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ABSTRACT |
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104 PFU by needle was required to attain the high early viremia observed in mosquito-infected chickens. Mosquitoes inoculated WNV at this level as estimated by feeding on a hanging drop of blood (mean: 102.5, range: 100.7–104.6 PFU). These results indicate that enhanced early infection in mosquito-infected chickens may be explained by higher viral dose delivered by mosquitoes. On the other hand, chickens infected by multiple mosquitoes (N = 3–11) had viremic titers that were 25–50 times higher at 6 and 12 hours PF than in chickens infected by a single mosquito, suggesting that viral dose is not the only factor involved in enhanced early infection. The likelihood that enhanced early infection in mosquito-infected chickens is due to a higher viral dose inoculated by mosquitoes and/or other factors (saliva, inoculation location, or viral source) is discussed. |
INTRODUCTION |
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Although needle inoculation has been accepted as a practical alternative to mosquito transmission of viruses, it is an imperfect substitute4 due to the mechanics of mosquito blood feeding. Mosquitoes alight on a potential host and actively probe for blood with their mouthparts throughout the dermal layer of skin. After a blood vessel is pierced, the mosquito begins feeding either directly from the vessel or from the resulting hemorrhagic pool. During the probing and feeding process, the mosquito ejects saliva that contains virus (if the mosquito is infectious), along with pharmacologically active molecules that counteract the host hemostatic response, reduce inflammation, and alter host immunity.5–12
Vertebrates infected with arboviruses by mosquito bite or when associated with mosquito saliva have been shown to exhibit enhanced infection compared with those infected by needle inoculation. Mice inoculated intradermally with WNV in an area where ~11 uninfected mosquitoes had fed had lower survival rates, higher and longer-lasting viremia, and accelerated neuroinvasion.13 Mice infected with vesicular stomatitis virus (VSV) by mosquitoes had higher seroconversion rates than those infected by needle inoculation.14 Chipmunks and deer infected with La Crosse virus by mosquito bite had higher and longer-lasting viremia than those infected by intramuscular inoculation.15 Mice inoculated subcutaneously with Cache Valley virus in an area where uninfected mosquitoes had recently fed exhibited higher infection rates and antibody response than those inoculated without prior mosquito feeding.16
On the other hand, other studies have shown no effect of mosquito transmission on vertebrate infection. One-week-old chickens and adult house finches infected by needle or mosquito bite with St. Louis encephalitis (SLE) or western equine encephalomyelitis (WEE) viruses did not differ significantly in viremic level or antibody response.17 Additionally, no difference in viremia was observed in hamsters18 or chickens19 infected with WNV by needle or mosquito inoculation, but early time points were not included in these studies.
The goal of this study was to further explore mosquito enhancement of arboviral infections in vertebrates. Young chickens were used as an avian model for the WNV enzootic transmission cycle. Viremia and viral shedding were compared for chickens infected with WNV by mosquito bite or subcutaneous needle inoculation of 103 plaque-forming units (PFU). In addition, the effect of viral dose on infection was examined.
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METHODS |
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Animals. We used a Cx. pipiens colony established in 2000 from mosquitoes collected in Albany, NY and a Cx. tarsalis colony that was derived from the Bakersfield, CA colony established in 1953 (kindly provided by Dr. William Reisen). Specific pathogen-free chickens (Charles River SPAFAS, North Franklin, CT) were housed in a BSL-3 animal facility. The use of chickens in this experiment was approved and conducted in accordance with the Wadsworth Center Institutional Animal Care and Use Committee.
Infection of chickens with West Nile virus by needle or mosquito. Mosquitoes were infected with WNV by intrathoracic inoculation of ~30 PFU WNV 7 days (Cx. pipiens) prior to feeding on 1-day-old chickens and 6 days (Cx. tarsalis) or 11 days (Cx. pipiens) prior to feeding on 5-day-old chickens. The amount of virus expelled by WNV-inoculated Cx. pipiens and Cx. tarsalis as they fed on hanging blood drops did not vary significantly between day 7–10 post-inoculation (PI) (data not shown). After inoculation, mosquitoes were held in 0.5L cardboard cartons with a mesh top at 27°C, and provided with 10% sucrose via a soaked cotton pad. Mosquitoes were starved by removing the sucrose pad from cartons for 24–48 hours prior to feeding on chickens.
