010;16(3):441-446. doi:10.3201/eid1603.091333.|
|APA||Williamson, P. C., Billingsley, P. M., Teltow, G. J., Seals, J. P., Turnbough, M. A., & Atkinson, S. F. (2010). Borrelia, Ehrlichia, and Rickettsia spp. in Ticks Removed from Persons, Texas, USA. Emerging Infectious Diseases, 16(3), 441-446. https://dx.doi.org/10.3201/eid1603.091333.|
Data regarding the type, frequency, and distribution of tick-borne pathogens and bacterial agents are not widely available for many tick species that parasitize persons in the southern United States. We therefore analyzed the frequency and identity of pathogens and bacterial agents in ticks removed from humans and subsequently submitted to the Texas Department of State Health Services, Zoonosis Control Program, from October 1, 2004, through September 30, 2008. The data showed associations of bacterial agents and potential vectors. Tick-related illnesses may pose unidentified health risks in areas such as Texas, where incidence of human disease related to tick bites is low but well above zero and where ticks are not routinely suspected as the cause of disease. Cause, treatment, and prevention strategies can be better addressed through collecting sufficient data to establish baseline assessments of risk.
Data concerning the full distribution of tick-borne agents and their potential relationship to both emerging and characterized illnesses in the southern United States are not widely available. Persons who become ill after a tick bite may be at increased risk because a tick bite may not be considered as the source of the pathogen and because of the length of time that febrile illness may elude effective treatment. Detailed knowledge of the causative agents, their distribution, and their relationship to potential vectors is also lacking. Most tick survey data for microorganisms in the genera Borrelia, Rickettsia, and Ehrlichia have been collected in areas where the associated diseases are considered endemic. Lyme disease, Rocky Mountain spotted fever, or human monocytotropic ehrlichiosis are not considered to be endemic to Texas. Studies of microorganisms carried in ticks in non–disease-endemic areas might provide information about potentially pathogenic organisms, their vectors, and reservoirs. These data might also provide an opportunity to examine the ecology of emerging zoonoses for which different ecologic determinants for disease transmission may be present.
In 2000, the 77th Texas Legislature Subcommittee on Administration prepared a report addressing the potentially severe nature of tick-borne disease in Texas. As of October 1, 2004, the Tick-Borne Disease Research Laboratory at the University of North Texas Health Science Center (UNTHSC) became the primary facility for testing ticks submitted to the Texas Department of State Health Services (TX DSHS).
From October 1, 2004, through September 30, 2008, tick specimens were submitted to UNTHSC through the Zoonosis Control Program of the TX DSHS. Only ticks that had been attached to a person were submitted to UNTHSC, where they were screened for the genera Borrelia, Rickettsia, and Ehrlichia with genus-specific PCRs. Ticks were identified to the species level by TX DSHS entomologists before being transferred to UNTHSC (1–3). Poor condition of some specimens made identification by morphologic examination difficult. Unidentified ticks were conclusively identified by molecular methods developed at UNTHSC, which used amplification of 12S rDNA (Table 1) and sequence determination (data not shown). Additionally, the identity of any tick containing an organism not previously reported in that species was also confirmed by the same molecular methods. Of all tick specimens, 10% were screened by the same molecular identification technique to verify the accuracy of morphologic identification.
Ticks were bisected laterally by using aseptic technique and a sterile scalpel blade. For independent verification of results, half of each tick was stored in 100% ethanol at –80°C. For larvae and nymphs, the entire tick was used for DNA extraction. Total DNA was isolated from the second half by using an E.N.Z.A. Mollusc DNA Isolation Kit (Omega Bio-Tek, Inc., Norcross, GA, USA) according to manufacturer’s recommended protocol. Extracted DNA was subjected to PCR that used primers for the amplification of the tick’s 12S rDNA or Borrelia spp., Ehrlichia spp., and Rickettsia spp. genes (Table 1).
