Survey for West Nile Virus Infection in Free-ranging American Alligators in Louisiana
Rachel M. McNew, Ruth M. Elsey, Thomas R. Rainwater, Eric J. Marsland, and Steven M. Presley
Southeastern Naturalist, Volume 6, Number 4 (2007): 737–742
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2007 SOUTHEASTERN NATURALIST 6(4):737–742
Survey for West Nile Virus Infection in Free-ranging
American Alligators in Louisiana
Rachel M. McNew1, Ruth M. Elsey2, Thomas R. Rainwater3,
Eric J. Marsland1, and Steven M. Presley1,*
Abstract - West Nile virus (WNV) is an endemic arboviral pathogen that occurs
throughout most of the United States and is typically maintained through a birdmosquito-
bird transmission cycle. The ecological significance of the virus is high due
to its ability to infect and cause disease in humans, livestock, and wildlife. West Nile
virus infection of many vertebrate species causes signs of viral illness, including encephalitis
that may result in mortality. Infection by WNV has recently been detected
in captive Alligator mississippiensis (American Alligators) in Georgia and Louisiana,
and in both captive and free-ranging alligators in Florida. However, additional surveys
for WNV in populations of free-ranging alligators within the southeastern USA have
not been conducted. The purpose of this study was to survey free-ranging alligators in
south Louisiana for active WNV infection. Blood samples were collected from 93 alligators
captured at Rockefeller Wildlife Refuge in Cameron Parish, LA, during May
2006 and were screened for WNV using reverse transcriptase polymerase chain reaction
(RT-PCR). All samples (100%) tested negative for WNV, indicating a lack of detectable
active infection in these animals. Additional surveys of the occurrence of WNV in
alligators throughout the southeastern USA are needed to determine the susceptibility of
these reptiles to the virus, effects on the health of infected populations, and the potential
role of alligators in the maintenance and transmission of the virus.
West Nile virus (WNV) is an arboviral (arthropod-borne viral) pathogen
(flavivirus) that is primarily maintained in a natural transmission cycle involving
ornithophillic mosquito species and birds, but other vertebrate hosts,
including humans and mammals, may be incidentally infected by mosquitoes
(Huchzermeyer 2002, Komar 2003); thus, this zoonotic disease poses
a threat to both wildlife and human health (Goddard 2000). The severity
of the disease associated with WNV infection ranges from mild fever and
malaise to life-threatening illness. Encephalitis and meningitis may occur in
humans and other animals, particularly mammals, as a result of WNV infection
(Marra et al. 2004). Other signs associated with the virus include fever,
internal hemorrhaging, and tissue necrosis, which may lead to eventual brain
damage (Marra et al. 2004).
Reptiles may play a significant role in arboviral disease transmission
cycles (Bowen 1977, Nevarez et al. 2005). Japanese encephalitis virus has
1The Institute of Environmental and Human Health, Box 41163, Lubbock, TX 79409.
2Louisiana Department of Wildlife and Fisheries, Rockefeller Wildlife Refuge, 5476
Grand Chenier Highway, Grand Chenier, LA 70643. 3The Institute of Environmental
and Human Health Field Station, Jeffersonian Institute, PO Box 764, Jefferson, TX
75657. *Corresponding author - email@example.com.
738 Southeastern Naturalist Vol. 6, No. 4
been reported in several reptilian species (Doi et al.1983, Mifune et al.1969,
Shortridge et al. 1977). Recently, WNV has been recovered from the blood
of multiple crocodilian species including Crocodylus niloticus Laurenti
(Nile Crocodile) in Africa and Crocodylus moreletii Dumeril and Bibron
(Morelet’s Crocodile) in Mexico (Steinman et al. 2003). In addition, WNV
has also been detected in both captive and free-ranging Alligator mississippiensis
(Daudin) (American Alligators) in the USA (Jacobson et al. 2005a,b,
Miller et al. 2003, Nevarez et al. 2005). Alligators are typically affected by
WNV in ways that are visually discernable, with the most common signs
being neurological changes such as tremors, tilting of the head, and swimming
in circles (Jacobson et al. 2005a, Nevarez et al. 2005). Human cases of
WNV infection have been documented in Louisiana since 2002 (Zohrabian
et al. 2004), and the virus has been detected in captive alligators at several
ranches in the state. However, free-ranging alligators in Louisiana have not
yet been examined for WNV infection. Thus, the objective of this study was
to conduct a preliminary survey of the incidence of active WNV infection in
an alligator population in southwestern Louisiana.