One-day-old and 5-day-old chickens were infected with WNV by subcutaneous needle inoculation of ~103 PFU WNV in the lateral neck, or by allowing WNV-infected mosquitoes to feed on a restrained chicken for ~1 hour. A previous study showed that mosquitoes inoculate a median of 103–104 PFU and mean of 104.3 PFU WNV during in vitro transmission assays.20 Chickens were exposed to either a single or multiple (up to 12) infected mosquitoes. After the feeding period, mosquitoes exposed to each chicken were frozen, and the number of mosquitoes with blood in their abdomen was determined using a dissecting microscope. This procedure allowed us to ascertain the minimum number of mosquito bites that each chicken received. We were not able to detect mosquitoes that only had probed. Three replicates of this experiment were completed using the previously described protocol: two with WNV-infected Cx. pipiens and one with WNV-infected Cx. tarsalis.
Sample collection and processing. At various times post-feeding (PF) or PI, blood, oral swab, and cloacal swab samples were collected from chickens. Blood samples were collected from the ulnar vein into a microtiter centrifuge tube, held at 4°C for up to 3 hours and centrifuged (8,000 rpm, 5 min). Serum was removed, diluted 1:10 with BA-1 diluent (M199H, 1% bovine serum albumin, 0.05 M Tris pH 7.6, 0.35 g/L sodium bicarbonate, 100 u/ml penicillin, 100 µg/ml streptomycin, 1 µg/ml fungizone), and stored at –80°C. Oral and cloacal swabs were obtained by swabbing the inside of the mouth or cloaca, respectively, with a cotton-tipped applicator stick, placing the swab into 500 µL BA-1 diluent, and storing at –80°C. All samples were titrated by plaque assay on Vero cells. After virus titers had been determined, serum samples were heat-inactivated at 56°C for 30 minutes, and tested for WNV-specific IgG and IgM antibodies by indirect IgG and IgM ELISA, respectively. We followed the IgG ELISA protocol of Ebel and coworkers,21 with a minor change in the blocking buffer from 2.0% casein to 5.0% skim milk. IgM ELISA followed the same protocol, except that horseradish peroxidase-conjugated goat anti-chicken IgM (Bethyl Laboratories, Montgomery, TX) was used. End point titrations were conducted by plaque reduction neutralization assay (PRNT90) on samples positive for IgG (positive antigen wells/negative antigen wells > 2.0).
Chickens inoculated with successively higher viral doses. We investigated the effect of viral dose on viral replication in chickens. Groups of five 5-day-old chickens were inoculated subcutaneously in the lateral neck with successively higher doses of WNV from 101–107 PFU. Serum and oral swab samples were taken as described previously at various times PI and were tested for infectious virus by plaque assay.
Viral titers inoculated by mosquitoes. Viral titers inoculated by mosquitoes were estimated by allowing mosquitoes to feed on hanging blood drops.22 Cx. tarsalis and Cx. pipiens were intrathoracically inoculated with ~30 PFU WNV, maintained in 0.5L cardboard cartons with a mesh top at 27°C for 6–8 days, and provided with 10% sucrose via a soaked cotton pad. Mosquitoes were starved by removing sucrose from the cartons at least 24 hours prior to blood feeding. On days 7–10 PI, mosquitoes were placed individually into 0.5L cardboard cartons, and a 30-uL drop of sweetened goose blood (1 part 50% sucrose, 24 parts defibrinated goose blood) was pipetted onto the mesh top of each carton. Mosquitoes were allowed 45–60 minutes to feed on the blood drop, after which the blood remaining in each drop was collected and placed into 500 µL of BA-1 diluent. Mosquitoes were observed throughout the feeding period; probing (mouthparts in contact with blood droplet) and engorgement status of each female was recorded. Blood drop samples were titrated by plaque assay on Vero cells.
Statistical analysis. Serum, swab, and hanging drop titers were log transformed and checked for normality using Shapiro-Wilk and Kolmogorov-Smirnov statistics. Negative serum samples were given a mathematical value of 50 PFU/ml in statistical analyses due to the high limit of detection for virus in serum samples (100 PFU/ml). However, negative samples were set at 0 PFU/ml in the graphs for clarity. Because multiple serum and swab samples were taken from individual chickens, we used repeated measures ANOVA (PROC GLM)23 to determine whether viral titers varied by infection method (needle versus mosquito bite), mosquito exposure level (single or multiple mosquito bites), and inoculum dose for each age group of chickens. The Tukey-Kramer method was used to adjust for multiple comparisons. Because of the small sample size, Fisher’s exact test was used to determine significant differences between proportions of positive IgM and IgG samples.