The locations of PCR setup and PCR product handling were physically separated. Reaction setup was performed in a class II type B2 biological safety cabinet that had been cleaned with 0.6% sodium hypochlorite daily and UV irradiated for 30 min before and after each use. To minimize risk for contamination, pipettor sets were dedicated to specific functions, i.e., reagent dispensing, template isolation, PCR setup, and template handling. Certified DNA/RNase-free filter barrier tips were used to prevent aerosol contamination. PCR setup was never performed in the presence of isolation materials, and reagent handling was separated both physically and temporally from templates. PCR assays were performed in duplicate with appropriate controls.
A typical, initial PCR was performed in a 25-µL reaction volume by using 5 pmol/L of each appropriate primer in conjunction with a final reaction concentration of 1× GeneAmp PCR Buffer II (Applied Biosystems, Foster City, CA, USA), 160 ng/μL bovine serum albumin, 1.0 mmol/L MgCl2, 200 μmol/L of each dNTP, 1.25 U Amplitaq (Applied Biosystems), and 10 µL of template. To establish the species of the tick specimen, we amplified 12S rDNA with the following cycle parameters: 95°C for 5 min; then 40 cycles each consisting of 95°C for 30 s, 45°C for 30 s, 72°C for 60 s; and a final 72°C extension for 5 min. Thermal cycling parameters for the initial PCRs of bacterial genes were 95°C for 5 min; then 40 cycles each consisting of 95°C for 60 s, 55°C for 60 s, 72°C for 30 s; and a final 72°C extension for 5 min. Nested PCR was performed by using the same reaction setup and 1.0 µL of amplified PCR mix as template. Nested PCR setup was performed in a dedicated dead air space cabinet that had been decontaminated in the same manner as the class II type B biosafety cabinet. The thermal cycling profile for the nested reactions was 95°C for 5 min; then 30 cycles each consisting of 95°C for 60 s, 55°C for 60 s, 72°C for 60 s; and a final 72°C extension for 5 min.
Verification of amplification was performed by agarose gel electrophoresis, followed by staining with 1X SYBR Green I (BioWhittaker Molecular Applications ApS, Rockland, ME, USA). Amplicons were examined with a UVP EC3 Imaging System (UVP, LLC, Upland, CA, USA) and subsequently analyzed by VisionworksLS Image Acquisition and Analysis Software (UVP, LLC). Secondary PCR systems (Table 1) were used to confirm positive results and did not contain primers that would amplify control DNA commonly used in the laboratory. Unincorporated primers were removed from samples producing amplicons before sequence determination by using ExoSAP-IT (USB Corporation, Cleveland, OH, USA).
DNA sequencing was performed for both strands of the PCR amplicons by using a BigDye Terminator Cycle Sequencing Kit, version 3.1 (Applied Biosystems). Unincorporated dye terminators were removed before electrophoresis by using Performa DTR Gel Filtration Cartridges (Edge BioSystems, Gaithersburg, MD, USA). Capillary electrophoresis was performed by using an ABI PRISM 310 Genetic Analyzer (Applied Biosystems). Final sequence analysis and editing was performed by using Sequencer 4.7 (Gene Codes Corporation, Ann Arbor, MI, USA). Using BLASTN, version 2.2.10 (www.ncbi.nlm.nih.gov/blast/Blast.cgi), we then compared edited sequence data with genetic sequences from characterized examples of Borrelia spp., Rickettsia spp., and Ehrlichia spp. published in GenBank.
