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Acquired resistance and antibody response of raccoons (Procyon lotor) to sequential feedings of Ixodes scapularis (Acari: Ixodidae)
veterinary parasitology ELSEVIER
Veterinary Parasitology63 (1996) 291-301
Acquired resistance and antibody response of raccoons ( Procyon lotor) to sequential feedings of lxodes scapularis ( Acari: Ixodidae) L.E. Craig a,*, D.E. Norris b,c,l, M.L. Sanders a, G.E. Glass a, B.S. Schwartz d a Department of Molecular Microbiology and Immunology, Johns Hopkins University School of Hygiene and Public Health, Baltimore, MD 21205, USA b Department of Entomology, College of Agriculture and Life Sciences, North Carolina State University, Raleigh, ArC 27606, USA e Department of Microbiology, Pathology, and Parasitology, College of Veterinary Medicine, North Carolina State University, Raleigh, NC 27606, USA d Department of Environmental Health Sciences, Division of Occupational Health, Johns Hopkins University School of Hygiene and Public Health, Baltimore, MD 21205, USA
Received 30 May 1995; accepted 6 October 1995
Abstract
Captive-bred raccoons (Procyon lotor) developed immune resistance to infestation by the larval stage of the ixodid tick, Ixodes scapularis, the vector of Borrelia burgdorferi, following repeated applications of both nymphs and larvae. Resistance was expressed as a significant decrease in the proportion of engorged larvae recovered from each cohort. Resistance to nymphs was not noted, but, only two such cohorts were applied. Utilizing an enzyme-linked immunosorbent assay (ELISA) developed to detect raccoon serum antibodies to tick salivary gland antigens, raccoons evidenced a two to ten-fold increase in anti-tick salivary extract antibody titer following the application of two cohorts of nymphs and eights cohorts of larvae. The tick saliva antigens recognized by both pre- and post-exposure raccoon sera were evaluated by Western blotting. The production of antibodies correlated with the development of resistance to infestation, suggesting that the resistance was immune-mediated and could be measured by anti-tick salivary extract antibody titers. Resistance in exposed raccoons prevents nearly 90% of larvae from prolonged
* Corresponding author at: Division of Comparative Medicine, Johns Hopkins University School of Medicine, Room G60 Traylor Building, Baltimore, MD 21205, USA. l Present address: Arthropod Borne Infectious Diseases Laboratory, Foothills Campus, Department of Microbiology,Colorado State University,Fort Collins, CO 80523, USA. 0304-4017/96/$15.00 © 1996 Elsevier Science B.V. All fights reserved SSDI 0304-4017(95)0091 1-6
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feeding. Prolonged feeding is required for engorgement and the transmission of various infectious agents, such as B. burgdorferi. Keywords: Raccoon; lxodes scapularis; Resistance
1. Introduction
Ixodes scapularis is the vector of Borrelia burgdorferi, the causative agent of Lyme disease. I. scapularis is a three-host tick that often feeds on white-footed mice (Peromyscus leucopus) in the larval and nymphal stages and white-tailed deer (Odocoileus virginianus) as an adult (Fish and Dowler, 1989). The ability of raccoons (Procyon lotor) to serve as reservoirs of B. burgdorferi in nature has been suggested by the presence of both immature and adult I. scapularis on wild raccoons (Fish and Dowler, 1989), by the presence of specific antibodies to B. burgdorferi in wild raccoons (Magnarelli et al., 1991), and by the recovery of B. burgdorferi from engorged I. scapularis larvae removed from wild raccoons (Fish and Daniels, 1990). However, the proportion of immature ticks that were infected with the spirochete was lower in raccoons than in white-footed mice, suggesting that the raccoons were less efficient reservoirs than mice (Fish and Daniels, 1990). One explanation for the lower proportion of infected ticks from raccoons may be the development of resistance to the ticks. The phenomenon of tick resistance has been reported in several animal species including guinea pigs (Trager, 1939a; Brown et al., 1982), rabbits (Latif et al., 1988; Clarke et al., 1989), mice (Need et al., 1991), and cattle (Roberts and Kerr, 1976; Wong and Opdebeeck, 1993). Resistance was demonstrated by reduced numbers of ticks feeding to repletion, reduced weights of feeding ticks, reduced feeding rates, reduced ability of fed larvae or nymphs to molt to the next stadium, or reduced progeny of fed female ticks (Allen, 1989). Experimental evidence has revealed that this resistance has an immunologic basis. Resistance is partially transferable to naive animals using serum from immune animals (Trager, 1939a; Brown et al., 1982). The serum from immune animals also has been shown to have an anti-feeding effect on ticks in vitro (Losel et al., 1992). The antigens eliciting the immunoglobulin production are in the saliva of the tick (Shapiro et al., 1987) and have been characterized in several tick species by Western blot analysis (Janse van Vuuren et al., 1992). Both mast cells and IgE are required for tick resistance in mice (Matsuda et al., 1990). Mast cell deficient mice develop a lower level of resistance than do mast cell sufficient mice (denHollander and Allen, 1986). Intact and degranulating mast cells, basophils and eosinophils are seen histologically surrounding the hypostome in tick resistant rabbits (Brossard and Fivaz, 1982). The role of T lymphocytes in tick resistance has been investigated in rabbits by treating them with anti-thymocyte serum. It was found that the T cell deficient rabbits developed less resistance and less of an antibody titer than the normal controls (Njau, 1989). However, the T cell deficient rabbits did develop a measurable titer and tick resistance (Njau, 1989), indicating that antibodies do have a role in tick resistance.
