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IDENTIFICATION OF POTENTIAL MOSQUITO VECTORS OF WEST NILE VIRUS
ON A FLORIDA ALLIGATOR FARM
SANDRA C. GARRETT
A THESIS PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
MASTER OF SCIENCE
UNIVERSITY OF FLORIDA
Sandra C. Garrett
This document is dedicated to my husband, Dr. Jose Carlos V. Rodrigues, and to my
father, Dr. Alfred J. Garrett, the two scientists who inspire me the most.
I thank Dr. Alejandra Maruniak for tireless technical help, encouragement, and
ideas. I thank Aissa Doumbouya also for her help in the lab and encouragement and I
thank both her and Leslie Rios for always being ready to bounce ideas around about our
work. I thank the people at the alligator farm for allowing me to roam around with my
strange-looking equipment and for supporting my collecting efforts. I thank Dr. Darryl
Heard of the vet school for providing alligator blood. I thank Dr. Goerning, David Hoel,
and Dr. Sandra Allan for lending collecting equipment. I thank my advisors, Dr. James
Maruniak, Dr. Jerry Butler, and Dr. Elliot Jacobson, for their guidance and support
without which the project would not have been possible.
TABLE OF CONTENTS
A C K N O W L E D G M E N T S ................................................................................................. iv
LIST OF TA BLE S .............................. ......... ..................... ............. vii
LIST OF FIGURES ............................... ................ ................ viii
ABSTRACT .............. .......................................... ix
1 IN TR O D U C T IO N ............................................................. .. ......... ...... .....
W N V in Farm ed A alligators .................................................................................. 6
Mosquitoes as Vectors of WNV on Alligator Farms ................................................9
Blood Meal Identification...................... ....... .............................. 10
Screening M osquitoes for W N V .................................. ..................................... 16
2 M ETHOD S AND M ATERIALS ........................................... .......................... 20
M mosquito Collecting .................................. ... .. ........ .............. 20
B lood M eal Identification ........................................... ...........................................26
V iru s D ete ctio n ...................................................................................................... 3 1
3 R E S U L T S .............................................................................3 6
M mosquito Collecting .................................. ... .. ........ .............. 36
B lood M eal Identification .......................................... ............................................40
V iru s D ete ctio n ...................................................................................................... 4 6
4 DISCUSSION .................. .................................... ...........................48
B lood M eal Identification .......................................... ............................................4 8
V irus D election .................................................................. ............. 51
V sector Incrim nation ......... .............................................................. ..... .... ..... 54
M osquito C ontrol........... .... .............................................. ............. .. .... 57
A alternative V ertebrate Reservoirs .............................................. ............... 58
5 CONCLUSIONS AND AREAS FOR FURTHER STUDY ..............................59
A PROTOCOL FOR QIAGEN QIAQUICK SPIN KIT, PURIFICATION OF DNA
FR OM A G A R O SE GEL ................................................. ............................... 61
B ABI PRISMTM DYE TERMINATOR CYCLE SEQUENCING KIT, PROTOCOL
FOR DN A SEQUEN CIN G .............................................. .............................. 62
C PROTOCOL FOR PGEM-T VECTOR LIGATION KIT,..................................... 64
D PROTOCOL FOR QIAPREP SPIN MINIPREP KIT, EXTRACTION OF
PL A SM ID ............. ..... .. ......... .............. ............................65
E PROTOCOL FOR RNA EXTRACTION FROM MOSQUITO POOL USING
T R IZ O L L S (G IB C O ) ........................................................................ .................. 66
F SEQUENCES OF PCR PRODUCTS USED TO IDENTIFY VERTEBRATE
HOST ORIGIN OF MOSQUITO BLOOD MEALS ...........................................67
L IST O F R E FE R E N C E S ....................................................................... .... ..................72
BIO GRAPH ICAL SK ETCH .................................................. ............................... 83
LIST OF TABLES
2-1 Primers sets in PCR used to amplify DNA from different vertebrate hosts. ...........31
2-2 Primers sets used in RT-PCR to test for the presence of WNV RNA....................33
2-3 Reagent concentrations and thermocycle conditions used for PCR with
vertebrate-specific primer sets and RT-PCR with WNV-specific primer sets.........34
3-1 Mosquitoes captured from CDC light traps during Trip one at Farm A ...............39
3-2 Mosquitoes collected in resting boxes and CDC light traps during the second
collecting trip to Farm A .. ............................. .... .......................................39
3-3 Mosquitoes collected from CDC light traps and resting boxes during the third
collecting trip to Farm A ..... ........................... ........................................40
3-4 Mosquitoes captured in CDC light traps and resting boxes on the fourth
collecting trip to Farm A ..... ........................... ........................................41
3-5 Identities of vertebrate hosts as determined by sequencing the PCR product, and
information about collection date and location on farm of the mosquito sample. ...45
LIST OF FIGURES
2-1 CDC light trap set up on the western margin of the farm. ............. ..................21
2-2 A 30 cm x 30 cm x 30 cm wooden resting box with black exterior and maroon
interior was used to attract blood fed mosquitoes. ............. .................................... 23
2-3 M ap depicting layout of Farm A. ........................................ ........................ 25
2-4 The membrane feeding system was used to feed alligator blood and alligator
meat juice to Cx. quinquefasciatus and Ae. aegypti mosquitoes ...........................27
3-1 Total mosquito numbers collected over four trips to Farm A...............................37
3-2 Portions of each mosquito species captured in CDC light traps set outside of
alligator pens versus inside of pens (for collecting trips 1,2, and 4)......................38
3-3 Products from PCR amplification of mosquito samples with alligator-specific
p rim e rs ...................................... ................................... ................ 4 3
3-4 Products from PCR amplifications with a mammalian-specific primer set (lanes
2-5) and an avian-specific primer set (lanes 6-8). .................................................44
3-5 Products from RT-PCR with WNV screening primer set (lanes 2-7) and WNV
confirm ation set (lanes 8-13)......................................................... ............... 47
Abstract of Thesis Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Master of Science
IDENTIFICATION OF POTENTIAL MOSQUITO VECTORS OF WEST NILE VIRUS
ON A FLORIDA ALLIGATOR FARM
Sandra C. Garrett
Chair: James Maruniak
Major Department: Entomology and Nematology
Over the past several years, alligator farms in Florida, Georgia, and Louisiana have
experienced sudden die-offs of juvenile and hatchling alligators (Alligator
mississippiensis). These events occurred in the fall and tended to last two or three weeks.
Histologic findings, virus culture, and RT-PCR evidence all suggest that the deaths were
caused, at least in part, by infection with West Nile virus (WNV), a virus which is
vectored by mosquitoes. Blood meal identification and virus screening were done in
order to determine which mosquito species, if any, were involved in transmission of
WNV on the farm. During September and October of 2003 four trips were made to an
alligator farm in central Florida to collect mosquitoes inside and around the alligator
pens. DNA was extracted from the abdomen of blood-fed individuals to test for the
presence of alligator, avian, and mammal blood using PCR with different primer sets.
Positives were confirmed with sequencing. The non-blood-fed mosquitoes were sorted
into pools of up to 50 individuals and screened for WNV by inoculation onto Vero cells
and by RT-PCR with WNV-specific primers sets. A total of 4484 mosquitoes (sixteen
different species and ten genera) were collected, 37 of which had visible blood meals.
Three species (seven individuals) were positive for alligator DNA: Culex erraticus,
Mansonia dyari, and Mansonia titillans. Other vertebrate blood meals were also
identified: raccoon, horse, turkey, and pig from Culex nigripalpus, Mansonia dyari,
Culex nigripalpus, and Anopheles quadrimaculatus and Mansonia dyari respectively. No
virus was detected in any of the pools. This study was able to identify three mosquito
species that fed on alligators, two of which (Mansonia spp.) have apparently not been
recorded feeding on reptiles before. Studies on vector competence will be necessary to
determine whether or not these mosquitoes are likely vectors of WNV on alligator farms.
West Nile Virus (WNV) is a Flavivirus (family Flaviviridae) and belongs to the
Japanese encephalitis serogroup. It is an enveloped, positive sense single stranded RNA
virus. WN virions are roughly spherical in shape and about 50 nm in diameter. WNV
infects a large range of vertebrates as well as invertebrate vectors, most notably
mosquitoes (Diptera: Culicidae) (Brinton, 2002).
West Nile virus was first isolated in 1937 in Uganda, from the blood of a woman
suffering mild febrile illness (Smithbum et al., 1940, as cited by Hubalek and Halouzka,
1999), and records show that it was present and infecting humans, birds, and mosquitoes
in Egypt in the 1950's (Melnick et al., 1951). Studies continued to expand the known
range of the virus, and WNV (or evidence of its transmission) has now been found in
many parts of Europe, the Middle East, Africa, China, and Southeast Asia. The closely
related Kunjin (KUN) virus is present in Southeast Asia and Australia. With this large
range, WNV is the most widespread flavivirus, although before 1999 it had not been
reported in the Americas. It has been isolated from over 40 different species of
mosquitoes in the Old World, with the genus Culex considered the primary enzootic and
epidemic vector and several species of Culex and Aedes demonstrated as competent
laboratory vectors. Culex univittatus Theobald is thought to be the principle vector in
Africa and Culexpipiens Linnaeus in Europe (Hubalek and Halouzka, 1999). The virus
is maintained in bird populations and spread with migrations (Rappole et al., 2000).
Vertical transmission in mosquitoes has been detected and may contribute to maintenance
of the virus (Miller et al., 2000). In Europe, transmission to humans occurs during
summer months (June to September) when mosquito vectors are most active (Hubalek
and Halouzka, 1999).
Each year in South Africa, there were sporadic cases of WN viral disease (WNVD)
often with mild illness. Two epidemics, one in 1974 and the other in 1984, marked a
change in that normal activity. These epidemics may have been due to unusually high
summer rains, which favored vector breeding and may have produced high vector
population densities, which in turn promoted feeding on non-avian hosts, especially with
the 1974 epidemic where more human cases were reported. Of all the WNV cases in
South Africa, only four have involved more serious illness, and only one
meningoencephalitis (Jupp, 2001).
In the late summer and fall of 1996, there was a major epidemic of WNVD in
southeastern Romania with the highest clinical incidence in the urban center of
Bucharest. WNV had been recorded in the area (by seroprevalence evidence) since the
1960's. This epidemic was the second largest recorded for Europe and was the first in
which many clinical cases showed involvement of the central nervous system (CNS).
Hospitals reported 17 deaths, and 400 cases of WN encephalitis, meningitis, or
meningoencephalitis. Sampling following the epidemic showed that eight percent of the
wild birds sampled and 41% of domestic birds had antibodies against WNV. Of about
6000 Culexpipienspipiens L. aspirated from man-made structures around Bucharest, one
was found positive for WNV, and the strain appeared to be most closely related to WNV
strains from sub-Saharan Africa. Among the factors that may have facilitated this
epidemic are the naivety of the population, the availability of flooded man-made
structures for mosquito breeding, and the summer drought that preceded the epidemic. In
the years following the Romanian epidemic, cases (some fatal) continued to occur and
seroconversions were observed in sentinel and domestic birds, although no WNV positive
mosquitoes (out of 23,000 tested over two years) were found (Campbell et al., 2001).
After the 1996 Romania outbreak, other epidemics of WNV-induced CNS disease
were reported in humans (including those in the United States, 1999-2004) (Lanciotti et
al., 2002). The large Romanian epidemic would turn out to be only a part of an
increasing trend of human and animal WNV outbreaks in Europe. Epidemics were
reported in Italy in 1998 and in Russia in 1999 (Brinton, 2002). In late summer through
fall of 2000, 131 WNV equine cases were reported in France, notably in an area with
colonies of migratory birds and plentiful mosquito breeding habitat (Murgue et al., 2001)
and during the fall of 2003 an outbreak caused disease in horses in Morocco
(Schuffeneker et al., 2005). In 2000, an epidemic of WNV in Israel led to 326
hospitalizations and 33 deaths. Severe cases were mostly in the elderly and involved the
CNS (Chowers et al., 2001). A study by Lanciotti et al. (2002) indicated that this
increased severity of disease was likely due to the greater virulence of the lineage 1 virus
responsible for these outbreaks.
In its Old World range, the virus appeared not to cause illness in wild birds with a
few exceptions (Bin et al., 2001). Similar to birds in the Old World, reptiles and
amphibians did not appear to suffer illness due to WNV, although evidence from multiple
studies demonstrated that they were subject to infection. Seropositive turtles were found
in Israel in the 1960's (Nir et al., 1969). Fourteen out of 20 healthy crocodiles
(Crocodylus niloticus: five males and 15 females between 1 and 2.5 years old) at a farm
in the Negev Desert in southern Israel were found to be seropositive for WNV, though no
deaths of crocodiles have been reported even during outbreaks of the virus in other
animals (humans, horses, and geese) (Steinman et al., 2003). Frogs (Rana sp.) were also
found with antibodies to WNV. Laboratory experiments showed that they could be
infected by the bite of an infective mosquito and could later re-infect biting mosquitoes,
thus demonstrating that they can be amplification hosts (Hubalek and Halouzka, 1999).
The first report of WNV in the Americas was from New York City in 1999. Since
then the virus has spread north, south, west and has now been detected in all 48 states in
the continental US except Washington (CDC, 2005), and has been reported in Canada
(Buck et al., 2003), the Caribbean (Quirin et al., 2004), Mexico, and Central America,
(Fernandez-Salas et al., 2003; Komar et al., 2003; Farfan-Ale et al., 2004; Cruz et al.,
2005). The transmission cycle has paralleled that of the Old World: bird and mosquito
(principally Culex) maintenance of the virus (Marfin et al., 2001; McLean et al., 2001)
spread of the virus with migrating birds, and illness in humans and horses (Huang et al.,
2002; Blackmore et al., 2003). The illness observed in humans and horses has been
similar to that seen during the more recent European epidemics with the virus affecting
the CNS in the more severe cases (Huang et al., 2002). Unlike in Africa and Europe,
WNV in North America has caused the death of many different species of bird (McLean
et al., 2001). Mortality in birds was so dependable that it actually became a warning
system for WNV activity (Mostashari et al., 2003). This greater mortality could be due in
part to the naivety of the birds in the New World, however, there is also experimental
evidence showing that the strain of WNV isolated in New York in 1999 is more
pathogenic to crows than Old World strains from Australia and Kenya (Brault et al.,
Sixty species of mosquito have been found infected with WNV thus far in the
United States (CDC, http://www.cdc.gov/ncidod/dvbid/westnile/mosquitoSpecies.htm,
2005) and many of these are competent laboratory vectors of the virus. Specifically
Culex stigmatosoma Dyar, Cx. erythrothorax Dyar, Cx. nigripalpus, Cx. pipiens, Cx.
quinquefasciatus Say, Cx. restuans Theobald, Cx. tarsalis Coquillett, and Cx. salinarius
Coquillett appear to be the most efficient enzootic vectors. Of these Cx. tarsalis, Cx.
salinarius, and Cx. erythrothorax appear to have the greatest potential as bridge vectors
although all have good potential. Other species like Ochlerotatus triseriatus (Say), Oc.
japonicus Theobald, and Aedes albopictus Skuse have a potential to serve as bridge
vectors (Turell et al., 2005). Not all species have been examined for their vector
competence; no member of the Melanoconion subgenus of Culex has yet been evaluated
(this subgenus is of special interest because some species are reptile-feeders). The impact
of WNV on North American reptiles has not been examined as closely as that of birds
and horses, and maybe there has been little impact overall. Common garter snakes
(Thamnophis sirtalis sirtalis (Linnaeus)) and red-ear sliders (Trachemys script elegans
(Wied-NeuWied)) did not develop detectable viremia after subcutaneous inoculation with
WNV. North American bullfrogs (Rana catesbeiana Shaw) and Green iguanas (Iguana
iguana (Linnaeus)) (infected by mosquito bite) did develop detectable viremia, although
not more than 103.2 PFU/mL (Plaque Forming Units, with one PFU equivalent to one
viable virus particle) serum which is lower than needed to efficiently infect a biting
mosquito such as Cx quinquefasciatus (Klenk and Komer, 2003; Jupp, 1974). Serious
morbidity was not noted (Klenk and Komer, 2003). In contrast, over the past several
years, alligator farms in Florida, Georgia, and Louisiana have experienced sudden die-
offs of juvenile and hatchling alligators (Alligator mississippiensis Daudin). These
events occurred in the fall and tended to last two or three weeks. Histological findings,
virus culture, and results from Reverse Transcriptase Polymerase Chain Reaction (RT-
PCR) all suggest that the deaths were caused, at least in part, by infection with WNV
(Miller et al., 2003; Jacobson et al., 2005a).
WNV in Farmed Alligators
In the US, alligators are grown commercially for their hide and meat, with the hide
being the more valuable raw product. The value of a 2 m alligator (about three years old
if grown in an intensive system) is about $US 150, with the major demand for hides and
meat coming from Japan, Europe, and North America (Florida Fish and Wildlife
Conservation Commission (FFWCC) report; Lane and King, 1989). About 1,500,000
crocodilian hides are traded per year with Florida, Texas, and Louisiana producing about
45,000 of that total, including hides from wild caught alligators. In 2003, Florida farms
produced 22,527 alligators at a value of about $3.3 million (FFWCC report). Alligators
are usually kept in temperature-controlled (ideally about 860 F, 300 C), dark pens and fed
pellet feeds and raw meats (Lane and King, 1989).
