Wild waterfowl are maintenance hosts of most influenza A virus (IAV) subtypes and are often the subjects of IAV surveillance and transmission models. While maternal antibodies have been detected in yolks and in nestlings for a variety of wild bird species and pathogens, the persistence of maternal antibodies to IAVs in mallard ducklings (Anas platyrhynchos) has not been previously investigated. Nonetheless, this information is important for a full understanding of IAV transmission dynamics because ducklings protected by maternal antibodies may not be susceptible to infection. In this study, we examined the transfer of IAV-specific maternal antibodies to ducklings. Blood samples were collected approximately every five days from ducklings hatched from hens previously infected with an H6 strain of IAV. Serum samples were tested for antibodies to IAV by an enzyme-linked immunosorbent assay. The median persistence of maternal antibodies in ducklings was 12.5 days (range: 4-33 days) post-hatch. The majority of ducklings (71%) had detectable maternal antibodies from 4 to 17 days post-hatch, while a small subset of individuals (29%) had detectable maternal antibodies for up to 21-33 days post-hatch. Antibody concentrations in hens near the time of egg laying were correlated with maternal antibody concentrations in the initial blood sample collected from ducklings (0-4 days post-hatch). Knowledge of the duration of maternal antibodies in ducklings will aid in the interpretation of IAV serological surveillance results and in the modeling of IAV transmission dynamics in waterfowl.

Influenza A viruses (IAVs) have been reported in wild lagomorphs in environments where they share resources with waterfowl. Recent studies have conclusively shown that a North American lagomorph, cottontail rabbits (Sylvilagus sp.), become infected following exposure to IAVs and can shed significant quantities of virus. However, the minimum infectious dose and the efficiency of various routes of infection have not been evaluated. Thirty-six cottontail rabbits were used in a dose response study assessing both the oral and nasal routes of infection. The nasal route of infection proved to be the most efficient, as all cottontail rabbits shed viral RNA following inoculation with doses as low as 102 EID50. The oral route of infection was less efficient, but still produced infection rates of ≥ 50% at relatively low doses (i.e., 103 and 104 EID50). These results suggest that cottontail rabbits are highly susceptible to IAVs at low exposure doses that have been routinely observed in environments contaminated by waterfowl. Furthermore, this study supports earlier observations that cottontail rabbits may pose a biosecurity risk to poultry operations, as a virus-contaminated water source or contaminated environment, even at low viral titers, could be sufficient to initiate viral replication in cottontail rabbits.

In November 2014, a Eurasian strain H5N8 highly pathogenic avian influenza virus was detected in poultry in Canada. Introduced viruses were soon detected in the United States and within six months had spread to 21 states with more than 48 million poultry affected. In an effort to study potential mechanisms of spread of the Eurasian H5 virus, the United States Department of Agriculture coordinated several epidemiologic investigations at poultry farms. As part of those efforts, we sampled synanthropic birds and mammals at five infected and five uninfected poultry farms in northwest Iowa for exposure to avian influenza viruses. Across all farms, we collected 2,627 samples from 648 individual birds and mammals. House mice were the most common mammal species captured while house sparrows, European starlings, rock pigeons, swallows, and American robins were the most commonly captured birds. A single European starling was positive for Eurasian H5 viral RNA and seropositive for antibodies reactive to the Eurasian H5 virus. Two American robins were also seropositive. No mammal species showed evidence of infection. These results indicate synanthropic species merit further scrutiny to better understand potential biosecurity risks. We propose a set of management practices aimed at reducing wildlife incursions.

Following a 2008 outbreak of North American low-pathogenic H5N8 influenza A virus at an upland gamebird farm, we sero-sampled rock doves (pigeons, Columba livia) at the outbreak site and conducted experimental inoculations of wild-caught pigeons using the H5N8 virus and another low-pathogenic virus (H4N6). While 13% of pigeons at the outbreak site were seropositive, none were positive for exposure to H5, and one was positive for N8. Challenged pigeons exhibited low susceptibility and limited viral RNA excretion for both viruses tested, but at least one individual had RNA loads indicative of the potential for viral transmission to other birds.