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Anti-tick saliva antibodies have been used as a marker in the development of host resistance in several species (Trager, 1939a) and the specific class of antibodies involved has been identified as IgG in rabbits (Brown, 1988) and IgG1 in guinea pigs (Brown et al., 1982). The detection of anti-tick saliva antibodies also has been used as a biologic marker of tick exposure in humans (Schwartz et al., 1991; Schwartz et al., 1993). The purpose of this experiment was threefold: (1) to determine if raccoons developed resistance to L scapularis following sequential exposure to tick cohorts that might affect their roles in maintaining tick populations; (2) to detect and quantitate any antibody response in the raccoons to the tick salivary antigens; and (3) to correlate the antibody response with resistance.
2. Materials and methods
2.1. Raccoons Three full-sibling, captive-born, parasite-free, tick-naive raccoons were used in this study. The animals were housed at the Zoonosis Laboratory at North Carolina State University College of Veterinary Medicine in cages (87 cm x 45 cm X 78 cm) over pans filled with water to facilitate the recovery of ticks. Raccoons were anesthetized with ketamine (15.0 mg k g - l ) / x y l a z i n e (0.5 mg k g - l ) prior to tick placement and blood sampling. Blood samples were collected by jugular venipuncture from each raccoon on a weekly or semi-weekly basis. The blood was allowed to clot at 4°C and the serum was harvested and stored at -70°C until testing. 2.2. Ticks I. scapularis larvae and nymphs obtained from Dr. J. Piesman (Centers for Disease Control, Vector Borne Infectious Diseases Laboratory, Fort Collins, CO) were used in this experiment. The first cohort of 20 nymphs was placed in the ears of each of the three raccoons on Day 0. One day after engorged nymphs were recovered (Day 4), a cohort of 300 larvae was applied to each raccoon. Subsequent cohorts of 300 larvae were placed on each raccoon on Days 20, 33, and 64. A second cohort of nymphs was placed on the raccoons on Day 175 and four subsequent cohorts of 300 larvae were applied on Days 181, 195, 209, and 237. 2.3. Antibody assay Anti-tick saliva antibodies were assayed by enzyme-linked immunosorbent assay (ELISA) performed at least in duplicate. Antigen was prepared by sonicating the salivary glands of laboratory-reared, uninfected adult I. scapularis (kindly provided by Dr. D. Fish) which had been allowed to feed for 3 days on rabbits. The sonication and storage methods have been previously described (Schwartz et al., 1990). Briefly, the salivary glands were dissected and stored in cold phosphate buffered saline (PBS), containing phenylthiourea (250 /zg 100ml-l PBS). The glands were sonicated for 5
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min. After centrifugation at 10 000 × g for 5 min, the protein content of the supernatant of the preparation was measured using the BCA Protein Assay (Pierce, Rockford, IL). The final protein concentration was 1.43 /xg /xl -l. Aliquots of 50 /xl were stored at - 70°C. The antigen was diluted 1:100 in a carbonate buffer (pH 9.6), and 5 0 / z l were added to the inner 60 wells of a 96 well microtiter plate. The plates were incubated at least 18 h at 4°C and then blocked with 5% fetal bovine serum (FBS) in phosphatebuffered saline containing 0.5% Tween-20 (PBST) for 1 h at 37°C. Serum samples were diluted 1:100 in PBST with 1% FBS. 50/~1 of this dilution was applied to the wells and incubated overnight at 4°C. Goat anti-raccoon IgG (heavy and light chain) conjugate labeled with alkaline phosphatase (Kirkegaard & Perry, Gaithersburg, MD) was diluted 1:1000 in phosphate-buffered saline (PBS) and applied to each well and incubated for 2 h at 37°C. 100 /zl of the substrate (5 mg ml-1 p-nitrophenyl phosphate) was added to each well and the plates were incubated for 2 h in the dark. The optical density was measured with a spectrophotometer at 405 nm. 2.4. Titer determination
The difference between the average of the two antigen positive wells and the average of the two antigen negative wells was determined for each sample. A late serum sample (positive control) from one of the raccoons was randomly selected to establish a dilutions series. Serum from this sample was diluted from 1:50 to 1:12 800 as a two-fold dilution series. Pre-exposure samples (1:100) from each of the raccoons were used to establish a baseline reading for negative sera. The titer for the positive control was determined to be the optical density reading that exceeded 95% of the negative (pre-exposure) samples. Three dilutions from the linear portion of the dilution series were included on each plate and used to calculate the titer of the remaining sera. The serum samples were tested on two separate days, but the three control dilutions were included on each plate to diminish day to day variation. 2.5. Western blotting lxodes scapularis salivary gland antigen was prepared and stored as described above (Antibody assay). Antigen preparation containing 240 /xg of protein per lane was separated electrophoretically by discontinuous sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) in a pre-cast, 4-20% gradient denaturing gel (Jules Biotechnology, New Haven, CT). Proteins were transferred to a nitrocellulose membrane (NC) (Bioblot-NC; Costar Corp., Cambridge, MA) in a MiniProtean II Electrophoresis/ Transfer Unit (Biorad Laboratories) containing 192 mM glycine, 89 mM Tris, and 20% ( v / v ) methanol in distilled water at 4°C, for 3.5 h at 170 mA. Following transfer, the NC membrane was cut into 4-mm wide strips and incubated overnight at 4°C in 2% non-fat milk in PBS. The NC strips were then incubated separately in a 1:100 dilution of raccoon sera overnight with rocking. All dilutions were made in 2% non-fat milk in PBS. The NC strips were washed five times with PBST and incubated for 1 h with horse radish peroxidase labeled goat anti-raccoon IgG (gamma chain) (The Binding Site, San Diego, CA) at a 1:1000 dilution. The NC strips were
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washed five times with PBST followed by five times with PBS and then incubated with 4-chloro-l-naphthol (4CN Peroxidase Substrate, Kirkegaard & Perry Laboratories Inc., Gaithersburg, MD). Western blots of raccoon sera were performed using pre-tick exposure sera and sera harvested from raccoons 223 days following first tick exposure.
