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West Nile virus PCR test for primates
primate assay data sheet

West Nile virus (WNV)

Test code:
S0048 - Ultrasensitive qualitative detection of West Nile virus by reverse transcription coupled real time polymerase chain reaction


West Nile virus (WNV) belongs to the genus Flavivirus of the family Flaviviridae and is an arthropod-borne virus. It possesses a single-stranded plus-sense RNA genome of approximately 11,000 nucleotides. It circulates in natural transmission cycles involving primarily Culex species mosquitoes and birds; humans and other mammals, such as primates, are thought to be incidental hosts.

Historically, WNV was found primarily in Africa, Asia, southern Europe, and Australia and was responsible for several significant epidemics, notably, in Israel (1950s), France (1962), South Africa (1974), and Romania (1996) (Hayes, 1989; Tsai et al., 1998;Savage et al., 1999). In 1999 and 2000, WNV was responsible for epidemics and epizootics in the northeastern United States, in which there were human fatalities and extensive avian mortality (Anderson et al., 1999; Lanciotta et al., 1999). On the basis of retrospective serosurveys conducted in New York City in 1999 and 2000, symptomatic illness develops in approximately 20% of persons infected with WNV and approximately 1 in 150 human infections results in meningoencephalitis, the most commonly reported form of WNV-associated illness.

In 2002, an outbreak of West Nile virus infection occurred in the state of Louisiana in which 319 human cases of WNV-associated illness were reported. Most of these cases happened in the southeastern portion of the state, including St. Tammany Parish. The Tulane National Primate Research Center (TNPRC) is located in St. Tammany Parish and houses large outdoor breeding colonies of baboons and macaques. A serological survey of primates in these breeding colonies indicated that approximately 36% of the nonhuman primates were infected with WNV during the 2002 transmission season (Ratterree et al., 2003). Implications of this study are that nonhuman primates can be as susceptible to West Nile virus infection as humans, and captive primate populations can be a potential source of viral carriers.

Surveillance for West Nile virus relies on the testing of field-collected mosquitoes and on the testing of dead birds for the presence of virus by isolation in cell culture. However, virus isolation followed by identification through immunofluorescence assays can take over a week to complete. In addition, virus isolation in cell culture from CSF or serum has generally been unsuccessful, likely due to the low level and short-lived viremia associated with infections with these viruses (Monath and Heinz, 1996; Southam and Moore, 1954).

Human WNV infections can be inferred by immunoglobulin M (IgM) capture and IgG enzyme-linked immunosorbent assays (ELISAs); however, confirmation of the type of infecting virus is possible only by detection of a fourfold or greater rise in virus-specific neutralizing antibody titers in either cerebrospinal fluid (CSF) or serum by performing the plaque reduction neutralization assay (PRNT) with several flaviviruses (Johnson et al., 2000; Martin et al., 2000). Thus serological detection of WNV infection is neither specific nor sensitive. PCR detection of West Nile virus is now considered to be a rapid, specific and sensitive detection method to identify this virus.


  • Help confirm the disease causing agent
  • Help ensure that animal colonies are free of West Nile virus
  • Early prevention of spread of the virus among a colony
  • Minimize personnel exposure to the virus
  • Safety monitoring of biological products and vaccines that derive from primates

Anderson, J. F., Andreadis, T.G., Vossbrinck, C.R., Tirrell, S.,Wakem, E.M., French, R.A., Garmendia, A.E. and Van Kruiningen, H.J. (1999) Isolation of West Nile virus from mosquitoes, crows, and a Cooper's hawk in Connecticut. Science 286:2331-2333.
Hayes, C. G. (1989). West Nile fever, p. 59-88. In T. P. Monath (ed.), The arboviruses: epidemiology and ecology, vol. V. CRC Press, Inc., Boca Raton, Fla.
Johnson, A. J., Martin, D.A., Karabatsos, N. and Roehrig, J.T.(2000) Detection of antiarboviral immunoglobulin G by using a monoclonal antibody-based capture enzyme-linked immunosorbent assay. J. Clin. Microbiol. 38:1827-1831.
Lanciotti, R. S., Roehrig, J.T., Deubel, V., Smith, J., Parker, M., Steele, K., Volpe, K.E., Crabtree, M.B., Scherret, J.H., Hall, R.A., MacKenzie, J.S., Cropp, C.B., Panigrahy, B., Ostlund, E., Schmitt, B., Malkinson, M., Banet, C., Weissman, J., Komar, N., Savage, H.M., Stone, W., McNamara, T. and Gubler, D.J.(1999) Origin of the West Nile virus responsible for an outbreak of encephalitis in the northeastern U.S. Science 286:2333-2337.
Martin, D. A., Muth, D.A., Brown, T., Johnson, A.J., Karabatsos, N. and Roehrig, J.T. (2000) Standardization of immunoglobulin M capture enzyme-linked immunosorbent assays for routine diagnosis of arboviral infections. J. Clin. Microbiol. 38:1823-1826.
Monath, T. P., and Heinz, F.X. (1996) Flaviviruses, p. 978-984. In B. N. Fields (ed.), Fields virology, 3rd ed., vol. 1. Lippincott-Raven Publishers, Philadelphia, Pa.
Ratterree, M.S., da Rosa, A.P., Bohm, R.P. Jr, Cogswell, F.B., Phillippi, K.M., Caillouet, K., Schwanberger, S., Shope, R.E. and Tesh, R.B.(2003) West Nile virus infection in nonhuman primate breeding colony, concurrent with human epidemic, southern Louisiana. Emerg Infect Dis. 9:1388-1394.
Southam, C. M., and Moore, A.E. (1954) Induced virus infections in man by the Egypt isolates of West Nile virus. Am. J. Trop. Med. Hyg. 3:19-50.
Savage, H. M., Ceianu, C., Nicolescu, G., Karabatsos, N.,Lanciotti, R., Vladimirescu, A., Laiv, L., Ungureanu, A., Romanca, C. and Tsai, T.F. (1999). Entomologic and avian investigations of an epidemic of West Nile fever in Romania, 1996, with serological and molecular characterization of a virus from mosquitoes. Am. J. Trop. Med. Hyg. 61:600-611.
Tsai, T. F., Popovici, F., Cernescu, C., Campbell, G.L. and Nedelcu, N.I. (1998) West Nile encephalitis epidemic in southeastern Romania. Lancet 352:767-771.

Specimen requirements:

Preferred specimens - 0.2 ml CSF, or 0.2 ml fresh or frozen CNS tissue.

Less preferred specimens - 0.2 ml whole blood in EDTA (purple top) tube, or 0.2 ml serum or plasma.

Contact Zoologix if advice is needed to determine an appropriate specimen type for a specific diagnostic application. For specimen types not listed here, please contact Zoologix to confirm specimen acceptability and shipping instructions.

For all specimen types, if there will be a delay in shipping, or during very warm weather, refrigerate specimens until shipped and ship with a cold pack unless more stringent shipping requirements are specified. Frozen specimens should be shipped so as to remain frozen in transit. See shipping instructions for more information.

Turnaround time: 2 business days

Methodology: Qualitative reverse transcription coupled real time PCR

Normal range: Nondetected

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