The first group to describe the epizootics of WNV in alligator farms was Miller et
al. (2003) when they investigated and reported on two die-offs occurring during the fall
of 2001 and 2002 at a farm in southern Georgia. They observed "stargazing" before
death, loss of leg control, and neck spasms in hatchling and juvenile alligators. Tissue
was collected from the eye, thyroid gland, lymph node, lung, heart, brain, spinal cord,
kidney, liver, spleen, pancreas, adrenal gland, gallbladder, tonsil, trachea, stomach,
intestines, and reproductive tract. Tissues and blood were subjected to RT-PCR, virus
isolation, and bacterial culture. The appearance of the tissues and the RT-PCR results
strongly suggested that West Nile virus was the cause of death, or had weakened the
animals' immune systems such that bacterial infection set in. Raw horsemeat is a part of
the alligator diet and was tested for WNV RNA. The meat that was fed to the alligators
during the epizootics was positive for WNV by RT-PCR but was negative after the
epizootic ended, leading the researchers to believe that virus in the horsemeat had caused
the epizootic. Supporting this idea are experiments that have demonstrated that mice and
hamsters can become infected when fed a fluid containing WNV (Odelola and Oduye,
1977; Sbrana et al., 2005). There have also been cases of predators becoming infected
with WNV after eating infected prey (Garmendia et al., 2000; Austgen et al., 2004).
In Florida, similar epizootics occurred on several farms and one farm was
investigated by Jacobson et al. (2005a). In 2002, an epizootic on a central Florida farm
(named Farm A from here on) killed 300 of the 9000 alligators at the farm. Clinical signs
in the alligators included anorexia, lethargy, tremors, swimming on the side, and
opisthotonus. Tissues of three alligators were examined and showed signs of CNS
disease and necrotizing hepatitis. Immunostaining revealed the presence of WNV
antigen in multiple tissues. There was no evidence of two other pathogens that have
previously been identified as disease agents in Crocodilians: Mycoplasma and
Chlamydia. In contrast to the findings from Georgia, no secondary bacterial infection
was apparent. Viremia in the infected alligators was greater than 105.o PFU/ml plasma
making the alligators capable of infecting mosquitoes like Cx. quinquefasciatus and Cx.
pipiens (Jacobson et al., 2005a). Unlike the farm investigated in Georgia, Farm A feeds
the alligators beef and alligator chow.
Illness occurred only in some pens on Farm A, with the affected pens containing
multiple sick animals. Jacobson et al. (2005b) found that all blood sampled alligators that
had shared a pen with sick animals during the epizootic carried WNV-neutralizing
antibodies three months later, while those sampled from pens where no disease was
recorded were not found to have such antibodies. This demonstrated that horizontal
transmission had likely occurred inside the pens and suggested that the sporadic pattern
of infection could be due to the infection of one alligator followed by viral shedding and
infection of all of the other alligators sharing that pen. Laboratory experiments
conducted by Klenk et al. (2004) confirmed the potential for horizontal transmission of
WNV between alligators. In the laboratory, American alligators were injected
subcutaneously with 7500 PFU of WNV or were fed viremic mice. All alligators
developed viremia within three to six days. The viremia persisted for about ten days and
reached approximately 106 PFU/mL. Uninoculated tank mates also became viremic
about a week after the inoculated alligators. Viral shedding from the cloacae was
detected and was suspected to be responsible for horizontal transmission between tank
mates. Two of the 29 infected alligators died, while the others developed WNV
neutralizing antibodies within 25 days of the onset of viremia. These experiments
demonstrated that horizontal transmission to tank mates does occur (100% in the study),
viral shedding does occur, alligators can become infected through the oral route, and that
the viremia of the alligators is high enough (Jupp, 1974) to infect biting mosquitoes
making them potential amplification hosts of the virus.
Alligator farmers in Florida are not required to report the cause of death of their
alligators, so there are no precise records of these epizootics, their impact, epidemiology,
and timing. Florida farmers are required to report all deaths annually to the Florida Fish
and Wildlife Conservation Commission, who then make this information available to the
public. While it is impossible to make many conclusions based on gross annual records,
it was clear in 2002 that some farms had virtually no unusual deaths while others
apparently lost 10-50% of their alligators due to causes other than intentional slaughter
(FFWCC 2002 annual report, and Dwayne Carbonneau personal communication).
Mosquitoes as Vectors of WNV on Alligator Farms
There are two basic explanations for the source of the outbreaks of WNV on
alligator farms, and they are not necessarily mutually exclusive. The first is that the virus
is introduced by the bite of an infective mosquito, and the second is that the virus is
introduced when the alligators are fed raw meat that contained active virus, an
explanation supported by the findings of Miller et al. (2003). As of yet, no studies have
been published that explore the potential for mosquito transmission of West Nile virus on
alligator farms. The search for potential vectors of WNV in farmed alligators can be
guided by a few criteria presented by Reeves (1957) (as reviewed in Turell et al. (2005))
and by Kilpatrick et al. (2005). A potential vector will repeatedly be found naturally
infected with the virus and will be found in association (during the time when
transmission is occurring) with the naturally infected vertebrate hosts, in this case, the
alligators. If the potential vector is found in large numbers around the infected host, this
should increase the chance that it is responsible for transmission (Kilpatrick et al., 2005).
A potential vector should also be able to transmit the virus efficiently as demonstrated
through laboratory competence studies. This study intended to identify potential
mosquito vectors of WNV in Florida farmed alligators by finding those mosquitoes that
were numerous, and associated with (specifically feeding on) farmed alligators and
determining if those associated mosquito species were also naturally infected with WNV.
Blood Meal Identification
A number of methods have been used to determine the hosts from which different
mosquito species take blood meals. Observation of feeding mosquitoes, capture of
mosquitoes in host baited traps, analysis of cytological characteristics of blood meals,
analysis of serological characteristics of blood meals, and genetic information contained
in blood cells have all been used to determine the host preferences of mosquitoes, with
the last two of these five methods being the most commonly used today (Tempelis, 1975;
Ngo and Kramer, 2003). The basic principle underlying the serological method is that
antiserum (made when blood from various hosts is injected into other animals) will react
with certain unidentified but unique elements in the blood of different hosts. Different
techniques use this principle. In precipitin tests a suspension of the blood meal is mixed
with antiserums against different vertebrates and if there is a reaction (portions of blood
meal binding with antiserum) a precipitate forms and the meal is considered positive for
that host type (Tempelis, 1975). The Enzyme-Linked ImmunoSorbent Assay (ELISA)
test uses an enzyme-linked color change to signal when binding has occurred between the
specific antibody and the reacting element in the blood meal. Fluorescent antibodies
again rely on serology, with the fluorescence enhancing visualization of positive matches.
The technique developed most recently uses genetic characteristics of a blood meal
to determine the host, in particular the technique relies on detection of specific regions of
host DNA (usually mitochondrial) in the blood cells. Primers have been designed to
amplify a region of the cytochrome b gene only for certain groups of vertebrates; there
are primer sets for all mammals, all birds, and for different orders of birds (Cicero and
Johnson, 2001; Ngo and Kramer, 2003). Sequencing the fragment, followed by matching
with known sequences in the BLAST database of GenBank, can confirm blood meal
identifications or take the identification further, to family, genus, or species. By using
these primers, host DNA could be detected in Cx. pipiens for up to 3 days after feeding
(at 270C) (Ngo and Kramer, 2003).
For these two techniques, naturally engorged females are collected from the field
and the blood meal analysis is done in the laboratory. Different methods can be used to
capture engorged females, and often the method chosen and the exact microhabitats
sampled will depend on which mosquito species the study is targeting. The three
collection methods used in this study were vacuum aspiration, CDC light traps (CDC =
Centers for Disease Control and Prevention), and wooden resting boxes. With vacuum
aspiration a battery-operated vacuum sucks mosquitoes against a screen until they can be
transferred to a separate container. Aspiration can be done in vegetation, animal burrows,
man-made objects/structures, and in natural and artificial crevices such as around tree
roots or mosquito resting boxes. Vacuum aspiration has been used in Florida to collect
Ae. albopictus, Culex of the subgenus Melanoconion, Cx. nigripalpus, Culex, Aedes,
Anopheles, Coquillettidia, Mansonia and Psorophora. (Nieblyski et al., 1994; Edman,
1979; Day and Curtis, 1993; Edman, 1971).
A CDC light trap makes use of light and CO2 to attract mosquitoes close to the trap
where a fan-generated air current draws them into a collection jar or bag (Sudia and
Chamberlain, 1988). In this study white incandescent lights were used. Field research in
Florida and Georgia has shown white lights to be attractive to (among others)
Uranotaenia sapphirina (Osten Saken), An. crucians (Wiedemann), Ae. vexans (Meigen),
An. quadrimaculatus Say, Ae. atlanticus Dyar and Knab, Cx. nigripalpus, and Culex of
the subgenus Melanoconion (Love and Smith, 1957; Burkett et al., 1998). The addition
of CO2 as bait dramatically increases overall catch numbers of most mosquitoes (Burkett
et al., 1998; Reisen et al., 1999). CDC traps are often left operating from before dusk
until dawn in order to attract mosquitoes when their flight activity is maximum
Resting boxes are containers designed to resemble mosquitoes' natural resting
places. They are often used to study host preferences because they attract females that
are seeking a dark place to remain while digesting the blood meal and developing eggs.
Resting boxes (with gray outside and red inside) set out on an island in the marshes near
Vero Beach, Florida attracted Melanoconion Culex and Uranotaenia in swampy areas,
and Culiseta melanura (Coquillett) and Anopheles near higher, hammock sites.
Mosquitoes tended to enter during the mornings and leave during the day although some
entered at all times (Edman et al., 1968).
A large body of work based on the different methods of host identification has
allowed some generalizations about the feeding habits of different mosquito genera and
species in North America. Species that fed exclusively on one class of vertebrate were
perhaps the exception rather than the rule. Regional variation, seasonal variation, and
habitat-linked variation in host preferences were observed. A number of different
mosquito genera and species feed on reptiles and/or amphibians ectothermss). Some
appear to feed mostly on reptiles or amphibians, or even particular orders of ectotherms.
Others appear to take meals from reptiles only occasionally, while primarily feeding on
mammals, birds, or both.
A number of studies from locations through out the eastern United States have
found that some mosquitoes will occasionally take meals from reptiles. Ae. atlanticus,
Ochlerotatus. triseriatus, and Oc. sollicitans (Walker) were found to feed on turtles,
although in general mosquitoes of the genus Aedes fed on mammals and to a lesser extent
birds (Tempelis, 1975). Turtle blood meals were identified from Cx. salinarius, Cx.
pipiens, and from Coquillettidiaperturbans (Walker) in New York. These three species
were also found to feed on mammals and birds in the same locations (Appersen et al.,
2002). In Florida, Oc. infirmatus Dyar and Knab, Ae. taeniorhynchus (Wiedemann), Ae.
albopictus, Ae. vexans, Culiseta melanura, Cx. territans, Cx. salinarius, and An. crucians
fed on one or more of the following reptiles: snake, turtle, and lizard (Edman, 1971;
Edman et al., 1972; Nieblyski et al., 1994). In North Carolina, Ae. atlanticus, Oc.
henderson, Ae. vexans, Psorophora columbiae (Dyar & Knab), Ps. ferox Humboldt, Ps.
howardii Coquillett, Cs. melanura, Cx. quinquefasciatus, and Cx. restuans were all found
with some reptile blood meals, though a majority of their meals were from non-reptilian
hosts (Irby and Apperson, 1988). Seventeen percent of the meals identified from Cs.
melanura in a Maryland study were from reptiles (Moussa et al., 1966). Of the engorged
mosquitoes collected during a study in central Alabama, about 2% of Cx. erraticus Dyar
and Knab were found to contain reptilian blood meals (Cupp et al., 2004). Animal baited
traps in Delaware showed that Oc. sollicitans, An. quadrimaculatus, and Cq. perturbans
occasionally fed on different reptiles but were better represented in traps with mammal or
bird hosts (Murphey et al., 1967). Mosquitoes in the genus Deinocerites appear to be
opportunistic feeders, taking meals from mammals, birds, amphibians, and reptiles
Many of these same studies also found species that took a majority or even all of
their meals from ectotherms. The Delaware (Murphey et al., 1967) study found that Cx.
territans Walker were frequently attracted to king snakes, water snakes, snapping turtles,
and Eastern box turtles, but were not attracted to the mammals and birds tested. In
Alabama Cupp et al. (2004) found that 75% of the Cx. peccator (Dyar & Knab) that they
collected had fed on ectotherms, including one Crocodilian. In their North Carolina
study, Irby and Apperson (1988) found that Cx. territans and Cx. peccator fed almost
exclusively on reptiles and amphibians (about 99% of meals from ectotherms and 1%
from birds). Culex erraticus and Cx. territans were collected from lizard (Anolis
carolinensis Voigt) baited traps in north central Florida and readily fed on the lizards
both in the traps and in the laboratory (Klein et al., 1987). Ocholerotatus canadensis
Theobald (=Aedes canadensis) was the most frequent mosquito encountered around wild
turtles in one study, and later research in North Carolina showed that 85% of the
individuals sampled had taken their meal from an ectotherm (Irby and Apperson, 1988).
With the wild turtles, most feeding took place around the head, neck, and legs, and
sometimes between the scutes of the turtle's carapace (Crans and Rockel, 1968). These
two studies also found that Ae. triseriatus was frequently attracted to or feeding on
A study in Panama (Tempelis and Galindo, 1975) examined Culex species in the
Neotropical subgenus Melanoconion (of which there are seven species in Florida) and
found that four species fed mostly on lizards: Cx. egcymon Dyar (81%), Cx. tecmarsis
Dyar (89%), Cx. elevator Dyar and Knab (90%) and Cx. dunni Dyar (63%) while the
other Melanoconion species in the study fed mostly on birds and mammals. This finding
in Panama, that multiple Culex species in the subgenus Melanoconion feed on reptiles, is
consistent with the findings in the United States.
Efforts to determine which species of mosquitoes) feed on alligators at a farm
would likely have the greatest chance of success if they concentrated on sampling blood
fed mosquitoes of the species that have already been identified feeding on reptiles. Based
on previous Florida studies mentioned before, the three sampling techniques used
(vacuum aspiration, CDC light traps, and resting boxes) should yield most, if not all, of
the species that have been recorded feeding on reptiles, assuming that they occur in the
vicinity of the farm.
In this study, the PCR-based method for analysis of blood meal was used, with
Crocodilian-specific primers designed by Yau et al. (2002) that amplify a segment of
chromosomal DNA and with Alligatoridae-specific primers based on work by Janke and
Arnason, (1997), Ray and Densmore, (2002), and Glenn et al. (2002). The Alligatoridae-
specific primers amplify a region of mitochondrial DNA, including portions of the
cytochrome b gene, and genes for transfer RNAs. The location within the genome and
the coding nature of the fragment amplified by the Crocodilian-specific primers are
A group of animals that contains enough viremic individuals to continually infect
mosquitoes constitutes the reservoir, and the vertebrate reservoirs of WNV are most often
birds (McLean et al., 2001). Thus a likely vector on the alligator farm would be a
mosquito species that fed on both birds and alligators, such that it could move virus from
populations of infected birds to the alligators. To determine if the mosquito species
found feeding on alligators were also feeding on birds, an avian-specific primer set was
used. In addition, a mammalian-specific primer set was used to gain more information
about the feeding habits of the mosquitoes captured around the alligator farm. These
primer sets amplify a region of cytochrome b gene in the mitochondrial DNA for birds
and mammals respectively (Ngo and Kramer, 2003).
Screening Mosquitoes for WNV
Potential vectors not only must feed on the host, but must also be infective.
Mosquitoes collected from the alligator farm were tested for the presence of WNV.
Work that is testing for viremic animals or for infected mosquitoes requires direct
evidence of the virus particles (as opposed to testing for virus-neutralizing antibodies).
Active virus from mosquito pools or tissues of viremic animals can be isolated in cell
culture or the presence of viral nucleic acid can be demonstrated with strain-specific
oligonucleotide primers and RT-PCR. In most recently published WNV research or
surveillance reports, two tests (some using two different techniques) were often done to
confirm a positive (and sometimes negatives as well). Often results from a real-time or
standard RT-PCR test were confirmed with a second RT-PCR with a different primer set
or with isolation of virus from the sample using cell culture (Kauffman et al., 2003;
Lanciotti et al., 2000; Bernard et al., 2001). In this study, mosquito pools were tested for
presence of virus by inoculation onto Vero cell monolayers, and by RT-PCR analysis
with WNV-specific primers.
