Volume 10, Number 4—April 2004
Ixodid and Argasid Tick Species and West Nile Virus
Control of West Nile virus (WNV) can only be effective if the vectors and reservoirs of the virus are identified and controlled. Although mosquitoes are the primary vectors, WNV has repeatedly been isolated from ticks. Therefore tick-borne transmission studies were performed with an ixodid (Ixodes ricinus) and an argasid tick species (Ornithodoros moubata). Both species became infected after feeding upon viremic hosts, but I. ricinus ticks were unable to maintain the virus. In contrast, O. moubata ticks were infected for at least 132 days, and the infection was maintained through molting and a second bloodmeal. Infected O. moubata ticks transmitted the virus to rodent hosts, albeit at a low level. Moreover, the virus was nonsystemically transmitted between infected and uninfected O. moubata ticks co-fed upon uninfected hosts. Although ticks are unlikely to play a major role in WNV transmission, our findings suggest that some species have the potential to act as reservoirs for the virus.
The first report of a West Nile virus (WNV) outbreak within the Western Hemisphere occurred in 1999 in New York City and resulted in human, equine, and avian deaths (1). Since 1999, WNV has been found in an additional 44 states of the United States as well as in parts of Canada, the Caribbean, and South America (2,3). During 2002 more than 4,000 people diagnosed with WNV and 284 deaths were reported in the United States (latest records available from: http://www.cdc.gov/ncidod/dvbid/westnile/index.htm).
WNV is a member of the genus Flavivirus that contains over 70 identified viruses. Most of these viruses are vectored by mosquitoes or ticks, although a few have no known vectors (4). WNV has been isolated from 43 species of mosquito in the United States (5), the most important of which is Culex pipiens (6). It has also been isolated from hard (ixodid) and soft (argasid) tick species in regions of Europe, Africa, and Asia (7–13) where WNV is endemic. Ticks rank second only to mosquitoes in their importance as vectors of human pathogens and transmit a greater variety of infectious agents than any other arthropod group (14). However, whether or not ticks are major vectors of WNV has not been adequately investigated.
Current strategies to control WNV in the United States are largely based on measures to avoid exposure and to control vector species, but at present only mosquito species are targeted by government surveillance and preventive control programs (15). Resident U.S. tick populations could also play a role in the WNV transmission cycle in the current outbreak. We investigated an argasid tick species and an ixodid tick species for their competence as vectors and reservoirs of the New York strain (NY99) of WNV.
Materials and Methods
We tested a hard tick species, Ixodes ricinus, and a soft tick species, Ornithodoros moubata, for their vector competence with WNV (NY99 strain). These species are not native to the United States and were chosen mainly for their availability. O. moubata ticks were considered potential vectors for the Eg101 strain of WNV in a study by Whitman and Aitken in 1960 (16). I. ricinus ticks are the primary vectors of Borrelia burgdorferi, the agent causing Lyme disease in Europe and are important vectors of the flaviviruses tick-borne encephalitis virus (TBEV) and Louping-ill virus (LIV) (17).
Ticks were taken from colonies reared and maintained for many generations at the Centre for Ecology and Hydrology Oxford according to standard methods (18). Colony ticks were WNV negative by reverse transcriptase–polymerase chain reaction (RT-PCR) testing (15 members of each species tested).
Virus and Viral Assays
The WNV strain used (NY99) was supplied by Robert Shope, University of Texas. High-titer mouse brain suspension stocks of WNV (2.9 x 107 PFU/mL-1) were diluted in phosphate-buffered saline (PBS) to a concentration of 105 PFU/mL-1 before use. Viral stocks and the serum samples from infected mice were tested for infectious virus by plaque assays on pig kidney epithelial cells as described previously (19), by using a 3% carboxymethylcellulose overlay.