2.6. Data analysis The development of resistance by raccoons to I. scapularis was examined by testing for trends in the proportion of larval and nymphal ticks that became engorged following placement on the raccoon using chi-square analysis. Temporal trends in antibody titers in the raccoons following sequential feedings were examined graphically.
3. R e s u l t s
3.1. Development of resistance There was a highly significant trend for the proportion of successful feeding larvae to decrease with successive feedings (Fig. 1) in each raccoon (X 2 = 120, P < 0.00001, Raccoon 1; X 2 = 761.7, P < 0.00001, Raccoon 2; X 2 = 736.4, P < 0.00001, Raccoon 3). Except for Raccoon 1, more than 90% of larvae successfully fed during the initial exposure. Successful feeding decreased to 40 to 60% by the second feeding (Fig. 1). By the fifth and all subsequent feedings, only 10 to 15% of larvae successfully engorged. In contrast, there was no evidence, based on the proportion of engorged nymphal ticks recovered, for the development of resistance to feedings by the second cohort of nymphs (Fig. 2). In each raccoon, the proportion of nymphs successfully feeding was slightly higher during the second exposure (P < 0.05). 100
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L.E. Craig et al./ Veterinary Parasitology 63 (1996) 291-301
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Fig. 4. Western blot of tick salivary gland proteins reacted with raccoon sera. Lanes 1, 2, and 3: sera from Raccoons 1, 2, and 3, respectively, bled on Day 0 (pre-tick exposure). Lanes 4, 5, and 6: sera from Raccoons 1, 2, and 3, respectively, bled on Day 223 (post-tick exposure).
3.2. Antibody production Antibody responses to tick feedings were variable among the three raccoons. Raccoon 1 had an elevated titer before ticks were applied and developed a significant antibody titer during the first four groups of larval feedings. Anti-tick salivary extract antibody titers then declined to baseline until the second exposure to nymphal ticks. Titers rose during the second cohort application, increasing to a peak on Day 251, 2 weeks after the last cohort of ticks were applied (Fig. 3). Raccoon 2 showed relatively little response to the initial feedings by larvae, but had a two to three-fold increase in antibody titer following the second nymphal feeding (Fig. 3). Responses to later larval feedings tended to be of a minor, though measurable nature. Raccoon 3 developed a significant antibody titer by the end of the initial larval feedings and then developed a very high titer following the second nymphal feeding that was boosted by subsequent larval feedings (Fig. 3).
3.3. Antigen recognition Following tick exposure the serum from all three raccoons recognized salivary antigens at 175, 145, 140, 118, and 90 kD. In addition, there were five to six high molecular weight bands ( > 205 kD) and 10-15 minor bands ranging from 83 to 30 kD (Fig. 4, Lanes 4, 5, and 6). Serum from Raccoon 1 recognized antigens of 175, 145, 140, ll8, and 90 kD in size prior to tick exposure (Fig. 4, Lane 1). The pre-exposure serum of Raccoons 2 and 3 (Lanes 2 and 3) developed no bands.
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4. Discussion
There was a dramatic reduction in the percentage of larvae recovered after sequential feedings (Fig. 1), indicating the development of resistance. Less than 20% of larvae applied were recovered fully engorged after the second nymphal exposure (Fig. 1). This reduction could be due to either an inflammatory response that interferes with feeding, or to an inflammatory response that increases grooming behavior resulting in removal by the host. The placement of the tick larvae and nymphs within the ears was thought to reduce the latter possibility. Either mechanism reduces parasite feeding time and prevents full engorgement. Full engorgement is required for molting to the next stadium (Allen, 1989) and 3 days of feeding is required for optimal transmission of B. burgdorferi (Piesman et al., 1987). Our findings suggest that previously exposed raccoons would be partially resistant to I. scapularis, preventing up to 85% of the feeding larvae from reaching full engorgement. The resistance to larval feeding correlates well with the development of anti-tick salivary extract antibody titers. The titer increased most dramatically following the second exposure to nymphs (Fig. 3), and the drop in the proportion of ticks feeding successfully also occurred following the second cohort of nymphs (Fig. 1). The pattem of antibody response varied somewhat among the three raccoons, which may be due to individual differences in the immune response in this outbred population. Naturally acquired immunity to ticks has been reported to vary widely in several species (Wong and Opdebeeck, 1993). Although, the correlation between tick resistance and antibody production is good, other immunologic mechanisms such as cell mediated immunity and hypersensitivity reactions are probably also involved. However, antibody titers to tick salivary gland antigens have been used as a marker of tick resistance in many species (Brown et al., 1982; Shapiro et al., 1986, 1987; Brown, 1988; Njau et al., 1988; Allen, 1989; Wheeler et al., 1989; Wong and Opdebeeck, 1993). In addition, antibodies produced by the host have been shown to pass across the gut epithelium of the tick serologically intact (Brossard and Rais, 1984; Fujisaki et al., 1984; Minoura et al., 1985). These anti-tick antibodies have been shown to retain binding activity by localizing to the tick salivary glands (Ackerman et al., 1981). Antibodies to tick gut components (regurgitated during feeding) have also been postulated to play a role in the development of resistance (Shapiro et al., 1986). However, inoculation with extracts of tick digestive tract, cephalic glands, and salivary glands showed that salivary gland extracts elicited the strongest resistance (Trager, 1939b). Raccoon 1 had an elevated anti-tick salivary gland extract antibody titer prior to tick application and was positive on Western blotting for the antigens recognized by the other raccoons only following tick exposure. This correlates well with the initial elevated titers seen on the ELISA (Fig. 3) and the reduced survival of larvae in the first four cohorts (Fig. 1) for Raccoon 1, suggesting that the antibodies detected in the Western blot and ELISA did interfere with larval feeding. Antibodies to mites have been reported to cross react with tick antigens in mice (denHollander and Allen, 1985). Interestingly, antibodies to ticks do not cross react with mite antigens (denHollander and Allen, 1986; Boyce et al., 1991). The occurrence of mite exposure in these raccoons is
