Rhipicephalus Sanguineus Sensu Lato (Latreille) and Prevalence of Pathogens in Peridomestic Populations

PERMETHRIN-ETOFENPROX CROSS-RESISTANCE STATUS IN THE BROWN DOG TICK, RHIPICEPHALUS SANGUINEUS SENSU LATO (LATREILLE) AND PREVALENCE OF PATHOGENS IN PERIDOMESTIC POPULATIONS

NICHOLAS S.G. TUCKER

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

2017

To Alyson, Fox, and Texas

ACKNOWLEDGMENTS

I would like to acknowledge the funding provided by the U.S. Army as part of the

Long Term Health Education and Training program. The experience I have gained at the University of Florida will allow me to better serve my fellow service members. I would like to thank my committee, Dr. Phillip E. Kaufman and Dr. Emma N.I. Weeks, for their patience, understanding, and tremendous support. They provided a challenging environment allowing me to develop as a scientist and provided numerous opportunities for professional development. I am extremely grateful for the instruction I received from

Jessica Rowland. She was instrumental in getting my molecular work underway.

I would also like to thank my fellow graduate students Jeff Hertz, Chris

Holderman, and Lary Reeves, who were always there to provide assistance and lead me in the right direction. I would have been lost without you. Thank you Lois Wood and all of the support staff in the UF Veterinary Entomology Laboratory for your dedication and willingness to assist with any task. I would also like to express my gratitude to the faculty who go to great lengths to provide well-referenced and relevant instruction. Your expertise and professionalism has provided an example to emulate.

Thank you Dr. Lorenza Beati at Georgia Southern University for species group typing and Lauren Schumaker at CDC Atlanta for providing typed samples. I would also like to thank Dr. Katherine Sayler for her assistance in pathogen detection by providing expert advice and supplying positive control samples. I am also grateful to Dr. David

Allred for the positive control samples provided and Glenn Cawthorne for providing susceptible ticks to assay. Thank you to the numerous individuals and organizations for providing samples to assay, your efforts are greatly appreciated.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS ...... 4

LIST OF TABLES ...... 7

LIST OF FIGURES ...... 8

ABSTRACT ...... 10

CHAPTER

1 LITERATURE REVIEW OF THE BROWN DOG TICK RHIPICEPHALUS SANGUINEUS SENSU LATO (LATREILLE) ...... 12

Introduction ...... 12 Description and Identification ...... 12 Biology and Ecology ...... 15 Tick Management ...... 20 On-Host Control ...... 20 Off-Host Control ...... 24 Pesticides ...... 26 Pesticide Resistance...... 28

2 CHARACTERIZATION OF A SODIUM CHANNEL MUTATION IN PERMETHRIN RESISTANT RHIPICEPHALUS SANGUINEUS SENSU LATO (LATREILLE) ...... 33

Introduction ...... 33 Materials and Methods...... 36 Ticks ...... 36 Polymerase Chain Reaction ...... 38 Statistical Analysis ...... 40 Results ...... 40 Discussion ...... 42

3 IDENTIFICATION OF PERMETHRIN AND ETOFENPROX CROSS- RESISTANCE IN RHIPCEPHALUS SANGUINEUS SENSU LATO (LATREILLE) ...... 50

Introduction ...... 50 Materials and Methods...... 51 Ticks ...... 51 Bioassay ...... 52 Polymerase Chain Reaction ...... 54 DNA isolation ...... 54

DNA amplification ...... 55 Statistics ...... 57 Results ...... 57 Discussion ...... 59

4 PREVALENCE AND DISTRIBUTION OF PATHOGEN INFECTION AND PERMETHRIN RESISTANCE IN TROPICAL AND TEMPERATE POPULATIONS OF RHIPICEPHALUS SANGUINEUS SENSU LATO (LATREILLE) COLLECTED FROM SITES AROUND THE WORLD ...... 73

Introduction ...... 73 Materials and Methods...... 76 Ticks ...... 76 Molecular Analysis ...... 78 DNA isolation ...... 78 DNA amplification ...... 78 Results ...... 81 Discussion ...... 82

5 IMPLICATIONS AND FUTURE DIRECTIONS FOR RHIPICEPHALUS SANGUINEUS RESEARCH ...... 91

APPENDIX

A IMPORTATION FORMS ...... 95

B TICK COLLECTION INSTRUCTIONS ...... 98

LIST OF REFERENCES ...... 104

BIOGRAPHICAL SKETCH ...... 118

LIST OF TABLES

Table page

2-1 Primers used to detect a sodium channel mutation at nucleotide 2,134 in Rhipicephalus sanguineus (Latreille) sensu lato...... 47

2-2 Association within populations of Rhipicephalus sanguineus (Latreille) sensu lato of ...... 48

3-1 Primers used to detect the sodium channel mutation in the brown dog tick, Rhipicephalus sanguineus sensu lato (Latreille)...... 66

3-2 Etofenprox lethal concentration (LC) values for a laboratory-reared Rhipicephalus sanguineus sensu lato (Latreille) (Ecto Services Inc.) colony evaluated using the Food and Agriculture Organization larval packet test...... 67

3-3 Association between phenotypic and genotypic resistance to a discriminating concentration of either permethrin or etofenprox in peridomestic Rhipicephalus sanguineus sensu lato (Latreille)...... 68

3-4 Toxicity of permethrin and etofenprox to peridomestic Rhipicephalus sanguineus sensu lato (Latreille) collected in Grenada and Mexico...... 69

3-5 Acaricide exposure in peridomestic adult Rhipicephalus sanguineus sensu lato (Latreille) populations evaluated with a larval packet test as reported by pet owners or kennel operators...... 70

4-1 Primers used to detect sodium channel mutations and pathogens in the brown dog tick, Rhipicephalus sanguineus sensu lato (Latreille)...... 88

4-2 Occurrence of genotypic pyrethroid resistance and infection with Rickettsia and Hepatozoon parasites in peridomestic Rhipicephalus sanguineus sensu lato (Latreille) collected in Africa, Asia, Europe, North America, and the Caribbean...... 89

LIST OF FIGURES

Figure page

2-1 Agarose gel visualization of sodium channel mutation presence in larval extracts of the homozygous-resistant St. Johns Rhipicephalus sanguineus (Latreille) sensu lato population...... 49

3-1 Survival (%) of Rhipicephalus sanguineus sensu lato (Latreille) larvae collected from five globally distributed locations following exposure to 1, 5, 16, and 53 times (x) the permethrin discriminating concentration (0.19% permethrin ...... 71

3-2 Survival of Rhipicephalus sanguineus sensu lato (Latreille) larvae collected from five globally distributed locations and exposed to 1, 5, 16, and 53 times (x) the etofenprox discriminating concentration (0.51% etofenprox) using the Food and Agriculture Organization larval packet test...... 72

LIST OF ABBREVIATIONS

AI Active ingredient

CDC Centers for Disease Control and Prevention

DC Discriminating Concentration

DEM Diethyl maleate

DDT Dichlorodiphenyltrichloroethane

DNA Deoxyribonucleic acid

EPA Environmental Protection Agency

FAO Food and Drug Organization

FIFRA Federal Insecticide, Fungicide, and Rodenticide Act

GABA Gamma-Aminobutyric Acid

GST Glutathione S-Transferase

IPM Integrated Pest Management

LC Lethal Concentration

L:D Light:Dark

LPT Larval Packet Test

SNP Single Nucleotide Polymorphism

PCR Polymerase Chain Reaction

RH Relative Humidity

RR Resistance Ratio

TCE Trichloroethylene

TPP Triphenyl phosphate

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

PERMETHRIN-ETOFENPROX CROSS-RESISTANCE STATUS IN THE BROWN DOG TICK, RHIPICEPHALUS SANGUINEUS SENSU LATO (LATREILLE) AND PREVALENCE OF PATHOGENS IN PERIDOMESTIC POPULATIONS

Nicholas S.G. Tucker

August 2017

Chair: Phillip E. Kaufman Major: Entomology and Nematology

The brown dog tick, Rhipicephalus sanguineus (Latreille) sensu lato, is an important ectoparasite of dogs and occasionally humans, capable of transmitting several pathogens, such as Rickettsia and Ehrlichia, which are of veterinary and medical importance. The brown dog tick is distributed worldwide and has an affinity for human habitations in much of its range. In some populations, lack of integrated pest management plans and overuse of pyrethroid pesticides and other sodium channel inhibitors has resulted in high levels of resistance to permethrin.

A molecular assay targeting a point mutation in the sodium channel was developed and optimized to separate ticks expressing permethrin resistance from those from a susceptible colony. Thereafter, multiple field-collected phenotypically permethrin resistant populations were evaluated using this molecular assay to determine genotype.

A point mutation was present in greater than 85% of phenotypically-permethrin resistant tick populations that was not present in the susceptible colony.

Following the establishment of the single nucleotide polymorphism (SNP) molecular assay, cross-resistance to two sodium channel-inhibiting pesticides,

permethrin and etofenprox, was investigated and to the mode of resistance was evaluated. The etofenprox discriminating concentration (DC) was established and mortality results following exposure to the DC for either etofenprox or permethrin from five brown dog tick populations were compared. Subsequently, using both bioassay and molecular methods with geographically disparate samples, etofenprox resistance expression was investigated at the location of the aforementioned sodium channel SNP established with permethrin. One tick population bearing the homozygous SNP in 95% of individuals screened also was resistant to etofenprox at the discriminating concentration.

Rhipicephalus sanguineus populations collected in North America, the

Caribbean, Africa, Europe, and Asia were tested to elucidate the relationship between tropical and temperate population types, resistance mechanisms, and pathogen-vector interactions. Using molecular assays, populations from 24 distinct locations were simultaneously screened for pathogen infection by Rickettsia, Ehrlichia, Hepatozoon,

Babesia and the presence of a sodium channel SNP known to confer permethrin resistance. A relationship between tropical type ticks and the resistance conferring SNP was observed. Rickettsia and Ehrlichia were the most commonly observed pathogens and both were found in 11 of 28 screened populations. Hepatozoon was isolated in one population but Babesia was not observed in any of the populations. Also, this study is the first report of Rickettsia rickettsii and Rickettsia masiliae in ticks in Florida.

CHAPTER 1 LITERATURE REVIEW OF THE BROWN DOG TICK RHIPICEPHALUS SANGUINEUS SENSU LATO (LATREILLE)

Introduction

Ticks are blood-feeding arthropods that can cause damage to their hosts through feeding behavior and the transmission of pathogens. Ticks are among the most important vectors of disease-causing pathogens affecting livestock, companion animals, and humans (Jongejan and Uilenberg 2004, Dantas-Torres 2008). Rhipicephalus sanguineus (Latreille) is a three host hard tick (Family: Ixodidae) that varies its habitat preference depending on geographic location. In temperate regions, R. sanguineus is more likely to be endophilic than in tropical regions (Dantas-Torres 2008). This tick prefers to feed on the domestic dog Canis lupus familiaris, but has been shown to occasionally feed on other mammalian hosts, including human beings. Rhipicephalus sanguineus likely was spread throughout the world by dogs, their primary host, which were either following or transported intentionally by humans. The brown dog tick is cosmopolitan in distribution but there is contention as to whether this species evolved north or south of the Mediterranean region. Rhipicephalus sanguineus is probably the most wide-spread tick in the world and commonly found between latitudes 50˚N and

30˚S (Walker et al. 2000).

Description and Identification

Ticks, which are characterized by having eight legs as adults, are members of the class Arachnida, along with spiders, scorpions, and mites. Ticks and mites constitute the order Metastigmata, which is further divided into two suborders separating ticks and mites. All ticks species are obligate blood feeding parasites of terrestrial

vertebrates at one or more stage in their life cycle. The suborder Ixodida includes the three families of ticks: Argasidae, the soft ticks, Ixodidae, the hard ticks, and

Nuttalliellidae, a monospecific family. Ixodidae, to which R. sanguineus belongs, are characterized by having a hard scleritized scutum, anterior mouthparts, eyes when present are located near the lateral margin of the scutum, and large spiracles located behind coxae IV. In adult males the scutum completely covers the dorsal surface but is much reduced in females and immature stages. The reduced scutum in adult females allows for greater expansion during blood meal acquisition and during the development of eggs. In addition, members of the other families within this suborder lack a hard scutum.

The family Ixodidae is composed of 13 genera, of which, Rhipicephalus is one of the most speciose. Ticks of the genus Rhipicephalus have been shown to be competent vectors of many pathogens. The genus Rhipicephalus is comprised of 79 species, several of which recently were reclassified and moved from the genus Boophilus. The most notable reclassified species are those that vector the protozoans Babesia bovis and Babesia bigemina, which are the etiological agents of bovine babesiosis: R. microplus (Canestrini), R. annulatus (Say), and R. decloratus (Koch). Species within the genus Rhipicephalus are characterized by having eyes, festoons, a short hypostome and short palps, and in males, adanal plates. The scutum is typically patternless in

Rhipicephalus species, hence the common name of ticks within the genus, brown ticks.

Rhipicephalus sanguineus was first described by Latreille in 1806 as Ixodes sanguineus. Synonyms include: Boophilus dugesi Donitz, Eurhipicephalus sanguineus

Stephens & Christophers, Ixodes dugesi Gervais, Ixodes haxagonus sanguineus

Seguy, Ixodes linnaei Audouin, Ixodes sanguineus (Latreille), Rhipicephalus beccarii

Pavesi, Rhipicephalus breviceps Warburton, Rhipicephalus brevicollis Neumann,

Rhipicephalus carinatus Frauenfeld, Rhipicephalus limbatus Koch, Rhipicephalus macropis Schulze, Rhipcephalus rubicundus Frauenfeld, Rhipicephalus rutilus Koch,

Rhipicephalus sanguineus brevicollis Neuman, Rhipicephalus sanguineus sanguineus

Neumann, Rhipicephalus siculus Koch, Rhipicephalus stigmaticus Gerstacker, and

Rhipicephalus texanus Banks (Nijhof et al. 2015).

Unfed adult R. sanguineus are comparatively small (2.28-3.18 mm) and elongate, with coxa I having deep clefts and having a hexagonal basis capituli. In males, spiracular plates are comma shaped (Walker et al. 2000). Morphologic identification of

R. sanguineus may not be sufficient to differentiate between closely related species such as R. turanicus. Molecular identification using mitochondrial 16S rDNA, 12S rDNA, and cox1 gene sequences aided the construction of a recent phylogenetic tree (Dantas-

Torres et al. 2013). Genetic analysis conducted by Burlini et al. (2010) and Moraes-

Filho et al. (2011) suggests that R. sanguineus diverged into two distinct lineages; those found in the tropics between the latitudes 22°S and 25°N and those found in temperate regions living below 30°S and above 29°N. Levin et al. (2012) conducted an experiment where three laboratory colonies of R. sanguineus from North America, Israel, and Africa were crossbred to determine the fertility of the offspring. Crosses between ticks from

Africa with both ticks from North America and Israel resulted in sterile offspring but crosses between ticks from North America and Israel resulted in fertile offspring. This indicates that the African population may represent a different taxon. The allopatric

establishment of these two groups may be in part due to differences in tolerance of annual temperature extremes (Moraes-Filho et al. 2011).

Rhipicephalus sanguineus is a capable vector of numerous pathogens of medical and veterinary importance. It has been known to vector the etiological agents of canine babesiosis, canine filariasis, Q fever, canine monocytic ehrlichiosis, canine hepatozoonosis, canine haemobartonellosis, Mediterranean spotted fever, Rocky

Mountain spotted fever, and has been implicated as a potential vector for numerous other pathogens. Some hosts will develop immunological defense mechanisms in response to feeding, which can reduce feeding success and overall fitness of the tick. A feeding tick can modulate the host response with saliva constituents that favor the transmission of pathogens (Ferreira and Silva 1998). When a tick is able to remain feeding and subsequently exchange fluids with the host the likelihood of pathogen transmission will increase.

Biology and Ecology

Ticks develop through four developmental stages: egg, larva, nymph, and adult.

It is typical for an adult female brown dog tick to uninterruptedly oviposit around 4,000 eggs, but clutches as large as 7,273 were reported by Koch (1982). At 25˚C and 100% relative humidity, R. sanguineus is capable of about four generations per year

(Srivastava and Varma 1964, Dantas-Torres 2008). A gravid female will deposit an egg mass in an inconspicuous location such as in a crack or crevice in a wall or between rocks in order to protect them from predation by spiders, ants, or wasps (Dantas-Torres

2010). Rhipicephalus sanguineus eggs are small, spherical, and dark brown. The egg incubation period can last from six days to over 30 days depending on humidity and

temperature. During their first week post-eclosion, larval ticks will aggregate together as their chitinous exoskeleton hardens and will not typically feed during this time

(Srivastava and Varma 1964). Larval ticks, often referred to as seed ticks, are distinguished from other life stages by their small size and the presence of three pairs of legs. Rhipicephalus sanguineus larvae are flat and measure approximately 0.54 mm in length and 0.39 mm in width and lack a genital aperture (Dantas-Torres 2008). Nymphal

R. sanguineus have four pairs of legs, measure from 1.14-1.30 mm in length and 0.57-

0.66 mm in width, and lack a genital aperture. Unengorged adult male R. sanguineus are flat, reddish-brown, have four pairs of legs, and range in length from 2.28-3.18 mm and in width from 1.11-1.68 mm. Unengorged adult female brown dog ticks resemble adult males but can grow up to 12 mm in length after taking a blood meal because their reduced dorsal scutum does not inhibit expansion as it does in males.

