ABSTRACT JOHNSON, CONNIE RAE. Molecular Characterization of Host

ABSTRACT

JOHNSON, CONNIE RAE. Molecular Characterization of Host Feeding Patterns and Pathogen Infection Prevalence among Ixodid Tick Vectors in the Piedmont and Coastal Plain of North Carolina. (Under the direction of Dr. Charles S Apperson).

Among tick-borne diseases reportable to the United States Centers for Disease Control and

Prevention (CDC), Lyme disease (LD) is the most commonly reported while Spotted Fever

Group (SFG) Rickettsioses cause the most severe and potentially fatal illness. North Carolina holds the distinction of being ranked among the states reporting the highest number of SFG

Rickettsioses to the CDC. While Rocky Mountain Spotted Fever (RMSF), caused by Rickettsia

rickettsii, is the cause of severe and potentially fatal disease, the majority of mild cases of SFG

Rickettsioses are likely caused by other SFG Rickettsia. Rickettsia parkeri is known to be

present in North Carolina ticks, transmitted by the Gulf Coast Tick, Amblyomma maculatum

Koch. The competent vertebrate reservoir for R. parkeri is not yet known. One objective of this

study was to identify the important vertebrate hosts of A. maculatum and determine if there was

an association with R. parkeri pathogen prevalence. Ticks were collected from a reconstructed

Piedmont prairie in Wake County, North Carolina. Molecular tools were employed to determine

pathogen prevalence and identify the blood meal remnants from host seeking ticks. Pathogen

prevalence was determined through PCR amplification and visualization of the R. parkeri outer

membrane protein A. The reverse line blot hybridization (RLBH) assay was used to detect

vertebrate blood meals through amplification of a 12S rDNA gene fragment and hybridization

against 38 vertebrate probes. Rickettsia parkeri was detected among 36.9% (45/122) A. maculatum adults and 33.3% (2/6) nymphs as well as in a single male lone star tick, Amblyomma americanum Linneaus (2.3%; 1/43). Although no statistically significant association was observed between blood meal host and pathogen prevalence, the cotton rat, Sigmodon hispidus

Say and Ord, was the most common blood meal host identified among 59.0% (36/61) of adult A.

maculatum illustrating its importance as a maintenance host for this tick species.

In the Coastal Plain region of the southeastern United States, two sympatric tick species,

Ixodes affinis Neumann and I. scapularis Say are infected with the LD spirochete, Borrelia

burgdorferi sensu lato (s.l.). Ixodes affinis is an enzootic vector of B. burgdorferi s.l., has a narrow feeding preference and is not thought to readily bite humans. Ixodes scapularis is a generalist feeder and is the primary bridge vector in the Eastern United States transmitting the

LD pathogen to humans. This study investigated if an association occurred between B.

burgdorferi s.l. infection prevalence and vertebrate hosts utilized by these two morphologically

similar and sympatric tick species in the Coastal Plain of North Carolina. Pathogen prevalence

was determined through amplification of the Borrelia burgdorferi s.l. flaB gene, rrfA-rrlB IGS,

and a B. burgdorferi sensu stricto (s.s.)-specific rrs-rrlA ITS fragment. The RLBH assay was

used to identify vertebrate hosts as well as molecularly confirm tick identifications. Statistically

significant differences were observed in B. burgdorferi s.l. infection between I. affinis (80%;

68/85) and I. scapularis (46.8%; 44/94). Two B. burgdorferi s.l. genospecies were identified

through Sanger sequencing of the flaB gene fragment, including the LD agent itself, B.

burgdorferi sensu stricto (s.s.) which was detected among both I. affinis (52.9%; 55/85) and I. scapularis (35.1%; 33/94) and Borrelia bissettii, only known to cause human illness in Europe, was detected at a lower prevalence among I. affinis (11.8%; 10/85) and I. scapularis (4.3%;

4/94). While no statistically significant association between blood meal host and pathogen prevalence was observed among I. affinis or I. scapularis, cotton rat and Canis spp. were the most frequently detected blood meals among both tick species indicating that these vertebrates may play an important role in LD transmission among these sympatric ticks.

Molecular Characterization of Host Feeding Patterns and Pathogen Infection Prevalence among Ixodid Tick Vectors in the Piedmont and Coastal Plain of North Carolina

by Connie Rae Johnson

A dissertation submitted to the Graduate Faculty of North Carolina State University in partial fulfillment of the requirements for the degree of Doctor of Philosophy

Entomology

Raleigh, North Carolina 2021

APPROVED BY:

______Charles S. Apperson R. Michael Roe Committee Chair

______Edward Breitschwerdt Edward Vargo

DEDICATION

To my mother, Wanda Johnson, who was my greatest champion and best friend.

BIOGRAPHY

Connie Rae Johnson is a native of Texas. She attended Texas State University and earned a Bachelor of Science in General Biology in 1996 and a Master of Science in Aquatic

Biology in 2001 under the direction of Dr. Alan Groeger; her thesis examined the longitudinal distribution of zooplankton and phytoplankton in a central Texas reservoir. She was then employed as an Aquatic Scientist by the Texas Commission on Environmental Quality where she worked in the Public Drinking Water Section’s Source Water Assessment and Protection team.

In 2006, she earned a second, non-thesis Master of Agriculture in Economic Entomology from

Texas A&M University under the direction of Dr. Jimmy Olsen where she completed a professional internship conducting West Nile virus surveillance for The Woodlands, Texas.

Connie joined the United States Navy in 2005 as a medical entomologist in the Medical Service

Corps. In 2009 she was selected for the Navy’s Duty Under Instruction program to pursue a PhD in Entomology and began her 3-year duty assignment as a graduate student at North Carolina

State University under Dr. Charles Apperson.

iii

ACKNOWLEDGMENTS

I am grateful to my friends, family and my Navy leadership who have supported and encouraged

me along the way in conducting this research and completing this doctoral program. In

particular, I express deep appreciation to my graduate advisor, Dr. Charles Apperson for his

patience, support and mentorship. I also thank my committee members for their time and

valuable direction, Dr. R. Michael Roe who served as my minor advisor, Dr. Edward

Breitschwerdt who served as my graduate representative, and Dr. Edward Vargo. To Dr.

Loganathan Ponnusamy for serving as special consultant on my committee, providing guidance

and direction and for his mentorship in helping me build my molecular skill set. Dr. Allen

Richards of the Navy Medical Research Command provided guidance and supplies for the

Rickettsia parkeri project. Mr. Alden Estep of the United States Department of Agriculture’s

Center for Medical, Agricultural and Veterinary Entomology in Gainesville, FL provided

guidance and technical support in nanopore sequencing. Haley Thornton Sutton and Dr. Sangmi

Lee, both formerly of the Dearstyne Laboratory, assisted in performing DNA extractions and

troubleshooting the RLBH assay, respectively. To the North Carolina Museum of Natural

Sciences (NCMNS) staff including Lisa Gatens who allowed me to volunteer in the Prairie Ridge

Ecostation small mammal survey and Charles Yelton who provided historical knowledge of that

site. Vertebrate tissues used in this research were obtained from the NCMNS tissue collection,

Louisiana State University Museum of Natural Science Collection of Genetic Resources and the

North Wildlife Resources Commission. Funding for this project was provided in part by a

student grant from the Armed Forces Pest Management Board’s Deployed Warfighter Protection

Fund.

TABLE OF CONTENTS

LIST OF TABLES ...... vii LIST OF FIGURES ...... viii Chapter 1: Molecular Analyses of Blood Meal Hosts and Prevalence of Rickettsia parkeri in the Gulf Coast Tick Amblyomma maculatum (Acari: Ixodidae) from a Reconstructed Piedmont Prairie Ecosystem, North Carolina ...... 1 Abstract ...... 2 Introduction ...... 3 Materials and Methods ...... 7 Study Site and Field Tick Collection ...... 7 DNA Extraction from Ticks and Vertebrate Tissues ...... 8 Pathogen Detection ...... 9 Identification of Host DNA ...... 11 Host Probe Development ...... 11 Reverse Line Blot Hybridization Assay ...... 12 Sensitivity and Specificity of Reverse Line Blot Hybridization ...... 15 Statistical Analysis ...... 15 Results ...... 16 Spatial and Temporal Distribution of Ticks ...... 16 Rickettsia parkeri Prevalence ...... 17 Host Blood meal Identification ...... 17 Host – Rickettsia parkeri Associations ...... 20 Discussion ...... 20 Identification of Blood meal Hosts ...... 20 RLB Probe Specificity ...... 24 Temporal Distribution of A. maculatum ...... 27 Spatial Distribution of Ticks ...... 27 Rickettsia parkeri Infection Rate ...... 29 Conclusion ...... 32 References ...... 34 Supplemental Material ...... 55 Appendix A: Sensitivity and Specificity of Reverse Line Blot Hybridization ...... 56 Appendix B: Supplemental Figure S1 ...... 58 Appendix C: Supplemental Table S1 ...... 59 Appendix D: Supplemental Figure S2 ...... 64 Appendix E: Supplemental Table S2 ...... 65 Appendix F: Supplemental Table S3 ...... 67 Appendix G: Supplemental Table S4 ...... 68

Chapter 2: Molecular identification of blood meals and Borrelia burgdorferi sensu lato infection prevalence among sympatric Ixodes affinis and Ixodes scapularis (Acari: Ixodidae) in the Coastal Plain of North Carolina ...... 69 Abstract ...... 70 Introduction ...... 71 Materials and Methods ...... 75

Study Site and Field Tick Collection ...... 75 DNA Extraction ...... 76 Pathogen Detection ...... 77 Nanopore Sequencing to Determine Co-Infections ...... 79 Identification of Host DNA ...... 82 Vertebrate Host Probe Development ...... 82 Tick Species Probe Development ...... 83 Reverse Line Blot Hybridization Assay ...... 84 Statistical Analysis ...... 86 Results ...... 86 Tick Collection...... 86 Molecular Identification of Tick Species ...... 87 Borrelia burgdorferi Prevalence ...... 88 Nanopore Sequencing to Determine Co-Infections ...... 89 Host Blood meal Determinations ...... 91 Host – Borrelia burgdorferi Associations ...... 92 Discussion ...... 92 Borrelia burgdorferi Prevalence ...... 92 Host – Borrelia burgdorferi Associations ...... 95 Conclusion ...... 99 References ...... 101 APPENDIX ...... 130 Appendix A: Supplemental Table S1 ...... 131

LIST OF TABLES

CHAPTER 1

Table 1. Oligonucleotide primers used in this study to amplify vertebrate and pathogen DNA ...... 47

Table 2. Results of the Reverse Line Blot Hybridization assay to identify blood meal hosts of Amblyomma maculatum, A. americanum and Dermacentor variabilis with pathogen analysis by amplification of Rickettsia parkeri specific ompA shown in parentheses ...... 49

CHAPTER 2

Table 1. Oligonucleotide primers and probes used in this study to detect Borrelia burgdorferi sensu lato, amplify vertebrate blood meals or to identify tick species ...... 118

Table 2. Prevalence of Borrelia burgdorferi s.l. among I. affinis and I. scapularis based on amplification of fragments of flaB, rrfA-rrlB, and Borrelia burgdorferi s.s. specific rrs-rrlA ...... 121

Table 3. Sanger sequencing of B. burgdorferi s.l. chromosomal gene fragments flaB, rrfA-rrlB, rrs-rrlA for I. affinis, I. scapularis and B. burgdorferi B31 positive template for ticks used in nanopore sequencing analysis ...... 122

Table 4. Nanopore sequencing of B. burgdorferi s.l. chromosomal gene fragments flaB, rrfA-rrlB, and rrs-rrlA among I. affinis, I. scapularis and B. burgdorferi s.l. positive template DNA ...... 124

Table 5. Host blood meals identified and B. burgdorferi s.l. genospecies infecting host- seeking I. affinis and I. scapularis...... 128

vii

LIST OF FIGURES

CHAPTER 1

Figure 1. Example probe validation with vertebrate tissues ...... 53

Figure 2. Seasonal distribution of A. maculatum, A Americanum and D. variabilis ...... 54

CHAPTER 2

Figure 1. Location of study sites in the Coastal Plain region of North Carolina ...... 113

Figure 2. Molecular phylogenetic analysis of Sanger sequencing of a 601 bp fragment of the B. burgdorferi s.l. flaB gene among Ixodes affinis and Ixodes scapularis ...... 114

Figure 3. Molecular phylogenetic analysis of Sanger and nanopore consensus sequences of B. burgdorferi ospC amplicons among positive template DNA, Ixodes affinis and Ixodes scapularis ...... 116

viii

CHAPTER 1

Molecular Analyses of Blood Meal Hosts and Prevalence of Rickettsia parkeri in the Gulf Coast

Tick Amblyomma maculatum (Acari: Ixodidae) from a Reconstructed Piedmont Prairie

Ecosystem, North Carolina

This chapter was formatted for submission to The Journal of Medical Entomology with the coauthors Loganathan Ponnusamy1,2, Allen L. Richards3, and

Charles S. Apperson1,2

1Dept. of Entomology and Plath Pathology, North Carolina State University, Raleigh, NC 27695

2Comparative Medicine Institute, North Carolina State University, Raleigh, NC 27695 USA

3Department of Preventive Medicine and Biostatistics, Uniformed Services University of the

Health Sciences, Bethesda, MD 20814

Abstract

Molecular methods were used to determine host feeding patterns and the prevalence of infection

with an emerging pathogen, Rickettsia parkeri among the primary vector, Amblyomma

maculatum Koch as well as sympatric tick species A. americanum (Linneaus) and Dermacentor

variabilis (Say) collected from a reconstructed prairie in the Piedmont region of North Carolina

during 2011 and 2012. The occurrence of R. parkeri among A. maculatum adults and nymphs

was 36.9% (45/122) and 33.3% (2/6), respectively. Rickettsia parkeri was detected in a single

male A. americanum 2.3% (1/43). A PCR-reverse line blot hybridization assay of a 12S rDNA

fragment amplified from host-seeking adult and nymphal ticks was used to identify vertebrate

hosts representing the remnant nymphal and larval blood meals, respectively. Blood meal host

identification was successful for 50% (61/122) of A. maculatum adults collected from vegetation

and 100% (4/4) of nymphs collected directly from cotton rats (Sigmodon hispidus Say and Ord).

Blood meal host identification was lower for adults of other species with host identification in

29.3% (12/41) of A. americanum and 39.2% (20/51) of D. variabilis. The cotton rat, Sigmodon

hispidus Say and Ord, was the most common blood meal host identified for 59.0% (36/61) of

adult A. maculatum. No statistically significant association was observed, however, between

blood meal host and pathogen prevalence for any tick species. While the cotton rat is an

important blood meal host for A. maculatum nymphs, this vertebrate does not appear to be the

primary source of R. parkeri infection for A. maculatum.

Keywords: Tick, Amblyomma maculatum, host blood meal identification, Sigmodon hispidus,

PCR-reverse line blot hybridization assay, Rickettsia parkeri

Short title: Host-pathogen associations in ticks from a restored Piedmont prairie

Introduction

Of tick-borne diseases in the United States, rickettsial diseases are second only to Lyme disease in the number of annual case reports to the CDC (https://www.cdc.gov/ticks/data- summary/index.html). The most severe of these is Rocky Mountain spotted fever (RMSF), a spotted fever group (SFG) rickettsiosis caused by infection with Rickettsia rickettsii, an obligate intracellular bacterium that historically has been transmitted by the Rocky Mountain wood tick

(Dermacentor andersoni Stiles) in the western United States and the American dog tick (D. variabilis Say) in the eastern United States (Burgdorfer 1975, McDade and Newhouse 1986).

Other tick species recently implicated as vectors include the brown dog tick (Rhipicephalus sanguineus Latreille) which is now recognized as an important vector along the U.S.-Mexico

Border region (Biggs et al. 2016), and the lone star tick (Amblyomma americanum (Linn.))

(Breitschwerdt et al. 2011, Levin et al. 2017).

The epidemiology of SFG rickettsioses in the United States is complicated by a growing number of SFG rickettsiae that have been recognized to cause mild illness in humans and cross- react in serologic tests (Vaughn et al. 2014) including R. amblyommatis (Karpathy et al. 2016) commonly found infecting the lone star tick A. americanum (Chapman et al. 2006), R. philipii

(strain 364D) in D. occidentalis Marx (Shapiro et al. 2010), R. akari in the house mouse mite

Liponyssoides (formerly Allodermanyssus) sanguineus (Hirst) (Eustis and Fuller 1952), and R. parkeri found in the Gulf Coast tick A. maculatum Koch (Paddock et al. 2004). Due to the emerging recognition of SFG rickettsioses with similar clinical features caused by these rickettisae, the CDC revised the case definition for RMSF reporting in 2010 to include ‘SFG rickettsioses including RMSF’ and added presence of an eschar among the signs as it is often associated with the mildly pathogenic infections with R. parkeri and R. akari (CSTE 2009).

Although geographically SFG rickettsioses are reported throughout the contiguous

United States, North Carolina is one of the top states reporting a consistently high number of

annual case reports of probable SFG rickettsioses; along with Arkansas, Missouri, Oklahoma,

and Tennessee, these five states collectively report more than 63% of probable SFG rickettsioses

in the United States annually (Dahlgren et al. 2012, CDC 2013). Rickettsia parkeri may be a

cause of probable SFG rickettsioses that have similar symptoms as RMSF yet are milder and

typically associated with an eschar at the bite site unlike RMSF (Paddock 2008).

Rickettsia parkeri was first isolated in 1937 from the Gulf Coast tick Amblyomma

maculatum removed from cows in Liberty County, Texas during an investigation into cases of

RMSF; the isolate was found to cause mild illness in guinea pigs and have complete cross

immunity with R. rickettsii and R. conorii, the agent of Boutonneuse fever (also known as

Mediterranean spotted fever) (Parker et al. 1939). It was not until 2002 that human infection

with R. parkeri was detected in a skin punch biopsy from a patient from southeastern Virginia with no recollection of tick bite who experienced fever, mild headache, malaise, myalgia,

arthralgia, multiple eschars on lower legs and maculopapular rash that spread from trunk to

extremities as described in Paddock (2004). Subsequently, Fornadel et al. (2011) reported a R.

parkeri infection prevalence of 41.4% for A. maculatum collected in Fairfax County, Virginia

and Wright et al. (2011) reported an infection prevalence of 43.1% for A. maculatum ticks

collected in southeastern Virginia. In North Carolina where some of the highest number of

annual cases of RMSF are reported, Varela-Stokes et al. (2011) detected an overall prevalence of

R. parkeri in 29% of A. maculatum. This prevalence rate in A. maculatum along with infrequent

occurrence of severe illness confirmed to be RMSF suggests that this re-emerging human

pathogen may be one of the agents responsible for the large number of probable RMSF cases

reported from North Carolina.

Ticks may become infected with pathogens through ingestion of a blood meal from an

infected vertebrate host (horizontal transmission) and/or through vertical transmission from an

infected female tick to her eggs (transovarial) or when molting from an infected larva to nymph

or infected nymph to adult (transstadial transmission). While many SFG rickettsiae are

transmitted through transstadial and transovarial transmission, the filial infection rate (proportion

of offspring infected) varies among SFG rickettsiae with R. rickettsii filial rates ranging from

30% to 100% (Burgdorfer and Brinton 1975, Ricketts 1907, Price 1954). In experimentally

infected A. maculatum, Harris et al. (2017) reported transovarial transmission of R. parkeri in

38% of female ticks to their eggs and observed transstadial transmission of 67% from larvae to nymph with no transstadial transmission of R. parkeri to the adult stage. In contrast, Wright et al. (2015) examined vertical transmission of R. parkeri among field collected A. maculatum,

finding 83.7% transovarial transmission and 100% transstadial transmission in all stages through

the adult. Under natural conditions vertical transmission may be less than that observed in

laboratory studies since survival and fecundity of tick vectors is greatly reduced when infected

with highly virulent strains (Burgdorfer and Brinton 1975, Azad and Beard 1998, Niebylski et al.

