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. Author manuscript; available in PMC: 2022 May 1.
Published in final edited form as: Ticks Tick Borne Dis. 2021 Jan 12;12(3):101650. doi: 10.1016/j.ttbdis.2021.101650

Passerine birds as hosts for Ixodes ticks infected with Borrelia burgdorferi sensu stricto in southeastern Virginia.

Alexandra N Cumbie a, Erin L Heller a,b, Zachary J Bement a, Anna Phan a, Eric L Walters a, Wayne L Hynes a,*, Holly D Gaff a,c
PMCID: PMC7979520  NIHMSID: NIHMS1662297  PMID: 33486431

Abstract

The ecology of vector-borne diseases in a region can be attributed to vector-host interactions. In the United States, tick-borne pathogens are the cause of the highest number of reported vector-borne diseases. In the mid-Atlantic region of the eastern United States, tick-borne diseases such as Lyme disease, have increased in incidence, with tick-host-pathogen interactions considered a contributing factor to this increase. Ticks become infected with pathogens after taking a blood meal from a systemically infected host or through a localized infection while co-feeding on a host with other infected ticks. The host not only plays a role in pathogen acquisition by the tick, but can also facilitate dispersal of the tick locally within a region or over greater distances into new geographical ranges outside of their historical distributional range. In this study conducted in southeastern Virginia (USA), we examined the interaction between both resident and migratory bird species and Ixodes ticks, the primary vectors of Borrelia burgdorferi sensu stricto (s.s.) the main causative agent of Lyme disease on the East coast of the United States. Over a two-year period (2012–2014), 1879 passerine birds were surveyed, and 255 Ixodes ticks tested for the presence of Borrelia spp. Eighty passerine birds (4.3%) representing 17 bird species were parasitized by at least one Ixodes tick, but only three bird species were parasitized by Ixodes ticks that tested positive for B. burgdorferi s.s. Twenty Ixodes ticks (7.8%) tested positive for B. burgdorferi s.s. with nearly all collected from resident bird species including the Carolina wren (Thryothorus ludovicianus) and brown thrasher (Toxostoma rufum). Given that millions of birds pass through southeastern Virginia during migration, even with the low number of Ixodes ticks parasitizing passerine birds and the low prevalence of B. burgdorferi s.s. found within Ixodes ticks collected, the sheer volume of passerine birds suggests they may play a role in the maintenance and dispersal of B. burgdorferi s.s. in southeastern Virginia.

Keywords: Borrelia burgdorferi, Ixodes, Lyme, ticks, birds

Introduction

The eastern half of the United States is known to harbor many hard tick species (family Ixodidae) and their associated pathogens. Most notably on the east coast, where Ixodes ticks are populous, there is a high incidence of Lyme disease cases (Kugeler et al., 2015). The blacklegged tick (Ixodes scapularis) is one of several Ixodes species established in southeastern Virginia and is a vector of Borrelia burgdorferi sensu stricto (s.s.), the primary causative agent of Lyme disease in North America (Keirans et al., 1996). The relapsing fever group spirochete Borrelia miyamotoi, the causative agent of Borrelia miyamotoi disease (CDC, 2015), is also spread by Ixodes ticks in the United States and is of interest because it can be associated with co-infections of B. burgdorferi sensu lato (s.l.), including B. burgdorferi s.s. (Barbour et al., 2009).

Lyme disease incidence in North America is changing (CDC, 2018), particularly in the northern mid-Atlantic region of the United States, where the number of infections has increased substantially over the past 20 yr (Brinkerhoff et al., 2014). This increase is likely mediated, at least in part, by tick-host interactions (Levy, 2013). Because B. burgdorferi s.s. is not transmitted transovarially (Patrican, 1997a), Ixodes ticks become infected through a blood meal from an infected host or through localized infection caused by another infected tick feeding on the same host (States et al., 2017). Once infected, an Ixodes tick will retain B. burgdorferi s.s. through subsequent life stages and can transmit the pathogen to additional hosts (Gauthier-Clerc et al., 1998; Gatewood et al., 2009).