A total of 903 ticks, representing 11 tick species, were submitted to UNTHSC from 138 of 254 Texas counties. Of these, 144 ticks contained the DNA of at least 1 of the agents in the genera Borrelia, Ehrlichia, or Rickettsia (Table 2). The most common tick species submitted were Amblyomma americanum, followed by Dermacentor variabilis. Spotted fever group Rickettsia spp. (SFGR) were the most common bacteria detected. Genetic material from SFGR was identified in A. americanum, A. cajennense, D. variabilis, Ixodes scapularis, and Rhipicephalus sanguineus ticks. Of all tick species submitted, minimum SFGR infection rates (MIRs) were highest for A. americanum (20.98%) and D. variabilis (47.37%) ticks. The most predominant SFGR sequences amplified were identical to those of Candidatus Rickettsia amblyommii (AY062007). Some contained a single-nucleotide difference relative to AY062007 (data not shown). SFGR amplicons produced from Ixodes spp. ticks were identical to those of I. scapularis endosymbiont isolates (EU544296, EF689740, EF689737) and shared >99% identity with Candidatus Rickettsia cooleyi (AF031535) (14) or an uncharacterized rickettsial endosymbiont previously reported for I. scapularis (AB002268) ticks (15). Amplicons with a DNA sequence identical to that of R. parkeri strains (U43802) (16), (EF102238) (17), and (FJ986616) were produced by 4 D. variabilis and 1 Rh. sanguineus tick samples. Amplicons identical to R. peacockii (CP001227) were produced by 2 A. americanum, 2 D. variabilis, and 1 I. scapularis tick samples. Amplicons identical to R. rhipicephali (U43803) and at least 99% similar to other R. rhipicephali strains (EU109175, EU109177, EU109178) (18) were produced by 1 Rh. sanguineus tick sample.
DNA sequences consistent with those of Borrelia spp. were derived from A. americanum, A. cajennense, D. variabilis, and I. scapularis ticks. The most commonly encountered Borrelia genetic material demonstrated at least 99% sequence identity or was identical to that of previously sequenced Candidatus Borrelia lonestari isolates (AY850063, AF538852) (19). Additionally, a borreliae flaB sequence was generated from 1 D. variabilis tick, which had 94% sequence similarity with that of Candidatus Borrelia texasensis (AF264901) (20) and sequences amplified from an uncultured Borrelia sp. from the bat tick Carios kelleyi (EF688577, EF688579) (21) and (EU492387). The flaB sequence contained 11 single-nucleotide polymorphisms relative to the corresponding section of AF264901. The Borrelia sp. 16S rDNA sequence generated from the same D. variabilis tick was also identical to that published for Candidatus B. texasensis (AF467976) (20,22). This tick was submitted from Webb County, the same Texas county from which the borreliae that produced GenBank sequence AF264901 were isolated. A single I. scapularis specimen produced the flaB sequence, which had 99% identity with B. burgdorferi (AE000783) (23).
Genetic data consistent with those from Ehrlichia spp. were observed for A. americanum, A. cajennense, and A. maculatum ticks. Amplicons produced from A. americanum and A. maculatum ticks were 99% similar to the homologous region of the E. chaffeensis disulfide oxidoreductase gene (dsb) sequences in GenBank (CP000236) (24). A single sample from A. cajennense ticks produced a DNA sequence that was 97% similar to that of the CP000236 sequence and contained 8 single-nucleotide polymorphisms relative to the similar sequence. Several single-nucleotide polymorphisms locations are at the same position as nucleotide differences identified between the dsb gene of E. ewingii (AY428950) (25) and E. canis (AF403710) (26). The nucleotide polymorphisms found within the dsb gene did not change the predicted amino acid sequence in relation to E. chaffeensis (data not shown).
By screening a diverse group of Texas tick species for a range of microorganisms and potential pathogens, we identified several novel associations: Candidatus B. lonestari in A. cajennense ticks, E. chaffeensis in A. cajennense ticks, and A. maculatum, and R. parkeri in D. variabilis ticks (Table 3). Because the geographic distribution of diseases caused by the agents is generally characterized by the distribution of the tick vectors, these findings provide insights regarding the distributions and endemicity of several potential emerging tick-borne agents.
SFGR were the most commonly observed agents in this survey. Both Candidatus R. amblyommii and Candidatus R. cooleyi are not well studied and are of undetermined pathogenicity. Current average SFGR seropositivity in Texas residents is also unknown, yet prior estimates indicate that it is higher than would be assumed from cases of Rocky Mountain spotted fever reported to the TX DSHS (27). Transmission through blood products has been noted previously (28,29). Unreported subclinical infections might cause concern about local blood products and could potentially compromise immunodeficient transfusion recipients. Additionally, detection of R. amblyommii in questing A. americanum larvae suggests transovarial transmission of the microbe, and the likelihood of pathogen transmission by larvae could be magnified by their habit of mass attack (huge numbers on a single host).
