Non-human Laboratory Diagnosis

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Laboratory Testing of Vectors

Identification and Pooling

Mosquitoes should be identified to species or lowest taxonomic unit. Specimens are placed into pools of 50 specimens or less based on species, sex, location, trap-type, and date of collection. Larger pool sizes can be used in some assays with loss of sensitivity (Sutherland and Nasci 2007). If resources are limited, testing of mosquitoes for surveillance purposes can be limited to the primary vector species.

Homogenizing and Centrifugation

After adding the appropriate media, mosquito pools can be macerated or ground by a variety of techniques including mortar and pestle, vortexing sealed tubes containing one or more copper clad BBs, or by use of tissue homogenizing apparatus that are commercially available (Savage et al. 2007). After grinding, samples are centrifuged, and an aliquot is removed for testing. Because mosquito pools may contain arboviruses and other pathogenic viruses, which may be aerosolized during processing, laboratory staff should take appropriate safety precautions including use of a Class II Type A biological safety cabinet and wearing appropriate personal protective equipment (PPE) and adhering to biosafety practices.

Virus Detection

Virus isolation in Vero cell culture remains the standard for confirmation of positive pools (Beaty et al. 1989, Savage et al. 1999, Lanciotti et al. 2000). Virus isolation provides the benefit of detecting other viruses that may be contained in the mosquitoes, a feature that is lost using test procedures that target virus-specific nucleotide sequence or proteins. However, Vero cell culture is expensive and requires specialized laboratory facilities; thus, nucleic acid assays have largely replaced virus isolation as detection and confirmatory assay methods of choice. Virus isolation requires that mosquito pools be ground in a media that protects the virus from degradation such as BA-1 (Lanciotti et al. 2000), and preservation of an aliquot at -70°C to retain virus viability for future testing.

Nucleic acid detection assays are the most sensitive assays for virus detection and confirmation of virus in mosquito pools (Lanciotti et al. 2000, Nasci et al. 2002). Real-Time RT-PCR assays with different primer sets may be used for both detection and confirmation of virus in mosquito pools. Standard RT-PCR primers are also available (Kuno et al 1998). Nucleic acids may be extracted from an aliquot of the mosquito pool homogenate by hand using traditional methods or with kits, or with automated robots in high-through-put laboratories (Savage et al 2007).

Virus antigen detection assays are available in ELISA format (Tsai et al. 1987, Hunt et al. 2002) and in commercial kits that employ lateral flow wicking assays, developed specifically for testing mosquitoes (Komar et al. 2002, Panella et al. 2001, Burkhalter et al. 2006). The antigen capture ELISA of Hunt et al. 2002 and the RAMP (Rapid Analyte Measurement Platform, Response Biomedical Corp, Burnaby, British Columbia, Canada) test are approximately equal in sensitivity and detect virus in mosquito pools at concentrations as low as 103.1 PFU/ml (Burkhalter et al. 2006). The VecTest (Medical Analysis Systems, Inc., Camarillo, CA) is less sensitive and detects virus in mosquito pools at concentrations of 105.17 PFU/ml. The VecTest (evaluated by Burkhalter et al. 2006) is no longer available but is similar to a lateral flow wicking assay marketed as VecTOR Test (VecTOR Test Systems, Inc., Thousand Oaks, CA). Although the antigen detection assays are less sensitive than nucleic acid detection assays, they have been evaluated in operational surveillance programs (Mackay et al. 2008. Lampman et al. 2006. Williges et al. 2009, Kesavaraju et al. 2012) and can provide valuable infection rate data when employed consistently in a mosquito surveillance program.


Burkhalter KL, Lindsay R, Anderson R, Dibernardo A, Fong W, Nasci RS. 2006. Evaluation of commercial assays for detecting West Nile virus antigen. J Am Mosq Control Assoc. 22:64-69.

Beaty BJ, Calisher CH, Shope RS. 1989 Arboviruses, p. 797-856. In Schmidt NJ, Emmons RW (ed.), Diagnostic procedures for viral, rickettsial and chlamydia infections. American Public Health Assoc, Washington DC.

Hunt AR, Hall RA, Kerst AJ, Nasci RS, Savage HM, Panella NA, et al. 2002. Detection of West Nile virus antigen in mosquitoes and avian tissues by a monoclonal antibody-based capture enzyme immunoassay. J Clin Microbiol. 40: 2023-30. 52

Kesavaraju B, Farajollahi A, Lampman RL, Hutchinson M, Krasavin NM, Graves SE, et al. 2012. Evaluation of a rapid analyte measurement platform for West Nile virus detection based on United States mosquito control programs. Am J Trop Med Hyg. 87(2):359-63.