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RESULTS |
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1 mosquito) (Table 1
).
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). Although it is possible that the delayed viremia was due to a low dose inoculated by a probing mosquito, a more likely scenario is that this chicken was infected via intracage transmission on day 1 PF. At that time, 3 cage mates of this chicken were shedding 103–104 PFU of virus orally and 101–103 PFU of virus via the cloaca.
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). The uninfected chicken, one of 12 uninfected chickens in the 1-day-old group, died on day 3 PF. This death was presumed to be due to failure to thrive. The infected chickens died on days 6–8 PF or PI; 4 were infected by mosquito bite, and 1 was infected by needle.
Age of the chicken had an impact on viral infection (see Figure 1). Peak titers in sera and swabs were ~10 fold lower in 5-day-old chickens than in 1-day-old chickens. Five-day-old chickens cleared virus ~2 days faster from sera and from oral and cloacal cavities. Age also had an impact on immune response. A greater percentage of 5-day-old chickens had detectable IgM and IgG antibodies at days 6–7 PF or PI compared with the younger chickens (IgM: 89% versus 17%, Fisher’s exact test, P = 0.01; IgG: 82% versus 38%, Fisher’s exact test, P = 0.07) (Figure 2A and 2B). Neutralizing antibody titers were also observed to rise later in 1-day-old chickens than in 5-day-old chickens (Figure 2C). Although a greater percentage of infected chickens from the 1-day-old group died (31%, N = 13) compared with the 5-day-old group (7%, N = 27), this difference was not statistically significant (Fisher’s exact test: P = 0.08).
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). Similarly, oral swab titers of chickens infected by Cx. pipiens were as much as 1,000 fold higher than titers in needle-inoculated chickens at 12, 24, and 48 hours PI (12h: P < 0.0001; 24h: P < 0.0001; 48h: P = 0.038). Cloacal swab titers of mosquito-infected chickens were significantly higher at 24 hours PF than those of needle-inoculated chickens (P < 0.0001).
A similar pattern of enhanced early infection was observed in older chickens. Serum titers in 5-day-old chickens infected by mosquito bite were significantly higher at 12 and 24 hours PF than in needle-inoculated chickens (Cx. pipiens versus needle 12h: P = 0.001, 24h: P < 0.0001; Cx. tarsalis versus needle 12h: P = 0.003; 24h: P < 0.0001) (see Figure 1). At 24 hours, oral and cloacal swab titers of 5-day-old chickens infected by mosquito bite were ~10 fold higher than in needle-inoculated chickens (oral: Cx. pipiens versus needle P = 0.0001; Cx. tarsalis versus needle P = 0.0007; cloacal: Cx. pipiens versus needle P = 0.0095).
There was some indication of more rapid viral clearance in chickens infected by mosquito bite. Viremia in 5-day-old chickens infected by Cx. pipiens was significantly lower at 48 and 72 hours post feeding than in needle-inoculated chickens (48h: P = 0.01, 72h: P = 0.02). Additionally, oral swab titers were significantly lower at 96 hours post feeding compared with 5-day-old chickens infected by needle (P = 0.0007). However, significantly faster viral clearance was not seen in 1-day-old chickens infected by Cx. pipiens or in 5-day-old chickens infected by Cx. tarsalis.
Five-day-old chickens infected by two different mosquito species had similar infection profiles (see Figure 1). There were no significant differences in serum titers between 5-day-old chickens infected by Cx. pipiens or by Cx. tarsalis at 12, 24, 72, and 96 hours PF. However, at 48 hrs PF, chickens infected by Cx. tarsalis had higher serum titers than chickens infected by Cx. pipiens (P = 0.0496). Oral swab titers were not significantly different between chickens infected by Cx. pipiens or by Cx. tarsalis, except at 96 hours PF when oral swab titers of chickens infected by Cx. tarsalis were higher than in those infected by Cx. pipiens (P = 0.0144). It is important to note that these differences in serum titers at 48 hours were confounded by the number of mosquitoes feeding (see analysis later in this article).