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unknown, however, mites are common parasites of laboratory animals (Shadduck and Pakes, 1978). We postulate that the antibodies detected in Raccoon 1 prior to exposure to the ticks were the result of previous mite infestation, since ticks are very uncommon in laboratory animals (Shadduck and Pakes, 1978). Some of the variation in antibody responses could be due to differences in antigens expressed by the different stages of ticks. Adult salivary glands were used as antigen in the ELISA, but larvae and nymphs were used in the experiment to measure feeding success. The raccoons developed resistance to larvae after four cohorts, but not to nymphs, which were only placed on the animals twice. Although data on this subject are scant, some prior research has suggested that many major antigens in adult salivary glands are not present in larval and nymphal salivary glands of ixodid ticks (Shapiro et al., 1986). The current study, in using adult salivary gland extract to evaluate antibody response to feeding by larvae and nymphs, suggests that there are either important proteins common to the glands of larvae and adults, or alternatively, that there are cross-reacting antibodies to antigenically similar, but not identical, proteins among the stages of I. scapularis. Many of the proteins secreted by feeding ticks have anti-coagulant, anti-histamine, and anti-inflammatory effects. These proteins facilitate blood feeding during the prolonged period of host attachment by antagonizing the chemical mediators of the host inflammatory response (Ribeiro et al., 1985). Resistance has not been found in the ticks' natural host species (Randolph, 1979), suggesting that the anti-inflammatory proteins in the tick saliva specifically antagonize the mediators of inflammation and resistance in the natural host (Ribeiro et al., 1985). In the absence of this specific interaction, i.e. in an unnatural host, inflammation and an immune response specific for tick salivary antigens results. The results of this study suggest that the raccoon is not a natural host for L scapularis. The role of raccoons in the maintenance of populations of I. scapularis and the transmission of B. burgdorferi is questionable. Although antibodies to B. burgdorferi have been found in raccoons (Magnarelli et al., 1991), the current study suggests that they develop resistance to the vector tick species. Specifically, the duration of feeding for most larvae would not be long enough in immune animals to efficiently transmit the spirochete. The longer feeding times on relatively naive raccoons and the ticks that do escape the immune mechanisms to feed to full engorgement may allow transmission of the spirochete, explaining the presence of antibodies in wild raccoon populations.
Acknowledgments The authors would like to thank Charles S. Apperson and Bruce F. Levine for their assistance. This research was supported by grants from the National Institute of Allergy and Infectious Diseases R55 AI 30042 and R29 AI 31608 and from the National Institutes of Health RR00130 and RR07002.
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Randolph, S.E., 1979. Population regulation in ticks: the role of acquired resistance in natural and unnatural hosts. Parasitology, 79: 141-156. Ribeiro, J.M., Makoul, G.T., Levine, J., Robinson, D.R. and Spielman, A., 1985. Antihemostatic, antiinflammatory, and immunosuppressive properties of the saliva of a tick, lxodes dammini. J. Exp. Med., 161: 332-344. Roberts, J.A. and Kerr, J.D., 1976. Boophilus microplus: Passive transfer of resistance in cattle. J. Parasitol., 62: 485-489. Schwartz, B.S., Ribeiro, J.M.C. and Goldstein, M., 1990. Anti-tick antibodies: an epidemiologic tool in Lyme disease research. Am. J. Epidemiol., 132: 58-66. Schwartz, B.S., Ford, D.P., Childs, J.E., Rothman, N. and Thomas, R.J., 1991. Anti-tick saliva antibody: A biological marker of tick exposure that is a risk factor for Lyme disease seropositivity. Am. J. Epidemiol., 134: 86-95. Schwartz, B.S., Nadelman, R.B., Fish, D., Childs, J.E., Forester, G. and Wormser, G., 1993. Entomologic and demographic correlates of anti-tick saliva antibody in a prospective study of tick bite subjects in Westchester County, New York. Am. J. Trop. Med. Hyg., 48: 50-57. Shadduck, J.A. and Pakes, S.P., 1978. Protozoal and metazoal diseases. In: K. Benirschke, F.M. Garner and T.C. Jones (Editors), Pathology of Laboratory Animals. Springer-Verlag, New York, pp. 1587-1696. Shapiro, S.Z., Voigt, W.P. and Fujisaki, K., 1986. Tick antigens recognized by serum from a guinea pig resistant to infestation with the tick Rhipicephalus appendiculatus. J. Parasitol., 72: 454-463. Shapiro, S.Z., Buscher, G, and Dobbelaere, D.A.E., 1987. Acquired resistance to Rhipicephalus appendiculatus (Acari: Ixodidae): Identification of an antigen eliciting resistance in rabbits. J. Med. Entomol., 24: 147-154. Trager, W., 1939a. Acquired immunity to ticks. J. Parasitol., 25: 57-81. Trager, W., 1939b. Further observations on acquired immunity to the tick Dermacentor variablilis Say. J. Parasitol., 25: 137-139. Wheeler, C.M., Coleman, J.L., Habicht, G.S. and Benach, J.L., 1989. Adult lxodes dammini on rabbits: Development of acute inflammation in the skin and immune responses to salivary gland, midgut, and spirochetal components. J. Infect. Dis., 159: 265-273. Wong, J.Y.M. and Opdebeeck, J.P., 1993. Immunity in vaccinated cattle exposed to experimental and natural infestations with Boophilus microplus. Int. J. Parasitol., 23: 689-692.