1999, Macaluso et al. 2002). Since even mildly pathogenic strains like R. parkeri have been

shown to have negative effects on the fitness of infected ticks, the importance of vertebrate hosts

in maintaining pathogenic SFG rickettsiae in a zoonotic cycle is of particular interest (Nieri-

Bastos et al. 2013).

There have been a limited number of specific studies conducted to determine the

reservoir competence of vertebrate hosts for R. parkeri in A. maculatum. Amblyomma

maculatum feeds on a variety of vertebrate hosts over its 3-host life cycle. While ground- dwelling birds and small mammals are preferred hosts of immature stages, larger mammals such as livestock and deer are hosts of adults (Bishopp and Hixson 1936, Bishopp and Trembley

1945, Samuel and Trainer 1970, Koch and Hair 1975, Barker et al. 2004, Ketchum et al. 2005).

Leydet and Liang (2013) found a high prevalence of R. parkeri in ticks collected from Louisiana black bears (Ursus americanus luteolus Griffith), including 65.7% of A. maculatum, 28% of D. variabilis and 11.1% of Ixodes scapularis, the blacklegged tick. Yabsley et al. (2009) detected

R. parkeri in 8.3% (1/12) of A. maculatum collected from bears in Florida and Georgia.

Domestic dogs have been found to be infected with R. parkeri (Grasperge et al. 2012); however, it is unlikely they serve as a reservoir in nature. Seroconversion studies have attempted to shed light on likely competent hosts for R. parkeri. Moraru et al. (2013b) examined 13 passerine bird species, 6 rodent species and the northern bobwhite quail (Colinus virginianus Linn.) from locations in Mississippi for evidence of SFG rickettsioses, only cotton rats (Sigmodon hispidus),

deer mice (Peromyscus sp.) and northern bobwhite quail exhibited seroreactivity against R.

parkeri antigen.

The purpose of this study was to identify vertebrate hosts and determine R. parkeri

prevalence among host seeking A. maculatum and sympatric tick species in order to characterize vertebrate reservoir host-pathogen associations. Identification of vertebrate hosts as well as infection rates of R. parkeri in A. maculatum and other tick vectors, including A. americanum

and D. variabilis from North Carolina, would increase understanding of the ecology of these tick

species and this emerging pathogen. We used molecular methods for identification of hosts,

eliminating the need to trap and handle wildlife and ensuring cryptic hosts were not overlooked.

Among hematophagous arthropods, molecular studies are particularly suited to tick blood meal

analysis since digestion is a slow intracellular process taking months to complete (Tarnowski and

Coons 1989). The reverse line blot hybridization (RLBH) assay was utilized in identification of tick blood meal hosts because up to 43 samples could be simultaneously screened against 43 oligonucleotide probes. This method enables the identification of blood meals taken from multiple hosts, which is a common limitation of other molecular techniques. Using 12S rDNA as the molecular marker, the assay allowed for species-level identification of pathogens and vertebrate hosts.

Materials and Methods

Study Site and Field Tick Collection

Our study was carried out in in Raleigh, Wake County, North Carolina at the Prairie Ridge

Ecostation (35°48’42.36” N, 78°42’50.00” W), a field station of the North Carolina Museum of

Natural Sciences (NCMNS) containing approximately 15.4 hectares of the prairie habitat restored from pastureland. The remaining pastureland is dominated by tall fescue (Festuca sp.) and Bermuda grass (Cynodon dactylon) and is bordered on the south and southeast with dry- mesic oak hickory forest and Piedmont bottomland forest (Yelton 2007). Field tick collections were made monthly from June to September 2011 and weekly from March to August 2012 with a few exceptions due to inclement weather. Beginning in 2012, collection efforts were divided into three areas of the study site in order to increase the likelihood of collecting ticks that had fed on diverse vertebrate hosts (Supp. Figure S1). Each area was approximately 4 hectares in size with a minimum of 20 m buffer between each area. Area “A” is a reconstructed native Piedmont prairie, a conversion which began from old pastureland in 2004. During peak tick activity the dominant plant species within area “A” were tallgrass prairie grasses switchgrass (Panicum

virgatum) and Indiangrass (Sorghastrum nutans). Area “A” will be subsequently referred to as

the “the prairie habitat”. Area “B” contained old pastureland and lowland and bottomland

forested areas. Tall fescue with patches of tickseed (Coreopsis sp.) and blackberry (Rubus sp.)

were dominant plants in area “B” during peak tick activity. Area “B” will subsequently be

referred to as “the old pastureland habitat”. Area “C” was predominately lowland and upland

forest although bottomland forest was located along the southern border running along Richland

Creek. Area “C” was dominated by invasive plant species Japanese stiltgrass (Microstegium

vimineum) and Chinese privet (Ligustrum sinense). Broomsedge (Andropogon virginicus) was

common within the lowland prairie of areas “B” and “C” during the peak of tick activity (C.

Yelton, personal communication, 2013). Area “C” will subsequently be referred to as “the forest

habitat”.

Host-seeking ticks were collected for one hour within each area using 1 m2 drag corduroy

cloths. Immediately after collection, ticks were placed in vials containing 95% ethanol. Nymphs

and adults were identified using the taxonomic keys of Keirans and Durden (1998) and Keirans

and Litwak (1989), respectively. Following identification, ticks were stored individually in vials

containing 95% ethanol, according to species, collection location and date, and held at -20°C.

DNA Extraction from Ticks and Vertebrate Tissues

To minimize contamination of samples with foreign DNA, extraction of total DNA, PCR

amplification, and product analysis were conducted in separate rooms. Extraction and PCR

workspaces along with equipment including pre-sterilized tubes, pipettes and filter tips were sterilized with UV light for 15 min prior to use. Separate pipettes and pipette tips were used for

DNA extraction and setup of PCR assays. Prior to DNA isolation, individual ticks were surface

sterilized by immersion and shaking for 30 s each in 1% sodium hypochlorite, 0.1 M phosphate buffered saline (PBS) and 95% ethanol followed by three additional 1 min washes in 0.1 M PBS.

One sample of the final wash in PBS was retained from each set of extractions to attempt amplification of vertebrate DNA to verify the effectiveness of surface sterilization.

DNA was extracted using a modified lysis buffer (Smith et al. 2010) and the phenol- chloroform method. Ticks were first minced individually with sterile #11 scalpel blades in 160

µl lysis buffer 1 [2% (w/v) sodium dodecyl sulfate, 100 mM tris(hydroxymethyl)-aminomethane hydrochloride (Tris-HCl), 200 mM sodium chloride (NaCl), 10 mM ethylenediaminetetraacetic acid (EDTA), pH 7.0] and 20 µl each of freshly prepared lysozyme (100 mg/mL) and proteinase

K (20 mg/mL). After a 1 h incubation at 37°C, 200 µL of lysis buffer 2 [1 % (w/v) cetyltrimethylammonium bromide, 1.5 M NaCl, 100 mM Tris-HCl and 100 mM EDTA] was added to each sample and incubated at 56°C for 2 h. Negative extraction controls were included with each round of tick extractions.

To remove PCR inhibitors, the DNA extract was subjected to an additional purification using silica spin columns (EconospinTM, Epoch Life Sciences, Sugarland, TX, USA) following the manufacturer recipe for DNA absorption and wash buffers and the protocol of Smith et al.

(2010). Purified DNA was eluted in a total volume of 100 µl sterile water by two centrifugations at 5900 x g for 1 min with 50 µl sterile water. DNA quantity and quality were measured using a

Nanodrop® spectrophotometer (Thermo Scientific, Waltham, MA, USA). Genomic DNA samples were stored at -20°C.

Pathogen Detection

A 744 bp fragment of the bacterial universal 16s rRNA gene was amplified using forward primer

0206 and biotin labeled reverse primer 0209 of Pichon et al. (2003) for use in RLBH detection of

R. parkeri (Table 1). During validation of the amine labeled probe developed to detect R. parkeri (amine-5’-CGGTCGTCTGGGCTACAA-3’), it was found to cross-hybridize with several rickettsiae, including R. rickettsii and R. amblyommatis, and therefore RLBH assay was not used to detect R. parkeri. Instead, a nested PCR assay described by Oliveira et al. (2008) was employed to amplify a 232 bp fragment of the Rickettsia 17 kDa antigen gene using outer primers 17kD1 and 17kD2 described by Webb et al. (1990) and inner primers 17kN1 and 17kN2 modified from Schriefer et al. (1994). Reaction mixture volumes of 20 µl contained 1x

AmpliTaq Gold® 360 Master Mix (Applied Biosystems, Carlsbad, CA), 0.4 µM of each primer and 1 µl of extract template DNA. PCR amplifications were run on a Bio-Rad PTC-100 thermal cycler, using an initial denaturation of 95°C for 10 min, 30 cycles of 95°C for 60 s, 50°C for 60 s, and 72°C for 2 min followed by a final extension at 72°C for 5 min. Reaction mixtures and

PCR parameters were the same for both outer and inner PCR reactions. The amplicons were purified and subjected to DNA sequencing (Sanger) using the reverse primer 17kN2 (Eton

Bioscience, Inc., Research Triangle Park, NC). Sequences were compared with rickettsial sequences from the RefSeq Genome Database using the BLASTn program (version 2.10.0 blast.ncbi.nlm.nih.gov).

Due to low sequence variability of the 17 kDa gene among SFG rickettsiae, PCR amplification of a 447 bp fragment of R. parkeri-specific outer membrane protein A (RpompA) was subsequently used to detect R. parkeri in tick extracts, using primers RpompAF and

RpompAR described and confirmed to specifically amplify R. parkeri among other SFG

rickettsiae by Varela-Stokes et al. (2011). PCR reactions were carried out on a Labnet

Multigene™ thermal cycler in 20 µl reaction mixtures containing 1x AmpliTaq Gold® 360

Master Mix (Applied Biosystems, Carlsbad, CA), 0.3 µM each primer and 2 µl DNA extract.

PCR thermal cycler conditions were 95°C for 10 min followed by 40 cycles of 94°C for 30 s,

54°C for 60 s, 72°C for 90 s and final extension at 72°C for 5 min. Successful amplification of the 447 bp ompA fragment of R. parkeri was determined by visualization of the PCR product of the expected size on an ethidium bromide stained 1% agarose gel.

Identification of Host DNA

Host Probe Development

The Prairie Ridge Ecostation is an ideal location to conduct host blood meal studies since the research staff has compiled an extensive list of vertebrate species found on the site. Probes were developed for potential vertebrate tick hosts that were confirmed or predicted to occur within the study area. Geographic information system (GIS) datasets available through the North Carolina

Gap Analysis Project (NC-GAP) were utilized to map predicted species distribution and habitat within the geographic extent of Prairie Ridge Ecostation and within each of the three study areas

(http://www.basic.ncsu.edu/ncgap/). The NC-GAP uses Landsat TM satellite imagery land cover data combined with existing range maps for vertebrate species to predict potential distribution of vertebrate species (McKerrow et al. 2006). Since the number of bird species known to serve as tick hosts is too large for the limited number of lanes available in the RLBH assay, an order level aves probe developed by Humair et al. (2007) was used to detect avian blood meals. Two species-level probes were targeted for ground dwelling birds that are known to be important tick hosts; namely, the northern bobwhite quail (Colinus virginianus) and wild turkey (Meleagris

gallopavo). Similarly, higher level probes to detect lizard and mammal DNA developed by

Humair et al. (2007) were used to detect presence of mammals and of selected lizard species, including the glass lizard (Ophisaurus sp.), five lined skink (Plestiodon sp.) and little brown skink (Scincella lateralis). All other probes were developed to complement the 145 bp fragment of vertebrate 12S rDNA of Humair et al. (2007). Sequences of the 12S rDNA fragment for target species identified from NC-GAP were downloaded from GenBank, trimmed to the fragment size and aligned using ClustalX 2.1 (Larkin et al. 2007). Each probe was developed to distinguish hosts at the lowest possible taxonomic level within the detection limits of the methodology. Probes were designed to minimize folding of the PCR amplicon under the RLBH assay conditions and to limit hybridization temperatures to within a few degrees of each other using OligoAnalyzer 3.1 (Integrated DNA Technologies, Inc., Give location). Developed probes were validated against vertebrate tissues obtained from the North Carolina Museum of Natural

Sciences (NCMNS), the Louisiana State University Museum of Natural Science Collection of

Genetic Resources and tissue specimens held by the Apperson lab. Northern bobwhite quail tissue was obtained from the North Carolina Wildlife Resources Commission.

Reverse Line Blot Hybridization Assay

A 145 bp fragment of the 12S rDNA gene was amplified following the nested PCR assay of

Humair et al. (2007) using outer primers 12S-12F and 12S-13R and inner primers 12S-6F and biotin labeled B-12S-9R. The reverse primer was modified from Humair et al. (2007) to amplify

North American vertebrates. The ~600 bp outer fragment was amplified on a Labnet

Multigene™ thermal cycler in 25 µl reaction mixture containing AmpliTaq Gold® 360 Master

Mix, 0.80 µM each primer and 10 µl DNA extract. Touchdown PCR conditions included an

initial denaturation step for 10 min at 95°C, followed by 10 cycles of 30s at 94°C, 30s at 65°C,

and 72°C for 60s with the annealing temperature lowered by 0.5°C each cycle followed by 25

cycles of 30 s at 94°C, 30 s at 60°C, and 1 min at 72°C with a final extension of 72°C for 10 min.

The 145 bp inner fragment was amplified as above using 1 µl of the outer PCR reaction as

template. Touchdown PCR conditions for the inner fragment included an initial denaturation for

10 min at 95°C followed by 8 cycles of 30 s at 94°C, 30 s at 60°C, and 30 s at 72°C, lowering the

annealing temperature by 1°C each cycle then 35 cycles using an annealing temperature of 52°C

and a final extension of 72°C for 7 min. PCR products were held at 4°C for up to 48 h before

RLBH analyses (Kong and Gilbert, 2007).

The PCR-RLB hybridization assay followed the protocol described by Kong and Gilbert

(2007). Thirty-eight oligonucleotide probes (Invitrogen, Carlsbad, CA) were diluted in 0.5 M

NaHCO3, pH 8.5 to achieve concentrations between 400-1400 pmol to compensate for

differences in hybridization intensity among probes (Suppl. Table S1). Probes were covalently

bound by their 5’ amino group to a Biodyne® C nylon membrane (Pall Corporation, Ann Arbor,

MI) by first activating the membrane in 16% (w/v) 1-ethyl-3-(3-dimethylaminopropyl) carbodiimide (EDAC) for 10 min. After a quick rinse with distilled water, the membrane was placed in a Miniblotter® 45 (Immunetics, Inc., Boston, MA) and 150 µl of each probe was added to each lane and allowed to incubate for 15 min. The membrane was then deactivated in 100 mM sodium hydroxide for no longer than 10 min, washed in 2x saline sodium phosphate-EDTA

(SSPE) and washed for 5 min in 2x SSPE / 0.1% SDS at 60°C. The membrane was stored until use in 20 mM EDTA at 4°C after a 20 min wash in 20 mM EDTA. All chemicals used in probe binding were obtained from Sigma-Aldrich, St. Louis, MO, USA.

Hybridization of biotin-labeled PCR products was initiated with a pre-hybridization wash of the stored membrane in 2x SSPE / 0.1% SDS for 5 min at 60°C. To decrease non-specific hybridizations, the membrane was blocked in 0.1 % casein (Fisher Scientific, Pittsburg, PA) in

2x SSPE / 0.1 % SDS for 60 s and then washed once again in 2x SSPE / 0.1 % SDS for 5 min at

60°C. Higher concentration of casein or longer incubation times resulted in diminished signal from positive controls. Ten µL of PCR products were diluted in 150 µl 2x SSPE/ 0.1% SDS.

The diluted PCR products were denatured at 99°C for 10 min and then submersed for 5 min in an ice bath. The membrane was placed in the miniblotter, rotated 90° to the bound probes and the diluted, denatured PCR products were applied to the membrane. PCR products were allowed to hybridize with probes for 1 h at 58°C after which the membrane was washed twice with 2x SSPE

/ 0.5% SDS for 10 min at 55°C. The membrane was incubated in a rolling hybridization oven in streptavidin-peroxidase conjugate diluted in 2x SSPE / 0.5% SDS for 45 min at 42°C. The enzyme-conjugated membrane was then washed twice with 2x SSPE / 0.5% SDS for 10 min at

42°C and then twice with 2x SSPE for 5 min at room temperature. The membrane was incubated for 2 min in electrochemiluminescence (ECL) detection liquid (GE Healthcare,

Otelfingen, Switzerland) and chemiluminescence was detected using a ChemiDoc-It® Imager

(UVP, Upland, CA, USA) after a 10 min exposure.

Hybridized PCR products were stripped from the membranes by two 30 min washes in

0.5 % SDS at 80°C followed by a 15 min wash in 20 mM EDTA. Membranes were stored at

4°C in 20 mM EDTA until the next use. Although Kong & Gilbert (2007) suggested membranes could be re-used at least 20 times, it was found that signal intensity diminished considerably after

14 uses at which point probes were bound to new membranes.

Sensitivity and Specificity of Reverse Line Blot Hybridization

Probes designed for this study effectively identified 38 host species that potentially occurred in the Prairie Ridge Ecostation (Fig. 1). Cross hybridizations between probes and vertebrate DNA extracts were observed during probe validation. Probes were re-designed if cross-hybridizations occurred with multiple vertebrate tissues. Some cross-hybridizations, however, did not preclude identification of the host(s) due to differential hybridization patterns. For example, tissue of the least shrew, Cryptotis parva (Say), showed a strong cross hybridization with the long-tailed shrew probe. Since long-tailed shrew (Sorex sp.) tissue did not cross hybridize with the probe for least shrew, a double hybridization for long-tailed shrew and least shrew would be most likely due to C. parva. Similarly, while rabbit tissue (Sylvilagus floridana) strongly hybridized to the mink probe, mink tissue (Neovison vison) would only weakly hybridize to rabbit probe.

Additional information on probe specificity is given in the supplementary materials.

Statistical Analysis

Statistical analyses were carried out using SPSS, Version 19.0 (IBM, Armonk, NY). Chi-square tests of association were used to determine if 1) there was an association between R. parkeri infection and tick demographics (sex, study area), 2) or between R. parkeri infection prevalence and blood meal host detected, or 3) between blood meal host detected and tick demographics

(sex, study area). Bonferroni correction was applied in multiple comparisons to decrease the likelihood of a type I error.

Results

Spatial and Temporal Distribution of Ticks

A total of 1,054 A. maculatum were collected during our study, including 6 nymphs, 579 female, and 469 male ticks. Four of these nymphs were collected from cotton rats trapped in the prairie habitat during a NCMNS quarterly small mammal survey, one in February and three in October

2012. In 2011, collection of questing ticks (n= 144) began in June and ended in mid-September when only two adult A. maculatum ticks were collected. In 2012, collection efforts began in

March. Adults (n= 904) began questing in mid-April and were active until the end of August with the collection of only two adult A. maculatum. We collected more males than females in the first four weeks of collection in 2012. Males were most abundant the first week of collection in April while females were most abundant in the first week of June (Fig. 2). Peak abundances overall were observed in the first week of June in both collection years.