Komar N, Lanciotti R, Bowen R, Langevin S, Bunning M. 2002. Detection of West Nile virus in oral and cloacal swabs collected from bird carcasses. Emerg Infect Dis. 8(7):741-2.

Kuno G, Chang G-J J, Tsuchiya KR, Karabatsos N, Cropp CB.1998. Phylogeny of the genus Flavivirus. J Virol. 72:73-83.

Lampman RL, Krasavin NM, Szyska M, Novak RJ. 2006. A comparison of two West Nile virus detection assays (TaqMan reverse transcriptase polymerase chain reaction and VecTest antigen assay) during three consecutive outbreaks in northern Illinois. J Am Mosq Control Assoc. 22(1):76-86.

Lanciotti RS, Kerst AJ, Nasci RS, Godsey MS, Mitchell CJ, Savage HM, et al. 2000. 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. 38:4066-71.

Mackay AJ, Roy A, Yates MM, Foil LD. 2008. West Nile virus detection in mosquitoes in East Baton Rouge Parish, Louisiana, from November 2002 to October 2004. J Am Mosq Control Assoc. 24(1):28-35.

Nasci RS, Gottfried KL, Burkhalter KL, Kulasekera VL, Lambert AJ, Lanciotti RS, Hunt AR, Ryan JR. 2002. Comparison of vero cell plaque assay, TaqMan reverse transcriptase polymerase chain reaction RNA assay, and VecTest antigen assay for detection of West Nile virus in field-collected mosquitoes. J Am Mosq Control Assoc.18:294-300.

Panella NA, Kerst AJ, Lanciotti RS, Bryant P, Wolf B, Komar N. 2001. Comparative West Nile virus detection in organs of naturally infected American crows (Corvus brachyrhynchos). Emerg Infect Dis. 7(4):754-5.

Savage HM, Ceianu C, Nicolescu G, Karabatsos N, Lanciotti R, Vladimirescu A, et al. 1999. Entomologic and avian investigations of an epidemic of West Nile fever in Romania in 1996, with serologic and molecular characterization of a virus isolate from mosquitoes. Am J Trop Med Hyg. 61:600-11.

Savage HM, Aggarwal D, Apperson CS, Katholi CR, Gordon E, Hassan HK, Anderson M, Charnetzky D, McMillen L, Unnasch EA, Unnasch TR. 2007. Host choice and West Nile virus infection rates in blood-fed mosquitoes, including members of the Culex pipiens complex, from Memphis and Shelby County, Tennessee, 2002-2003. Vector Borne Zoonotic Dis. 7:365-86.

Sutherland GL, Nasci RS. 2007. Detection of West Nile virus in large pools of mosquitoes. J Am Mosq Control Assoc. 23:389-95.

Tsai TF, Bolin RA, Montoya M, Bailey RE, Francy DB, Jozan M, et al. 1987. Detection of St. Louis encephalitis virus antigen in mosquitoes by capture enzyme immunoassay. J Clin Microbiol. 25:370-6.

Williges E, Farajollahi A, Nelder MP, Gaugler R. 2009. Comparative field analyses of rapid analyte measurement platform and reverse transcriptase polymerase chain reaction assays for West Nile virus surveillance. J Vector Ecol. 34(2):324-8.

Laboratory Testing of Non-human Vertebrates


Diagnostic kits for serologic diagnosis of WNV infection in clinically ill domestic animals are not commercially available. IgM-capture ELISA has been developed for use in horses and can be readily adapted to other animal species where anti-IgM antibody reagents are commercially available. Alternatively, seroconversion for IgG, neutralizing antibodies, and haemagglutinin inhibiting (HAI) assays in acute and convalescent serum samples collected 2-3 weeks apart can be used as screening assays. The latter two approaches do not require species-specific reagents and thus have broad applicability. The ELISA format may be used when employed as inhibition or competition ELISAs, which avoids the use of species-specific reagents. A popular blocking ELISA has been applied to a variety of vertebrate species with very high specificity and sensitivity, reducing the necessity of a second confirmatory test (Blitvich et al 2003a, 2003b). Similarly, the microsphere immunoassay, when used comparatively with WNV antigen-coated beads and St. Louis encephalitis virus (SLEV) antigen-coated beads, performs with high specificity and sensitivity (Johnson et al. 2005). Typically, a confirmatory 90% plaque-reduction neutralization test (PRNT90) with end-point titration is used to confirm serology in non-human vertebrates. Plaque-reduction thresholds below 80% are not recommended. Because of the cross-reactive potential of anti-flavivirus antibodies, the PRNT must be comparative, performed simultaneously with SLEV.