An overall Borrelia spp. MIR of 1.1% was observed for the entire 4-year collection. Prevalence of Candidatus B. lonestari in ticks from Texas was low. However, Candidatus B. lonestari sequences were detected in A. americanum ticks regardless of geographic origin. The MIR was slightly higher for A. americanum (2.53%) ticks during periods when that tick was the most abundant species parasitizing humans (October 1, 2007 through October 1, 2008). These rates are within ranges previously established in the literature (30–32). A single isolate of Candidatus B. lonestari was observed in A. cajennense ticks. This represents the potential for Candidatus B. lonestari to use hard ticks of species other than A. americanum in its maintenance cycle and suggests that Candidatus B. lonestari may occur in areas outside the natural distribution of A. americanum ticks. An MIR of 1.3% for Borrelia spp. was found for in D. variabilis and may indicate the presence of uncharacterized borreliae strains in Texas tick populations.
A. cajennense ticks have been associated with E. ruminantium (33) and spotted fever group Rickettsia spp. (34). According to seropositivity in a human population in Argentina, these ticks have also been suspected of transmitting ehrlichiosis (35). The presence of E. chaffeensis in an A. cajennense tick seems novel. Long et al. (13) suggest an E. ewingii MIR of 7.6% in southcentral Texas A. americanum ticks. Similar results for Ehrlichia spp. in A. cajennense tick populations may be plausible.
Screening ticks for a range of bacterial agents has provided several additional associations. These findings provide insights regarding the distributions and endemicity of potentially pathogenic and emerging tick-borne agents. Some of these tick-borne agents may pose an unknown health risk. Because of the wide distribution of these ticks, accurate assessments of the frequency of bacterial agents in these tick populations, their potential for causing human disease, and the ability for these tick species to act as competent vectors are warranted. Continued study and monitoring will play a vital role in public health assessment for related disease risks.
Dr Williamson is an assistant professor in the Institute of Investigative Genetics at the University of North Texas Health Science Center and director for the Tick-borne Disease Research Laboratory in the Center for Biosafety and Biosecurity. His primary research focus is the development of methods and tools for rapid assessment of disease outbreak and the study of efficient mechanisms for researching the epidemiology, genetics, and associated clinical manifestations of emerging pathogens.
We thank Chris Paddock and Bruce Budowle for their review of the manuscript.
This project was supported by the State of Texas.
- United States Department of Agriculture. Ticks of veterinary importance. Agriculture Handbook no. 485. Rockville (MD): The Department; 1976. p. 21–35.
- Cooley RA, Kohls GM. The genus Ixodes in North America. Washington: US Government Printing Office; 1945. p. 7–11.
- Keirans JE, Litwak TR. Pictorial key to the adults of hard ticks, family Ixodidae (Ixodida: Ixodoidea), east of the Mississippi River. J Med Entomol. 1989;26:435–48.
- Barbour AG, Maupin GO, Teltow GJ, Carter CJ, Piesman J. Identification of an uncultivable Borrelia species in the hard tick Amblyomma americanum: possible agent of a Lyme disease-like illness. J Infect Dis. 1996;173:403–9.
- Regnery RL, Spruill CL, Plikaytis BD. Genotypic identification of rickettsiae and estimation of intraspecies sequence divergence for portions of two rickettsial genes. J Bacteriol. 1991;173:1576–89.
- Eremeeva M, Yu X, Raoult D. Differentiation among spotted fever group rickettsiae species by analysis of restriction fragment length polymorphism of PCR-amplified DNA. J Clin Microbiol. 1994;32:803–10.
- Kollars TM Jr, Kengluecha A. Spotted fever group Rickettsia in Dermacentor variabilis (Acari: Ixodidae) infesting raccoons (Carnivora: Procyonidae) and opossums (Marsupialia: Didelphimorphidae) in Tennessee. J Med Entomol. 2001;38:601–2.
- Webb L, Carl M, Malloy DC, Dasch GA, Azad AF. Detection of murine typhus infection in fleas by using the polymerase chain reaction. J Clin Microbiol. 1990;28:530–4.