After taking a successful blood meal, larval and nymphal ticks undergo ecdysis, also known as molting, which is the process in which the outer cuticular layer is shed in order to allow for cuticle growth and transition to the next life stage. Preceding and during ecdysis, structural changes to the subsequent life stage regulated by ecdysteroids occur in the integument, salivary glands, dermal glands, and mouthparts

(Rees 2004). After feeding, larval and nymphal ticks will drop off the host and seek a concealed location as they enter the pre-molting period. The pre-molting period may last several weeks depending on life stage and environmental conditions. Higher temperatures typically correlate with shorter molting periods (Koch and Tuck 1986).

Ecdysis takes several hours to complete and occurs when the cuticle ruptures and the old integument is shed by abdominal peristaltic waves (Dantas-Torres 2010). Similar to

the aggregation of larvae after eclosion, nymphs undergoing ecdysis have been shown to aggregate, which acts a mechanical stimulus and accelerates the molting process

(Dantas-Torres 2010). Adult female brown dog ticks will aggregate in response to conspecific faeces which indicate favorable environmental conditions (Yoder et al.

2013).

Rhipicephalus sanguineus is a three-host tick, meaning that each active developmental stage feeds only once and ecdysis occurs in the environment, requiring changing hosts between blood meals. A successful blood meal is necessary for development and reproduction. The domestic dog is the primary host for R. sanguineus, but this tick species has been known to parasitize small rodents, wild canids, and human beings (Goddard 1989, Venzal et al. 2003, Brouqui et al. 2005, Dantas-Torres

2008). Immature stages are more likely than adults to be found on other hosts, including human beings (Demma et al. 2005). There appears to be a correlation between parasitism by R. sanguineus on humans and the close association of those humans with tick-infested dogs (Burgdorfer et al. 1975, Uspensky and Ioffe-Uspensky 2002,

Demma et al. 2005).

Ticks utilize two approaches to locate hosts, questing and hunting. Ticks can wait along a path used by potential hosts with their front legs outstretched; this is called questing. The brown dog tick is considered a hunter tick because it actively pursues its host. It modifies its host-seeking behavior based on environmental conditions and the availability of hosts (Dantas-Torres 2008). Ticks can employ a variety of mechanisms in order to detect a suitable host. All ticks have sensory setae on the appendages and body surface that detect vibrations, heat, or chemicals like CO2 or ammonia (Waladde

and Rice 1982). Ticks also possess photoreceptors on the marginal edge of the scutum that aid in the location of their hosts. In R. sanguineus, these photoreceptors are in the form of eyes. Haggart and Davis (1980) found that R. sanguineus have ammonia- sensitive neurons located in the anterior pit of the Haller’s organ, which confer the ability to detect trace amounts of ammonia, such as the amount typically present in the sweat or excrement of mammals.

The feeding behavioral sequence found in all ticks consists of nine distinct events: hunting or seeking a host, physical contact with the host, searching on the host for an attachment site, insertion of the mouthparts into the host integument, securing to the feeding site with hardening attachment proteins, the slow uptake of host body fluids, the rapid uptake of host body fluids to complete engorgement, withdrawal of the mouthparts from the host integument, and leaving the host (Waladde and Rice 1982).

Brown dog ticks typically will feed only once during each life stage but adult males will take multiple blood meals. The feeding period of R. sanguineus can last from three to

11 days depending on the developmental stage and environmental conditions

(Troughton and Levin 2007). Unfed larvae can survive for up to eight months, unfed nymphs can survive for up to six months, and adults have been observed under laboratory conditions to survive up to 19 months without taking a blood meal (Dantas-

Torres 2008). Like other Ixodidae, adult R. sanguineus females die after oviposition.

Males of this species have been observed to survive up to 568 days under laboratory conditions (Little et al. 2007).

Ticks of the genus Rhipicephlaus are metastriate, meaning that both males and females require a blood meal before becoming sexually mature. Adult males will feed

intermittently and prior to each mating. Little et al. (2007) found that male R. sanguineus readily move between hosts, presumably in search of reproductively receptive females.

Due to their multiple-host feeding strategy, male metastriate ticks have been shown to intrastadially vector pathogens to multiple hosts (Kocan et al. 1992, Bremer et al. 2005).

Rhipicephalus sanguineus can attach anywhere on the host but typically are found on the back, inguinal region, axilla, interdigital spaces, and on the head, particularly the ears (Dantas-Torres 2010). When it has found a suitable feeding site, R. sanguineus will probe with its mouthparts and form a pool from which to feed by the laceration of blood vessels and capillaries. It then secretes a protein to firmly attach itself to the host. Little or no blood feeding occurs during the first 24 to 36 hours of attachment (Snyder et al.

2009). After the tick has established the feeding site, it begins to slowly feed and digest a blood meal. The slow uptake and digestion period is followed by a period of rapid ingestion of blood and expansion before dropping off the host to complete digestion in a secluded place. The salivation and regurgitation that occurs during early stages of blood feeding is important in the transmission of pathogens to the host. Tick saliva acts to regulate the immune response of its host so that the tick can continue to feed for a longer period undisturbed. Dogs do not develop an immune response to R. sanguineus re-infestation as seen in other hosts (Ferreira et al. 2003). In tick-resistant hosts the presence of R. sanguineus saliva and antigens induce an immediate and strong delayed-type hypersensitivity. This illustrates the close evolutionary relationship that R. sanguineus has with its primary host.

Mating of R. sanguineus occurs on the host and feeding initiates the sexual maturation of adults. After two days of feeding, females release the attractant sex

pheromone 2,6-dichlorophenol (Sonenshine 2006). A male will locate a feeding female by detecting the attractant sex pheromone, and when in close contact, will be stimulated by the mounting sex pheromone to position so that its ventral surface faces the female ventral surface. The mounting sex pheromone is composed of cholesterol esters and allows the male to ensure intraspecific mating. The male will then insert a spermatophore using its mouthparts and tarsi into the female gonopore. After completing a successful blood meal, the mated female will then drop off the host to begin the pre-oviposition period before depositing her eggs near a host resting site.

Tick Management

An integrated pest management approach should be applied when controlling any pest. Integrated pest management (IPM), which is defined as the selection, integration and implementation of pest control based on predicted economic, ecological and sociological consequences, makes maximum use of naturally occurring control agents, including weather, disease organisms, predators, and parasites (Bottrell 1979).

Unlike most ticks, the brown dog tick is endophilic in most regions where it occurs, meaning that it has an affinity for living inside human-made structures (Dantas-Torres

2008). It spends the majority of its life off-host, hiding in the cracks and crevices of structures. In heavily infested homes, adults and immatures can be seen climbing walls or curtains. On-host or off-host IPM approaches should be used simultaneously in the control of R. sanguineus.

On-Host Control

In cases where few ticks are present on host animals, manual removal is recommended. Ticks can be removed with tweezers or commercially available tick removal devices. Gloves should be worn to avoid contact with any potential pathogens.

Safe removal can be achieved by firmly grasping the mouthparts of the tick as close to the attachment site as possible and slowly pulling the tick out, ensuring that it is pulled without twisting. Care should be taken to ensure the mouthparts of the tick do not detach and remain in the host, which could lead to an inflammatory response and secondary infection. Following these extraction instructions will reduce the likelihood that the gut contents of the tick are emptied into the host, which would increase the potential for pathogen transmission. Topical application of any substance like petroleum jelly or fingernail polish in an attempt to suffocate or smother the tick should be avoided, as it is generally ineffective and can cause expulsion of the gut contents into the host

(Blagburn and Dryden 2009).

The ability of pesticides to repel or kill ticks before they attach and successfully feed on a host is important in the prevention of tick-borne pathogen transmission

(Young et al. 2003). Commercially available on-host tick control products include: spot- on treatments, pour-on treatments, impregnated collars, shampoos, and oral treatments. Spot-on treatments are pesticide formulations that are applied at the base of the neck between the shoulders of the dog monthly or as directed by the label. A 37-day trial on the effectiveness of a spot-on treatment composed of 10% imidacloprid and 50% permethrin was conducted by Epe et al. (2003). The treatment was applied to dogs previously infested with 50 R. sanguineus and 50 Ixodes ricinus L. at a dosage of 0.1 mL/kg. The treatment had a curative efficacy at day two of 74% against R. sanguineus and 67% against I. ricinus. At day 30, the preventive efficacy against re-infestation from

R. sanguineus and I. ricinus was over 95% (Epe et al. 2003). In order to test the effect of water and shampooing on spot-on treatments, Schuele et al. (2008) treated dogs

infested with adult R. sanguineus using a 12.5% pyriprole spot-on formulation and washed the dogs at varying intervals. It was concluded that neither repeated washing nor shampooing had a detrimental effect on the efficacy of the applied treatment against

R. sanguineus.

Another host-targeted pest control option is the use of pesticide-impregnated collars. Pesticide collars are molded plastic impregnated with a combination of slow release pesticides and are typically longer lasting when compared to spot-on treatments. Stanneck et al. (2012) found that an impregnated collar containing 4.5% flumethrin was 96% effective against R. sanguineus and I. ricinus when ticks were applied immediately after the collar was placed and 99% effective against infestation 6 hours after collar placement. The curative efficacy against R. sanguineus was between

32.2% and 46.5% and the preventive efficacy remained above 90% for 35 weeks.

Additionally, the study reported that neither shampooing nor immersing in water had a negative effect on the efficacy of flumethrin-impregnated collars against R. sanguineus

(Stanneck et al. 2012).

Pesticide-containing shampoos are another effective method to control ticks on dogs, although their use typically has a relatively short residual effect. Franc and

Cadiergues (1999) found that deltamethrin shampoo was effective at controlling R. sanguineus infestations and re-infestations. Dogs were infested with 50 R. sanguineus and left for 72 hours, before treatment with a 0.07% deltamethrin shampoo formulation on day 0, and re-infested on days 2, 7, 9, 14, 16, and 20. The treatment was 95% effective at controlling existing tick infestations, 99% effective against re-infestation during the first week, and 96% effective against re-infestation during the second week.

However, by day 22 of the study, the efficacy of the product had dropped to 86.3%, indicating that the shampoo formulation may be less persistent and require more frequent application than other treatment methods.

On-host R. sanguineus control measures also can be administered orally and have been shown to be at least as effective as spot-on treatments (Snyder et al. 2009,

Dumont et al. 2014, Letendre et al. 2014, Rohdich et al. 2014). Snyder et al. (2009) evaluated the efficacy of orally-administered spinosad, which is produced by the fermentation of the actinomycete, Saccharopolyspora spinosa. At 100 mg/kg, spinosad was found to be effective against existing R. sanguineus infestations, causing a 97.2% reduction in attached ticks (Snyder et al. 2009). Dumont et al. (2014) reported that the oral treatment Nexguard® was 100% effective against pre-existing tick infestations and remained 96.4% effective against re-infestation through day 30 of the study. The active ingredient of Nexguard® is afoxolaner, which is an isoxazoline that disrupts the arthropod specific gamma-aminobutyric acid (GABA) receptor causing irreversible hyperexcitation of the central nervous system (Letendre et al. 2014). GABA receptors are found in the central and peripheral nervous system and are responsible for the transportation of chloride ions across the cell membrane. Rohdich et al. (2014) compared the oral treatment Bravecto™ with the spot-on treatment Frontline™, over 12 weeks. The active ingredient of Bravecto™ is fluralaner, which is also an isoxazoline.

The active ingredient of Frontline™ is the phenylpyrazole fipronil. Dogs naturally infested by R. sanguineus or several other species of Ixodidae were treated with either

Bravecto™ once, or Frontline™ once per month for three months. Bravecto™ was

shown to be not significantly different in protection, but provided greater longevity than the commonly used spot-on treatment Frontline™ (Rohdich et al. 2014).

There is no commercially-available brown dog tick vaccine for canines.

However, research has been conducted to characterize the constituents of R. sanguineus salivary secretions to aid in vaccine development (Anatriello et al. 2010).

Vaccination against ticks of the genus Rhipicephalus has been shown to be an effective means of control in cattle and small mammals (de la Fuente et al. 1999, Trimnell et al.

2005). De la Fuente et al. (1999) evaluated Gavac™ (Heber Biotech), a vaccine designed to produce protective immune responses in cattle against Rhipicephalus

(Boophilus). In this vaccine, the cattle produced antigen, Bm86, inhibits bloodmeal digestion, reduces survival and egg laying capacity in surviving ticks, and thus reduces vectorial capacity (Willadsen et al. 1989, de la Fuente et al. 1999). These GavacTM- vaccinated cattle produced Bm86 in sufficient quantities to reduce pathogen transmission by reducing feeding success of R. microplus, R. decloratus, and R. annulatus (de la Fuente et al. 1999). Trimnell et al. (2005) found that vaccinating guinea pigs with incomplete portions of 64P salivary cement proteins derived from R. appendiculatus Neumann conferred protection by causing local inflammatory immune responses against R. sanguineus and other ticks. Rhipicephalus sanguineus avoided feeding on vaccinated guinea pigs under a free-release experimental design study wherein ticks were placed in the host’s enclosure and showed post-feeding mortality when ticks were enclosed in a feeding chamber attached to a host.

Off-Host Control

Considering that R. sanguineus lives off the host for the majority of its life, treatment of the home or kennel is required for successful management. Off-host

control of R. sanguineus typically includes all aspects of IPM. A highly recommended mechanical control method is the use of a high-efficiency particulate air filtered vacuum to capture ticks inside the home and immediately dispose of the ticks outside of the home in a sealed plastic bag in a trash receptacle. In highly infested homes, washing and drying of sheets and dog bed covers with high heat may be required. The removal and destruction of furniture and dog beds also may be necessary. Sealing cracks and crevices along baseboards and other inconspicuous locations with spray insulating foam or caulk will eliminate resting sites in the structure of the home. Other methods of physical control include removing harborage near the home and keeping grass trimmed around the home, especially in areas where dogs frequent.

Entomophagous fungi have been evaluated as a potential biological control agent of ticks. Unlike most pathogens, which have to be ingested in order to affect arthropods, fungi are capable of penetrating the host cuticle. Samish et al. (2001) tested the pathogenicity of four species of entomopathogenic fungi against the various life stages of R. sanguineus. The study found that in a laboratory setting Metarhizium anisopliae was transmissible between ticks at the same or different life stage and caused 92-100% mortality of unfed and engorged larvae, nymphs, and adults. The study also found that infection by M. anisopliae caused a secondary infection, which can prevent engorged females from egg laying (Samish et al. 2001).

Synthetic acarcides are used frequently in the control of R. sanguineus. Indoor residual pesticide sprays can remain active for up to six months, depending on the pesticide formulation and concentration used. In highly infested homes the use of on- host control and residual spraying is recommended.

Pesticides

Pesticides are synthetic or natural compounds that are applied to control damaging or annoying pests. Pesticides are used in various formulations to protect human and companion animal health, crops, and livestock. Through the use of pesticides and anti-malarial drugs, worldwide annual deaths caused by malaria were reduced from 6 million in 1939 to less than 600,000 in 2013 (Yu 2008, WHO 2014).

Pesticides used to control injurious arthropods work by various modes of action. Many pesticides affect the nervous system by binding to synapses or through nerve polarization. Other modes of action include: cell respiration disruption, cell osmosis disruption, chitin formation inhibition, or disruption of the development of immature target organisms. Commonly used pesticides to control arthropods include: pyrethroids, carbamates, neonicotinoids, organochlorines, organophosphates, insect growth regulators, microbials, and inorganic compounds. Pyrethroids, organophosphates, and neonicotinoids account for over 60% of global pesticide use (Yu 2008). Different pesticide classes can have the same mode of action. An example of this can be seen with research conducted on Aedes aegypti (L.) and the sodium channel inhibitors dichlorodiphenyltrichloroethane (DDT) and permethrin. In this study, cross-resistance to permethrin was documented in mosquito populations before it was used to control the vector (Rongsriyam and Busvine 1975). It was determined that prior exposure to DDT, an organochlorine class insecticide, had preselected the mosquitoes to be resistant to permethrin, a pyrethroid-class insecticide.

Pyrethroids are a synthetic photostable analog of pyrethrum, the toxin found in the flower of Chrysanthemum cinerariaefolium. Due to low vertebrate toxicity,

pyrethroids are the active ingredient in many pesticide formulations used in agriculture, home and garden, and ectoparasite control on humans and animals. Pyrethroids affect arthropods by altering voltage-gated sodium channels located on neurons. Neurons transmit electrical impulses to other sensory and motor neurons by a rapid polarization and depolarization along the axon between the cell body and synapse. When at rest, the neuron contains a high concentration of potassium ions and a low concentration of sodium ions; in contrast, the fluid surrounding the neuron contains a low concentration of potassium ions and a high concentration of sodium ions. Sodium exchange is regulated by two voltage-sensitive gates, whereas potassium exchange is regulated by one voltage-sensitive gate. When a nerve impulse travels along the axon of a neuron, a sodium channel activation gate opens, a sodium channel inactivation gate closes slowly, and a potassium channel gate opens slowly allowing for the depolarization of the neuron. This is followed by repolarization in which both sodium channels are closed and the potassium channel remains open until original ion distribution is achieved. The entire nerve impulse takes 0.001 seconds. Pyrethroids and DDT bind to the sodium channel site and delay the closure of the gates, which prevents the return to resting ion concentration. The exposed arthropod loses motor control and enters a state of rigid paralysis (Yu 2008). Pyrethroids are classified as type I or type II. The pyrethroid types differ in their molecular configuration, binding site on the sodium channel, symptomology, active duration in vivo, and effective temperature (Breckenridge et al.