PRNTs require the use of a biosafety cabinet within a containment laboratory utilizing Vero cell culture. As of 2020, WNV was recommended in the Biomedical and Microbiological Laboratory guide (BMBL; v6 Biosafety in Microbiological and Biomedical Laboratories (BMBL) 6th Edition | CDC Laboratory Portal | CDC) to be handled under BSL-2 standards. Modification to the standard PRNT using a recombinant chimeric virus featuring the WNV envelope glycoprotein gene in a yellow fever virus backbone (Chimeravax®, originally developed as a live-attenuated vaccine candidate) can be used for an increased safety profile for lab staff. For PRNTs, the Chimeravax provided equivalent results for bird sera, and 10-100 fold lower titers for equine sera (Komar et al. 2009).

The same serologic techniques applied to clinically ill animals may also be used for healthy subjects for vertebrate serosurveys or for healthy sentinel animals serially sampled as sentinels. Serologic techniques for WNV diagnosis should not be applied to carcasses, as in many cases of fatal WNV infection, the host will die before a detectable immune response develops. Furthermore, some morbid or moribund animals that have WNV antibodies due to past infection may be currently infected with a pathogen other than WNV. Fatal cases should have readily detectable WNV in their tissues.

As with human diagnostic samples, serologic results from non-human vertebrates must be interpreted with caution and with an understanding of the cross-reactive tendencies of WNV and other flaviviruses. For primary WNV infections, a low rate of cross-reactivity is expected (<5%) and misdiagnoses are avoided by the requirement that the reciprocal anti-WNV titer be a minimum of 4-fold greater than the corresponding anti-SLEV titer. In rare cases, a secondary flavivirus infection due to WNV in a host with a history of SLEV infection may boost the older anti-SLEV titer to greater levels than the anti-WNV titer, resulting in a misdiagnosis of SLEV infection, a phenomenon known as “original antigenic sin”. Some serum samples will have endpoint titers for WNV and SLEV that are the same or just 2-fold different. While it is possible that this serologic result is due to past infections with both of these viruses, it is impossible to rule out cross-reaction from one or the other, or even from a third indeterminate flavivirus. Such a result should be presented as “undifferentiated flavivirus infection.”

Virus Detection

Methods for WNV detection, isolation, and identification are the same as described for human and mosquito diagnostics. Specimens typically used are tissues and/or fluids from acutely ill and/or dead animals. Virus detection in apparently healthy animals is very low-yield and inefficient, and therefore not cost-effective, and should not be considered for routine surveillance programs. In bird, mammal, and reptile carcasses, tissue tropisms have varied among individuals within a species, and across species. Some animals, like humans, have few tissues with detectable virus particles or viral RNA at necropsy, such as horses. Others, such as certain bird species, may have fulminant infections with high viral loads in almost every tissue.


Blitvich BJ, Marlenee NL, Hall RA, Calisher CH, Bowen RA, Roehrig JT. 2003a. Epitope-blocking enzyme-linked immunosorbent assays for the detection of serum antibodies to west nile virus in multiple avian species. J Clin Microbiol. 41(3):1041-7.

Blitvich BJ, Bowen RA, Marlenee NL, Hall RA, Bunning ML, Beaty BJ. 2003b. Epitope-blocking enzyme-linked immunosorbent assays for detection of west nile virus antibodies in domestic mammals. J Clin Microbiol. 41(6):2676-9.

Johnson AJ, Noga AJ, Kosoy O, Lanciotti RS, Johnson AA, Biggerstaff BJ. 2005. Duplex microsphere-based immunoassay for detection of anti-West Nile virus and anti-St. Louis encephalitis virus immunoglobulin m antibodies. Clin Diagn Lab Immunol. 12(5):566-74.

Komar, N., Langevin, S., Monath, T. P. 2009. Use of a surrogate chimeric virus to detect West Nile virus-neutralizing antibodies in avian and equine sera. Clinical and Vaccine Immunology, 16(1), 134-135.