There were no significant differences in PRNT90 titers between chickens infected by mosquito and those infected by needle in each age group (see Figure 2C). However, a greater proportion of 5-day-old chickens infected by mosquito had detectable IgM antibody on day 4 PI (50%, N = 14) compared with 5-day-old chickens infected by needle (9%, N = 11) (Fisher’s exact test: P = 0.04) (see Figure 2A).
Chickens infected by a single mosquito or multiple mosquitoes.
We compared serum and swab titers of 5-day-old chickens infected by a single mosquito or multiple mosquitoes, as defined in Table 1. Chickens infected by multiple mosquitoes had serum titers that were ~25 times higher at 12 hours (P = 0.014) and ~3 times higher at 24 hours (P = 0.029), compared with chickens infected by a single mosquito (Figure 3). Serum titers of chickens infected by multiple mosquitoes were 100 fold lower at 72 hours (P = 0.042). When this restricted data set was used, differentiating single from multiple mosquito bites, there was no difference between serum titers of chickens infected by Cx. pipiens and by Cx. tarsalis. Similarly, oral swab titers of chickens infected by multiple mosquitoes were lower at 72 hours (P = 0.034) and higher at 24 hours than those infected by a single mosquito; however, the difference at 24 hours was not significant. Chickens infected by multiple mosquitoes also had significantly higher cloacal swab titers at 24 hours (P = 0.039) compared with chickens infected by single mosquitoes.
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). When compared with data from the previous experiment, serum and oral swab titers of chickens infected by a single mosquito were most similar to those from chickens inoculated with 107 PFU by needle. There was no statistical difference, however, between WNV titers in chickens infected by a single mosquito and chickens inoculated with 104 PFU at 6–24 hours PI. Serum titers in chickens infected by multiple mosquitoes were 25–50 times higher than all other groups at 6 and 12 hours PI (P < 0.04); there was no significant difference between chickens infected by multiple mosquitoes and those inoculated with 104 PFU at 24 hours PI (see Fig. 4).
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). Geometric mean virus titer and range of titers expelled by blood fed mosquitoes did not differ significantly by species (Cx. pipiens: mean = 102.8, range = 100.7–104.3 PFU, Cx. tarsalis: mean = 102.0, range = 100.7–104.6 PFU). Median titer expelled by blood fed Cx. pipiens (103.1 PFU) was higher than that expelled by blood fed Cx. tarsalis (101.8 PFU).
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DISCUSSION |
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These results are in contrast to two previous studies with WNV. In a hamster model, no difference in viremia was observed on days 1–3 in hamsters infected with WNV by needle or mosquito inoculation, but time points earlier than 24 hours were not evaluated.18 Another study reported no difference in viremia in older chickens (17–60 weeks) infected with WNV by needle or mosquito bite; however, viremic levels were low (< 104 PFU/ml) and were not measured at early time points post-inoculation (< 24 hr).19 Thus, the differences between these two studies and our current results are most likely explained by differences in sampling times and/or animal models.
Enhanced early WNV infection in mosquito-infected chickens may partially be explained by higher viral dose delivered by mosquitoes. Higher doses inoculated by needle resulted in more rapid development of viremia and oral shedding. On the other hand, a needle inoculation of 107 PFU was required to attain the high early viremia observed in chickens infected by single mosquitoes (although there was no statistical difference between chickens infected by a single mosquito and those inoculated with 104 PFU at 6–24 hours). This result suggests that mosquitoes need to inoculate at least 104 PFU consistently while probing and feeding if more rapid development of viremia is due only to high viral dose.