- Doyle CK, Labruna MB, Breitschwerdt EB, Tang YW, Corstvet RE, Hegarty BC, Detection of medically important Ehrlichia by quantitative multicolor TaqMan real-time polymerase chain reaction of the dsb gene. J Mol Diagn. 2005;7:504–10.
- Dawson JE, Stallknecht DE, Howerth EW, Warner C, Biggie K, Davidson WR, Susceptibility of white-tailed deer (Odocoileus virginianus) to infection with Ehrlichia chaffeensis, the etiologic agent of human ehrlichiosis. J Clin Microbiol. 1994;32:2725–8.
- Dawson JE, Biggie KL, Warner CK, Cookson K, Jenkins S, Levine JF, Polymerase chain reaction evidence of Ehrlichia chaffeensis, an etiologic agent of human ehrlichiosis, in dogs from southeast Virginia. Am J Vet Res. 1996;57:1175–9.
- Anderson BE, Sumner JW, Dawson JE, Tzianabos T, Greene CR, Olson JG, Detection of the etiologic agent of human ehrlichiosis by polymerase chain reaction. J Clin Microbiol. 1992;30:775–80.
- Long SW, Pound JM, Yu XJ. Ehrlichia prevalence in Amblyomma americanum, central Texas. Emerg Infect Dis. 2004;10:1342–3.
- Billings AN, Teltow GJ, Weaver SC, Walker DH. Molecular characterization of a novel Rickettsia species from Ixodes scapularis in Texas. Emerg Infect Dis. 1998;4:305–9.
- Weller SJ, Baldridge GD, Munderloh UG, Noda H, Simser J, Kurtti TJ. Phylogenetic placement of rickettsiae from the ticks Amblyomma americanum and Ixodes scapularis. J Clin Microbiol. 1998;36:1305–17.
- Roux V, Fournier PE, Raoult D. Differentiation of spotted fever group rickettsiae by sequencing and analysis of restriction fragment length polymorphism of PCR-amplified DNA of the gene encoding the protein rOmpA. J Clin Microbiol. 1996;34:2058–65.
- Silveira I, Pacheco RC, Szabo MP, Ramos HG, Labruna MB. Rickettsia parkeri in Brazil. Emerg Infect Dis. 2007;13:1111–3.
- Wikswo ME, Hu R, Dasch GA, Krueger L, Arugay A, Jones K, Detection and identification of spotted fever group rickettsiae in Dermacentor species from southern California. J Med Entomol. 2008;45:509–16.
- Moore VA IV, Varela AS, Yabsley MJ, Davidson WR, Little SE. Detection of Borrelia lonestari, putative agent of southern tick-associated rash illness, in white-tailed deer (Odocoileus virginianus) from the southeastern United States. J Clin Microbiol. 2003;41:424–7.
- Lin T, Gao L, Seyfang A, Oliver JH Jr. ‘Candidatus Borrelia texasensis’ from the American dog tick Dermacentor variabilis. Int J Syst Evol Microbiol. 2005;55:685–93.
- Gill JS, Ullmann AJ, Loftis AD, Schwan TG, Raffel SJ, Schrumpf ME, Novel relapsing fever spirochete in bat tick. Emerg Infect Dis. 2008;14:522–3.
- Lin T, Oliver JH Jr, Gao L. Comparative analysis of Borrelia isolates from southeastern USA based on randomly amplified polymorphic DNA fingerprint and 16S ribosomal gene sequence analyses. FEMS Microbiol Lett. 2003;228:249–57.
- Fraser CM, Casjens S, Huang WM, Sutton GG, Clayton R, Lathigra R, Genomic sequence of a Lyme disease spirochaete, Borrelia burgdorferi. Nature. 1997;390:580–6.
- Hotopp JC, Lin M, Madupu R, Crabtree J, Angiuoli SV, Eisen JA, Comparative genomics of emerging human ehrlichiosis agents. PLoS Genet. 2006;2:e21.