2009). Due to the difference in molecular configuration, type II pyrethroids are thought to have enhanced acute neurotoxicity relative to type I pyrethroids. Another type of pesticide related to this class are the non-ester pyrethroids or pseudopyrethroids. These

pyrethroid-like pesticides also are sodium channel inhibitors, but are advantageous due to low fish and mammal toxicity, while maintaining high efficacy against insects

(Schleier III and Peterson 2011).

Pesticide Resistance

Any application of pesticides invariably selects for resistant individuals within a population. When pesticides are applied, genetically resistant individuals are more likely to survive, reproduce, and pass on these resistance-expressing genes. Resistance to pesticides is not uncommon; 80 members of the subclass Acari have been reported as resistant to an acaricide (Whalon 2015). Pesticide resistance is considered either behavioral or physiological. Behavioral resistance may be expressed as hypersensitivity or irritability to certain compounds. Wang et al. (2004) found that the German cockroach

Blatella germanica (L.), avoided the inert ingredients in gel baits; when the formulation was altered, the behavioral resistance to the formulation within the resistant population was lost. In that study, the resistant population exhibited reduced fecundity when compared to susceptible populations in the absence of selection, demonstrating that there is often a tradeoff between resistance and overall fitness. Another example of behavioral resistance is illustrated in the study of Anopheles farauti Laveran biting rhythm in Papua New Guinea. Anopheles mosquitoes, the vector for the protozoan that causes malaria, feed at night when people are typically asleep. The study found that An. faraunti shifted feeding activity to earlier in the night due to the use of permethrin- treated bed nets by the local population (Charlwood and Graves 1987).

Physiological resistance includes reduced cuticular penetration, metabolic detoxification, and target site insensitivity. The level of resistance expressed is

compounded when multiple modes of resistance are present. An example of cuticular resistance was observed by Noppun (1989) in laboratory conditions with an important crop pest, the diamond back moth, Plutella xylostella (L.). Noppun (1989) discovered that with resistant individuals, more than 65% of the applied fenvalerate was found on the surface of the cuticle, indicating that it did not penetrate, but less than 40% was found on the surface of susceptible individuals.

Metabolic resistance occurs when an arthropod metabolizes a toxic substance into a less harmful compound before the toxic effect manifests at the target site. One widely recognized group of metabolic detoxification enzymes are the cytochrome

P450’s, which are thought to have evolved in response to natural chemicals produced by plants (Després et al. 2007, Yu S.J. 2008). Metabolic resistance is typically nonspecific and has been shown to confer resistance in multiple arthropods to many pesticide classes (Carino et al. 1994, Scott 1999, Daborn et al. 2002).

Mutations on the sodium channel causing target site insensitivity are common among arthropods and seen in many dipteran and ixodid pests. An example of target site insensitivity is the inability of a pyrethroid to properly bind to the sodium channel target site due to an amino acid-coding point mutation that results in an altered confirmation of the target site. A classic example of this is knock-down resistance (kdr) in the house fly Musca domestica L., which was first identified in the 1950’s. Knock down resistance in M. domestica is caused by a point mutation associated with the sodium channel causing an amino acid change from leucine to phenylalanine that confers cross-resistance to DDT and pyrethroid pesticides (Soderlund and Knipple.

2003).

A study on resistant and super resistant laboratory reared strains of the horn fly

Haematobia irritans (L.), an important cattle pest, was conducted by Guerrero et al.

(1997) to determine the resistance mechanisms to pyrethroids. The resistant strain did not receive any additional selection pressure after the establishment of the colony. The super resistant strain was a subset of the resistant strain that was exposed to cyhalothrin weekly for over three years. The results of a filter paper bioassay showed that the resistant and super resistant strains were 17 and 11,300 times, respectively, more resistant to cyhalthrin than the susceptible colony. The resistant and super resistant colonies were also cross-resistant to permethrin at a rate of 17 and 688 times, respectively, compared to the susceptible colony. Genetic analysis using real time polymerase chain reaction (PCR) showed that the super resistant strain had two point mutations associated with pyrethroid resistance, whereas the resistant strain had only one. It was postulated that the weekly selective pressure of the super resistant strain by cyhalthrin selected for individuals with multiple sodium channel mutations and enhanced detoxification mechanisms (Guerrero et al. 1997). The super resistant strain in this study with multiple sodium channel mutations is termed super knock down resistance or super-kdr.

The southern cattle tick, R. microplus is a major pest of cattle in many parts of the world. This tick and the pathogens associated with it were so damaging that the

U.S. eradicated this tick and continually monitors cattle entering the U.S. from Mexico.

Rhipicephlaus microplus was investigated by Guerrero et al. (2001) to determine the molecular basis of acaricide resistance in this tick. Using larvae from permethrin susceptible and resistant strains of R. microplus, a mutation at nucleotide number 2,134

of the sodium channel in resistant ticks was verified using traditional PCR. The mutation from thymine to adenine in the S6 transmembrane segment of domain III of the para- like sodium channel caused a change in the coded amino acid. This change in amino acid from phenylalanine to isoleucine resulted in target site insensitivity to permethrin

(Guerrero et al. 2001).

Acaricide resistance was investigated in the brown dog tick in a laboratory study conducted by Eiden et al. (2015). Known susceptible larval ticks were exposed to various levels of permethrin or fipronil using a larval packet test in order to determine a discriminating concentration. The discriminating concentration is two times the LC99 value, which is the concentration at which 99% of the larvae did not survive. The discriminating concentration was determined to be 0.19% for permethrin and 0.15% for fipronil (Eiden et al. 2016). Ticks from infested homes and kennels in Florida and Texas were then assessed for permethrin and fipronil reistance using a larval packet test. In one population of R. sanguineus, the level of resistance to permethrin was so high that

72% of larvae survived when challenged with 30% permethrin (Eiden et al. 2017).

Resistance ratios for fipronil were much lower than permethrin but all four populations assessed showed some tolerance to the active ingredient in Frontline™ spot-on products. The Eiden et al. (2017) study used three synergists, piperonyl butoxide, diethyl maleate, and triphenyl phosphate to determine the presence of metabolic resistance within sampled populations. In several populations, two or more metabolic resistance mechanisms were observed. Cytochrome P450 activity was observed in all resistant populations but permethrin susceptibility was not restored through the use of synergists. This implicates that other resistance mechanisms such as target site

insensitivity could be present in these permethrin resistant populations (Eiden et al.

2017).

Based on previous resistance work with R. microplus, it is likely that a mutation on the sodium channel is responsible for permethrin resistance not accounted for by metabolic resistance. Currently there is no published research covering this topic in R. sanguineus. Considering that this tick is distributed worldwide and an important vector of many pathogens, further investigation is warranted.

CHAPTER 2 CHARACTERIZATION OF A SODIUM CHANNEL MUTATION IN PERMETHRIN RESISTANT RHIPICEPHALUS SANGUINEUS SENSU LATO (LATREILLE)

Introduction

The brown dog tick, Rhipicephalus sanguineus (Latreille) sensu lato, is a three- host tick, but is highly restricted in its preferred host, the domestic dog. It is perhaps this close association that has led the brown dog tick to be the only tick known to complete its lifecycle indoors, particularly in temperate climates (Dantas-Torres 2008). This synanthropic adaptation has allowed for increased exposure to acaricides used both in the environment (residence or kennel) and on the canine host when compared to exophilic tick species. Brown dog ticks spend greater than 95% of their lifespan in the environment and dogs serve as the primary dispersal mechanism of this species

(Dantas-Torres 2010). Thus, exchange of ticks among locations is largely restricted to dog visitations to common areas, such as veterinary clinics, or when uninfested dogs visit infested homes or vice versa. This restricted genetic exchange along with long- lasting or prophylactic use of acaricides with a similar mode of action provides tremendous opportunity for selection of pesticide resistant populations. Acaricide resistance in R. sanguineus has been observed in several North America populations

(Miller et al. 2001, Eiden et al. 2015) and samples from around the world are currently being tested in the Veterinary Entomology Laboratory, Entomology and Nematology

Department, University of Florida (Gainesville, FL, U.S.).

Brown dog ticks are not unique in their expression of pesticide resistance.

Worldwide, over 440 instances of arthropods expressing resistance to one or more pesticides have been recorded (Roush and Tabashnik 2012). Physiological pesticide

resistance can occur through metabolic detoxification, reduced cuticular penetration, and target site insensitivity. A point mutation conferring target site insensitivity to pyrethroids is common and has been observed in multiple subphyla of Arthropoda

(Guerrero et al. 1997, He et al. 1999, Martinez‐Torres et al. 1999, Ranson et al. 2000,

Fallang et al. 2005). The negative impact generated by important arthropod vectors can be exacerbated by the presence of pesticide resistance, especially when a reliable alternative to the failing active ingredient is not available.

Permethrin is a commonly used pesticide both in and around the home. Many formulations containing permethrin exist that target numerous arthropod species; its popularity is due to its quick mode of action, high efficacy, and low vertebrate toxicity

(Yu 2008). Frequent exposure to permethrin products designed for other indoor pests and pesticides with similar modes of action can inadvertently select for resistance in brown dog ticks. Widespread availability along with relatively low costs have undoubtedly contributed to the selection for permethrin resistant populations of R. sanguineus. In addition, the use of cyclodienes, such as dichlorodiphenyltrichloroethane

(DDT), has been shown to confer cross-resistance to pyrethroids including permethrin, in some arthropods due to target site insensitivity and metabolic detoxification (Scott et al. 1999, Brengues et al. 2003).

Homeowners and pest control operators have reported difficulties in managing brown dog tick infestations using conventional pesticide applications, although the underlying reasons for this remain unclear. Permethrin resistance was recorded in nearly all field-collected Texas and Florida populations of R. sanguineus, with several populations thought to express multiple resistance mechanisms (Eiden et al. 2015). This

conclusion was based on certain populations expressing a resistance level that could not be completely overcome by metabolic mechanism-blocking synergists. Thus, permethrin resistance in these highly resistant populations was suspected to result from, in part, target site insensitivity of the sodium channel. Target site insensitivity has been investigated previously in the brown dog tick. A study on a Panama population of

R. sanguineus implicated a sodium channel mutation as a potential resistance mechanism in ticks regularly exposed to DDT and pyrethroids (Miller et al. 2001).

A molecular-based technique was developed by Guerrero et al. (2001) to detect the presence of a single nucleotide polymorphism (SNP) in the sodium channel of the cattle fever tick, Rhipicephalus (Boophilus) microplus (Canestrini). The reported point mutation in pyrethroid-resistant R. microplus individuals is located on domain III segment VI of the sodium channel at 2,134 and is a substitution of adenine in place of thymine. Due to the close evolutionary relationship between R. microplus and R. sanguineus, Klafke et al. (2017) sequenced this highly conserved region, i.e. domain III segment VI of the voltage gated sodium channel gene cDNAs, from phenotypically susceptible and pyrethroid resistant R. sanguineus strains to determine amino acid and nucleotide sequences. Sequencing revealed four SNP’s, three of which (T2024C,

C2033T, and G2048A), were missense mutations and did not cause an amino acid change. The SNP of cytosine in place of thymine at nucleotide 2,134 conferred an amino acid change from phenylalanine to leucine. As such, the goal of this study was to utilize this presumptive mutation site to develop a molecular assay to detect pyrethroid resistance with the following objectives. Firstly, to optimize a polymerase chain reaction

(PCR) assay to determine the resistance genotype of brown dog ticks, which could be

used to screen populations collected in the field. A secondary objective was to determine the prevalence of the mutation among previously collected samples of R. sanguineus of known phenotypic pesticide resistance status and to confirm genotypic correlation with phenotypic resistance-expressing individual brown dog ticks.

Materials and Methods

Ticks

Resistant and susceptible R. sanguineus were used to develop the PCR assay.

All ticks were phenotypically evaluated by Eiden et al. (2015) using the Food and

Agricultural Organization (FAO) larval packet test (LPT) (FAO 2004) and held at -80°C.

The susceptible ticks were laboratory reared R. sanguineus (Ecto Services Inc.,

Henderson, NC). Resistant ticks were bioassay-screened field collected R. sanguineus from Brevard county FL, which were collected from a private residence and showed high resistance to 10% permethrin. Thereafter, resistant and susceptible ticks were used to evaluate the PCR assay for the presence of a sodium channel target site insensitivity mechanism associated with permethrin resistance.

Resistant ticks were field collected by pest control operators from kennels and homes that were experiencing tick infestations between June 2010 and August 2013, originating in Florida and Texas. Tick populations were named by the county in which they were collected, with multiple county collections from unique residences numbered sequentially. The seven populations from Florida evaluated in this study were: Broward,

Gilchrist, Palm Beach-1, Palm Beach-2, Palm-Beach-3, Sarasota-2, and St. Johns. An additional population was evaluated from Texas, named Webb, TX. For the purposes of this study, we define phenotypic resistance as the ability of an individual to survive the

acaricide discriminating concentration or higher. Phenotypic resistance testing was conducted by Eiden et al. (2016) using technical grade permethrin. In brief, larval ticks,

12-14 days old, were exposed to 1, 5, 16, or 53 times the permethrin discriminating concentration (0.19%) using the FAO LPT (FAO 2004). The discriminating concentration is double the LC99 value for susceptible populations. Phenotypically resistant ticks were stored dry at -80˚C and used in this study.

Metabolic resistance was assessed through the use of synergists in the Broward,

Gilchrist, Palm Beach-2, and Sarasota-2 brown dog tick populations included in this study by Eiden et al. (2017). Larval ticks were exposed to 1, 5, and 16 times the permethrin discriminating concentration with the simultaneous addition of a synergist in the LPT. The synergists used were triphenyl phosphate (TPP) an esterase inhibitor, piperonyl butoxide (PBO) a cytochrome P450 inhibitor, and diethyl maleate (DEM) a glutathione-S-transferase inhibitor. Each synergist was applied separately at a constant rate.

To demonstrate the frequency of the point mutation believed to confer resistance

(Klafke et al., 2017), 20 individual R. sanguineus larvae from the susceptible colony and eight distinct populations were tested using our PCR protocol. Larvae used were those that survived the highest exposure dose of the previously described bioassays. As presented by Eiden et al. (2015), all tested strains displayed greater than 70% survival at the discriminating concentration. Three strains in particular, Broward, Palm Beach-1, and St. Johns, showed 100% survival at the permethrin discriminating concentration.

The Gilchrist and Palm Beach-2 populations displayed 94%, and 99% survival, respectively, at the discriminating concentration with the addition of TPP.

Polymerase Chain Reaction

DNA was extracted from individual larvae using a modified technique described by Guerrero et al. (2001). In brief, larvae were snap frozen in liquid nitrogen. After 1 minute of cooling, larvae were transferred into a Petri dish held on ice. Individual larvae were transferred to bead beating tubes held in liquid nitrogen (Fisher Scientific,

Waltham, MA) containing three 2.0 mm diameter beads (BioSpec Products, Bartlesville,

OK). Bead beating tubes containing larvae were held in liquid nitrogen until disruption by bead beating was conducted.

Ticks were pulverized in bead beating tubes using a Precellys® Evolution homogenizer (Bertin Technologies, île de France, France) for three iterations of 20 seconds with 30 second pauses in between. After pulverizing, tubes containing ticks were returned to liquid nitrogen. Pulverized samples received 25 µL of sterile DNA isolation buffer (3.16 g of 1.0 M Tris HCL to 7.5 g of 1.0 M KCL). Tubes were placed in a

100˚C water bath for five minutes. Lastly, sample tubes were placed in a 4˚C centrifuge at 14,000 rpm for five minutes. Extracted DNA was stored at -20˚C if not immediately subjected to PCR amplification.

DNA amplification methods developed for the cattle fever tick, R. microplus (He et al. 1999) were modified to develop a novel PCR at the target site (GENBANK

KU886032) (Klafke et al. 2017). Sense and anti-sense degenerate primers were designed to be specific to the brown dog tick sodium channel. Each 20 µL reaction received: 2 µL of DNA, 20 pmol of each primer, 0.7 µL of 50 mM MgCl2 (Invitrogen,

Carlsbad, CA), 0.5 µL of 10 mM dNTP mix (Thermo Fisher Scientific, Waltham, MA), 0.1

µL of Platinum® Taq polymerase (Invitrogen, Carlsbad, CA), and 2 µL of 10x PCR reaction buffer (Invitrogen, Carlsbad, CA). Two reactions were used for each sample to

visualize the SNP on a gel due to the identical amplicon size produced by both primer sets. Therefore, the primers BDT-227 and either RSSC-SUS-F to amplify phenotypic susceptible tick DNA or RSSC-RES-F to amplify phenotypic resistant tick DNA were used to generate a 69 base pair amplicon at the site of the point mutation (Table 1).