Three times as many adult A. maculatum were collected in the prairie habitat than the other two areas (Supp. Figure S2). Fewer A. americanum and D. variabilis were collected and had a more limited distribution and seasonal activity than A. maculatum. Only 41 adult and 2 nymphal A. americanum were collected during our study, primarily within the old pastureland and forest habitats. The majority of the 68 D. variabilis were collected in July 2011 when collection area was not recorded. Although questing activity of both species began in April 2012 along with A. maculatum, A. americanum were no longer collected after the first week of July and D. variabilis were absent by the third week of July. Populations of A. americanum peaked in mid-May when host-seeking nymphs were also collected. Dermacentor variabilis reached peak abundance in late June/early July as the A. maculatum population began to decline. Although both D. variabilis and A. americanum showed evidence of a bimodal seasonality, this

distribution was more distinct for D. variabilis, which were absent in collections in mid-May.

Both A. americanum and D. variabilis were rarely collected in the prairie habitat.

Rickettsia parkeri Prevalence

Six A. maculatum nymphs along with 122 A. maculatum, 41 A. americanum and 51 D. variabilis

adults were used in pathogen and blood meal analyses. Because A. maculatum adults were

abundant, at least three of each sex for each collection date and 15 of each sex from each area

were analyzed.

Of 128 A. maculatum used in blood meal analysis, 36.7% (47/128) produced a RpompA amplicon, including two nymphs (2/6; 33.3%), 25 females (25/61; 41.0%) and 20 males (20/61;

32.8%). Female and male ticks did not differ significantly in pathogen infection (χ2 = 0.88, df =

1, p = 0.348). One male A. americanum collected in the forest habitat was infected with R.

parkeri (1/41; 2.4%). No R. parkeri was detected among the 51 D. variabilis tested. In

comparing infection prevalence across the three areas of the study site, only prairie and old

pastureland habitats differed significantly with more A. maculatum adults infected in the prairie

habitat (20/36; 55.6%) than in the old pastureland habitat (8/31; 25.8%) (χ2 = 6.228, df = 2, p =

0.044). Infection prevalence among A. maculatum in the forest habitat was 46.9% (15/32).

Host Blood meal Identification

Blood meal determinations were possible for 100% (4/4) engorged A. maculatum nymphs removed from cotton rats and 50.0% (61/122) host seeking adults. Blood meal identification success was lower in adult A. americanum (12/41; 29.3%) and D. variabilis (20/51; 39.2%). No blood meals were identified in two host-seeking nymphal A. maculatum and A. americanum.

Among blood meals, multiple hosts were detected in adult A. maculatum (3/65; 4.6%), A.

americanum (2/12; 16.7%) and D. variabilis (4/20; 20.0%) (Table 2).

No statistically significant difference was observed in blood meal host identification

success or host determination between female and male A. maculatum ticks (χ2 = 0.03, df = 1, p

= 0.856). Success of blood meal host identification was lowest 20 June 2012 (1/13; 7.7%) and

highest on 27 April 2012 (7/9; 77.8%).

Amplified DNA from blood meals of all four A. maculatum nymphs removed from cotton

rats hybridized correctly to host probes for cricetidae and cotton rat. Of 10 blood meal hosts

identified, the majority of adult A. maculatum amplifiable blood meals hybridized to cotton rat

probes (36/61; 59.0%), Canis sp. (10/61; 16.4%) and bird (9/61; 14.8%). Other hosts identified

included five-lined skink (Plestiodon sp.), rabbit (Sylvilagus sp.), chipmunk (Tamias striatus L.),

small rodent (cricetidae), Peromyscus sp., gray fox (Urocyon cinereoargenteus Schreber) and

white-tailed deer (Odocoileus virginianus Zimmermann) (Table 2). Of the multiple blood meals, one male and one female tick showed hybridizations with both Canis sp. and cricetidae + cotton rat probes and one female contained blood meals of Canis sp. and Peromyscus sp.

Eight hosts were identified among A. americanum blood meals amplified. Raccoon was the only blood meal host not identified among A. maculatum. The most frequent blood meal hosts detected among A. americanum were cotton rat and Canis sp. (3/12; 25%). Multiple blood meals were detected in one female (cotton rat + raccoon) and one male (cotton rat + Canis sp.).

Eight hosts were identified among D. variabilis blood meals, with only opossum and raccoon identified as blood meal hosts that were not identified for A. maculatum. The majority of D. variabilis blood meals hybridized to cotton rat probe (14/24; 58.3%). Multiple blood meals

were detected, including a male and a female that each had fed on rabbit + cotton rat, one female

that fed on rabbit + gray fox and one male that fed on cotton rat + raccoon.

Differences were observed in A. maculatum blood meal host identification success among

areas. Identifications of blood meal hosts were significantly higher in adult A. maculatum

collected within the prairie habitat (25/36; 69.4%) than in the forest habitat (11/32; 34.4%) (χ2 =

8.36, df = 1, p = 0.004). Blood meal identification was successful in 45.2% (14/31) A. maculatum adults collected in the old pastureland habitat. No difference in blood meal identification success among study areas was observed for A. americanum or D. variabilis. No blood meals were identified among the few A. americanum and D. variabilis collected within the prairie habitat. Of blood meals identified among A. americanum, 53.8% (7/13) were from adults collected in the old pastureland habitat and 17.4% (4/23) were collected in the forest habitat.

Blood meals were identified for 28.6% (4/14) of D. variabilis collected in the old pastureland habitat and in 52.9% (9/17) of adults collected in the forest habitat.

Associations between host identified and study area were only observed in A. maculatum.

Among A. maculatum adults with identifiable blood meals, significantly higher numbers of cotton rat blood meals were identified in the prairie habitat (27/29; 93.1%) than old pastureland

(1/14; 7.1%) or forest (3/11; 27.3%), (χ2 = 33.66, df = 2, p = 0.000). Birds were the most

common hosts of ticks collected in the old pastureland habitat (7/14; 50.0%). Identification of

bird DNA in blood meals were significantly higher in the old pastureland habitat than in the

prairie habitat where no birds were identified among blood meals (0/29; 0%), (χ2 = 15.234, df =

1, p = 0.000) but were not significantly different from bird identifications in the forest habitat

(2/11; 18.2%), (χ2 = 2.707, df = 1, p = 0.100). No other differences were found between hosts identified and area of collection.

Host – Rickettsia parkeri Associations

There was no difference in R. parkeri infection between A. maculatum adults with or without identified blood meals (χ2 = 2.85, df = 1, p = 0.091). Rickettsia parkeri was detected in 44.3%

(27/61) of adult A. maculatum ticks with identified blood meals and 29.5% (18/61) without identified blood meals. Among A. maculatum with identified blood meals, no difference was observed between infection status and sex (χ2 = 0.788, df = 1, p = 0.375) or host (χ2 = 10.47, df =

9, p = 0.314). Of the nymphs removed from cotton rats, 50% were infected with R. parkeri (2/4).

Among infected A. maculatum, cotton rats were the most frequently identified blood meal host

(13/34; 38.2%), followed by birds (4/9; 44.4%) and Canis sp. (3/7; 42.9%). The majority of infected A. maculatum that fed on cotton rat were collected in the prairie habitat (11/13; 84.6%), while the majority that fed on birds were collected in the old pastureland habitat (3/4; 75.0%) and those that fed on Canis sp. were mainly collected in the forest habitat (2/3; 66.7%).

Discussion

Identification of Blood meal Hosts

Our present study represents the first molecular analysis of remnant blood meals from host- seeking A. maculatum and D. variabilis. Blood meals identified in host seeking adult ticks represented remnant nymphal blood meals. In our study, the majority of A. maculatum and D. variabilis adults amplified blood meals that hybridized to the cotton rat probe. The most frequent blood meals identified in adult A. americanum were from cotton rat and Canis sp. The

Canis sp. blood meals were likely from coyote (Canis latrans Say). Pet dogs (C. lupus familiaris

Linn.) are excluded from entry into Prairie Ridge Ecostation and have only been rarely caught on camera traps whereas coyotes are more commonly caught on camera traps and evidence of

coyote scat containing rodent ear tags was observed during this study (L. Gatens, personal

communication, 2021).

Our study is the first report of A. maculatum feeding on eastern chipmunk and five-lined

skink. Consistent with the known feeding patterns of A. maculatum immatures, the majority of

blood meals from host seeking adults representing the remnant of the nymphal blood meal were

identified as avian [14.8% (9/61)] and small mammal [59.0% (36/61)]. It is likely that immature

A. maculatum regularly feed on birds at Prairie Ridge Ecostation. Because the list of potential

bird hosts is prohibitively large for RLBH analysis to work at the species level, a better

understanding of avian host usage could be gained through amplification and sequencing of

DNA in tick blood meals using the approach of Gariepy et al. (2012) targeting the COI gene.

While the majority of blood meals were derived from cotton rats, blood meals from other small

mammals included deer mice, cricetidae, chipmunk and rabbit. At the time of this study, the

only species of Peromyscus recorded from the study site was the white-footed mouse

(Peromyscus leucopus Rafinesque). Other rodents in the family Cricetidae that are found in the study area included the eastern harvest mouse (Reithrodontomys humulis Audubon & Bachman) and the pine vole (Microtus pinetorum Le Conte).

The majority of identified A. americanum blood meals came from medium sized mammals (8/14; 57.1%) including coyote, gray fox, and raccoon while only one tick contained a blood meal from white-tailed deer although they were commonly observed on the study site during tick collections. A surprising number of overall blood meal identifications were derived from rodents (5/14; 35.7%) given what is known of the feeding predilection of all stages to feed on medium to large mammals (Bishopp and Trembley 1945, Kollars et al. 2000a, Kollars et al.

2000b).

Immature D. variabilis are known to utilize rodents and rabbits as hosts (Kollars et al.,

2000b). Our study reflects this feeding predilection, blood meals from rodents and rabbits represented 62.5% (15/24) and 16.7% (4/24) of the host blood meals identified. Among rodent blood meals, 93.3% (14/15) and 6.7% (1/15) were identified as cotton rat and Peromyscus mice, respectively. This compares well with what has been reported in host trapping in Oklahoma from similar grassland habitat showing 88.7% and 10.8% of D. variabilis nymphs fed on cotton rat and deer mice, respectively (Gage et al., 1992).

Multiple blood meals were detected in all three tick species examined. In this study multiple blood meal detections ranged from 4.9% (3/61) in A. maculatum to 25% (3/12) in A. americanum and compared well with adult ticks from RLBH studies conducted in Europe using the same fragment, which ranged from 3.1% (1/32) to 18.7% (40/214) in I. ricinus (Cadenas et al. 2007, Humair et al. 2007). Several researchers have suggested multiple blood meals may be due to interrupted feeding or from external contamination of the tick from physical contact with the host (Pichon et al. 2005, Cadenas et al. 2007). The sterilization technique used in our study eliminated external contamination of the tick surface with environmental DNA. Interrupted feeding and reattachment to a new host has been documented among several species of tick; the reasons for interrupted feeding include death of the host, host removal by scratching or grooming and the host’s immune response (Tahir et al. 2020). Interestingly, of the nine multiple blood meals detected in this study, all but two (rabbit + cotton rat) involved a prey + predator combination (either rabbit, cotton rat or Peromyscus sp. mouse + Canis sp., gray fox or raccoon).

These multiple blood meals may represent feeding interrupted by predation as nymphs left bodies of dead prey to feed on the predator. An alternative explanation for multiple blood meal hosts is that they may represent remnants of both the larval and nymphal blood meals in the host

seeking adult. This is unlikely, however, due to the degradation of the blood meal that would

occur over the period of time between engorgement of larvae and molting to adult.

In our study, blood meal host detection success was defined as amplification of the 12S

rDNA gene fragment. Detection of blood meal amplicons among adult tick species was highest

in A. maculatum (50.0%) and lowest in A. americanum (29.3%). Other blood meal studies that

used the same fragment of the 12S rDNA gene were successful in identifying hosts among

49.9% to 59.3% of adult I. ricinus in Europe (Cadenas et al. 2007, Humair et al. 2007) and in

50.3% A. americanum and 53.1% I. scapularis in North America (Scott et al. 2012). Possible explanations for the relatively lower detection success among A. americanum and D. variabilis

(39.2%) compared to A. maculatum may be the smaller sample size of these ticks analyzed, by

the conservative approach of excluding samples showing multiple non-specific hybridizations

and the age of the nymphal blood meal detected among host seeking adults.

Successful amplification of DNA in a tick blood meal is affected by the time since

engorgement. Kirstein and Gray (1996) successfully amplified cytochrome b fragments from the

blood meal of nymphs fed on mice as larvae just over 6 months post-engorgement but only 40%

of blood meals could be amplified after 9 months post-engorgement. Similarly, Pichon et al.

(2003) amplified a 120-150 bp fragment of 18S rDNA from the blood meal of nymphs fed as

larvae on mice and rabbit, successfully identifying both host blood meals in nymphs up to 7

months post-engorgement and only 40% of blood meals were detected from rabbits after 9

months; attempts failed to amplify DNA from a blood meal 10 months post-engorgement. In

our study, blood meal identification was lowest in June and began to rise just as both A.

americanum and D. variabilis populations showed a second abundance peak in June and July,

suggesting blood meals identified after June were from recently molted adults that began host-

seeking.

During this study, only two A. americanum nymphs were collected in May 2012. In

North Carolina, Ouellette et al. (1997) observed questing activity of A. americanum nymphs to

begin in early May with peak parasitization of hosts, such as raccoon, observed in July.

Assuming A. americanum adults fed as nymphs from May to July, a conservative estimate of the

age of blood meals over the course of our study would have ranged from 9 to 12 months; a

length of time over which blood meals would be expected to have been severely degraded. Thus

age related degradation may explain the relatively lower success in amplifying blood meals for

adult A. americanum than A. maculatum.

RLB Probe Specificity

In this study, 31.9% of PCR amplifications resulted in non-specific hybridizations on the RLBH

assay that appeared as clusters of hybridizations of three or more probes. Despite these non-

specific hybridizations, blood meal identifications could still be made by examining the pattern

of hybridizations on the membrane. The nested PCR approach used in our current study showed

a reduction in the number of non-specific amplification products but likely increased the

amplification of contaminating human DNA. Non-specific amplification of tick, human and

other extrinsic DNA has been observed in other molecular blood meal analyses. Using the same

12S rDNA fragment in their RLBH assay of A. americanum and I. scapularis blood meals, Scott

et al. (2012) adjusted many PCR and RLBH conditions to control nonspecific background reactions and to ensure ‘extraneous PCR products amplified from tick DNA’ were minimized.

Both Alcaide et al. (2009) and Gariepy et al. (2012) sequenced the DNA barcoding region of

COI to identify blood meals from engorged ticks removed from known hosts and observed

ambiguous sequences that could not easily be identified to host species. Alcaide et al. (2009)

believed these ambiguous sequences were multiple blood meals and matched several to

hypothetical hosts but also suggested some amplification of nuclear inserts or heteroplasmy

could have occurred. Gariepy et al. (2012) found that 6% of amplification products could not be

identified due to amplification of tick DNA or other extrinsic DNA and even found one sequence

that matched closely to a fungus. Given that the genomic DNA of both tick and prokaryotic

organisms is in far higher concentration in the DNA extract than the vertebrate DNA from a

remnant blood meal, it is likely non-specific PCR amplification using mitochondrial primers

occurs frequently. PCR reagents may also be a source of exogenous DNAs since they contain

bacteria and vertebrate derived DNAs that may amplify. Leonard et al. (2007) reported evidence

that 5% of PCR reactions from four labs processing ancient DNA samples contained vertebrate

DNA likely derived from PCR reagents. The contaminating DNA source was identified as

primarily cow (Bos taurus), pig (Sus scrofa) or chicken (Gallus gallus) although goat (Capra hircus), rabbit (Oryctolagus cuniculus) and guinea pig (Cavia porcellus) were also detected.

Cross hybridizations are common in RLBH assays because the potential for re-designing

probes is limited by the variability of the fragment amplified. Goessling et al. (2012), using the

same 12S rDNA fragment designed redundant vertebrate probes to assist in resolving cross

hybridizations, 23.8% (5/21) of their probes showed cross-hybridizations among eight vertebrate

tissues. Our study had a comparable rate of cross-hybridizations with 18.4% (7/38) probes

showing cross hybridizations with seven vertebrate tissues. Probes that cannot be redesigned

may still be able to identify host species if patterns of probe hybridizations can be established as

in Goessling et al. (2012) for distinguishing between cross-hybridizing squirrel species probes.

Scott et al. (2012) were able to distinguish between Peromyscus probes based on observed

patterns of cross-hybridization.

Blocking with casein proved to be the most useful in eliminating background

hybridizations but did not remove hybridizations with tick or other extrinsic DNA that appeared

as clusters of hybridizations. Similar to Scott et al. (2012), we found that a narrow range of

hybridization and post-hybridization wash temperatures were able to retain blood meal products while eliminating most non-specific hybridizations. Hybridization temperatures above 60°C and post-hybridization wash temperatures above 58°C may eliminate all hybridization signals on the membrane. Hybridization temperature did not seem to be as important as wash temperatures in reducing non-specific hybridizations. In initial trials of the assay, hybridization temperatures as low as 42°C showed similar non-specific hybridizations as hybridization at the upper limit of

58°C. Only blood meal identifications in ticks were counted if there were no ambiguous hybridizations evident over one or more analysis of the tick blood meal. Therefore, blood meal host determinations in our study are conservative and are more likely to have excluded some correct blood meal host identifications than included some false-positive identifications. The digital imaging system utilized in this study may have been more sensitive to non-specific

hybridizations than previous RLBH studies of tick blood meals which all relied on x-ray film to

detect chemiluminescence. The benefit of using the imaging system outweighed the costs in that

results that could be obtained in a short amount of time and allowed changes to be made to

exposure length and aperture settings to increase sensitivity of detection without the use of film

processing chemicals.

Temporal Distribution of A. maculatum

Weekly tick collections initiated in March 2012 showed that seasonal host-seeking activity of adult A. maculatum began abruptly in the third week of April in 2012, peaking in the first week of June and was absent by the beginning of September. This seasonal distribution compared favorably to reports of adult populations from Georgia, Oklahoma and South Carolina although questing began in May in Oklahoma and South Carolina and extended through October in

Georgia with peak abundance occurring in early June (Hixson 1940, Semtner and Hair 1973b,

Clark et al. 1998).

Spatial Distribution of Ticks

Over all study areas, the highest number of A. maculatum were collected from the prairie habitat. This area is typical of lowland prairie associated with A. maculatum populations in

Oklahoma (Semtner and Hair 1973b). Few A. americanum and D. variabilis were collected in the prairie habitat. Amblyomma americanum are typically most abundant in woodland/forested areas, likely due to larval requirements for higher humidity in shaded areas (Semtner and Hair

1973a, Goddard 2007). Sonenshine et al. (1966) classified vegetative habitat on a 16.2 hectare study site in the Piedmont of Virginia into three types: evergreen, woody deciduous and grasses and herbs. These authors correlated tick presence and host preference to small mammal communities within these habitat types. Dermacentor variabilis was found to be most abundant in woody deciduous habitat and moderately abundant in grasses and herbs. Amblyomma americanum preferred evergreen habitat and were found most abundant in all wooded areas than the grasses and herbs habitat (Sonenshine et al. 1966). Since A. americanum is a woodland tick

species, the low numbers collected in our study may be due to the landcover being primarily

grassland/old field habitat with less than 30% woodland/forest habitat.