The availability of a validated commercial assay is an asset for any wildlife investigation. However, commercial products are often developed for use in livestock and are not optimized for wildlife. Consequently, it is incumbent upon researchers and managers to apply commercial products appropriately to optimize program outcomes. We tested more than 800 serum samples from mallards for antibodies to influenza A virus with the IDEXX AI MultiS-Screen Ab test to evaluate assay performance. Applying the test per manufacturer's recommendations resulted in good performance with 84% sensitivity and 100% specificity. However, performance was improved to 98% sensitivity and 98% specificity by increasing the recommended cut-off. Using this alternative threshold for identifying positive and negative samples would greatly improve sample classification, especially for field samples collected months after infection when antibody titers have waned from the initial primary immune response. Furthermore, a threshold that balances sensitivity and specificity reduces estimation bias in seroprevalence estimates.

The potential role of wild mammals in avian influenza A virus (IAV) transmission cycles has received some attention in recent years and cases where birds have transmitted IAV to mammals have been documented. However, the contrasting cycle, wherein a mammal could transmit an avian IAV to birds, has been largely overlooked. We experimentally tested the abilities of two mammalian species to transmit avian IAV to mallards (Anas platyrhynchos) in simulated natural environments. Results suggested that striped skunks (Mephitis mephitis) can successfully transmit avian IAV to mallards through indirect contact with shared resources, as transmission was noted in 1 of 4 of the mallards tested. Cottontail rabbits (Sylvilagus sp.) exhibited a similar pattern, as one of five cottontail rabbits successfully transmitted IAV to a mallard, likely through environmental contamination. For each mammalian species tested, the mallards that became infected were those paired with the individual mammals with the lowest shedding levels but were anecdotally observed to be the most active animals. Mammals associated with and around poultry rearing facilities should be taken into consideration in biosecurity plans.

Background Striped skunks (Mephitis mephitis) are susceptible to infection with some influenza A viruses. However, the viral shedding capability of this peri-domestic mammal and its potential role in influenza A virus ecology are largely undetermined. Methodology/Principal Findings Striped skunks were experimentally infected with a low pathogenic (LP) H4N6 avian influenza virus (AIV) and monitored for 20 days post infection (DPI). All of the skunks exposed to H4N6 AIV shed large quantities of viral RNA, as detected by real-time RT-PCR and confirmed for live virus with virus isolation, from nasal washes and oral swabs (maximum ≤106.02 PCR EID50 equivalent/mL and ≤105.19 PCR EID50 equivalent/mL, respectively). Some evidence of potential fecal shedding was also noted. Following necropsy on 20 DPI, viral RNA was detected in the nasal turbinates of one individual. All treatment animals yielded evidence of a serological response by 20 DPI. Conclusions/Significance These results indicate that striped skunks have the potential to shed large quantities of viral RNA through the oral and nasal routes following exposure to a LP AIV. Considering the peri-domestic nature of these animals, along with the duration of shedding observed in this species, their presence on poultry and waterfowl operations could influence influenza A virus epidemiology. For example, this species could introduce a virus to a naive poultry flock or act as a trafficking mechanism of AIV to and from an infected poultry flock to naive flocks or wild bird populations.

Background Cottontails (Sylvilagus spp.) are common mammals throughout much of the U.S. and are often found in peridomestic settings, potentially interacting with livestock and poultry operations. If these animals are susceptible to avian influenza virus (AIV) infections and shed the virus in sufficient quantities they may pose a risk for movement of avian influenza viruses between wildlife and domestic animals in certain situations. Methodology/Principal Findings To assess the viral shedding potential of AIV in cottontails, we nasally inoculated fourteen cottontails with a low pathogenic AIV (H4N6). All inoculated cottontails shed relatively large quantities of viral RNA both nasally (≤106.94 PCR EID50 equivalents/mL) and orally (≤105.09 PCR EID50 equivalents/mL). However, oral shedding tended to decline more quickly than did nasal shedding. No animals showed any obvious signs of disease throughout the study. Evidence of a serological response was found in all infected rabbits at 22 days post infection in convalescent sera. Conclusions/Significance To our knowledge, cottontails have not been previously assessed for AIV shedding. However, it was obvious that they shed AIV RNA extensively via the nasal and oral routes. This is significant, as cottontails are widely distributed throughout the U.S. and elsewhere. These mammals are often found in highly peridomestic situations, such as farms, parks, and suburban neighborhoods, often becoming habituated to human activities. Thus, if infected these mammals could easily transport AIVs short distances.