Specificity of the PCR was optimized at 59˚C using an annealing temperature gradient of 55˚C to 65˚C. Both susceptible and resistant DNA was used with both primer sets to ensure the correct temperature was used in order to prevent non-specific priming. PCR conditions with an optimized annealing temperature were as follows: 1 cycle at 96˚C for

3 minutes, followed by 40 cycles at 94˚C for 1 minute, 59˚C for 1 minute, and 72˚C for 1 minute with a final 7 minute extension step at 72˚C. Using this approach, samples were determined to be either heterozygous, or homozygous resistant or susceptible. Primers

FG-228 and BDT-227 were used to produce a 126 base pair PCR product that was used for sequencing (Table 1). All 20 of the susceptible larvae and 10% of randomly selected samples from each population were sequenced. The PCR reaction for sequencing used the same master mix as described above. The 20 µL mixture was pipetted into a 96 well plate and placed into a Bio-Rad DNAEngine™ Peltier Thermal

Cycler (Bio-Rad Laboratories, Hercules, CA). PCR conditions were the same as described earlier except the annealing temperature was 50˚C. Samples were held at

4˚C until used for gel electrophoresis.

A 3% agarose gel with Synergel™ agarose clarifier additive (Diversified Biotech,

Dedham, MA) and ethidium bromide solution was mixed. Using FastRuler Ultra Low

Range DNA Ladder (Thermo Fisher Scientific, Waltham, MA) and 6x DNA loading dye

(Thermo Fisher Scientific, Waltham, MA), DNA fragments were visualized and photographed under UV light.

DNA was purified using QIAquick® PCR Purification Kit (Qiagen, Hilden,

Germany) following the manufacturer’s protocol. Susceptible colony and resistant population DNA was sequenced to determine if the mutation was present at the exact location of the SNP on domain III segment VI of the sodium channel that confers permethrin resistance. Sequencing results were analyzed using the pairwise function in

Bioedit to confirm the presence of the susceptible sequence or the resistant sequence with the mutation at nucleotide 2,134 on domain III segment VI of the sodium channel

(Bioedit Sequence Alignment Editor Version 7.0, Ibis Bioscience, Carlsbad, CA).

Statistical Analysis

The sequencing results were used to quantify the frequency of the point mutation believed to confer resistance within the susceptible colony and each population. For each colony or population the frequency of the resistant allele was calculated by summing the number across all genotypes. The proportion of resistant alleles was then calculated by dividing that number by the total number of alleles (number of individuals tested multiplied by two). The Pearson product-moment correlation coefficient was calculated between percentage survival at 0.19% permethrin, the discriminating concentration according to Eiden et al. (2016), and the proportions of the resistant allele.

Results

The PCR was successfully optimized to identify the SNP on domain III segment

VI of the sodium channel, allowing for the screening of eight phenotypically-resistant brown dog tick populations. Sequencing of a random 10% of samples from each

phenotypically permethrin-resistant population, with the resistant band as visualized on a gel, confirmed that the point mutation of thymine to cytosine conferring permethrin resistance was present in all eight populations tested (100%; 16/16). Sequencing of 20 individual Ecto Services phenotypically susceptible colony larvae, with the susceptible band as visualized on a gel, also confirmed the susceptible genotype (100%; 20/20). Of the eight field populations evaluated, 95% of individual tick samples were homozygous for the resistance-conferring sodium channel mutation (RR), while 1% were homozygous susceptible (SS), and the remaining 4% were heterozygotes (SR). A comparison with the results from the PCR assays and the LPT data collected by Eiden et al. (2017) showed an apparent association between genotypic and phenotypic resistance (Table 2). Gilchrist, Palm Beach-3, Sarasota-2, and St. Johns populations displayed 100% homozygous resistance and the observed survival at the discriminating concentration, 0.19% permethrin, was also high.

Broward, Palm Beach-1, Palm Beach-2, and Webb, TX populations included homozygous susceptible individuals. Heterozygous individuals accounted for 4% and

22%, of the phenotypically-resistant Palm Beach-1 and Webb, TX populations, respectively. Of the populations evaluated in this study, the Webb, TX population had the highest percentage of heterozygous individuals and the lowest survival at the discriminating concentration. Combined, these results suggest that this population had the lowest permethrin resistance expression among the eight sampled.

The correlation between the proportions of the resistant allele of a population or colony and the percentages of survival at 0.19% permethrin, the discriminating concentration, was 0.98 with a significant p-value of <0.001. As the percentage survival

at the discriminating concentration increased so did the proportions of the resistant alleles in the population. The correlation remained significant (0.72; p<0.05) even if the results of the susceptible colony were excluded from the analysis. This confirms the presence of a significant association between phenotypic resistance and the presence of the mutation.

Discussion

Using sequences published by Klafke et al. (2017), a molecular PCR assay was successfully optimized to detect a R. sanguineus sodium channel SNP and was used to demonstrate the presence of the mutation among eight previously collected brown dog tick sample populations with known phenotypic permethrin resistance expression.

Bioassay, molecular assay, and sequencing data were consistent and showed a significant association between phenotypic resistance to permethrin and the substitution of cytosine for thymine on domain III segment VI at nucleotide 2,134 of the R. sanguineus sodium channel.

The co-occurrence of enhanced metabolic detoxification and target site insensitivity in injurious arthropod pests is not uncommon (Scott and Georghiou 1986,

Jamroz et al. 2000, Guerrero et al. 2002, Brengues et al. 2003, Ochomo et al. 2013).

Scott and Georghiou (1986) investigated the resistance mechanisms occurring in a known pyrethroid-resistant strain of the house fly, Musca domestica L. Using electrophysiological, in vitro metabolism, in vivo penetration, and synergism studies they characterized enhanced metabolic detoxification, target site insensitivity, and reduced cuticular penetration activity in this highly resistant population. As in the present study, metabolic detoxification was the most important mechanism in permethrin resistance, but target site insensitivity also played a role in the ability of the house fly strain to

overcome the desired insecticidal effect. Populations of the dengue vector Aedes aegypti (L.), collected throughout the world, were analyzed by Brengues et al. (2003) to understand the distribution of resistance mechanisms within this species. Eleven out of thirteen of these populations exhibited some form of pyrethroid resistance. Four novel sodium channel mutations were identified, all conferring varying levels of resistance.

Brengues et al. (2003) also reported that the relative contribution of metabolic detoxification and target site insensitivity were strain dependent, which has interesting implications to the current, as well as future studies. Two Aedes strains were identified as utilizing both metabolic detoxification and at least one target site mutation. Like Ae. aegypti, R. sanguineus is distributed worldwide and likely exhibits similar diversification of resistance mechanisms.

A study investigating esterase-mediated and target site insensitivity mechanisms in two permethrin-resistant strains of R. (Boophilus) microplus collected in Mexico was conducted by Guerrero et al. (2002). The mutation investigated is at the same location on domain III segment VI, as the mutation detected in our R. sanguineus populations.

Both R. microplus populations demonstrated survival at low doses of permethrin, but through different mechanisms. One population of those ticks utilized metabolic resistance as the primary survival mechanism, while the other population utilized the sodium channel mutation. It is important to note, however, that both mechanisms were present in each population. Contrary to our results with R. sanguineus, the sodium channel mutation expressed in R. microplus conferred much higher resistance expression than did the expression of enhanced esterase activity. This may have been due to the difference in the coded amino acid conferred by the mutation; the R.

sanguineus mutation codes for leucine, whereas the R. microplus mutation codes for isoleucine. This comparison may suggest that the change to leucine only provides partial resistance to permethrin in R. sanguineus.

The United States Department of Agriculture (USDA), Agricultural Research

Service (ARS) laboratory (Kerrville, TX) established a colony of brown dog ticks from the Corozal Army Veterinary Quarantine Center in Panama where a severe infestation was showing resistance to pyrethroids, carbamates, and organophosphates (Miller et al.

2001). Aside from being a nuisance to the kenneled transient dog population, the ticks were transmitting Ehrlichia canis among the dogs. The laboratory-reared ticks were assayed against five acaricides and two synergists. Ticks were shown to be highly resistant to permethrin alone but susceptible following the addition of TPP. Enzyme activity gel staining showed high esterase activity indicating that metabolic detoxification through enhanced esterase activity was an important resistance mechanism. However, resistance conferred by a sodium channel SNP could still not be ruled out as an additional resistance mechanism.

Studies by Eiden et al. (2017) indicated that metabolic resistance is an important mechanism used by the brown dog tick to overcome permethrin toxicity. It was apparent that increased esterase activity had the greatest effect on permethrin resistance, followed by cytochrome P450’s, then minimally detoxification by glutathione-S- transferase. None of the synergists fully restored the acaricidal effect of permethrin at the discriminating concentration in any of the four populations (Eiden et al. 2017). The addition of TPP had the greatest effect in restoring permethrin toxicity in the Broward population where 54% mortality was observed at the permethrin discriminating dose.

However, the Palm Beach-2 strain demonstrated little change when TPP was added, displaying only 1% mortality at both the discriminating concentration and at five times the discriminating concentration. The most interesting data were observed at the discriminating concentration when the cumulative effects of each metabolic resistance mechanism are viewed collectively. There remains a gap between 100% mortality and the observed data within each population, indicating that another mechanism is contributing to permethrin resistance expression. Homozygous mutation rates within these populations exceeded 95%, suggesting that the SNP at 2,134 on domain III segment VI confers permethrin resistance in these populations at the discriminating concentration.

In a closed environment, such as a residence, selective pressure by permethrin is likely to drive the increased occurrence of resistance. The molecular assay developed herein provides a reliable method for accurately determining genotypic pyrethroid resistance associated with target site insensitivity in R. sanguineus. Additionally, this technique provides a means to rapidly screen a submitted brown dog tick population for permethrin resistance and provide clientele with feedback on the likelihood that a permethrin application will be efficacious. The potential difficulty in controlling the brown dog tick on the host and in the environment is demonstrated by the lack of combined acaricidal and synergist efficacy in populations possessing the SNP. Further research should investigate other potential mutations on the sodium channel of the brown dog tick that may be acting to provide resistance. Additionally, recent studies suggest that R. sanguineus sensu lato systematics are reflective of perhaps four species types, several of which may be geographically dependent on temperature (summarized in Zemtsova et

al. 2016). The impact of these clades in relation to acaricide resistance expression remains unknown.

Table 2-1. Primers used to detect a sodium channel mutation at nucleotide 2,134 in Rhipicephalus sanguineus (Latreille) sensu lato. Purpose Primer Name Direction Sequence

Na+ Channel Mutationa RSSC-SUS-F Forward 5’-ATT ATC TTC GGC TCC TTC T-3’

RSSC-RES-F Forward 5’-ATT ATC TTC GGC TCC TTC C-3’

BDT-227 Reverse 5’-TTG TTC ATT GAA ATT GTC AA-3’

Na+ Channel Sequencing FG-228b Forward 5’-CTA ACA TCT ACA TGT ACC-3’

BDT-227a Reverse 5’-GCA ATC CTC CAG CCT TC-3’ a Primers developed in this study using GENBANK KU886032 as a target sequence. b Primers from Guerrero et al. (2001).

Table 2-2. Association within populations of Rhipicephalus sanguineus (Latreille) sensu lato of phenotypic and genotypic resistance to permethrin. Strain % survival at % survival at 0.19% No. Genotypec Resistant allele 0.19% permethrina permethrin + TPPb individuals proportionf tested SS SR RR Ecto Services 0 NT 20 20 0 0 0.00

Broward 100 46 20 1 0 19 0.95

Gilchrist 100 94 21 0 0 21 1.00

Palm Beach-1d NT NT 21 1 1 19 0.93

Palm Beach-2 100 99 22 1 0 21 0.95

Palm Beach-3 88 NT 21 0 0 21 1.00

Sarasota-2 100 50 20 0 0 20 1.00

St. Johnse 100 NT 20 0 0 20 1.00

Webb, TXe 74 NT 20 0 5 15 0.88 a Survival at 0.19%, the discriminating concentration (Eiden et al. 2015b). b Survival at 0.19%, the discriminating concentration, with and without the esterase blocking synergist triphenyl phosphate (TPP) (Eiden et al. 2017). c Genotypic expression of a point mutation resulting in an amino acid substitution of cytosine for thymine in domain III segment VI at nucleotide 2,134 of the sodium channel, RR=homozygous resistant, SR=heterozygous, and SS=homozygous susceptible. d Palm Beach-1 displayed high survival (78%) at 10% permethrin (Eiden et al. 2015a), but was not tested at the discrimating concentration, 0.18% permethrin. e Assayed at 0.15% permethrin (Eiden et al. 2015a). f Proportion of resistant allele (R) including heterozygotes as compared to susceptible alleles (S). NT= Not tested

Figure 2-1. Agarose gel visualization of sodium channel mutation presence in larval extracts of the homozygous-resistant St. Johns Rhipicephalus sanguineus (Latreille) sensu lato population. The upper portion of the gel (S) shows that the non-mutated wild-type (susceptible) allele was not present in the tick population using the primer set, RSSC-SUS-F and BDT-227. The lower portion of the gel (R) shows presence of the mutation, displayed as white bands, using the resistant primer set, RSSC-RES-F and BDT-227. L=Ladder, B=Blank sample, +=Positive control for either the susceptible or resistant genotype.

CHAPTER 3 IDENTIFICATION OF PERMETHRIN AND ETOFENPROX CROSS-RESISTANCE IN RHIPCEPHALUS SANGUINEUS SENSU LATO (LATREILLE)

Introduction

In the United States, pesticides require a registration certification by the U.S.

Environmental Protection Agency (EPA) in accordance with the guidelines set forth by the Federal Insecticide, Fungicide, and Rodenticide Act (FIFRA) before a pesticide can be applied legally to control pests. The EPA must evaluate scientific data in order to generate a risk assessment of the pesticide. While the registration process is in place to protect human and environmental health, delays due to the registration process or subsequent EPA cancellation of registrations can result in limited pesticide availability for control of injurious pests.

Considering that many pesticides, including those in different classes, have a similar mode of action, the loss of any one pesticide can reduce the available effective pesticides additively. Cross-resistance to multiple pesticides has been recorded in numerous arthropod pests (Cahill et al. 1995, Rodríguez et al. 2002, Brengues et al.

2003). The non-ester pyrethroid, permethrin, and the pyrethroid-like pesticide, etofenprox, are both sodium channel inhibitors that disrupt normal nerve cell function.

Both pesticides are available in formulations to control arthropods in and around the home. Although permethrin etofenprox cross-resistance has been investigated in other arthropods, such information, to our knowledge, does not exist for any tick species. A study by Fonseca-González et al. (2011) found four of 12 surveyed Colombian populations of Aedes aegypti (L.) were cross-resistant to permethrin and etofenprox at varying levels. Enhanced metabolic detoxification was detected in these resistant populations, but sodium channel mutations were not investigated (Fonseca‐González et

al. 2011). However, mosquito behavior and consequent contact with pesticide residues would be quite different to that experienced by an endophilic tick, such as the brown dog tick. Resistance to numerous classes of insecticides was investigated in the endophilic hemipterans Cimex lectularis (L.) and C. hemipterus (F.), which were collected in several Thailand hotels (Tawatsin et al. 2011). Resistance to permethrin and etofenprox was observed in several of these populations and was conferred by a mutation of the voltage-gated sodium channel.

The aims of this study were to investigate the presence of cross-resistance to these two sodium channel-inhibiting pesticides in the brown dog tick, Rhipicephalus sanguineus sensu lato (Latreille), and to evaluate the mode of resistance. The primary goal was to establish the discriminating concentration for etofenprox with a secondary goal of comparing the acaricidal action of both pesticides. Subsequently, using both bioassay and molecular methods and geographically disparate specimen samples, etofenprox resistance was investigated at the location of a sodium channel single nucleotide polymorphism (SNP) known to confer permethrin resistance (Tucker et al. in press).

Materials and Methods

Ticks

Laboratory reared and presumably etofenprox susceptible brown dog ticks,

Rhipicephalus sanguineus (Ecto Services Inc., Henderson, NC) (hereafter referred to as the susceptible colony) were used in a bioassay to determine the discriminating concentration of etofenprox for this species. Two additional, presumably susceptible, populations of laboratory-reared brown dog ticks from the Centers for Disease Control and Prevention (hereafter referred to as the CDC colony; Atlanta, GA) and Oklahoma

State University (hereafter referred to as the OSU colony; Stillwater, OK) were used to confirm bioassay and molecular assay results. None of the laboratory-reared populations were selected for resistance using acaricides prior to this study. Field- collected peridomestic ticks were obtained from 11 residences experiencing infestations in Europe, North America, Africa, and the Caribbean between May 2016 and February

2017 under U.S. Department of Agriculture-Animal and Plant Health Inspection Service

(USDA-APHIS) (Permit #129047) and Florida (Permit #2015-055) importation permits.

Populations were named by the country from which they were collected or, if more than one collection occurred in a country, the state or city was added. Populations that produced viable eggs and thus were screened in this study were named: Grenada, Italy,

Mexico-Guerrero, Mexico-Merida, and Mexico-Nayarit. Once received, engorged females were placed in plastic vials in an incubator maintained at 85% RH, with temperature held at 25 ± 1°C, and a photoperiod of 12:12 (L:D). After transfer to the incubator, engorged females were left undisturbed to encourage oviposition. Egg masses from the same location were mixed and separated into 500 mg aliquots. Larvae were held at the above parameters for 14 to 16 days after eclosion before testing for acaricide resistance. Once acaricide phenotype was evaluated, individual larvae were used in a molecular assay to determine resistance genotype characteristics.

Bioassay

All larval ticks were kept alive until the Food and Agriculture Organization larval packet test (FAO LPT), first described by Stone and Haydock (1962), was conducted.

