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Identification of Potential Mosquito Vectors of West Nile Virus on a Florida Alligator Farm

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Title:
Identification of Potential Mosquito Vectors of West Nile Virus on a Florida Alligator Farm
Copyright Date:
2008

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Subjects / Keywords:
Alligators ( jstor )
Animal feeding behavior ( jstor )
Birds ( jstor )
Boxes ( jstor )
DNA ( jstor )
Infections ( jstor )
Polymerase chain reaction ( jstor )
Reverse transcriptase polymerase chain reaction ( jstor )
Species ( jstor )
West Nile virus ( jstor )

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University of Florida
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University of Florida
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4/17/2006

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IDENTIFICATION OF POTENTIAL MOSQUITO VECTORS OF WEST NILE VIRUS
ON A FLORIDA ALLIGATOR FARM















By

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


2005

































Copyright 2005

by

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.















ACKNOWLEDGMENTS

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

page

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

CHAPTER

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



v









APPENDIX

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


Table page

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


Figure pge

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

By

Sandra C. Garrett

December 2005

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.














CHAPTER 1
INTRODUCTION

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.,

2004).

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

(Bidlingmayer, 1967).

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

(Tempelis, 1975).

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

reptiles.

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

unknown.

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

potential vectors.









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.














CHAPTER 2
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.

Mosquito Collecting

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

(Fig. 2-2).























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

days.






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.


Wastewater


0
0
*0

*0
4H


o*
*0 0@0
000
000
*oo


*+I +

O0

f


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

use.










E...











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

before.

Table 2-1. Primers sets in PCR used to amplify DNA from different vertebrate hosts.
Product
Host Primer Sequence size(bp) Reference

Alligator Forward CGCTTCACTGCCCTACACTT 850 Current study
Reverse GCTTTAGTGTTTAAGCTACGATAACTG

Yau et al.
Crocodilian Forward GATGTGGACCTTCAGGATGC 209 (2002)
Reverse CAGAGGTTCAATCCACGGTT

Cicero and
Avian Forward GACTGTGACAAAATCCCNTTCCA 508 Johnson (2001)
Reverse GGTCTTCATCTYHGGYTTACAAGAC

Ngo and
MammalianForward CGAAGCTTGATATGAAAAACCATCGTTG 772 Kramer (2003)
Reverse TGTAGTTRTCWGGGTCHTCTA


Virus Detection

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).









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
Envelop
Set WN1229 GGGTCAGCACGTTTGTCATTG glycoprotein

NS5: RNA-
dependent
RNA
Screening WN9483 CACCTACGCCCTAAACACTTTCACC 311 polymerase
NS5: RNA-
dependent
RNA
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


Mammalian


Avian


Crocodilian







Alligator


WNV


Buffer
dNTP's
each primer
MgCl2
Taq polymerase
template DNA

Buffer
dNTP's
each primer
MgCl2
Taq polymerase
template DNA

Buffer
dNTP's
each primer
MgCl2
Taq polymerase
template DNA

Buffer
dNTP's
each primer
MgCl2
Taq polymerase
template DNA

Buffer
dNTP's
each primer
MgSO4
Taq polymerase
Reverse
transcriptase
Template RNA


1X
0.2 mM each
5 pmol/rxn
4 mM
1 unit/rxn
1 pl

1X
0.2 mM
15 pmol/rxn
2.0 mM
1 unit/rxn
1 pl

1X
0.2 mM
10 pmol/rxn
2.5 mM
1 unit/rxn
2 pl1

1X
0.2 mM
10 pmol/rxn
2 mM
0.2 pl/rxn
1 pl

1X
0.2 mM
0.4 pmol/pl
2 mM
1 unit/rxn

1 unit/rxn
1 pl


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.














CHAPTER 3
RESULTS

Mosquito Collecting

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 [4] 272
Culex erraticus [3] 273
Culex nigripalpus [5] 535
Anopheles crucians [1] 737
Mansonia dyari [9] 1170
Anopheles quadrimaculatus [15] 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

aspiration.

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).











900

800

700

600

500
0 outside pens
400 inside pens

300

200

100

0 T







Mosquito species

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

trips.

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[1] 6[1] 323[3] 338 338
Ma. titillans 63[2] 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 [1] 54[1] 122[1] 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[1] 8
Ma. dyari 2 7 22 0 31
Ma. titillans 4 [2] 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 [1] 0 [2] 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.
CDC
trap Resting TOTAL
trap 1 trap2 trap3 trap4 trap5 trap6 box

An. quadrimaculatus 7 9 36 6 7 109 [9] 183
An. crucians 2 9 30 0 0 0 0 41
Ma. dyari 4 0 58 3 4 39[3] 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

collected.

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


trap

An. quadrimaculatus
An. crucians
Ma. dyari
Ma. titillans
Ps. columbiae
Ur. sapphirina
Ur. lowii
Ae. alobopictus
Cx. erraticus
Cx. nigripalpus
Cq. perturbans
Ps. ferox
Oc. infirmatus
Cs. melanura
Cx. quinquefasciatus
Wy. vanduzeei
TOTAL


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[1] 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

database.

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


4 box

3
15
30
9
1
4
1
1
2
10
0
0
2
1
0
0
79


TOTAL

171
142
318
68
9
26
2
3
49
229
10
0
8
3
2
1
1041


25[6]
3
0
0
0
0
0
0
2
1
0
0
0
0
1
0
32









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






43
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


44

5 6 7 8 9 10


-
uNiK
c/l


.ii ..





a.Y


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.


^^j^kn

*>i~


t*~j


W









Table 3-5. Identities of vertebrate hosts as determined by sequencing the PCR product,


Primer set


Mammal


and information about collection
sample.
Mosquito


Species


Ma. dyari




An.
quadrimaculatus


Cx. nigripalpus


Cx. nigripalpus


Alligator


Cx. erraticus


Ma. dyari








Ma. titillans


date and location on farm of the mosquito


Vertebrate host


Collection
date


Trap
location


ID number


1MBF4 Equus callabus
(horse)a
1MBF1(2) Sus scrofa (wild
boar)b

3AnBF
Sus scrofa
3An3BF
Sus scrofa
3An8BF ***


4CuNBF


Procyon lotor
(raccoon)c


1CuNBF2 Meleagris
gallopavo
(turkey)d

Alligator
mississippiensise
2ABF
Alligator
2CBF mississippiensis
Alligator
2DBF mississippiensis
Alligator
1MBF3 mississippiensis
Alligator
3AABF mississippiensis
Alligator
3AAABF mississippiensis
Alligator
2MaTBF2 mississippiensis


References for identification of sequences:
1994); b = AY237534 (Alves et al., 2003);:


(Kornegay et al., 1993);e
identified.


Sept. 12

Sept. 12


Oct. 17

Oct. 17

Oct. 17

Oct. 24


Inside pen

In trees on
West
margin
Resting
box
Resting
box
Resting
box
Eastern
margin on
tree


Sept. 12 In trees on
West
margin


Resting
Sept. 26 box


Sept. 26

Sept. 26

Sept. 12

Oct. 17

Oct. 17

Sept. 26


Inside pen

Inside pen

Inside pen

CDC 8

CDC 8

Inside pen


: GenBank accession # D32190 (Chikuni,
U12853 (Lento et al., 1995); d = L08381


AF318572 (Glen et al., 2002), ***


specific host not


Bird









Virus Detection

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

of 0/4447.

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.














CHAPTER 4
DISCUSSION

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

study.

Virus Detection

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

species.

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.

Vector Incrimination

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.

Mosquito Control

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).














CHAPTER 5
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.














APPENDIX A
PROTOCOL FOR QIAGEN QIAQUICK SPIN KIT, PURIFICATION OF DNA FROM
AGAROSE GEL

(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.















APPENDIX B
ABI PRISMTM DYE TERMINATOR CYCLE SEQUENCING KIT, PROTOCOL FOR
DNA SEQUENCING

(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
ladder.
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
tube:

REAGENT QUANTITY
Terminator Ready Reaction Mix 2.0 [tL
ICBR dNTP mix 2.0 [tL
Template
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).

Thermocycle program:
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
10 min.
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
DNA.
16. Dry under vacuum.
17. Store in dark freezer until ready to read.













APPENDIX C
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
tube.
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.















APPENDIX D
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
microcentrifuge tube.
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
min.
15. Centrifuge for one min at 13,000 to elute DNA.














APPENDIX E
PROTOCOL FOR RNA EXTRACTION FROM MOSQUITO POOL USING TRIZOL
LS (GIBCO)

(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
-800C.
4. To the 200 tl, add 600 [tl Trizol LS, then mix and incubate at room temperature for
five min.
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.
















APPENDIX F
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 #
AY237534):

ATCCGAAAATCACACCCACTAATAAAAATTATCAACAACGCATTCATTGACCTCCCAGC
CCCCTCAAACATCTCATCATGATGAAACTTCGGTTCCCTCTTAGGCATCTGCCTAATCT
TGCAAATCCTAACAGGCCTGTTCTTAGCAATACATTACACATCAGACACAACAACAGCT
TTCTCATCAGTTACACACATTTGTCGAGACGTAAATTACGGATGAGTTATTCGCTATCT
ACATGCAAACGGAGCATCCATATTCTTTATTTGCCTATTCATCCACGTAGGCCGAGGTC
TATACTACGGATCCTATATATTCCTAGAAACATGAAACATTGGAGTAGTCCTACTATTT
ACCGTTATAGCAACAGCCTTCATAGGCTACGTCCTGCCCTGAGGACAAATATCATTCTG
AGGAGCTACGGTCATCACAAATCTACTATCAGCTATCCCTTATATCGGAACAGACCTCG
TAGAATGAATCTGAGGGGGCTTTTCCGTCGACAAAGCAACCCTCACACGATTCTTCGCC
TTCCACTTTATCCTGCCATTCATCATTACCGCCCTCGCAGCCGTACAT

Ma. dyari (1MBF4); 100% identity with Equus caballus (horse) (accession # D32190):

TGGAATGGGATTTTGTCCATATCGGATGGGATTCCTGAGGGGTTGTTAGATCCTGTTTC
GTGAAGAAATAGTAAATGTACGACTACCAGGGCTGTGATGATGAAGGGTAGGATGAAGT
GGAAAGCAAAAAATCGGGTAAGGGTGGCTTTGTCTACTGAGAATCCACCTCAGATTCAC
TCGACGAGGGTAGTACCGATGTAGGGAATTGCTGATAGGAGGTTCGTGATGACTGTTGC
TCCTCAAAAGGATATTTGGCCTCATGGTAGGACATAGCCCATGAATGCTGTAGCTATAA
CTGTGAAAAGTAGGATGATTCCAATGTTTCATGTCTCTAGGAATGTGTAAGAGCCGTAG
TAGAGGCCGCGTCCTACGTGAATGAAGAGGCAGATAAAAAATATTGATGCTCCGTTGGC
ATGGAGGTAGCGAATAATTCATCCGTAGTTAACGTCTCGGCAGATGTGAGTGACGGATG
AGAAGGCAGTTGTCGTGTCTGATGTGTAGTGTATGGCTAGGAATAGGCC