Tick Infection and Co-feeding Transmission Experiments
Seven groups of six BALB/c mice (female, 4–6 weeks old) were injected subcutaneously with 104 PFU of WNV. Three of the mice were bled daily from the tail to follow the course of viremia by plaque assay. Two groups of mice were infested with I. ricinus nymphs (20 per mouse); one group was infested 3 days before infection, the other 4 days after infection. The other five groups of mice were infested with second instar O. moubata ticks (10 per mouse) on either the same day (day 0) or 1, 2, 3, or 4 days after infection. After the initial experiment, and to increase the number of positive ticks available for experimentation, 12 additional mice were infested with O. moubata 2 days after infection with WNV.
Ticks housed in gauze-covered neoprene feeding chambers on mice (18) were removed when fully engorged, 24 hours after infestation in the case of O. moubata ticks and 6 days after infestation in the case of I. ricinus nymphs. The engorged ticks were stored at 20°C in KCl-saturated desiccators until testing for WNV or until ready for a further bloodmeal, as indicated in Table 1. After storage, the ticks (pools and individual ticks) were homogenized in 500 μL of PBS by using plastic homogenizers under sterile conditions. The homogenates were frozen and stored at –70°C until analyzed. Tick homogenates were assayed for infectious virus antigen (by immunofluorescence assay) and viral RNA (by RT-PCR) as shown in Table 1.
Co-feeding transmission experiments were carried out by infesting clean BALB/c mice (n = 7, Harlan, UK) with 10 third instar O. moubata ticks 57 days after they had taken an infectious bloodmeal, and 10 uninfected ticks (second instar) in separate feeding chambers. The two feeding chambers were separated by at least 1 cm.
To investigate tick-to-host transmission, 10 BALB/c mice were infested with cohorts of 5, 10, 15, or 20 third instar O. moubata ticks 57 days after an infectious bloodmeal. Fifteen days after infestation, the mice (including those used for co-feeding) were euthanized; brains were removed, homogenized in 1 mL of sterile PBS, and stored at –70°C until they were tested for WNV.
Samples of tick (or mouse brain) homogenate (100 μL) were used to infect 2 x 106 C6/36 mosquito cells in a total of 3 mL L-15 medium containing 7% fetal calf serum (Gibco-BRL, Paisley, UK) in 30 mm Petri dishes that contained glass coverslips. Infected cells were incubated at 28°C for 3 days. Cells were fixed in cold acetone and treated according to standard methods (19). Infected cells were fluorescein isothiocyanate–labelled with a broadly reactive flavivirus E-protein monoclonal antibody (MAb 813) (20) or a WNV-specific monoclonal antibody (MAb 546) (21). Labelled cells were visualized by using an Olympus epifluorescence microscope. Uninfected cells were used as negative controls and cells infected with the original viral stock as positive controls. Tick samples were deemed positive when more than 10% of the cells showed specific fluorescence with both monoclonal antibodies.
Nested RT-PCR Assay
RNA was extracted from homogenized samples (100 μL) by using RNAgents total RNA extraction kit in accordance with the manufacturer’s instructions (Promega, Madison, WI). cDNA synthesis was carried out with Superscript II reverse transcriptase (Invitrogen, San Diego, CA) and 3′(1) primer (Table 2) for 50 min at 42°C, in a total volume of 20 μL. PCR was carried out on the cDNA (1 μL) by using 5′(1) and 3′(1) primers. Nested PCR was carried out on 1 μL of the first-round PCR product using the nested primers 5′(2) and 3′(2). All PCR reactions were carried out in a 50-μL volume with REDTaq DNA polymerase (Sigma Chemical Co., St. Louis, MO). A Hybaid Touchdown thermal cycler was used with the following program: 94.5°C for 1 min, 30 cycles of 94°C for 40 s, 56°C for 1 min, and 72°C for 1 min, followed by a final extension step of 72°C for 10 min. Viral stock, RNA extracted from uninfected ticks, and PBS-only samples were used as control reactions. Positive samples gave a PCR product of approximately 1.2 kbp. This method could detect RNA from a viral stock equivalent of 9 PFU (data not shown).