During May 2006, 93 free-ranging American Alligators were captured at
night from impoundments on the Louisiana Department of Wildlife and Fisheries’
Rockefeller Wildlife Refuge in Cameron Parish, LA. Alligators were
located with high-intensity spotlights, approached by airboat, and captured
with a self-locking cable snare (Decker Manufacturing, Keokuk, IA). The alligators
ranged in size from 49.5 cm to 208.3 cm total length. A 10-mL blood
sample was immediately collected from the spinal vein of the larger alligators
into a heparin-coated syringe and placed on ice for transport. After blood was
collected, measurements and sex were recorded, and then the larger alligators
were released at the capture site. The smallest alligators were retained in
holding boxes and later bled with smaller bore needles not available during the
field capture. Of the 10-mL blood sample taken from each alligator, 2–3 mL of
whole blood was placed into a vial and frozen for analyses.
Reverse-transcriptase polymerase chain reaction (RT-PCR) was used to
detect viral infection in the blood at the time of capture. RT-PCR is an assay
that has been documented to provide accurate and timely results in cell
cultures and infected mosquito pools (Hadfield 2001). Other studies have
utilized RT-PCR for blood meal analyses in mosquitoes that fed on reptilian
blood as a means of detecting other flavivirus species (Cupp et al. 2004).
RNA extraction and assay
Total RNA was extracted from 0.2 mL of whole blood using TriReagent®
BD (Sigma-Aldrich), per manufacturer’s directions. Approximate RNA yield
was determined to be 3–4 μg for the majority of samples, and the concentrations
were adjusted to 0.2 ng/μL for RT-PCR screening. RT-PCR was performed on
2007 R.M. McNew, R.M. Elsey, T.R. Rainwater, E.J. Marsland, and S.M. Presley 739
the total RNA extracted from the alligator blood to determine presence of WNV.
Specimens were assayed using the primer set FLV1/FLV2, developed by Platonov
et al. (2001), which targets the West Nile/Kunjin virus’ conserved region
of the envelope (E) gene; FLV1 consists of 5’- GGI AGC AGI GCC ATI TGG
T(A/T)C ATG TGG - 3’ and FLV2 consists of 5’- C(G/T)I GTG TCC CAI CCI
GCI GTG TCA TC - 3’. Forty-five μL of a prepared RT-PCR mixture was combined
with 5 μL (1 ng) of the blood sample and dispensed into a 0.2-mL PCR
tube. Using a PTC-100 thermal cycler (MJ Research Waltham, MA), cycling
procedures for the RT-PCR reactions included a cycle at 49 °C for 45 minutes to
synthesize the first-strand cDNA; a second cycle at 95 °C for 3 minutes for initial
denaturation; 40 cycles at 94 °C for 45 seconds, 56 °C for 45 seconds, and 72
°C for 1 minute for PCR amplification; and then a final elongation step at 72 °C
for 10 minutes. Twenty-five μL of the RT-PCR product from each sample was
then loaded along with bromophenol blue onto a 2% agarose gel in Tris-borate-
EDTA (TBE) buffer and processed for approximately 1.5 hr at 50 volts.
All of the 93 alligator blood samples assayed (one per animal) proved
negative for the presence of active WNV infection by RT-PCR (Fig. 1).
Results from this study suggest that alligators tested for WNV did not
have a detectable viremia at the time of sampling, and that active infection
by WNV was not occurring in free-ranging alligators in the Rockefeller
Figure 1. Gel electrophoresis of an RT-PCR assay for WNV from whole blood samples
of alligators. Shown are the results of 12 samples. Lane 1 is the molecular ladder.
Lanes 2–13 are negative alligator samples. Lane 14 is the WNV positive control at
220 bp, and lane 15 is the negative control.
740 Southeastern Naturalist Vol. 6, No. 4
Wildlife Refuge. This may be due to a lack of WNV presence in mosquito
species in the specific area sampled during this study. These results support
the low prevalence of WNV-infected mosquitoes in early summer months, as
peak viral activity has been reported in the months of July through October
in the United States (Hayes et al. 2005). Alligators are known to be potential
WNV amplifiers, and one source of viral infection may be the consumption
of infected prey species (Klenk et al. 2004).
Although birds are most often used as indicator species for WNV
(Guptill et al. 2003), they may not be likely the most appropriate choice
for Louisiana (LA Department of Health and Hospitals 2006). Several
species of mosquitoes occurring in the New Orleans area prefer ectothermic
blood meals, including Uranotaenia sapphirina (Osten Sacken) and
Culex erraticus (Dyar and Knab) (Darsie and Ward 2005). Both of these
species have tested positive for WNV infection, and Culex spp. mosquitoes
have been shown to be the principal vectors of the virus (CDC 2005,
Hayes et al. 2005).