- Labruna MB, McBride JW, Camargo LM, Aguiar DM, Yabsley MJ, Davidson WR, A preliminary investigation of Ehrlichia species in ticks, humans, dogs, and capybaras from Brazil. Vet Parasitol. 2007;143:189–95.
- McBride JW, Ndip LM, Popov VL, Walker DH. Identification and functional analysis of an immunoreactive DsbA-like thio-disulfide oxidoreductase of Ehrlichia spp. Infect Immun. 2002;70:2700–3.
- Taylor JP, Tanner WB, Rawlings JA, Buck J, Elliott LB, Dewlett HJ, Serological evidence of subclinical Rocky Mountain spotted fever infections in Texas. J Infect Dis. 1985;151:367–9.
- Arguin PM, Singleton J, Rotz LD, Marston E, Treadwell TA, Slater K, An investigation into the possibility of transmission of tick-borne pathogens via blood transfusion. Transfusion-associated Tick-borne Illness Task Force. Transfusion. 1999;39:828–33.
- McQuiston JH, Childs JE, Chamberland ME, Tabor E. Transmission of tick-borne agents of disease by blood transfusion: a review of known and potential risks in the United States. Transfusion. 2000;40:274–84.
- Stromdahl EY, Williamson PC, Kollars TM Jr, Evans SR, Barry RK, Vince MA, Evidence of Borrelia lonestari DNA in Amblyomma americanum (Acari: Ixodidae) removed from humans. J Clin Microbiol. 2003;41:5557–62.
- Taft SC, Miller MK, Wright SM. Distribution of borreliae among ticks collected from eastern states. Vector Borne Zoonotic Dis. 2005;5:383–9.
- Mixson TR, Campbell SR, Gill JS, Ginsberg HS, Reichard MV, Schulze TL, Prevalence of Ehrlichia, Borrelia, and rickettsial agents in Amblyomma americanum (Acari: Ixodidae) collected from nine states. J Med Entomol. 2006;43:1261–8.
- Mahan SM, Peter TF, Simbi BH, Kocan K, Camus E, Barbet AF, Comparison of efficacy of American and African Amblyomma ticks as vectors of heartwater (Cowdria ruminantium) infection by molecular analyses and transmission trials. J Parasitol. 2000;86:44–9.
- de Lemos ER, Machado RD, Coura JR, Guimaraes MA, Freire NM, Amorim M, Epidemiological aspects of the Brazilian spotted fever: seasonal activity of ticks collected in an endemic area in São Paulo, Brazil. Rev Soc Bras Med Trop. 1997;30:181–5.
- Ripoll CM, Remondegui CE, Ordonez G, Arazamendi R, Fusaro H, Hyman MJ, Evidence of rickettsial spotted fever and ehrlichial infections in a subtropical territory of Jujuy, Argentina. Am J Trop Med Hyg. 1999;61:350–4.
- Table 1. Nucleotide sequence of primers used for PCR screening of tick specimens removed from humans, Texas, October 1, 2004, to September 30, 2008
- Table 2. Number and identity of ticks submitted to University of North Texas Health Science Center by the Texas Department of State Health Services Zoonosis Control Program, October 1, 2004, to September 30,...
- Table 3. No. ticks containing bacterial DNA sequences, Texas, October 1, 2004, to September 30, 2008
Please use the form below to submit correspondence to the authors or contact them at the following address:
Phillip C. Williamson, University of North Texas Health Science Center, Institute of Investigative Genetics, 3500 Camp Bowie Blvd, Fort Worth, TX 76107-2699, USA
Comment submitted successfully, thank you for your feedback.
The conclusions, findings, and opinions expressed by authors contributing to this journal do not necessarily reflect the official position of the U.S. Department of Health and Human Services, the Public Health Service, the Centers for Disease Control and Prevention, or the authors' affiliated institutions. Use of trade names is for identification only and does not imply endorsement by any of the groups named above.
- Page created: December 14, 2010
- Page last updated: December 14, 2010
- Page last reviewed: December 14, 2010
- Centers for Disease Control and Prevention,
National Center for Emerging and Zoonotic Infectious Diseases (NCEZID)
Office of the Director (OD)