Veterinary Parasitology63 (1996) 291-301
Acquired resistance and antibody response of raccoons ( Procyon lotor) to sequential feedings of lxodes scapularis ( Acari: Ixodidae) L.E. Craig a,*, D.E. Norris b,c,l, M.L. Sanders a, G.E. Glass a, B.S. Schwartz d a Department of Molecular Microbiology and Immunology, Johns Hopkins University School of Hygiene and Public Health, Baltimore, MD 21205, USA b Department of Entomology, College of Agriculture and Life Sciences, North Carolina State University, Raleigh, ArC 27606, USA e Department of Microbiology, Pathology, and Parasitology, College of Veterinary Medicine, North Carolina State University, Raleigh, NC 27606, USA d Department of Environmental Health Sciences, Division of Occupational Health, Johns Hopkins University School of Hygiene and Public Health, Baltimore, MD 21205, USA
Received 30 May 1995; accepted 6 October 1995
Abstract
Captive-bred raccoons (Procyon lotor) developed immune resistance to infestation by the larval stage of the ixodid tick, Ixodes scapularis, the vector of Borrelia burgdorferi, following repeated applications of both nymphs and larvae. Resistance was expressed as a significant decrease in the proportion of engorged larvae recovered from each cohort. Resistance to nymphs was not noted, but, only two such cohorts were applied. Utilizing an enzyme-linked immunosorbent assay (ELISA) developed to detect raccoon serum antibodies to tick salivary gland antigens, raccoons evidenced a two to ten-fold increase in anti-tick salivary extract antibody titer following the application of two cohorts of nymphs and eights cohorts of larvae. The tick saliva antigens recognized by both pre- and post-exposure raccoon sera were evaluated by Western blotting. The production of antibodies correlated with the development of resistance to infestation, suggesting that the resistance was immune-mediated and could be measured by anti-tick salivary extract antibody titers. Resistance in exposed raccoons prevents nearly 90% of larvae from prolonged
* Corresponding author at: Division of Comparative Medicine, Johns Hopkins University School of Medicine, Room G60 Traylor Building, Baltimore, MD 21205, USA. l Present address: Arthropod Borne Infectious Diseases Laboratory, Foothills Campus, Department of Microbiology,Colorado State University,Fort Collins, CO 80523, USA. 0304-4017/96/$15.00 © 1996 Elsevier Science B.V. All fights reserved SSDI 0304-4017(95)0091 1-6
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feeding. Prolonged feeding is required for engorgement and the transmission of various infectious agents, such as B. burgdorferi. Keywords: Raccoon; lxodes scapularis; Resistance
1. Introduction
Ixodes scapularis is the vector of Borrelia burgdorferi, the causative agent of Lyme disease. I. scapularis is a three-host tick that often feeds on white-footed mice (Peromyscus leucopus) in the larval and nymphal stages and white-tailed deer (Odocoileus virginianus) as an adult (Fish and Dowler, 1989). The ability of raccoons (Procyon lotor) to serve as reservoirs of B. burgdorferi in nature has been suggested by the presence of both immature and adult I. scapularis on wild raccoons (Fish and Dowler, 1989), by the presence of specific antibodies to B. burgdorferi in wild raccoons (Magnarelli et al., 1991), and by the recovery of B. burgdorferi from engorged I. scapularis larvae removed from wild raccoons (Fish and Daniels, 1990). However, the proportion of immature ticks that were infected with the spirochete was lower in raccoons than in white-footed mice, suggesting that the raccoons were less efficient reservoirs than mice (Fish and Daniels, 1990). One explanation for the lower proportion of infected ticks from raccoons may be the development of resistance to the ticks. The phenomenon of tick resistance has been reported in several animal species including guinea pigs (Trager, 1939a; Brown et al., 1982), rabbits (Latif et al., 1988; Clarke et al., 1989), mice (Need et al., 1991), and cattle (Roberts and Kerr, 1976; Wong and Opdebeeck, 1993). Resistance was demonstrated by reduced numbers of ticks feeding to repletion, reduced weights of feeding ticks, reduced feeding rates, reduced ability of fed larvae or nymphs to molt to the next stadium, or reduced progeny of fed female ticks (Allen, 1989). Experimental evidence has revealed that this resistance has an immunologic basis. Resistance is partially transferable to naive animals using serum from immune animals (Trager, 1939a; Brown et al., 1982). The serum from immune animals also has been shown to have an anti-feeding effect on ticks in vitro (Losel et al., 1992). The antigens eliciting the immunoglobulin production are in the saliva of the tick (Shapiro et al., 1987) and have been characterized in several tick species by Western blot analysis (Janse van Vuuren et al., 1992). Both mast cells and IgE are required for tick resistance in mice (Matsuda et al., 1990). Mast cell deficient mice develop a lower level of resistance than do mast cell sufficient mice (denHollander and Allen, 1986). Intact and degranulating mast cells, basophils and eosinophils are seen histologically surrounding the hypostome in tick resistant rabbits (Brossard and Fivaz, 1982). The role of T lymphocytes in tick resistance has been investigated in rabbits by treating them with anti-thymocyte serum. It was found that the T cell deficient rabbits developed less resistance and less of an antibody titer than the normal controls (Njau, 1989). However, the T cell deficient rabbits did develop a measurable titer and tick resistance (Njau, 1989), indicating that antibodies do have a role in tick resistance.