Mark-recapture studies investigating the horizontal movement of several species of

ixodid ticks in response to CO2-baited traps show adult ticks move less than 10 meters during questing activities with some species, such as I. scapularis and I. ricinus moving no more than 2

meters (Koch and McNew 1982, Gray 1985, Falco and Fish 1991). In a mark-recapture study of

adult A. maculatum in Mississippi, movement did not exceed 7.5 meters and was often 2.5

meters or less (Goddard et al. 2011). Our study site is just over 15.4 hectares in size while each

collection area was roughly 4 hectares in size with a minimum spacing between areas of 20 m.

Given these distances, it is likely that small mammals found within study areas could travel

between study areas as they move about within their respective home ranges; however, it is

unlikely ticks moved between study areas unassisted.

Ticks may be transported over large areas while on larger vertebrates and even between

continents by migrating birds (Anderson and Magnarelli 2008). Home ranges for small

mammals such as rodents, moles, and shrews are generally less than 1 hectare in size, cotton- tailed rabbits average 1.6 hectare. Medium and large sized mammals may have large home ranges that extend over 60 hectares for opossum, in the 100’s for bobcat, red fox, and white- tailed deer and in the 1000’s of hectares for bear and coyote (Harestad and Bunnel 1979).

Lizards have smaller home ranges with all but the largest species having home ranges less than

0.5 hectares (Verwaijen and Van Damme 2008). Howell (1954) examined the home ranges of small mammals in old-field habitats in Tennessee and recorded the home range of the eastern harvest mouse as 0.67 hectares, cotton rat male home ranges averaged 0.34 hectares while those of females averaged 0.18 hectares, and white-footed mouse had the smallest home range size of

0.12 hectares. Movement of ticks during host seeking is far more restricted. Because small

mammals would move less throughout the study areas than larger mammals, small mammal

blood meals should show characteristic spatial distributions.

The only significant spatial distribution observed over the study site was in cotton rat and

bird blood meal host identifications. Cotton rat blood meals were detected at higher frequency in

the prairie habitat while bird blood meals were higher in the old pastureland habitat than in the

prairie habitat. The majority of non-cotton rat rodent blood meals (Peromyscus sp. and

cricetidae) were detected from ticks collected in the forest habitat. These findings may be

explained by habitat preferences of small rodents within the study area. Peromyscus sp. blood

meals were only detected in the forest habitat which is expected since woodland and forested

habitat is preferred by deer mice (Matlack et al. 2008). The larger home range of eastern harvest

mice along with a preference for grasslands and old fields with areas of thick vegetation may

explain the single cricetid blood meal that was detected within the old pastureland habitat

(Cawthorn and Rose 1989).

Rickettsia parkeri Infection Rate

Detection of R. parkeri in adult A. maculatum in our study (45/122; 36.9%) is higher than the

statewide infection rate reported by Varela-Stokes et al. (2011) for North Carolina (68/234; 29%)

but is comparable to data for Wake County, North Carolina (21/52; 40.4%). Fornadel et al.

(2011) reported a similar R. parkeri infection prevalence of 41.4% for A. maculatum collected in

Fairfax County, Virginia and (Wright et al. 2011) reported 43.1% from collections of the tick

made in southeastern Virginia. Lower infection prevalence have been reported in other states by

Paddock (2005) with R. parkeri among 15% of A. maculatum populations from Georgia, Florida,

Kentucky, Mississippi and Oklahoma and among 28% of A. maculatum tested from several

locations in Florida and Mississippi (Paddock et al. 2010).

The lack of infection in D. variabilis and low infection prevalence among A. americanum

is not surprising given that previous studies have also noted low natural infection prevalence of

R. parkeri among both tick species. In our present study, no D. variabilis were found infected

with R. parkeri, which is consistent with the findings reported by Varela-Stokes et al. (2011) for

D. variabilis sympatric with A. maculatum in North Carolina. Although D. variabilis is not a

recognized vector of R. parkeri they have been found infected with the pathogen at low rates.

Fornadel et al. (2011) found 0.3% prevalence among D. variabilis from Fairfax County,

Virginia. Similarly, Fritzen et al. (2011) found only 0.4% of D. variabilis collected from animals

in Kentucky were PCR positive for R. parkeri. Rickettsia parkeri was detected in 2.3% human-

attached D. variabilis submitted to the Texas Department of State Health Services (Williamson

et al. 2010). Remarkably, Kakumanu et al. (2018) detected R. parkeri among 14.7% of North

Carolina D. variabilis using RLBH of the 23S-5S IGS region of Rickettsia.

The low infection rate of 2.4% among adult A. americanum reported here is consistent

with Goddard and Norment (1986) who first reported R. parkeri in 0.7% of A. americanum

individually tested by hemolymph and indirect fluorescent antibody assay in ticks collected from

Kentucky, Mississippi, Oklahoma and Texas. Cohen et al. (2009) found 0.2% A. americanum

collected from humans and animals in Georgia and host seeking ticks from Tennessee to be

positive for R. parkeri. Using RLBH of the 23S-5S intergenic spacer (IGS) region of Rickettsia,

Lee et al. (2014) detected R. parkeri in 2.1% of A. americanum that had been removed from outdoor workers in North Carolina.

The low infection prevalence among A. americanum may be due to the effects of the bacteria on the tick survival. In established R. parkeri infected A. americanum lines, only 50%

(2/4) survived and only one of the surviving lines passed on the bacteria transovarially (Goddard

2003). The South American vector of R. parkeri, A. triste, has been shown to transmit R. parkeri through transstadial and transovarial transmission, negative effects were observed as higher nymphal mortality. Negative effects among nymphs are thought to have less of an effect on maintenance of rickettsiae in the population than negative effects on females that may otherwise lay thousands of rickettsia-infected eggs (Nieri-Bastos et al. 2013).

Low infection prevalence of a disease pathogen in A. americanum, an abundant tick in the southeastern and midwestern United States noted for its aggressive biting behavior, is epidemiologically important. Amblyomma americanum is a vector of several pathogenic bacteria including Francisella tularensis, R. rickettsii, Coxiella burnetti, Ehrlichia chaffeensis and E. ewingii (Childs and Paddock, 2003). Studies have described similarly low infection prevalence of R. rickettsii (Levin et al. 2017) and R. parkeri among A. americanum and a comparably high rate of infection with R. amblyommatis, a SFG rickettsia with unknown pathogenicity that may be responsible for cases of probable SFG rickettsia reported to the CDC annually (Egizi et al.

2020). While prevalence of pathogenic bacteria may be low, A. americanum is more likely to encounter and bite humans and therefore is an important vector regardless of the low prevalence of some pathogenic bacteria.

No statistical association was detected between blood meal host identifications and R. parkeri infection even though infection prevalence was higher among adult A. maculatum with an identifiable blood meal (27/61; 44.3%) than without (18/61; 29.5%). Infection is known to occur when high levels of rickettsia are circulating in the blood (Burgdorfer and Brinton 1975).

Although cotton rats were frequently fed upon in our study, it may be that they do not become

rickettsemic at a high enough level to infect naïve ticks. Moraru et al. (2013a) fed A. maculatum

larvae and nymphs on R. parkeri inoculated cotton rats and bobwhite quail; R. parkeri could not be detected in bobwhite quail tissues and while cotton rat tissues maintained rickettsia for 7 days,

R. parkeri was not detected in the nymphs feeding upon rickettsemic cotton rats. The lack of an association between R. parkeri infected ticks and potential vertebrate reservoirs is not surprising given transovarial transmission of SFG rickettsiae. While the cotton rat is an important host for immature A. maculatum, with rates of 83.7% transovarial and 100% transstadial transmission reported by Wright et al (2015) among field collected A. maculatum, it is likely R. parkeri is maintained among A. maculatum at Prairie Ridge Ecostation through transovarial and transstadial transmission.

Conclusion

Our study represents the first use of molecular methods to identify blood meal hosts in field- collected A. maculatum and D. variabilis in North America. The spatial distribution of tick species as determined in our study was concordant with host blood meal molecular identifications among tick species. Our investigation was limited in scope to one prairie habitat in the Piedmont of North Carolina and a single pathogen, Rickettsia parkeri. Multiple SFG rickettsia are known to infect the ticks in North Carolina including Candidatus Rickettsia andeanae (Varela-Stokes et al. 2011), Rickettsia amblyommatis, R. bellii, R. massiliae, R. montanensis, R. rickettsii and there is evidence that new (Lee et al. 2014) and potentially pathogenic rickettsia (Wilson et al. 2020) are present. Studies from other states have found

Candidatus Rickettsia andeanae infecting A. maculatum although it is of unknown pathogenicity

(Ferrari et al. 2012, Jiang et al. 2012, Paddock et al. 2015, Lee et al. 2017). In order to fully characterize host feeding patterns and pathogen infection among A. maculatum, blood meals would have to be characterized from additional populations and with a more comprehensive investigation into all SFG rickettsia likely to infect this tick and other important vector species such as A. americanum and D. variabilis.

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Table 1. Oligonucleotide primers used in this study to amplify vertebrate and pathogen DNA.

Target Primer Name 5’-3’ Sequence Amplicon Size (bp)

Bacterial 16S rDNA 0206 CCTACGGGAGGCAGC 744

0209 TCGTTGCGGGACTTAAC

Rickettsia genus 17-kDa protein 17kD1 GCTCTTGCAACTTCTATGTT 434

- Outer primers 17kD2 CATTGTTCGTCAGGTTGGCG

Rickettsia genus 17-kDa protein 17kN1 CATTACTTGGTTCTCAATTCGGT 232

- Inner primers

17kN2 GTTTTATTAGTGGTTACGTAA

Table 1 (continued).

447

Rickettsia parkeri outer RpompAF AATGCAGCATTTAGTGATGATGTTAA

membrane protein A RpompAR TCCTCCATTTATATTGCCTG

Vertebrate 12S rDNA - Outer 12S-12F TGCCAGCCACCGCGGTCA 600

primers 12S-13R AGGAGGGTGACGGGCGGT

Vertebrate 12S rDNA - Inner 12S-6F CAAACTGGGATTAGATAC 145

primers B-12S-9R* ACAGGCTCCTCTARR

* developed for this study

Host(s) Identified Amblyomma Amblyomma Dermacentor

Maculatum americanum variabilis

No. No. % b % No. No. % b % No. % b

blood RpompA+ blood RpompA+ blood RpompA+ blood RpompA+ blood blood

meal meal meal meal meal meal

Aves 9 (4)a 13.9 (8.5)c 1 8.3 1 5.0

Plestiodon inexpectatus 1 1.5

Table 2 (continued).

Cricetidae (small 2 (2) 3.1 (4.3) 1 8.3 rodent)

Peromyscus sp. 1 8.3 1 5.0

Sigmodon hispidus 38 (15) 58.5 (31.9) 1 8.3 11 55.0

Tamias striatus 1 1.5

Sylvilagus sp. 2 (1) 3.1 (2.1) 1 5.0

Didelphis virginiana 1 5.0

Canis sp. 7 (3) 10.8 (6.4) 2 16.6 1 5.0

Urocyon 1 (1) 1.5 (2.1) 2 (1) 16.6 (100.0) cinereoargenteus

Procycon lotor 1 8.3

Table 2 (continued).

Odocoileus virginianus 1 1.5 1 8.3

Sylvilagus sp.+ U. 1 5.0 cinereoargenteus

Sylvilagus sp.+ S. 2 10.0 hispidus

Procycon lotor + S. 1 8.3 1 5.0 hispidus

Canis sp. + S. hispidus 2 (2) 3.1 (4.3) 1 8.3

Canis sp. + Peromyscus 1 (1) 1.5 (2.1) sp.

Table 2 (continued).

Total blood meal 65 (29) 50.8 (61.7) 12 (1) 27.9 (100) 20 (0) 39.2 (0) identified

Total no blood meal 63 (18) 49.2 (38.3) 31 (0) 72.1 (0) 31 (0) 60.8 (0) detected

aNumber of ticks infected with Rickettsia parkeri. bPercentage of the total number of blood meal hosts identified. cPercentage of total number of ticks infected with Rickettsia parke

Figure 1. Example probe validation with vertebrate tissues. Probes that were combined in lanes on this membrane were individually validated against tissues and occupied separate lanes during the assay. Most cross-hybridizations shown in this illustration have since been resolved by re- extracting DNA from fresh tissue samples (Ophisaurus ventralis, Anolis carolinensis, Sceloporus undulatus, Sorex longirostris, Condylura cristata, and Bos taurus).

(a)

(b)

(c)

Figure 2. Seasonal distribution of (a) A. maculatum, (b) A. americanum and (c) D. variabilis.

Ticks were collected by 1m2 tick drag cloth for three hours on each date. Host seeking A.

americanum nymphs were collected in May 2012. Amblyomma maculatum nymphs were found

engorged on cotton rat in February and October 2012, host seeking nymphs were collected in

September 2011 and June 2012.

Supplemental Material

Appendix A

Sensitivity and Specificity of Reverse Line Blot Hybridization

During probe validation, human DNA extract cross-hybridized weakly with short-tailed shrew

(Blarina carolinensis) and swine probes and hybridized strongly with the rat probe. These cross- hybridizations were useful in evaluating the extent of human DNA contamination since they occurred together in a pattern of mammal-short-tailed shrew-rat-swine. From the cross hybridizations noted with human DNA, only the short-tailed shrew has been recorded on the site, therefore blood meals from this species may have been underestimated using this approach by their misidentification as blood meals taken from a human.

In addition, during the RLB assay, patterns of non-specific hybridizations were noted that did not appear during probe validation using known vertebrate DNA extracts. These non- specific hybridizations diminished slightly after blocking the membrane with casein; however, they were still visible and thus likely represented exogenous DNA and not true vertebrate blood meals. Since these patterns were fairly consistent, they were not considered as true blood meals and were ignored. The patterns included aves-turkey-skink, woodchuck and/or squirrel-flying squirrel-cricetidae-rat, and weasel-felis-swine-white tailed deer. Only single hybridizations of one of these probes was counted as a true host blood meal. No turkeys had been recorded at

Prairie Ridge Ecostation prior to or during this study, which suggested that other extrinsic DNA, such as tick DNA, may have resulted in these hybridizations. To evaluate non-specific PCR product amplification from tick DNA using 12S rDNA primers, legs from all three tick species were removed and extracted separately. Amplicons of 12S rDNA from tick leg extracts showed strong hybridized to aves, turkey and ground skink probes and weakly with the cricetidae probe.

Attempts to sequence these non-specific PCR products from two A. maculatum from this study

resulted in sequence matches of 78% similarity to Aspergillus sp. (NW_001884680).

Regardless of the source of these non-specific DNA hybridizations, all turkey probe hybridizations were excluded from this analysis. Bird probe hybridizations were only accepted if no other hybridizations occurred with turkey or skink probes. Thus, like short-tailed shrew, the number of bird blood meals was likely underestimated.

Appendix B

Figure S1. Prairie Ridge Ecostation, Raleigh, Wake County, North Carolina study site showing

2012 tick collection areas A, B and C. Collection areas were approximately 10 acres in size.

The study site is dominated by former agricultural land containing herbaceous vegetation

(77.2%) with woodland and forested areas (22.8%). Area A is old pastureland that has undergone reconstruction to piedmont prairie. Area B is old pastureland with lowland and bottomland forest. Area C contains lowland and upland forest with bottomland forest.

Appendix C

Table S1. Sequences of 5’ amine labeled probes designed to complement a 145 bp fragment of 12S rDNA. All vertebrate tissues listed were evaluated against the entire suite of probes in order to validate probes and evaluate cross-hybridizations with vertebrate tissues.

Probe Name Probe Sequences (5’-3’) Probe Conc. Vertebrate species

(pmol)

Avesac TACGAGCACAAACGCTTAA 450 Meleagris gallopavo, Colinus virginianus,

Coturnix japonica, Zenaida macroura,

Strix varia, Thryothorus ludovicianus,

Junco hyemalis

Turkeyc ATCTTGATACTAATATACTCACGTATCC 450 Meleagris gallopavo

Quail TCCAGATACCCCCAATACCAAC 450 Colinus virginianus, Coturnix japonica*

Lacertillaa GAGAACTACAAGTGAAAAACT 1200 Ophisaurus attenuatus, O. ventralis,

Plestiodon fasciatus, P. laticeps, Scincella

lateralis†

Glass Lizard GCTTAGTCCTAAYCTCAGATATTAAC 1050 Ophisaurus attenuatus, O. ventralis

Table S1 (continued).

Anole ACTAAAGTGTTCGCCAGAATATTACG 750 Anolis carolinensis

Fence Lizard CAGAAAACTACGAGCGAAAAGCTTA 750 Sceloporus undulatus

Plestiodon sp. TGAAAAACTTAAAACTCCAAGGACTT 1200 Plestiodon fasciatus, P. laticeps

Skinkc

Ground Skinkc AACAACGATCTTATCTYACAATATTATCC 750 Scincella lateralis

Racerunner TAACAATTTGTCCGCCAGAAAATT 1200 Cnemidophorus sexlineatus

Mammaliabc AAACTCAAAGGACTTGGC 1200 All mammal tissues listed below, Homo

sapiens, Lasiurus borealis, Eptesicus

fuscus

Opossum GCTTAGTAATAAACTAAAATAATTTAACAAACA 1200 Didelphis virginiana

Long-tailed CTAACAAAAATACCCGCCAGAGAA 900 Sorex cinereus, S. dispar, S. palustris, S.

Shrewd longirostris, S. hoyi

Short-tailed & AATTAACAAAACTACTCGCCAGAGGA 900 Blarina brevicauda, B. carolinensis,

Least Shrewcd Cryptotis parva

Mole ACTAAGACAATCCAACTAACAAGATT 900 Scalopus aquaticus, Condylura cristata

Table S1 (continued).

Rabbitd ACTTAAATAATTCCATAACAAAATTACTCG 900 Sylvilagus palustris, S. floridanus, S.

obscurus

Chipmunk CTTAAACACAAATACTTAATAAACAAGAGTATT 1200 Tamias striatus

Woodchuckc AGAGTACTACTAGCAATAGCCTGAA 600 Marmota monax

Squirrelcd AGAGAACTACTAGCCACTGCTTA 900 Sciurus carolinensis, S. niger

Flying Squirrelc GCTTAGCCCTAAACACAAATATTTAA 900 Glaucomys volans

Cricetidaec ATTTGCCYGAGAACTACTGGC 1200 Oryzomys palustris, Reithrodontomys

humulis, Ochrotomys nuttalli, Peromyscus

gossypinus, P. leucopus, P. maniculatus,

Microtus chrotorrhinus, M.

pennsylvanicus, M. pinetorum, Myodes

gapperi, Ondatra zibethicus, Synaptomys

cooperi

Peromyscus mice CTAAACCTYAAAGATTAAATAACAAAATCAT 1050 Peromyscus gossypinus, P. leucopus, P.

maniculatus

Table S1 (continued).