To confirm the identity of RT-PCR products, PCR products were gel purified with QIAquick (Qiagen, Crawley, UK) columns in accordance with manufacturer’s instructions. The purified DNA was sequenced with an ABI automatic sequencer and the nested primers 5′(2) and 3′(1) and a primer based on the internal sequence of the E gene of WNV (not shown).
BALB/c mice infected with WNV were weakly viremic 2 and 3 days after injection, with mean titers of 6 x 103 and 3 x 103 PFU/mL-1 blood respectively. After 4 days, viremia was no longer detectable by plaque assay, although severe neurologic disease developed in the mice after 5 or 6 days, and they were euthanized. O. moubata ticks that had fed on mice on days corresponding to the viremic period (i.e., days 2 and 3 after infection), but not those fed outside this period, contained viral antigen as measured by immunofluorescence assay (IFA) (Table 1). Two days after engorgement, 17% (n = 12) I. ricinus ticks that started to feed on hosts 3 days before WNV injection, but not those that had started to feed 4 days after injection, were positive for WNV RNA. When the former group of ticks was tested 28 days later, no evidence of infection was found. Infected O. moubata ticks, in contrast, maintained the virus after molting into the next instar (i.e., third instar); following a second, noninfectious bloodmeal; and after molting a second time into fourth instars. Fifty percent of the individual ticks (n = 14) tested by RT-PCR were positive for WNV RNA when examined 132 days after the initial infectious bloodmeal.
Five days after engorgement, 23% (n = 66) of uninfected second instar O. moubata ticks that had co-fed with infected cohorts of third instar ticks on noninfected mice were positive for WNV RNA (Table 1). The remaining unfed ticks (n = 15) were tested after they had molted into third instars, 45 days after co-feeding. Four of these ticks (26%) were positive for WNV RNA. The identities of the PCR products obtained from three positive samples were confirmed by sequence analysis.
Infected cohorts of O. moubata ticks (third instar) were fed on uninfected mice to investigate tick-to-host transmission. Of the 17 uninfected mice used (including mice used in co-feeding experiments), none showed clinical signs of infection. One of the brains tested, from a mouse infested with an infected cohort of 20 ticks, was positive by RT-PCR but negative when tested by IFA (Table 1). The PCR product was sequenced to confirm the identity of WNV.
Laboratory studies from the 1950s suggested that some tick species might serve as competent vectors for WNV. Hurlbut and Taylor (1956) showed that O. savignyi and O. erraticus ticks were infected after feeding on mice inoculated with the Ar-248 strain of WNV, but transmission from infected ticks to mice was not observed (22,23). Vermeil et al. (1959) infected O. maritimus and O. erraticus ticks by feeding on inoculated (Uganda 28B strain) chickens, guinea-pigs, mice, or gerbils. Infected ticks transmitted the virus to uninfected mice (24). More recently, an artificial membrane system was used to infect Argas arboreus ticks, which were then able to transmit the virus to uninfected hosts, although transstadial transmission of WNV was not observed (25,26).
Our study demonstrated that both I. ricinus and O. moubata ticks become infected with WNV (NY99 strain) through feeding on virus-infected rodent hosts, but only when these hosts were viremic (i.e., systemic transmission). Thirty days after engorgement, we no longer found any evidence of WNV infection in the I. ricinus ticks. This finding suggests that nymphs of this tick species do not support replication of the virus, and therefore are not competent vectors for WNV. By extrapolation, the closely related tick species, I. scapularis (the main U.S. Lyme disease vector) is also unlikely to be a competent vector of WNV, although this hypothesis will need to be confirmed experimentally.