The amount of WNV inoculated by mosquitoes while probing or feeding on a live host is not known. Our data using the hanging drop method suggest that mosquitoes inoculate a wide range of viral titers (100.7–104.6, mean = 102.5 PFU) while blood feeding. Similar results were reported by Vanlandingham and colleagues,20 who observed that mosquitoes inoculate a range of 100.5–105.3 PFU (mean = 104.3) of WNV as measured by in vitro capillary tube transmission assay. Other studies with WEE virus,17 SLE virus,17 and VSV,14 using in vitro capillary tube transmission assays, have reported wide ranges and similar average salivary secretion titers of 102–104 PFU. Despite these consistent results for several arboviruses, in vitro assays may underestimate the inoculated dose and introduce variability because mosquitoes are not feeding normally in these assays. The hanging drop method that was used in this study may be more accurate because mosquitoes are able to probe and feed, but this method still does not mimic natural feeding completely because mosquitoes do not need to find a blood vessel or overcome the host hemostatic response. A previous study showed that significantly fewer Cx. tritaeniorhynchus transmitted WNV when feeding on a hanging blood drop than they did when feeding directly on a suckling mouse.24 In addition, mosquitoes probe for a shorter time (and presumably excrete less saliva) when feeding through a membrane than they do when feeding on an intact host.25 One study quantified the amount of Rift Valley fever virus inoculated by Cx. pipiens into a live host (median: 102.5, range: < 100.7–103.7 PFU); however, mosquitoes in this study were only allowed to feed or probe for 30 seconds and this study did not account for virus that may have bound to the cells prior to assay and thus was not detectable as infectious virus.26,27 We are currently conducting studies to determine the amount of WNV inoculated into live hosts. Preliminary results suggest that mosquitoes routinely inoculate between 104 and 105 PFU and can inoculate as much as 106.6 PFU while probing and feeding on a live host (unpublished data).
Overall our current data and studies by others indicate that mosquitoes routinely inoculate between 102 and 105 PFU and may inoculate as much as 106.6 PFU of various arboviruses, suggesting that the enhanced early infection in mosquito-infected chickens may be due to viral dose. However, two lines of evidence suggest that other factors are involved. First, viral dose cannot explain the high sera titers seen in chickens exposed to multiple infected mosquitoes (N = 3–11). Assuming that a single mosquito delivers a dose of ~106 PFU while feeding (a high estimate), the maximum dose inoculated by 11 mosquitoes is ~107 PFU. However, in our study chickens exposed to multiple mosquitoes developed sera titers that were 25–50 times higher than chickens inoculated with 107 PFU at 6 and 12 hours PI. Second, all chickens infected by mosquito feeding, or by mosquito probing without feeding, exhibited consistently high viremia and viral shedding titers at early time points. One would expect more variation in viremia and shedding titers if dose was the only cause for earlier development of viremia, especially considering that some mosquitoes only probed, whereas others both probed and fed during a 1-hour period.
Other differences, besides inoculum dose, exist between WNV infection via needle and mosquito bite that could explain the enhanced early infection observed in mosquito-infected chickens. First, mosquito saliva has been shown in previous studies to cause potentiation of viral infection,7,13,16 perhaps through its ability to modulate host immune response at the inoculation site. Mouse fibroblast cells treated with salivary gland homogenate had lower interferon 
/ß production and higher VSV growth kinetics than did untreated cells.7 Mosquito salivary gland extract and saliva also suppress murine splenocyte proliferation and alter cytokine production.8–12 A recent study demonstrated the important effect of mosquito feeding and salivary gland extracts on WNV infection. Mice fed upon by uninfected Aedes aegypti females prior to intradermal inoculation of WNV had higher WNV RNA titers at the inoculation site and draining lymph node, higher and longer lasting viremia, and more rapid neuroinvasion compared with mice inoculated with WNV without mosquito feeding.13 It is not known if potentiation due to mosquito saliva also occurs within the enzootic WNV cycle that involves Culex mosquitoes and avian hosts.
A second difference between mosquitoes and needles is viral inoculation site. Mosquitoes salivate (and, therefore, deposit virus) while probing and feeding throughout the dermal tissue, whereas subcutaneous needle inoculation deposits the entire inoculum in one location below the dermis. Although studies have suggested that mosquitoes inject most virus extravascularly,26,27 it is possible that some virus is injected intravascularly by mosquitoes while blood feeding. Faster introduction of virus into the blood stream by mosquitoes could result in earlier dissemination and higher early viremia in mosquito-infected chickens. In addition, multiple mosquitoes feeding on a single chicken may inoculate virus into multiple locations, resulting in more rapid initial viral replication. Increased viral replication at multiple sites could explain the high initial viremia that we observed in chickens bitten by multiple mosquitoes.
The final difference between viral infection by mosquito bite and needle inoculation is viral source. In this study, virus injected by needle was harvested from Vero cell culture, whereas virus inoculated by mosquitoes had replicated in mosquito cells. Differences in viral glycoprotein maturation have been observed between virus derived from invertebrate cells and from vertebrate cells.28,29 If these differences in viral glycoproteins alter viral entry into primary infection sites, it could affect viral amplification rates and thus early levels of viremia and viral shedding. In addition, mutations may occur during replication in the mosquito that can affect early viral replication.