Cotton Rat CTAAACCACAATAACTTAAAAACAAAGTT 900 Sigmodon hispidus

Woodrat CTAAACCCTAATAATTCAATAACAAAAT 1200 Neotoma floridana

Ratcd GCTTAGCCCTAAACCTTAATAATTA 750 Rattus norvegicus

House Mouse CCATAAACCTAAATAATTAAATTTAACAAAACT 750 Mus musculus

ATTT

Canis sp. AACATAGATAATTTTACAACAAAATAATTCG 900 Canis latrans, Canis lupus familiaris

Red Fox CATAAATAGTTCTATAACAAAACAATTCG 1200 Vulpes vulpes

Gray Fox ACATAAACAGTTCTATAAACAAAATAGTT 1200 Urocyon cinereoargenteus

Bear AGCCTTAAACATAAGTAATTTATTAAACAAA 1200 Ursus americanus

Raccoonc ATTAACGTAACAAAATTATTTGCCA 1050 Procyon lotor

Minkd ATTCACATAACAAAATTACTTGCCA 1050 Neovison vison

Weaselc ATTTACATAACAAAATTATTTGCCA 1050 Mustela frenata, M. nivalis

Skunk CCATAAACACAGACAATTAATATAACAAA 1350 Mephitis mephitis

Felidc AAACAAAACTATCCGCCAGAGAA 900 Lynx rufus

Swinecd TAAACCCAAATAGTTACATAACAAAACTAT 600 Sus scrofa

Table S1 (continued).

White-tailed AACATAAATAGTTATATAAACAAAACTATTCG 1350 Odocoileus virginianus

Deerc

Cow CCTAAACACAGATAATTACATAAACAAAAT 1200 Bos taurus aProbe designed by Humair et al. (2007). bModified from Humair et al. (2007). cProbe hybridized with exogenous DNA in PCR amplicon. Since these occurred in patterns of three or more probes, only single hybridizations with this probe were counted as a true bloodmeal. dCross hybridization noted. Long-tailed shrew probe strongly cross-hybridizes with Least shrew (Cryptotis parva) tissue; Short- tailed shrew probe weakly cross-hybridizes with Human (Homo sapiens); Rabbit probe weakly cross-hybridizes with Mink (Neovison vison); Squirrel probe strongly x-hybridizes with Marsh rice rat (Oryzomys palustris). Rat probe strongly x-hybridizes with human.

Mink probe strongly x-hybridizes with Eastern cottontail rabbit (Sylvilagus floridanus). Swine probe weakly cross-hybridizes with human.

*DNA did not hybridize to probe.

†DNA hybridizes weakly to probe.

Appendix D

Female Male Nymph

Figure S2. Spatial distribution of (a) A. maculatum, (b) A. americanum and (c) D. variabilis

collected by 1 m2 tick drag for one hour in each area (three hours total for the entire study site).

Collection areas were not designated until 2012. Area habitats included restored prairie (A), old pastureland (B) and forest (C). Nymphs removed from cotton rat were collected in area A. Host

seeking nymphs of A. maculatum were collected in areas A and B while those of A. americanum

were collected in areas B and C.

Appendix E

Table S2. Amblyomma maculatum host bloodmeal determinations by stage and year/area collected (Number R. parkeri infected).

Female Male Host Identified Nympha Total 2011 A B C 2011 A B C

Bird 5 (2) 1 (1) 2 (1) 1 9 (4)

Five-lined skink 1 1

Small rodent (cricetidae)b 2 (2) 2 (2)

Cotton rat 4 (2) 4 (1) 10 (5) 2 (1) 5 11 (6) 1 1 38 (15)

Chipmunk 1 1

Rabbit (Sylvilagus sp.) 1 1 (1) 2 (1)

Canis sp. 1 1 (1) 1 3 (1) 1 (1) 7 (3)

Gray fox 1 (1) 1 (1)

White tailed deer 1 1

Canis sp. + Peromyscus sp. 1 (1) 1 (1)

Canis sp. + cotton rat 1 (1) 1 (1) 2 (2)

Table S2 (continued).

No bloodmeal DNA amplified 2 7(1) 5(4) 9(2) 10(3) 5(0) 6(2) 8(1) 11(5) 63(18) aBloodmeals from nymphs found engorged on cotton rat, unfed nymphs collected in areas A and B did not amplify a bloodmeal. bSmall rodent (Cricetidae) represents members of the family that did not hybridize to Peromyscus sp. or cotton rat probes, likely eastern harvest mice (Reithrodontomys humulis) or woodland vole (Microtus pinetorum).

Appendix F

Table S3. Amblyomma americanum host bloodmeal determinations by tick life stage and year/area collected (Number R. parkeri infected).

Host Identified Nymph Female Male Total

2011 A B C 2011 A B C

Bird 1 1

Small rodent (cricetidae) 1 1

Peromyscus sp. 1 1

Cotton rat 1 1

Canis sp. 1 1 2

Gray fox 1 (1) 1 2 (1)

Raccoon 1 1

White tailed deer 1 1

Cotton rat + Raccoon 1 1

Cotton rat + Canis sp. 1 1

No bloodmeal DNA amplified 2 1 2 3 9 0 1 3 10 31

Appendix G

Table S4. Dermacentor variabilis host bloodmeal determinations by tick sex and year/area collected.

Host Identified Female Male Total

2011 A B C 2011 A B C

Bird 1 1

Peromyscus sp. 1 1

Cotton rat 2 1 2 2 1 3 11

Rabbit (Sylvilagus sp.) 1 1

Opossum 1 1

Canis sp. 1 1

Rabbit + Gray fox 1 1

Rabbit + cotton rat 1 1 2

Cotton rat + raccoon 1 1

No bloodmeal DNA amplified 2 3 8 2 8 0 2 6 31

CHAPTER 2

Molecular Identification of Blood meals and Borrelia burgdorferi sensu lato Infection

Prevalence among Sympatric Ixodes affinis and Ixodes scapularis (Acari: Ixodidae) in the

Coastal Plain of North Carolina

This chapter was formatted for submission to The Journal of Medical Entomology with the coauthors Loganathan Ponnusamy1,2, Alden S. Estep3, and Charles S. Apperson1,2

1Dept. of Entomology and Plath Pathology, North Carolina State University, Raleigh, NC 27695

2Comparative Medicine Institute, North Carolina State University, Raleigh, NC 27695 USA

3USDA ARS Center for Medical, Agricultural, and Veterinary Entomology, Gainesville, FL

32608

Abstract

Molecular techniques were employed in an effort to identify important associations between vertebrate hosts and Lyme borreliosis group spirochetes prevalence among the enzootic vector, Ixodes affinis Neumann and the bridge vector, I. scapularis Say from the Coastal Plain of

North Carolina. Pathogen prevalence was determined by PCR amplification of the Borrelia burgdorferi sensu lato (s.l.) flaB gene, rrfA-rrlB IGS, and a B. burgdorferi sensu stricto (s.s.)- specific rrs-rrlA ITS fragment while blood meal hosts were identified using a reverse line blot hybridization (RLBH) assay of a 12S rDNA gene fragment against 38 vertebrate probes.

Statistically significant differences were observed in B. burgdorferi s.l. infection between I. affinis (80%; 68/85) and I. scapularis (46.8%; 44/94). Based on amplification and Sanger sequencing of the flaB gene fragment, we report a higher prevalence of B. burgdorferi sensu stricto (s.s.) among both I. affinis (52.9%;55/85) and I. scapularis (35.1%; 33/94) than previously reported. Borrelia bissettii, known to be a human pathogen in Europe, was detected at a lower prevalence among I. affinis (11.8%; 10/85) and I. scapularis (4.3%; 4/94). Sanger and nanopore sequencing of multiple loci indicate these sympatric ticks are likely co-infected with multiple B. burgdorferi s.l. genospecies. Although no statistically significant association between blood meal host and pathogen prevalence was observed among I. affinis or I. scapularis, we found cotton rat, Sigmodon hispidus Say and Ord, and Canis spp. were the most frequently detected blood meals in both ticks; these shared hosts likely play an important role in LD transmission in the Coastal Plain of North Carolina.

Keywords: blood meal identification, Ixodes affinis, Ixodes scapularis, Borrelia burgdorferi

Short title: Host-pathogen associations in sympatric enzootic vectors of Lyme borreliosis

Introduction

Since Lyme borreliosis (Lyme disease) became a reportable disease in 1991, it has

remained the most commonly reported arthropod-borne disease in the United States (Barbour

and Fish, 1993, Bacon et al. 2008, CDC, 2014) with approximately 300,000 cases reported each

year (Meade et al. 2015). Lyme disease (LD) is caused by a bacterium, Borrelia burgdorferi

Burgdorfer. The LD pathogen belongs to a group of 20 described genospecies in the B. burgdorferi sensu lato (s.l.) complex, 11 of which are known human pathogens (Franke et al.

2013, Pritt et al. 2016). Eight genospecies in B. burgdorferi s.l. complex have been identified in

North America including B. burgdorferi sensu stricto (s.s.), B. andersonii (Marconi et al. 1995),

B. bissettii (Postic et al. 1998), B. californiensis (Postic et al. 2007), B. carolinensis (Rudenko et

al. 2009a), B. americana (Rudenko et al. 2009b), B. kurtenbachii (Margos et al. 2010) and most recently, B. mayonii (Pritt et al. 2016). Only B. burgdorferi s.s. and B. mayonii are known to

cause LD in the United States. However, evidence for B. bissettii pathogenicity has been

reported in Europe. Recent evidence suggests that a B. bissettii-like pathogen along with B.

andersonii and B. americana may be clinically significant, having been associated with patients

exhibiting signs and symptoms of LD in the United States (Rudenko et al. 2009c, Girard et al.

2011, Clark et al. 2014). The other three genospecies, B. californiensis, B. kurtenbachii and B.

carolinensis have not yet been associated with illness. The majority of LD case reports in the

United States are distributed in the northeastern and upper midwestern states. Despite the low

incidence of LD reports from southern states, Lyme Borrelia have long been known to infect

ticks and vertebrates in this region (Oliver et al. 1993a) and the genospecies circulating in the

southern states represent greater genetic variability than northern strains (Rudenko et al. 2009a).

The incidence of indigenous cases of LD from North Carolina and other southern states

has been historically low since the first LD cases from North Carolina have been reported as

early as 1983 (Pegram et al. 1983). As of 2019, the incidence of LD in North Carolina was 3.3

cases per 100,000 population and although the state itself is considered to have a low incidence

of endemicity, case numbers have continued to rise annually with several counties exceeding the

10 cases per 100,000 population that is considered high incidence according to the CDC’s 2017

case definition for LD (CDC, 2017, NCDHHS, 2020).

The primary vector of B. burgdorferi s.s. in the eastern United States is the blacklegged

tick, Ixodes scapularis Say and it is distributed east of the 100th meridian with highest densities in the northeast, mid-Atlantic and upper Midwest states (Dennis et al. 1998, Brownstein et al.

2003, Brownstein et al. 2005, Diuk-Wasser et al. 2006). In addition, an important enzootic vector of B. burgdorferi s.s. in Georgia and South Carolina, I. affinis, has been reported in

Florida, Georgia, South Carolina, North Carolina, and Virginia (Harrison et al. 2010, Nadolny and Gaff, 2018). Unlike I. scapularis, a generalist feeder, I. affinis has a limited host range of 15 mammal and one bird species (Oliver et al. 1987, Harrison et al. 2010). It is reported that this tick may only rarely bite humans (Oliver, 1996).

Studies of B. burgdorferi s.l. prevalence among I. affinis and I. scapularis in the southern

United States have reported significantly higher infection prevalence among I. affinis (26%) than

I. scapularis (1.4%) from South Carolina (Clark et al. 2002) and among I. affinis (63.2%) and I. scapularis (0%) in North Carolina (Maggi et al. 2010). The narrow host range of I. affinis coupled with the previously reported B. burgdorferi s.l. infection prevalence potentially makes this species an important maintenance host for the LD pathogen in the southeastern United

States. Determining the host feeding patterns of these sympatric species may provide insight

into why LD is not commonly reported from North Carolina and identify reservoir hosts that sustain enzootic pathogen transmission in the coastal plain of North Carolina.

Maintenance hosts provide blood meals to sustain vector populations while competent reservoir hosts sustain pathogen transmission. Maintenance hosts may act as dilution hosts when they are fed upon more frequently than competent reservoir hosts, thus diluting the pathogen within the vector population. Ixodes scapularis is known to feed on 125 species including 54 mammal, 57 bird and 14 lizard species (Lane et al. 1991, Keirans et al. 1996). In the southern

United States, three groups of animals have been considered as holding important roles in maintaining populations of I. scapularis and B. burgdorferi s.l.: deer, rodents and lizards.

Although not thought to be competent reservoirs for B. burgdorferi s.l. (Telford et al.

1988), white-tailed deer (Odocoileus virginianus Zimmermann) are hosts for all three life stages

of I. scapularis and with large home ranges, these animals are capable of transporting adult females and their progeny into areas uncolonized by the tick (Madhav et al. 2004, Piesman,

2002). In LD endemic areas, rodents are hosts for larval and nymphal I. scapularis, and many

are amplifying hosts including chipmunks, rats and mice, in particular the white-footed mouse

(Peromyscus leucopus Rafinesque). In North Carolina, I. scapularis have rarely been found attached to rodents, including white-footed mice (Apperson et al. 1993, Ginsberg et al. 2021).

Reptiles have been reported to be important hosts of immature I. scapularis in the southern

United States (Rogers, 1953).

Ixodes scapularis are known to parasitize a variety of lizards in the southern states

(Oliver et al. 1993b; Levine et al. 1997). Like white-tailed deer, lizards are not considered competent reservoirs of B. burgdorferi s.l. and may function as dilution hosts. In the southeast, rodents are less available than lizards during months that immature ticks are active. In Onslow

County, North Carolina, I. scapularis were observed to heavily parasitize lizards during summer months when infestation of rodents was low (Apperson et al. 1993).

Many studies have utilized molecular methods to identify tick pathogens and vertebrate hosts from tick blood meals, eliminating the need to trap and handle wildlife (Kent, 2009).

Molecular methods have the potential to give a broader insight into host species that are difficult to trap or due to timing may miss tick attachment on hosts and thus may give a less biased view of host feeding preferences of ticks. Identification of pathogens and host species from tick blood meals using molecular methods is well suited to ticks. Unlike other hematophagous arthropods, digestion in ticks is a slow intracellular process, taking months to complete (Balashov, 1972,

Tarnowski and Coons, 1989). Of the molecular methods for the identification of arthropod blood meals, reverse line blot hybridization (RLBH) has been identified as a valuable method for screening multiple samples simultaneously against up to 43 pathogen or vertebrate host probes on each run of the assay (Kong and Gilbert, 2007). This method may allow clear identification of individual blood meal hosts when ticks feed on multiple hosts, avoiding the ambiguous results in mixed genomic samples often encountered in conventional PCR followed by Sanger sequencing.

The principal objective of our study was to evaluate the potential for zoonotic transmission of the LD spirochete in the Coastal Plain region of North Carolina by determining the prevalence of B. burgdorferi s.l. infection and identifying blood meal hosts of sympatric I. affinis and I. scapularis using molecular methods. We aimed to determine if an association between vertebrate host and B. burgdorferi s.l. pathogen infection prevalence for each Ixodes tick species could be identified. Determining shared hosts between I. affinis, an enzootic vector of B. burgdorferi s.l. in the southern states, and the bridge vector transmitting the pathogen to

humans, I. scapularis would shed more light into the ecoepidemiology of Lyme in the southern

states and identify important vertebrate reservoirs for LD management.

Materials and Methods

Study Sites and Field Tick Collection

Study sites were located in Martin, Onslow and Wayne Counties within the Coastal Plain of North Carolina where sympatric populations of I. affinis and I. scapularis have been observed

(Figure 1). The study site in Martin County (35 º 59'11''N, 77º17'47''W) covered a 12 hectare area along a gravel road and utility right of way located in the Conoho Farms tract of the Lower

Roanoke River Wetlands Game Land in the Devereaux Swamp. Land cover at the Martin county site was primarily cypress-gum floodplain forest, with agriculture fields, successional deciduous forest, pocosin woodlands and shrublands and piedmont mixed successional forest. Onslow

County collection sites covered an area of 22 hectares located on Marine Corps Base Camp

Lejeune (34º34'58''N, 77 º21'40''W). Land cover at collection sites on Camp Lejeune were composed primarily of managed coniferous forest plantation and coastal plain nonriverine wet flat forest with Coastal Plain mixed bottomland and little wet longleaf or slash pine savannas, dry mesic oak pine forest and pocosin woodlands and shrublands. The Wayne County study site collection area covered approximately 13 hectares along hiking trails in the Cliffs of the Neuse

State Park (35°13'31''N, 17°53'37''W). Land cover was composed primarily of Coastal Plain mixed bottomland, coastal plain dry to dry-mesic oak forest, coastal plain mixed bottomland, coniferous cultivated plantation and dry mesic oak pine forests.

Host-seeking ticks were collected over thirteen dates from April 2011 to May 2013, no collection attempts were made in October, December or January of any year. Ticks were

collected for one hour at each location using 1 m2 drag cloths made of corduroy cloth. In the field, ticks were transferred into vials of 95% ethanol and stored at -20°C until identification and

DNA extraction. Tick species and stages collected were identified morphologically using relevant taxonomic keys (Keirans and Litwak, 1989, Keirans and Durden, 1998). Ixodes spp. immatures were identified using Oliver et al. (1987) and Durden and Keirans (1996), and adults using Keirans and Clifford (1978).

DNA Extraction

To prevent contamination of samples with foreign DNA, extraction of total DNA from ticks, preparation of samples for PCR amplification, and product analysis were conducted in separate rooms. Extraction and PCR workspaces along with equipment thoroughly cleaned prior to use.

Separate pipettes and filtered tips were used for DNA extraction and setup of PCR assays. Prior to DNA isolation, individual ticks were surface sterilized by immersion and shaking for 30 s each in 1% sodium hypochlorite, 0.1 M phosphate buffered saline (PBS) and 95% ethanol followed by three additional 1 min washes in 0.1 M PBS. One sample of the final wash in PBS was retained from each set of extractions to attempt amplification of vertebrate DNA to evaluate surface sterilization effectiveness.

DNA was extracted using the phenol-chloroform method after processing in a modified

lysis buffer composition based on Smith et al. (2010). Ticks were minced individually with

sterile #11 scalpel blades in 160 µL lysis buffer 1 [2% (w/v) sodium dodecyl sulfate, 100 mM

tris(hydroxymethyl)-aminomethane hydrochloride (Tris-HCl), 200 mM sodium chloride (NaCl),

10 mM ethylenediaminetetraacetic acid (EDTA), pH 7.0] and 20 µL each of freshly prepared

lysozyme (100 mg/mL) and proteinase K (20 mg/mL). After a 1 h incubation at 37°C, 200 µL of

lysis buffer 2 [1 % (w/v) cetyltrimethylammonium bromide, 1.5 M NaCl, 100 mM Tris-HCl and

100 mM EDTA] was added to each sample and incubated at 56°C for 2 h. Negative extraction

controls were included with each round of tick extractions.

The extracted DNA was subjected to an additional purification using silica spin columns

(EconospinTM, Epoch Life Sciences, Sugarland, TX, USA) following the protocol of Smith et al.

(2010) and manufacturer recipes for DNA absorption and wash buffers. Purified DNA was then

eluted in a total volume of 100 µL sterile water by two centrifugations at 5900 g for 60 s using

50 µL sterile water each. DNA quantity and quality were evaluated using a NanoDrop ND1000

spectrophotometer (Thermo Scientific, Waltham, MA, USA). Genomic DNA extracts were

stored at -20°C.