In contrast, infected O. moubata ticks maintained infectious virus for at least 132 days (length of experiment), and WNV persisted transstadially through at least two developmental stages. Evidence for tick-to-host transmission of WNV was found in our study, although the level of infection observed (subclinical) makes assessing its importance without further investigation difficult. Whitman and Aitken (1960) observed much higher levels of transmission from WNV-infected (Eg101 strain) O. moubata ticks to day-old chicks but only when very high feeding densities were used (an average of 49 ticks per chick) (16). Although ticks often feed in large numbers on individual hosts (27), tick-to-host transmission appears to be very inefficient when compared to mosquito transmission of WNV (23). Consequently, this mode of transmission is unlikely to be important in the natural transmission cycle of WNV. Perhaps higher levels of infection (and therefore transmission) would be found with ticks that feed on birds, the natural reservoir hosts of WNV. Some avian species exhibit much higher (>1010 PFU/mL serum) and more prolonged viremia when infected with WNV than the mice used for this investigation (28,29). Although neither of the tick species that we tested are obligate bird feeders, I. ricinus ticks often feed on pheasants in the United Kingdom (30), and several species of Ornithodoros ticks feed almost exclusively on birds, for example, the O. capensis group of ticks that are established along the southern coast of the United States (31). As members of this group have been shown to be competent vectors for WNV (24), these ticks could represent a reservoir of the virus in the United States.
The transmission of flaviviruses such as TBEV and LIV from infected to noninfected ixodid ticks through co-feeding on nonviremic hosts (nonsystemic transmission) is a well-established phenomenon (32). Indeed, this mode of transmission is believed to play a substantial role in the epidemiology of these diseases (27). We tested for co-feeding transmission of WNV between infected and uninfected O. moubata ticks. More than 22% of the uninfected ticks were positive for WNV RNA 5 days after co-feeding. A similar percentage of ticks were positive 40 days later, after having molted to the next developmental stage. As co-fed ticks were in contact with the mice for <24 hours, this finding strongly suggests that WNV was nonsystemically transmitted between infected and uninfected ticks, since viremia had insufficient time to develop. Our study represents the first unequivocal report of co-feeding transmission by an argasid tick species. Argasid ticks, unlike ixodid ticks, typically feed for <2 hours. Vesicular stomatitis virus has been transmitted between infected and noninfected co-feeding black flies (Simulium vittatum), insects that typically feed for 4–5 min (33). Langerhans cells are believed to be the agents of viral transmission between feeding sites of infected and noninfected co-feeding hard ticks (32,34). Langerhans cells, which are susceptible to WNV infection (35), have been shown to migrate rapidly (within 2 hours) from localized antigen-stimulated epidermal sites (36). Therefore, these cells could possibly play a similar role in the co-feeding transmission of WNV by soft tick species.
Although this study is not exhaustive, it does demonstrate that tick species can become infected with the U.S. strain of WNV through feeding upon infected hosts and through co-feeding with infected ticks on noninfected hosts. In some tick species, WNV can be maintained through the transstadial stages of the tick lifecycle, and infected ticks may be capable of infecting hosts through further feeding. When compared to experimental studies with mosquito species (37–39), ticks are clearly not efficient vectors of WNV and therefore are unlikely to be important vectors for WNV in the current U.S. epidemic. However, our results demonstrate that WNV can persist for a comparatively long time in infected ticks and be transmitted between vertebrate hosts; this finding suggests a reservoir potential of ticks for WNV that justifies further investigation.
Dr. Lawrie is currently a postdoctoral researcher working in the Nuffield Department of Clinical Laboratory Sciences, University of Oxford. His research interests include identifying and characterizing cancer-associated antigens that are recognized by autologous antibody responses, molecular aspects of the tick-host interface, and transmission of flaviviruses in tick species.
- Lanciotti RS, Roehrig JT, Deubel V, Smith J, Parker M, Steele K, Origin of the West Nile virus responsible for an outbreak of encephalitis in the northeastern United States. Science. 1999;286:2333–7.
- Centers for Disease Control and Prevention. Provisional surveillance summary of the West Nile virus epidemic—United States, January-November 2002. MMWR Morb Mortal Wkly Rep. 2002;51:1129–33.
- Aiken L. Health Canada “nearly blindsided” by West Nile virus incidence. CMAJ. 2003;168:756.
- Monath TP, Heinz FX. Flaviviruses. In: Fields BN, Knipe DM, Howley PM, editors. Fields virology. New York: Lippincott-Raven; 1996. p. 961–1034.