BACKGROUND:Wild raccoons have been shown to be naturally exposed to avian influenza viruses (AIV). However, the mechanisms associated with these natural exposures are not well-understood.METHODOLOGY/PRINCIPAL FINDINGS:We experimentally tested three alternative routes (water, eggs, and scavenged waterfowl carcasses) of AIV transmission that may explain how raccoons in the wild are exposed to AIV. Raccoons were exposed to 1) water and 2) eggs spiked with an AIV (H4N6), as well as 3) mallard carcasses experimentally inoculated with the same virus. Three of four raccoons exposed to the high dose water treatment yielded apparent nasal shedding of >10(2.0) PCR EID50 equivalent/mL. Little to no shedding was observed from the fecal route. The only animals yielding evidence of serologic activity during the study period were three animals associated with the high dose water treatment.CONCLUSIONS/SIGNIFICANCE:Overall, our results indicate that virus-laden water could provide a natural exposure route of AIV for raccoons and possibly other mammals associated with aquatic environments. However, this association appears to be related to AIV concentration in the water, which would constitute an infective dose. In addition, strong evidence of infection was only detected in three of four animals exposed to a high dose (e.g., 10(5.0) EID50/mL) of AIV in water. As such, water-borne transmission to raccoons may require repeated exposures to water with high concentrations of virus.

Highly pathogenic avian influenza virus A/H5N1 has been reported in 11 African countries. Migratory waterbirds have the potential of introducing A/H5N1 into east Africa through the Rift Valley of Kenya. We present the results of a wild bird surveillance system for A/H5N1 and other avian influenza viruses based on avian fecal sampling in Kenya. We collected 2630 fecal samples in 2008. Viral RNA was extracted from pools of 3-5 fecal samples and analyzed for presence of avian influenza virus RNA by real-time RT-PCR. Twelve (2.3%) of the 516 sample pools were positive for avian influenza virus RNA, 2 of which were subtyped as H4N6 viruses. This is the first report of avian influenza virus in wild birds in Kenya. This study demonstrates the success of this approach in detecting avian influenza virus in wild birds and represents an efficient surveillance system for avian influenza virus in regions with limited resources.

In aquatic bird populations, the ability of avian influenza (AI) viruses to remain infectious in water for extended periods provides a mechanism that allows viral transmission to occur long after shedding birds have left the area. However, this also exposes other aquatic organisms, including freshwater invertebrates, to AI viruses. Previous researchers found that AI viral RNA can be sequestered in snail tissues. Using an experimental approach, we determined whether freshwater snails (Physa acuta and Physa gyrina) can infect waterfowl with AI viruses by serving as a means of transmission between infected and naïve waterfowl via ingestion. In our first experiment, we exposed 20 Physa spp. snails to an AI virus (H3N8) and inoculated embryonated specific pathogen-free (SPF) chicken eggs with the homogenized snail tissues. Sequestered AI viruses remain infectious in snail tissues; 10% of the exposed snail tissues infected SPF eggs. In a second experiment, we exposed snails to water contaminated with feces of AI virus-inoculated Mallards (Anas platyrhynchos) to evaluate whether ingestion of exposed freshwater snails was an alternate route of AI virus transmission to waterfowl. None of the immunologically naïve Mallards developed an infection, indicating that transmission via ingestion likely did not occur. Our results suggest that this particular trophic interaction may not play an important role in the transmission of AI viruses in aquatic habitats.

Low-pathogenicity avian influenza virus (LPAIV) can lead to epizootics that cause economic losses in poultry or the emergence of human-infectious strains. LPAIVs experience a complex immunity landscape as they are endemic in numerous host species, and many antigenically distinct strains co-circulate. Prevention and control of emergence of detrimental strains requires an understanding of infection/transmission characteristics of the various subtypes in different hosts, including interactions between subtypes. In order to develop analytical frameworks for examining control efficacy, quantification of heterosubtypic immunity interactions is fundamental. However, these data are scarce, especially for wild avian subtypes in natural hosts. Consequently, in this study, three host species (mallards, quail and pheasants) were infected with two LPAIV subtypes isolated from wild birds: H3N8 and H4N6. The recovered hosts were also reinfected with the alternate subtype to measure the effects of heterosubtypic immunity. Oropharyngeal and cloacal swabs were collected and viral RNA load was quantified by real-time RT-PCR. For secondary infections in recovered hosts, peak viral load was up to four orders of magnitude lower and shedding length was up to 4 days shorter. However, both the magnitude and presence of heterosubtypic immunity varied across specific host species/subtype combinations. Using a mathematical model of virus replication, the variation in virus replication dynamics due to host individuals was quantified. It was found that accounting for individual heterogeneity is important for drawing accurate conclusions about treatment effects. These results are relevant for developing epidemiological models to inform control practices and for analysing virus replication data.