Pathogen Detection

Ticks used in bloodmeal analysis were evaluated for Borrelia burgdorferi s.l. infection by

PCR amplification of three molecular markers. Positive (B. burgdorferi B31 from a positive

tick), negative (nuclease free water), and negative DNA extraction (nuclease free water without

tick) controls were run alongside each PCR reaction. PCR amplification of the chromosomal flagellin gene (flaB) followed methods described by Clark et al. (2005); 5S (rrfA) - 23S (rrlB)

intergenic spacer (IGS) region of Borrelia spp. and B. burgdorferi s.s. specific sequence within

the 16S-23S (rrs-rrlA) internal transcribed spacer (ITS) region followed Maggi et al. (2010). In

addition, all B. burgdorferi s.l. positive ticks were submitted for Sanger sequencing using purified amplicons of a larger, 840 bp flaB fragment using primers designed for this study, forward primer flaB 103F and reverse primer flaB 943R (Macrogen, USA). Table 1 lists all

primers used in this study. Sequence quality was evaluated by reviewing chromatograms with

4Peaks Software Version 1.7.1 available at www.nucleobytes.com (Griekspoor and Groothuis,

2006) and aligned using CLUSTALX 2.1 (Larkin et al. 2007). Sequences were compared to sequences deposited in the GenBank® database (http://blast.ncbi.nlm.nih.gov) using the Basic

Local Alignment Search Tool (BLASTn). Sequences were aligned using CLUSTALX 2.1

(Larkin et al. 2007) and trimmed to 601 bp. Phylogenetic trees were constructed using the partial flaB by the maximum likelihood method based on the Tamura-Nei model (Tamura and Nei,

1993) using MEGAX (Kumar et al. 2018, Stecher et al. 2020).

A nested PCR of a fragment of flaB was used to screen all ticks for B. burgdorferi s.l.

The outer 497 bp flaB fragment was amplified in a Bio-Rad PTC-100 thermal cycler in a 25 µL reaction volume using 0.5 µM each of flaB outer primers, 1× AmpliTaq Gold 360 Master Mix

(Applied Biosystems, Carlsbad, CA), and 5 µL of DNA extract. An initial denaturation at 95°C for 10 min was followed by 40 cycles of 94°C for 30 s, 52°C for 30 s, 68°C for 45 s and a final cycle of 68°C for 5 min. The inner 389 bp fragment of flaB was amplified for an additional 30 cycles using similar PCR conditions as above with 2 µL of the outer fragment PCR product as template DNA and a 55°C annealing temperature.

An 840 bp flaB fragment was amplified using primers flaB 103F and flaB 943R in a

Mastercycler Gradient PCR thermal cycler (Eppendorf North America, Inc.; Hauppage, NY) using a 25 µL reaction volume containing 0.5 U Platinum™ Taq DNA Polymerase (Thermo

Fisher Scientific, Waltham, MA), 0.2 mM dNTPs, 1.5 mM MgCl2, 1x Buffer (IDT, Coralville,

IA), and 0.40 µM of each primer and 2 µL DNA extract. Touchdown PCR conditions included an initial denaturation step for 3 min at 94°C, followed by 10 cycles of 30 s at 94°C, 30 s at

60°C, and 72°C for 60 s with the annealing temperature lowered by 1°C each cycle followed by

30 cycles of 30 s at 94°C, 30 s at 52°C, and 60 s at 72°C with a final extension of 72°C for 7

min.

Amplification of a 302 bp rrfA-rrlB IGS fragment for Borrelia spp. was carried out in a

25 µL reaction volume containing 1× AmpliTaq Gold 360 Master Mix, 0.4 µM each of primers

BoIGSA and BoIGSB and 5 µL DNA extract following Maggi et al. (2010). An initial

denaturation of 95°C for 10 minutes was followed by 55 cycles of 94°C for 15 s, 61°C for 15 s,

72 °C for 20 s and a final cycle of 72°C for 5 min.

A 596 bp fragment of Borrelia burgdorferi s.s. rrs-rrlA ITS was amplified using primers

BobuITS120s and BoBuITS720as developed by Maggi et al. (2010) following the PCR

conditions of the Borrelia spp. IGS using a 71°C annealing temperature. PCR products were

visualized on a 2% agarose gel stained with ethidium bromide.

Nanopore Sequencing to Determine Co-Infections

Next generation sequencing (NGS) was used to determine if ambiguous sequencing

results observed in sequencing chromatograms among some flaB Sanger sequences could be due

to co-infection with multiple B. burgdorferi s.l. genospecies. Sanger sequencing produces a

single sequence from a PCR sample whereas NGS produces millions of individual sequences

from a PCR sample and thus would better identify mixtures of B. burgdorferi s.l. genospecies.

Nanopore sequencing was conducted on a sample of eight tick DNA extracts and one B.

burgdorferi s.s. positive control template. Ticks selected for this analysis had previously produced flaB amplicons that were Sanger sequenced and found to be >98% similar to either B. burgdorferi s.s. B31 (NC_001318.1) or B. bissettii DN127 (NC_015921.1). Several of the chromatograms showed evidence of few or many single nucleotide polymorphisms as well as

chromatograms that appeared to be mixtures of the two B. burgdorferi s.l. genospecies.

MinION™ sequencing libraries were prepared from barcoded PCR amplicons of flaB, rrfA-rrlB, and rrs-rrlA and outer surface protein C (ospC).

A 665 bp fragment of flaB was amplified using the flaB Inner 1 and flaB 943R primers in a Mastercycler Gradient PCR thermal cycler in a 20 µL reaction volume containing 0.5 U

Platinum™ Taq DNA Polymerase (Thermo Fisher Scientific, Waltham, MA), 0.2 mM dNTPs,

1.5 mM MgCl2, 1x Buffer (IDT, Coralville, IA), 0.20 µM each primer and 2 µL DNA extract.

Touchdown PCR conditions included an initial denaturation step for 4 min at 94°C, followed by

10 cycles of 30 s at 94°C, 30 s at 60°C, and 72°C for 60 s with the annealing temperature lowered by 1°C each cycle followed by 30 cycles of 30 s at 94°C, 30 s at 51°C, and 60 s at 72°C with a final extension of 72°C for 7 min.

Amplification of the rrfA-rrlB IGS and rrs-rrlA ITS were carried out as previously described above in a Mastercycler Gradient PCR thermal cycler in a 25 µL reaction volume containing 0.5 U Platinum™ Taq DNA polymerase, 0.2 mM dNTPs, 1.5 mM MgCl2, 1x Buffer,

0.40 µM each primer and 3 µL DNA extract. Amplification of a 642 bp ospC fragment followed the methods of Di et al. (2018) using primers Oc-Fwd and Oc-Rev using the respective PCR thermal cycler and reaction mixture outlined above using 0.2 µM each primer and 2 µL DNA extract. Amplification conditions included an initial denaturation step for 4 min at 95°C, followed by 36 cycles of 30 s to denature at 95°C, 30 s anneal at 58°C, and extend at 72°C for 60 s with a final extension of 72°C for 5 min. All amplification products were verified by visualization on a 2% agarose gel and were Sanger sequenced to confirm the quality of the amplification products (Microgen, USA). The quantity of amplicons were each measured for quantity using an Agilent Tapestation 4200 (Agilent, Mississauga, ON, Canada). Two I.

scapularis ticks used in this analysis only produced an amplicon on the rrfA-rrlB primers in this nanopore sequencing analysis. Sequencing libraries were prepared using the one-dimensional ligation sequencing kit SQK-LSK109 (Oxford Nanopore Technologies, Oxford, UK).

Each set of amplicons from each of the eight ticks and positive control template (B. burgdorferi B31) were barcoded using the native barcoding expansion kit (EXP-NBD104) following the manufacturer’s protocol (Oxford Nanopore Technologies, Oxford, UK). Three

MinION™ flow cells were loaded with barcoded sequencing libraries from three ticks each consisting of 14 µL barcoded DNA, 25.5 µL loading beads and 37.5 µL sequencing buffer.

Sequencing was carried out over 48 h using the GridION™ nanopore sequencing platform

(Oxford Nanopore Technologies, Oxford, UK). Fast5 reads generated by nanopore sequencing were base called and demultiplexed using the high accuracy setting in Guppy program v3.2.2

(https://community.nanoporetech.com) followed by adapter trimming. FASTQ files were evaluated for quality using MultiQC and filtered for a 200-800 bp fragment size and PHRED scores above 20 (https://usegalaxy.org/). FASTQ files for each barcode were merged into single bulk FASTQ files and run individually through Nanopipe (http://bioinformatics.uni- muenster.de/tools/nanopipe) to align the FASTQ sequences against reference sequence targets and produce consensus sequences (Shabardina et al. 2019), which were then compared by

BLASTn in GenBank. Target sequences used in Nanopipe included B. burgdorferi s.s. B31

(CP019767) and B. bissettii DN127 (CP002746) chromosomes and the respective B. burgdorferi s.s. B31 and B. bissettii DN127 circular plasmid 26 (cp26) on which the ospC gene resides

(CP002747 and NC001903, respectively). Target sequences encompassing all B. burgdorferi s.l. genospecies found in North America were initially used in Nanopipe analyses, however they were excluded from the final analysis when no consensus sequences matching these targets were

produced. A phylogenetic tree was constructed using the partial ospC obtained from Sanger and

nanopore consensus sequences by the maximum likelihood method based on the Tamura-Nei

model (Tamura and Nei, 1993) using MEGAX (Kumar et al. 2018, Stecher et al. 2020).

Identification of Host DNA

Vertebrate Host Probe Development

Reverse Line Blot Hybridization probes were developed for vertebrate species that were potential tick hosts and confirmed or predicted to occur within the study areas (Supp. Table S1).

North Carolina Gap Analysis Project (NC-GAP) Geographic information system datasets were used to identify vertebrate species that are known or predicted to be distributed within the geographic area of each of the three study areas (McKerrow et al. 2006). Vertebrate probes were developed based on aligned sequences of North Carolina vertebrate species to complement 145 bp fragments of vertebrate 12S rDNA of Humair et al. (2007). Sequences of the 12S rDNA fragment for target species identified from NC-GAP were downloaded from GenBank, trimmed to the desired fragment size and aligned using ClustalX 2.1. Each probe was developed to distinguish hosts at the lowest possible taxonomic level. OligoAnalyzer 3.1 (Integrated DNA

Technologies, Inc) was used to ensure probes were designed to minimize folding of the PCR amplicon under the RLBH assay conditions and limit hybridization temperatures to within a few degrees of each other. Developed probes were validated against vertebrate tissues acquired from

North Carolina Museum of Natural Sciences, the Louisiana State University Museum of Natural

Science Collection of Genetic Resources and tissues specimens held by the Apperson Lab.

Northern bobwhite quail tissue was acquired from the North Carolina Wildlife Resources

Commission.

Tick Species Probe Development

In addition to vertebrate probes, two probes were developed based on a 441 bp fragment of tick mitochondrial 16S rDNA to distinguish I. affinis and I. scapularis (Table 1). Sequences of 16S rDNA from selected North American and European Ixodes species, including 12 of 15 species of

Ixodes ticks known to have a distribution east of the Mississippi river and having sequences available in the GenBank database were aligned using CLUSTALX. To confirm morphologic identifications, probes with a similar melting temperature to that of vertebrate probes and that would distinguish between I. scapularis and I. affinis molecularly were then developed.

Although the I. affinis probe shares 100% sequence similarity with a strain of I. minor used in the alignment, all life stages of I. affinis and I. minor are distinguishable morphologically. Tick

DNA was amplified from the 16S rDNA fragment using primers modified from Black and

Piesman (1994), 16S+1 and 16S – 1: Tick-16S-F and biotin labeled Tick-16S-R. Amplifications were run in a Labnet Multigene™ thermal cycler in 25 µL reaction mixture containing Amplitaq

Gold® 360 polymerase Master Mix, 0.40 µM each primer and 2 µL DNA extract. Amplification conditions included an initial denaturation step for 10 min at 95°C, followed by 40 cycles of 30 s each to denature at 95°C, anneal at 50°C, and extend at 72°C with a final extension of 72°C for 7 min. Tick amplification products were held at 4°C until use in RLBH assays. Ticks that were not clearly identified to species based on morphological characters and the RLBH assay were

Sanger sequenced using the 16S amplicon or a cytochrome c oxidase I (COI) gene fragment for unambiguous identifications. Amplification of the COI gene fragment using primers LCO1490 and HCO2198 (Folmer et al. 1994) were conducted in a Mastercycler Gradient PCR thermal cycler in 20 µL reaction mixture containing 0.5 U Platinum™ Taq DNA Polymerase, 0.2 mM dNTPs, 1.5 mM MgCl2, 1x Buffer, 0.40 µM each primer and 1 µL DNA extract. Amplification

conditions included an initial denaturation step for 3 min at 94°C, followed by 39 cycles to denature at 94°C for 30 s, anneal at 50°C for 50 s, and extend at 72°C for 60 s with a final extension of 72°C for 10 min. Sequences were compared to nucleotide sequences in GenBank using BLASTn to confirm tick species morphological identifications.

Reverse Line Blot Hybridization Assay

The RLBH assay was used to identify tick blood meals through amplification and hybridization of a 12S rDNA gene fragment with probes developed for potential vertebrate hosts.

Amplification of the 145 bp 12S rDNA gene fragment followed the nested PCR assay of Humair et al. (2007) using outer primers 12S-12F and 12S-13R and inner primers 12S-6F and biotin labeled B-12S-R. The biotin labeled reverse primer in our study was modified from Humair et al. (2007) with degenerate bases to amplify North American vertebrates. The outer fragment was amplified on a Labnet Multigene™ thermal cycler in a 25 µL reaction mixture containing

AmpliTaq Gold® 360 Master Mix, 0.80 µM each primer and 10 µL DNA extract. Touchdown

PCR conditions included an initial denaturation step for 10 min at 95°C, followed by 10 cycles of 30s at 94°C, 30s at 65°C, and 72°C for 60s with the annealing temperature lowered by 0.5°C each cycle followed by 25 cycles of 30 s at 94°C, 30 s at 60°C, and 60 s at 72°C with a final extension of 72°C for 10 min. The inner fragment was amplified as above using 1 µL of the outer PCR reaction as template. Touchdown PCR conditions for the inner fragment included an initial denaturation for 10 min at 95°C followed by 8 cycles of 30 s at 94°C, 30 s at 60°C, and 30 s at 72°C, lowering the annealing temperature by 1°C each cycle then 35 cycles using an annealing temperature of 52°C and a final extension of 72°C for 7 min. PCR products were held at 4°C up to 48 h before RLBH analyses.

The PCR-RLBH assay used followed the protocol described by Kong and Gilbert (2007).

Forty oligonucleotide probes (Invitrogen, Carlsbad, CA) including thirty-eight vertebrate probes and two tick probes were diluted in 0.5 M NaHCO3, pH 8.5 to achieve concentrations between

400-1400 pmol to compensate for differences in hybridization intensity among probes (Suppl.

Table S1). Probes were covalently bound to a Biodyne® C nylon membrane (Pall Corporation,

Ann Arbor, MI) by first activating the membrane in 16% (w/v) 1-ethyl-3-(3- dimethylaminopropyl) carbodiimide (EDAC) for 10 min. After a quick rinse with distilled water, the membrane was placed in a Miniblotter® 45 (Immunetics, Inc., Boston, MA) and 150

µL of each probe was added to individual lanes and allowed to incubate for 15 min. The membrane was then deactivated in 100 mM sodium hydroxide for up to 10 min, washed in 2x saline sodium phosphate-EDTA (SSPE) and washed for 5 min in 2x SSPE / 0.1% SDS at 60°C.

After a 20 min wash in 20 mM EDTA, the membrane was stored in 20 mM EDTA at 4°C. All chemicals used in probe binding were obtained from Sigma-Aldrich, St. Louis, MO, USA.

Hybridization of biotin-labeled PCR products was initiated with a pre-hybridization wash of the stored membrane in 2x SSPE / 0.1% SDS for 5 min at 60°C. To prevent non-specific hybridizations, the membrane was blocked in 0.1 % casein (Fisher Scientific, Pittsburg, PA) in

2x SSPE / 0.1 % SDS for 60 s and then washed once again in 2x SSPE / 0.1 % SDS for 5 min at

60°C. Higher concentration of casein or longer incubation times resulted in diminished signals from positive controls. Ten-µL of PCR products were diluted in 150 µL 2x SSPE/ 0.1% SDS.

42°C and then twice with 2x SSPE for 5 min at room temperature. The membrane was incubated for 2 min in electrochemiluminescence (ECL) detection liquid (GE Healthcare,

Otelfingen, Switzerland) and chemiluminescence was detected using a ChemiDoc-It® Imager

(UVP, Upland, CA, USA) after a 10 min exposure.

Statistical Analysis

Statistical analyses were carried out using SPSS, Version 27 (IBM, Armonk, NY). Chi-square tests of association were used to determine if 1) there was an association between B. burgdorferi s.l. detected by amplification of either flaB, rrfA-rrlB or rrs-rrlA gene fragments and tick demographics (species, sex, county), 2) between flaB sequenced B. burgdorferi s.l. genospecies prevalence among tick species and host blood meal identification, and 3) between bloodmeal host detected and tick demographics (species, sex, county). Results were considered to be statistically significant at P < 0.05. Bonferonni correction was applied in multiple comparisons to decrease the likelihood of type I error.

Results

Tick Collection

A total of 157 I. affinis (89 female, 68 male), 195 I. scapularis (91 female, 102 male, 2 nymph) and 2 I. brunneus nymphs were collected from three North Carolina counties. Adult I. affinis were collected from April through June while adult I. scapularis were collected from

November to May. The majority of I. affinis (75.8%; 119/157) were collected from Martin

county while 9.6% (15/157) were collected from Onslow county and 14.6% (23/157) were

collected in Wayne county. The majority of I. scapularis (76.4%; 149/195) were collected from

Onslow county while 10.3% (20/195) were collected from Martin county and 14.4% (28/195)

were collected from Wayne county. Two I. brunneus nymphs were collected in Onslow county

in the month of April. Two I. scapularis nymphs were collected; one from Onslow county in

April and one from Wayne county in June.

Molecular Identification of Tick Species

In addition to morphologic identifications, ticks utilized in RLBH assays and pathogen

analyses were molecularly identified by hybridization with RLBH probes specific for I. affinis or

I. scapularis. Of 182 ticks selected for pathogen and blood meal analysis, identifications of 25

ticks could not be confirmed by the molecular probes used in the RLBH assay due to failure of hybridization to either probe; therefore, 16S or COI gene markers amplified from the DNA of these ticks were Sanger sequenced. Molecular identification by either RLBH or Sanger sequencing revealed 13.7% (25/182) of ticks had been misidentified by morphology alone. Two nymphs that had been incorrectly identified as I. scapularis were > 99.4% similar to I. brunneus

(KX360364.1). One adult that had been incorrectly identified as I. scapularis was 99.7% similar to I. affinis (KX360422.1). Of the remaining 21 I. scapularis adults and one nymph that were initially identified as I. affinis, all but four adults and one nymph were identified using RLBH; these five ticks were sequenced for unambiguous identification. Two adults were > 99.4% similar to I. scapularis isolate Cor2-15 (KT831601), 2 adults were 100% similar to I. scapularis from Georgia (L43855 and L34294) and the nymph was 99.4% similar to I. scapularis voucher

FS52 (MN359328). Among I. scapularis morphological misidentifications as I. affinis, twice as

many males were misidentified as females.