Susceptible ticks were pooled into groups of approximately 100 individuals, which were then exposed to one of five concentrations of etofenprox in an olive oil-trichloroethylene mixture (TCE) or a control treatment (olive oil-TCE) with four replicates at each

concentration. This test was conducted four times using the susceptible colony to successfully locate the appropriate concentration range and determine the etofenprox

LC99. The generated LC99 was subsequently evaluated against the susceptible CDC and OSU colonies. Thereafter, the LC99 value was doubled to establish the discriminating concentration (Eiden et al. 2015).

Once the discriminating concentration was established, field-collected ticks were assessed for etofenprox permethrin cross-resistance using the FAO LPT at 1 times (1x),

5x, 16x, and 53x the discriminating concentration of each acaricide. Five populations of field-collected tick strains were screened for etofenprox and permethrin resistance, similar to methods described in Eiden et al. (2015). Three of the five strains, Grenada,

Mexico-Guerrero, and Mexico-Nayarit, produced an adequate quantity of larvae to test the toxicity of permethrin and etofenprox at all screening concentrations. Larval packet test data from wild strains were compared to the susceptible colony to calculate resistance ratios (RR). Assessment of the tick strains for permethrin resistance were conducted using the established 1x (0.19% active ingredient [AI]), 5x (0.95% AI), 16x

(3% AI), and 53x (10% AI) discriminating concentration values for the Ecto Services tick colony previously determined by Eiden et al. (2016).

Technical grade etofenprox (97%; Mitsui Chemicals Inc., Tokyo, Japan) and permethrin (98.8%; ChemService Inc., West Chester, PA) were used concurrently to conduct the LPT. Acaricides were dissolved into a solution of two parts trichloroethylene

(Sigma-Aldrich, St. Louis, MO) and one part olive oil (Spectrum, Gardena, CA). To ensure the acaricide would suspend into solution, acaricide was added to the flask first,

followed by TCE, and lastly olive oil. The solution was vortexed to ensure even acaricide distribution.

Filter papers (Whatman® No. 1, Maidstone, UK) were cut into 7.6 × 8.9 cm pieces and 1 mL of test dilution was applied to a paper. The control filter papers received 1 mL of a 2:1 TCE:olive oil solution. Treated filter papers were suspended from an aluminum restaurant ticket order-holding bar (Alegacy®, Santa Fe Springs, CA) and placed under a fume hood to allow the TCE to evaporate. After a two-hour drying period the papers were folded in half, treated side inwards, and sealed on two sides with bulldog clips

(Boston number 2; Sparco™, Atlanta, GA). Using a watercolor paintbrush, approximately 100 larval ticks were transferred to the packet, which was sealed on the remaining side with a third bulldog clip. The packets were suspended on a metal rod and held at 25 ± 1°C, 92% relative humidity (RH), with a photoperiod of 12:12 (L:D) for

24 hours. Following incubation, packets were opened and mortality assessed over a light box. Dead ticks were categorized by lack of movement and being flat and desiccated in appearance. After completion of the LPT assay, surviving and dead ticks were separated and pooled by acaricide concentration exposure and placed in 1.5 mL microcentrifuge tubes. Ticks were stored at -80°C by post-exposure mortality status within a dose until DNA was extracted.

Polymerase Chain Reaction

DNA isolation

Tick larvae, which previously were sorted into groups of alive or dead at the various etofenprox exposure concentrations and held at -80°C, were removed from group storage vials and prepared for DNA isolation. DNA was extracted from 24 samples (individual ticks) in each population using the technique previously described in

Tucker et al. (in press). Briefly, pooled larvae, from alive or dead larval microcentrifuge tubes, were cooled using liquid nitrogen then transferred to a Petri dish held on ice.

Individual larvae were then transferred from the Petri dish to bead beating tubes held in liquid nitrogen (Fisher scientific, Waltham, MA) containing three 2.0 mm diameter beads

(Biospec Products, Bartlesville, OK). Tubes containing larvae were snap frozen using liquid nitrogen for one minute and larvae were pulverized using a Thermo Savant

FastPrep 120 Cell Disruptor System (Qbiogene Inc., Carlsbad, CA) for three iterations of 20 seconds with 30-second pauses. Pulverized samples received 25 µL of sterile

DNA isolation buffer (3.16 g of 1.0 M Tris HCL to 7.5 g of 1.0 M KCL) and were transferred to a 100°C water bath for five minutes. Lastly, sample tubes were centrifuged at 14,000 rpm and 4°C for five minutes. Extracted DNA was stored at -20°C if not immediately used for amplification by PCR.

DNA amplification

DNA amplification followed the methods developed by Tucker et al. (in press).

Each 20 µL reaction received: 20 pmol of each primer, 0.7 µL of 50 mM MgCl2

(Invitrogen, Carlsbad, CA), 0.5 µL of 10 mM dNTP mix (Thermo Fisher Scientific,

Waltham, MA), 0.1 µL of Platinum® Taq polymerase (Invitrogen, Carlsbad, CA), and 2

µL of 10x PCR reaction buffer (Invitrogen, Carlsbad, CA). Primers BDT-227 and either

RSSC-SUS-F or RSSC-RES-F were used to generate a 69 base pair (bp) amplicon at the site of the point mutation. Two reactions for each sample were needed due to the shared reverse primer that generated a 69 bp ampicon for both primer sets. Primers

FG-228 and BDT-SC-R were used to produce a PCR product for sequencing (Table 3-

1).

The 20 µL mixture was pipetted into a 96 well plate and placed into a Bio-Rad

DNAEngine™ Peltier Thermal Cycler (Bio-Rad Laboratories, Hercules, CA). PCR conditions were as follows: one cycle at 96.0°C for three minutes, followed by 37 cycles of denaturation at 94.0°C for one minute, annealing at 56.5°C for one minute, and extension at 72.0°C for one minute. The conditions also included a final seven minute extension step at 72.0°C. The master mix and PCR conditions for the sequencing reaction were the same as described above except the annealing temperature was

50.0°C. Samples were held at 4.0°C until used for gel electrophoresis.

A 3.5% agarose gel was made by adding 1.4 g of agarose (Bioline, Taunton,

MA), 2.8 g of Synergel™ agarose clarifier additive (Diversified Biotech, Dedham, MA), and 4 µL of 10 mg/mL ethidium bromide solution (Invitrogen, Carlsbad, CA), to 196 mL deionized water mixed with 4 mL of TAE buffer 50x. The mixture was heated until the solution was clear and then the solution was poured into a gel box. The agarose gel was allowed to cool for 30 minutes. An aliquot (5 µL) of exACTGene 100 bp ladder

(Thermo Fisher Scientific, Waltham, MA) was used to determine amplicon size.

PCR reaction product (5 µL) was added to 1 µL of 6x DNA loading dye (Thermo

Fisher Scientific, Waltham, MA), mixed, and added to an individual well in the agarose gel. The gel was run in a Bio-Rad Mini-Sub Cell GT (Bio-Rad Laboratories, Hercules,

CA) at 100 volts for 20 minutes then 110 volts for 10 minutes and photographed under

UV light.

By using both primer sets separately on each sample, presence of a sodium channel mutation conferring cross-resistance to permethrin and etofenprox and/or the wild-type amplicon could be confirmed. Cross-resistance was evaluated by comparing

phenotypic and genotypic results of permethrin and etofenprox bioassays and molecular assays within each tested population.

Statistics

The susceptible strain was used to generate a concentration-response curve for etofenprox, establishing a baseline for comparisons. Larval packet test data for all tested populations were analyzed by standard probit analysis (Finney 1971). Resistance ratios (RR) were calculated to determine the level of resistance within each population, as compared to the laboratory susceptible strain. Resistance ratios were calculated as:

LCx field population/LCx susceptible population. Resistance was defined by a RR of ≥10 and tolerance as a RR of ≥2 and <10 (Gondhalekar et al. 2011).

Results

A total of 10,090 laboratory-reared and presumably susceptible ticks were used in 24 replicates to establish the etofenprox LC99 and discriminating concentration values. The mean LC99 value was 0.254% active ingredient (AI), which, when doubled, provided an etofenprox discriminating concentration of 0.510% AI (Table 3-2). Brown dog tick samples were received from 11 global locations: one population from Cuba,

California, Florida, Grenada, Nigeria, and St. Kitts, two populations from Italy, and three populations from Mexico. Of the 11 peridomestic R. sanguineus populations received, only the Grenada, Italy, Mexico-Guerrero, Mexico-Merida, and Mexico-Nayarit strains provided sufficient numbers of larvae to perform concurrent permethrin and etofenprox bioassays.

Relative to etofenprox, resistance expression to permethrin exposure was higher.

Ticks from Grenada, Mexico-Guerrero, and Mexico-Nayarit expressed resistance at 53 times the permethrin discriminating concentration with 3%, 12%, and 15% of the screened individuals surviving, respectively (Figure 3-1). Resistance to permethrin was not detected in the Italy population where survival was not observed at the discriminating concentration.

Etofenprox proved to be more toxic to tested tick populations than permethrin.

Although four of the five brown dog tick populations tested expressed resistance at the etofenprox discriminating concentration (Table 3-3), only two populations, Grenada

(2%) and Mexico-Guerrero (3%), expressed survival when tested with the 5x etofenprox concentration (Figure 3-2). The individuals screened from the Italy population did not exhibit resistance to etofenprox as they did not survive the discriminating concentration.

Resistance ratios for permethrin and etofenprox were calculated for the Grenada,

Mexico-Guerrero, and Mexico-Nayarit populations (Table 3-4). RR90 values were consistently lower for etofenprox than permethrin across all populations tested. The highest permethrin RR90 of 848.2 was observed in the Mexico-Nayarit population, which corresponded to an etofenprox RR90 of 5.1. The highest etofenprox RR90 of 10.1 was observed in the Mexico-Guerrero population, which had a permethrin RR90 of 215.6, indicating resistance to both chemicals. Resistance ratio values could not be calculated for the Italy population, due to low survival at the discriminating concentration, or the

Mexico-Merida population, because this population was screened only at the discriminating concentration due to a small sample size.

The permethrin resistance-conferring sodium channel SNP was present in the populations collected from Grenada, Mexico-Merida, and Mexico-Nayarit. The mutation was most frequent in the Grenada population where 95% of the screened individuals that survived at the etofenprox discriminating concentration presented a homozygous resistant genotype. Homozygous expression of the mutation also was detected in non- surviving individuals from the Grenada population screened with permethrin and etofenprox at 25% and 10%, respectively. Homozygous expression of the SNP in non- surviving individuals was not observed in any other population.

Discussion

Resistance-conferring mutations or physiological traits are present at low levels prior to the introduction of a new effective acaricide (Food and Agriculture Organization

2004). After several generations of repeated acaricide exposure, heterozygous and subsequently homozygous resistant genotypes can increase in frequency until the genetic makeup of that population is predominantly resistant. However, when an acaricide with a shared mode of action has been used prior to the introduction of a new acaricide, resistance preselection already has occurred and efficacy could be diminished prior to first use (Rongsriyam and Busvine 1975). This is costly not only for the homeowner, but also for companies developing new pesticides. The development of a new acaricide has been estimated to cost US $100 million (Graf et al. 2004). Acaricide resistance can be accelerated by many factors such as the overuse of one class of pesticide, using ineffective concentrations, or using acaricides that remain in the environment at sub-lethal concentrations for a prolonged period (Food and Agriculture

Organization 2004). Limited influx of genetics, and refugia within a home also may contribute to resistance development in R. sanguineus.

The Food and Agriculture Organization larval packet test (FAO LPT) has been used to screen for resistance to multiple pesticides in numerous tick species and is considered a valuable tool in the rapid diagnosis of resistance (Miller et al. 2002,

Ducornez et al. 2005, Mendes et al. 2011). With the establishment of the permethrin, fipronil (Eiden et al.2016), and now etofenprox discriminating concentrations, presumably resistant strains of peridomestic R. sanguineus can be screened against three acaricides using the FAO LPT to make more informed pest management decisions. A limited number of pesticides are currently available for in-home and on- canine host use. Permethrin and to a lesser extent, etofenprox have been used as active ingredients in homes against numerous arthropod pests due to low mammalian toxicity, and high efficacy. Cross-resistance would exacerbate the difficulty in controlling the brown dog tick in and around homes given the relative lack of active ingredients with these restricted application needs.

Resistance ratios (RR) are valuable for comparisons of resistance levels between populations relative to a known susceptible population. The calculated permethrin RR90 values for the Grenada, Mexico-Guerrero, and Mexico-Nayarit populations exceeded a value of 10, which indicates genetically-based resistance. Only the Mexico-Guerrero population had an etofenprox RR90 value of greater than 10. The

Grenada and Mexico-Nayarit RR90 values of 9.1 and 5.1 respectively, indicate a tolerance to etofenprox (Gondhalekar et al. 2011).

The tick populations used in this study were exposed to synthetic pyrethroids

(SP’s), organophosphates (OP’s), or the formamidine amitraz (Table 3-5) and in the case of the Mexico-Guerrero and Mexico-Nayarit populations, ticks were exposed to more than one class of acaricide. The simultaneous resistance to amitraz, SP’s, and

OP’s has been reported in field-collected populations and laboratory-reared colonies of

Rhipicephalus microplus (Canestrini) (Li et al. 2004, Guerrero et al. 2012). Synergism bioassays using triphenylphosphate and piperonyl butoxide indicated that cross- resistance could have been caused by the enhancement of esterases and multifunction oxidases, respectively. However, the resistance mechanism for amitraz has not been elucidated fully and is thought to be polygenic due to the unstable nature of amitraz resistance and diverse larval exposure responses (Jonsson and Hope. 2007).

Permethrin was used infrequently at the feral dog kennel where the Italy population ticks were collected and only as an on premise treatment. The lack of selective pressure on this population likely conserved the susceptible phenotype. The permethrin-resistant and etofenprox-tolerant Grenada population was previously treated with amitraz through on-host applications in a home that experienced a brown dog tick infestation intermittently over five years. The Mexico-Merida and Mexico-Nayarit populations also were collected from homes where an on-host amitraz treatment was used and long-term infestations occurred. Coumaphos, an organophosphate, was used on-host against the Mexico-Merida population. For the Mexico-Guerrero population, when ticks were present in the home, the organophosphate parathion was used on-host and the synthetic pyrethroids cyfluthrin and imiprothrin were used off-host.

Amitraz is a formamidine acaricide that targets octopamine receptors causing tremors and convulsions and has been used to control ectoparasites of dogs, livestock, and parasitic mites of honey bees (Estrada-Pena and Ascher 1999, Wallner 1999, Li et al. 2004). In the Grenada, Mexico-Merida, and Mexico-Nayarit populations that were habitually exposed to amitraz, it is possible that the up-regulation of esterases and oxidases could have contributed to the resistance to permethrin and etofenprox. Amitraz tolerance has been reported in a permethrin resistant Panama-collected R. sanguineus population where a presumed sodium channel mutation and increased esterase activity were present (Miller et al. 2001). In this Panama population, pyrethroids and organophosphates were applied off-host every two weeks for multiple years at the study site in an attempt to control the tick population. Eiden et al. (2017) investigated resistance mechanisms in peridomestic R. sanguineus collected in Florida and Texas that showed high survival at the permethrin discriminating concentration. Through the use of synergist assays, esterase activity and cytochrome P450’s were shown to be the most important resistance mechanisms among the four most resistant of the screened populations. These findings suggest that multi-acaricide cross-resistance caused by metabolic detoxification may be present in the Grenada, Mexico-Merida, and Mexico

Nayarit populations.

Organophosphates irreversibly bind acetylcholinesterase, which is important for normal nerve cell function (Fukuto 1990). Miller et al. (2001) found resistance to coumaphos in R. sanguineus, but synergist bioassays suggested that resistance was primarily conferred by a target site mechanism. The organophosphate resistance mechanism in R. microplus is unknown, but synergist bioassays also suggest target site

resistance as a major factor (Guerrero et al. 2012). A cloning and sequencing study on

R. sanguineus acetylcholinesterase coding regions found them to be homologous to one of the R. microplus coding regions (Xu et al. 2003). Organophosphate metabolic resistance mediated by the up-regulation of esterase and cytochrome P450 has been reported in R. microplus, but typically in the presence of a presumed target site mutation

(Li et al. 2004, Baffi et al. 2008). Although synthetic pyrethroids and organophosphates do not have a shared target site, the on-host use of organophosphates in the Mexico-

Guerrero and Mexico-Merida populations could have contributed to the pyrethroid resistance observed in this study through metabolic detoxification.

The segment VI domain III SNP was not present in all populations tested, but all populations that carried the mutation showed resistance to both permethrin and etofenprox at the respective discriminating concentrations. However, it is perplexing that none of the populations bearing the SNP were from homes that reported synthetic pyrethroid use. It is important to note that only the most recent pesticide use was reported by homeowners and given the common use of synthetic pyrethroids around the world, it is possible that ticks were exposed, and thus selection occurred through previously targeted or environmental pyrethroid treatments allowing for the mutation expression in the Grenada, Mexico-Merida, and Mexico-Nayarit populations. As pyrethroids became ineffective, homeowners likely moved to another product. A previous study has indicated that the segment VI domain III mutation at location 2,134 confers resistance to permethrin at the discriminating concentration (Tucker et al. in press). The Grenada population expressed this mutation in 80% and 95% of surviving ticks exposed to the discriminating concentrations of permethrin and etofenprox,

respectively. However, the mutation was present in 25% and 10% (respectively) of the non-surviving individuals screened at the same concentrations. This suggests multiple additive sodium channel mutations, as has been reported in the horn fly, Haematobia irritans (L.) (Guerrero et al. 1997), and the house fly, Musca domestica L. (Farnham et al. 1987). In H. irritans a knockdown resistance (kdr) mutation conferred 17-fold resistance to permethrin whereas the kdr and super-kdr mutations conferred 688-fold resistance.