Various other arboviral infections have been reported in other reptiles
(Bowen 1977, Doi et al. 1983). The ability for a pathogen to overwinter in
an area is essential to the persistence of a disease in temperate environments,
and is a critical factor in the study and understanding of endemic diseases
(Kitron et al. 1998). The potential for reptiles to serve as overwintering
reservoirs for specific arboviruses has been previously proposed. Bowen
(1977) reported Gopherus berlandieri (Agassiz) (Texas Tortoise) served
as an overwintering reservoir for western equine encephalitis (WEE), and
Doi et al. (1982) reported Takydromus sp. (lacertid lizards) and Eumeces sp.
(skinks) as overwintering reservoirs for Japanese encephalitis virus (JEV).
However, the results of the present study do not support such a relationship
between WNV and the alligator population.
A recent report (Jacobson et al. 2005b) of WNV antibodies detected
in blood of free-ranging alligators in Florida (a low seroprevalence of
1.5%) suggests the potential for exposure to and infection of alligators
with WNV in other areas within the species’ range. In the present study,
antibodies were not examined, as we were focused on active WNV infection
at the time of sampling. Recent studies have reported that the serum
of American Alligators exhibit antibacterial, amoebacidal, and antiviral
properties (Merchant et al. 2003, 2004, 2005). The alligators sampled in
this study may have never been infected with WNV, or they may have
been exposed to the virus, but both innate and acquired immunity may
have prevented progression of infection.
It is clear that reptiles are susceptible to infection by WNV, and they
may also be involved in the maintenance and transmission dynamics of the
virus (Bowen 1977, Nevarez et al. 2005). The lack of WNV detection in alligators
tested in this study suggests that active infection among alligators
in this geographic area was not occurring at the time of sampling. However,
given the high density of alligator populations in many areas of the southeastern
US and the documented occurrence of WNV in free-ranging and
2007 R.M. McNew, R.M. Elsey, T.R. Rainwater, E.J. Marsland, and S.M. Presley 741
captive alligators from numerous geographic locations, additional research
is needed to more closely examine the epizoological role of these reptiles
in WNV transmission dynamics. Future studies should include IgG, IgM,
and protein electrophoresis so that the history of infection in sampled animals
can be determined.
We thank Phillip “Scooter” Trosclair, Dwayne LeJeune, Jeb Linscombe, George
Melancon, and Vida Landry of the Louisiana Department of Wildlife and Fisheries
for assistance with capture and sampling of alligators. Justin Pinkston, Undergraduate
Scholar with the Howard Hughes Medical Institute’s Undergraduate Science
Education Program, is thanked for assistance with laboratory assays. We also thank
Mr. W. Parke Moore III for administrative assistance during this study.
Bowen, S.G. 1977. Prolonged western equine encephalitis viremia in the Texas
Tortoise (Gopherus berlandieri). American Journal of Tropical Medical Hygiene
Center for Disease Control (CDC). 2005. Infectious disease information. Available
online at http://www.cdc.gov/ncidod/diseases/list_mosquitoborne.htm. Accessed
October 25, 2006.
Cupp, E.W., D. Zhang, X. Yue, M.S. Cupp, C. Guyer, T.R. Sprenger, and T.R. Unnasch.
2004. Identification of reptilian and amphibian blood meals from mosquitoes
in an eastern equine encephalomyelitis virus focus in central Alabama.
American Journal of Tropical Medical Hygiene 71:272–276.
Darsie, R.F., and R.A. Ward. 2005. Identification and Geographical Distribution of
the Mosquitoes of North America, North of Mexico. University Press of Florida,
Doi, R., A. Oya, A. Shirasaka, S. Yabe, and M. Sasa. 1983. Studies on Japanese encephalitis
virus infection of reptiles II. Role of lizards on hibernation of Japanese
encephalitis virus. Japanese Journal of Experimental Medicine. 53:125–134.
Goddard, J. 2000. Infectious Diseases and Arthropods. Humana Press, New York, NY.
Guptill, S.C., K.G. Julian, G.L. Campbell, S.D. Price, and A.A. Marfin. 2003.
Early-season avian deaths from West Nile virus as warnings of human infection.
Emerging Infectious Diseases 9:483–484.
Hadfield, T.L., M. Turell, M.P. Dempsey, J. David, and E.J. Park. 2001. Detection
of West Nile virus in mosquitoes by RT-PCR. Molecular and Cellular Probes.
Hayes, E.B., N. Komar, R.S. Nasci, S.P. Montgomery, D.R. O’Leary, and G.L.
Campbell. 2005. Epidemiology and transmission dynamics of West Nile virus
disease. Emerging Infectious Diseases 11:1167–1173.
Huchzermeyer, F.W. 2002. Diseases of farmed crocodiles and ostriches. Revue Scientifi
que et Technique. 2:265–276.