An. quadrimaculatus (3AnBF), clone 2; 100% identity with Sus scrofa (pig) (accession #
AY237534):

CCACTAATAAAAATTATCAACAACGCATTCATTGACCTCCCAGCCCCCTCAAACATCTC
ATCATGATGAAACTTCGGTTCCCTCTTAGGCATCTGCCTAATCTTGCAAATCCTAACAG
GCCTGTTCTTAGCAATACATTACACATCAGACACAACAACAGCTTTCTCATCAGTTACA
CACATTTGTCGAGACGTAAATTACGGATGAGTTATTCGCTATCTACATGCAAACGGAGC
ATCCATATTCTTTATTTGCCTATTCATCCACGTAGGCCGAGGTCTATACTACGGATCCT
ATATATTCCTAGAAACATGAAACATTGGAGTAGTCCTACTATTTACCGTTTAGCAACA
GCCTTCATAGGCTACGTCCTGCCCTGAGGACAAATATCATTCTGAGGAGCTACGGTCAT
CACAAATCTACTATCAGCTATCCCTTATATCGGAACAGACCTCGTAGAATGAATCTGGG










GGGGCTTTTCCGTCGACAAAGCAACCCTCACACGATTCTTCGCCTTCCACTTTATCCTG
CCATTCATCATTACCGCCCTCGCAGCCGTACATCTCCTATTCCTGCACGAAACCGGATC
C

Cx. nigripalpus (1CuNBF2), clone 3; 100% identity with Meleagris gallopavo (turkey)
(accession #L08381):

CTTACTCACATTAACCCTATTCTCACCTAACCTCTTAGGAGACCCAGAAAACTTTACCC
CAGCAAATCCACTAGTAACCCCCCCACACATTAAACCAGAGTGATACTTTCTATTTGCC
TACGCAATCCTACGCTCAATCCCAAACAAACTTGGAGGTGTCCTAGCCTTAGCAGCATC
AGTACTCATTCTTCTCCTTATCCCCTTCCTTCATAAATCTAAACAACGGGCATAAAT
TCCGGCCACTCTCACAAACCTTATTCTGACTCTTAGTAGCAAACCTCCTCATCCTAACC
TGAGTAGGAAGCCAAACGTAGAACACCCATTCATCATCATGGCCAAATAGCATCCCT
TTCCTACTTCACTATCTTACTAATCCTCTTCCCCTTAATCGGAGCCCTAGAAAACAAAA
TACTCAACCTCTAAGTACTCTAATAGTTTATGAAAAAC

Cx. nigripalpus (4CuNBF); 98.22% identity with Procyon lotor (raccoon) cyt.b gene
(accession # U12853):

ATCCGAAAAACTCACCCATTAGCTAAAATCGTCAACAACTCATTCATTGATCTACCCAC
CCCCTCAAACATCTCAGCATGATGAAATTTCGGCTCCCTCCTCGGAATTTGTTTGCTTC
TACAGATCGCAACAGGTTTATTCTTAGCCATGCACTACACACCAGATACAGCCACAGCT
TTCTCATCAGTGACCCACATTTGCCGAGATGTAAATTATGGCTGAATTATCCGATATAT
ACACGCTAACGGAGCTTCTATATTCTTTATATGCCTATTCTTACACGTAGGACGAGGCT
TATACTATGGCTCCTATACATTCTCTGAAACATGAAATATTG

An. quadrimaculatus (3An3BF); 100% identity with Sus scrofa (accession # AY237534):

ATCCGAAAATCACACCCACTAATAAAAATTATCAACAACGCATTCATTGACCTCCCAGC
CCCCTCAAACATCTCATCATGATGAAACTTCGGTTCCCTCTTAGGCATCTGCCTAATCT
TGCAAATCCTAACAGGCCTGTTCTTAGCAATACATTACACATCAGACACAACAACAGCT
TTCTCATCAGTTACACACATTTGTCGAGACGTAAATTACGGATGAGTTATTCGCTATCT
ACATGCAAACGGAGCATCCATATTCTTTATTTGCCTATTCATCCACGTAGGCCGAGGTC
TATACTACGGATCCTATATATTCCTAGAAACATGAAACATTGGAGTAGTCCTACTATTT
ACCGTTATAGCAACAGCCTTCATAGGCTACGTCCTGCCCTGAGGACAAATATCATTCTG
AGGAGCTACGGTCATCACAAATCTACTATCAGCTATCCCTTATATCGGAACAGACCTCG
TAGAATGAATCTGAGGGGGCTTTTCCGTCGACAAAGCAACCCTCACACGA

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:

ACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCT
GACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGC
TGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAA
ACTTCACCCCAGCAAATTCCATAATTACCCAACACCATCAAACCCGAATGGTACTTC










CTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAAT
ATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAAC
CAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTC
TTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAAC
TGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAG
AGAACA

Cx. erraticus (2CBF):

ACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCT
GACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGC
TGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAA
ACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGTACTTC
CTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAAT
ATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAAC
CAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTC
TTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCATATATTTTAATTGGCCAAAC
TGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAG
AG

Cx. erraticus (2ABF)

CACCCACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAA
ATGCTGACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCA
CTAGCTGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCC
TGAAAACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGT
ACTTCCTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTA
GCAATATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACA
ACAACCAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACT
TCCTCTTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGC
CAAACTGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAAAGCAGGCCT
CTTAGAGAACAAAATAGTTGAACCCACCTATGTTACCCC

Ma. dyari (3AAABF):

CTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCTGA
CAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGCTG
CCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAAAC
TTCACCCCAGCAAATTCCATAATTACCCAACACACATCAAACCCGAATGGTACTTCCT
ATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAATAT
TCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAACCA
ATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTCTT
ACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAACTG
CCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAGAG
AACAAAATAGTTGAACCCACCTATGTTACC


Ma. dyari (1MBF3):











ACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCT
GACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGC
TGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAA
ACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAAACCCGAATGGTACTTC
CTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAAT
ATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAAC
CAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTC
TTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAAC
TGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAG
AGAACAAAATAGTTGAACCCACCTATGTTAC

Ma. dyari (3AABF):

ACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCT
GACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGC
TGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAA
ACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAAACCCGAATGGTACTTC
CTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAAT
ATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAAC
CAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTC
TTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAAC
TGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAG
AGAACAAAATAGTTGAACCCACCTATGTTA

Ma. dyari (2DBF):

CCACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATG
CTGACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTA
GCTGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGA
AAACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGTACT
TCCTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCA
ATATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACA
ACCAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCC
TCTTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAA
ACTGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTT

Ma. titillans (2MaTBF2):

CCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCTG
ACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGCT
GCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAAA
CTTCACCCCAGCAAATTCCATAATTACCCAACACACATCAAACCCGAATGGTACTTCC
TATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAATA
TTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAACC
AATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTCT
TACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAACT







71


GCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAGA
GAACAAAATAGT TGAACCCACCTATGTTAC
















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BIOGRAPHICAL SKETCH

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.




Full Text

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IDENTIFICATION OF POTENTIAL MOSQUI TO VECTORS OF WEST NILE VIRUS ON A FLORIDA ALLIGATOR FARM By SANDRA C. GARRETT A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLOR IDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2005

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Copyright 2005 by Sandra C. Garrett

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This document is dedicated to my husband, Dr Jos Carlos V. Rodrigues, and to my father, Dr. Alfred J. Garrett, the two scientists who inspire me the most.

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iv ACKNOWLEDGMENTS I thank Dr. Alejandra Maruniak for tire less technical help, encouragement, and ideas. I thank Aissa Doumbouya also for he r 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 fa rm 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 alligato r blood. I thank Dr. Goerning, David Hoel, and Dr. Sandra Allan for lending collecting e quipment. I thank my advisors, Dr. James Maruniak, Dr. Jerry Butler, and Dr. Ellio t Jacobson, for their guidance and support without which the project would not have been possible.

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v TABLE OF CONTENTS page ACKNOWLEDGMENTS.................................................................................................iv LIST OF TABLES............................................................................................................vii LIST OF FIGURES.........................................................................................................viii ABSTRACT....................................................................................................................... ix CHAPTER 1 INTRODUCTION........................................................................................................1 WNV in Farmed Alligators..........................................................................................6 Mosquitoes as Vectors of WNV on Alligator Farms....................................................9 Blood Meal Identification....................................................................................10 Screening Mosquitoes for WNV.........................................................................16 2 METHODS AND MATERIALS...............................................................................20 Mosquito Collecting...................................................................................................20 Blood Meal Identification...........................................................................................26 Virus Detection...........................................................................................................31 3 RESULTS...................................................................................................................36 Mosquito Collecting...................................................................................................36 Blood Meal Identification...........................................................................................40 Virus Detection...........................................................................................................46 4 DISCUSSION.............................................................................................................48 Blood Meal Identification...........................................................................................48 Virus Detection...........................................................................................................51 Vector Incrimination...................................................................................................54 Mosquito Control.................................................................................................57 Alternative Vertebrate Reservoirs.......................................................................58 5 CONCLUSIONS AND AREAS FOR FURTHER STUDY......................................59

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vi APPENDIX A PROTOCOL FOR QIAGEN QIAQUICK SPIN KIT, PURIFICATION OF DNA FROM AGAROSE GEL............................................................................................61 B ABI PRISMTM DYE TERMINATOR CYCLE SEQUENCING KIT, PROTOCOL FOR DNA SEQUENCING........................................................................................62 C PROTOCOL FOR PGEM -T VECTOR LIGATION KIT,......................................64 D PROTOCOL FOR QIAPREP SPIN MI NIPREP KIT, EXTRACTION OF PLASMID...................................................................................................................65 E PROTOCOL FOR RNA EXTRACTION FROM MOSQUITO POOL USING TRIZOL LS (GIBCO)................................................................................................66 F SEQUENCES OF PCR PRODUCTS USED TO IDENTIFY VERTEBRATE HOST ORIGIN OF MOSQUITO BLOOD MEALS.................................................67 LIST OF REFERENCES...................................................................................................72 BIOGRAPHICAL SKETCH.............................................................................................83

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vii LIST OF TABLES Table page 2-1 Primers sets in PCR used to amplif y 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 boxe s 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 de termined by sequencing the PCR product, and information about collection date and locat ion on farm of the mosquito sample....45

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viii LIST OF FIGURES Figure page 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 Map 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 ca ptured in CDC light traps set outside of alligator pens versus inside of pe ns (for collecting trips 1,2, and 4)........................38 3-3 Products from PCR amplification of mo squito samples with alligator-specific primers......................................................................................................................43 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 scr eening primer set (l anes 2-7) and WNV confirmation set (lanes 8-13)....................................................................................47

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ix Abstract of Thesis Presen ted 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 MOSQUI TO VECTORS OF WEST NILE VIRUS ON A FLORIDA ALLIGATOR FARM By Sandra C. Garrett December 2005 Chair: James Maruniak Major Department: Entomology and Nematology Over the past several years, alligator farm s in Florida, Georgia, and Louisiana have experienced sudden die-offs of j uvenile and hatchling alligators ( Alligator mississippiensis ). These events occurred in the fall a nd tended to last two or three weeks. Histologic findings, virus culture and RT-PCR evidence all sugge st 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 identi fication 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 Fl orida 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 bl ood 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 screen ed for WNV by inoculat ion onto Vero cells

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x and by RT-PCR with WNV-specific primers sets A total of 4484 mosquitoes (sixteen different species and ten genera) were coll ected, 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 ar e likely vectors of W NV on alligator farms.