BackgroundAvian influenza viruses are known to productively infect a number of mammal species, several of which are commonly found on or near poultry and gamebird farms. While control of rodent species is often used to limit avian influenza virus transmission within and among outbreak sites, few studies have investigated the potential role of these species in outbreak dynamics.Methodology/principal findingsWe trapped and sampled synanthropic mammals on a gamebird farm in Idaho, USA that had recently experienced a low pathogenic avian influenza outbreak. Six of six house mice (Mus musculus) caught on the outbreak farm were presumptively positive for antibodies to type A influenza. Consequently, we experimentally infected groups of naïve wild-caught house mice with five different low pathogenic avian influenza viruses that included three viruses derived from wild birds and two viruses derived from chickens. Virus replication was efficient in house mice inoculated with viruses derived from wild birds and more moderate for chicken-derived viruses. Mean titers (EID(50) equivalents/mL) across all lung samples from seven days of sampling (three mice/day) ranged from 10(3.89) (H3N6) to 10(5.06) (H4N6) for the wild bird viruses and 10(2.08) (H6N2) to 10(2.85) (H4N8) for the chicken-derived viruses. Interestingly, multiple regression models indicated differential replication between sexes, with significantly (p<0.05) higher concentrations of avian influenza RNA found in females compared with males.Conclusions/significanceAvian influenza viruses replicated efficiently in wild-caught house mice without adaptation, indicating mice may be a risk pathway for movement of avian influenza viruses on poultry and gamebird farms. Differential virus replication between males and females warrants further investigation to determine the generality of this result in avian influenza disease dynamics.

Veterinary RecordVolume 166, Issue 1 p. 22-23 Short Communication Antibodies to influenza and West Nile viruses in horses in Mexico M. A. Loroño-Pino PhD, M. A. Loroño-Pino PhD Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorJ. A. Farfan-Ale PhD, MD, J. A. Farfan-Ale PhD, MD Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorJ. E. Garcia-Rejon PhD, J. E. Garcia-Rejon PhD Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorM. Lin PhD, M. Lin PhD Department of Veterinary Microbiology and Preventive Medicine, College of Veterinary Medicine, Iowa State University, Ames, IA, 50011 USASearch for more papers by this authorE. Rosado-Paredes BSc, E. Rosado-Paredes BSc Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorF. I. Puerto MSc, F. I. Puerto MSc Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorA. Bates DVM, A. Bates DVM Centro Médico Veterinario del Oriente, Tizimin, Yucatán, CP, 97702 MexicoSearch for more papers by this authorJ. J. Root PhD, J. J. Root PhD United States Department of Agriculture, Wildlife Services, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorA. B. Franklin PhD, A. B. Franklin PhD United States Department of Agriculture, Wildlife Services, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorH. J. Sullivan MSc, H. J. Sullivan MSc United States Department of Agriculture, Wildlife Services, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorB. J. Blitvich PhD, Corresponding Author B. J. Blitvich PhD [email protected] Department of Veterinary Microbiology and Preventive Medicine, College of Veterinary Medicine, Iowa State University, Ames, IA, 50011 USACorrespondence to Dr Blitvich, e-mail: [email protected]Search for more papers by this author M. A. Loroño-Pino PhD, M. A. Loroño-Pino PhD Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorJ. A. Farfan-Ale PhD, MD, J. A. Farfan-Ale PhD, MD Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorJ. E. Garcia-Rejon PhD, J. E. Garcia-Rejon PhD Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorM. Lin PhD, M. Lin PhD Department of Veterinary Microbiology and Preventive Medicine, College of Veterinary Medicine, Iowa State University, Ames, IA, 50011 USASearch for more papers by this authorE. Rosado-Paredes BSc, E. Rosado-Paredes BSc Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorF. I. Puerto MSc, F. I. Puerto MSc Laboratorio de Arbovirologia, Centro de Investigaciones Regionales ‘Dr Hideyo Noguchi’, Universidad Autónoma de Yucatán, Mérida, Yucatán, CP, 97000 MexicoSearch for more papers by this authorA. Bates DVM, A. Bates DVM Centro Médico Veterinario del Oriente, Tizimin, Yucatán, CP, 97702 MexicoSearch for more papers by this authorJ. J. Root PhD, J. J. Root PhD United States Department of Agriculture, Wildlife Services, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorA. B. Franklin PhD, A. B. Franklin PhD United States Department of Agriculture, Wildlife Services, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorH. J. Sullivan MSc, H. J. Sullivan MSc United States Department of Agriculture, Wildlife Services, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorB. J. Blitvich PhD, Corresponding Author B. J. Blitvich PhD [email protected] Department of Veterinary Microbiology and Preventive Medicine, College of Veterinary Medicine, Iowa State University, Ames, IA, 50011 USACorrespondence to Dr Blitvich, e-mail: [email protected]Search for more papers by this author First published: 02 January 2010 https://doi.org/10.1136/vr.b5586Citations: 4 Provenance: not commissioned; externally peer reviewed Read the full textAboutPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinkedInRedditWechat No abstract is available for this article.Citing Literature Volume166, Issue1January 2010Pages 22-23 RelatedInformation