Passerine birds are frequently parasitized by ticks, including several Ixodes species, but the role of such hosts in the maintenance and transmission of B. burgdorferi in southeastern Virginia has, to our knowledge, not been examined previously. This region serves as a major stopover area to millions of migratory birds each autumn and spring (Hinshaw et al., 1985; Eggeman and Johnson, 1989). Thus, understanding the potential role of birds in tick-borne pathogen maintenance and transfer has far-reaching implications given the large concentration of birds in the area and their propensity to move long distances in short periods of time. Migrating birds likely play the greatest role among vertebrate hosts as long-distance dispersal agents of ticks and their associated pathogens when compared with mammalian, reptilian, or amphibian hosts that disperse ticks across much shorter distances (Smith et al., 1996; Scott et al., 2001; Loss et al., 2016; Norte et al., 2020). The goal of this study was to assess the role of passerine birds, both resident and migratory, as hosts of B. burgdorferi s.l.-infected ticks in a region of the eastern United States where passerine birds are concentrated during autumn and spring migration.

Methods

Host Collection and Tick Removal

Birds were caught and sampled at 10 locations across southeastern Virginia from August 2012 to August 2014 (Figure 1) using 12 m x 2.5 m mist nets erected for 4 h, typically beginning at local sunrise. Mist net sites consisted of 5 core locations that were sampled on one day every other week over the two years of this study (750+ net hours per site) and 5 sites that were sampled on an ad hoc basis at least once per month (≤350 net hours per site). Each bird was banded and checked for ticks, primarily around the head and vent. Ticks were removed using forceps, placed in individually labeled tubes, and stored at −20°C prior to identification and DNA extraction. We calculated the mean infestation of Ixodes per individual host species by taking the sum of all Ixodes ticks found on a particular host species divided by the total number of individual hosts of that species.

Fig. 1.

Fig. 1.

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Map showing location of mist-netting sites sampled during the 2-yr study period (2012–2014) in southeastern Virginia, United States.

We defined migratory birds as those that had seasonal movement from one region to another; however, we categorized some birds as “resident” even though they may exhibit some seasonal movement throughout parts of their range as our recapture data suggested some individuals did not exhibit such seasonal movements (e.g., American robin and brown thrasher); therefore, we conservatively classified them as residents.

Tick Identification and DNA Extraction

Ticks were identified morphologically to genus and life stage (Keirans and Clifford, 1978). All ticks identified as Ixodes sp. were separated from other tick species, labeled individually, and DNA was extracted using a DNeasy Blood and Tissue Kit (Qiagen, Inc., Valencia, CA, USA) following the manufacturer’s instructions. Larval and nymphal ticks were extracted whole, while adult ticks were cut bilaterally (one half of the tick was stored at −80°C and the other half pulverized for DNA extraction). Pulverization was carried out using 1 mm and 5 mm glass beads in a Mini Beadbeater (BioSpec, Inc., Bartlesville, OK, USA); extracted DNA was eluted in 200 μL elution buffer according to the manufacturer’s instructions.