Jacobson, E.R., P.E. Ginn, J.M. Troutman, L. Farina, L. Stark, K. Klenk, K.L.
Burkhalter, and N. Komar. 2005a. West Nile virus infection in farmed American
Alligators (Alligator mississippiensis) in Florida. Journal of Wildlife Diseases
742 Southeastern Naturalist Vol. 6, No. 4
Jacobson, E.R., A.J. Johnson, J.A. Hernandez, S. Tucker, A.P. Dupuis II, R. Steven,
D. Carbonneau, and L. Stark. 2005b. Validation and use of an indirect enzymelinked
immunosorbent assay for detection of antibodies to West Nile virus in
American Alligators (Alligator mississippiensis) in Florida. Journal of Wildlife
Kitron, U., J. Swanson, M. Crandell, P.J. Sullivan, J. Anderson, R. Garro, L.D. Haramis,
and P.R. Grimstad. 1998. Introduction of Aedes albopictus into a La Crosse
virus-enzootic site in Illinois. Emerging Infectious Diseases 4:627–630.
Klenk, K., J. Snow, K. Morgan, R. Bowen, M. Stephens, F. Foster, P. Gordy, S. Beckett,
N. Komar, D. Gubler, and M. Bunning. 2004. Alligators as West Nile virus
amplifiers. Emerging Infectious Diseases 10(12): 2150–2155.
Komar, N. 2003. West Nile virus: Epidemiology and ecology in North America.
Advanced Virus Research 61:185–234.
Louisiana Department of Health and Hospitals. 2006. West Nile virus and other
arboviruses. Available online at http://www.dhh.louisiana.gov/offices/?ID=282.
Accessed October 25, 2006.
Marra, P.P., S. Griffing, C. Caffrey, A.M. Kilpatrick, R. McLean, C. Brand, E. Saito,
A.P. Dupuis, L. Kramer, and R. Novak. 2004. West Nile virus and wildlife. Bio-
Merchant, M.E., C.M. Roche, R.M. Elsey, and J. Prudhomme. 2003. Antibacterial
properties of serum from the American Alligator (Alligator mississippiensis).
Comparative Biochemistry and Physiology. 136(3):505–513.
Merchant, M.E., D. Thibodeaux, K. Loubser, and R.M. Elsey. 2004. Amoebacidal effects
of serum from the American Alligator (Alligator mississippiensis). Journal
of Parasitology 90(6):1480–1483.
Merchant, M.E., R.M. Elsey, P.L. Trosclair III, and G. Diamond. 2005. (Abstract).
Broad spectrum antimicrobial activities of leukocyte extracts from the American
Alligator. Presented at the Louisiana Chapter of the American Fisheries Society
Meeting, Baton Rouge, LA. 4 February 2005.
Mifune, K., A. Shichijo, Y. Ueda, O. Suenaga, and I. Miyagi. 1969. Low susceptibility
of common snakes in Japan to Japanese encephalitis virus. Tropical Medicine.
Miller, D.L., M.J. Mauel, C. Baldwin, G. Burtle, D. Ingram, M.E. Hines II, and S.
Frazier. 2003. West Nile virus in farmed alligators. Emerging Infectious Diseases
Nevarez, J.G., M.A. Mitchell, D.Y. Kim, R. Poston, and H.M. Lampinen. 2005. West
Nile virus in alligator, Alligator mississippiensis, ranches from Louisiana. Journal
of Herpetology Medicine and Surgery. 15:10–15.
Platonov, A.E., G.A. Shipulin, O.Y. Shipulina, E.N. Tyutyunnik, T.I. Frolochkina,
R.S. Lanciotti, S. Yazyshina, O.V. Platonova, I.L. Obukhov, A.N. Zhukov, Y.Y.
Vengerov, and V.I. Pokrovskii. 2001. Outbreak of West Nile virus infection, Volgograd
Region, Russia, 1999. Emerging Infectious Diseases 7:128–132.
Shortridge, K.F., A. Oya, M. Kobayashi, and R. Duggan. 1977. Japanese encephalitis
virus antibody in cold-blooded animals. Trans. R. Soc. Transactions of the Royal
Society of Tropical Medicine and Hygiene. 71:261–262.
Steinman, A., C. Banet-Noach, S. Tal, O. Levi, L. Simanov, S. Perk, M. Malkinson,
and N. Shpigel. 2003. West Nile virus infection in crocodiles. Emerging Infectious
Zohrabian, A., M.I. Meltzer, R. Ratard, K. Billah, N.A. Molinari, K. Roy, R.D. Scott
II, and L.R. Petersen. 2004. West Nile virus economic impact, Louisiana, 2002.
Emerging Infectious Diseases 10:1736–1744.