Kidney cells from the African Green monkey (Vero cells) are used in WNV
isolation because they show cytopathic effects when infected by the virus, usually visible
after three days (Odelola and Fabiyi, 1977). The virus binds to cells and enters by
receptor-mediated endocytosis (Chu et al., 2005). The capsid releases the positive single
stranded RNA which is treated as messenger RNA by the cells and the single -10,000
base pair open reading frame is translated into a single protein which is then cleaved by
cellular and viral proteases (Brinton, 2002). Translation of the viral proteins is associated
with the rough endoplasmic reticulum (Lee and Ng, 2004). The seven resultant non-
structural proteins can then make a negative strand copy of the viral RNA, which serves
as a template for new positive strand RNAs that can associate with structural proteins to
form new virions. New virions move to the cell's margin in membrane vesicles, and are
released by budding, individually at first and later in "bags" (Brinton, 2002). Budding of
new virions starts within 10-12 hours after infection and is at maximum about 24 hours
after infection (Ng et al., 2001). This process may perceivably slow the growth and
division of the Vero cells, however, distinct cytopathic effects are usually first visible
three days post-inoculation (Odelola and Fabiyi, 1977). Cells appear more rounded, with
thicker, more distinct margins. They may appear "grainy" with vacuoles. As the cells
die, they disconnect from the substrate. Vero cells are usually monitored for seven days
after inoculation with mosquito homogenate (Kauffman et al., 2003). With virus
isolation in cell culture, only active virus can be detected, and some work has suggested
that it may not be as sensitive as RT-PCR (Nasci et al., 2002).
RT-PCR with primers specific for WNV was used to detect viral RNA in the
samples. Two primer sets were used. Set one was used to screen the pools for WNV
RNA and the second was used to confirm any positive bands from the first set. The first
set, WN9483 and WN9794, were based on suggestions made by the CDC (based on work
by Lanciotti). These primers amplify a 311 base-pair region within the NS5 gene, the
gene which codes for the viral RNA-dependent RNA polymerase (Lanciotti et al., 1999).
This polymerase is the most highly conserved protein of West Nile virus (and of
flaviviruses in general) (Brinton, 2002). As a consequence of the conserved nature of the
region, WN9483 and WN9794 should readily bind to any potential strain of WNV.
The second set of primers, WN212 and WN1229, is based on suggestions of the
CDC and work by Lanciotti et al., (2002). WN212 binds to a region within the gene for
the viral nucleocapsid protein and WN1229 binds to a region within the envelope
glycoprotein gene. The envelope protein gene is the more variable region of the WNV
genome, but little genetic variation has been observed among US strains of WNV up to
2003 (Ebel et al., 2004), and new strains isolated since 2002 still have around 99.7%
homology to strains isolated in New York in 1999 (Davis et al., 2004). Consequently this
primer set will also likely bind to all potential strains of WNV.
The sensitivity of these approaches should allow for detection of mosquitoes that
are potentially infective. To vector an arbovirus, a mosquito must have a minimum of
about 105 virions disseminated within its body (Hardy et al., 1983), and a fully
disseminated infection in a mosquito with WNV is often more than this, about 106.5
virions in the whole mosquito when measured 14 days after oral inoculation (Johnson et
al., 2003). Mosquitoes encountered in the field may have lower titers than the minimum
105 virions, titers that may be below the detection limit of the techniques applied in this
study. However, because mosquitoes with such low titers are unlikely to be capable of
efficiently transmitting WNV (Hardy et al., 1983), they are relevant to the search for
In general, the numbers of mosquitoes in a field collection that are found positive
for WNV are low (Bernard et al., 2001), and it appears that the number of positives out of
the total number collected (the Minimum Infection Rate = MIR) is not greater than 1 in a
1000 unless the collection was made in the vicinity of transmission (as demonstrated by
human, horse, or bird cases) (Bernard et al., 2001.). However, it is difficult to make a
generalization. MIR's vary considerably between studies and surveillance reports, and
are likely influenced by the time of collection, the proportion of the collection comprised
by "high risk" species like Culex, the age composition of the collections, and other
factors that are difficult to quantify.
The objectives in this study were to identify potential vectors of WNV on Farm A
using three of the four criteria described above. Mosquitoes were captured around the
farm to determine which species fit the following criteria:
1. Species is present around host (alligators) during the time of transmission
2. Species is feeding on host
3. Species is infected with WNV
The fourth criteria, vector competence, was not addressed in this study.
METHODS AND MATERIALS
Mosquitoes were captured, identified, and counted to determine which species were
common around the farm. Mosquito blood meals were tested for presence of alligator
DNA to determine which species were feeding on the alligators and were tested for the
presence of avian and mammalian DNA to see if the alligator-feeding species were also
feeding on other animals around the farm. Unengorged mosquitoes were screened for
WNV to determine if any species had a high MIR.
Four over night collecting trips were made to Farm A.
Trip 1. On September 9, 2003, the first collecting trip was made to Farm A
alligator farm in Christmas, FL (Orange County, east of Orlando, on highway 50). At the
time of this trip there had been multiple alligator deaths, many consistent with WNV
infection. Equipment included one battery-powered backpack aspirator, plastic bags for
collecting samples of feed and aspirator samples, a cooler with ice to keep samples cold
during transit, and aerial nets for sweep net collecting in the vegetation around the farm.
Active collecting began mid-morning. The interior and exterior walls of pens and
other buildings were visually scanned for resting mosquitoes. Insects were aspirated
from vegetation, buildings and construction debris. Insects were also collected from
vegetation using sweep nets. Around mid-day four CDC light traps (Sudia and
Chamberlain, 1988) baited with CO2 from dry ice were set. The dry ice was contained in
an insulated plastic box with a small opening for outflow (MEDUSA Patent # 5,228,233
and # 5,272,179). Plastic tubing directed the flow of carbon dioxide from the metal box,
through a bottle of water, and to the light trap. Small plastic vials with a sugar solution
and a cotton stopper were taped inside of the collection jars of the CDC traps. Two traps
were hung from low branches about 1 meter above the ground on trees along a chain-link
fence that separated the farm from adjacent property (Fig 2-1).
Figure 2-1. CDC light trap set up on the western margin of the farm. CO2 came from dry
ice inside the insulated white plastic box.
The adjacent property was mostly wooded and was home to several pigs and at
least one horse.
The other two CDC traps were hung inside of alligator pens where some alligator
deaths had occurred in the past two years. They were hung from support pipes close to
the door, also about 1 meter from the ground. The weather was sunny and warm when
traps were set out and when collected.
Samples from sweep netting and aspirating were transferred to plastic bags, put on
ice in the cooler, and taken to the lab in Gainesville where they were stored in -700C until
processed. The CDC traps were left over night. The traps were removed and samples
collected the next day around the same time that they had been originally set up.
Samples were put on ice, taken to Gainesville, transferred to plastic bags, and stored at -
700C until processed.
Trip two. A second collecting trip was made on September 24, 2003. Four CDC
light traps were set inside four separate pens, each of which had housed alligators that
died from illness consistent with WNV within the past two years. In addition, eight
resting boxes (Moussa et al., 1966) were set up around the farm: four along the eastern
margin of the farm, abutting a body of freshwater, three along the western margin of the
farm close to the chain-link fence, and one inside of an alligator pen, where a CDC trap
had also been placed. The resting boxes were wooden cubes roughly one foot on each
side (30 cm) and open on one face. The open side of each box was fitted with a square of
mesh and Velcro such that the mesh could be pulled down and secured over the opening
to trap any mosquitoes that had gone inside the box. The outer surfaces of the box were
painted black with acrylic paint, and the inside surfaces were painted a maroon color
Figure 2-2. A 30 cm x 30 cm x 30 cm wooden resting box with black exterior and
maroon interior was used to attract blood fed mosquitoes.
The CDC traps and resting boxes were set in the early afternoon on Sept. 25, left
over night, and collected at about the same time the following day. There was light rain
when the traps were set out and the weather was overcast with showers in the area when
the traps were collected. The following steps were conducted to collect the mosquitoes
from the resting boxes:
1. Boxes were approached from "behind" (the side opposite the open face);
2. From behind, screen was secured over the open face;
3. Boxes were then brought one at a time into the cab of a truck;
4. The screen was carefully pulled back and any mosquitoes aspirated with a
Dustbuster vacuum fitted with a plastic tube. Any mosquitoes that escaped into the
cab of the truck were also aspirated. Gauze was secured over the mouth of the
Dustbuster vacuum so that mosquitoes that were aspirated into the plastic tube
would not be sucked into the Dustbuster.;
5. A gauze stopper was put in both ends of the plastic tube to trap mosquitoes, and
tubes were then placed inside a cooler with ice.
Trip three. The third trip was made on October 17, 2003. Traps were set around
2:30 PM inside and outside of alligator pens, although records showing the exact
locations of the traps were lost while in transit from Farm A to Gainesville. Traps were
collected the following afternoon. Resting boxes were collected first, starting around
1:00 PM. All traps had been collected by 4:00 PM. All samples were kept on ice during
the trip. For this trip and the following trip, the source of CO2 bait was switched from
dry ice to compressed gas in tanks. The regulators on the tanks were set to a flow rate of
500 mL/min. The weather was clear and warm both days.
Trip four. A fourth and final trip was made on October 24, 2003. Traps were set
out around 3:00 PM. Two CDC traps were hung from trees on the eastern side of the
farm, adjacent to the adult alligator lagoon. Three were hung inside of alligator pens: pen
# 14 with small alligators and recorded deaths, pen # 15 with medium alligators and
recorded deaths, and pen # 10 with medium alligators and no recorded deaths. One CDC
trap was hung from the gate to the enclosure with the rectangular pens housing large
alligators. One was hung from a tree in the middle of the farm and another was hung in
the trees along the western margin of the farm adjacent to the neighboring property. Of
the eight resting boxes, two were placed along the eastern edge of the farm adjacent to
the lagoon, and the other six were set up along the western edge of the farm (Fig. 2-3).
Traps were collected the following afternoon. The weather was clear and warm both
In Gainesville, mosquitoes were separated into pools of 1 50 individuals based on
presence of blood meals, species (Darsie, 1998), trap type and number, and trap date.
Figure 2-3. Map depicting layout of Farm A. Alligator pens where deaths had occurred
are blue; pens with no history of deaths are yellow. Large red stars indicate
where resting boxes were placed. Smaller green stars indicate where CDC
light traps were placed. The structures indicated with brown outlines were
buildings used for purposes like storage of maintenance equipment, housing
for water heaters, basins for wastewater, and an office.
All identification and sorting was done on top of a chill table, and all pools were placed
in tubes (blood fed mosquitoes singly in 1.5 mL microcentrifuge tubes, and non blood fed
pools in 2 mL graduated microcentrifuge tubes (OPS, Petaluma, CA) with 1-2 copper-
clad steel beads (BB-caliber airgun shot)) and stored at -700C until processed.
Each pool was assigned a code name. For the pools ofunengorged mosquitoes, the
code names started with a digit one through four that corresponded to the collecting trip
when mosquitoes were captured. Letters were used to designate each pool, and the code
ended with a digit that indicated the trap the mosquitoes were from. For the engorged
mosquitoes the first digit also designated the collection trip and the letters were shorthand
for the genus and/or species of mosquito, BF stood for "blood fed", and the end digits
described either the trap number or were used to distinguish multiple mosquitoes that
were from the same species, date, and trap number.
Blood Meal Identification
Prior to working with field-collected mosquitoes, extraction and PCR procedures
were tested and optimized on positive controls. To form a positive control for the
alligator bloodmeal study, Ae. aegypti Linneaus and Cx. quinquefasciatus mosquitoes
were obtained from the USDA (United States Department of Agriculture), Gainesville
colonies. These mosquitoes were starved for 24 hours and then offered one of two
liquids using the membrane feeding system (Davis et al., 1983; McKenzie, 2003).
Briefly, one mL of the liquid was placed into the depression in the bottom of a film
canister lid, a square of membrane (bridal veil with a layer of silicon) was placed so that
it covered the depression, and then the membrane was secured over the canister lid using
a plastic ring. This membrane feeding system was then inverted and put on the top the
wire-mesh mosquito cages such that mosquitoes could insert their proboscis through the
mesh of their cages, through the silicon layer of the membrane, and into the liquid. The
two liquids offered to the mosquitoes in this manner were: heparinized alligator blood
and meat juice from previously frozen alligator tail meat that was sweetened with 10%
sucrose sugar (Figure 2-4). The sugar was added to encourage feeding (Aissa
Doumbouya, personal communication). The alligator blood was drawn from the sinus
vein of an alligator patient at the large animal clinic of the University of Florida School
of Veterinary medicine, and was provided by Dr. Darryl Heard (use of blood approved,
UF animal use protocol #D687). Mosquitoes were allowed to feed for 24 hours after
which they were frozen at -200C, and the engorged individuals were separated for later
Figure 2-4. The membrane feeding
system was used to feed
alligator blood and alligator meat juice to Cx. quinquefasciatus and Ae.
aegypti mosquitoes to be used as positive controls when testing field-collected
mosquitoes for the presence of alligator blood.
The positive controls for testing the avian-specific and mammal-specific primers
were a mosquito fed on a live chicken (feeding that is a normal part of colony
maintenance at the USDA) and a Coquillitidiaperturbans captured after it had fed on the
investigator. After it was established that the avian primers worked for the chicken-fed
mosquito, DNA from a rock dove (Columba livia G.F. Gmelin) was used as the avian
positive control for PCR reactions.
The abdomens of both the engorged positive control mosquitoes and an un-
engorged negative control mosquito were removed, placed separately into 1.5 mL plastic
tubes, and homogenized in 250 ptl of buffer 1 (buffer 1 contains 0.32 M sucrose, 50 mM
Tris at pH 7.25, 10 mM MgC12, and 0.5% NP 40 detergent) using a plastic mortar. The
tubes were then centrifuged at 6000 rpm for four minutes to pellet the mosquito cells and
parts, and the supernatant was discarded. The pellet was resuspended in a second buffer
(75 mM NaC1, 25 mM EDTA, and 10 mM Tris at pH 7.8) to lyse the cells. Fifteen pl of
0.5 M EDTA, 15 pl of 20% SDS, and 8 ptl of proteinase K (20 mg/mL) were added and
the mixture was incubated overnight in a 55 C water bath. The following day the tubes
were centrifuged at 13,000 rpm for 10 minutes and the supernatant was transferred to a
new tube. Twenty ptl of RNAse (5 mg/ml) was added and the mixture was incubated at
37 C for one hour. Following incubation, DNA was separated using phenol and
chloroform followed by a second precipitation using only chloroform. DNA was
precipitated from the aqueous phase with 600 ptl of cold 95% ethanol followed by
centrifugation (13,000 rpm at 4 C for 10 minutes). The ethanol was then removed and
the pellet was vacuum dried. The DNA was resuspended in 30 ptl of 10 mM Tris and
stored at -20 C until used in PCR reactions. Three other DNA extraction protocols
(TRIZOL, C-TAB, and DNeasy) were tried on the positive controls. Only the one
described above was used on the field samples because it was the easiest protocol that
consistently gave good final DNA concentrations.
The DNA was tested to determine its vertebrate origin with multiple primer sets:
Crocodilian-specific primers described by Yau et al. (2002), mammal-specific primers
described in Ngo and Kramer (2002), and bird-specific primers described by Cicero and
Johnson (2001). In addition primers for alligators were designed for this study based on
the suggestions of Glen et al. (2002). Primers were made using the Primer3 program and
the mitochondrial genome ofA. mississippiensis from GenBank, accession number
Y13113, (Janke and Arnason, 1997) (Table 2-1). This primer set will be referred to as
the alligator-specific set, although they may also amplify DNA from other members of
the Alligatoridae family or Crocodilian order. The Crocodilian-specific primers amplify
a region of chromosomal DNA, while the alligator, avian, and mammalian primers
amplify a mitochondrial region including parts of the cytochrome b gene. The conditions
used with the mammal and bird primers closely followed those described in the original
papers. For the Crocodilian and the alligator primers several optimization experiments
were done to find the conditions under which the positive controls would consistently
amplify. These experiments tested for optimal concentrations of MgC12, primers,
template DNA, and for the optimal annealing temperature. Once positive controls were
working consistently, DNA was extracted from the field-collected mosquitoes using the
same protocol as before, and each mosquito sample was tested for vertebrate DNA using
each of the four primer sets. Reaction conditions and the thermocycle program for each
of the primer sets are described in Table 2-3.
The PCR products were run on 1% agarose gels stained with ethidium bromide.
Any bands were excised from the gel, DNA was purified using the QIAquick Spin kit for
gel extraction (Quiagen, Valencia CA) following the handbook protocol (Appendix A),
and the fragments were sequenced using the BigDye Terminator Cycle Sequencing kit
(PE Biosystems, Foster City CA) following a protocol modified from the kit instructions
(Appendix B). Sequences were run at the University of Florida ICBR (Interdisciplinary
Center for Biotechnology Research) core facility in Gainesville, Florida. The sequences
were then edited using SequencherTM version 4.1 software (Gene Codes Co., Ann Arbor,
MI) and compared to all those on the BLAST database (GenBank) to identify hosts with
more certainty and specificity.