Each individual Ixodes tick was identified to species by first using an I. scapularis I Ixodes affinis real-time PCR assay (Wright et al., 2014). The I. scapularis / I. affinis real-time PCR assay amplified a fragment of the tick internal-transcribed spacer 2 (ITS2) region of the nuclear rDNA transcriptional unit; amplification methodology followed that described by Wright et al. (2014). Melt curve real-time PCR reactions were carried out using 15 μL reaction volumes consisting of 1X Bio-Rad iQ SYBR green supermix, 0.5 μL of each primer (10 μM) (Table 1), and 2 μL of template DNA. Melt temperatures of samples with C(q) values ≤ 30 were used to separate the ticks into groups: I. scapularis, I. affinis, or Ixodes sp. for those that did not fit into either the I. scapularis or I. affinis groups. A 454-bp fragment of tick mitochondrial 16S rRNA gene was amplified using primers 16S+1 and 16S−1 (Table 1) to identify any Ixodes ticks that were neither I. scapularis nor I. affinis. PCR reactions were carried out using 12.5 μL of 2X EconoTaq PLUS (Lugien Corp., Middleton, WI, USA), 2 μL of each primer (10 μM), and 5 μL template DNA (de la Fuente et al., 2001; Nadolny et al., 2011). Samples not identified through mitochondrial 16S rRNA gene sequencing were further characterized by sequencing the 12S rDNA gene and the tick internal transcribed spacer (ITS) region (Table 1). PCR products for use in sequencing reactions were purified using Wizard PCR preps DNA Purification System (Promega, Madison, WI, USA) according to the manufacturer’s instructions. Sequencing reactions were performed using a BigDye Terminator v.3.1 Cycle Sequencing Kit (Applied Biosystems, Foster City, CA, USA). DNA sequences were assembled using Geneious Bioinformatics Software for Sequence Analysis (https://www.geneious.com/) and compared against known DNA sequences using NCBI BLAST (http://blast.ncbi.nlm.nih.gov).

Table 1.

Primers and probes used to determine Ixodes species.

Primer/Probe Primer/Probe Sequence Amplicon Size Reference
Tick 16S gene
16S−1 5’-GTCTGAACTCAGATCAAGT-3’ 454 bp de la Fuente et al., 2001
16S+1 5’-CTGCTCAATGATTTTTTAAATTGCTGT-3’ Nadolny et al., 2011
Tick 12S gene
85F 5’-TTAAGCTTTTCAGAGGAATTTGCTC-3’ 140 bp Williamson et al., 2010
225R 5’-TTTWWGCTGCACCTTGACTTAA-3’ Williamson et al., 2010
Ixodes ITS2 region
IsSeq 5.8F 5’-TCGATGAAGAACGCAGCCAG-3’ 800 bp Zahler et al., 1995
IsSeq 28SR 5’-TTCTATGCTTAAATTCAGGGGGTTGTC-3’ McLain et al., 1995
I. scapularis / I. affinis assay
aff_f8 5’-TGGAAATCCCGCAAATCT-3’ 75 bp Wright et al., 2014
aff_r8 5’-CCGTTCCAATCTCCGTTTA-3’ Wright et al., 2014
Scap_f2.2 5′-GCGTTAGAAACGGAGATTTGA-3′ 142 bp Wright et al., 2014
Scap_r2.2 5′-CCACGAGATTTACATTTGCC-3′ Wright et al., 2014
Ixodes actin
Actin-R 5’-CCGTCGGGAAGCTCGTAGG-3’ 76 bp Hojgaard et al., 2014
Actin-F 5’-GCCCTGGACTCCGAGCAG-3’ Hojgaard et al., 2014
Actin-Pr 5’-HEX-705-CCACCGCCGCCTCCTCTTCTTCC-BHQ3-3’ Hojgaard et al., 2014
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Borrelia Identification and Sequencing

To verify the quality of the extracts, an Ixodes actin real-time assay was performed as described by Graham et al. (2018). The real-time assay protocol was 95°C for 3 min, followed by 40 cycles of 95°C for 10 s, 60°C for 45 s, with a plate reading after the extension step. Real-time PCR reactions were carried out using 20 μL aliquots consisting of 10 μL of 2X EconoTaq PLUS master mix, 1 μL of each primer (10 μM), 0.5 μL of probe (10 μM) (Table 1), and 5 μL of template DNA. The quality of all extracts was verified as good with C(q) values below 30 for every sample. Tick DNA was screened for Borrelia sp. using a real-time PCR assay detecting the 23S rRNA gene as described in Courtney et al. (2004). Real-time PCR reactions were carried out using 20 μL aliquots consisting of 10 μL of 2X EconoTaq PLUS master mix, 1.4 μL of each primer (10 μM), 0.35 μL of probe (10 μM) (Table 2), 1.2 μL of MgCl2 (50 μM), and 5 μL of template DNA.