An investigation was performed to describe the responses of naturally acquired antibodies to influenza A virus in raccoons (Procyon lotor) over time. Seven wild raccoons, some of which had been exposed to multiple subtypes of influenza A virus, were held in captivity for 279 days, and serum samples were collected on 10 occasions during this interval. Serum samples from 9 of 10 bleeding occasions were tested using an epitope-blocking enzyme-linked immunosorbent assay for the presence of antibodies to influenza A virus. Although titer declines were noted in most animals over time, all animals maintained detectable antibodies for the duration of the study. These data indicate that naturally acquired antibodies to influenza A virus can remain detectable in raccoons for many months, with the actual duration presumably being much longer because all animals had been exposed to influenza A virus before this study commenced. This information is important to surveillance programs because the duration of naturally acquired antibodies to influenza A virus in wildlife populations is largely unknown.

BackgroundWild mallards (Anas platyrhychos) are considered one of the primary reservoir species for avian influenza viruses (AIV). Because AIV circulating in wild birds pose an indirect threat to agriculture and human health, understanding the ecology of AIV and developing risk assessments and surveillance systems for prevention of disease is critical.Methodology/principal findingsIn this study, mallards were experimentally infected with an H4N6 subtype of AIV by oral inoculation or contact with an H4N6 contaminated water source. Cloacal swabs, oropharyngeal swabs, fecal samples, and water samples were collected daily and tested by real-time RT-PCR (RRT-PCR) for estimation of viral shedding. Fecal samples had significantly higher virus concentrations than oropharyngeal or cloacal swabs and 6 month old ducks shed significantly more viral RNA than 3 month old ducks regardless of sample type. Use of a water source contaminated by AIV infected mallards, was sufficient to transmit virus to naïve mallards, which shed AIV at higher or similar levels as orally-inoculated ducks.ConclusionsBodies of water could serve as a transmission pathway for AIV in waterfowl. For AIV surveillance purposes, water samples and fecal samples appear to be excellent alternatives or additions to cloacal and oropharyngeal swabbing. Furthermore, duck age (even within hatch-year birds) may be important when interpreting viral shedding results from experimental infections or surveillance. Differential shedding among hatch-year mallards could affect prevalence estimates, modeling of AIV spread, and subsequent risk assessments.