For the three samples that did not produce clear sequences, the PCR products were
inserted into pGEM-T vector (Promega, Madison, WI) according to the kit instructions
(Appendix C) with an overnight incubation at 4 C. Following incubation the vectors
were prepared for transformation into Escherichia coli bacteria by heat inactivating the
ligase at 65 C for 10 min, diluting the DNA (x three) with sterile water, and sterilizing
the DNA with 300 [tl of ether. Vectors were then transformed into bacteria. Five il of
the DNA ligation mixture was mixed with 50 [l of competent bacterial cells, and the
combination was incubated for 30 min on ice and then heat shocked for 30 s at 37 C.
The cell mixture was left for 2 min on ice, then 0.95 mL of medium deionizedd water
with 2% bacto-tryptone, 0.5% bacto-yeast extract, and 0.05% NaC1) was added and the
bacteria were set in a 37 C water bath and shaken at 225 rpm for 1 h. This mixture was
then plated onto LB agar plates with 100 [g/mL ampicillin and 20 [g/mL X-gal, plates
were incubated at 37 C overnight, transformed colonies were selected, and transformed
bacteria were grown overnight in media (4 mL LB medium with 5 mg/mL ampicillin) at
55 C. The plasmid was removed using the QIAprepTM Spin Miniprep kit (QIAGEN,
Valencia, CA) following kit instructions (Appendix D), and the insert was removed from
the plasmid by digestion with EcoRI at 37 C in a mixture containing 7 tl water, 1 tl
reaction buffer, 0.3 [il EcoRI, and 2 [il plasmid DNA. The insert was then sequenced as
Table 2-1. Primers sets in PCR used to amplify DNA from different vertebrate hosts.
Host Primer Sequence size(bp) Reference
Alligator Forward CGCTTCACTGCCCTACACTT 850 Current study
Yau et al.
Crocodilian Forward GATGTGGACCTTCAGGATGC 209 (2002)
Avian Forward GACTGTGACAAAATCCCNTTCCA 508 Johnson (2001)
MammalianForward CGAAGCTTGATATGAAAAACCATCGTTG 772 Kramer (2003)
About 12 hours prior to virus work, each well of 24-cluster well plates was
inoculated with 5.0 x 104 Vero cells in 1 mL cell culture media (media: Lebovitz L-15
media, 10% fetal bovine serum, 100 U of penicillin/streptomycin, 100 [tg/mL gentimicin,
and 1 [tg/mL amphotericin B (Fungizon)). Plates were kept over-night in a 37 C
incubator and used for virus isolation the following day.
Mosquitoes were processed in a Biosafety Level 3 laboratory (BSL-3 lab). Pools
were homogenized for 1 minute in 1 ml of diluent (Phosphate Buffered Saline (PBS,
contents: 0.8% NaC1, 0.02% KC1, 0.144% Na2HPO4, and 0.024% KH2PO4 in distilled
H20, pH of 7.4) with 4% Fetal Bovine Serum (FBS)) using a laboratory mixer.
Following homogenization tubes were centrifuged at 13,700 rpm for 10 minutes to pellet
mosquito solids. The mosquito supernatant was removed to a new tube and 200 [tl and
10 [tl were removed for use in screening. Any remaining homogenate was frozen at -70
C until needed further.
For the RNA extraction, the 200 [tl of homogenate was then added to a tube
containing 600 [tl of Trizol LS reagent (Life Technologies, Gaithersburg, MD), and the
mixture was incubated for 5 minutes to inactivate the virus. After incubation, tubes were
removed from the BSL-3, stored at -70 C, and later RNA was extracted as described in
the Trizol manufacturer's instructions (Appendix E) and was resuspended in 30 [tl of
nuclease-free water. All RNA samples were then tested for WNV RNA using Promega
Access RT-PCR System (Promega, Madison, WI) with the following concentrations of
reagents: 5 pmol of each primer, 1X kit reaction buffer, 0.2 mM each dNTPs, 2 mM
MgSO4, 1 unit/reaction of both Taq polymerase and Reverse Transcriptase, and 1 [tl of
template for 25 [tl of reaction mix (Table 2-3). The primers used were WN9483 and
WN9794, (Table 2-2) and the thermocycle was run in a PTC-200 (Table 2-3). The RT-
PCR products were visualized on 1% agarose gels with ethidium bromide staining. All
bands were excised from the gel, cleaned, and sequenced as described for the vertebrate
primers (see Appendix A and B). The samples showing positive bands with WN9483
and WN9794 were also confirmed using a second primer set, WN212 and WN1229
Table 2-2. Primers sets used in RT-PCR to test for the presence of WNV RNA.
Amplicon size Annealing
Primer* Sequence (bp) region
Confirmation WN212 TTGTGTTGGCTCTCTTGGCGTTCTT 1071 Capsid protein
Set WN1229 GGGTCAGCACGTTTGTCATTG glycoprotein
Screening WN9483 CACCTACGCCCTAAACACTTTCACC 311 polymerase
Set WN9794 GGAACCTGCTGCCAATCATACCATC polymerase
WNV-specific primer sets
Ten [tl of the mosquito supernatant was mixed with 100 [tl of cell culture media to
be used as an inoculum for the Vero cells. Media was removed from the prepared wells
of the 24 cluster well plates and the inoculum was added. Inoculated plates were
incubated for one hour at 37 C with gentle hand rocking every ten minutes. After
incubation 500 [tl of cell culture media was added (media: Lebovitz L-15 media, 10%
fetal bovine serum, 100 U of penicillin/streptomycin, 100 [tg/mL gentimicin, and 1
[tg/mL amphotericin B (Fungizon)) to each well and plates were placed inside a plastic
box with moistened paper towels and kept in a 37 C incubator. Cells were checked daily
for 7 days for cytopathic effect (CPE) using an inverted compound microscope with
WNV CPE expected to begin on days 3 or 4 post-inoculation.
Samples that showed signs of bacterial contamination were recorded as such, and
the homogenate for that sample was thawed and the inoculation was repeated with a new
well of Vero cells. In these cases, it was assumed that the homogenate was the source of
the contamination and for the new well, the inoculum was removed after the one-hour
Table 2-3. Reagent concentrations and thermocycle conditions used for PCR with
vertebrate-specific primer sets and RT-PCR with WNV-specific primer sets.
Primer set PCR reagents Concentration Thermocvcle
0.2 mM each
93 for 3 min
94 for 30 sec
50 for 30 sec
72 for 1 min 30 sec
Goto 2 45 times
72 for 3 min
93 for 3 min
94 for 30 sec
50 for 30 sec
72 for 1 min 30 sec
Goto 2 34 times
72 for 3 min
94 for 3 min
94 for 30 sec
53 for 30 sec
72 for 30 sec
Goto 2 40 times
10 for ever
93 for 3 min
94 for 30 sec
55 for 30 sec
72 for 1 min 30 sec
Goto 2 34 times
72 for 3 min
48 for 5 min
94 for 5 min
95 for 30 sec
58 for 45 sec
68 for 2 min
Goto 3 39 times
68 for 10 min
incubation period in an attempt to remove the contaminated homogenate after the
inoculum had inoculated.
Assuming one infective mosquito in the pool contained 106.5 virions, this method
would produce an inoculum with about 104.5 virions. This inoculum placed into a well
with 5 x 104 cells would yield a Multiplicity of Infection (MOI) of approximately 0.63.
Three positive controls were conducted simultaneously with samples. Each control
used a previously frozen Florida isolate of West Nile virus, WNV-FL01-JC2-3C2P2,
which had been at a titer of approximately 107.5 TCID5o/mL before freezing. In one
control well, 50 [tl of the virus was added directly. For the other two controls, 2 and 100
[tl of the virus were added to tubes each containing 38 Ae. aegypti colony mosquitoes,
and these controls were then processed in the same manner as the field samples. For
these two controls the inoculation contained approximately 102.5 and 104.5 virions
respectively. This gave an MOI of about 0.0063 and 0.63 respectively. For the direct
inoculation of 50 il, the MOI was about 32.
During collecting no mosquitoes were seen resting inside of the alligator pens or on
other buildings. Mosquitoes were observed in the brush and woods along the western
margin of the farm. These swarmed if one entered the woods but did not attempt to feed
on collectors in the open. In several instances, mosquitoes, Mansonia sp., pursued and
bit the investigator in the open during daylight hours (1:00 to 3:00 PM). Inspection of the
pens revealed that while they were mostly closed, there were cracks and spaces around
the doors and pipes that would be sufficient for mosquitoes to enter and exit.
Collection bags on several of the CDC traps that were hung inside alligator pens
were apparently torn down by the alligators some time during the night. These samples
could not be recovered. Taking into account the losses due to alligator interference and
one disturbed collection bag there was a total of 20 trap nights for the CDC light traps
and 24 trap nights for the resting boxes over the four collecting trips.
A total of 4484 unfed and 37 blood fed mosquitoes was collected from CDC traps,
resting boxes, and aspiration. There were 16 species (10 genera) represented in the
collection. The species were An. quadrimaculatus, An. crucians, Mansonia dyari, Ma.
titillans, Cx.. nigripalpus, Cx. erraticus, Cx. quinquefasciatus, Uranotaenia sapphirina,
Ur. lowii, Psorophora columbiae, Ps. ferox, Coquillettidiaperturbans, Wyeomyia
vanduzeei, Culiseta. melanura, Ae. albopictus, and Oc. infirmatus (Fig 3-1). The
numbers of mosquitoes collected varied from one trip to the next.
Wyeomyia vanduzeei 1
Psorophora ferox 1
Culex quinquefasciatus 2
Culiseta melanura 2
Uranotaenia lowii 8
Aedes albopictus 10
Coquillettidia perturbans 20
Ochlerotatus infirmatus 23
Psorophora columbiae 115
Uranotaenia sapphirina L 142
Mansonia titillans  272
Culex erraticus  273
Culex nigripalpus  535
Anopheles crucians  737
Mansonia dyari  1170
Anopheles quadrimaculatus  1172
0 200 400 600 800 1000 1200 1400
Figure 3-1. Total mosquito numbers collected over four trips to Farm A. Numbers in [ ]
indicate engorged mosquitoes.
Five different species were captured in resting boxes and overall the percent of
blood-fed individuals was greater in resting boxes than in CDC light trap collections
(31.5% versus 0.4%). In the resting boxes there were An. quadrimaculatus (37 total, 15
blood-fed), An. crucians (6, 1), Cx. erraticus (4, 2), Cx. nigripalpus (9), and Cx.
quinquefasctiatus (1). One blood-fed Cx. nigripalpus was captured with vacuum
Seven different species were collected from CDC traps that were located inside of
alligator pens. These seven species were An. quadrimaculatus, An. crucians, Ma. dyari,
Ma. titillans, Cx. nigripalpus, Cx. erraticus, and Cq. perturbans. Based on average
numbers of the different species in traps located inside and outside of the pens, it
appeared that some species more readily entered pens than others (Fig. 3-2).
0 outside pens
400 inside pens
Figure 3-2. Portions of each mosquito species captured in CDC light traps set outside of
alligator pens versus inside of pens (for collecting trips 1,2, and 4).
Trip one. A total of 2665 mosquitoes was collected from CDC traps (Table 3-1)
during trip one, and this represented more than half of the total collected during the four
Trip two. A total of 150 mosquitoes was collected during this trip (Table 3-2). Of
the four CDC light traps inside of alligator pens, one was damaged by the alligators. The
collection bag was removed from the trap and pushed into the water of the alligator pen,
thus no mosquitoes were obtained from that CDC trap.
Trip three. A total of 628 mosquitoes was collected (Table 3-3). The pen or
position were samples were collected can not be stated with certainty because the records
of trap placement were lost while in transit between Farm A and Gainesville.
Table 3-1. Mosquitoes captured from CDC light traps during Trip one at Farm A.
Numbers in [ ] indicate the number blood-fed mosquitoes.
CDC inside pen CDC outside pen TOTAL
trap 1 trap 2 trap 1 trap2
An. quadrimaculatus 343 3 184 251 781
An. crucians 0 0 179 364 543
Ma. dyari 46 6 323 338 338
Ma. titillans 63 0 4 23 27
Ps. columbiae 0 0 39 62 101
Ur. sapphirina 0 0 28 39 67
Ur. lowii 0 0 3 1 4
Ae. alobopictus 0 0 0 6 6
Cx. erraticus 7 0 56 84 147
Cx. nigripalpus 19  54 122 19
Cq. perturbans 2 0 5 1 8
Ps. ferox 0 0 0 1 1
Oc. infirmatus 0 0 0 8 8
Cs. melanura 0 0 0 0 0
Cx. quinquefasciatus 0 0 0 0 0
Wy. vanduzeei 0 0 0 0 0
TOTAL 480 10 875 1300 2665
Table 3-2. Mosquitoes collected in resting boxes and CDC light traps during the second
collecting trip to Farm A. Numbers in [ ] indicate blood-fed mosquitoes.
CDC inside pen
trap 1 trap 2 trap 3 Resting box TOTAL
An. quadrimaculatus 0 9 25 3 37
An. crucians 0 0 8 3 8
Ma. dyari 2 7 22 0 31
Ma. titillans 4  40 0 44
Ps. columbiae 0 0 0 0 0
Ur. sapphirina 0 0 0 0 0
Ur. lowii 0 0 0 0 0
Ae. alobopictus 0 0 0 0 0
Cx. erraticus 4  0  4
Cx. nigripalpus 12 3 3 0 18
Cq. perturbans 0 0 0 0 0
Ps. ferox 0 0 0 0 0
Oc. infirmatus 0 0 0 0 0
Cs. melanura 0 0 0 0 0
Cx. quinquefasciatus 0 0 0 0 0
Wy. vanduzeei 0 0 0 0 0
TOTAL 22 22 98 8 150
Table 3-3. Mosquitoes collected from CDC light traps and resting boxes during the third
collecting trip to Farm A. Numbers in [ ] indicate blood fed individuals.
trap Resting TOTAL
trap 1 trap2 trap3 trap4 trap5 trap6 box
An. quadrimaculatus 7 9 36 6 7 109  183
An. crucians 2 9 30 0 0 0 0 41
Ma. dyari 4 0 58 3 4 39 0 108
Ma. titillans 0 5 3 2 12 46 0 68
Ps. columbiae 0 2 3 0 0 0 0 5
Ur. sapphirina 5 9 35 0 0 0 0 49
Ur. lowii 0 0 2 0 0 0 0 2
Ae. alobopictus 0 0 1 0 0 0 0 1
Cx. erraticus 9 10 50 0 0 1 0 70
Cx. nigripalpus 20 19 40 0 0 5 8 92
Cq. perturbans 1 0 0 0 0 1 0 2
Ps. ferox 0 0 0 0 0 0 0 0
Oc. infirmatus 0 0 7 0 0 0 0 7
Cs. melanura 0 0 0 0 0 0 0 0
Cx. quinquefasciatus 0 0 0 0 0 0 0 0
Wy. vanduzeei 0 0 0 0 0 0 0 0
TOTAL 48 63 265 11 23 201 17 628
Trip four. On the last collecting trip 1041 mosquitoes were collected (Table 3-4).
The battery failed on one of the CDC traps although some mosquitoes were still
Numbers and proportions of different mosquito species varied from one collecting
trip to the next, however, statistical analysis of these differences was not done as the
sampling size and system did not allow it.
Blood Meal Identification
The Crocodilian-specific primers produced a PCR product band of the correct size
for six of the 37 blood-fed mosquito samples (two Cx. erraticus and fourMa. dyari). Of
these six positives, one was a mosquito from a resting box and the others were from CDC
traps. There were also DNA bands of the wrong size (about 180 bp) for two
Table 3-4. Mosquitoes captured in CDC light traps and resting boxes on the fourth
collecting trip to Farm A. Numbers in [ ] indicate blood-fed individuals.
CDC inside CDC outside
pen pen Resting
1 trap 2 trap 3 trap
3 18 0
1 0 0
0 2 2
22 10 2
0 0 0
0 0 0
0 0 0
0 0 0
1 0 0
0 0 0
1 0 0
0 0 0
0 0 0
0 0 0
0 0 0
0 0 0
28 30 4
1 trap 2 trap 3 trap
20 51 51
28 54 41
152 102 30
12 1 12
1 1 6
3 16 3
0 1 0
1 0 1
9 29 6
43 148 27
7 2 0
0 0 0
6 0 0
1 0 1
1 0 0
0 0 1
284 405 179
Cx. nigripalpus and one Ma. titillans (the Ma. titillans had a second, very faint band of
approximately the correct size). Sequences from the correct-sized bands matched that of
the positive control (193 out of 200, or 96.5% homology). The other bands produced
sequences that matched neither the positive control nor any entry on the GenBank
Of the 37 mosquitoes that had apparent blood meals, 14 reacted with one of the
mitochondrial primer sets (mammal, alligator, bird) giving an identification rate of
37.8%. Seven individuals (representing three species) were positive for alligator DNA
(Fig. 3-3), six (three species) were positive for mammalian DNA, and one individual was
positive for avian DNA (Fig. 3-4). For the alligator primer set, the seven positives
included the six samples that were positive for the Crocodilian primer set and the seventh
was the Ma. titillans for which the Crocodilian primers had produced two bands, the
fainter of which was the appropriate size for a positive.