Table 2.

Primers and probes used to identify Borrelia spp.

Primer/Probe Primer/Probe Sequence Amplicon Size Reference
Borrelia 23S
Bb23SF 5’-CGAGTCTTAAAAGGGCGATTTAGT-3’ 75 bp Courtney et al., 2004
Bb23SR 5’-GCTTCAGCCTGGCCATAAATAG-3’ Courtney et al., 2004
Bb23SPr 5’-FAM-AGATGTGGTAGACCCGAAGCCGAGTG-3BHQ-3’ Courtney et al., 2004
Borrelia 16S
16S-F 5’-AGCYTTTAAAGCTTC GCTTGTAG-3’ 148 bp Kingry et al., 2018
16S-R 5’-GCCTCCCGTAGGAGTCTG G-3’ Kingry et al., 2018
16S-Pr 5’-FAM-CCGGCCTGAGAGGGTGAWCGG-3IABkFQ-3’ Kingry et al., 2018
Borrelia oppA2
Bb-F 5’-AATTTTTGGTTCCATACCC-3’ 162 bp Graham et al., 2018
Bb-R 5’-CTGTCAATAGCAAGAGTTAA-3’ Graham et al., 2018
Bb-Pr 5’-FAM-CGTTCAATACACACATCAAACCACT-3IABkFQ-3’ Graham et al., 2018
B. miyamotoi glpQ
BmglpQ-F 5’-GACCCAGAAATTGACAAACCACAA-3’ 108 bp Graham et al., 2018
BmglpQ-R 5’-TGATTTAAGTTCAGTTAGTGTGAAGTCAGT-3’ Graham et al., 2018
BmqlpQ-Pr 5’-FAM-CAATCGAGCTAGAGAAAACGGAAGATATTACG-3IABkFQ-3’ Graham et al., 2018
Borrelia flagellin
flaB outer 1 5’-AAGTAGAAAAAGTCTTAGTAAGAATGAAGGA-3’ 475 bp Johnson et al., 1992
flaB outer 2 5’-AATTGCATACTCAGTACTATTCTTTATAGAT -3’ Johnson et al., 1992
flaB inner 1 5’-CACATATTCAGATGCAGACAGAGGTTCTA- 3’ 437 bp Johnson et al., 1992
flaB inner 2 5’-GAAGGTGCTGTAGCAGGTGCTGGCTGT- 3’ Johnson et al., 1992
Borrelia ospC
  ospC outer-forward 5’-ATGAAAAAGAATACATTAAGTGC- 3’ 658 bp Tsao et al., 2013
  ospC outer-reverse 5’-ATTAATCTTATAATATTGATTTTAATTAAGG- 3’ Tsao et al., 2013
  ospC inner-forward 5’-TATTAATGACTTTATTTTTATTTATATCT- 3’ 618 bp Tsao et al., 2013
  ospC inner-reverse 5’-TTGATTTTAATTAAGGTTTTTTTGG- 3’ Tsao et al., 2013
Borrelia IGS
rrs-rrIA IGS F 5’-GTATGTTTAGTGAGGGGGGTG- 3’ 1028 bp Bunikis et al., 2004
rrs-rrIA IGS R 5’-GGATCATAGCTCAGGTGGTTAG- 3’ Bunikis et al., 2004
rra-rrIA IGS Fn 5’-AGGGGGGTGAAGTCGTAACAAG- 3’ 812 bp Bunikis et al., 2004
rra-rrIA IGS Rn 5’-GTCTGATAAACCTGAGGTCGGA- 3’ Bunikis et al., 2004
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Samples positive for the 23S real-time assay were also screened for Borrelia sp. using the Borrelia 16S real-time PCR assay described in Graham et al. (2018) but modified by running the samples as single reactions (using FAM-labeled probes) rather than as multiplex reactions (Table 2). Real-time PCR reactions were carried out in 20 μL aliquots consisting of 10 μL of 2X EconoTaq PLUS master mix, 1 μL of each primer (10 μM), 0.5 μL of probe (10 μM) (Table 2), and 5 μL of template DNA. The real-time assay was performed using the same protocol as the Ixodes actin assay (Graham et al., 2018). Borrelia-positive samples were further screened for B. burgdorferi s.s. and B. miyamotoi using the B. burgdorferi s.s. oppA2 real-time PCR assay modified as above (Graham et al., 2018) and a B. miyamotoi glpQ real-time PCR assay (Reiter et al., 2015), respectively. Each reaction for the B. burgdorferi s.s. oppA2 assay was carried out in 20 μL aliquots consisting of 10 μL of 2X EconoTaq PLUS master mix, 1 μL of each primer (10 μM), 0.5 μL of probe (10 μM) (Table 2), and 5 μL of template DNA. Each reaction for the B. miyamotoi glpQ was conducted using the same protocol as the Borrelia 16S assay. The positive control used in each B. burgdorferi assay was DNA from B. burgdorferi s.s. cell culture. The positive control used in the B. miyamotoi assay was tick DNA from a B. miyamotoi-infected adult I. scapularis. All negative controls used in these assays consisted of nuclease-free water.