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1 CHAPTER 1 INTRODUCTION 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 roughl y spherical in shape and about 50 nm in diameter. WNV infects a large range of vertebrates as we ll as invertebrate vectors, most notably mosquitoes (Diptera: Cu licidae) (Brinton, 2002). West Nile virus was first isolated in 1937 in Uganda, from the blood of a woman suffering mild febrile illness (Smithburn et al., 1940, as cited by Hubalek and Halouzka, 1999), and records show that it was present a nd infecting humans, bi rds, and mosquitoes in Egypt in the 1950s (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 Sout heast Asia and Australia. With this large range, WNV is the most widespread flaviv irus, although before 1999 it had not been reported in the Americas. It has been is olated 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 Culex pipiens Linnaeus in Europe (Hubalek and Halouzka, 1999). The virus is maintained in bird populations and spr ead with migrations (Rappole et al., 2000). Vertical transmission in mosquitoes has been detected and may contribute to maintenance

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2 of the virus (Miller et al., 2000). In Eu rope, transmission to humans occurs during summer months (June to September) when mo squito vectors are most active (Hubalek and Halouzka, 1999). Each year in South Africa, there were s poradic cases of WN viral disease (WNVD) often with mild illness. Two epidemics, one in 1974 a nd the other in 1984, marked a change in that normal activity. These epidem ics may have been due to unusually high summer rains, which favored vector breed ing 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 involve d more serious illness, and only one meningoencephalitis (Jupp, 2001). In the late summer and fall of 1996, th ere was a major epidemic of WNVD in southeastern Romania with the highest cl inical incidence in the urban center of Bucharest. WNV had been recorded in the area (by seroprevalence evidence) since the 1960s. 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 bi rds had antibodies agai nst WNV. Of about 6000 Culex pipiens pipiens L. aspirated from man-made st ructures around Bucharest, one was found positive for WNV, and the strain app eared to be most cl osely related to WNV strains from sub-Saharan Africa. Among the factors that may have facilitated this epidemic are the naivety of the populati on, the availability of flooded man-made

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3 structures for mosquito breeding, and the summ er 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 ye ars) were found (Campbell et al., 2001). After the 1996 Romania outbreak, other ep idemics of WNV-induced CNS disease were reported in humans (including those in the United States, 19992004) (Lanciotti et al., 2002). The large Romanian epidemic w ould turn out to be only a part of an increasing trend of human and animal W NV 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 reporte d in France, notably in an area with colonies of migratory birds and plentiful mo squito breeding habitat (Murgue et al., 2001) and during the fall of 2003 an outbreak cau sed disease in horses in Morocco (Schuffeneker et al., 2005). In 2000, an ep idemic of WNV in Israel led to 326 hospitalizations and 33 deaths. Severe cases were mostly in the el derly and involved the CNS (Chowers et al., 2001). A study by Lancio tti et al. (2002) i ndicated 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 1960s (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

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4 in the Negev Desert in southern Israel we re found to be seropositive for WNV, though no deaths of crocodiles have been reported ev en 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-inf ect biting mosquitoes, thus demonstrating that they can be amplif ication 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 a nd 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 Caribb ean (Quirin et al., 2004), Me xico, 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 de pendable 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 st rain of WNV isolated in New York in 1999 is more

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5 pathogenic to crows than Old World strains from Australia and Kenya (Brault et al., 2004). Sixty species of mosquito have been f ound 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 laborato ry 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 inte rest because some species are reptile-feeders). The impact of WNV on North American reptil es has not been examined as closely as that of birds and horses, and maybe there has been little impact overall. Co mmon garter snakes ( Thamnophis sirtalis sirtalis (Linnaeus)) and red-ear sliders ( Trachemys scripta elegans (Wied-NeuWied)) did not develop detectable vi remia 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

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6 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 dieoffs 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 Tr anscriptase Polymerase Chain Reaction (RTPCR) all suggest that the deaths were cause d, 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 va lue 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 No rth America (Florida Fish and Wildlife Conservation Commission (FFWCC) re port; Lane and King, 1989). About 1,500,000 crocodilian hides are traded per year with Fl orida, Texas, and L ouisiana producing about 45,000 of that total, including hi des from wild caught alligators. In 2003, Florida farms produced 22,527 alligators at a value of a bout $3.3 million (FFWCC report). Alligators are usually kept in temperatur e-controlled (ideally about 86 F, 30 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 repor ted on two die-offs occurring during the fall of 2001 and 2002 at a farm in southern Georgi a. 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,

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7 kidney, liver, spleen, pancreas, adrenal gl and, gallbladder, tonsil, trachea, stomach, intestines, and reproductive tract. Tissues and blood were subjected to RT-PCR, virus isolation, and bacterial culture. The appear ance of the tissues a nd 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 in fection set in. Raw hor semeat is a part of the alligator diet and was test ed for WNV RNA. The meat that was fed to the alligators during the epizootics was positive for W NV by RT-PCR but was negative after the epizootic ended, leading the researchers to be lieve that virus in the horsemeat had caused the epizootic. Supporting this idea are experime nts 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 predat ors becoming infected with WNV after eating infected prey (G armendia et al., 2000; Austgen et al., 2004). In Florida, similar epizootics occurre d on several farms and one farm was investigated by Jacobson et al. (2005a). In 2002, an epizoo tic on a central Florida farm (named Farm A from here on) killed 300 of th e 9000 alligators at the farm. Clinical signs in the alligators included anorexia, le thargy, tremors, swimming on the side, and opisthotonus. Tissues of thr ee alligators were examined and showed signs of CNS disease and necrotizing hepatitis. Immunos taining revealed th e presence of WNV antigen in multiple tissues. There was no evidence of two other pathogens that have previously been iden tified as disease agents in Crocodilians: Mycoplasma and Chlamydia In contrast to the findings from Ge orgia, no secondary bacterial infection was apparent. Viremia in the infected alligators was greater than 105.0 PFU/ml plasma making the alligators capable of infecting mosquitoes like Cx. quinquefasciatus and Cx.

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8 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 duri ng the epizootic carried WNV-neutralizing antibodies three months later, while thos e sampled from pens where no disease was recorded were not found to have such antibod ies. This demonstrated that horizontal transmission had likely occurred inside the pe ns 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. La boratory experiments conducted by Klenk et al. (2004) confirmed th e 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 inoc ulated alligators. Viral shedding from the cloacae was detected and was suspected to be responsib le for horizontal transmission between tank mates. Two of the 29 infected alligator s 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 e nough (Jupp, 1974) to infect biting mosquitoes making them potential amplifica tion hosts of the virus.

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9 Alligator farmers in Florida are not required to report the cause of death of their alligators, so there ar e no precise records of these epizootics, their impact, epidemiology, and timing. Florida farmers are required to re port 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 ma ny conclusions based on gross annual records, it was clear in 2002 that some farms had virtually no unusual d eaths 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 th e 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 mo squito, 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 (1 957) (as reviewed in Turell et al. (2005)) and by Kilpatrick et al. (2005). A potential vector will repeated ly 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 (Kilpat rick et al., 2005). A potential vector should also be able to tr ansmit the virus efficiently as demonstrated through laboratory competence studies. Th is study intended to identify potential

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10 mosquito vectors of WNV in Florida farmed alligators by finding those mosquitoes that were numerous, and associated with (specifi cally feeding on) farmed alligators and determining if those associated mosquito spec ies 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. Observ ation of feeding mosquitoes, capture of mosquitoes in host baited traps, analysis of cytological characteri stics of blood meals, analysis of serological charac teristics of blood meals, and genetic information contained in blood cells have all been used to determ ine 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 react ion (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 reac ting 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 techni que 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 onl y for certain groups of vertebrates; there

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11 are primer sets for all mammals, all birds, a nd for different orders of birds (Cicero and Johnson, 2001; Ngo and Kramer, 2003). Seque ncing the fragment, followed by matching with known sequences in the BLAST databa se of GenBank, can confirm blood meal identifications or take the iden tification 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 27 C) (Ngo and Kramer, 2003). For these two techniques, na turally engorged females are collected from the field and the blood meal analysis is done in the la boratory. Different methods can be used to capture engorged females, and often the method chosen and the exact microhabitats sampled will depend on which mosquito spec ies 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 mosq uitoes against a screen until they can be transferred to a separate cont ainer. 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 aspi ration 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 incandes cent lights were used. Field research in Florida and Georgia has shown white light s to be attractive to (among others)

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12 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 ca tch 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 (Bidlingmayer, 1967). Resting boxes are containers designed to resemble mosquitoes natural resting places. They are often used to study host pref erences because they attract females that are seeking a dark place to remain while di gesting the blood meal and developing eggs. Resting boxes (with gray outside and red insi de) 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 leav e during the day although some entered at all times (Edman et al., 1968). A large body of work based on the differe nt methods of host identification has allowed some generalizations about the feedi ng 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 observe d. A number of different mosquito genera and species feed on reptiles and/or amphi bians (ectotherms). Some appear to feed mostly on reptiles or amphibians, or even particular or ders of ectotherms.