Sequencing confirmed that all seven of the alligator-positive PCR bands were from
Alligator mississippiensis. The mosquitoes feeding on alligators were Cx. erraticus (two
individuals), Ma. dyari (four individuals), and Ma. titillans (one individual). All of these
individuals except for one Cx. erraticus were obtained from CDC traps that were inside
of alligator pens. The exception was from a resting box (Table 3-5).
Sequencing allowed species identification of the mammalian and avian blood
meals. The single avian positive was from a Cx. nigripalpus that had fed on a turkey,
(Meleagris gallopavo). For the mammalian positives, one Cx. nigripalpus fed on a
raccoon (Procyon lotor), two An. quadrimaculatus fed on pigs (Sus scrofa), one Ma.
dyari fed on a pig, and another Ma. dyari fed on a horse (Equus callabus). The blood
meal of one An. quadrimaculatus could not be identified further (Table 3-5). The
GenBank E value for all of the matches was 0.0 except for the match with the raccoon,
where the E value was 5e-175 (indicating that there is zero or almost zero probability that
these matches were due to chance). Sequences are in Appendix E.
The mammalian-specific primers consistently amplified mosquito DNA in the
negative control (an un-engorged mosquito). Repetition of the DNA extraction from a
new un-engorged mosquito reduced the possibility of contamination of the negative
control with mammal DNA. The sequence of the brightest DNA PCR fragment did not
match closely with any of the GenBank entries, however one 64 bp portion of the region
1 2 3 4 5 6 7 8 9 10 11 1213 1415
1 2 3 4 5 6 7 8 9 10 11 12 13 1415
S2 3 4 5 6 7 8 9 10 11 1213 1415
Figure 3-3. Products from PCR amplification of mosquito samples with alligator-specific
primers. Gel a: lane 1 contained a 1 kb ladder, lanes 3,4, and 8 contained Ma.
dyari, lanes 5 and 6 contained Cx. erraticus, and lane 15 contained the
alligator positive control. Gel b: lane 1 contained a 1 kb ladder, lane 6
contained Ma. dyari, and lane 10 contained Ma. titillans.
(out of the 422 bp sequence) did match closely with chromosomal DNA (from partial
mRNA) from An. gambiae. Within this 64 bp portion there were 57 bases shared with
the database's An. gambiae sequence and the E-value assigned for this match was 3e-10
1 2 3 4
5 6 7 8 9 10
1. 100 bp ladder
2. Avian negative control
3. Cx. nigripalpus
4. Avian positive
5. Mammal negative control
6. Ma. dyari
7. Cx. nigripalpus
8. An. quadrimaculatus
9. Mammal positive control
10. 1 kb ladder
Figure 3-4. Products from PCR amplifications with a mammalian-specific primer set
(lanes 2-5) and an avian-specific primer set (lanes 6-8).
Sequencing results from the correct-sized bands often had multiple overlapping
peaks, suggesting that the mammal primers may sometimes bind and amplify a portion of
the chromosomal DNA of mosquitoes. When one of these bands was cloned into
bacteria, only some of the clones were of fragments whose sequences matched with
vertebrates. The other clones yielded sequences for which there were no close matches
on the database.
Table 3-5. Identities of vertebrate hosts as determined by sequencing the PCR product,
and information about collection
date and location on farm of the mosquito
1MBF4 Equus callabus
1MBF1(2) Sus scrofa (wild
References for identification of sequences:
1994); b = AY237534 (Alves et al., 2003);:
(Kornegay et al., 1993);e
In trees on
Sept. 12 In trees on
Sept. 26 box
: GenBank accession # D32190 (Chikuni,
U12853 (Lento et al., 1995); d = L08381
AF318572 (Glen et al., 2002), ***
specific host not
Isolation of virus in cell cultures. In cell culture two of the three WNV positive
control wells showed obvious CPE before day seven. For the direct inoculation (MOI =
32), strong CPE was apparent on day two. For the positive control with MOI = 0.63,
small foci of infection were observed on day three and by day five infected cells were
apparent through out the well. In these two positive controls, all cells appeared infected
by day seven and many had detached from the substrate and were floating in the media.
No CPE was apparent in the well that received the inoculum with MOI = 0.0063.
No apparent viral CPE was observed in any of the wells containing homogenate
from field-collected mosquitoes. This and the absence of positive bands from RT-PCR
indicated that there were no WNV positive field-collected mosquito pools, giving an MIR
Two of the wells had bacterial contamination that became apparent after two days.
When these samples were repeated (as described above) one had bacterial contamination
again, and the other did not.
The cells in many of the wells that had been inoculated with homogenate from a
pool of 50 Ma. dyari showed some effects that were believed to be due to non-viral
components of the mosquitoes. The cells did not grow as well in the center of the well,
and many had a "lacey" appearance, i.e. the cells appeared to have more vacuoles and
margins of the cells became less smooth. However this condition was not progressive,
and there were still many healthy cells in the well at day seven. The addition of 100 lil of
cysteine to the diluent prior to homogenization appeared to prevent this effect in
subsequent pools of 50 Ma. dyari.
Detection of viral RNA. Both of the positive controls that were tested for WNV
RNA showed clear, bright positive bands with both the screening primers (about 300 bp)
and the confirmation primers (about 1000 bp) (Fig 3-5). The results from the positive
1 2 3 4 5 6 7 8 9 10 11 12 13 (lane#)
Figure 3-5. Products from RT-PCR with WNV screening primer set (lanes 2-7) and
WNV confirmation set (lanes 8-13). WNV positive controls are in lanes 2 and
8. Lane 1 contains a 1 kb molecular weight standard ladder. Lanes 3 and 10
were no-template negative controls. All other lanes contain samples (4C7,
4G7, 4A3, and 4L1) for which there was some amplification in the original
screening, but not in any subsequent RT-PCR reactions.
controls demonstrated that the cell culture screening was sensitive enough to detect
mosquitoes containing a "normal" titer of WNV (Johnson et al., 2003), but may have
missed mosquitoes with the minimum titer of 105. However RT-PCR would have
detected infected mosquitoes with more sensitivity, including those with titers well below
105. No field-collected mosquito pools were positive for WNV RNA.
Blood Meal Identification
While encouraging, the results from the Crocodilian-specific primers (Yau et al.
2002), did not seem conclusive, because the primers were not specific. Because there are
mitochondrial sequences for many more different organisms published and available on
the GenBank database, it is easier to identify an unknown mitochondrial sequence than an
unknown chromosomal sequence. The database contains chromosomal DNA entries for
fewer reptiles and nothing was known about the region of DNA that the Crocodilian-
specific primers amplified, therefore it was difficult to determine whether or not the
homology found between the mosquito samples and the alligator positive control
constituted a real match. It was these unknowns that led to the selection of the second
primer set, one designed to amplify alligator mitochrondrial DNA from a well-studied
region, the cytochrome b gene.
It appeared that the mammalian-specific primers sometimes amplified
chromosomal DNA from the mosquito. The bands in the negative control (where only
mosquito DNA was present) shared some homology with a mosquito and even where
there was vertebrate host DNA present, sequencing results (overlapping peaks and cloned
fragments whose sequences were not from a known region of vertebrate DNA) showed
that the primers annealed in multiple places apparently on both host and mosquito DNA.
Of the three mosquito species found to feed on alligators, only one, Cx. erraticus,
has been reported to feed on reptiles and in general has been identified as an
opportunistic feeder taking meals from mammals, birds, and reptiles or amphibians
(Robertson et al., 1993; Irby and Apperson, 1988). Species in the genus Mansonia are in
general considered mammal and some times bird feeders (Edman, 1971). Studies of this
genus in other parts of the world have found them attracted to or feeding on cow and
human "baits" (Tuno et al., 2003; Khan et al., 1997). The normal feeding habits ofMa.
titillans and Ma. dyari (both of the neotropical subgenus Mansonia) have not received
much attention probably because they do not seem to be implicated in transmission of
pathogens in the United States. While Mansonia do not appear to be reptile feeders, this
would not be the first instance of feeding "patterns" being strongly influenced by
availability of hosts. Edman (1971) found that the number of mosquitoes with squirrel
blood meals increased dramatically on a night when caged squirrels happened to be
placed near the collection site. On Farm A, thousands of alligators are captive in pens
with water depths insufficient to allow the alligators to submerge. They may present a
blood source so readily available that a range of species takes advantage.
The mitochondrial PCR product from the alligator positive control was distinctly
fainter than the bands from the field collected mosquitoes. This may have been due to
the presence of heparin in the positive control alligator blood. Yokota et al. (1999) found
that heparin interfered with PCR when template DNA was from heparinized blood, and
the degree of interference was related to the concentration of heparin and the type of
polymerase enzyme used.
The total blood meal host identification percentage (38%) was lower than that of
other studies (65%) where host determination was done using DNA probes and a PCR
reaction (Ngo and Kramer, 2003; Leslie Rios, personal communication). This may be
due to components within the mosquito or processes during digestion that interfere with
the PCR or rapidly degrade the DNA. Cupp et al. (2004) found that their overall
identification percentage (for two Culex species) was 65%, but was much lower for Ur.
sapphirina with only two individuals out of the 35 (about 6%) blood fed collected
yielding a result. While Uranotaenia are quite small mosquitoes, it seems unlikely that
the size of the blood meal alone would be responsible for the dramatically smaller
identification rate, especially considering that smaller blood meals (incomplete
engorgements in "normal" sized mosquito species) were successfully amplified in this
study and that in other studies there was no negative correlation between blood meal size
and success of amplification (Mukabana et al., 2002). In a study working only with An.
gambiae, Gokool et al. (1993) had a 31% positive identification rate. For this study,
when Anopheline and Culicine mosquitoes are considered separately, the identification
rates are 18.8% and 52% respectively. The idea that differences in mosquito digestive
physiology might influence the success rate of PCR-based host identification studies
warrants further study. After a blood meal is ingested it clumps inside the mosquito
midgut yielding separated serum and a clot containing the erythrocytes. After that (and
for the next several hours) enzymes are secreted which begin to digest the surface of the
clot. Components of the separate serum are absorbed and used for nutrition or egg
development (Nayar and Sauerman, 1977). Nayar and Sauerman (1977) showed that in
An. quadrimaculatus, the mean clotting time was significantly greater than that of five
Culicine mosquitoes. The average clotting time (based on results from five different
blood hosts) was 203 minutes for An. quadrimaculatus, compared with 45, 40, 31, 21,
and about 8 minutes for Ae. taeniorhynchus, Oc.. sollicitans, Ae. aegypti, Ps. columbiae,
and Cx. nigripalpus, respectively. The delay in clotting (likely due to differences in
salivary anticoagulants) may allow enzymes to more readily reach and digest the
erythrocytes in an Anopheles blood meal. In addition, An. quadrimaculatus blood is
sometimes excreted within a few hours of feeding (Nayar and Sauerman, 1977). The net
effect of these differences may result in host DNA being degraded more quickly in
Anopheles than in some other mosquito genera.
In this study mosquito collections were gathered and placed on ice in the early
afternoon. Assuming mosquitoes were active and thus captured at dusk (Bidlingmayer,
1967), then many of the blood fed individuals would have been placed on ice about 18 h
after they had taken a blood meal (estimated dusk at 9:00 PM). In other studies (Cupp et
al., 2004; Ngo and Kramer, 2003) mosquitoes were collected at dawn, or about 10 h after
taking a dusk blood meal. The additional eight h of time may have allowed greater
breakdown of DNA, thus making the positive identification percentage lower in this
Since there were no virus isolations and no WNV RNA detected in mosquito pools,
the MIR was 0 in 4447 or 0 in 270, 268, and 1161 for Cx. erraticus, Ma. titillans, and
Ma. dyari (the three species that fed on alligators). The virus isolation results neither
support nor diminish the possibility of mosquito-transmitted WNV on the alligator farm,
nor can they help in incriminating any one of the three alligator-feeding species found. In
a study done in New York, WNV isolations tended to occur in the vicinity of greater
transmission such that the authors made the following generalization: the greatest
number of human cases and dead crows corresponded to a mosquito MIR of 5.27/1000, a
few human cases and moderate number dead crows corresponded to an MIR of 0.18 to
2.36/1000, and no human cases and few dead crows corresponded to an MIR of 0 to
0.86/1000 (White et al., 2001). Some studies have had similar MIR's (Rutledge et al.,
2003; Reisen et al., 2004), and others have had much lower MIR's, even when there has
been evidence of WNV transmission, such as dead birds (Meece et al., 2003; Andreadis
et al., 2001). The MIR for collections made in Ohio by Mans et al. (2004) was higher (8
out of 1000), but they tested only those mosquitoes collected from gravid traps, thus
biasing the results towards a higher MIR by testing only older females. In a Florida
study, an MIR of 1.2 in 1000 was found, and results also indicated that viral activity was
very focal (Rutledge et al., 2003). In this study, almost 12,000 mosquitoes were collected
and two species, Cx. nigripalpus and Cx. quinquefasciatus made up about 78% of the
collection. Fourteen pools from these two species were positive for WNV, and a single
Cx. nigripalpus was responsible for infecting a sentinel chicken. This species was
present around and feeding on the host (chicken) and was most frequently infected with
WNV, showing that these criteria can be helpful in identifying possible vectors. This
study also found that the number of WNV positive mosquitoes around the chickens was
greater than the number of transmissions to chickens. Thus an infected mosquito pool is
not a sure way to identify the species responsible for transmission. Alternatively, the
study found that there were no infected mosquitoes at a site where a horse had become
infected a month prior to collecting. In this case, the mosquito infection rates
underestimated transmission rates, probably because collecting was started after the
transmission had taken place.
Many studies also found Culex mosquitoes to be the most frequently infected with
WNV, so the MIR of these species may be more meaningful for comparisons to the
situation at the alligator farm. On the alligator farm the mosquito collections were only
about 18% Culex (808 out of 4484). The collection was predominately (75%) Anopheles
and Mansonia, genera from which WNV is much more rarely isolated. So although there
were almost 4500 mosquitoes collected, an "average" MIR of 1 in 1000 could not be
expected as many of the collections that this MIR is based on were dominated by Culex
Based on the above information, it appears that the current study may have
"missed" any WNV positive mosquitoes on the alligator farm because: 1) the mosquito
collection was not large enough, 2) the mosquito collections were predominately non-
Culex mosquitoes which are less likely to vector and be infected with WNV or 3) the
collecting began during the epidemic of disease and after transmission had occurred.
With the last explanation, it is possible that the transmission of WNV had taken place
about two weeks before the majority of the alligators began to die. Work by Klenk et al.
(2004) showed that alligators developed viremia about 5 days after infection and that they
in turn infected their tank mates about a week after that viremia developed. In one
possible scenario, mosquitoes infected several alligators during a brief period of intense
WNV activity. These alligators then developed viremia and infected their tank mates,
and a week later the situation progressed to what was observed during the first collecting
trip: multiple alligators sick and dying from WNV-like disease. In this scenario the
WNV transmission was very focal (both temporally and spatially) and had subsided by
the time mosquito collecting had begun. If this were the case, the practice of pre-
epizootic surveillance would not only help predict when an epizootic might start, but
would also be important for identifying the mosquito species involved.
Surveillance reports were used to fill in information about the state of transmission
in the area around the time of the outbreaks. If the alligator epidemics were isolated, it
may suggest a cause separate from the surrounding virus activity, (i.e. infection due to
WNV contaminated meat). However, if there was transmission, as demonstrated by
infected horses, humans, birds, or sentinel chickens, this supports the idea that the
outbreaks on the alligator farms were related to the virus activity occurring in the area.
The reports posted by the Florida Mosquito Control Association and the United States
Geological Service's maps (http://westnilemaps.usgs.gov/index.html, created with
information from CDC) give information about the level of WNV activity in the vicinity
of the alligator farm during the fall of 2003. During that year, the county had 64
conversions in sentinel chicken flocks, one osprey (Pandion haliaetus Linnaeus) positive
for antibodies to WNV, and three cases of WNV reported by veterinarians. However,
there were no isolations of virus from mosquito pools in the county that year. This
surveillance information indicates that there was mosquito transmission of WNV in the
county during the time of the epizootic on the alligator farm, even though no isolations
were made from mosquitoes. The possibility remains that the WNV on Farm A was
mosquito transmitted even though no positive mosquitoes were detected.
The information gained in this study can be considered in the context of the criteria
established by Reeves (1954) and expanded on by Kilpatrick et al. (2005) and used to
identify potential mosquito vectors. Because this study did not identify WNV in any
mosquito pool or identify any competent vectors for WNV, information from other
studies can be incorporated to identify potential vectors. In this study it was established
that Cx. erraticus, Ma. titillans, and Ma. dyari feed on alligators at the farm and that
these species are relatively numerous around the farm (6%, 6%, and 26% of the total
catch, respectively). Other studies have shown that they are in greatest abundance during
the season when the alligator epidemics occur (Bidlingmayer, 1968; Zhong et al., 2003).
Information from other studies and surveillance reports will be necessary to answer the
remaining questions about these potential vectors: are they competent vectors for WNV,
and are they repeatedly found infected with the virus?