Rhipicephalus microplus employs several sodium channel mutations to overcome the toxic effects of synthetic pyrethroids (Morgan et al. 2009, Kumar et al.

2013) and shares a segment VI domain III sodium channel mutation location with R. sanguineus but this mutation differs in the coded amino acid (Guerrero et al. 2001). A domain II SNP, known as super-kdr, which confers resistance to pyrethroids at very high levels in H. irritans and M. domestica and provides a multiplicative effect with other

SNPs, has been identified in R. microplus (Stone et al. 2014). Although Stone et al.

(2014) has identified the super-kdr SNP in R. microplus, no one has shown a synergistic interaction in any Rhipicephalus species. Considering the lack of reported synthetic pyrethroid resistance selection in the Grenada population and the SNP presence in the non-surviving larvae, it is possible that target site insensitivity in R. sanguineus is polygenic. Other sodium channel mutations that work in conjunction with the SNP at location 2,134 also may be present at varying frequencies, which would explain the presence of the resistance conferring mutation in non-surviving larvae.

Based on bioassay and molecular results, it is clear that the SNP previously identified by Tucker et al. (in press) was not the primary resistance mechanism in any of

the Mexico-collected strains. While the mutation was present in the Mexico-Merida and

Mexico-Nayarit surviving larvae, homozygous expression levels were low indicating some other resistance mechanism is acting to provide resistance to the applied acaricides. However, the high level of SNP expression in the Grenada larvae surviving exposure to etofenprox suggests that this mutation confers resistance to both permethrin and etofenprox. Further research should include synergist bioassays to investigate the level of metabolic detoxification of etofenprox in resistant populations and the occurrence of other resistance conferring sodium channel SNP’s.

Table 3-1. Primers used to detect the sodium channel mutation in the brown dog tick, Rhipicephalus sanguineus sensu lato (Latreille). Purpose Primer Name Direction Sequence

Na+ Channel Mutationa RSSC-SUS-F Forward 5’-ATT ATC TTC GGC TCC TTC T-3’

RSSC-RES-F Forward 5’-ATT ATC TTC GGC TCC TTC C-3’

BDT-227 Reverse 5’-TTG TTC ATT GAA ATT GTC AA-3’

Na+ Channel Sequencing FG-228b Forward 5’-CTA ACA TCT ACA TGT ACC-3’

BDT-227a Reverse 5’-GCA ATC CTC CAG CCT TC-3’

Sodium channel mutation located at segment VI domain III is a single nucleotide substitution of cytosine for thymine known to confer permethrin resistance. aGuerrero et al. (2001) bTucker et al. (in press)

Table 3-2. Etofenprox lethal concentration (LC) values for a laboratory-reared Rhipicephalus sanguineus sensu lato (Latreille) (Ecto Services Inc.) colony evaluated using the Food and Agriculture Organization larval packet test. Discriminating a a 2 Test n (replicates) LC50 (95% CI) LC99 (95% CI) Slope (SE) χ (df) concentration 1 2,745 (6) 0.171 (0.167-0.176) 0.280 (0.257-0.318) 10.89 (0.50) 67.10 (18) 0.56

2 3,058 (6) 0.149 (0.138-0.156) 0.209 (0.192-0.253) 15.68 (0.80) 211.47(16) 0.42

3 2,431 (6) 0.123 (0.089-0.144) 0.260 (0.245-0.290) 7.14 (1.00) 23.76 (18) 0.52

4 1,856 (6) 0.195 (0.189-0.199) 0.231 (0.224-0.246) 31.11 (2.20) 82.65 (18) 0.51

Pooled total 10,090 (24) 0.162 (0.157-0.166) 0.254 (0.241-0.273) 11.89 (0.26) 936.35 (76) 0.51

LC values determined using probit analysis. Discriminating concentration is double the LC99 value. a Percent (%) active ingredient applied to filter papers 7.5 × 8.5 cm. n=number of larvae tested. Replicates=total number of packets. CI=confidence interval, SE=standard error, df=degrees of freedom, χ2=Chi square.

Table 3-3. Association between phenotypic and genotypic resistance to a discriminating concentration of either permethrin or etofenprox in peridomestic Rhipicephalus sanguineus sensu lato (Latreille).

% Survival at Tick exposure resulta discriminating Died Survived Colony/Population Acaricide concentration (n) SS SR RR SS SR RR CDCb Permethrin 0 (410) 100 0 0 NT NT NT Etofenprox 0 (352) 100 0 0 NT NT NT Ecto Servicesb,c Permethrin 0 (ND) 100 0 0 NT NT NT Etofenprox 0 (ND) 100 0 0 NT NT NT OSUb Permethrin 0 (389) 100 0 0 NT NT NT Etofenprox 0 (421) 100 0 0 NT NT NT Grenada Permethrin 77 (624) 75 0 25 10 10 80 Etofenprox 41 (530) 85 5 10 0 5 95 Italy Permethrin 0 (444) 100 0 0 NT NT NT Etofenprox 0 (429) 100 0 0 NT NT NT Mexico-Guerrero Permethrin 69 (862) 100 0 0 100 0 0 Etofenprox 74 (793) 100 0 0 100 0 0 Mexico-Merida Permethrin 98 (232) NT NT NT 100 0 0 Etofenprox 61 (267) 100 0 0 70 20 10 Mexico-Nayarit Permethrin 78 (646) 100 0 0 90 5 5 Etofenprox 48 (435) 80 20 0 NT NT NT Tick resistance status was evaluated using the Food and Agriculture Organization larval packet test. Permethrin discriminating concentration (0.19% active ingredient). Etofenprox discriminating concentration (0.51% active ingredient). aGenotype (%) was determined using 20 individuals from each survival status in molecular assay developed by Tucker et al. (in press). bLaboratory reared, presumably acaricide susceptible. cEcto Services Inc. colony was used to calculate discriminating concentration and, therefore, not tested specifically against the discriminating concentration. SS=homozygous susceptible. SR=heterozygous. RR=homozygous resistant. n=sample size. NT=not tested. ND=no data. CDC=Centers for Disease Control and Prevention. OSU=Oklahoma State University.

Table 3-4. Toxicity of permethrin and etofenprox to peridomestic Rhipicephalus sanguineus sensu lato (Latreille) collected in Grenada and Mexico.

a b a Strain Acaricide n LC50 (95% CI) LC90 (95% CI) RR50 RR90 Slope (SE) Ecto Services Permethrin† 5,240 0.027 (0.026-0.028) 0.051 (0.048-0.054) 1.0 1.0 4.64 (0.15) Etofenprox 10,090 0.162 (0.157-0.166) 0.254 (0.241-0.273 1.0 1.0 11.89 (0.26) Grenada Permethrin 2,137 0.641 (0.462-0.861) 3.877 (2.595-7.006) 23.7 76.0 1.64 (0.06) Etofenprox 1,518 0.470 (0.169-0.761) 1.894 (1.155-5.885) 2.9 9.1 1.92 (0.13) Mexico-Guerrero Permethrin 1,280 4.382 (3.745-5.198) 10.994 (8.435-17.324) 162.3 215.6 3.21 (0.18) Etofenprox 2,203 0.848 (0.725-1.009) 2.105 (1.647-3.035) 5.2 10.1 3.25 (0.15) Mexico-Nayarit Permethrin 1,984 1.885 (1.333-2.832) 43.262 (19.709-154.861) 69.8 848.2 0.94 (0.05) Etofenprox 1,710 0.506 (0.470-0.541) 1.053 (0.922-1.289) 3.1 5.1 4.03 (0.46) Tick resistance status was evaluated using the Food and Agriculture Organization larval packet test. a Permethrin lethal concentration (LC) values determined by Eiden et al. (2015). b Values represent % active ingredient applied to filter papers 7.5 × 8.5 cm. LC values determined using probit analysis. † Values determined by Eiden et al. (2015). n=number of ticks tested. RR=resistance ratio. CI=confidence interval, SE=standard error.

Table 3-5. Acaricide exposure in peridomestic adult Rhipicephalus sanguineus sensu lato (Latreille) populations evaluated with a larval packet test as reported by pet owners or kennel operators.

Population Acaricide (application type) Acaricide class Frequency Grenada Amitraz (on-host) Formamidine Every other month Italy Permethrin (off-host) Synthetic pyrethroid Annually Mexico-Guerrero Parathion (on-host) Organophosphate When pests present Cyfluthrin (off-host) Synthetic pyrethroid When pests present Imiprothrin (off-host) Synthetic pyrethroid When pests present Mexico-Merida Amitraz (on-host) Formamidine Every other month Coumaphos (on-host) Organophosphate Every other month Mexico-Nayarit Amitraz (on-host) Formamidine Every other month

100

%SURVIVAL 40

0 Grenada Italy Mexico-Guerrero Mexico-Merida Mexico-Nayarit

Permethrin 53x Permethrin 16x Permethrin 5x Permethrin 1x

Figure 3-1. Survival (%) of Rhipicephalus sanguineus sensu lato (Latreille) larvae collected from five globally distributed locations following exposure to 1, 5, 16, and 53 times (x) the permethrin discriminating concentration (0.19% permethrin; Eiden et al. 2016) using the Food and Agriculture Organization larval packet test.

100

%SURVIVAL 40

0 Grenada Italy Mexico-Guerrero Mexico-Merida Mexico-Nayarit

Etofenprox 53x Etofenprox 16x Etofenprox 5x Etofenprox 1x

Figure 3-2. Survival of Rhipicephalus sanguineus sensu lato (Latreille) larvae collected from five globally distributed locations and exposed to 1, 5, 16, and 53 times (x) the etofenprox discriminating concentration (0.51% etofenprox) using the Food and Agriculture Organization larval packet test.

CHAPTER 4 PREVALENCE AND DISTRIBUTION OF PATHOGEN INFECTION AND PERMETHRIN RESISTANCE IN TROPICAL AND TEMPERATE POPULATIONS OF RHIPICEPHALUS SANGUINEUS SENSU LATO (LATREILLE) COLLECTED FROM SITES AROUND THE WORLD

Introduction

The brown dog tick, Rhipicephalus sanguineus (Latreille), is an endophilic pest of dogs and occasionally humans, and is found worldwide (Dantas-Torres 2008). Likely due to its cosmopolitan distribution, strong association with its canine host, and general hardiness, this tick is the primary vector for several pathogens of medical and veterinary importance and it has been suggested as a potential vector for many others (Dantas-

Torres 2010, Dantas-Torres and Otranto 2015). There is contention over the classification of the brown dog tick as a single species, due in part to its worldwide distribution (Gray et al. 2013, Nava et al. 2015). Morphological and molecular comparisons have been completed and indicate that there are multiple lineages within the R. sanguineus species complex, however, more research is needed to fully understand the relationships within this group (Jones et al. 2017). The current understanding, from phylogenetic analysis, is that R. sanguineus may be a species group within which separate clades occupy distinct latitudinal ranges: temperate or tropical. A cross-breeding study on R. sanguineus between populations from North

America, Israel, and Africa found that ticks from North America and Israel successfully interbred, but when either was bred with the African tick population, the resulting offspring were infertile (Levin et al. 2012). Furthermore, phylogenetic analysis of ribosomal DNA placed the African ticks from the cross-breeding study in a separate

clade. Such relationships may have considerable impact on pathogen transmission and global distribution of acaricide resistance mechanisms.

The brown dog tick is a capable vector of Rickettsia conorii in the Eastern

Hemisphere and Ri. rickettsii in the Western Hemisphere. Both pathogens can be transovarially transmitted to their progeny. Socolovschi et al. (2009) found that 94.3% of larvae were infected with Ri. conorii after four filial generations without re-inoculation.

Rickettsia conorii, the etiological agent of Mediterranean spotted fever, is the most important tick-borne pathogen vectored in southern Europe and northern Africa (Parola et al. 2009). Rickettsia conorii is typically vectored by R. sanguineus, whereas Ri. rickettsii, the causative agent of Rocky Mountain spotted fever, is typically vectored by

Dermacentor spp. and seldom by R. sanguineus. Rocky Mountain spotted fever is an important tick-borne disease in the U.S. with case fatality rates as high as 20% in untreated patients (Demma et al. 2005). The first account of Ri. rickettsii vectored by R. sanguineus was reported by Demma et al. (2005).

Another rickettsial pathogen of medical and veterinary importance is Ehrlichia canis. Canine monocytotrophic ehrlichiosis is a potentially fatal, globally distributed canine disease caused by E. canis and primarily vectored by R. sanguineus (Murphy et al. 1998, Harrus and Waner. 2011). Dogs are considered the main reservoir for E. canis as this pathogen is not typically transmitted transovarially by its tick host (Aguiar et al.

2007). This pathogen also has zoonotic potential causing symptoms similar to those observed with E. chaffeensis, the etiological agent of human monocytic ehrlichiosis, in humans (Perez et al. 2006). In highly endemic areas, serology has confirmed ehrlichial exposure rates among dogs as high as 43.8% (Yabsley et al. 2008). A study by Koh et

al. (2015) investigated ehrlichial infection rates of R. sanguineus collected in peninsular

Malaysia from stray dogs and found 51.5% of tick samples to be infected by E. canis.

Canine babesiosis is a tickborne disease of worldwide importance. The causative agents of canine babesiosis are strongly associated with a specific tick vector and each protozoan parasitic subspecies presents unique clinical symptoms. The babesial pathogen typically vectored by the brown dog tick is Babesia canis vogeli, which often results in mild disease symptoms without clinical signs (Solano-Gallego et al. 2008).

Babesia canis vogeli is the least clinically-significant pathogen reviewed here, but is relevant considering the distribution and prevalence of its vector.

Rhipicephalus sanguineus is also the primary vector of Hepatozoon canis. Unlike many other tick-borne pathogens, H. canis is not transmitted in the ectoparasite saliva but instead enters the host by consumption of the infected tick during grooming (Baneth et al. 2007). While the pathogen is not typically transovarially transmitted, transtadial transmission in the tick, and vertical transmission in the canine host have been observed (Murata et al. 1993).

Important for the preclusion of the aforementioned pathogens is the use of pesticides to kill or deter their vector. The use of non-selective pyrethroid treatments to control other domestic arthropod pests likely has contributed to the selection for resistance conferring traits in R. sanguineus. Pyrethroids have been used extensively in the control of injurious arthropods on canine hosts, particularly for flea control. Fleas, ticks, and mosquitoes, can pose a risk beyond annoyance and psychological distress due to the microorganisms they often harbor and transmit. Unfortunately, repeated efforts to control an arthropod using a pesticide often leads to pesticide resistance

selection. Due to its common use and widespread availability, resistance to permethrin at doses as high as 53x the concentration expected to kill R. sanguineus larvae were observed in several Florida populations (Eiden et al. 2015). A single nucleotide polymorphism (SNP) conferring resistance to permethrin was identified on domain III segment VI of the R. sanguineus sodium channel (Klafke et al. 2017). This mutation was found to be present in eight North American populations with demonstrated phenotypic resistance (Tucker et al. in press).

Due to the severity of rickettsial infections (Treadwell et al. 2000), zoonotic potential of these pathogens, and reported occurrences of R. sanguineus acaricide resistance, further investigation of pathogen and resistance prevalence was warranted.

Using brown dog tick samples submitted from locations around the world, I compared pathogen presence, and a presumed pyrethroid resistance mechanism, among ticks obtained from temperate and tropical latitudes. Thus, this study is a temporal snapshot examining permethrin resistance and pathogen presence in peridomestic populations of

R. sanguineus collected worldwide.

Materials and Methods

Ticks

A tick request form and data sheet (Appendix A) were distributed through the

U.S. Army Veterinary Corps, the U.S. Department of Agriculture, universities, and pest control companies located throughout the western hemisphere in: the Caribbean,

Central and South America, Canada, and several U.S. states. A more restricted request was made to individuals in Asia, Europe, Africa, and Australia. Participants were asked to collect adult blood-fed ticks from infested homes and kennels or preferably, from dogs, and ship these live ticks with completed importation forms (USDA-APHIS Permit

#129047 and Florida Permit #2015-055) to the Veterinary Entomology Laboratory,

Entomology and Nematology Department, University of Florida (Gainesville, FL, U.S.).

Live tick samples from 14 distinct locations representing the Caribbean, Europe, and

North America were received. An additional eight Florida and Texas populations previously described by Tucker et al. (in press) and six populations provided by the

Centers for Disease Control and Prevention (Atlanta, GA) as DNA extracts and preserved samples from Africa, Asia, Europe, and North America were used in this study. Of the 14 live populations received, nine produced viable offspring in sufficient numbers for resistance and pathogen molecular testing. Only the adult females were tested in the remaining five populations.

Once received, engorged females were placed individually in vials held in an incubator maintained at 85% relative humidity and 25 ± 1°C, and a photoperiod of 12:12

(L:D). After transfer to the vial, engorged females were left undisturbed to encourage oviposition. Upon completion of oviposition, females were stored at -80°C until DNA was extracted. Larvae were held for 14 to 16 days post-eclosion prior to conducting DNA extraction procedures.