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Anti-tick saliva antibodies have been used as a marker in the development of host resistance in several species (Trager, 1939a) and the specific class of antibodies involved has been identified as IgG in rabbits (Brown, 1988) and IgG1 in guinea pigs (Brown et al., 1982). The detection of anti-tick saliva antibodies also has been used as a biologic marker of tick exposure in humans (Schwartz et al., 1991; Schwartz et al., 1993). The purpose of this experiment was threefold: (1) to determine if raccoons developed resistance to L scapularis following sequential exposure to tick cohorts that might affect their roles in maintaining tick populations; (2) to detect and quantitate any antibody response in the raccoons to the tick salivary antigens; and (3) to correlate the antibody response with resistance.
2. Materials and methods
2.1. Raccoons Three full-sibling, captive-born, parasite-free, tick-naive raccoons were used in this study. The animals were housed at the Zoonosis Laboratory at North Carolina State University College of Veterinary Medicine in cages (87 cm x 45 cm X 78 cm) over pans filled with water to facilitate the recovery of ticks. Raccoons were anesthetized with ketamine (15.0 mg k g - l ) / x y l a z i n e (0.5 mg k g - l ) prior to tick placement and blood sampling. Blood samples were collected by jugular venipuncture from each raccoon on a weekly or semi-weekly basis. The blood was allowed to clot at 4°C and the serum was harvested and stored at -70°C until testing. 2.2. Ticks I. scapularis larvae and nymphs obtained from Dr. J. Piesman (Centers for Disease Control, Vector Borne Infectious Diseases Laboratory, Fort Collins, CO) were used in this experiment. The first cohort of 20 nymphs was placed in the ears of each of the three raccoons on Day 0. One day after engorged nymphs were recovered (Day 4), a cohort of 300 larvae was applied to each raccoon. Subsequent cohorts of 300 larvae were placed on each raccoon on Days 20, 33, and 64. A second cohort of nymphs was placed on the raccoons on Day 175 and four subsequent cohorts of 300 larvae were applied on Days 181, 195, 209, and 237. 2.3. Antibody assay Anti-tick saliva antibodies were assayed by enzyme-linked immunosorbent assay (ELISA) performed at least in duplicate. Antigen was prepared by sonicating the salivary glands of laboratory-reared, uninfected adult I. scapularis (kindly provided by Dr. D. Fish) which had been allowed to feed for 3 days on rabbits. The sonication and storage methods have been previously described (Schwartz et al., 1990). Briefly, the salivary glands were dissected and stored in cold phosphate buffered saline (PBS), containing phenylthiourea (250 /zg 100ml-l PBS). The glands were sonicated for 5
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min. After centrifugation at 10 000 × g for 5 min, the protein content of the supernatant of the preparation was measured using the BCA Protein Assay (Pierce, Rockford, IL). The final protein concentration was 1.43 /xg /xl -l. Aliquots of 50 /xl were stored at - 70°C. The antigen was diluted 1:100 in a carbonate buffer (pH 9.6), and 5 0 / z l were added to the inner 60 wells of a 96 well microtiter plate. The plates were incubated at least 18 h at 4°C and then blocked with 5% fetal bovine serum (FBS) in phosphatebuffered saline containing 0.5% Tween-20 (PBST) for 1 h at 37°C. Serum samples were diluted 1:100 in PBST with 1% FBS. 50/~1 of this dilution was applied to the wells and incubated overnight at 4°C. Goat anti-raccoon IgG (heavy and light chain) conjugate labeled with alkaline phosphatase (Kirkegaard & Perry, Gaithersburg, MD) was diluted 1:1000 in phosphate-buffered saline (PBS) and applied to each well and incubated for 2 h at 37°C. 100 /zl of the substrate (5 mg ml-1 p-nitrophenyl phosphate) was added to each well and the plates were incubated for 2 h in the dark. The optical density was measured with a spectrophotometer at 405 nm. 2.4. Titer determination
The difference between the average of the two antigen positive wells and the average of the two antigen negative wells was determined for each sample. A late serum sample (positive control) from one of the raccoons was randomly selected to establish a dilutions series. Serum from this sample was diluted from 1:50 to 1:12 800 as a two-fold dilution series. Pre-exposure samples (1:100) from each of the raccoons were used to establish a baseline reading for negative sera. The titer for the positive control was determined to be the optical density reading that exceeded 95% of the negative (pre-exposure) samples. Three dilutions from the linear portion of the dilution series were included on each plate and used to calculate the titer of the remaining sera. The serum samples were tested on two separate days, but the three control dilutions were included on each plate to diminish day to day variation. 2.5. Western blotting lxodes scapularis salivary gland antigen was prepared and stored as described above (Antibody assay). Antigen preparation containing 240 /xg of protein per lane was separated electrophoretically by discontinuous sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) in a pre-cast, 4-20% gradient denaturing gel (Jules Biotechnology, New Haven, CT). Proteins were transferred to a nitrocellulose membrane (NC) (Bioblot-NC; Costar Corp., Cambridge, MA) in a MiniProtean II Electrophoresis/ Transfer Unit (Biorad Laboratories) containing 192 mM glycine, 89 mM Tris, and 20% ( v / v ) methanol in distilled water at 4°C, for 3.5 h at 170 mA. Following transfer, the NC membrane was cut into 4-mm wide strips and incubated overnight at 4°C in 2% non-fat milk in PBS. The NC strips were then incubated separately in a 1:100 dilution of raccoon sera overnight with rocking. All dilutions were made in 2% non-fat milk in PBS. The NC strips were washed five times with PBST and incubated for 1 h with horse radish peroxidase labeled goat anti-raccoon IgG (gamma chain) (The Binding Site, San Diego, CA) at a 1:1000 dilution. The NC strips were
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washed five times with PBST followed by five times with PBS and then incubated with 4-chloro-l-naphthol (4CN Peroxidase Substrate, Kirkegaard & Perry Laboratories Inc., Gaithersburg, MD). Western blots of raccoon sera were performed using pre-tick exposure sera and sera harvested from raccoons 223 days following first tick exposure.