Culex erraticus is a member of the neotropical subgenus Melanoconion, and is
found all over the eastern United States, as far north as Connecticut and New York
(Andreadis, 2003; Kulasekera et al., 2001), south of the great lakes, through out the
southeast (Darsie and Ward, 1981), and has been found in California (Lorthrop et al.,
1995). Culex erraticus specimens have been found positive for West Nile virus each year
from 2002 to 2004 (CDC,
http://www.cdc.gov/ncidod/dvbid/westnile/mosquitoSpecies.htm). They have also been
found infected with other arboviruses in the United States such as Eastern Equine
Encephalomyelitis virus (EEE, Togaviridae: Alphavirus) (Wozniak et al., 2001; Cupp et
al., 2003) and St. Louis Encelphalitis virus (Cupp et al., 2004a). St. Louis Encephalitis
(SLE) is also a flavivirus of the Japanese encephalitis (JE) serogroup (Poidinger et al.
1996). Culex erraticus are considered competent vectors of EEE (Cupp et al., 2004b).
However, competence for one type of arbovirus, often does not correlate with
competence for another (Hardy et al., 1983). Some reports have given a WNV minimum
infection rate for this species (Gaines, Virginia Department of Health), although in many
cases this species was pooled and/or reported together with other Culex species under the
general heading "Culex sp.". This makes it difficult to know the minimum infection rate
although it is possible to say that WNV has been repeatedly isolated from these
mosquitoes and that other members of the genus are the most commonly found WNV-
positive mosquitoes. As of July 2005, no WNV vector competence studies have been
published for Cx. erraticus or for any other North American Melanoconion. The studies
that have been done indicate that all of the Culex species tested have moderate to
excellent vector competence and moderate to excellent potential to vector WNV (Turell
et al., 2005). Based on laboratory experience, Cx. erraticus appears to be a long-lived
species (Klein et al., 1987), and this could contribute to its potential vector competence.
In the United States, Ma. dyari is found in Florida and parts of Georgia and South
Carolina (Darsie and Ward, 1981; Darsie and Hager, 1993). Mansonia titillans has been
found in central and south Florida, in southern Texas, and in Mississippi (Darsie and
Ward, 1981; Goddard and Harrison, 2005). These species are also found in south and
Central America, where Ma. titillans is likely involved in the transmission of Venezuelan
Equine Encephalomyelitis virus, an alphavirus (Mendez et al., 2001; Turell et al., 2000)
and Ma. dyari is a maintenance vector of SLE (Gorgas Memorial Laboratory 1979, as
cited in Lounibos et al., 1990). As with Cx. erraticus, no vector competence studies with
WNV have been done for Mansonia species. WNV has been detected in pools of several
species of Mansonia in Africa (Traore-Lamizana et al., 2001), and other members of the
genus appear to be involved in transmission of JE in Asia (Arunachalam et al., 2002;
Arunachalam et al., 2004). Regardless of their presumed vector competence based on
these other diseases, Ma. dyari has never been positive for WNV in the United States and
WNV has been detected in Ma. titillans only in 2004
(http://www.cdc.gov/ncidod/dvbid/westnile/mosquitoSpecies.htm). Based on this input,
Cx. erraticus seems the most likely potential vector of WNV on Farm A, although
Mansonia are at least nuisance species.
Culex erraticus, Ma. dyari, and Ma. titillans are all associated with vegetated
aquatic habitats (Alfonzo et al., 2005; Lounibos et al., 1990). Culex erraticus females
prefer to oviposit where there are aquatic plants (Klein et al., 1987) and all Mansonia
larvae are associated with aquatic plants from which they derive oxygen and possibly
cover from predation (Lounibos et al., 1990). Conceivably the numbers of these
mosquitoes could be controlled on Farm A by reducing the amount of aquatic vegetation
present in the bodies of water. Especially with the Mansonia, mosquito populations are
closely related to the availability of the preferred larval host plants (Lounibos and Esher,
1985), which are water lettuce, Pistia sp. for Ma. dyari, and common water hyacinth,
Eichhornia crassipes (Mart.), forMa. titillans (Slaff and Haefner, 1985). These plants
can be controlled with herbicides (Slaff and Haefner, 1985). The practicality of serious
water plant control in this case would need to be investigated. First, both Ma. titillans
and Cx. erraticus have been reported as traveling greater than 2 km in mark and recapture
studies (Morris et al., 1991), so control may have to include all vegetated water bodies
within a 1 to 2 km radius. Second, farmers would need to consider the risks associated
with managing vegetation in water bodies that are occupied by a number of large
alligators, especially because these alligators are accustomed to receiving food from
humans. Alternatively, control could be aimed at adult mosquitoes. For all three
species, adulticides could be applied for several months in the late summer and early fall
when populations are at their peak (Slaff and Haefner, 1985; Bidlingmayer, 1968;
Roberson et al., 1993; Zhong et al., 2003).
Alternative Vertebrate Reservoirs
The other blood meal hosts (pigs, a horse, a turkey, and a raccoon) identified on the
farm were probably not involved in maintenance or amplification of WNV. In a study
where three-week-old turkeys were inoculated subcutaneously with NY99 WNV, none
displayed illness, and viremia, while detectable, was very low. From the results
researchers concluded that turkeys would not be severely effected by WNV nor would be
they important amplification hosts (Swayne et al., 2000). In another study, pigs were
subjected to mosquitoes infected with New York 99 strain of West Nile virus, and while
adult pigs seroconverted, most of the animals did not have sufficient viremia to allow
reisolation of the virus from serum. Weanling pigs developed viremia less than or equal
to 1031 PFU/mL. No signs of clinical disease were observed (Teehee et al., 2005). The
low viremia found in horses and their failure to infect mosquitoes in experiments also
makes them unlikely amplification hosts (Bunning et al., 2002).
CONCLUSIONS AND AREAS FOR FURTHER STUDY
In conclusion, the study found that of the 16 species collected in CDC light traps
and resting boxes on Farm A, three species contained blood from alligators: Cx.
erraticus, Ma. dyari, and Ma. titillans. Based on its known feeding habits Cx. erraticus
would also feed on birds near the farm (Roberson et al., 1993). If Cx. erraticus has
vector competence similar to what has been found for many of the other members of its
genus (Turrel et al., 2005), then it could serve as a vector of WNV to alligators on Farm
A. Additional laboratory and fieldwork, such as vector competence studies and efforts to
screen for WNV, can further clarify the potential role of this species in WNV
transmission on alligator farms. In addition, this study found Mansonia mosquitoes
feeding on alligators, and this appears to be the first report of these two species of
mosquito feeding on reptiles.
It may also be interesting and informative to study mosquitoes' responses to
potentially attractive or repellent compounds associated with the alligators. In one trap
placed inside of an alligator pen there were over 400 mosquitoes collected in one 24 h
period, suggesting that the mosquitoes were attracted to compounds coming from the pen.
A 1 kg alligator at rest should excrete about 7 mL of CO2 per min (Farmer and Carrier,
2000). An alligator pen containing 200 such individuals could be putting out CO2 at a
rate of about 1400 mL/min, thus representing a very strong attractant for many mosquito
species (Kline and Mann, 1998). However, not all of the species that were found inside
of the alligator pens had blood meals from alligators. This may mean that there are
repellents or missing attractants (or other stimuli) that prevent feeding in many of the
mosquitoes that were initially attracted to the alligator pens. Additional collections that
are carried out more regularly and systematically may provide more information about
what mosquitoes are attracted to the alligators, and whether or not these mosquitoes
proceed to feed. Traps that do not have a CO2 bait could be set in the alligator pens to
single out species of mosquito attracted to the alligators in the absence of additional
attractants. Also laboratory experiments with an olfactometer (McKenzie, 2003) could
be used to determine the attractiveness of different aromatic compounds present in
alligator hide to mosquitoes, thus further adding to our understanding of how mosquitoes
respond to alligators as a potential blood host. The collecting results suggested that some
species are more inclined to enter alligator pens than others. Investigating these
differences could not only help predict vectors of WNV in alligators, but could also be
useful in the continued effort to describe mosquito host seeking behavior.
PROTOCOL FOR QIAGEN QIAQUICK SPIN KIT, PURIFICATION OF DNA FROM
(modified from QIAquick Spin Handbook, S. C. Garrett)
1. Add 96-100% ethanol to buffer PE before beginning.
2. Weigh the excised piece of agarose gel containing the PCR product and place in a 1.5
mL microfuge tube.
3. Add 3 [tl of buffer QG for each 1 mg of gel.
4. Incubate at 500C for 10 min (tapping tube to mix every 2 minutes) to dissolve gel.
5. Once the gel is dissolved add 1 [tl of isopropanol for each 1 mg of gel and mix.
6. Place a QIAquick spin column into a 2 ml plastic collection tube.
7. Transfer the dissolved gel solution to the column and centrifuge at 13,000 rpm for one
min in a microcentrifuge.
8. Remove the column, discard the flow-through from the collection tube, and place the
column back into the tube.
9. Add 0.5 ml of buffer QG to the column and centrifuge for one min (13,000 rpm).
10. Discard the flow-through and return column to tube.
11. Add 0.75 ml of buffer PE and centrifuge for one min. (13,000 rpm).
12. Discard flow-through, return column to tube, and centrifuge for an additional
minute at the same speed.
13. Transfer the column to a clean, labeled microcentrifuge tube.
14. Add 30 [tl of buffer EB to the center of the column membrane (white material in the
center of the column), allow the buffer to soak in for one min, and then centrifuge for
1 min (13,000 rpm).
15. Eluted DNA can be stored at 40C until needed for sequencing or other purposes.
ABI PRISMTM DYE TERMINATOR CYCLE SEQUENCING KIT, PROTOCOL FOR
(Modified from PERKIN ELMER PROTOCOL, revised July, 2005)
NOTE: Keep all reagents on ice
1. Estimate concentration of template DNA by running 5 til in an agarose gel
and comparing intensity to known concentration of molecular weight
2. Calibrate the thermocycler.
3. Dilute templates to recommended concentration (See table below).
4. Remove Terminator Ready Reaction Mix and ICBR dNTP mix from
freezer and thaw on ice.
5. For each reaction, mix the following reagents in a microfuge
Terminator Ready Reaction Mix 2.0 [tL
ICBR dNTP mix 2.0 [tL
single-stranded DNA (100ng/ul) 50-100 ng
double-stranded DNA (500 ng/ul) 200-500 ng
PCR products (100-200 bp) 1-3 ng
(200-500 bp) 3-10 ng
(500-1000 bp) 5-20 ng
(1000-2000 bp) 10-40 ng
(> 2000 bp) 40-100 ng
Primer (3.5 pmol) tL
Deionized Water Bring final volume to 10.0 gL
Final Reaction Volume 10.0 gL
6. Gently pipette to mix the reagents.
7. Place tubes in thermocycler and start thermocycle (See below).
1. Ramp to 960C and hold for 30 s (denaturation)
Ramp to 500C and hold for 15 s (primer annealing)
Ramp to 600C and hold for 4 min (product extension)
2. Repeat step 1. For 25 cycles
3. Ramp to 40C and hold.
8. Remove tubes from the thermocycler.
9. For each reaction, prepare a 1.5 mL microfuge tube by adding:
1.0 ptL 3M Sodium acteate, pH 4.6
30.0 tL 95% cold ethanol
10. Transfer the sample to the prepared microfuge tube and place on ice for
11. Centrifuge for 15 min (13,000 rpm).
12. Carefully and completely remove ethanol solution, without disturbing
the pellet of DNA.
13. Rinse the pellet with 250 tL of 70% ethanol.
14. Centrifuge for 1 min to secure the pellet onto the bottom of the tube.
15. Carefully remove the 70% ethanol without disturbing the pellet of
16. Dry under vacuum.
17. Store in dark freezer until ready to read.
PROTOCOL FOR PGEM-T VECTOR LIGATION KIT,
(modified from the Promega pGEM-T and pGEM-T Easy Vector Systems Technical
Manual, S. C. Garrett, July 2005)
1. Estimate the concentration (ng/[tl) of the PCR product to be cloned by comparing the
intensity of the band in a gel to the intensity of the standardized bands of the molecular
weight ladders with known DNA concentrations.
2. Calculate the amount (in ng) of PCR product needed for the amount of pGEM-T vector
by using the following equation:
(ng of vector)(kb size of insert) = x ng of PCR product
3. Based on the above calculations/estimations, calculate the volume of PCR product that
should be added to the vector.
4. Centrifuge the pGEM-T vector for 4 s to concentrate contents at the bottom of the
5. Vortex the 2X rapid ligation buffer before use.
6. Combine the following in a 0.5 ml microfuge tube:
5 [tl 2X rapid ligation buffer
1 tl pGEM-T vector
x tl PCR product
1 tl T4 DNA ligase
deionized water to a final volume of 10 [tl
7. Mix the reagents.
8. Incubate the mixture for 1 h at room temperature or overnight at 40C if less than the
recommended amount (see equation in step 2) of PCR product was added.
PROTOCOL FOR QIAPREP SPIN MINIPREP KIT, EXTRACTION OF PLASMID
(modified from QIAprep Miniprep Handbook, S. C. Garrett, July 2005)
1. Centrifuge three to five ml of bacteria from overnight bacterial culture.
2. Add RNAase A to Buffer P1.
3. Resuspend pelleted bacterial cells in 250 [tl of buffer PI and transfer to a 1.5 ml plastic
4. Add 250 [tl of buffer P2 and mix by gently inverting 4-6 times.
5. Add 350 [tl of buffer N3 and mix by gently inverting 4-6 times.
6. Centrifuge the extracted DNA for 10 min at 13,000 rpm.
7. Pipette supernatant into a QIAprep column and place column into a collection tube.
8. Centrifuge for 60 s (13,000 rpm).
9. Remove column from collection tube, discard flow-through, and place column back
into collection tube.
10. Add 0.75 ml of buffer PE to column.
11. Centrifuge for 60 s (13,000 rpm).
12. Discard flow-through and then centrifuge for an additional min at 13,000.
13. Transfer the column to a clean 1.5 microcentrifuge tube.
14. Add 50 [tl of buffer EB to the center of the column and let the buffer soak in for one
15. Centrifuge for one min at 13,000 to elute DNA.
PROTOCOL FOR RNA EXTRACTION FROM MOSQUITO POOL USING TRIZOL
(modified (6/7/04) and July/05 (S. C. Garrett) from Leslie Rios's protocol)
1. Homogenize mosquito pool with 1-4 beebees (copper-clad metal airgun shot) in lml
PBS medium with 4% Fetal Bovine Serum.
2. Centrifuge at 15,000 rpm for 10 min.
3. Remove 200 [tl of supernatant into a new tube and save the rest of the supernatant at
4. To the 200 tl, add 600 [tl Trizol LS, then mix and incubate at room temperature for
5. Add 160 [tl of chloroform, mix by inverting for about 15 seconds, and then incubate
at room temperature for 15 min.
6. Centrifuge at 12,500 rpm for one min.
7. Remove the upper aqueous layer to a new tube.
8. Add isopropanol such that the isopropanol is about 0.7 times the volume of the
solution then mix.
9. Centrifuge at 12,500 rpm for 15 min.
10. Pipette off the isopropanol carefully and then add 300 [tl of 70% EtOH.
11. Centrifuge at 12,500 rpm for 5 min.
12. Carefully pipette EtOH and vacuum dry for 10-15 min.
13. Resuspend the dried pellet with 10 [tl of RNAse-free water.
SEQUENCES OF PCR PRODUCTS USED TO IDENTIFY VERTEBRATE HOST
ORIGIN OF MOSQUITO BLOOD MEALS
Ma. dyari (1MBF1(2)); 100% identity with Sus scrofa (wild boar) cyt.b gene (accession #
Ma. dyari (1MBF4); 100% identity with Equus caballus (horse) (accession # D32190):
An. quadrimaculatus (3AnBF), clone 2; 100% identity with Sus scrofa (pig) (accession #
Cx. nigripalpus (1CuNBF2), clone 3; 100% identity with Meleagris gallopavo (turkey)
Cx. nigripalpus (4CuNBF); 98.22% identity with Procyon lotor (raccoon) cyt.b gene
(accession # U12853):
An. quadrimaculatus (3An3BF); 100% identity with Sus scrofa (accession # AY237534):
All of the following sequences (from PCR bands of alligator mitochondrial primers) had
100% identity with Alligator mississippiensis sequence, accession number AF318572:
Alligator positive, bases 39-575:
Cx. erraticus (2CBF):
Cx. erraticus (2ABF)
Ma. dyari (3AAABF):
Ma. dyari (1MBF3):
Ma. dyari (3AABF):
Ma. dyari (2DBF):
Ma. titillans (2MaTBF2):
LIST OF REFERENCES
Alves, E., C. Orilo, M. C. Rodriguez, and L. Solio. 2003. Mitochondrial DNA sequence
variation and phylogenetic relationships among Iberian pigs and other domestic and
wild pig populations. Anim. Genet. 34: 319-324.
Andreadis, T. G. 2003. A checklist of the mosquitoes of Connecticut with new state
records. J. Am. Mosq. Control Assoc. 19: 79-81.