Borrelia burgdorferi Prevalence

Borrelia burgdorferi s.l. DNA amplified, using all three molecular targets in both tick

species, with a higher detection prevalence among I. affinis than I. scapularis (Table 2). Of the three amplicons, B. burgdorferi s.l. was detected more frequently using the flaB amplicon than the rrfA-rrlB IGS or rrs-rrlA ITS. Statistical differences were observed in prevalence of B. burgdorferi s.l. between adult I. affinis and I. scapularis based on amplification of flaB (χ2 =

20.902, df = 1, P = 0.0000), rrfA-rrlB (χ2 = 80.03, df = 1, P = 0.000) and rrs-rrlA ITS sequence

(χ2 = 17.762, df = 1, P = 0.000). No difference was observed in the prevalence of B.

burgdorferi s.l. among females and males of either tick species regardless of the molecular

target.

Of the flaB amplicons that were submitted for Sanger sequencing, 17 of 111 amplicons

either failed to return a sequence or were of low sequence quality. Among I. affinis adults,

81.8% (45/55) of amplicons were 97.8 to 100% sequence identity matches to B. burgdorferi s.s.

strains (Accession numbers AB035616, AB035617, AB052665, AE000783, CP002228,

CP002228, CP002312, KF422803, KR782218, MF150054, KF422810, MK604312) and 18.2%

(10/55) were 97.9 to 99.4% sequence identity matches to a B. bissettii (CP002746) sequence in

GenBank (Figure 2). One I. affinis male returned a sequence that was only a 95.9% identity match to B. burgdorferi s.s. strain N40 (CP002228) and was excluded from flaB sequencing

statistical analysis. Among I. scapularis, 91.7% (33/36) of amplicons were 98.4 to 100% similar

to B. burgdorferi s.s. strains (AB035616, AB052665, AE000783, AY374137, CP002228,

CP002312, CP005925, CP017201, KM269447, KR782218, MF150054, MG967652) and 8.3%

(3/36) were 98.3 to 99.4% identity matches to B. bissettii (CP002746, MK684400, MG944964) sequences in GenBank. Among nymphs, one I. scapularis flaB sequence had 98.1% identity to

B. bissettii (CP002746) and one I. brunneus nymph flaB sequence had 99.6% identity to B.

burgdorferi s.s. (AE000783). There were no significant differences observed among tick

demographics and numbers of ticks with partial flaB gene sequences that clustered with either B.

burgdorferi s.s. or B. bissettii like-genospecies (species χ2 = 1.517, df = 1, P = 0.218; stage χ2 =

0.473, df = 1, P = 0.491; county χ2 = 1.010, df = 1, P = 0.604).

Nanopore Sequencing to Determine Co-Infections

Sanger sequencing was used to determine quality of PCR products used in the barcoded

nanopore sequencing analysis. Two I. scapularis (nos. M26 and M49) did not produce visible

amplicons on the 2% agarose gel electrophoresis for the rrs-rrlA or flaB fragments thus these

samples were not submitted for Sanger sequencing; these PCR products were nevertheless used

in the nanopore analysis. Sanger sequencing of the rrfA-rrlB target showed the majority of ticks

(7/8) were 96.2% to 100.0% similar to B. burgdorferi s.s. strains (Table 3). One I. affinis (no.

F92) was 93.9% similar to B. bissettii strain HLJ-233 (MG557646). Sanger sequencing of

amplicons of the rrs-rrlA reveal that the majority of ticks (6/6) showed 99.3% to 99.6%

similarity B. burgdorferi s.s. strains. Amplicons of flaB showed more variability in Sanger

sequences with four ticks 99.0 to 99.8% similar to B. burgdorferi s.s. strains and two I. affinis

(nos. F23 and F92) 98.9% and 99.0% similar to B. bissettii DN127 (CP002746.1), respectively.

Since B. burgdorferi ospC types may be found across B. burgdorferi s.l. genospecies, a

phylogenetic analysis of the ospC Sanger and nanopore consensus sequences was carried out to

determine similarity to ospC types deposited in GenBank (Figure 3). Three I. affinis sequences

grouped with ospC type L (EU375832), two I. affinis sequences grouped with type Fa

(AY275225), one I. scapularis grouped with type E3 (EF592545) and the B. burgdorferi B31 positive template grouped with type Ba (EF537413).

Nanopore data presented in Table 4 show that reads mapped to both B. burgdorferi s.s.

B31 and B. bissettii DN127 target sequences. Among all barcoded tick samples, the majority

(5/8) showed a high proportion of reads (92.5% to 99.3%) mapping to B. burgdorferi s.s. B31.

Ixodes affinis F23 and F92 mapped 58.7% and 54.8%, respectively, of their total mapped reads to B. burgdorferi s.s. B31 while I. affinis F22 mapped 82.3% of total reads to B. burgdorferi s.s.

B31.

Consensus sequences produced in Nanopipe from nanopore sequences of flaB amplicons

were 98.5% to 99.9% similar to multiple B burgdorferi s.s. strains including B31 (CP019767.1),

JD1 (CP002312.1) and B331 (CP017201.1) and 98.1% to 99.1% similar to B. bissettii DN127

(CP002746.1). The positive control (B. burgdorferi B31) produced a flaB consensus sequence

that was 99.0% similar to B. burgdorferi s.s. strain B331 (CP017201.1) and did not produce any

consensus sequence against the B. bissettii target. Consensus sequences produced from rrfA-rrlB

amplicons were 96.0% to 98.2% similar to B. burgdorferi s.s. strains 297 (JX564636.1), JD1

(CP002312.1), B31 NRZ (CP019767.1) and only 93.4% to 96.3% similar to B. bissettii DN127

(CP002746.1) and B. bissettii isolate HLJ-233 (MG557646.1). The positive control produced

rrfA-rrlB consensus sequences against both B. burgdorferi B31 and B. bissettii targets that were

97.6% similar to B. burgdorferi s.s. B31 (CP019767.1) and 96.4% similar to B. bissettii isolate

HLJ-233 (MG557646.1) respectively. Consensus sequences produced from rrs-rrlA amplicons

were only produced against the B. burgdorferi s.s. B31 target and were 97.4% to 99.7% similar

to B. burgdorferi s.s. strains HWT37 (JQ356546.1), JD1 (CP002312.1), m11p (KM269456.1),

and BTW16 (JQ308239.1). The positive control produced a rrs-rrlA consensus sequence that

was 99.4% similar to B. burgdorferi s.s. strain M11p (KM269456.1).

Based on the results of Sanger sequencing of flaB and nanopore read mapping, I. affinis

F23 and F92 appear to be co-infected with B. burgdorferi s.s. and B. bissettii. A proportion of reads that matched to B. bissettii in the B. burgdorferi B31 control may be due to a small number of polymorphic sites within B. burgdorferi rrfA-rrlB gene, however this positive template DNA

was derived from an I. minor tick and thus possible it could be co-infected as well. It is not clear

if the remaining ticks were co-infected since they had matches to both B. burgdorferi s.s. B31

and B. bissettii DN127 target sequences and polymorphic sites are abundant in many of the

consensus sequences.

Host Blood meal Determinations

Based on vertebrate 12S rDNA amplicons that could be visualized by 2% agarose gel

electrophoresis, amplifications of tick blood meal DNA was successful for 33.3% (58/174) of

tick extracts. Despite a lack of visual gel bands for 116 amplification products, all 174 vertebrate

12S rDNA amplification products were run in RLBH assays. Amplicons that failed to produce a

gel band likewise did not hybridize to any RLBH probe. Of the visualized amplified blood

meals, 45.5% (10/22) I. affinis and 47.2% (17/36) I. scapularis could be identified in RLBH

assays. Of those that could not be identified by RLBH, 30 blood meals could not be resolved to

host species due to non-specific hybridizations. One amplified blood meal that failed to

hybridize to any vertebrate probe was Sanger sequenced. This 12S rDNA amplicon from an I.

scapularis nymph collected from Martin county in May 2011 showed 98% sequence similarity to a pseudogene of Canis lupus familiaris (AB048587).

Two I. affinis females showed evidence of multiple blood meals, hybridizing to cotton rat

(Sigmodon hispidus Say and Ord) and gray fox (Urocyon cinereoargenteus Schreber) and to cotton rat and eastern chipmunk (Tamias striatus Linnaeus) probes in RLBH assays (Table 5).

The prevalent blood meal detected in I. affinis was cotton rat (40.0%; 4/10) followed by Canis sp. (20.0%; 2/10). Likewise, cotton rat (33.3%; 6/18) and Canis spp. (27.8%; 5/18) were also the most prevalent blood meals detected among I. scapularis. Three potentially new vertebrate hosts were identified for I. affinis, including little brown skink (Scincella lateralis Say), eastern chipmunk (Tamias striatus Linnaeus) and gray fox (Urocyon cinereoargenteus Schreber).

Host – Borrelia burgdorferi Associations

Among ticks with identified blood meals, 44.4% (12/27) were infected with B. burgdorferi s.l. based on the amplification of the flaB, rrfA-rrlB or rrs-rrlA gene targets.

Borrelia burgdorferi s.l. prevalence did not differ significantly between tick extracts that amplified vertebrate 12S rDNA and those that did not (χ2 = 0.007, df = 1, P = 0.934. Nor was there a significant difference between B. burgdorferi s.l. prevalence and vertebrate blood meal host among I. affinis or I. scapularis (χ2 = 2.085, df = 7, P = 0.955).

Discussion

Borrelia burgdorferi Prevalence

We found B. burgdorferi s.l. prevalence, based on amplification of rrfA-rrlB intergenic spacer sequence primers used by Maggi et al. (2010), to be higher overall than previously

reported for I. affinis and I. scapularis collected in 2008 and 2009 from four North Carolina

Coastal Plain counties. Compared to Maggi et al. (2010), who reported 63.2% I. affinis and

0.0% of I. scapularis infected with B. burgdorferi s.l., we found 77.8% of I. affinis and 9.1% I. scapularis to be infected with B. burgdorferi s.l. Notably, B. burgdorferi s.l. infection rates were substantially higher for the flaB gene, with 80.0% of I. affinis and 46.8% of I. scapularis infected with B. burgdorferi s.l. The prevalence of infection among I. scapularis is comparable to that reported in Smith et al. (2010) who amplified a B. burgdorferi s.l. flaB gene fragment in 40%

(6/15) I. scapularis nymphs collected from Chatham county, North Carolina.

One explanation for the higher prevalence of flaB gene amplicons is that it is a more sensitive and specific gene target when compared to other commonly utilized molecular markers, including rrs and rrs-rrlA, to detect Borrelia spp. (Wodecka et al. 2010). There have been reports of apparent non-specific amplification using the flaB gene (Mays et al. 2014) and in this study 15.3% of flaB amplicons submitted for Sanger sequencing did not return readable sequences. Nevertheless, sequencing results for the remaining flaB amplicons confirmed that this target was a more sensitive molecular target than the rrfA-rrlB and rrs-rrlA genes in detecting B. burgdorferi s.l. infections and may explain the higher infection prevalence for ticks in these North Carolina Coastal Plain counties than previously reported.

The sensitivity of a PCR amplification can be influenced by the DNA extraction method

(Mauel et al. 1999, Pruvot et al. 2013). In our study, phenol-chloroform extraction was utilized and has been proven to produce high quality DNA facilitating its amplification by PCR. Our extraction methods differed from Maggi et al. (2010) who used the Qiagen DNeasy tissue kit.

We conducted a side-by-side comparison of both extraction methods in a sample of ticks cut

lengthwise; no differences in terms of DNA quality and quantity or PCR amplification were

noted.

Accurate identification of ticks with varying roles in pathogen transmission is crucial.

Incorrect identifications can misrepresent overall infection prevalence of pathogens in important

bridge vectors that alter perceived disease risk to the public. The misidentifications of

morphologically similar ticks reported in this study highlight the need for molecular

confirmation in order to report accurate pathogen prevalence. Nadolny et al. (2011) found

sympatric I. affinis and I. scapularis in the Coastal Plain of southeastern Virginia with both

species active throughout the year with the exception of August. Since I. affinis is a relatively

newly detected species in North Carolina and Virginia, studies of Borrelia spp. infection

prevalence among these sympatric ticks should include both morphologic identification followed

by molecular confirmation.

Co-infections of pathogens in ticks are difficult to detect and identify using conventional

PCR and Sanger sequencing. Based on preliminary analysis of nanopore sequencing of multiple

loci in this study, it appears that at least two I. affinis are co-infected with B. burgdorferi s.s. and

B. bissettii based on the proportions of reads mapped to either target and to Sanger sequence

results that showed similarity to B. bissettii and B. burgdorferi s.s. across multiple loci. This

finding is consistent with Maggi et al. (2010) that reported coinfection among 4% of I. affinis from Coastal Plain counties. Further analysis using the EPI2ME bioinformatics platform and

What’s In My Pot (WIMP) workflow (Juul et al. 2015) will allow species level classification and quantification of reads in order to fully examine the extent of co-infection among these ticks. As with any next generation sequencing identification strategy, defining appropriate thresholds using known samples is critical for determining positivity versus sequencing error in the

technology or background variation due to natural polymorphism which is not captured in

available database sequences deposited in GenBank.

The B. burgdorferi ospC gene is expressed during tick feeding and facilitates spirochete

infection of vertebrate hosts (Grimm et al. 2004) thus playing an important role in the process of

infection with some ospC types being more invasive than others. Interestingly, three out of eight

of the ospC sequences grouped phylogenetically with ospC type L, which is rare in areas of highly endemic LD in the United States but has been reported to be widely distributed in the southeastern United States. OspC type L has been associated with I. affinis and has invasive potential comparable to ospC type B, which is associated with severe LD (Golovchenko et al.

2014). Other ospC types observed in our study among I. affinis and I. scapularis include type Fa which has been previously reported from the northern United States and E3 previously reported as associated with I. pacificus in the north-central and western United States, respectively

(Travinsky et al. 2010, Rudenko et al. 2013).

Host – Borrelia burgdorferi Associations

Only a third of tick extracts amplified the 12S rDNA fragment and half of those yielded an identifiable blood meal host in RLBH assays. The age of the blood meal is an important factor in amplification success. In laboratory studies Kirstein and Gray (1996) found only 40% of I. ricinus larval blood meals amplified a cytochrome b gene fragment after 9 months post- engorgement. Given field conditions would likely be harsher with respect to blood meal degradation than laboratory conditions, it is likely that age related degradation occurred at increased rates, limiting amplification success in our study. To amplify vertebrate species from

North America primers used in this study were slightly altered from Humair et al. (2007), a study

which was successful in identifying blood meal hosts in half of host-seeking I. ricinus ticks from

Switzerland with the majority identified to genus or species level. Using the same 12S rDNA

gene target in an RLBH assay to identify hosts of nymphal I. ricinus in north-central Spain,

Estrada-Pena et al. (2005) reported similar difficulties in amplifying a product with 40.1%

yielding “unambiguous” identifications to order and genus level. The authors felt that blood

meal degradation explained the lower than expected ability to amplify and identify blood meals.

Using RLBH and similar modifications to the primers of Humair et al. (2007) to amplify North

American species, Scott et al. (2012) were able to successfully identify the hosts for 53.1% of blood meals in host-seeking adult I. scapularis but reported many of the challenges faced in using RLBH, including cross-reacting probes, non-specific binding to probes causing false positive signals, and the possibility that extraneous PCR products amplified from tick DNA.

Added to these challenges is the well-known and unavoidable contamination of tick extracts with human DNA. It was for these reasons that a very conservative approach in interpreting hybridizations in RLBH assays was undertaken in our study and likely contributed to a reduction in our overall success in identifying host blood meals among Ixodes spp. ticks.

Although useful in differentiating blood meals taken from multiple hosts, the success of the RLBH in identifying the vertebrate host blood meals was confounded by non-specific and ambiguous hybridizations that prevented the identification of the blood meal hosts for some ticks. Despite these challenges in identifying tick blood meals, our present study results do not vary in success from those reported previously by other researchers using the RLBH assay, including non-specific amplification, cross- and non-specific hybridizations (Goessling et al.

2012, Scott et al. 2012). Our study was likely too conservative in assessing the remnant blood meals from the nymphal and larval feeding since it excluded ambiguous hybridizations. Future

studies of remnant blood meals of ticks may benefit from the ever evolving next generation

sequencing technologies that have workflows that do not require PCR amplification and thus

eliminate potential non-specific amplification.

Molecular blood meals in host-seeking adult ticks represents the remnants of the nymphal

blood meals. Given that the nymphal stage of I. scapularis is recognized as the most important

stage in the transmission cycle of LD, these blood meal determinations have the greatest

potential for elucidating important maintenance and competent reservoir hosts. In our study,

RLBH analyses of the remnant blood meals from the nymphal feeding showed that the majority

of blood meals identified in adults of both I. affinis and I. scapularis were cotton rat (Sigmodon

hispidus Say and Ord) and Canis spp.

Although results of our study revealed showed no significant association between tick blood meal host and pathogen infection, cotton rats are likely an important competent reservoir for B. burgdorferi s.l. genospecies in the south as they are known to develop lasting spirochetemias when experimentally infected (Burgdorferi and Gage, 1987). Cotton rats, along with cotton mice (Peromyscus gossypinus Le Conte), and eastern woodrat (Neotoma floridana

Ord), are known as important hosts of I. affinis in sandhill and coastal areas of South Carolina

(Clark et al. 1998). In an intensive trapping study in North Carolina, Apperson et al. (1993) did

not find any I. scapularis attached to cotton rat; however, four rodent species were fed upon by I.

scapularis including white-footed mouse, cotton mouse, golden mouse (Ochrotomys nuttalli

Harlan) and house mouse (Mus musculus Linnaeus). In addition to the cotton rat probe, there

were two detections of I. scapularis feeding on small rodents, having hybridized to the

Cricetidae probe which is designed to detect, but not differentiate, between the marsh rice rat

(Oryzomys palustris Harlan), eastern harvest mouse (Reithrodontomys humulis Audubon and

Bachman), golden mouse, deer mouse (Peromyscus maniculatus Gloger), several species of voles (Microtus chrotorrhinus Miller, M. pennsylvanicus Ord and M. pinetorum Le Conte,

Myodes gapperi Vigors), muskrat (Ondatra zibethicus Linnaeus) and the southern bog lemming

(Synaptomys cooperi Baird).

The Canis spp. probe used in this study is designed to detect DNA from domestic or feral dog (Canis familiaris Linnaeus), coyote (C. latrans Say) and red wolf (C. lupus Linnaeus). The only wild red wolf population in the United States is in northeastern North Carolina, however, with very low population numbers it is unlikely that the detections in this study were from red wolf. Serological tests have previously detected Lyme borrelia antibodies in both red wolves and coyotes in North Carolina (Brzeski et al. 2015) and among both domestic and stray dogs from North Carolina (Greene et al. 1986). Three of the tick samples with Canis sp. blood meal detections were positive for B. burgdorferi s.s. based on flaB sequencing which indicates that although canines in North Carolina may have a low seroprevalence of B. burgdorferi B31 antigen (Duncan et al. 2004), they are likely important maintenance hosts and due to their large home ranges may be important in the movement of B. burgdorferi s.l. infected ticks.

Some of the earliest descriptions of I. scapularis from the southern United States describe lizards as important blood meal hosts (Rogers, 1953; Spielman et al, 1984). Of all hosts identified, only three tick blood meals were identified from lizards while the majority of nymphal blood meals identified were taken from rodents or canines. While Apperson et al.