- Centers for Disease Control and Prevention. West Nile virus—entomology [monograph on the Internet]. 2002 [cited 2004 Feb 11]. Available from: http://www.cdc.gov/ncidod/dvbid/westnile/mosquitospecies.htm
- Campbell GL, Marfin AA, Lanciotti RS, Gubler DJ. West Nile virus. Lancet Infect Dis. 2002;2:519–29.
- Platonov AE. West Nile encephalitis in Russia 1999–2001: Were we ready? Are we ready? Ann N Y Acad Sci. 2001;951:102–16.
- Mathiot CC, Georges AJ, Deubel V. Comparative analysis of West Nile virus strains isolated from human and animal hosts using monoclonal antibodies and cDNA restriction digest profiles. Res Virol. 1990;141:533–43.
- Iakimenko VV, Bogdanov II, Tagil’tsev AA, Drokin DA, Kalmin OB. The characteristics of the relationships of arthropods of the refuge complex with the causative agents of transmissible viral infections in bird rookeries. Parazitologiia. 1991;25:156–62.
- L’Vov DK, Dzharkenov AF, L’Vov DN, Aristova VA, Kovtunov AI, Gromashevskii VL, Isolation of the West Nile fever virus from the great cormorant Phalacrocorax carbo, the crow Corvus corone, and Hyalomma marginatum ticks associated with them in natural and synanthroic biocenosis in the Volga delta (Astrakhan region, 2001). Vopr Virusol. 2002;47:7–12.
- Lvov DK, Timopheeva AA, Smirnov VA, Gromashevsky VL, Sidorova GA, Nikiforov LP, Ecology of tick-borne viruses in colonies of birds in the USSR. Med Biol. 1975;53:325–30.
- Hoogstraal H. Birds as tick hosts and as reservoirs and disseminators of tickborne infectious agents. Wiad Parazytol. 1972;18:703–6.
- Hoogstraal H, Clifford CM, Keirans JE, Kaiser MN, Evans DE. The Ornithodoros (Alectorobius) capensis group (Acarina: Ixodoidea: Argasidae) of the palearctic and oriental regions. O. (A.) maritimus: identity, marine bird hosts, virus infections, and distribution in western Europe and northwestern Africa. J Parasitol. 1976;62:799–810.
- Sonenshine DE. Biology of ticks. Oxford: Oxford University Press; 1991.
- Centers for Disease Control and Prevention. Epidemic/epizootic West Nile virus in the United States: Guidelines for surveillance, prevention, and control. Atlanta: The Centers; 2003.
- Whitman L, Aitken THG. Potentiality of Ornithodorus moubata Murray (Acarina, Argasidae) as a reservoir-vector of West Nile virus. Ann Trop Med Parasitol. 1960;54:192–204.
- Sonenshine DE, Mather TN. Ecological dynamics of tick-borne zoonoses. Oxford: Oxford University Press; 1994.
- Jones LD, Davies CR, Steele GM, Nuttall PA. The rearing and maintenance of ixodid and argasid ticks in the laboratory. Anim Technol. 1988;39:99–106.
- Gould EA, Clegg JCS. Growth, titration and purification of alphaviruses and flaviviruses. In: Mahy BWJ, editors. Virology—a practical approach: Oxford: IRL; 1985. p. 43–78.
- Gould EA, Buckley A, Cammack N, Barrett ADT, Clegg JCS, Ishak R, Examination of the immunological relationships between flaviviruses using yellow fever virus monoclonal antibodies. J Gen Virol. 1985;66:1369–82.
- Gould EA, Buckley A, Higgs S, Gaidamovich SY. Antigenicity of flaviviruses. In: Archives of virology. Supplementum 1. Calisher CH, editor. Vienna: Springer-Verlag; 1990. p. 137–52.
- Taylor RM, Work TH, Hurlbut HS, Rizk F. A study of the ecology of West Nile virus in Egypt. Am J Trop Med Hyg. 1956;5:579–620.
- Hurlbut HS. West Nile virus infection in arthropods. Am J Trop Med Hyg. 1956;5:76–85.