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13 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 eas tern United States have found that some mosquitoes will occa sionally take meals from reptiles. Ae. atlanticus, Ochlerotatus. triseriatus, and Oc. sollicitans (Walker) were found to feed on turtles, although in general mos quitoes 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 Coquillettidia perturbans (Walker) in New York. These three species were also found to feed on mammals and bird s in the same locations (Appersen et al., 2002). In Florida, Oc. infirmatus Dyar and Knab Ae. taeniorhynchus (Wiedemann), Ae. albopictus, Ae. vexans, Culiseta mel anura, Cx. territans, Cx. salinarius, and An. crucians fed on one or more of the following reptile s: 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 major ity of their meals we re from non-reptilian hosts (Irby and Apperson, 1988). Seventeen percent of the meal s 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 bl ood 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 be tter represented in traps with mammal or bird hosts (Murphey et al., 1967) Mosquitoes in the genus Deinocerites appear to be

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14 opportunistic feeders, taking meals from ma mmals, birds, amphi bians, and reptiles (Tempelis, 1975). Many of these same studies also found species that took a majority or even all of their meals from ectotherms. The Delawa re (Murphey et al., 1967) study found that Cx. territans Walker were frequently attracted to ki ng snakes, water snakes snapping turtles, and Eastern box turtles, but we re not attracted to the mamma ls 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 Flor ida 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, a nd later research in North Caro lina showed that 85% of the individuals sampled had taken their meal fr om 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 reptiles. A study in Panama (Tempelis and Galindo, 1975) examined Culex species in the Neotropical subgenus Melanocon ion (of which there are seven species in Florida) and found that four species fed mostly on lizards: Cx. egcymon Dyar (81%), Cx. tecmarsis

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15 Dyar (89%), Cx. elevator Dyar and Knab (90%) and Cx. dunni Dyar (63%) while the other Melanoconion species in the study fed mo stly 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 mo squito(es) feed on alligators at a farm would likely have the greatest chance of su ccess if they concentrated on sampling blood fed mosquitoes of the species that have alr eady been identified feed ing on reptiles. Based on previous Florida studies mentioned befo re, the three sampling techniques used (vacuum aspiration, CDC light traps, and restin g 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-spe cific primers based on work by Janke and Arnason, (1997), Ray and Densmore, (2002), an d Glenn et al. (2002). The Alligatoridaespecific primers amplify a region of mito chondrial DNA, including portions of the cytochrome b gene, and genes for transfer RNAs. The location w ithin the genome and the coding nature of the fragment amplif ied by the Crocodilian-specific primers are unknown. A group of animals that contains enough vi remic individuals to continually infect mosquitoes constitutes the rese rvoir, and the vertebrate rese rvoirs 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 bot h birds and alligators, such th at it could move virus from

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16 populations of infected birds to the alligator s. 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 mu st 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 pa rticles (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 tech niques) 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 samp le using cell culture (Kauffman et al., 2003; Lanciotti et al., 2000; Bernard et al., 2001). In this study, mos quito 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 m onkey (Vero cells) are used in WNV isolation because they show cytopathic effect s when infected by the virus, usually visible after three days (Odelola and Fabiyi, 1977). The virus binds to cells and enters by

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17 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 in to a single protein which is then cleaved by cellular and viral proteas es (Brinton, 2002). Translation of th e viral proteins is associated with the rough endoplasmic reticulum (Lee and Ng, 2004). The seven resultant nonstructural 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 cells margin in membrane vesicles, and are released by budding, individually at first and la ter 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 pr ocess 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 Fa biyi, 1977). Cells appear more rounded, with thicker, more distinct margins. They may a ppear grainy with vacuoles. As the cells die, they disconnect from the substrate. Ve ro cells are usually monitored for seven days after inoculation with mosquito homogena te (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. Se t one was used to scr een the pools for WNV RNA and the second was used to confirm any po sitive bands from the first set. The first set, WN9483 and WN9794, were based on sugge stions made by the CDC (based on work by Lanciotti). These primers amplify a 311 base-pair region within the NS5 gene, the

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18 gene which codes for the viral RNA-dependent RNA polymerase (Lanci otti et al., 1999). This polymerase is the most highly conser ved protein of West Nile virus (and of flaviviruses in general) (Bri nton, 2002). As a consequence of the conserved nature of the region, WN9483 and WN9794 should readily bind to any potential stra in of WNV. The second set of primers, WN212 and WN 1229, is based on suggestions of the CDC and work by Lanciotti et al., (2002). WN2 12 binds to a region within the gene for the viral nucleocapsid protein and WN1229 bi nds 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 b een observed among US strains of WNV up to 2003 (Ebel et al., 2004), and new strains is olated 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 ar bovirus, 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 encounter ed in the field may have lo wer titers than the minimum 105 virions, titers that may be below the detec tion limit of the techni ques 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 potential vectors.

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19 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 Infec tion Rate = MIR) is not greater than 1 in a 1000 unless the collection was made in the vici nity of transmission (as demonstrated by human, horse, or bird cases) (B ernard et al., 2001.). However, it is difficult to make a generalization. MIRs vary considerably be tween studies and surv eillance reports, and are likely influenced by the time of collecti on, the proportion of th e collection comprised by high risk species like Culex the age composition of th e collections, and other factors that are diffi cult to quantify. The objectives in this study were to id entify potential vectors of WNV on Farm A using three of the four crit eria described above. Mosqu itoes were captured around the farm to determine which species fit the following criteria: 1. Species is present around host (alligat ors) 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.

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20 CHAPTER 2 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. Un engorged mosquitoes were screened for WNV to determine if any species had a high MIR. Mosquito Collecting Four over night collecting trip s 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 C ounty, 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-power ed backpack aspirator, plastic bags for collecting samples of feed and aspirator samples, a cooler with ice to keep samples cold during transit, and aerial ne ts for sweep net collecting in the vegetation around the farm. Active collecting began mid-morning. The in terior and exterior walls of pens and other buildings were visually scanned for re sting mosquitoes. Insects were aspirated from vegetation, buildings and construction de bris. Insects were also collected from vegetation using sweep nets. Around midday 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 ope ning for outflow (MEDUSA Patent # 5,228,233

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21 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 tree s along a chain-link fence that separated the farm from adjacent property (Fig 2-1). Figure 2-1. CDC light trap set up on th e western margin of the farm. CO2 came from dry ice inside the insulated white plastic box.

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22 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 pa st two years. They were hung from support pipes close to the door, also about 1 meter from the gr ound. The weather was sunny and warm when traps were set out and when collected. Samples from sweep netting and aspirating we re transferred to plastic bags, put on ice in the cooler, and taken to the lab in Gainesville where they were stored in -70 C until processed. The CDC traps were left over ni ght. 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 70 C until processed. Trip two. A second collecting trip was ma de on September 24, 2003. Four CDC light traps were set inside f our separate pens, each of wh ich 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 freshw ater, three along the we stern 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 si de 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

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23 painted black with acrylic paint, and the inside surfaces were painted a maroon color (Fig. 2-2). Figure 2-2. A 30 cm x 30 cm x 30 cm w ooden 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 aftern oon on Sept. 25, left over night, and collected at a bout the same time the followi ng 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 step s 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 tim e 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

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24 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 th e plastic tube to trap mosquitoes, and tubes were then placed inside a cooler with ice. Trip three. The third trip was made on Oc tober 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 box es 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 regulat ors 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 ma de 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 enclosur e with the rectangula r 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 fa rm adjacent to the neighboring property. Of the eight resting boxes, two we re placed along the eastern ed ge of the farm adjacent to the lagoon, and the other six were set up along th e western edge of the farm (Fig. 2-3). Traps were collected the following after noon. The weather was clear and warm both days.

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25 In Gainesville, mosquitoes were separate d into pools of 1 50 individuals based on presence of blood meals, species (Darsie, 1998 ), trap type and numb er, 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. Sma ller green stars indicate where CDC light traps were placed. The structures indicated with brown outlines were buildings used for purposes like storag e of maintenance equipment, housing for water heaters, basins fo r wastewater, and an office. All identification and sorting wa s 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 t ubes (OPS, Petaluma, CA) with 1-2 copperclad steel beads (BB-caliber airgun shot)) and stored at C until processed. 200 20 15 Wastewater N

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26 Each pool was assigned a code name. Fo r the pools of unengorged mosquitoes, the code names started with a digit one through f our that corresponded to the collecting trip when mosquitoes were capture d. Letters were used to desi gnate each pool, and the code ended with a digit that indicat ed 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 us ed to distinguish multiple mosquitoes that were from the same species, date, and trap number. Blood Meal Identification Prior to working with field-collected mo squitoes, 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 in to 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 syst em was then inverted and put on the top the wire-mesh mosquito cages such that mosqu itoes could insert their proboscis through the mesh of their cages, through th e 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 alli gator tail meat that was sweetened with 10% sucrose sugar (Figure 2-4). The sugar was added to encourage feeding (Aissa

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27 Doumbouya, personal communication). The al ligator 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 C, and the engorged individuals were separated for later use. 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 avia n-specific and mammal-specific primers were a mosquito fed on a live chicken (feed ing that is a normal part of colony maintenance at the USDA) and a Coquillitidia perturbans captured after it had fed on the investigator. After it was established that the avian primers worked for the chicken-fed

PAGE 38

28 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 unengorged negative control mosquito were rem oved, placed separately into 1.5 mL plastic tubes, and homogenized in 250 l of buffer 1 (buffer 1 contains 0.32 M sucrose, 50 mM Tris at pH 7.25, 10 mM MgCl2, 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 NaCl, 25 mM EDTA, and 10 mM Tris at pH 7.8) to lyse the cells. Fifteen l of 0.5 M EDTA, 15 l of 20% SDS, and 8 l of proteinase K (20 mg/mL) were added and the mixture was incubated overnight in a 55 oC water bath. The following day the tubes were centrifuged at 13,000 rpm for 10 minutes and the supernatant wa s transferred to a new tube. Twenty l of RNAse (5 mg/ml) was added and the mixture was incubated at 37 oC for one hour. Following incubation, DNA was separated using phenol and chloroform followed by a second precipit ation using only chloroform. DNA was precipitated from the aqueous phase with 600 l of cold 95% ethanol followed by centrifugation (13,000 rpm at 4 oC for 10 minutes). The ethanol was then removed and the pellet was vacuum dried. The DNA was resuspended in 30 l of 10 mM Tris and stored at oC 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 sample s because it was the easiest protocol that consistently gave good final DNA concentrations.