Morphological identification was conducted using keys published by Filippova

(1997) and Walker et al. (2000). Eleven of the tick populations were previously typed by geographical zone using morphological identification and molecular techniques by the

Centers for Disease Control and Prevention (Atlanta, GA) or Georgia Southern

University (Statesboro, GA) using procedures outlined in Zemtsova et al. (2016).

Polymerase chain reactions (PCRs) were performed using primers and cycling

conditions for the 12S and 16S gene targets as described by Beati and Keirans (2001) and Mangold et al. (1998), respectively.

Molecular Analysis

DNA isolation

Using 20 individual larvae from each population, DNA was extracted using a technique described by Tucker et al. (in press). In brief, larvae were snap frozen in liquid nitrogen for 1 minute and transferred onto a Petri dish held on ice. Bead beating tubes (Fisher Scientific, Waltham, MA) containing three 2.0 mm diameter beads

(BioSpec Products, Bartlesville, OK) were chilled on liquid nitrogen for one minute prior to the transfer of individual larvae. Tubes containing larvae were then returned to liquid nitrogen and held for 1 minute until disruption by bead beating was conducted.

Ticks were pulverized in bead beating tubes using a Thermo Savant FastPrep

120 Cell Disruptor System (Qbiogene Inc., Carlsbad, CA) for three iterations of 20 seconds with 30 second pauses in between. Samples were returned to liquid nitrogen after disruption until receiving 25 µL of sterile DNA isolation buffer (3.16 g of 1.0 M Tris

HCL to 7.5 g of 1.0 M KCL) followed by being placed in a 100°C water bath for five minutes. Lastly, sample tubes were placed in a 4°C centrifuge at 14,000 rpm for five minutes. Extracted DNA was stored at -20°C if not immediately used in a molecular assay.

DNA amplification

To determine genotypic resistance status, amplification of a SNP known to confer resistance to permethrin in R. sanguineus followed methods developed by Tucker et al.

(in press). Each 20 µL reaction received: 2 µL of DNA, 20 pmol of each primer, 0.7 µL of

50 mM MgCl2 (Invitrogen, Carlsbad, CA), 0.5 µL of 10 mM dNTP mix (Thermo Fisher

Scientific, Waltham, MA), 0.1 µL of Platinum® Taq polymerase (Invitrogen, Carlsbad,

CA), and 2 µL of 10x PCR reaction buffer (Invitrogen, Carlsbad, CA). Each sample required two reactions due to the shared reverse primer binding site that each produced a 69 base pair (bp) amplicon. BDT-227 was used in both reactions, whereas RSSC-

SUS-F was used to screen for susceptible alleles and RSSC-RES-F was used to screen for resistant alleles (Table 4-1). The 20 µL mixture was pipetted into a 96 well plate and placed into a Bio-Rad DNAEngine™ Peltier Thermal Cycler (Bio-Rad Laboratories,

Hercules, CA). PCR conditions were as follows: 1 cycle at 96°C for 3 minutes, followed by 40 cycles at 94°C for 1 minute, 59°C for 1 minute, and 72°C for 1 minute with a final

7 minute extension step at 72°C. Samples were held at 4°C until used for gel electrophoresis. A 3% agarose gel with Synergel™ agarose clarifier additive (Diversified

Biotech, Dedham, MA) and ethidium bromide solution was mixed. Using FastRuler Ultra

Low Range DNA Ladder (Thermo Fisher Scientific, Waltham, MA) and 6x DNA loading dye (Thermo Fisher Scientific, Waltham, MA), DNA fragments were visualized and photographed under UV light.

A traditional PCR was used to screen for Rickettsia based on methods developed by Kidd et al. (2008). Each 50 µL reaction received: 5 µL of DNA, 30 pmol of each primer, and 25 µL of PCR mastermix® (Promega, Madison, WI). Primers 107F and

299R were used to produce a 209 to 212 bp amplicon of the conserved Rickettsia ompA region (Table 4-1). The 50 µL mixture was pipetted into a 96 well plate and placed into a

Bio-Rad DNAEngine™ Peltier Thermal Cycler (Bio-Rad Labratories, Hercules, CA).

PCR conditions were as follows: an initial denaturation step at 95°C, followed by 40 cycles of 95°C for 30 seconds, 56°C for 45 seconds, 72°C for 30 seconds, and one final

cycle of 72°C for 7 minutes. Samples were held at 4°C until used for gel electrophoresis. PCR product and Trackit™ 50 bp ladder (Invitrogen, Carlsbad, CA) were loaded onto a 2% agarose gel with ethidium bromide and photographed under UV light. Positive samples were submitted for Sanger sequencing to Genewiz (South

Plainfield, NJ) and sequenced in forward and reverse directions and compared to known sequences in GenBank using the Basic Local Alignment Search Tool (BLAST).

Primers developed by Doyle et al. (2005) were used in a traditional PCR to screen for Ehrlichia canis. The primers DSB-321 and DSB-671 were used to produce a

378 bp amplicon of the Ehrlichia dsb gene. Each 20 µL reaction received: 2 µL of DNA,

400 nmol/L of each primer, 10 µL of iQ Supermix (Bio-Rad Laboratories, Hercules, CA), and final concentration of 4 mmol/L MgCl2. The 20 µL mixture was pipetted into a 96 well plate and placed into a Bio-Rad DNAEngine™ Peltier Thermal Cycler (Bio-Rad

Labratories, Hercules, CA). PCR conditions consisted of initial denaturation of one cycle at 95°C for 5 minutes, followed by 40 cycles of denaturation at 95°C for 15 seconds, annealing at 60°C for 1 minute, and extension at 72°C for 1 minute. The conditions also included a final seven minute extension step at 72°C. Samples were held at 4°C until used for gel electrophoresis. The samples were loaded onto a 2% agarose gel with ethidium bromide and TrackIt™ 50 bp DNA ladder (Invitrogen, Carlsbad, CA) was used to approximate amplicon size. Positive samples were submitted for Sanger sequencing to Genewiz (South Plainfield, NJ) and sequenced in forward and reverse directions and compared to known sequences in GenBank using BLAST.

Primers PIRO-A1 and PIRO-B were used in a traditional PCR (Földvári et al.

2005) to screen for Hepatozoon and Babesia to amplify a 450 bp region of the 18s

rRNA (Table 4-1) (Promega, Madison, WI). Each 50 µL reaction received: 2 µL of DNA,

25 pmol of each primer, and 25 µL of PCR mastermix® (Promega, Madison, WI). The 50

µL mixture was pipetted into a 96 well plate and placed into a Bio-Rad DNAEngine™

Peltier Thermal Cycler (Bio-Rad Labratories, Hercules, CA). PCR conditions consisted of initial denaturation of one cycle at 94°C for 10 minutes, followed by 40 cycles of denaturation at 94°C for 30 seconds, annealing at 59°C for 45 seconds, and extension at 72°C for 30 seconds. The conditions also included a final seven minute extension step at 72°C. Samples were held at 4°C until used for gel electrophoresis. The samples were loaded onto a 2% agarose gel with ethidium bromide and TrackIt™ 50 bp DNA ladder (Invitrogen, Carlsbad, CA) was used to approximate amplicon size. Positive samples were submitted for Sanger sequencing to Genewiz (South Plainfield, NJ) and sequenced in forward and reverse directions and compared to known sequences in

GenBank using BLAST.

Results

A total of 458 individual peridomestic R. sanguineus from 28 populations were screened for pyrethroid resistance and Rickettsia, Ehrlichia, Babesia and Hepatozoon presence (Table 4-2). The pyrethroid resistance conferring SNP was isolated in ticks from Florida, Texas, Cuba, and St. Kitts populations. These resistant populations were genetically-typed as tropical. However, the susceptible allele was present in the tropical ticks tested from Africa (i.e. Burkina Faso and Ghana), and Asia (i.e. Thailand). The resistance-conferring SNP was not isolated in any temperate field-collected tick population.

Sequencing provided verification of pathogen presence in 17 tick strains.

Pathogen sequences showed homology of 98% or above with those stored in GenBank. 81

Ehrlichia canis (GenBank Accession # KY576856.1) was the most common pathogen detected and was present in the Burkina Faso, Ghana, Thailand, Italy 1, Italy 2,

California, Florida-Maitland, Mexico-Merida 1, Mexico-Merida 2, Mexico-Mexicali,

Mexico-Nayarit 2, St. Kitts 1, and St. Kitts 3 populations. Rickettsia massiliae (GenBank

Accession # KY440239.1) was detected and isolated in eight strains, three of which were from Florida-originating tick populations. Ehrlichia canis and Ri. massilae were detected in both tropical and temperate tick types. Rickettsia rickettsii (GenBank

Accession # KX363464.1) was isolated in the Florida-Palm Beach 2 strain in one of the

20 larval ticks tested and one of the 10 adults tested from the Mexico-Mexicali strain.

Rickettsia conorii was identified in two Thailand samples. Hepatozoon sp. was detected in three out of 20 Burkina Faso samples screened, two of which sequenced as

Hepatozoon sp. (GenBank Accession # KJ499492.1) and the third as H. canis

(GenBank Accession # KT246304.1). No tick samples tested positive for Babesia.

Discussion

In order to fully understand epidemiological interactions of arthropod-borne pathogens, it is important to accurately identify the vector. In the case of R. sanguineus, this can be difficult and time consuming, as morphological characters are not always reliable and an investigator must rely on multi-step molecular techniques. To add to the confusion, the R. sanguineus species complex has been divided into six groups consisting of: R. sanguineus sensu lato (s.l.), R. turanicus, and Rhipicephalus spp. I, II,

III, and IV (Dantas-Torres and Otranto 2015). Rhipicephlaus sanguineus s.l. is stated as corresponding to the tropical lineage and R. sanguineus II corresponds to the temperate lineage. However, not all taxonomists share this view and some authors maintain that the species complex should be identified as R. sanguineus s.l. temperate and R. 82

sanguineus s.l. tropical (Zemtsova et al. 2016). The current study conforms to the latter and more simplified two-group description.

Other authors have investigated the R. sanguineus geographic species groups and associated pathogens. Latrofa et al. (2014) investigated the presence of multiple pathogens including Circopithfilaria species, H. canis, and Anaplasma platys in association with the tick collection geographic zone. In that study, H. canis was found in several temperate samples, but was not detected in any tropical samples. The detection of H. canis in tropical-type samples in the current study contradicts the findings of

Latrofa et al. (2014). It is important to note that the Latrofa et al. (2014) study did not investigate more central African-collected tropical ticks from Burkina-Faso, only Algeria and South Africa, which could explain the differences in pathogen detection.

An interesting finding was the geographical distribution of detected rickettsial pathogens. Rickettsia and Ehrlichia were detected in samples from all five sampled geographic regions and were in both the temperate and tropical lineages. The isolation of Rickettsia rickettsii in the Florida-Palm Beach 2 strain is a first report of this pathogen in Florida. A foci of Ri. rickettsii with documented R. sanguineus transmission in

Mexicali, Mexico was investigated by Eremeeva et al. (2011). The authors placed both the R. sanguineus and Ri. ricketsii in clades geographically distinct from cases in nearby Arizona, which is interesting considering the frequent movement of migrants and animals across the United States-Mexico border (Eremeeva et al. 2011). Additionally, the Mexicali ticks were identified as tropical, while the Arizona ticks were temperate.

The brown dog tick is also a capable vector of Ri. massiliae in the Mediterranean and the western hemisphere and the pathogen can be transovarially transmitted to

progeny. Matsumoto et al. (2005) reported that 98% of larvae were infected with Ri. massiliae after four filial generations without re-inoculation. Rickettsia massiliae is typically vectored by R. sanguineus or R. turanicus and has been isolated in ticks in

Argentina (Cicuttin et al. 2014), France (Beati and Raoult 1993), Israel (Harrus et al.

2011), Switzerland (Bernasconi et al. 2002), Arizona (Eremeeva et al. 2006), and

California (Beeler et al. 2011). To our knowledge, the isolation of Ri. massiliae DNA in three of the Florida populations is a first report of this pathogen in Florida. The pathogenicity of Ri. massiliae in dogs is unknown. Beeler et al. (2011) reported on two dogs in California showing clinical signs of spotted fever group rickettsiae that tested seropositive for Rickettsia antibodies and were exposed to heavy infestations of Ri massiliae-infected R. sanguineus. Although rickettsial DNA was not detected in the dogs’ serum, the authors suggested that R. massiliae may cause infection in dogs.

Rickettsia massiliae has been isolated from humans showing clinical signs of rickettsial infection in Italy (Vitale 2006), France (Parola et al. 2008), and Argentina

(García-García et al. 2010). The first isolation of Ri. massiliae from a human in Italy was presumed be a R. conorii infection until molecular analysis of the isolate 20 years later.

Considering that Ri. massiliae presents similar symptoms in humans to other rickettsioses and has been isolated in ticks where Ri. rickettsii and Ri. conorii are reported it is possible that this widespread pathogen is under-reported in spotted fever group cases. Also, considering that spotted fever group rickettsioses have been reported in Florida (Drexler et al. 2016) it is possible that R. sanguineus is contributing to the prevalence of these rickettsial pathogens.

This study represents the first evaluation of the geographic distribution of the domain III segment VI pyrethroid resistance conferring SNP. Considering that the SNP was isolated only in tropical ticks from North America and the Caribbean it is possible that with further research, a correlation could be drawn between the two factors.

However, without detailed treatment records of the evaluated tick populations, it is difficult to confirm if the lack of SNP presence is truly associated with the geographic zone or due to lack of pyrethroid exposure. It is important to note that when the Mexico-

Merida and Mexico-Nayarit populations were bioassayed using permethrin or etofenprox (Chapter 3), the SNP was subsequently detected at low levels using PCR.

Whereas, when those same populations were tested using individuals not previously screened for resistance, the mutation was not detected. Thus it is possible that the SNP may be present at low levels in other populations.

Both temperate and tropical ticks were identified in Texas. The Texas-Webb population, which was collected near the United States-Mexico border in southern

Texas, was typed as tropical and the resistance conferring SNP was detected. The

Texas-Central strain, which typed as temperate, was collected near San Antonio,

Texas, and did not express the SNP. This provides support to the hypothesis that the domain III segment VI SNP is present only in tropical R. sanguineus.

Target site insensitivity is not the only acaricide resistance mechanism employed by R. sanguineus. Several studies have implicated metabolic detoxification through up- regulation of esterases and cytochrome P450’s as major contributors to pyrethroid resistance in R. sanguineus (Miller et al. 2001, Eiden et al. 2017). However, a molecular assay has not yet been developed to identify these mechanisms.

While the development of resistance mechanisms provides protection against lethal acaricides, they also likely incur a fitness cost. Rivero et al. (2010) discussed factors that influence insect vector competence and its interaction with resistance. They found that in some instances, resistance likely results in a decrease in vector longevity, decrease in infectiousness, or a change in behavior (Rivero et al. 2010). The upregulation of detoxifying esterase enzymes requires a shift in protein and lipid resources that would otherwise be used for normal insect metabolism (Rivero et al.

2011). In the house fly, Musca domestica L., oxidative stress due to increased activity of

P450 monoxygenases has been observed (Murataliev et al. 2008). Pesticide resistance also may affect the vector-pathogen interaction by rendering the vector toxic to the pathogen through excess esterase production, making pesticide-resistant vectors immune to the pathogen, or reducing growth of the pathogen due to a reduction in available resources (Rivero et al. 2010). Target site resistance expression alters the arthropod nervous system and can disrupt normal behavior of the vector. Sodium channel target site resistance has been observed to disrupt oviposition and predator cue detection in anopholine mosquitoes (Rowland 1991). Metabolic resistance also may affect vector behavior by decreasing locomotive performance (Berticat et al. 2004).

Interestingly, pathogens were detected in tropical and temperate strains and strains with and without the resistance conferring SNP. There does not appear to be an association between the presence of the resistance conferring SNP and pathogen infection in R. sanguineus. Although the metabolic resistance status is unknown in some of the populations bearing the SNP, the eight Florida and Texas populations described by Tucker et al. (in press) and assayed with synergists by Eiden et al. (2017)

also employed metabolic mechanisms to overcome toxicants. The data described in

Table 4-2 suggests that the domain III segment VI SNP is isolated to North America and the Caribbean in tropical-type R. sanguineus and that pyrethroid resistance expression does not affect pathogen infection. The R. sanguineus sodium channel should be investigated further to allow description of other resistance conferring mutations that may be present.

Table 4-1. Primers used to detect sodium channel mutations and pathogens in the brown dog tick, Rhipicephalus sanguineus sensu lato (Latreille).

Purpose Primer Name Direction Sequence

Na+ Channel Mutationa RSSC-SUS-F Forward 5’-ATT ATC TTC GGC TCC TTC C-3’

RSSC-RES-F Forward 5’-ATT ATC TTC GGC TCC TTC T-3’

BDT-227 Reverse 5’-TTG TTC ATT GAA ATT GTC AA-3’

Rickettsia detectionb 107F Forward 5’-GCT TTA TTC ACC ACC TCA AC-3’

299R Reverse 5’-TRA TCA CCA CCG TAA GTA AAT-3’

Babesia and Hepatozoon PIRO-A1 Forward 5’-AGG GAG CCT GAG AGA CGG CTA CC-3’ detectionc

PIRO-B Reverse 5’-TTA AAT ACG AAT GCC CCC AAC-3’

Ehrlichia detectiond DSB-321 Forward 5’-TTG CAA AAT GAT GTC TGA AGA TAT GAA ACA-3’

DSB-671 Reverse 5’-GCT GCT CCA CCA ATA AAT GTA TCY CCT A-3’ a Primers developed by Tucker et al. (in press). b Primers developed by Kidd et al. (2008). c Primers developed by Földvári et al. (2005). d Primers developed by Doyle et al. (2005).

