Volume 8, Number 7—July 2002
Dispatch
Detection of West Nile Virus in Oral and Cloacal Swabs Collected from Bird Carcasses
, Robert Lanciotti*, Richard Bowen†, Stanley Langevin*, and Michel Bunning*‡
Abstract
We evaluated if postmortem cloacal and oral swabs could replace brain tissue as a specimen for West Nile virus (WNV) detection. WNV was detected in all three specimen types from 20 dead crows and jays with an average of >105 WNV PFU in each. These findings suggest that testing cloacal or oral swabs might be a low-resource approach to detect WNV in dead birds.
Since 1999, surveillance of bird deaths has become a standard epidemiologic method for detecting the spread and continued presence of West Nile virus (formal name: West Nile virus [WNV]) transmission throughout the eastern United States (1). In 2000 alone, approximately 13,000 bird carcasses were tested for WNV (2). Substantial resources are required to accomplish the tasks associated with this novel type of arbovirus surveillance: transport of the avian carcasses to a laboratory (often distinct from the microbiology laboratory where diagnostic testing will be performed), organ removal, tissue maceration and clarification, and testing of tissue homogenates. We considered ways to simplify these tasks.
Given that birds with acute WNV infection frequently shed the virus in cloacal or oral cavities (3–6) and that we have detected very high WNV titers (e.g. 106 PFU) on cloacal and oral (nasopharyngeal) swabs of corvid1 and other passerine birds with experimentally induced, acute WNV infections (N. Komar, unpub. data), we hypothesized that cloacal swabs or oral swabs from carcasses could replace brain samples, the preferred tissues to test for WNV infection in corvid carcasses (7).
We collected postmortem specimens from 20 corvids, including 12 American Crows (Corvus brachyrhynchos), 4 Fish Crows (Corvus ossifragus), and 4 Blue Jays (Cyanocitta cristata), that had died (or in one case had been euthanized after being moribund for 24 hours) after experimental infection with the New York 1999 strain of WNV. (The modes of infection1, sampling protocol, and resulting pathogenesis will be described separately.) Brain and other organs were harvested, and postmortem cloacal and oral swabs were collected (using standard cotton- or Dacron-tipped applicators) in 0.5-mL physiological buffer containing antibiotics, within 24 hours of death. All specimens were frozen at -70°C until assayed for virus content by Vero plaque assay and for WNV-specific RNA by TaqMan reverse transcriptase–polymerase chain reaction (RT-PCR), as previously described (8).
We detected WNV RNA in all postmortem brain tissue samples as well as cloacal and oral swabs. Infectious WNV particles were detected in all but one specimen, a cloacal swab taken from a Fish Crow. Viral titrations and quantitative TaqMan RT-PCR indicated that the concentrations of WNV averaged >105 in all three specimen types (Table).
Avian mortality surveillance for WNV targets fresh carcasses (generally dead <24 h), especially corvids, for detection of infectious virus particles or RNA in brain or other viscera. We have shown that postmortem oral and cloacal swabs, in addition to brain, are effective samples to collect for WNV detection in experimentally infected corvids. A potential implication of these findings, pending field trials using corvids and other species routinely collected as part of avian mortality surveillance, is that WNV may be detected by simply collecting swabs from carcasses and forwarding the swabs (frozen) to a virology laboratory for testing. Eliminating multiple steps currently necessary for WNV testing of bird carcasses may conserve valuable public health resources and reduce the risk of exposure for laboratory personnel.
Nicholas Komar is the Vertebrate Ecologist for the Centers for Disease Control and Preventions' Arbovirus Diseases Branch, Division of Vector-Borne Infectious Diseases, in Fort Collins, Colorado. His major research interest is the role of vertebrate hosts in arbovirus transmission cycles.
Acknowledgment
We thank Bruce Cropp for assisting with laboratory testing; Carol Snarey, Robert Craven, Grant Campbell, John Roehrig, Duane Gubler, and Lyle Petersen for critically reviewing the manuscript; and the Maryland Department of Natural Resources for providing the crows used in this study.
References
- Eidson M, Komar N, Sorhage F, Nelson R, Talbot T, Mostashari F, Crow deaths as a sentinel surveillance system for West Nile virus in the northeastern United States, 1999. Emerg Infect Dis. 2001;7:615–20.PubMedGoogle Scholar
- Marfin AA, Petersen LR, Eidson M, Miller J, Hadler J, Farello C, Widespread West Nile virus activity, eastern United States, 2000. Emerg Infect Dis. 2001;7:730–5.PubMedGoogle Scholar
- Senne DA, Pedersen JC, Hutto DL, Taylor WD, Schmitt BJ, Panigrahy B. Pathogenicity of West Nile virus in chickens. Avian Dis. 2000;44:642–9. DOIPubMedGoogle Scholar
- Swayne DE, Beck JR, Zaki S. Pathogenicity of West Nile virus for turkeys. Avian Dis. 2000;44:932–7. DOIPubMedGoogle Scholar
- Langevin SA, Bunning M, Davis B, Komar N. Experimental infection of chickens as candidate sentinels for West Nile virus. Emerg Infect Dis. 2001;7:726–9.PubMedGoogle Scholar
- Swayne DE, Beck JR, Smith CS, Shieh WJ, Zaki SR. Fatal encephalitis and myocarditis in young domestic geese (Anser anser domesticus) caused by West Nile virus. Emerg Infect Dis. 2001;7:751–3.PubMedGoogle Scholar
- Panella NA, Kerst AJ, Lanciotti RS, Bryant P, Wolf B, Komar N. Comparative West Nile virus detection in organs of naturally infected American Crows (Corvus brachyrhynchos). Emerg Infect Dis. 2001;7:754–5.PubMedGoogle Scholar
- Lanciotti RS, Kerst AJ, Nasci RS, Godsey MS, Mitchell CJ, Savage HM, Rapid detection of West Nile virus from human clinical specimens, field-collected mosquitoes, and avian samples by a TaqMan reverse transcriptase-PCR assay. J Clin Microbiol. 2000;38:4066–71.PubMedGoogle Scholar
Table
Cite This Article1 Pertaining to the family Corvidae, including crows, jays and magpies.
2 These birds were infected either by mosquito bite or by direct contact with infected cagemates, both of which are potentially natural modes of infection.
Table of Contents – Volume 8, Number 7—July 2002
| EID Search Options |
|---|
Advanced Article Search – Search articles by author and/or keyword.
|
Articles by Country Search – Search articles by the topic country.
|
Article Type Search – Search articles by article type and issue.
|
Please use the form below to submit correspondence to the authors or contact them at the following address:
Nicholas Komar, Centers for Disease Control and Prevention, P.O. Box 2087, Fort Collins, CO 80522, USA; fax: 970-221-6476
Top