Andreadis, T. G., J. F. Anderson, and C. R. Vossbrinck. 2001. Mosquito surveillance for
West Nile virus in Connecticut, 2000: Isolation from Culexpipiens, Cx. restuans,
Cx. salinarius, and Culiseta melanura. Emerg. Infect. Dis. 7: 670-674.
Alfonzo, D., M. E. Grillet, J. Liria, J.-C. Navarro, S. C. Weaver, and R. Barrera. 2005.
Ecological characterization of the aquatic habitats of mosquitoes (Diptera:
Culicidae) in enzootic foci of Venezuelan Equine Encephalitis virus in western
Venezuela. J. Med. Entomol. 42: 278-284.
Apperson, C. S., B. A. Harrison, T. R. Unnasch, H. K. Hassan, W. S. Irby, H. M. Savage,
S. E. Aspen, D. W. Watson, L. M. Rueda, B. R. Engber, and R. S. Nasci. 2002.
Host-feeding habits of Culex and other mosquitoes (Diptera: Culicidae) in the
borough of Queens in New York City, with characters and techniques for
identification of Culex mosquitoes. J. Med. Entomol. 39: 777-785.
Arunachalam, N., P. Philip Smauel, J. Hiriyan, V. Thenmozhi, A. Balasubramanian, A.
Gajanana, and K. Satyanarayana. 2002. Vertical transmission of Japanese
encephalitis virus in Mansonia species, in an epidemic-prone area of southern
India. Ann. Trop. Med. Parasit. 96: 419-420.
Arunachalam, N., P. Philip Samuel, J. Hiriyan, V. Thenmozhi, and A. Gajanara. 2004.
Japanese Encephalitis in Kerala, South India: Can Mansonia (Diptera: Culicidae)
play a supplemental role in transmission? J. Med. Entomol. 41: 456-461.
Austgen, L. E., R. A. Bowen, M. L. Bunning, B. S. Davis, C. J. Mitchell, and G. J.
Chang. 2004. Experimental infection of cats and dogs with West Nile virus. Emerg.
Infect. Dis. 10: 82-86.
Bernard, K. A., J. G. Maffei, S. A. Jones, E. B. Kauffman, G. D. Ebel, A. P. Dupuis II, K.
A. Ngo, D. C. Nicholas, D. M. Young, P. Shi, V. L. Kulasekera, M. Edison, D. J.
White, W. B. Stone, NY state West Nile virus surveillance team, and L. D. Kramer.
2001. West Nile virus infection in birds and mosquitoes, New York State, 2000.
Emerg. Infect. Dis. 7:679-685.
Bidlingmayer, W. L., 1967. A comparison of trapping methods for adult mosquitoes:
species response and environmental influence. J. Med. Entomol. 4: 200-220.
Bidlingmayer, W. L. 1968. Larval development ofMansonia mosquitoes in central
Florida. Mosq. News 28: 51-57.
Bin, H., Z. Grossman, S. Polamunski, M. Malkinson, L. Weiss, P. Duvdevani, C. Banet,
Y. Weisman, E. Annis, D. Gandaku, V. Yahalom, M. Hindyieh, L. Shulman, and E.
Mendelson. 2001. West Nile fever in Israel 1999-2000: from geese to humans.
Ann. NY Acad. Sci. 951: 127-142.
Blackmore, C. G. M., L. M. Stark, W. C. Jeter, R. L. Oliveri, R. G. Brooks, L. A. Conti,
and S. T. Wiersma. 2003. Surveillance results from the first West Nile virus
transmission season in Florida, 2001. Am. J. Trop. Med. Hyg. 69: 141-150.
Brault, A. C., S. A. Langevin, R. A. Bowen, N. A. Panella, B. J. Biggerstaff, B. R. Miller,
and N. Komar. 2004. Differential virulence of West Nile strains for American
crows. Emerg. Infect. Dis. 10: 2161-2167.
Brinton, M. A. 2002. The molecular biology of West Nile virus: a new invader of the
Western hemisphere. Annu. Rev. Microbiol. 56: 371-402.
Buck, P. A., P. Sockett, I. K. Barker, M. Drebot, R. Lindsay, and H. J. Artsob. 2003.
West Nile virus: surveillance activities in Canada. Ann. Epidemiol. 13: 582.
Bunning, M. L., R. A. Bowen, C. B. Cropp, K. G. Sullivan, B. S. Davis, N. Komar, M. S.
Godsey, D. Baker, D. L. Hettler, D. A. Holmes, B. J. Biggerstaff, and C. J.
Mitchell. 2002. Experimental infection of horses with West Nile virus. Emerg.
Infect. Dis. 8: 380-386.
Burkett, D. A., J. E. Butler, and D. L. Kline. 1998. Field evaluation of colored light
emitting diodes as attractants for woodland mosquitoes and other Diptera in north
central Florida. J. Am. Mosq. Control Assoc. 14: 186-195.
Campbell, G. L., C. S. Ceianu, and H. M. Savage. 2001. Epidemic West Nile encephalitis
in Romania. Ann. NY Acad. Sci. 951: 94-101.
Center for Disease Control and Prevention (CDC), 2005. West Nile Virus Mosquito
species. (last accessed May 12th 2005)
Chu, J. J. H., R. Rajamanonmani, J. Li, R. Bhuvanakantham, J. Lescar, and M.-L. Ng.
2005. Inhibition of West Nile virus entry by using a recombinant domain III from
envelope glycoprotein. J. Gen. Virol. 86: 405-412.
Chowers, M. Y., R. Lang, F. Nassar, D. Ben-David, M. Giladi, E. Rubinshtein, A.
Itzhaki, J. Mishal, Y. Siegman-Igra, R. Kitzes, N. Pick, Z. Landau, D. Wolf, H.
Bin, E. Mendelson, S. D. Pitlik, and M. Weinberger. 2001. Clinical characteristics
of the West Nile fever outbreak, Israel, 2000. Emerg. Infect. Dis. 7: 675-678.
Cicero, C., and N. K. Johnson. 2001. Higher-level phylogeny of New World vireos
(Aves: Vireonidae) based on sequences of multiple mitochondrial DNA genes.
Mol. Phylogenet. Evol. 20: 27-40.
Crans, W. J., and E. G. Rockel. 1968. The mosquitoes attracted to turtles. Mosq. News
Cruz, L, V. M. Cardenas, M. Abarca, T. Rodriguez, R. Flores Reyna, M. V. Serpas, R. E.
Fontaine, D. W. C. Beasley, A. P. A. Travassos Da Rosa, S. C. Weaver, R. B. Tesh,
A. M. Powers, and G. Suarez-Rangel. 2005. Short report: serological evidence of
West Nile virus activity in El Salvador. Am. J. Trop. Med. Hyg. 72: 612-615.
Cupp, E. W., K. Klingler, H. K. Hassan, L. M. Viguers, and T. R. Unnasch. 2003.
Transmission of Eastern Equine Encephalomyelitis virus in central Alabama. Am.
J. Trop. Med. Hyg. 68: 495-500.
Cupp, E. W., K. J. Tennessen, W. K. Oldland, H. K. Hassan, G. E. Hill, C. R. Katholi,
and T. R. Unnasch. 2004a. Mosquito and arbovirus activity during 1997-2002 in a
wetland in northeastern Mississippi. J. Med. Entomol. 41: 495-501.
Cupp, E. W., D. Zhang, X. Yue, M. S. Cupp, C. Guyer, T. R. Sprenger, and T. R.
Unnasch. 2004b. Identification of reptilian and amphibian blood meals from
mosquitoes in an Eastern Equine Encephalomyelitis virus focus in central Alabama.
Am. J. Trop. Med. Hyg. 71: 272-276.
Darsie, R. F. 1998. Keys to the adult females and fourth instar larvae of mosquitoes of
Florida (Diptera: Culicidae)/ R. F. Darsie and C. D. Morris. ,Ft. Meyers, Fla.>:
Florida Mosq. Control. Assoc.
Darsie, R. F., and E. J. Hager. 1993. New mosquito records for South Carolina. J. Am.
Mosq. Control Assoc. 9: 472-473.
Darsie, R. F., Jr., and R. A. Ward. 1981. Identification and geographic distribution of
mosquitoes of North America, north of Mexico. Mosq. Syst. Suppl. 1: 1-313.
Davis, C. T., D. W. C. Beasley, H. Guzman, M. Siirin, R. E. Parsons, R. B. Tesh, and A.
D. T. Barrett. 2004. Emergence of attenuated West Nile virus variants in Texas,
2003. Virology 330: 342-350.
Davis, E. L., J. F. Butler, R. H. Roberts, J. F. Reinert, and D. L. Kline. 1983. Laboratory
feeding of Culicoides mississippiensis (Diptera: Ceratopogonidae) through a
reinforced silicone membrane. J. Med. Entomol. 20: 177-182.
Day, J.F., and G. A. Curtis. 1993. Annual emergence patterns of Culex nigripalpus
females before, during and after a widespread St. Louis Encephalitis epidemic in
South Florida. J. Am. Mosq. Control Assoc. 9: 249-255.
Ebel, G. D., J. Carricaburu, D. Young, K. A. Bernard, and L. D. Kramer. 2004. Genetic
and phenotypic variation of West Nile virus in New York, 2000-2003. Am. J. Trop.
Med. Hyg. 71: 493-500.
Edman, J. D. 1971. Host-feeding patterns of Florida mosquitoes I. Aedes, Anopheles,
Coquillettidia, Mansonia and Psorophora. J. Med. Entomol. 8: 687-695.
Edman, J. D. 1979. Host-feeding patterns of Florida mosquitoes (Diptera: Culicidae) VI.
Culex (Melanoconion). J. Med. Entomol. 15: 521-525.
Edman, J. D., F. D. S. Evans, and J. A. Williams. 1968. Development of a diurnal resting
box to collect Culiseta melanura (COQ.). Am. J. Trop. Med. Hyg. 17: 451-456.
Edman, J. D., L. A. Webber, and H. W. Kale II. 1972. Host-feeding patterns of Florida
mosquitoes II. Culiseta. J. Med. Entomol. 9: 429-434.
Farfan-Ale, J. A., B. J. Blitvich, M. A. Lorono-Pino, N. L. Marlenee, E. P. Rosado-
Paredes, J. E. Garcia-Rejon, L. F. Flores-Flores, L. Chulim-Perera, M. Lopez-
Uribe, G. Perez-Mendoza, I. Sanchez-Herrera, W. Santamaria, J. Moo-Huchim, D.
J. Gubler, B. C. Cropp, C. H. Calisher, and B. J. Beaty. 2004. Longitudinal studies
of West Nile virus infection in avians, Yucatan State, Mexico. Vector-Borne
Zoonot. 4: 3-14.
Farmer, C. G., and D. R. Carrier. 2000. Ventilation and gas exchange during treadmill
locomotion in the American alligator (Alligator mississippiensis). J. Exp. Biol. 203:
Fernandez-Salas, I., J. F. Contreras-Cordero, B. J. Blitvich, J. I. Gonzalez-Rojas, A.
Cavazos-Alvarez, N. L. Marlenee, A. Elizondo-Quiroga, M. A. Lorono-Pino, D. J.
Gubler, B. C. Cropp, C. H. Calisher, and B. J. Beaty. 2003. Serologic evidence of
West Nile virus infection in birds, Tamaulipas State, Mexico. Vector-Borne
Zoonot. 3: 209-213.
Garmendia, A. E., H. J. Van Kruiningen, R. A. French, J. F. Anderson, T. G. Andreadis,
A. Kumar, and A. B. West. 2000. Recovery and identification of West Nile virus
from a hawk in winter. J. Clin. Microbiol. 38: 3110-3111.
Glenn, T. C., J. L. Staton, A. T. Vu, L. M. Davis, J. R. Alvarado Bremer, W. E. Rhodes, I
L. Brisbin JR, and R. H. Sawyer. 2002. Low mitochondrial DNA variation among
American alligators and a novel non-coding region in Crocodilians. J. Exp. Zoo.
Goddard, J., and B. A. Harrison. 2005. New, recent, and questionable mosquito records
from Mississippi. J. Am. Mosq. Control Assoc. 21: 10-14.
Gokool, S., C. F. Curtis, and D. F. Smith. 1993. Analysis of mosquito bloodmeals by
DNA profiling. Med. Vet. Entomol. 7: 208-215.
Hardy, J. L., E. J. Houk, L. D. Kramer, and W. C. Reeves. 1983. Intrinsic factors
affecting vector competence of mosquitoes for arboviruses. Annu. Rev. Entomol.
Huang, C., B. Slater, R. Rudd, N. Parchuri, R. Hull, M. Dupuis, and A. Hindenburg.
2002. First isolation of West Nile virus from a patient with encephalitis in the
United States. Emerg. Infect. Dis. 8: 1367-1371.
Hubalek, Z., and J. Halouzka. 1999. West Nile fever-a reemerging mosquito-borne viral
disease in Europe. Emerg. Infect. Dis. 5: 643-650.
Irby, W. S. and C. S. Apperson. 1988. Hosts of mosquitoes in the coastal plain of North
Carolina. J. Med. Entomol. 25: 85-93.
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. J. Wildlife Dis. 41: 96-106.
Jacobson, E.R., Johnson, A.J., Hernandez, J.A., Tucker, S., Dupuis, A., Stevens, R.,
Carbonneau, D., Stark, L. 2005b. Validation and use of an indirect enzyme-linked
immunosorbent assay for detection of antibodies of West Nile virus in American
alligators (Alligator mississippiensis) in Florida. J. Wildlife Dis. 41: 107-114.
Janke, A., and U. Arnaston. 1997. The complete mitochondrial genome of Alligator
mississippiensis and separation between recent archosauria (birds and crocodiles).
Mol. Biol. Evol. 14: 1266-1272.
Johnson, B. W., T. V. Chambers, M. B. Crabtree, J. Arroyo, T. P. Monath, and B. R.
Miller. 2003. Growth characteristics of the veterinary vaccine candidate
ChimeriVaxTM-West Nile (WN) virus in Aedes and Culex mosquitoes. Med. Vet.
Entomol. 17: 235-243.
Jupp, P. G. 2001. The ecology of West Nile virus in South Africa and the occurrence of
outbreaks in humans. Ann. NY Acad. Sci. 951: 143-152.
Jupp, P. G. 1974. Laboratory studies on the transmission of West Nile virus by Culex
(Culex) univittatus Theobald; factors influencing the transmission rate. J. Med. Ent.
Kauffman, E. B., S. A. Jones, A. P. Dupuis II, K. A. Ngo, K. A. Bernard, and L. D.
Kramer. 2003. Virus detection protocols for West Nile virus in vertebrate and
mosquito specimens. J. Clin. Microbiol. 41:3661-3667.
Khan, S. A., K. Narain, P. Dutta, R. Handique, V. K. Srivastava, and J. Mahanta. 1997.
Biting behavior and biting rhythm of potential Japanese encephalitis vectors in
Assam. J. Communicable Dis. 29: 109-120.
Kilpatrick, A. M., L. D. Kramer, S. R. Campbell, E. O. Alleyne, A. P. Dobson, and P.
Daszak. 2005. West Nile virus risk assessment and the bridge vector paradigm.
Emerg. Infect. Dis. 11: 425-428.
Klein, T. A., D. G. Young, and S. R. Telford Jr. 1987. Vector incrimination and
experimental transmission of Plasmodiumfloridense by bites of infected Culex
(Melanoconion) erraticus. J. Am. Mosq. Control Assoc. 3: 165-175.
Klenk, K. and N. Komer. 2003. Poor replication of West Nile Virus (New York 1999
strain) in three reptilian and one amphibian species. Am. J. Trop. Med. Hyg. 69:
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. Emerg. Infect. Dis. 10: 2150-2155.
Kline, D. L., and M. O. Mann. 1998. Evaluation ofbutanone, carbon dioxide, and 1-
octen-3-OL as attractants for mosquitoes associated with north central Florida bay
and cypress swamps. J. Am Mosq. Control Assoc. 14: 289-297.
Komar, O., M. B. Robbins, K. Klenk, B. J. Blitvich, N. L. Marienee, K. L. Burkhalter, D.
J. Gubler, G. Gonzalvez, C. J. Pefia, A. Townsend Peterson, and N. Komar. 2003.
West Nile virus transmission in resident birds, Dominican Republic. Emerg. Infect.
Dis. 9: 1299-1302.
Kornegay, J. R., T. D. Kocher, L. A. Williams, and A. C. Wilson. 1993. Pathways of
lysozyme evolution inferred from sequences of cytochrome b in birds. J. Mol. Evol.
Kulasekera, V. L., L. Kramer, R. S. Nasci, F. Mostashardi, B. Cherry, S. C. Trock, C.
Glaser, and J. R. Miller. 2001. West Nile virus infection in mosquitoes, birds,
horses, and humans, Staten Island, New York, 2000. Emerg. Infect. Dis. 7: 722-
Lanciotti, R. S., G. D. Ebel, V. Deubel, A. J. Kerst, S. Murri, R. Meyer, M. Bowen, N.
McKinney, W. E. Morrill, M. B. Crabtree, L. D. Kramer, and J. T. Roehrig. 2002.