West Nile virus infection in chickens is age dependent, perhaps due to age-related differences in the immune response. One-day-old chickens exhibited levels of viremia and viral shedding that peaked ~1 day later and ~10 fold higher than in 5-day-old chickens. Increased production of virus may have been due to slower antibody response in 1-day-old chickens; IgG, IgM, and neutralizing antibody titers of 1-day-old chickens were detectable 1–6 days later than when they were detectable in 5-day-old chickens. In studies by others, older chickens (7–60 weeks old) infected with WNV exhibited viral titers that peaked at a lower level (~105 PFU/ml).19,30 In addition, no significant difference was found between older chickens (17–60 weeks) infected by needle and those infected by mosquito bite; perhaps this lack of difference was due to lower overall viremic levels.19
One 1-day-old chicken, whose level of viral viremia and viral shedding rose 1–2 days later than other chickens, was presumably infected by intracage transmission. This chicken was housed with 3 chickens that were shedding 103–104 PFU of virus orally and 101–103 PFU of virus through the cloaca at the presumed time of infection. A previous study reported intracage transmission of WNV to a 20-week-old chicken; however, cloacal and oral swab titers of the infected cage mate were not recorded prior to the transmission event.19 Intracage transmission of WNV has also been reported in geese, ring-billed gulls, blue jays, black-billed magpies, and American crows.31,32 These species have been shown to shed up to 103.5–105.7 PFU orally and 102.4–106.0 PFU through the cloaca.32
Mosquitoes in our study were infected with WNV by intrathoracic inoculation to obtain mosquitoes that transmitted virus with ~100% efficiency. Because every mosquito was theoretically able to transmit virus, fewer animals were needed per treatment and mosquito-to-mosquito variability was reduced, leading to a more controlled study. Despite virus reaching the salivary glands more quickly in inoculated mosquitoes, no difference was found in the amount of virus expelled by perorally infected and inoculated mosquitoes into hanging drops or into live hosts (data not shown).
Previous studies showing mosquito enhancement of arbovirus infection used primarily negative-sense RNA viruses belonging to the virus families Rhabdoviridae and Bunyaviridae. Our study demonstrated mosquito bite enhancement of a positive-sense RNA virus (Flaviviridae) using a natural host and mosquito vector, and it is the first to demonstrate significant differences in viremia and viral shedding between needle-inoculated and mosquito-infected hosts. Although higher viral doses inoculated by mosquitoes may partially explain enhanced early WNV infection in chickens, other factors (mosquito saliva, differences in inoculum site, and viral source) could also play an important role. Future studies are planned to evaluate these possible factors.
Received April 6, 2005. Accepted for publication April 17, 2006.
Acknowledgments: The authors acknowledge the excellent technical assistance provided by Matthew Jones, Jennifer Longacker, Amy Lovelace, Christine Lussier, Sarah Sperry, and David Young. The authors thank the Wadsworth Center Tissue Culture Facility for providing cell culture support.
Financial support: This project has been funded in part with Federal funds from the National Institute of Allergy and Infectious Disease, National Institutes of Health, under Contract No. N01-AI-25490. The BSL-3 animal facility at the Wadsworth Center was used, which is funded in part by the animal core on the NIH/NIAID award U54A17158.
* Address correspondence to Laura D. Kramer, Arbovirus Laboratories, Wadsworth Center, New York State Dept of Health, 5668 State Farm Road, Slingerlands, NY 12159. E-mail: Kramer{at}wadsworth.org 
Authors’ addresses: Laura D. Kramer, Arbovirus Laboratories, Wadsworth Center, New York State Dept of Health, 5668 State Farm Road, Slingerlands, NY 12159, Telephone: (518) 869-4524, Fax: (518) 869-4530, E-mail: kramer{at}wadsworth.org. Linda M. Styer, Arbovirus Laboratories, Wadsworth Center, New York State Dept of Health, 5668 State Farm Road, Slingerlands, NY 12159, Telephone: (518) 862-4306, Fax: (518) 869-4530, E-mail: lstyer{at}wadsworth.org. Kristen A. Bernard, Arbovirus Laboratories, Wadsworth Center, New York State Dept of Health, 5668 State Farm Road, Slingerlands, NY 12159, Telephone: (518) 869-4519, Fax: (518) 869-4530, E-mail: kbernard{at}wadsworth.org.
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