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29 The DNA was tested to determine its vertebra te 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 desc ribed by Cicero and Johnson (2001). In addition primers for alligat ors were designed for this study based on the suggestions of Glen et al. (2002). Prim ers were made using the Primer3 program and the mitochondrial genome of A. mississippiensis from GenBank, accession number Y13113, (Janke and Arnason, 1997) (T able 2-1). This primer se t will be referred to as the alligator-specific set, a lthough they may also amplify DNA from other members of the Alligatoridae family or Crocodilian orde r. 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 closel y 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 MgCl2, primers, template DNA, and for the optimal annealing temperature. Once pos itive 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 condi tions and the thermocycle program for each of the primer sets are described in Table 2-3. The PCR products were run on 1% agarose ge ls stained with ethidium bromide. Any bands were excised from the gel, DNA wa s purified using the QI Aquick Spin kit for gel extraction (Quiagen, Valencia CA) fo llowing the handbook protocol (Appendix A),

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30 and the fragments were sequenced using th e BigDye Terminator Cycle Sequencing kit (PE Biosystems, Foster City CA) following a pr otocol modified from the kit instructions (Appendix B). Sequences were run at the Un iversity 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 produ ce 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 oC. Following incubation the vectors were prepared for transformation into Escherichia coli bacteria by heat inactivating the ligase at 65 oC for 10 min, diluting the DNA (x three) with sterile water, and sterilizing the DNA with 300 l of ether. Vectors were then tran sformed into bacteria. Five l 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 oC. The cell mixture was left for 2 min on ice, then 0.95 mL of medi um (deionized water with 2% bacto-tryptone, 0.5% bacto-yeast ex tract, and 0.05% NaCl) was added and the bacteria were set in a 37 oC 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 oC overnight, transformed colonies were selected, and transformed bacteria were grown overnight in media (4 mL LB medium with 5 mg/mL ampicillin) at 55 oC. The plasmid was removed using the QIAprepTM Spin Miniprep kit (QIAGEN, Valencia, CA) following kit instructions (Appendix D), and the insert was removed from

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31 the plasmid by digestion with EcoRI at 37 oC in a mixture containi ng 7 l water, 1 l reaction buffer, 0.3 l EcoRI, and 2 l plasmi d DNA. The insert was then sequenced as before. Table 2-1. Primers sets in PCR used to am plify DNA from differ ent vertebrate hosts. Virus Detection 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 g/mL gentimicin, and 1 g/mL amphotericin B (Fungizon)). Pl ates were kept over-night in a 37 oC incubator and used for virus isolation the following day. Mosquitoes were processed in a Biosafet y Level 3 laboratory (BSL-3 lab). Pools were homogenized for 1 minute in 1 ml of diluent (Phosphate Buffered Saline (PBS, contents: 0.8% NaCl, 0.02% KCl, 0.144% Na2HPO4, and 0.024% KH2PO4 in distilled H2O, pH of 7.4) with 4% Fetal Bovine Serum (FBS)) using a laboratory mixer. Host Primer Sequence Product size(bp) Reference Alligator Forward CGCTTCACTG CCCTACACTT 850 Current study Reverse GCTTTAGTGTTT AAGCTACGATAACTG Crocodilian Forward GATGTGGACCTTCAGGATGC 209 Yau et al (2002) Reverse CAGAGGTTCAATCCACGGTT Avian Forward GACTGTGACAAAATCCCNTTCCA 508 Cicero and Johnson (2001) Reverse GGTCTTCATCTYHGGYTTACAAGAC Mammalian Forward CGAAGC TTGATATGAAAAACCATCGTTG772 Ngo and Kramer (2003) Reverse TGTAGTTRTCWGGGTCHTCTA

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32 Following homogenization tubes were centrif uged at 13,700 rpm for 10 minutes to pellet mosquito solids. The mosquito supern atant was removed to a new tube and 200 l and 10 l were removed for use in screening. A ny remaining homogenate was frozen at oC until needed further. For the RNA extraction, the 200 l of homogenate was th en added to a tube containing 600 l of Trizol LS reagent (Life Tech nologies, Gaithersburg, MD), and the mixture was incubated for 5 minutes to inactiv ate the virus. After incubation, tubes were removed from the BSL-3, stored at oC, and later RNA was extracted as described in the Trizol manufacturers instructions (Appendix E) and was resuspended in 30 l of nuclease-free water. All RNA samples were then tested for WNV RNA using Promega Access RT-PCR System (Promega, Madison, W I) 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 polymeras e and Reverse Transcriptase, and 1 l of template for 25 l 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 RTPCR products were visualized on 1% agarose gels with ethidium bromide staining. All bands were excised from the gel, cleaned, a nd sequenced as described for the vertebrate primers (see Appendix A and B). The sa mples showing positive bands with WN9483 and WN9794 were also confirmed using a second primer set, WN212 and WN1229 (Table 2).

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33 Table 2-2. Primers sets used in RT-PCR to test for the presence of WNV RNA. WNV-specific primer sets Ten l of the mosquito supernatant was mixed with 100 l 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 inoc ulum was added. Inoculated plates were incubated for one hour at 37 oC with gentle hand rocking every ten minutes. After incubation 500 l of cell culture media was added (media: Lebovitz L-15 media, 10% fetal bovine serum, 100 U of penicillin/streptomycin, 100 g/mL gentimicin, and 1 g/mL amphotericin B (Fungizon)) to each well and plates were placed inside a plastic box with moistened paper towels and kept in a 37 oC incubator. Cells were checked daily for 7 days for cytopathic effect (CPE) us ing an inverted compound microscope with WNV CPE expected to begin on da ys 3 or 4 post-inoculation. Samples that showed signs of bacterial c ontamination were recorded as such, and the homogenate for that sample was thawed a nd the inoculation was repeated with a new well of Vero cells. In these cases, it was a ssumed that the homogenate was the source of the contamination and for the new well, th e inoculum was removed after the one-hour Primer* Sequence Amplicon size (bp) Annealing region Confirmation WN212 TTGTGTTGGCTCTCTTGGCGT TCTT1071 Capsid protein Set WN1229 GGGTCAGCACGTTTGTCATTG Envelop glycoprotein Screening WN9483 CACCTACGCCCTAAACACTTTCACC311 NS5: RNAdependent RNA polymerase Set WN9794 GGAACCTGCTGCCAATCATACCATC NS5: RNAdependent RNA polymerase

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34 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 Thermocycle Mammalian Buffer 1X 93 for 3 min dNTP's 0.2 mM each 94 for 30 sec each primer 5 pmol/rxn 50 for 30 sec MgCl2 4 mM 72 for 1 min 30 sec Taq polymerase 1 unit/rxn Goto 2 45 times template DNA 1 l 72 for 3 min Avian Buffer 1X 93 for 3 min dNTP's 0.2 mM 94 for 30 sec each primer 15 pmol/rxn 50 for 30 sec MgCl2 2.0 mM 72 for 1 min 30 sec Taq polymerase 1 unit/rxn Goto 2 34 times template DNA 1 l 72 for 3 min Crocodilian Buffer 1X 94 for 3 min dNTP's 0.2 mM 94 for 30 sec each primer 10 pmol/rxn 53 for 30 sec MgCl2 2.5 mM 72 for 30 sec Taq polymerase 1 unit/rxn Goto 2 40 times template DNA 2 l 10 for ever Alligator Buffer 1X 93 for 3 min dNTP's 0.2 mM 94 for 30 sec each primer 10 pmol/rxn 55 for 30 sec MgCl2 2 mM 72 for 1 min 30 sec Taq polymerase 0.2 l/rxn Goto 2 34 times template DNA 1 l 72 for 3 min WNV Buffer 1X 48 for 5 min dNTP's 0.2 mM 94 for 5 min each primer 0.4 pmol/l 95 for 30 sec MgSO4 2 mM 58 for 45 sec Taq polymerase 1 unit/rxn 68 for 2 min Reverse transcriptase 1 unit/rxn Goto 3 39 times Template RNA 1 l 68 for 10 min incubation period in an attempt to remove the contaminated homogenate after the inoculum had inoculated.

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35 Assuming one infective mosqu ito 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 Multip licity of Infection (MO I) of approximately 0.63. Three positive controls were conducted simu ltaneously with samples. Each control used a previously frozen Fl orida isolate of West Nile virus, WNV-FL01-JC2-3C2P2, which had been at a titer of approximately 107.5 TCID50/mL before freezing. In one control well, 50 l of the virus was added directly. For the other two controls, 2 and 100 l 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 inoculat ion contained approximately 102.5 and 104.5 virions respectively. This gave an MOI of a bout 0.0063 and 0.63 respectively. For the direct inoculation of 50 l, the MOI was about 32.

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36 CHAPTER 3 RESULTS Mosquito Collecting During collecting no mosquitoes were seen re sting 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 en tered 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 sufficien t for mosquitoes to enter and exit. Collection bags on several of the CDC trap s that were hung inside alligator pens were apparently torn down by the alligators some time during the night. These samples could not be recovered. Taki ng into account the losses due to alligator interference and one disturbed collection bag there was a tota l of 20 trap nights for the CDC light traps and 24 trap nights for the resting boxe s 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. cruci ans, Mansonia dyari, Ma. titillans, Cx.. nigripalpus, Cx. erraticus, Cx. quinquefasciatus, Uranotaenia sapphirina, Ur. lowii, Psorophora columbiae, Ps. ferox Coquillettidia perturbans, Wyeomyia vanduzeei Culiseta. melanura Ae. albopictus and Oc. infirmatus (Fig 3-1). The numbers of mosquitoes collected va ried from one trip to the next.

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37 1172 1170 737 535 273 272 142 115 23 20 10 8 2 2 1 1 0200400600800100012001400 Anopheles quadrimaculatus [15] Mansonia dyari [9] Anopheles crucians [1] Culex nigripalpus [5] Culex erraticus [3] Mansonia titillans [4] Uranotaenia sapphirina Psorophora columbiae Ochlerotatus infirmatus Coquillettidia perturbans Aedes albopictus Uranotaenia lowii Culiseta melanura Culex quinquefasciatus Psorophora ferox Wyeomyia vanduzeei 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 over all 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 aspiration. 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).

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38 0 100 200 300 400 500 600 700 800 900An. quadrimaculatus An c r u c i a n s Ma. d y ari M a. ti till a ns P s. colu m biae Ur. s a pp hir ina U r lowii A e. a lo b opictu s Cx. erra t icus C x n i g r ipa lpu s C q perturbans Ps f er ox Oc. infir m at u s Cs m ela nu r a Cx quinq u efasciatu s Wy. v a n du ze eiMosquito species outside pens inside pens Figure 3-2. Portions of each mosquito species captured in CDC light traps set outside of alligator pens versus inside of pe ns (for collecting trips 1,2, and 4). Trip one. A total of 2665 mosquitoes was co llected from CDC traps (Table 3-1) during trip one, and this represented more than half of the total co llected during the four trips. Trip two A total of 150 mosquito es 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 obt ained from that CDC trap. Trip three. A total of 628 mosquitoes was co llected (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.