Veterinary RecordVolume 166, Issue 18 p. 565-567 Short Communication Seroprevalence of equine influenza virus in northeast and southern Mexico B. J. Blitvich PhD, Corresponding Author B. J. Blitvich PhD blitvich@iastate.edu Department of Veterinary Microbiology and Preventive Medicine, College of Veterinary Medicine, Iowa State University, Ames, IA, 50011 USAE-mail for correspondence: blitvich@iastate.eduSearch for more papers by this authorL. A. Ibarra-Juarez MSc, L. A. Ibarra-Juarez MSc Laboratorio de Entomologia Medica, Facultad de Ciencias Biologicas, Universidad Autonoma de Nuevo Leon, San Nicolas de los Garza, Nuevo Leon, 66450 MexicoSearch for more papers by this authorA. J. Cortes-Guzman MSc, A. J. Cortes-Guzman MSc Departamento de Control de Vectores, Secretaria de Salubridad y Asistencia, Chilpancingo, Guerrero, 39000 MexicoSearch for more papers by this authorJ. J. Root PhD, J. J. Root PhD Wildlife Services, United States Department of Agriculture, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorA. B. Franklin PhD, A. B. Franklin PhD Wildlife Services, United States Department of Agriculture, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorH. J. Sullivan MSc, H. J. Sullivan MSc Wildlife Services, United States Department of Agriculture, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorI. Fernandez-Salas PhD, I. Fernandez-Salas PhD Laboratorio de Entomologia Medica, Facultad de Ciencias Biologicas, Universidad Autonoma de Nuevo Leon, San Nicolas de los Garza, Nuevo Leon, 66450 MexicoSearch for more papers by this author B. J. Blitvich PhD, Corresponding Author B. J. Blitvich PhD blitvich@iastate.edu Department of Veterinary Microbiology and Preventive Medicine, College of Veterinary Medicine, Iowa State University, Ames, IA, 50011 USAE-mail for correspondence: blitvich@iastate.eduSearch for more papers by this authorL. A. Ibarra-Juarez MSc, L. A. Ibarra-Juarez MSc Laboratorio de Entomologia Medica, Facultad de Ciencias Biologicas, Universidad Autonoma de Nuevo Leon, San Nicolas de los Garza, Nuevo Leon, 66450 MexicoSearch for more papers by this authorA. J. Cortes-Guzman MSc, A. J. Cortes-Guzman MSc Departamento de Control de Vectores, Secretaria de Salubridad y Asistencia, Chilpancingo, Guerrero, 39000 MexicoSearch for more papers by this authorJ. J. Root PhD, J. J. Root PhD Wildlife Services, United States Department of Agriculture, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorA. B. Franklin PhD, A. B. Franklin PhD Wildlife Services, United States Department of Agriculture, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorH. J. Sullivan MSc, H. J. Sullivan MSc Wildlife Services, United States Department of Agriculture, National Wildlife Research Center, Fort Collins, CO, 80521 USASearch for more papers by this authorI. Fernandez-Salas PhD, I. Fernandez-Salas PhD Laboratorio de Entomologia Medica, Facultad de Ciencias Biologicas, Universidad Autonoma de Nuevo Leon, San Nicolas de los Garza, Nuevo Leon, 66450 MexicoSearch for more papers by this author First published: 01 May 2010 https://doi.org/10.1136/vr.b4845Citations: 3 Provenance: not commissioned; externally peer reviewed Read the full textAboutPDF ToolsRequest permissionExport citationAdd to favoritesTrack citation ShareShare Give accessShare full text accessShare full-text accessPlease review our Terms and Conditions of Use and check box below to share full-text version of article.I have read and accept the Wiley Online Library Terms and Conditions of UseShareable LinkUse the link below to share a full-text version of this article with your friends and colleagues. Learn more.Copy URL Share a linkShare onFacebookTwitterLinked InRedditWechat No abstract is available for this article.Citing Literature Volume166, Issue18May 2010Pages 565-567 RelatedInformation

Cliff swallows (Petrochelidon pyrrhonota) were inoculated with differing doses of West Nile virus (WNV) to evaluate their potential role as reservoir hosts in nature. Swallows often nest in large colonies in habitats and months associated with high mosquito abundance and early WNV transmission in North America. Additionally, cliff swallow diet consists of insects, including mosquitoes, leading to an additional potential route of WNV infection. The average peak viremia titer among infected cliff swallows was 106.3, plaque-forming units (PFU)/mL serum and the reservoir competence index was 0.34. There was no correlation between dose and probability of becoming infected or viremia peak and duration. Oral shedding was detected from 2 to 14 days post-inoculation with an average peak titer of 10(4.4) PFU/swab. These results suggest that cliff swallows are competent reservoir hosts of WNV and therefore, they may play a role in early seasonal amplification and maintenance of WNV.