Complete genome sequence and phylogenetic analysis of West Nile virus strains
isolated from the United States, Europe, and the Middle East. Virology 298: 96-
Lanciotti, R. S., A. J. Kerst, R. S. Nasci, M. S. Godsey, C. J. Mitchell, H. M. Savage, N.
Komar, N. A. Panella, B. C. Allen, K. E. Volpe, B. S. Davis, and J. T. Roehrig.
2000. Rapid detection of West Nile virus from human clinical specimens, field-
collected mosquitoes, and avian samples by a Taqman reserve transcriptase-PCR
assay. J. Clin. Microbiol. 38: 4066-4071.
Lanciotti, R. S., J. T. Roehrig, V. Deubel, J. Smith, M. Parker, K. Steele, B. Crise, K. E.
Volpe, M. B. Crabtree, J. H. Scherret, R. A. Hall, J. S. MacKenzie, C. B. Cropp, B.
Panigrahy, E. Ostlund, B. Schmitt, M. Malkinson, C. Banet, J. Weissman, N.
Komar, H. M. Savage, W. Stone, T. McNamara, and D. J. Gubler. 1999. Origin of
the West Nile virus responsible for an outbreak of encephalitis in the northeastern
United States. Science. 286: 2333-2337.
Lane, Thomas J., and King, Wayne F. 1989. Alligator production in Florida. Cooperative
Extension Service publication. University of Florida.
Lee, J. W. M. and M. L. Ng. 2004. A nano-view of West Nile virus-induced cellular
changes during infection. Journal of Nanobiotechnology. Open access:
(http://www.jnanobiotechnology.com/content/2/1/6), last accessed February, 2005.
Lento, G. M. 1995. Use of spectral analysis to test hypotheses on the origin of pinnipeds.
Mol. Biol. Evol. 12: 28-52.
Lothrop, B. B., R. P. Meyer, W. K. Reisen, and H. Lothrop. 1995. Occurrence of Culex
(Melanoconion) erraticus (Diptera: Culicidae) in California. J. Am. Mosq. Control
Assoc. 11: 367-368.
Lounibos, L. P., and Escher, R. L. 1985. Mosquitoes associated with water lettuce (Pistia
stratiotes) in Southeastern Florida. Fla. Entomol. 68: 169-178.
Lounibos, L. P., V. L. Larson, and C. D. Morris. 1990. Parity, fecundity, and body size of
Mansonia dyari in Florida. J. Am. Mosq. Control Assoc. 6: 121-126.
Love, G. J., and W. W. Smith. 1957. Preliminary observations on the relation of light trap
collections to mechanical sweep net collections in sampling mosquito populations.
Mosq. News 17: 9-14.
Mans, N. Z., S. E. Yurgionas, M. C. Garvin, R. E. Gary, J. D. Bresky, A. C. Galaitsis,
and O. A. Ohajuruka. 2004. West Nile virus in mosquitoes of northern Ohio, 2001-
2002. Am. J. Trop. Med. Hyg. 70: 562-565.
Marfin, A. A., L. R. Petersen, M. Eidson, J. Miller, J. Hadler, C. Farello, B. Werner, G.
L. Campbell, M. Layton, P. Smith, E. Bresnitz, M. Cartter, J. Scaletta, G. Obiri, M.
Bunning, R. C. Craven, J. T. Roehrig, K. G. Julian, S. R. Hinten, D. J. Gubler, and
the ArboNET Cooperative Surveillance Group. 2001. Widespread West Nile virus
activity, eastern United States, 2000. Emerg. Infect. Dis. 7: 730-735.
McKenzie, K. E. 2003. Determining factors of preferential host selection by Aedes
aegypti. Ph.D. dissertation. University of Florida, Gainesville, 148 pp.
McLean, R. G., S. R. Ubico, D. E. Docherty, W. R. Hansen, L. Sileo, and T. S.
McNamara. 2001. West Nile virus transmission and ecology in birds. Ann. NY
Acad. Sci. 951: 54-57.
Meece, J. K., J. S. Henkel, L. Glaser, and K. D. Reed. 2003. Mosquito surveillance for
West Nile virus in southeastern Wisconsin 2002. Clin. Med. Res. 1: 37-42.
Melnick, J. L., J. R. Paul, J. T. Riordan, V. H. Bamett, N. Goldblum, and E. Zabin. 1951.
Isolation from human sera in Egypt of a virus apparently identical to West Nile
virus. P. Soc. Exp. Biol. Med. 77: 661-665.
Mendez, W., J. Liria, J. C. Navarro, C. Z. Garcia, J. E. Freier, R. Salas, S. C. Weaver, and
R. Barrera. 2001. Spatial dispersion of adult mosquito (Diptera: Culicidae) in a
sylvatic focus of Venezuelan equine encephalitis virus. J. Med. Entomol. 38: 813-
Miller, B. R., R. S. Nasci, M. S. Godsey, H. M. Savage, J. J. Lutwama, R. S. Lanciotti, C.
J. Peters. 2000. First field evidence for natural vertical transmission of West Nile
virus in Culex univittatus complex mosquitoes from Rift Valley Province, Kenya.
Am. J. Trop. Med. Hyg. 62: 240-246.
Miller, D.L., Mauel, M.J., Baldwin, C., Burtle, G., Ingram, D., Hines, M.E. II, Frazier,
K.S. 2003. West Nile virus in farmed alligators. Emerg. Infect. Dis. 9: 794-799.
Morris, C. D., V. L. Larson, and L. P. Lounibos. 1991. Measuring mosquito dispersal for
control programs. J. Am. Mosq. Control Assoc. 7: 608-615.
Mostashari, F., M. Kulldorff, J. J. Hartman, J. R. Miller, and V. Kulasekera. 2003. Dead
bird clusters as an early warning system for West Nile virus activity. Emerg. Infect.
Dis. 9: 641-646.
Mukabana, W. R., W. Takken, and B. G. J. Knols. 2002. Analysis of arthropod
bloodmeals using molecular genetic markers. Trends Parasitol. 18: 505-509.
Moussa, M. A., D. J. Gould, M. P. Nolan Jr., and D. E. Hayes. 1966. Observations on
Culiseta melanura (Coquillett) in relation to encephalitis in southern Maryland.
Mosq. News 26: 385-393.
Murgue, B., S. Murri, S. Zientara, B. Durand, J.P. Durand, and H. Zeller. 2001. West
Nile outbreak in horses in southern France 2000: the return after 35 years. Emerg.
Infect. Dis. 7: 692-696.
Murphey, F. J., P. P. Burbutis, and D. F. Bray. 1967. Bionomics of Culex salinarius
Coquillett. II. Host acceptance and feeding by adult females of Cx. salinarius and
other mosquito species. Mosq. News 27: 366-374.
Nasci, R. S., K. L. Gottfried, K. L. Burkhalter, V. L. Kulasekera, A. J. Lambert, R. S.
Lanciotti, A. R. Hunt, and J. R. Ryan. 2002. Comparison ofvero cell plaque assay,
Taqman reverse transcriptase polymerase chain reaction RNA assay, and Vectest
anigen assay for detection of West Nile virus in field-collected mosquitoes. J. Am.
Mosq. Control Assoc. 18: 294-300.
Nayer, J. K., and D. M. Sauerman. 1977. The effects of nutrition on survival and
fecundity in Florida mosquitoes: Effects of blood source on oocyte development. J.
Med. Entomol. 14: 167-174.
Ng, M. L., S. H. Tan, and J. J. Chu. 2001. Transport and budding at two distinct sites of
visible nucleocapsids of West Nile (Sarafend) virus. J. Med. Virol. 65: 758-764.
Ngo, K. A., and L. D. Kramer. 2003. Identification of mosquito bloodmeals, using
polymerase chain reaction (PCR) with order-specific primers. J. Med. Entomol. 40:
Nieblyski, M. L., H. M. Savage, R. S. Nasci, and G. B. Craig. 1994. Blood hosts ofAedes
albopictus in the United States. J. Am. Mosq. Control Assoc. 10: 447-450.
Nir, Y., Y. Lasowski, A. Avivi, R. Cgoldwasser, 1969. Survey for antibodies to
arboviruses in the serum of various animals in Israel during 1965-1966. Am. J.
Trop. Med. Hyg. 18: 416-422.
Odelola, H. A., and A. Fabiyi. 1977. Biological characteristic of Nigerian strains of West
Nile virus in mice and cell cultures. Acta Virol. 21: 161-164.
Odelola, H. A., and 0. O. Oduye. 1977. West Nile virus infection of adult mice by oral
route. Arch. Virol. 54: 251-253.
Poidinger, M., R. A. Hall, and J. S. Mackenzie. 1996. Molecular characterization of the
Japanese Encephalitis serocomplex of the Flavivirus genus. Virology 218: 417-421.
Quirin, R., M., M. Salas, S. Zientara, H. Zeller, J. Labie, S. Murri, T. Lefrancois, M.
Petitclerc, and D. Martinez. 2004. West Nile virus, Guadeloupe. Emerg. Infect. Dis.
Rappole, J. H., S. R. Derrickson, and Z. Hubalek. 2000. Migratory birds and spread of
West Nile virus in the Western hemisphere. Emerg. Infect. Dis. 6: 319-328.
Ray, D. A., and L. Densmore. 2002. The Crocodilian mitochondrial control region:
general structure, conserved sequences, and evolutionary implications. J. Exp. Zoo.
Reeves, W. C. 1957. Arthropods as vectors and reservoirs of animal pathogenic viruses.
pp. 177-202. In C. Hallauer and K. F. Meyer [eds.], Handbuch der Virus
Forschung, vol 4, Suppl. 3, Springer, Vienna, Austria.
Reeves, W. C., E. L. French, N. Marks, and N. E. Kent. 1954. Murray Valley
encephalitis: a survey of suspected mosquito vectors. Am. J. Trop. Med. Hyg. 3:
Reisen, W. K., K. Boyce, R. C. Cummings, O. Delgado, A. Gutierrez, R. P. Meyer, and
T. W. Scott. 1999. Comparative effectiveness of three adult mosquito sampling
methods in habitats representative of four different biomes of California. J. Am.
Mosq. Control Assoc. 15: 24-31.
Reisen, W., H. Lothrop, R. Chiles, M. Madon, C. Cossen, L. Woods, S. Husted, V.
Kramer, and J. Edman. 2004. West Nile virus in California. Emerg. Infect. Dis. 10:
Roberson, L. C., S. Prior, C. S. Apperson, and W. S. Irby. 1993. Bionomics of Anopheles
quadrimaculatus and Culex erraticus (Diptera: Culicidae) in the Falls Lake basin,
North Carolina: seasonal changes in abundance and gonotrophic status, and host-
feeding patterns. J. Med. Entomol. 30: 689-698.
Rutledge, C. R., J. F. Day, C. C. Lord, L. M. Stark, and W. J. Tabachnick. 2003. West
Nile virus infection rates in Culex nigripalpus (Diptera: Culicidae) do not reflect
transmission rates in Florida. J. Med. Entomol. 40: 253-258.
Sbrana, E., J. H. Tonry, S-Y. Xiao, A. P. A. Travassos Da Rosa, S. Higgs, and R. B.
Tesh. 2005. Oral transmission of West Nile virus in a hamster model. Am. J. Trop.
Med. Hyg. 72: 325-329.
Schuffenecker, I., C. N. Peyrefitte, M. el Harrak, S. Murri, A. Leblond, and H. G. Zeller.
2005. West Nile virus in Morocco, 2003. Emerg. Infect. Dis. 11: 306-309.
Slaff, M. and J. D. Haefner. 1985. Seasonal and spatial distribution ofMansonia dyari,
Mansonia titillans, and Coquillettidiaperturbans (Diptera: culicidae) in the central
Florida, USA, phosphate region. J. Med. Entomol. 22: 624-629.
Smithburn, K. C., T. P. Hughes, A. W. Burke, and J. H. Paul. 1940. A neurotropic virus
isolated from the blood of a native of Uganda. Am. J. Trop. Med. Hyg. 20: 471-
Steinman, A., C. Banet-Noach, S. Tal, O. Levi, L. Simanov, S. Perk, M. Malkinson, N.
Shpigel. 2003. Letter to Editor: West Nile virus infection in crocodiles. Emerg.
Infect. Dis. 9: 887.
Sudia, W. D., and R. W. Chamberlain. 1988. Classic paper: Battery-operated light trap,
an improved model. J. Am. Mosq. Control Assoc. 4: 536-538.
Swayne, D. E., J. R. Beck, and S. Zaki. 2000. Pathogenicity of West Nile virus for
turkeys. Avian Dis. 44: 932-937.
Teehee, M. L., M. L. Bunning, S. Stevens, and R. A. Bowen. 2005. Experimental
infection of pigs with West Nile virus. Arch. Virol. 150: 1249-1256.
Tempelis, C. H. and P. Galindo. 1975. Host-feeding patterns of Culex (Melanoconion)
and Culex (Aedinus) mosquitoes collected in Panama. J. Med. Entomol. 12: 205-
Tempelis, C. H. 1975. Host-feeding patterns of mosquitoes, with a review of advances in
analysis of blood meals by serology. J. Med. Entomol. 11: 635-653.
Traore-Lamizana, M., D. Fontenille, M. Diallo, Y. Ba, H. G. Zeller, M. Mondo, F. Adam,
J. Thonon, and A. Maiga. 2001. Arbovirus surveillance from 1990 to 1995 in the
Barkedji area (Ferlo) of Senegal, a possible natural focus of Rift Valley Fever
virus. J. Med. Entomol. 38: 480-492.
Tuno, N., Y. Tsuda, M. Takagi, and W. Swonderd. 2003. Pre-and postprandial mosquito
resting behavior around cattle hosts. J. Am. Mosq. Control Assoc. 19: 211-219.
Turell, M. J., D. J. Dohm, M. R. Sardelis, M. L. O'Guinn, T. G. Andreadis, and J. A.
Blow. 2005. An update on the potential of North American mosquitoes (Diptera:
Culicidae) to transmit West Nile virus. J. Med. Entomol. 42: 57-62.
Turell, M. J., J. W. Jones, M. R. Sardelis, D. M. Dohm, R. E. Coleman, D. M. Watts, R.
Fernandez, C. Calampa, and T. A. Klein. 2000. Vector competence of Peruvian
mosquitoes (Diptera: Culicidae) for Epizootic and Enzootic strains of Venezuelan
Equine Encephalomyelitis virus. J. Med. Entomol. 37: 835-839.
White, D. J., L. D. Kramer, P. B. Backenson, G. Lukacik, G. Johnson, J. Oliver, J. J.
Howard, R. G. Means, M. Eidson, I. Gotham, V. Kulasekera, S. Campbell,
Arbovirus Research Laboratory, and the Statewide West Nile virus response teams.
2001. Mosquito surveillance and polymerase chain reaction detection of West Nile
virus, New York State. Emerg. Infect. Dis. 7: 643-649.
Wozniak, A., H. E. Dowda, M. W. Tolson, N. Karabatsos, D. R. Vaughan, P. E. Turner,
D. I. Oritz, and W. Wills. 2001. Arbovirus surveillance in South Carolina, 1996-
1998. J. Am. Mosq. Control Assoc. 17: 73-78.
Yau., F. C.-F., K. Wong, J. Wang, P. Pui-Hay But, and P. Shaw. 2002. Generation of a
sequence characterized amplified region probe for authentication of Crocodilian
species. J. Ex. Zoo. 294: 382-386.
Yokota, M., N. Tatsumi, O. Nathalang, T. Yamada, and I. Tsuda. 1999. Effects of heparin
on Polymerase Chain Reaction for blood white cells. J. Clin. Lab. Anal. 13: 133-
Zhong, H., Z. Yan, F. Jones, and C. Brock. 2003. Ecological analysis of mosquito light
trap collections from west central Florida. Environ. Entomol. 32: 807-815.
Sandra Coral Garrett was born on the windy, cold morning of March 8, 1981 in
Aiken, South Carolina, to Dr. Alfred J. Garrett and Susan Hersey Garrett. She has three
siblings: an older brother Travis, a younger sister Allison, and a younger brother
Benjamin. Sandra and her siblings grew up in Aiken, but also spent time on Hilton Head
Island, a barrier island near the Georgia-South Carolina border. Both places presented
the children with opportunity for outdoor exploration, and thus allowed Sandra to
develop a strong interest in biology in addition to outdoor sports and art.
Sandra and her siblings all attended the South Carolina Governor's School for
Science and Mathematics for the last two years of their high school education. This
unique school and its outstanding teachers helped Sandra explore her interests in the
biological sciences and prepared her for college and research pursuits. It was during a
school tour of the Clemson entomology department that Sandra decided entomology
might be an exciting and rewarding area of biology to study. She became interested in
the University of Florida's strong entomology department and was able to attend with
financial assistance from UF's National Merit Scholar program.
Sandra received a BS in entomology from UF. Experiences like her senior thesis
work with Dr. Howard Frank and the Tropical Entomology field trip to Venezuela further
strengthened her interest in entomology. She graduated summa cum laude from UF and
decided to stay for a master's degree. She met her future husband, Dr. Jose Carlos V.
Rodrigues, in the department and was married in May of 2005.