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39 Table 3-1. Mosquitoes captured from CDC li ght 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 3433184251781 An. crucians 00179364543 Ma. dyari 46[1] 6[1] 323[3]338338 Ma. titillans 63[2]042327 Ps. columbiae 003962101 Ur. sapphirina 00283967 Ur. lowii 00314 Ae. alobopictus 00066 Cx. erraticus 705684147 Cx. nigripalpus 19 [1] 54[1] 122[1]19 Cq. perturbans 20518 Ps. ferox 00011 Oc. infirmatus 00088 Cs. melanura 00000 Cx. quinquefasciatus 00000 Wy. vanduzeei 00000 TOTAL 4801087513002665 Table 3-2. Mosquitoes collected in resti ng boxes and CDC light tr aps during the second collecting trip to Farm A. Number s in [ ] indicate blood-fed mosquitoes. CDC inside pen trap 1trap 2 trap 3 Resting box TOTAL An. quadrimaculatus 09253 37 An. crucians 008 3[1] 8 Ma. dyari 27220 31 Ma. titillans 4 [2]400 44 Ps. columbiae 0000 0 Ur. sapphirina 0000 0 Ur. lowii 0000 0 Ae. alobopictus 0000 0 Cx. erraticus 4 [1]0 [2] 4 Cx. nigripalpus 12330 18 Cq. perturbans 0000 0 Ps. ferox 0000 0 Oc. infirmatus 0000 0 Cs. melanura 0000 0 Cx. quinquefasciatus 0000 0 Wy. vanduzeei 0000 0 TOTAL 2222988 150

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40 Table 3-3. Mosquitoes collect ed from CDC light traps and resting boxes during the third collecting trip to Farm A. Numbers in [ ] indicate blood fed individuals. CDC trap Resting TOTAL trap 1 trap 2 trap 3 trap 4 trap 5 trap 6 box An. quadrimaculatus 793667109 [9] 183 An. crucians 29300000 41 Ma. dyari 405834 39[3] 0 108 Ma. titillans 053212460 68 Ps. columbiae 0230000 5 Ur. sapphirina 59350000 49 Ur. lowii 0020000 2 Ae. alobopictus 0010000 1 Cx. erraticus 910500010 70 Cx. nigripalpus 2019400058 92 Cq. perturbans 1000010 2 Ps. ferox 0000000 0 Oc. infirmatus 0070000 7 Cs. melanura 0000000 0 Cx. quinquefasciatus 0000000 0 Wy. vanduzeei 0000000 0 TOTAL 4863265112320117 628 Trip four. On the last collecting trip 1041 mos quitoes were collected (Table 3-4). The battery failed on one of the CDC trap s although some mos quitoes were still collected. Numbers and proportions of different mosqu ito species varied from one collecting trip to the next, however, statistical analys is 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 four Ma. dyari ). Of

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41 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 li ght traps and resting boxes on the fourth collecting trip to Farm A. Numbers in [ ] indicate bloodfed individuals. CDC inside pen CDC outside pen Resting trap 1 trap 2 trap 3 trap 1 trap 2 trap 3 trap 4 box TOTAL An. quadrimaculatus 31802051513 25[6]171 An. crucians 100285441153142 Ma. dyari 02215210230300318 Ma. titillans 22102121129068 Ps. columbiae 000116109 Ur. sapphirina 00031634026 Ur. lowii 000010102 Ae. alobopictus 000101103 Cx. erraticus 10092962249 Cx. nigripalpus 00043 148[1]27101229 Cq. perturbans 1007200010 Ps. ferox 000000000 Oc. infirmatus 000600208 Cs. melanura 000101103 Cx. quinquefasciatus 000100012 Wy. vanduzeei 000001001 TOTAL 2830428440517979321041 Cx. nigripalpus and one Ma. titillans (the Ma. titillans had a second, very faint band of approximately the correct size). Sequences fr om 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 database. Of the 37 mosquitoes that had apparent blood meals, 14 reacted with one of the mitochondrial primer sets (mammal, alligato r, bird) giving an identification rate of 37.8%. Seven individuals (representing thr ee species) were positive for alligator DNA (Fig. 3-3), six (three species) were pos itive for mammalian DNA, and one individual was

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42 positive for avian DNA (Fig. 3-4). For the a lligator primer set, the seven positives included the six samples that were positive fo r 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 th e 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 identifica tion 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 fu rther (Table 3-5). The GenBank E value for all of the matches wa s 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 consiste ntly amplified mosquito DNA in the negative control (an un-engorged mosquito). Repetition of the DNA extraction from a new un-engorged mosquito reduced the possibi lity 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 entr ies, however one 64 bp portion of the region

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43 Figure 3-3. Products from PCR am plification of mosquito samp les 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 cl osely with chromosomal DNA (from partial mRNA) from An. gambiae Within this 64 bp portion there were 57 bases shared with the databases An. gambiae sequence and the E-value assigned for this match was 3e-10. 12345678 9101112131415 123456789101112131415 a b

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44 12345678910 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 ma mmal 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 clone s yielded sequences for which there were no close matches on the database. 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

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45 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 sample. Primer set Mosquito Vertebrate host Collection date Trap location Species ID number Mammal Ma. dyari 1MBF4 Equus callabus (horse)a Sept. 12 Inside pen 1MBF1(2) Sus scrofa (wild boar)b Sept. 12 In trees on West margin An. quadrimaculatus 3AnBF Sus scrofa Oct. 17 Resting box 3An3BF Sus scrofa Oct. 17 Resting box 3An8BF *** Oct. 17 Resting box Cx. nigripalpus 4CuNBF Procyon lotor (raccoon)c Oct. 24 Eastern margin on tree Bird Cx. nigripalpus 1CuNBF2 Meleagris gallopavo (turkey)d Sept. 12 In trees on West margin Alligator Cx. erraticus 2ABF Alligator mississippiensiseSept. 26 Resting box 2CBF Alligator mississippiensis Sept. 26 Inside pen Ma. dyari 2DBF Alligator mississippiensis Sept. 26 Inside pen 1MBF3 Alligator mississippiensis Sept. 12 Inside pen 3AABF Alligator mississippiensis Oct. 17 CDC 8 3AAABF Alligator mississippiensis Oct. 17 CDC 8 Ma. titillans 2MaTBF2 Alligator mississippiensis Sept. 26 Inside pen References for identification of sequences: a = GenBank accession # D32190 (Chikuni, 1994); b = AY237534 (Alves et al., 2003); c = U12853 (Lento et al., 1995); d = L08381 (Kornegay et al., 1993); e = AF318572 (Glen et al., 2002), *** = specific host not identified.

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46 Virus Detection 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 pos itive controls, all cell s appeared infected by day seven and many had detached from the s ubstrate and were floating in the media. No CPE was apparent in the well that r eceived 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 of 0/4447. Two of the wells had bacterial contaminati on that became apparent after two days. When these samples were repeated (as desc ribed above) one had bacterial contamination again, and the other did not. The cells in many of the wells that had b een 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 cel ls 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 l of cysteine to the diluent prior to homogeniza tion appeared to prev ent this effect in subsequent pools of 50 Ma. dyari

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47 Detection of viral RNA. Both of the positive contro ls that were tested for WNV RNA showed clear, bright positive bands w ith 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 W NV screening primer set (lanes 2-7) and WNV confirmation set (lanes 8-13). WNV positive cont rols are in lanes 2 and 8. Lane 1 contains a 1 kb molecular we ight 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 s ubsequent 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 sens itivity, including those with titers well below 105. No field-collected mosquito pools were positive for WNV RNA.

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48 CHAPTER 4 DISCUSSION Blood Meal Identification While encouraging, the results from the Cr ocodilian-specific primers (Yau et al. 2002), did not seem conclusive, because the prim ers were not specific. Because there are mitochondrial sequences for many more differe nt organisms published and available on the GenBank database, it is easier to iden tify 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 Crocodilianspecific primers amplified, therefore it was difficult to determine whether or not the homology found between the mosquito samp les 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 alligat or mitochrondrial DNA from a well-studied region, the cytochrome b gene. It appeared that the mammalian-sp ecific primers sometimes amplified chromosomal DNA from the mosquito. The ba nds in the negative control (where only mosquito DNA was present) shared some ho mology with a mosquito and even where there was vertebrate host DNA present, seque ncing results (overlapping peaks and cloned fragments whose sequences were not from a known region of vertebrate DNA) showed that the primers anneal ed in multiple places apparently on both host and mosquito DNA. Of the three mosquito species f ound to feed on alligators, only one, Cx. erraticus has been reported to feed on reptiles a nd in general has been identified as an

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49 opportunistic feeder taking m eals from mammals, birds, a nd reptiles or amphibians (Robertson et al., 1993; Irby and Appe rson, 1988). Species in the genus Mansonia are in general considered mammal and some times bi rd feeders (Edman, 1971). Studies of this genus in other parts of the world have f ound them attracted to or feeding on cow and human baits (Tuno et al., 2003; Khan et al ., 1997). The normal feeding habits of Ma. titillans and Ma. dyari (both of the ne otropical subgenus Mansonia ) have not received much attention probably because they do not se em 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 feedi ng 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 alligator s are captive in pens with water depths insufficient to allow the al ligators to submerge. They may present a blood source so readily available that a range of species takes advantage. The mitochondrial PCR product from the alli gator positive control was distinctly fainter than the bands from the field collected mosquitoes. This may have been due to the presence of heparin in th e positive control alligator bl ood. 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 th e concentration of heparin and the type of polymerase enzyme used. The total blood meal host id entification percentage (38 %) was lower than that of other studies (65%) where host determin ation was done using DNA probes and a PCR reaction (Ngo and Kramer, 2003; Leslie Rios personal communication). This may be

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50 due to components within the mosquito or pr ocesses during digestion that interfere with the PCR or rapidly degrade the DNA. C upp et al. (2004) found that their overall identification per centage (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 dr amatically smaller identification rate, especially consider ing that smaller blood meals (incomplete engorgements in normal sized mosquito spec ies) 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% positiv e identification rate. For this study, when Anopheline and Culicine mosquitoes are considered separatel y, the identification rates are 18.8% and 52% respectively. The id ea that differences in mosquito digestive physiology might influence the success rate of PCR-based host id entification studies warrants further study. After a blood meal is ingested it clumps inside the mosquito midgut yielding separated serum and a clot c ontaining the erythrocyt es. After that (and for the next several hours) enzymes are secret ed which begin to digest the surface of the clot. Components of the separate serum ar e absorbed and used for nutrition or egg development (Nayar and Sauerman, 1977). Na yar and Sauerman (1977) showed that in An. quadrimaculatus the mean clotting time was significan tly greater than that of five Culicine mosquitoes. The average clotting ti me (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

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51 and Cx. nigripalpus respectively. The dela y 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 f eeding (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 w ould 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 ei ght h of time may ha ve allowed greater breakdown of DNA, thus making the positive identification percentage lower in this study. Virus Detection 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 fe d on alligators). The viru s isolation results neither support nor diminish the possibility of mos quito-transmitted WNV on the alligator farm, nor can they help in incriminating any one of the three alligator-feed ing species found. In a study done in New York, WNV isolations tende d to occur in the vi cinity of greater transmission such that the authors made the following generalization: the greatest number of human cases and dead crows co rresponded 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

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52 2.36/1000, and no human cases and few dead cr ows corresponded to an MIR of 0 to 0.86/1000 (White et al., 2001). Some studies have had similar MIRs (Rutledge et al., 2003; Reisen et al., 2004), and others have ha d much lower MIRs, 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 mosqu itoes collected from gravid traps, thus biasing the results towards a higher MIR by te sting only older females. In a Florida study, an MIR of 1.2 in 1000 was found, and resu lts 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 sp ecies were positive for WNV, and a single Cx. nigripalpus was responsible for infecting a sentinel chicken. This species was present around and feeding on th e host (chicken) and was most frequently infected with WNV, showing that these criteria can be help ful in identifying possi ble vectors. This study also found that the numb er of WNV positive mosquitoes around the chickens was greater than the number of transmissions to ch ickens. Thus an infected mosquito pool is not a sure way to identify the species res ponsible for transmissi on. Alternatively, the study found that there were no infected mosqu itoes at a site where a horse had become infected a month prior to co llecting. In this case, th e 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

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53 situation at the alligator farm. On the alli gator farm the mosquito collections were only about 18% Culex (808 out of 4484). The collec tion 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 species. Based on the above information, it app ears 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 mos quito collections were predominately nonCulex mosquitoes which are less likely to vect or and be infected with WNV or 3) the collecting began during the epidemic of dis ease and after transmission had occurred. With the last explanation, it is possible th at the transmission of WNV had taken place about two weeks before the majority of the alli gators 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 severa l alligators during a brief period of intense WNV activity. These alligators then develope d viremia and infected their tank mates, and a week later the situation progressed to what was obser ved during the fi rst collecting trip: multiple alligators sick and dying from WNV-like disease. In this scenario the WNV transmission was very focal (both tempor ally and spatially) and had subsided by the time mosquito collecting had begun. If this were th e case, the practice of preepizootic surveillance would not only help predict when an epizootic might start, but would also be important for identifyi ng the mosquito species involved.