Table 4-2. Occurrence of genotypic pyrethroid resistance and infection with Rickettsia and Hepatozoon parasites in peridomestic Rhipicephalus sanguineus sensu lato (Latreille) collected in Africa, Asia, Europe, North America, and the Caribbean.

Samples Tick Genotype (%)† Detected (prevalence) Region Strain (Stage) type SS SR RR Rickettsia - Hepatozoon - Ehrlichia Africa Burkina Faso 17(A) 3(N) Tropicala 100 0 0 R. massiliae (55%), H. canis (30%), E. canis (20%) Ghana 10(A) Tropicala 100 0 0 E. canis (10%) Asia Thailand 10(A) Tropicala 100 0 0 R. conorii (20%), R. amblyommii (10%) E. canis (30%) Europe Italy 1 10(A) Temperateb 100 0 0 R. massiliae (90%), E. canis (50%) Italy 2 11(A) Temperateb 100 0 0 R. massiliae (10%), E. canis (45%) Spain 20(A) Temperatea 100 0 0 R. massiliae (20%) North California 1(A) Temperateb 100 0 0 E. canis (100%) America Florida-Broward* 20(L) Tropicala 5 0 95 R. massiliae (5%) Florida-Dunnellon 22(L) Tropicalc 0 0 100 - Florida-Gilchrist* 21(L) Tropicala 0 0 100 - Florida-Maitland 20(A) Tropicalc 0 5 95 R. massiliae (30%), E. canis (55%) Florida-Palm Beach 1* 21(L) Tropicalc 5 5 90 - Florida-Palm Beach 2* 22(L) Tropicala 5 0 95 R. rickettsii (4%), R. massiliae (4%) Florida-Palm Beach 3* 21(L) Tropicala 0 0 100 - Florida-Port Richey 22(L) Tropicalc 0 0 100 - Florida-Sarasota 2* 20(L) Tropicalc 0 0 100 - Florida-St. Johns* 20(L) Tropicala 0 0 100 - Mexico-Merida 1 1(A) 20(L) Tropicalb 100 0 0 E. canis (100%) Mexico-Merida 2 1(A) 20(L) Tropicalb 100 0 0 E. canis (100%) Mexico-Mexicali 10(A) Tropicala 100 0 0 R. rickettsii (10%), E. canis (10%) Mexico-Nayarit 2 1(A) 20(L) Tropicalb 100 0 0 E. canis (100%) Mexico-Nayarit 4 1 (A) 20 (L) Tropical 100 0 0 R. amblyommii (5%) Texas-Central 10(A) Temperatea 100 0 0 -

Table 4-2. Continued

Samples Tick Genotype (%)† Detected (prevalence) Region Strain (Stage) type SS SR RR Rickettsia - Hepatozoon - Ehrlichia North Texas-Webb* 20(L) Tropicala 0 25 75 - America Caribbean Cuba 1(A) Tropicalb 0 0 100 - St. Kitts 1 1(A) 20(L) Tropicalb 0 0 100 E. canis (100%) St. Kitts 2 1(A) 20(L) Tropicalb 0 0 100 - St. Kitts 3 1(A) 20(L) Tropicalb 0 0 100 R. massiliae (5%), E. canis (100%) a Tick species typing conducted by Centers for Disease Control and Prevention (Atlanta, GA) or Georgia Southern University (Statesboro, GA). b Tick type classification determined by latitude of collection site. c Presumed tick classification based on proximity to previously typed samples. † Molecular assay for screening sodium channel permethrin resistance developed by Tucker at al. (in press). * Previously described by Tucker et al. (in press). A=adult, N=nymph, L=larva. SS=homozygous susceptible. SR=heterozygous. RR=homozygous resistant. -=pathogen not detected All larvae reared from submitted blood-fed females. Adults and nymphs were field-collected.

CHAPTER 5 IMPLICATIONS AND FUTURE DIRECTIONS FOR RHIPICEPHALUS SANGUINEUS RESEARCH

Rhipicephaus sanguineus sensu lato (Latreille), the brown dog tick, is a peridomestic pest of veterinary and medical importance with a cosmopolitan distribution

(Dantas-Torres 2008). Based on morphological and molecular identification, R. sanguineus is classified as a species group and divided into at least two types. The most widely accepted definition is a split between temperate, or Mediterranean origin, and tropical, or African origin. Although both tick sub-types are found worldwide, differences in vector competence and resistance status have not been fully elucidated.

Additionally, the brown dog tick is the only known tick to complete its life cycle indoors, particularly in temperate regions (Dantas-Torres 2008). This endophilic behavior has allowed for increased exposure to acaricides used both in the environment (residence or kennel) and on the canine host. These ticks spend greater than 95% of their lifespan in their non-host environment and dogs serve as this ticks’ primary dispersal mechanism (Dantas-Torres 2010). Movement of ticks into new areas is largely restricted to dog visitations to common areas, such as kennels, or when uninfested dogs visit infested homes or vice versa. This restricted genetic exchange, along with long-lasting or prophylactic use of acaricides with a similar mode of action, provides tremendous opportunity for selection of pesticide resistant populations.

In this study a PCR assay was designed to isolate a domain III segment VI sodium channel single nucleotide polymorphism (SNP) in eight populations of peridomestic R. sanguineus collected in Florida and Texas. The Florida and Texas ticks were originally collected by homeowners and kennel operators experiencing difficult-to- 91

eliminate infestations and subsequently screened for acaricide resistance using a larval packet test bioassay with synergists (Eiden et al. 2017). High levels of resistance to permethrin through metabolic detoxification were observed using synergist assays, but it was clear that another mechanism also was providing resistance. The current study identified the SNP at high levels in these populations suggesting that this mutation was contributing to resistance expression at the permethrin discriminating concentration.

After the confirmation of target site insensitivity in R. sanguineus, cross- resistance between permethrin and etofenprox was investigated. In order to do this, the discriminating concentration for etofenprox was established to compare the acaricdal efficacy of both pesticides. Then using the larval packet test and PCR, specimens collected in North America, Europe, and the Caribbean were screened using permethrin and etofenprox concurrently. Resistance ratios generated from these data suggested cross-resistance in the Mexico-Guererro population, however, only tolerance was observed to etofenprox in the Mexico-Nayarit and Grenada populations, negating cross- resistance in these populations at this time. Thereafter, etofenprox target site insensitivity was investigated at the location of a sodium channel SNP known to confer permethrin resistance. The permethrin resistance-conferring sodium channel SNP was present in the populations collected from Grenada, Mexico-Merida, and Mexico-Nayarit.

The mutation was most frequent in the Grenada population where 95% of the screened individuals that survived at the etofenprox discriminating concentration presented a homozygous resistant genotype. Interestingly, homozygous expression of the mutation also was detected in 25% and 10% of non-surviving individuals from the Grenada population screened with permethrin and etofenprox, respectively. This indicates that

there may be other mutations in the R. sanguineus sodium channel that interact with this mutation.

In Chapter 4 evaluations utilized preserved brown dog tick samples that had not been exposed to acaricides and were obtained from populations in North America, the

Caribbean, Africa, Europe, and Asia as well as North American samples from Chapter

2. These populations were tested to elucidate the relationship between latitude, resistance mechanisms, and pathogen-vector interactions. Using molecular assays, populations from these 28 distinct locations were screened for pathogen infection by

Rickettsia, Ehrlichia, Hepatozoon, and Babesia and the presence of the SNP known to confer permethrin resistance. Following DNA amplification, sequencing was used to identify pathogens to species. Sequencing confirmed the isolation of one or more pathogen from the Rickettsia, Ehrlichia, or Hepatozoon groups in samples from 17 distinct locations. The most common pathogen detected was E. canis, which was isolated in 12 populations. Rickettsia massiliae was the most common rickettsial pathogen detected and was isolated in five populations. Also, first reports of Ri. massiliae and Ri. rickettsii in Florida were recorded. Rickettsia masiliae is a known etiological agent in humans (Vitale 2006) and is suspected of causing illness in dogs as well (Beeler et al. 2011). Rickettsia rickettsii was isolated in one Mexico-Mexicali sample. Three samples from Burkina Faso were positive for Hepatozoon, one of which definitively sequenced as H. canis.

Given the global distribution of the brown dog tick, pathogen interactions, and reported acaricide resistance, ample opportunities for investigation are available. Future work should focus on differences in vector competence between the tropical and

temperate lineage. Acquisition of samples from geographic areas where both groups are present, like Texas, would be beneficial to this investigation. The presence of a pyrethroid resistance-conferring SNP was only found in tropical ticks. Further work should investigate the temperate/tropical interaction with other resistance mechanisms by use of synergist bioassays. Another avenue of research could be investigation into the genetic basis for metabolic resistance. This has been studied in insects (Li et al.

2004) but much work needs to be done before fully understanding the genetic processes that drive metabolic resistance in ticks.

As described in Chapter 4, amitraz usage was reported in several Mexico populations of R. sanguineus that were resistant to pyrethroids. Rodriguez-Vivaz et al.

(2016) recently reported amitraz and cypermethrin resistance in Mexico-collected R. sanguineus. Amitraz resistance in R. microplus was investigated by Li et al. (2004) and detected Esterase and glutathione-S-tranferase activity , but target site insensitivity was thought to also be acting to confer resistance. Molecular techniques should be used to investigate amitraz target site insensitivity in future studies.

Several authors have recently investigated the phylogenetics of R. sanguineus using molecular techniques and morphological identification (Hekimoğlu et al. 2016,

Zemstova et al. 2016). The species group interaction could be investigated further through the use of physiological experiments. This would entail rearing tropical and temperate type R. sanguineus at a gradient of temperature and humidity to replicate natural conditions. Physiological data such as fecundity and growth rate could help further define these groups.

APPENDIX A IMPORTATION FORMS

APPENDIX B TICK COLLECTION INSTRUCTIONS Brown Dog Tick Collection and Shipping Instructions

Fig. 1. Wanted! Fully- and nearly fully-fed brown dog tick females, about the size of a raisin. Need at least 5 from a site.

Are you treating a home or kennel for a brown dog tick infestation?

Do you have a home with an infestation?

Researchers at the University of Florida are seeking live, blood-fed (engorged) female brown dog ticks, Rhipicephalus sanguineus, to continue their research on insecticide resistance. Recent research has found most tested populations were highly resistant to permethrin, a common insecticide used on dogs and in area treatments. In the United States, the brown dog tick is the only tick that is capable of completing its entire life-cycle indoors.

If your home or account has a brown dog tick infestation, we would greatly appreciate the submission of at least five or more (>5) fully-engorged ticks (Fig. 1, raisin-sized). Only this size tick will lay eggs. Unfortunately, flat (unfed) ticks are not useful for this study. Engorged ticks are most likely found on the dog’s ears, neck/head, and between the toes, or crawling on the floor or near dog beds.

If you have at least five (5) live, engorged ticks, they can be mailed to the following address. Please see “General collection” and “Shipping instructions ”(pg. 2). Collected ticks must be kept alive. We cannot use dead ticks for the study.

Dr. Phillip Kaufman Entomology and Nematology Dept. PO Box 110620 1881 Natural Area Drive University of Florida Gainesville, FL 32611

If you are outside of Florida (even those of you still in the U.S.) you need to contact us for permitting materials. Instructions for these shipments are

provided in a second document, as are the importation permits that will be needed.

Along with the live ticks, please include the collection data sheet or a note with the following: • City and state (or country, if not U.S.) of the collection • Pesticide products used (on pets or in the home/kennel) and the frequency of pesticide application. • If the homeowner agrees, please include their name and contact information. At a minimum, we need the city and state/country. • It would be most helpful if the homeowner contacts Dr. Kaufman at [email protected] or 352-273-3975 to provide a treatment history for the dog. If enough ticks are submitted alive, we will return resistance status information to providers.

General collection • To remove ticks from animals, firmly grasp the tick as close to the dog’s skin as possible and pull gently. o Fully and nearly-fully engorged ticks (the size we most want) often fall off easily when you are trying to remove them. This is a good clue that you are getting the size of tick that will be most useful. • If needed, tweezers can be used, however, the ticks can be injured. • Do not cover the ticks in any substances to get them to release. We need the ticks to remain alive.

Shipping instructions (Florida only) • Place ticks into something rigid, like an empty prescription pill bottle or clean deli take-out container with a loosely wadded tissue or piece of paper to prevent ticks from being crushed. o No air holes are needed as long as the tissue/paper is not packed too tightly. • Place container in a zipper-type sealable bag, seal, and place into a second zipper-type sealable bag that is then placed into a box container for shipping. • It is best if shipments (from within the U.S.) are on Mondays through Wednesdays to ensure tick survival. • Please do not mail ticks in envelopes, as automatic mail sorting systems will crush the ticks. • Please contact either Dr. Phillip Kaufman ([email protected] /352-273-3975 or graduate student Nick Tucker ([email protected]) to notify us when you ship ticks so that we can prepare for their arrival or if you have questions.

More information on the brown dog tick: https://pmu.ifas.ufl.edu/news-info/2012/brown-dog-ticks https://pmu.ifas.ufl.edu/sites/ufpmu/files/TickBMPs.pdf http://news.ifas.ufl.edu/2015/05/ufifas-scientists-zero-in-on-brown-dog-tick-control/

Brown Dog Tick Collection Data Sheet

Name: ______

Company:______

City:______State/Province:______

Country:______

Phone:______

Email:______

Number of ticks shipped:______

Pesticide(s) applied to dogs:______

Frequency:______

______

Pesticide(s) applied to area/home

(indoor):______

Frequency:______

______

How long have ticks been in the home:______

100

Where have you seen ticks in or around the home:______

Additional information:______

______

101

Shipments from within the U.S., but from outside of Florida.

The state of Florida has an extensive history of invasive species, therefore, the state has rigid importation permits for living organisms from outside its borders. As such, an import permit is required to ship live animals into the state. We have obtained this permit for the shipping of brown dog ticks to our laboratory. We have included in this packet a PDF version of the State of Florida importation permit. You are required to place a copy within the box that you ship to us, as well as a copy on the outside of the box (within the sleeve that holds your mailing label). Use the mailing information above. We recommend an express service for delivery.

Because of the likelihood of your package travelling by air and the exceedingly cold conditions in the cargo area of an airplane, we strongly suggest that you ship the ticks to us in an insulated box. If you do not have such a box, we can send you a box, but they can be obtained from many local sources. If you work near a university or any type of laboratory (including hospitals), you can ask for discarded boxes. It takes these ticks several weeks to lay their eggs, so if provided enough lead notice, we can ship you a box or help to locate one in your area.

Please contact either Dr. Phillip Kaufman ([email protected] /352-273-3975 or graduate student Nick Tucker ([email protected]) if you have any questions and notify when you ship ticks so that we can anticipate your shipments arrival.

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Shipments from outside the U.S.

The United States Department of Agriculture - Animal Plant Health Inspection Service (USDA-APHIS) has oversight on the importation of living organisms into the United States. They require importation permits to be attached to shipments for customs inspection.

The state of Florida has an extensive history of invasive species; therefore, the state also has rigid importation permits for living organisms from outside its borders. As such, another import permit (issued by the state) is required to ship live animals into the state.

We have obtained both of these permits that will allow shipment of brown dog ticks to our laboratory. We have included in this packet a PDF version of the both the USDA- APHIS and the State of Florida importation permits. You are required to place a copy of both permits within the box that you ship to us, as well as a copy of each on the outside of the box (within the sleeve that holds your mailing label).

These shipments must be sent via airline, as other transportation systems are too slow. Because of the exceedingly cold conditions in the cargo area of an airplane, we strongly suggest that you ship the ticks to us in an insulated box. If you do not have such a box we can send you a box, but they can be obtained from many local sources. If you work near a university or any type of laboratory (including hospitals), you can ask for discarded boxes. It takes these ticks several weeks to lay their eggs, so if provided enough lead notice, we can ship you a box or help to locate one in your area.

Please contact either Dr. Phillip Kaufman ([email protected]) or graduate student Nick Tucker ([email protected]) if you have any questions and notify us when you ship ticks so that we can anticipate your shipments arrival.

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BIOGRAPHICAL SKETCH

Nicholas Tucker is a sixth generation Floridian and was born and raised in

Naples, Florida. He received his Bachelor of Science in zoology from Kentucky

Wesleyan College. Nick has been a military service member for over 14 years including three combat deployments to Kuwait, Jordan, Iraq, Afghanistan, and the United Arab

Emirates. After spending several years as a field artillery officer, he was selected for transfer to the medical service corps as a U.S. Army entomologist. His first assignment as a medical service corps officer was serving as an entomologist in a preventive medicine detachment. While deployed, he focused on conducting occupational and environmental health site assessments at numerous base camps throughout southwest

Asia. Nick was then selected to attend graduate school and his first choice was the

University of Florida. He attained a Master of Science in entomology and nematology with an emphasis on medical and veterinary entomology. Nick’s research focused on pesticide resistance and pathogens associated with the brown dog tick. The laboratory experience gained while at the University of Florida will be vital to his success at one of the many U.S. Army research facilities that he may be assigned to. Nick currently serves as a staff entomologist at the Defense Heath Headquarters, Office of The Army

Surgeon General in Falls Church, VA. He plans on returning to the University of Florida to further his education and pursue a Ph. D.

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