2.6. Data analysis The development of resistance by raccoons to I. scapularis was examined by testing for trends in the proportion of larval and nymphal ticks that became engorged following placement on the raccoon using chi-square analysis. Temporal trends in antibody titers in the raccoons following sequential feedings were examined graphically.
3. R e s u l t s
3.1. Development of resistance There was a highly significant trend for the proportion of successful feeding larvae to decrease with successive feedings (Fig. 1) in each raccoon (X 2 = 120, P < 0.00001, Raccoon 1; X 2 = 761.7, P < 0.00001, Raccoon 2; X 2 = 736.4, P < 0.00001, Raccoon 3). Except for Raccoon 1, more than 90% of larvae successfully fed during the initial exposure. Successful feeding decreased to 40 to 60% by the second feeding (Fig. 1). By the fifth and all subsequent feedings, only 10 to 15% of larvae successfully engorged. In contrast, there was no evidence, based on the proportion of engorged nymphal ticks recovered, for the development of resistance to feedings by the second cohort of nymphs (Fig. 2). In each raccoon, the proportion of nymphs successfully feeding was slightly higher during the second exposure (P < 0.05). 100
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L.E. Craig et al./ Veterinary Parasitology 63 (1996) 291-301
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3.2. Antibody production Antibody responses to tick feedings were variable among the three raccoons. Raccoon 1 had an elevated titer before ticks were applied and developed a significant antibody titer during the first four groups of larval feedings. Anti-tick salivary extract antibody titers then declined to baseline until the second exposure to nymphal ticks. Titers rose during the second cohort application, increasing to a peak on Day 251, 2 weeks after the last cohort of ticks were applied (Fig. 3). Raccoon 2 showed relatively little response to the initial feedings by larvae, but had a two to three-fold increase in antibody titer following the second nymphal feeding (Fig. 3). Responses to later larval feedings tended to be of a minor, though measurable nature. Raccoon 3 developed a significant antibody titer by the end of the initial larval feedings and then developed a very high titer following the second nymphal feeding that was boosted by subsequent larval feedings (Fig. 3).
3.3. Antigen recognition Following tick exposure the serum from all three raccoons recognized salivary antigens at 175, 145, 140, 118, and 90 kD. In addition, there were five to six high molecular weight bands ( > 205 kD) and 10-15 minor bands ranging from 83 to 30 kD (Fig. 4, Lanes 4, 5, and 6). Serum from Raccoon 1 recognized antigens of 175, 145, 140, ll8, and 90 kD in size prior to tick exposure (Fig. 4, Lane 1). The pre-exposure serum of Raccoons 2 and 3 (Lanes 2 and 3) developed no bands.
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4. Discussion
There was a dramatic reduction in the percentage of larvae recovered after sequential feedings (Fig. 1), indicating the development of resistance. Less than 20% of larvae applied were recovered fully engorged after the second nymphal exposure (Fig. 1). This reduction could be due to either an inflammatory response that interferes with feeding, or to an inflammatory response that increases grooming behavior resulting in removal by the host. The placement of the tick larvae and nymphs within the ears was thought to reduce the latter possibility. Either mechanism reduces parasite feeding time and prevents full engorgement. Full engorgement is required for molting to the next stadium (Allen, 1989) and 3 days of feeding is required for optimal transmission of B. burgdorferi (Piesman et al., 1987). Our findings suggest that previously exposed raccoons would be partially resistant to I. scapularis, preventing up to 85% of the feeding larvae from reaching full engorgement. The resistance to larval feeding correlates well with the development of anti-tick salivary extract antibody titers. The titer increased most dramatically following the second exposure to nymphs (Fig. 3), and the drop in the proportion of ticks feeding successfully also occurred following the second cohort of nymphs (Fig. 1). The pattem of antibody response varied somewhat among the three raccoons, which may be due to individual differences in the immune response in this outbred population. Naturally acquired immunity to ticks has been reported to vary widely in several species (Wong and Opdebeeck, 1993). Although, the correlation between tick resistance and antibody production is good, other immunologic mechanisms such as cell mediated immunity and hypersensitivity reactions are probably also involved. However, antibody titers to tick salivary gland antigens have been used as a marker of tick resistance in many species (Brown et al., 1982; Shapiro et al., 1986, 1987; Brown, 1988; Njau et al., 1988; Allen, 1989; Wheeler et al., 1989; Wong and Opdebeeck, 1993). In addition, antibodies produced by the host have been shown to pass across the gut epithelium of the tick serologically intact (Brossard and Rais, 1984; Fujisaki et al., 1984; Minoura et al., 1985). These anti-tick antibodies have been shown to retain binding activity by localizing to the tick salivary glands (Ackerman et al., 1981). Antibodies to tick gut components (regurgitated during feeding) have also been postulated to play a role in the development of resistance (Shapiro et al., 1986). However, inoculation with extracts of tick digestive tract, cephalic glands, and salivary glands showed that salivary gland extracts elicited the strongest resistance (Trager, 1939b). Raccoon 1 had an elevated anti-tick salivary gland extract antibody titer prior to tick application and was positive on Western blotting for the antigens recognized by the other raccoons only following tick exposure. This correlates well with the initial elevated titers seen on the ELISA (Fig. 3) and the reduced survival of larvae in the first four cohorts (Fig. 1) for Raccoon 1, suggesting that the antibodies detected in the Western blot and ELISA did interfere with larval feeding. Antibodies to mites have been reported to cross react with tick antigens in mice (denHollander and Allen, 1985). Interestingly, antibodies to ticks do not cross react with mite antigens (denHollander and Allen, 1986; Boyce et al., 1991). The occurrence of mite exposure in these raccoons is