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54 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 surround ing 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 Services maps (http://westn ilemaps.usgs.gov/index.html, created with information from CDC) give information abou t 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 ch icken flocks, one osprey ( Pandion haliaetus Linnaeus) positive for antibodies to WNV, and th ree cases of WNV reported by veterinarians. However, there were no isolations of virus from mo squito pools in the county that year. This surveillance information indicates that ther e was mosquito transmission of WNV in the county during the time of the epizootic on th e alligator farm, even though no isolations were made from mosquitoes. The possibili ty remains that the WNV on Farm A was mosquito transmitted even though no positive mosquitoes were detected. Vector Incrimination The information gained in this study can be considered in the cont ext of the criteria established by Reeves (1954) and expanded on by Kilpatrick et al. (2005) and used to identify potential mosquito vectors. Becau se 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

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55 these species are relatively numerous around th e farm (6%, 6%, and 26% of the total catch, respectively). Other studies have show n that they are in gr eatest abundance during the season when the alligator epidemics occur (Bidlingmayer, 1968; Zhong et al., 2003). Information from other studies and surveillanc e reports will be necessary to answer the remaining questions about these potential vect ors: 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 b een 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/mosquito Species.htm). They have also been found infected with other ar boviruses in the United States such as Eastern Equine Encephalomyelitis virus (EEE, Togaviridae: Alphavirus) (Wozniak et al., 2001; Cupp et al., 2003) and St. Louis Encelpha litis 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

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56 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 WNVpositive mosquitoes. As of July 2005, no WN V vector competence studies have been published for Cx. erraticus or for any other North Ameri can Melanoconion. The studies that have been done i ndicate 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 po tential 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 southe rn Texas, and in Mi ssissippi (Darsie and Ward, 1981; Goddard and Harrison, 2005). Thes e species are also found in south and Central America, where Ma. titillans is likely involved in the transmission of Venezuelan Equine Encephalomyelitis virus, an alphavi rus (Mendez et al., 2001; Turell et al., 2000) and Ma. dyari is a maintenance vector of SLE (Gor gas 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 (Traor-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 th eir 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/mosqu itoSpecies.htm). Based on this input,

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57 Cx. erraticus seems the most likely potential vector of WNV on Farm A, although Mansonia are at least nuisance species. Mosquito Control 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 th ere are aquatic plants (Kle in 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 wate r. Especially with the Mansonia mosquito populations are closely related to the availab ility 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.), for Ma. titillans (Slaff and Haefner, 1985). These plants can be controlled with herbicides (Slaff and Haefner, 1985). The pr acticality of serious water plant control in this case would need to be i nvestigated. 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 alligat ors are accustomed to receiving food from humans. Alternatively, contro l could be aimed at adult mosquitoes. For all three species, adulticides could be applied for severa l months in the late summer and early fall when populations are at their peak (Sla ff and Haefner, 1985; Bidlingmayer, 1968; Roberson et al., 1993; Zhong et al., 2003).

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58 Alternative Vertebrate Reservoirs The other blood meal hosts (pigs, a horse, a turkey, and a racc oon) identified on the farm were probably not involved in maintenan ce or amplification of WNV. In a study where three-week-old turkeys were inocul ated subcutaneously with NY99 WNV, none displayed illness, and viremia, while dete ctable, was very low. From the results researchers concluded that tu rkeys 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 Yo rk 99 strain of West Nile virus, and while adult pigs seroconverted, most of the animal s did not have sufficient viremia to allow reisolation of the virus from serum. Weanli ng pigs developed viremi a less than or equal to 103.1 PFU/mL. No signs of clinical diseas e were observed (Teehee et al., 2005). The low viremia found in horses and their failure to infect mosquitoes in experiments also makes them unlikely amplificati on hosts (Bunning et al., 2002).

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59 CHAPTER 5 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 speci es 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 comp etence 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 inform ative to study mosquitoes responses to potentially attractive or repell ent compounds associated with the alligators. In one trap placed inside of an alligator pen there we re 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 su ch 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 th at were found inside of the alligator pens had blood meals from a lligators. This may mean that there are

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60 repellents or missing attractants (or other stimuli) that pr event feeding in many of the mosquitoes that were initially attracted to th e alligator pens. Additional collections that are carried out more regularly and systema tically may provide more information about what mosquitoes are attracted to the alligators, and whet her 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 olfactom eter (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 alligato r pens than others. Investigating these differences could not only help predict vectors of WNV in al ligators, but could also be useful in the continued effort to de scribe mosquito host seeking behavior.

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61 APPENDIX A PROTOCOL FOR QIAGEN QIAQUICK SPIN KIT, PURIFICATION OF DNA FROM AGAROSE GEL (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 co ntaining the PCR product and place in a 1.5 mL microfuge tube. 3. Add 3 l of buffer QG for each 1 mg of gel. 4. Incubate at 50 C for 10 min (tapping tube to mix every 2 minutes) to dissolve gel. 5. Once the gel is dissolved add 1 l 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 clea n, labeled microcentrifuge tube. 14. Add 30 l of buffer EB to the center of the co lumn 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 4 C until needed for sequencing or other purposes.

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62 APPENDIX B ABI PRISMTM DYE TERMINATOR CYCLE SEQUENCING KIT, PROTOCOL FOR DNA SEQUENCING (Modified from PERKIN ELMER PROTOCOL, revised July, 2005) NOTE: Keep all reagents on ice 1. Estimate concentration of template DNA by running 5 l in an agarose gel and comparing intensity to known concentration of molecular weight ladder. 2. Calibrate the thermocycler. 3. Dilute templates to recommende d concentration (See table below). 4. Remove Terminator Ready Reacti on Mix and ICBR dNTP mix from freezer and thaw on ice. 5. For each reaction, mix the follo wing reagents in a microfuge tube: REAGENT QUANTITY Terminator Ready Reaction Mix 2.0 L ICBR dNTP mix 2.0 L Template 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) ___ L Deionized Water Bring final volume to 10.0 L Final Reaction Volume 10.0 L 6. Gently pipette to mix the reagents. 7. Place tubes in thermocycler a nd start thermocycle (See below). Thermocycle program : 1. Ramp to 96 o C and hold for 30 s (denaturation) Ramp to 50 o C and hold for 15 s (primer annealing) Ramp to 60 o C and hold for 4 min (product extension) 2. Repeat step 1. For 25 cycles

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63 3. Ramp to 4 o C and hold. 8. Remove tubes from the thermocycler. 9. For each reaction, prepare a 1. 5 mL microfuge tube by adding: 1.0 L 3M Sodium acteate, pH 4.6 30.0 L 95% cold ethanol 10. Transfer the sample to the prepared microfuge tube and place on ice for 10 min. 11. Centrifuge for 15 min (13,000 rpm). 12. Carefully and completely remove et hanol solution, without disturbing the pellet of DNA. 13. Rinse the pellet with 250 L 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 DNA. 16. Dry under vacuum. 17. Store in dark freezer until ready to read.

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64 APPENDIX C 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/ l) of the PCR product to be cloned by comparing the intensity of the band in a gel to the intens ity 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 tube. 5. Vortex the 2X rapid ligation buffer before use. 6. Combine the following in a 0.5 ml microfuge tube: 5 l 2X rapid ligation buffer 1 l pGEM -T vector x l PCR product 1 l T4 DNA ligase deionized water to a final volume of 10 l 7. Mix the reagents. 8. Incubate the mixture for 1 h at room temperature or overnight at 4 C if less than the recommended amount (see equation in step 2) of PCR product was added.

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65 APPENDIX D PROTOCOL FOR QIAPREP SPIN MINIPR EP KIT, EXTRACTION OF PLASMID (modified from QIAprep Miniprep Handbook, S. C. Garrett, July 2005) 1. Centrifuge three to five ml of bact eria from overnight bacterial culture. 2. Add RNAase A to Buffer P1. 3. Resuspend pelleted bacterial cells in 250 l of buffer PI and transfer to a 1.5 ml plastic microcentrifuge tube. 4. Add 250 l of buffer P2 and mix by gently inverting 4-6 times. 5. Add 350 l 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, di scard 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 centri fuge for an additional min at 13,000. 13. Transfer the column to a cl ean 1.5 microcentrifuge tube. 14. Add 50 l of buffer EB to the center of the colu mn and let the buffer soak in for one min. 15. Centrifuge for one mi n at 13,000 to elute DNA.

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66 APPENDIX E PROTOCOL FOR RNA EXTRACTION FROM MOSQUITO POOL USING TRIZOL LS (GIBCO) (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 1ml PBS medium with 4% Fetal Bovine Serum. 2. Centrifuge at 15,000 rpm for 10 min. 3. Remove 200 l of supernatant into a ne w tube and save the rest of the supernatant at -80 C. 4. To the 200 l, add 600 l Trizol LS, then mix and incuba te at room temperature for five min. 5. Add 160 l of chloroform, mix by inverting fo r 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 l 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 l of RNAse-free water.