(1993) reported I. scapularis to parasitize lizards in summer months in Onslow County, North

Carolina far fewer tick-infested rodents were collected during the study period, suggesting that immature I. scapularis were diverted from Peromyscus sp. mice by skinks that made up the majority of tick-infested lizard species. Ginsberg et al. (2021) conducted trapping in North

Carolina as part of a larger study looking at Ixodes host preference from north to south in the eastern United States and found I. scapularis larvae and nymphs infesting mice, medium sized mammals and skinks. In our study, the RLBH resulted in some ambiguous binding patterns that could not be resolved for some tick blood meals and thus excluded some host identifications which may have included ground dwelling birds and some lizards, making our host identifications reported here conservative. Molecular methods used in blood meal analyses continue to be useful in uncovering host-pathogen relationships that avoid biases derived from trapping of vertebrates. Third generation sequencing methods such as nanopore sequencing or qPCR-based bloodmeal analyses (Kim et al. 2021) may improve our ability to distinguish multiple blood meals.

Conclusion

The aim of our study was to determine if an association between vertebrate host and B. burgdorferi s.l. infection prevalence could be identified among sympatric I. affinis, an enzootic vector of the LD pathogen, and I. scapularis, the bridge vector capable of infecting humans.

While no host-pathogen associations could be made due to the limited number of blood meals amplified and identified in RLBH assays, cotton rat and Canis spp. appear to be important hosts of both tick species and are likely important in the transmission in the Coastal Plain of North

Carolina. Our study found a higher prevalence of B. burgdorferi s.s. among both I. affinis and I. scapularis than has previously been reported in North Carolina, which may be the result of a combination of accurate molecular identification of tick species and a more sensitive flaB molecular target. Several of the ospC sequences among I. affinis were phylogenetically similar to ospC type L, a type that is rare but commonly associated with I. affinis and has invasive

potential. Where I. affinis and I. scapularis are sympatric and utilize the same vertebrate host species, these more invasive strains may be a cause of increasing cases of LD. Molecular techniques remain one of the most promising tools to fully characterize the infection prevalence and blood meal hosts of I. affinis and I. scapularis and avoid bias introduced through trapping methods and timing of field captures of animal hosts. Future tick blood meal analyses would benefit from NGS techniques that have evolved to be both less costly and time consuming than other blood meal identification methods.

100

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112

Figure 1. Location of study sites in the Coastal Plain region of North Carolina from north to south are Martin, Wayne and Onslow counties.

113

Figure 2. Molecular phylogenetic analysis of Sanger sequencing of a 601 bp fragment of the B. burgdorferi s.l. flaB gene among Ixodes affinis () and Ixodes scapularis (). The evolutionary relationships were inferred using the Maximum Likelihood method and Tamura-Nei model. The

tree with the highest log likelihood (-1906.16) is shown. Initial tree(s) for the heuristic search

were obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix of

pairwise distances estimated using the Tamura-Nei model and then selecting the topology with

superior log likelihood value. This analysis involved 36 tick and eight reference nucleotide

sequences. There were a total of 604 positions in the final dataset.

114

115

Figure 3. Molecular phylogenetic analysis of Sanger and nanopore consensus sequences of B.

burgdorferi ospC amplicons among positive template DNA (), Ixodes affinis () and Ixodes

scapularis (). Evolutionary analysis by Maximum Likelihood method. The evolutionary

relationships were inferred by using the Maximum Likelihood method and Tamura-Nei model.

The tree with the highest log likelihood (-9287.57) is shown. Initial tree(s) for the heuristic search were obtained automatically by applying Neighbor-Join and BioNJ algorithms to a matrix

of pairwise distances estimated using the Tamura-Nei model, and then selecting the topology with superior log likelihood value. The tree is drawn to scale, with branch lengths measured in

the number of substitutions per site. This analysis involved 14 nanopore and six Sanger

sequences from six tick extracts and one B. burgdorferi B31 template DNA and 31 ospC reference nucleotide sequences. Codon positions included were 1st+2nd+3rd+Noncoding. There

were a total of 555 positions in the final dataset. Evolutionary analyses were conducted in

MEGA X.

116

117

Table 1. Oligonucleotide primers and probes used in this study to detect Borrelia burgdorferi sensu lato, amplify vertebrate blood meals or identify tick species.

Target Primer / 5’-3’ Sequence Amplicon Reference

Probe Size (bp)

Name

flaB Outer 1 AARGAATTGGCAGTTCAATC 497 Clark et al. (2005)

Outer 2 GCATTTTCWATTTTAGCAAGTGATG

Inner 1 ACATATTCAGATGCAGACAGAGGTTCTA 389

Inner 2 GAAGGTGCTGTAGCAGGTGCTGGCTGT

flaB 103F AGAATTAATCGAGCTTCTGATGATGC 840 This study

flaB 943R GCAATCATTGCCATTGCAGATTG

rrfA-rrlB (5S- BoIGSA CGACCTTCTTCGCCTTAAAGC 302 Maggi et al. (2010)

23S) GTTAAGCTCTTATTCGCTGATGGTA BoIGSB

118

Table 1 (continued).

rrs-rrlA (16S- Bobu AGGTCATTTTGGGGGTTTAGCTCAGTTGGCT 596 Maggi et al. (2010)

23S) ITS120s

of B. Bobu AGTGTCGGGCAAATCCAAAACTGAAAATCTG

burgdorferi s.s. ITS720as

ospC Oc-Fwd AATAAAAAGGAGGCACAAATTAATG 642 Di et al. (2018)

Oc-Rev ATATTGACTTTATTTTTCCAGTTAC

COI LCO1490 GGTCAACAAATCATAAAGATATTGG 648 Folmer et al. (1994)

HCO2198 TAAACTTCAGGGTGACCAAAAAATCA

Vertebrate 12S 12S-12F TGCCAGCCACCGCGGTCA 600 Humair et al. (2007)

rDNA 12S-13R AGGAGGGTGACGGGCGGT

12S-6F CAAACTGGGATTAGATAC 145 Humair et al. (2007)

B-12S-R* ACAGGCTCCTCTARR This study

119

Table 1 (continued).

Tick 16S Tick-16S-F TAAATTGCTGTGGTATTTTGA Black and Piesman (1994)

Tick-16S-R GGTCTGAACTCAGATCA

Tick 16S I. affinis GGTGATGGAAAAAGAATAAATATCTT This study

probes I. GGCGATAAAAAAAGAATAAGTATCTTT This Study

scapularis

* Biotin labeled probe.

120

Table 2. Prevalence of Borrelia burgdorferi s.l. among I. affinis and I. scapularis based on amplification of fragments of flaB, rrfA-rrlB, and Borrelia burgdorferi s.s. specific rrs-rrlA.

Sample sizes of ticks analyzed are given in parentheses.

flaB rrfA-rrlB rrs-rrlA

I. affinis

Female 80.0% (45) 75.6% (45) 37.8% (45)

Male 80.0% (40) 80.6% (36) 22.2% (36)

80.0% (85) 77.8% (81) 30.9% (81)

I. scapularis

Female 45.5% (44) 7.7% (39) 5.1% (39)

Male 47.9% (48) 10.6% (47) 6.3% (47)

Nymph 50.0% (2) 0.0% (2) 0.0% (2)

46.8% (94) 9.1% (88) 5.7% (88)

121

Sample rrfA-rrlB rrs-rrlA flaB

I. affinis F22 96.2% B. burgdorferi JD1 99.3% B. burgdorferi M11p 99.5% B. burgdorferi B31

(CP002312) (KM269456.1) (AE000783.1)

I. affinis F23 97.8% B. burgdorferi 426905 99.5% B burgdorferi HWT37 98.9% B. bissettii DN127

(GQ463611) (JQ356546.1) (CP002746.1)

I. affinis F25 99.3% B. burgdorferi JD1 99.5% B. burgdorferi JD1 99.0% B. burgdorferi B31

(CP002312) (CP002312.1) (AE000783.1)

I. affinis F92 93.9% B. bissettii HLJ-233 99.6% B. burgdorferi JD1 99.0% B. bissettii DN127

(MG557646) (CP002312.1) (CP002746.1)

I. affinis M29 100.0% B. burgdorferi MM1 99.5% B. burgdorferi Y14 99.8% B. burgdorferi JD1

(CP031412.1) (KC416425.1) (CP002312.1)

122

Table 3 (continued).

I. scapularis M26 99.7% B. burgdorferi B31 . .

(AE000783.1)

I. scapularis M46 100.0% B. burgdorferi MM1 99.5% B. burgdorferi Y14 99.8% B. burgdorferi JD1

(CP031412.1) (KC416425.1) (CP002312.1)

I. scapularis M49 100.0% B. burgdorferi JD1 . .

(CP002312.1)

B. burgdorferi s.s. 100.0% B. burgdorferi B31 99.5% B. burgdorferi M11p 99.7% B. burgdorferi B31 (KM269456.1) (AE000783.1) Positive Template (AE000783.1)

123

Table 4. Nanopore sequencing of B. burgdorferi s.l. chromosomal gene fragments flaB, rrfA-rrlB, and rrs-rrlA among I. affinis, I. scapularis and B. burgdorferi s.s. positive template DNA.

Sample Number Percent Percent Consensus Sequence GenBank BLASTn matches

of Reads Reads Reads (B. burgdorferi B31 / B. bissettii DN127) passing Mapped to Mapped rrfA-rrlB rrs-rrlA flaB QC B. to B.

burgdorfe bissettii

ri s.s. B31 DN127

(%) (%)

I. affinis F22 558,902 82.3 20.0 98.1% B. burgdorferi 98.7% B 99.9% B. burgdorferi 297 (JX564636.1) / burgdorferi B31 (CP0197667.1) /

95.0% B. bissettii HWT37 99.1% B. bissettii

DN127 (CP002746.1) (JQ356546.1) DN127 (CP002746.1)

124

Table 4 (continued).

I. affinis F23 444,387 58.7 29.1 98.2% B. burgdorferi 97.4% B. 99.0% B. burgdorferi JD1 (CP002312.1) / burgdorferi JD1 B31 (CP0197667.1) /

95.3% B. bissettii (CP002312.1) 98.2% B. bissettii

DN127 (CP002746.1) DN127 (CP002746.1)

I. affinis F25 414,840 94.2 4.3 97.7% B. burgdorferi 99.5% B. 99.6% B. burgdorferi JD1 (CP002312.1) / burgdorferi JD1 B331 (CP017201.1) /

95.9% B. bissettii (CP002312.1) 99.1% B. bissettii

HLJ-233 DN127 (CP002746.1)

(MG557646.1)

I. affinis F92 2,955,445 54.8 38.5 96.0% B. burgdorferi 99.7% B. 98.5% B. burgdorferi B31_NRZ burgdorferi M11p B331 (CP017201.1) /

(CP019767.1) / 93.4% (KM269456.1) 98.1% B. bissettii

B. bissettii DN127 DN127 (CP002746.1)

(CP002746.1)

125

Table 4 (continued).

I. affinis M29 406,402 92.8 5.7 96.3% B. burgdorferi 99.2% B. 99.3% B. burgdorferi JD1 (CP002312.1) / burgdorferi JD1 (CP002312.1) /

95.6% B. bissettii BTW16 98.7% B. bissettii

HLJ-233 (JQ308239.1) DN127 (CP002746.1)

(MG557646.1)

I. scapularis M26 1,115,383 92.5 0.9 97.9% B. burgdorferi 97.5% B. 98.5% B. burgdorferi JD1 (CP002312.1) / burgdorferi JD1 (CP002312.1) /

95.2% B. bissettii BTW16 99.1% B. bissettii

DN127 (CP002746.1) (JQ308239.1) DN127 (CP002746.1)

I. scapularis M46 1,204,267 95.2 0.5 97.1% B. burgdorferi 99.4% B. 99.4% B. burgdorferi JD1 (CP002312.1) / burgdorferi JD1 JD1 (CP002312.1) /

95.6% B. bissettii (CP002312.1) 98.7% B. bissettii

DN127 (CP002746.1) DN127 (CP002746.1)

126

Table 4 (continued).

I. scapularis M49 455,066 99.3 0.1 97.2% B. burgdorferi 98.7% B. 99.9% B. burgdorferi B31_NRZ burgdorferi JD1 (CP002312.1) /

(CP019767.1) / 95.5% BTW16 99.0% B. bissettii

B. bissettii DN127 (JQ308239.1) DN127 (CP002746.1)

(CP002746.1)

B. burgdorferi s.s. 354,714 98.5 2.5 93.5% B. burgdorferi 99.4% B. 99.0% B. burgdorferi JD1 (CP002312.1) / burgdorferi M11p B331 (CP017201.1) Positive Template 96.0% B. bissettii (KM269456.1)

HLJ-233

(MG557646.1)

127

Table 5. Host blood meals identified and B. burgdorferi s.l. genospecies infecting host-seeking I. affinis and I. scapularis.

*Represents identifications of two host blood meals from individual female I. affinis ticks

Species, Stage No 12S Not Eastern Little Rabbit Eastern Wood- Small Cotton Canis Gray Total

and B. amplicon identified fence brown chip- chuck rodent Rat sp. Fox Ticks burgdorferi s.l. lizard skink munk

infection

I. affinis

Female

Not infected 9 1 0 0 0 0 0 0 1* 0 1* 11

B bissettii 4 1 0 0 0 0 0 0 0 0 0 5

B burgdorferi 21 4 0 1 0 1* 0 0 2* 0 1 29

Male

Not detected 12 1 0 1 0 0 0 0 1 1 0 16

B bissettii 3 2 0 0 0 0 0 0 0 0 0 5

B burgdorferi 12 3 0 0 1 0 0 0 0 1 0 17

Total 61 12 0 2 1 1 0 0 4 2 2 83

128

Table 5 (continued).

I. scapularis

Female

Not detected 13 6 1 0 1 0 0 1 0 1 0 23

B bissettii 1 0 0 0 0 0 0 0 0 0 0 1

B burgdorferi 9 5 0 0 0 0 1 0 0 1 0 16

Male

Not detected 18 4 0 0 1 0 0 1 3 1 0 28

B bissettii 2 0 0 0 0 0 0 0 0 0 0 2

B burgdorferi 10 4 0 0 0 0 0 0 2 1 0 17

Nymph

Not detected 0 0 0 0 0 0 0 0 1 0 0 1

B bissettii 0 0 0 0 0 0 0 0 0 1 0 1

Total 62 19 1 0 2 0 1 2 6 5 0 89

129

APPENDIX

130

Appendix A

Probe Name Probe Sequences (5’-3’) Probe Conc. Vertebrate species

(pmol)

Avesac TACGAGCACAAACGCTTAA 450 Meleagris gallopavo, Colinus virginianus,

Coturnix japonica, Zenaida macroura,

Strix varia, Thryothorus ludovicianus,

Junco hyemalis

Turkeyc ATCTTGATACTAATATACTCACGTATCC 450 Meleagris gallopavo

Quail TCCAGATACCCCCAATACCAAC 450 Colinus virginianus, Coturnix japonica*

Lacertillaa GAGAACTACAAGTGAAAAACT 1200 Ophisaurus attenuatus, O. ventralis,

Plestiodon fasciatus, P. laticeps, Scincella

lateralis†

Glass Lizard GCTTAGTCCTAAYCTCAGATATTAAC 1050 Ophisaurus attenuatus, O. ventralis

131

Table S1 (continued).

Anole ACTAAAGTGTTCGCCAGAATATTACG 750 Anolis carolinensis

Fence Lizard CAGAAAACTACGAGCGAAAAGCTTA 750 Sceloporus undulatus

Plestiodon sp. TGAAAAACTTAAAACTCCAAGGACTT 1200 Plestiodon fasciatus, P. laticeps

Skinkc

Ground Skinkc AACAACGATCTTATCTYACAATATTATCC 750 Scincella lateralis

Racerunner TAACAATTTGTCCGCCAGAAAATT 1200 Cnemidophorus sexlineatus

Mammaliabc AAACTCAAAGGACTTGGC 1200 All mammal tissues listed below, Homo

sapiens, Lasiurus borealis, Eptesicus

fuscus

Opossum GCTTAGTAATAAACTAAAATAATTTAACAAACA 1200 Didelphis virginiana

Long-tailed CTAACAAAAATACCCGCCAGAGAA 900 Sorex cinereus, S. dispar, S. palustris, S.

Shrewd longirostris, S. hoyi

Short-tailed & AATTAACAAAACTACTCGCCAGAGGA 900 Blarina brevicauda, B. carolinensis,

Least Shrewcd Cryptotis parva

Mole ACTAAGACAATCCAACTAACAAGATT 900 Scalopus aquaticus, Condylura cristata

132

Table S1 (continued).

Rabbitd ACTTAAATAATTCCATAACAAAATTACTCG 900 Sylvilagus palustris, S. floridanus, S.

obscurus

Chipmunk CTTAAACACAAATACTTAATAAACAAGAGTATT 1200 Tamias striatus

Woodchuckc AGAGTACTACTAGCAATAGCCTGAA 600 Marmota monax

Squirrelcd AGAGAACTACTAGCCACTGCTTA 900 Sciurus carolinensis, S. niger

Flying Squirrelc GCTTAGCCCTAAACACAAATATTTAA 900 Glaucomys volans

Cricetidaec ATTTGCCYGAGAACTACTGGC 1200 Oryzomys palustris, Reithrodontomys

humulis, Ochrotomys nuttalli, Peromyscus

gossypinus, P. leucopus, P. maniculatus,

Microtus chrotorrhinus, M.

pennsylvanicus, M. pinetorum, Myodes

gapperi, Ondatra zibethicus, Synaptomys

cooperi

Peromyscus mice CTAAACCTYAAAGATTAAATAACAAAATCAT 1050 Peromyscus gossypinus, P. leucopus, P.

maniculatus

133

Table S1 (continued).

Cotton Rat CTAAACCACAATAACTTAAAAACAAAGTT 900 Sigmodon hispidus

Woodrat CTAAACCCTAATAATTCAATAACAAAAT 1200 Neotoma floridana

Ratcd GCTTAGCCCTAAACCTTAATAATTA 750 Rattus norvegicus

House Mouse CCATAAACCTAAATAATTAAATTTAACAAAACT 750 Mus musculus

ATTT

Canis sp. AACATAGATAATTTTACAACAAAATAATTCG 900 Canis latrans, Canis lupus familiaris

Red Fox CATAAATAGTTCTATAACAAAACAATTCG 1200 Vulpes vulpes

Gray Fox ACATAAACAGTTCTATAAACAAAATAGTT 1200 Urocyon cinereoargenteus

Bear AGCCTTAAACATAAGTAATTTATTAAACAAA 1200 Ursus americanus

Raccoonc ATTAACGTAACAAAATTATTTGCCA 1050 Procyon lotor

Minkd ATTCACATAACAAAATTACTTGCCA 1050 Neovison vison

Weaselc ATTTACATAACAAAATTATTTGCCA 1050 Mustela frenata, M. nivalis

Skunk CCATAAACACAGACAATTAATATAACAAA 1350 Mephitis mephitis

Felidc AAACAAAACTATCCGCCAGAGAA 900 Lynx rufus

Swinecd TAAACCCAAATAGTTACATAACAAAACTAT 600 Sus scrofa

134

Table S1 (continued).

White-tailed AACATAAATAGTTATATAAACAAAACTATTCG 1350 Odocoileus virginianus

Deerc

Mink probe strongly x-hybridizes with Eastern cottontail rabbit (Sylvilagus floridanus). Swine probe weakly cross-hybridizes with human.

*DNA did not hybridize to probe.

†DNA hybridizes weakly to probe.

135