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unknown, however, mites are common parasites of laboratory animals (Shadduck and Pakes, 1978). We postulate that the antibodies detected in Raccoon 1 prior to exposure to the ticks were the result of previous mite infestation, since ticks are very uncommon in laboratory animals (Shadduck and Pakes, 1978). Some of the variation in antibody responses could be due to differences in antigens expressed by the different stages of ticks. Adult salivary glands were used as antigen in the ELISA, but larvae and nymphs were used in the experiment to measure feeding success. The raccoons developed resistance to larvae after four cohorts, but not to nymphs, which were only placed on the animals twice. Although data on this subject are scant, some prior research has suggested that many major antigens in adult salivary glands are not present in larval and nymphal salivary glands of ixodid ticks (Shapiro et al., 1986). The current study, in using adult salivary gland extract to evaluate antibody response to feeding by larvae and nymphs, suggests that there are either important proteins common to the glands of larvae and adults, or alternatively, that there are cross-reacting antibodies to antigenically similar, but not identical, proteins among the stages of I. scapularis. Many of the proteins secreted by feeding ticks have anti-coagulant, anti-histamine, and anti-inflammatory effects. These proteins facilitate blood feeding during the prolonged period of host attachment by antagonizing the chemical mediators of the host inflammatory response (Ribeiro et al., 1985). Resistance has not been found in the ticks' natural host species (Randolph, 1979), suggesting that the anti-inflammatory proteins in the tick saliva specifically antagonize the mediators of inflammation and resistance in the natural host (Ribeiro et al., 1985). In the absence of this specific interaction, i.e. in an unnatural host, inflammation and an immune response specific for tick salivary antigens results. The results of this study suggest that the raccoon is not a natural host for L scapularis. The role of raccoons in the maintenance of populations of I. scapularis and the transmission of B. burgdorferi is questionable. Although antibodies to B. burgdorferi have been found in raccoons (Magnarelli et al., 1991), the current study suggests that they develop resistance to the vector tick species. Specifically, the duration of feeding for most larvae would not be long enough in immune animals to efficiently transmit the spirochete. The longer feeding times on relatively naive raccoons and the ticks that do escape the immune mechanisms to feed to full engorgement may allow transmission of the spirochete, explaining the presence of antibodies in wild raccoon populations.
Acknowledgments The authors would like to thank Charles S. Apperson and Bruce F. Levine for their assistance. This research was supported by grants from the National Institute of Allergy and Infectious Diseases R55 AI 30042 and R29 AI 31608 and from the National Institutes of Health RR00130 and RR07002.
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Randolph, S.E., 1979. Population regulation in ticks: the role of acquired resistance in natural and unnatural hosts. Parasitology, 79: 141-156. Ribeiro, J.M., Makoul, G.T., Levine, J., Robinson, D.R. and Spielman, A., 1985. Antihemostatic, antiinflammatory, and immunosuppressive properties of the saliva of a tick, lxodes dammini. J. Exp. Med., 161: 332-344. Roberts, J.A. and Kerr, J.D., 1976. Boophilus microplus: Passive transfer of resistance in cattle. J. Parasitol., 62: 485-489. Schwartz, B.S., Ribeiro, J.M.C. and Goldstein, M., 1990. Anti-tick antibodies: an epidemiologic tool in Lyme disease research. Am. J. Epidemiol., 132: 58-66. Schwartz, B.S., Ford, D.P., Childs, J.E., Rothman, N. and Thomas, R.J., 1991. Anti-tick saliva antibody: A biological marker of tick exposure that is a risk factor for Lyme disease seropositivity. Am. J. Epidemiol., 134: 86-95. Schwartz, B.S., Nadelman, R.B., Fish, D., Childs, J.E., Forester, G. and Wormser, G., 1993. Entomologic and demographic correlates of anti-tick saliva antibody in a prospective study of tick bite subjects in Westchester County, New York. Am. J. Trop. Med. Hyg., 48: 50-57. Shadduck, J.A. and Pakes, S.P., 1978. Protozoal and metazoal diseases. In: K. Benirschke, F.M. Garner and T.C. Jones (Editors), Pathology of Laboratory Animals. Springer-Verlag, New York, pp. 1587-1696. Shapiro, S.Z., Voigt, W.P. and Fujisaki, K., 1986. Tick antigens recognized by serum from a guinea pig resistant to infestation with the tick Rhipicephalus appendiculatus. J. Parasitol., 72: 454-463. Shapiro, S.Z., Buscher, G, and Dobbelaere, D.A.E., 1987. Acquired resistance to Rhipicephalus appendiculatus (Acari: Ixodidae): Identification of an antigen eliciting resistance in rabbits. J. Med. Entomol., 24: 147-154. Trager, W., 1939a. Acquired immunity to ticks. J. Parasitol., 25: 57-81. Trager, W., 1939b. Further observations on acquired immunity to the tick Dermacentor variablilis Say. J. Parasitol., 25: 137-139. Wheeler, C.M., Coleman, J.L., Habicht, G.S. and Benach, J.L., 1989. Adult lxodes dammini on rabbits: Development of acute inflammation in the skin and immune responses to salivary gland, midgut, and spirochetal components. J. Infect. Dis., 159: 265-273. Wong, J.Y.M. and Opdebeeck, J.P., 1993. Immunity in vaccinated cattle exposed to experimental and natural infestations with Boophilus microplus. Int. J. Parasitol., 23: 689-692.