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67 APPENDIX F 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 # AY237534): ATCCGAAAATCACACCCACTAATAAAAATTATCAACAACGCATTCATTGACCTCCCAGC CCCCTCAAACATCTCATCATGATGAAACTTCGGTTCCCTCTTAGGCATCTGCCTAATCT TGCAAATCCTAACAGGCCTGTTCTTAGCAATACATTACACATCAGACACAACAACAGCT TTCTCATCAGTTACACACATTTGTCGAGACGTAAATTACGGATGAGTTATTCGCTATCT ACATGCAAACGGAGCATCCATATTCTTTATTTGCCTATTCATCCACGTAGGCCGAGGTC TATACTACGGATCCTATATATTCCTAGAAACATGAAACATTGGAGTAGTCCTACTATTT ACCGTTATAGCAACAGCCTTCATAGGCTACGTCCTGCCCTGAGGACAAATATCATTCTG AGGAGCTACGGTCATCACAAATCTACTATCAGCTATCCCTTATATCGGAACAGACCTCG TAGAATGAATCTGAGGGGGCTTTTCCGTCGACAAAGCAACCCTCACACGATTCTTCGCC TTCCACTTTATCCTGCCATTCATCATTACCGCCCTCGCAGCCGTACAT Ma. dyari (1MBF4); 100% identity with Equus caballus (horse) (accession # D32190): TGGAATGGGATTTTGTCCATATCGGATGGGATTCCTGAGGGGTTGTTAGATCCTGTTTC GTGAAGAAATAGTAAATGTACGACTACCAGGGCTGTGATGATGAAGGGTAGGATGAAGT GGAAAGCAAAAAATCGGGTAAGGGTGGCTTTGTCTACTGAGAATCCACCTCAGATTCAC TCGACGAGGGTAGTACCGATGTAGGGAATTGCTGATAGGAGGTTCGTGATGACTGTTGC TCCTCAAAAGGATATTTGGCCTCATGGTAGGACATAGCCCATGAATGCTGTAGCTATAA CTGTGAAAAGTAGGATGATTCCAATGTTTCATGTCTCTAGGAATGTGTAAGAGCCGTAG TAGAGGCCGCGTCCTACGTGAATGAAGAGGCAGATAAAAAATATTGATGCTCCGTTGGC ATGGAGGTAGCGAATAATTCATCCGTAGTTAACGTCTCGGCAGATGTGAGTGACGGATG AGAAGGCAGTTGTCGTGTCTGATGTGTAGTGTATGGCTAGGAATAGGCC An. quadrimaculatus (3AnBF), clone 2; 100% identity with Sus scrofa (pig) (accession # AY237534): CCACTAATAAAAATTATCAACAACGCATTCATTGACCTCCCAGCCCCCTCAAACATCTC ATCATGATGAAACTTCGGTTCCCTCTTAGGCATCTGCCTAATCTTGCAAATCCTAACAG GCCTGTTCTTAGCAATACATTACACATCAGACACAACAACAGCTTTCTCATCAGTTACA CACATTTGTCGAGACGTAAATTACGGATGAGTTATTCGCTATCTACATGCAAACGGAGC ATCCATATTCTTTATTTGCCTATTCATCCACGTAGGCCGAGGTCTATACTACGGATCCT ATATATTCCTAGAAACATGAAACATTGGAGTAGTCCTACTATTTACCGTTATAGCAACA GCCTTCATAGGCTACGTCCTGCCCTGAGGACAAATATCATTCTGAGGAGCTACGGTCAT CACAAATCTACTATCAGCTATCCCTTATATCGGAACAGACCTCGTAGAATGAATCTGGG

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68 GGGGCTTTTCCGTCGACAAAGCAACCCTCACACGATTCTTCGCCTTCCACTTTATCCTG CCATTCATCATTACCGCCCTCGCAGCCGTACATCTCCTATTCCTGCACGAAACCGGATC C Cx. nigripalpus (1CuNBF2), clone 3; 100% identity with Meleagris gallopavo (turkey) (accession #L08381): CTTACTCACATTAACCCTATTCTCACCTAACCTCTTAGGAGACCCAGAAAACTTTACCC CAGCAAATCCACTAGTAACCCCCCCACACATTAAACCAGAGTGATACTTTCTATTTGCC TACGCAATCCTACGCTCAATCCCAAACAAACTTGGAGGTGTCCTAGCCTTAGCAGCATC AGTACTCATTCTTCTCCTTATCCCCTTCCTTCATAAATCTAAACAACGGGCAATAACAT TCCGGCCACTCTCACAAACCTTATTCTGACTCTTAGTAGCAAACCTCCTCATCCTAACC TGAGTAGGAAGCCAACCAGTAGAACACCCATTCATCATCATTGGCCAAATAGCATCCCT TTCCTACTTCACTATCTTACTAATCCTCTTCCCCTTAATCGGAGCCCTAGAAAACAAAA TACTCAACCTCTAAGTACTCTAATAGTTTATGAAAAAC Cx. nigripalpus (4CuNBF); 98.22% identity with Procyon lotor (raccoon) cyt.b gene (accession # U12853): ATCCGAAAAACTCACCCATTAGCTAAAATCGTCAACAACTCATTCATTGATCTACCCAC CCCCTCAAACATCTCAGCATGATGAAATTTCGGCTCCCTCCTCGGAATTTGTTTGCTTC TACAGATCGCAACAGGTTTATTCTTAGCCATGCACTACACACCAGATACAGCCACAGCT TTCTCATCAGTGACCCACATTTGCCGAGATGTAAATTATGGCTGAATTATCCGATATAT ACACGCTAACGGAGCTTCTATATTCTTTATATGCCTATTCTTACACGTAGGACGAGGCT TATACTATGGCTCCTATACATTCTCTGAAACATGAAATATTG An. quadrimaculatus (3An3BF); 100% identity with Sus scrofa (accession # AY237534): ATCCGAAAATCACACCCACTAATAAAAATTATCAACAACGCATTCATTGACCTCCCAGC CCCCTCAAACATCTCATCATGATGAAACTTCGGTTCCCTCTTAGGCATCTGCCTAATCT TGCAAATCCTAACAGGCCTGTTCTTAGCAATACATTACACATCAGACACAACAACAGCT TTCTCATCAGTTACACACATTTGTCGAGACGTAAATTACGGATGAGTTATTCGCTATCT ACATGCAAACGGAGCATCCATATTCTTTATTTGCCTATTCATCCACGTAGGCCGAGGTC TATACTACGGATCCTATATATTCCTAGAAACATGAAACATTGGAGTAGTCCTACTATTT ACCGTTATAGCAACAGCCTTCATAGGCTACGTCCTGCCCTGAGGACAAATATCATTCTG AGGAGCTACGGTCATCACAAATCTACTATCAGCTATCCCTTATATCGGAACAGACCTCG TAGAATGAATCTGAGGGGGCTTTTCCGTCGACAAAGCAACCCTCACACGA 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: ACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCT GACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGC TGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAA ACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGTACTTC

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69 CTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAAT ATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAAC CAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTC TTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAAC TGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAG AGAACA Cx. erraticus (2CBF): ACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCT GACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGC TGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAA ACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGTACTTC CTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAAT ATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAAC CAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTC TTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAAC TGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAG AG Cx. erraticus (2ABF) CACCCACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAA ATGCTGACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCA CTAGCTGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCC TGAAAACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGT ACTTCCTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTA GCAATATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACA ACAACCAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACT TCCTCTTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGC CAAACTGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCT CTTAGAGAACAAAATAGTTGAACCCACCTATGTTACCCC Ma. dyari (3AAABF): CTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCTGA CAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGCTG CCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAAAC TTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGTACTTCCT ATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAATAT TCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAACCA ATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTCTT ACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAACTG CCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAGAG AACAAAATAGTTGAACCCACCTATGTTACC Ma. dyari (1MBF3):

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70 ACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCT GACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGC TGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAA ACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGTACTTC CTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAAT ATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAAC CAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTC TTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAAC TGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAG AGAACAAAATAGTTGAACCCACCTATGTTAC Ma. dyari (3AABF): ACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCT GACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGC TGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAA ACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGTACTTC CTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAAT ATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAAC CAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTC TTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAAC TGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAG AGAACAAAATAGTTGAACCCACCTATGTTA Ma. dyari (2DBF): CCACCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATG CTGACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTA GCTGCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGA AAACTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGTACT TCCTATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCA ATATTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACA ACCAATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCC TCTTACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAA ACTGCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTT Ma. titillans (2MaTBF2): CCTGATCTTCCTCCATGAACGAGGATCATTTAACCCCCTAGGAATTAGCCCAAATGCTG ACAAAATCCCATTCCACCCCTACTTCACCATAAAAGACGCCCTAGGAGCAGCACTAGCT GCCTCCTCACTACTCATCTTAGCTCTCTACCTACCAGCCCTATTAGGGGACCCTGAAAA CTTCACCCCAGCAAATTCCATAATTACCCCAACACACATCAAACCCGAATGGTACTTCC TATTTGCTTATGCCATTCTACGATCTATTCCAAATAAGTTAGGAGGAGTACTAGCAATA TTCTCATCCATTTTAGTCCTATTCCTAATACCCGCCCTACACACAGCAAAACAACAACC AATATCAATACGCCCTATATCTCAGCTTCTATTTTGAGCCCTTACCCTGGACTTCCTCT TACTCACATGAATCGGAGGCCAACCAGTAAACCCCCCATATATTTTAATTGGCCAAACT

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71 GCCTCCCTATTCTACTTCATCATCATCCTAATCCTCATACCAATAGCAGGCCTCTTAGA GAACAAAATAGTTGAACCCACCTATGTTAC

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72 LIST OF REFERENCES Alves, E., C. Orilo, M. C. Rodriguez, and L. Solio. 2003. Mitochondrial DNA sequence variation and phylogenetic rela tionships among Iberian pigs and other domestic and wild pig populations. Anim. Genet. 34: 319-324. Andreadis, T. G. 2003. A checklist of the mosq uitoes 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 Culex pipiens Cx. restuans Cx. salinarius and Culiseta melanura Emerg. Infect. Dis. 7: 670-674. Alfonzo, D., M. E. Grillet, J. Liri a, J.-C. Navarro, S. C. Weaver, and R. Barrera. 2005. Ecological characterization of the aquatic habitats of mosquitoes (Diptera: Culicidae) in enzootic foci of Venezuel an 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. Hiriya n, 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. Hiri yan, V. Thenmozhi, and A. Gajanara. 2004. Japanese Encephalitis in Kerala, South India: Can Mansonia (Diptera: Culicidae) play a supplemental role in transm ission? 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 a nd 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. Sh i, V. L. Kulasekera, M. Edison, D. J. White, W. B. Stone, NY state West Nile vi rus surveillance team, and L. D. Kramer. 2001. West Nile virus infec tion in birds and mosquitoes, New York State, 2000. Emerg. Infect. Dis. 7:679-685.

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83 BIOGRAPHICAL SKETCH Sandra Coral Garrett was born on the wi ndy, 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 sib lings grew up in Aiken, but al so spent time on Hilton Head Island, a barrier island near th e 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 GovernorÂ’s School for Science and Mathematics for the last two y ears 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 coll ege and research pursuit s. It was during a school tour of the Clemson entomology de partment that Sandra decided entomology might be an exciting and rewarding area of bi ology to study. She b ecame interested in the University of FloridaÂ’s strong entomol ogy department and was able to attend with financial assistance from UFÂ’s Na tional 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 inte rest in entomology. She graduate d summa cum laude from UF and decided to stay for a masterÂ’s degree. Sh e met her future husband, Dr. Jos Carlos V. Rodrigues, in the department and was married in May of 2005.