- Vermeil C, Lavillaureix J, Reeb E. Sur la conservation et la transmission du virus West Nile par quelques arthropodes. Bull Soc Pathol Exot. 1960;53:273–9.
- Abbassy MM, Stein KJ, Osman M. New artificial feeding technique for experimental infection of Argas ticks (Acari: Argasidae). J Med Entomol. 1994;31:202–5.
- Abbassy MM, Osman M, Marzouk AS. West Nile virus (Flaviviridae:Flavivirus) in experimentally infected Argas ticks (Acari:Argasidae). Am J Trop Med Hyg. 1993;48:726–37.
- Randolph SE, Miklisova D, Lysy J, Rogers DJ, Labuda M. Incidence from coincidence: patterns of tick infestations on rodents facilitate transmission of tick-borne encephalitis virus. Parasitology. 1999;118:177–86.
- Komar N, Langevin S, Hinten S, Nemeth N, Edwards E, Hettler D, Experimental infection of north american birds with the New York 1999 strain of West Nile virus. Emerg Infect Dis. 2003;9:311–22.
- McLean RG, Ubico SR, Docherty DE, Hansen WR, Sileo L, McNamara TS. West Nile virus transmission and ecology in birds. Ann N Y Acad Sci. 2001;951:54–7.
- Hoodless AN, Kurtenbach K, Nuttall PA, Randolph SE. The impact of ticks on pheasant territoriality. Oikos. 2002;96:245–50 Available from http://www.ingenta.com.
- Keirans JE, Hutcheson HJ, Oliver JH Jr. Ornithodoros (Alectorobius) capensis Neumann (Acari: Ixodoidea: Argasidae), a parasite of seabirds, established along the southeastern seacoast of the United States. J Med Entomol. 1992;29:371–3.
- Randolph SE, Gern L, Nuttall PA. Co-feeding ticks: epidemiological significance for tick-borne pathogen transmission. Parasitol Today. 1996;12:472–9 Available from http://www.sciencedirect.com.
- Mead DG, Ramberg FB, Besselsen DG, Mare CJ. Transmission of vesicular stomatitis virus from infected to noninfected black flies co-feeding on nonviremic deer mice. Science. 2000;287:485–7.
- Labuda M, Austyn JM, Zuffova E, Kozuch O, Fuchsberger N, Lysy J, Importance of localized skin infection in tick-borne encephalitis virus transmission. Virology. 1996;219:357–66.
- Johnston LJ, Halliday GM, King NJ. Langerhans cells migrate to local lymph nodes following cutaneous infection with an arbovirus. J Invest Dermatol. 2000;114:560–8.
- Weinlich G, Sepp N, Koch F, Schuler G, Romani N. Evidence that Langerhans cells rapidly diasappear from the epidermis in response to contact sensitizers but not to tolerogens/nonsensitizers. In: Arbeitsgemeinschaft Dermatologische Forschung (ADF) in cooperation with Deutsche Dermatologische Gesellschaft. XVII meeting. 1989 Nov 17–19. Hamburg, Federal Republic of Germany. Abstracts. Arch Dermatol Res. 1990;281:544–91.
- Sardelis MR, Turell MJ, Dohm DJ, O’Guinn ML. Vector competence of selected North American Culex and Coquillettidia mosquitoes for West Nile virus. Emerg Infect Dis. 2001;7:1018–22.
- Turell MJ, Sardelis MR, Dohm DJ, O’Guinn ML. Potential North American vectors of West Nile virus. Ann N Y Acad Sci. 2001;951:317–24.
- Goddard LB, Roth AE, Reisen WK, Scott TW. Vector competence of California mosquitoes for West Nile virus. Emerg Infect Dis. 2002;8:1385–91.
Suggested citation for this article: Lawrie CH, Uzcátegui NY, Gould EA, Nuttall PA. Ixodid and argasid tick species and West Nile virus. Emerg Infect Dis [serial online] 2004 Apr [date cited]. Available from: http://wwwnc.cdc.gov/eid/article/10/4/03-0517.htm