Samples positive for B. burgdorferi s.s. from real-time screening were amplified using standard end point PCR for B. burgdorferi s.s.-specific genes: flaB, ospC, and IGS (Table 2). Amplified DNA was visualized following gel electrophoresis and purified using a Wizard PCR preps DNA Purification System (Promega) according to the manufacturer’s instructions. Sequencing reactions were performed using a BigDye Terminator v.3.1 Cycle Sequencing Kit using the primers for the specific genes (Table 2). DNA sequences were assembled in Geneious and compared against known sequences using NCBI BLAST.

Results

Host-Tick Interactions

During this study (2012–2014), 1879 birds were banded and checked for ticks at field sites in southeastern Virginia (Supplemental Table 1). The number of banded birds per bird species provided a relative measure of abundance at each sampling site, given that sampling effort was roughly equivalent among sites. Eleven percent of the birds sampled (207 of 1879) were parasitized by ticks (Ixodes and non-Ixodes) with approximately 4.3% of birds (80 of 1879) being parasitized by at least one Ixodes tick. Host species where >10% of individuals were parasitized by at least one Ixodes tick (Table 3) included Carolina wrens (Thryothorus ludovicianus; 32 of 186), brown thrashers (Toxostoma rufum; 12 of 75), gray-cheeked thrushes (Catharus minimus; 1 of 5), hermit thrushes (Catharus guttatus; 4 of 37), ovenbirds (Seiurus aurocapilla; 2 of 8), swamp sparrows (Melospiza georgiana; 3 of 15), and winter wrens (Troglodytes hiemalis; 1 of 9). The other bird hosts surveyed (10 additional species) with Ixodes ticks had infestation prevalences ≤ 10% (Figure 2).

Table 3.

Mean infestation of Ixodes per bird by host bird species; host bird species listed had >10% of individuals from that species parasitized by Ixodes spp. The number of birds examined parasitized by Ixodes spp. is listed as well as the range of the number of ticks parasitizing individual hosts from a species.

Host Bird Species Number of birds examined Number of birds parasitized by Ixodes ticks Total number of Ixodes ticks from each host species (range of ticks per bird) Mean infestation of Ixodes per bird (+/− SD)
Carolina wren 186 32 180 (1–50) 0.97 (+/− 4.58)
Thryothorus ludovicianus
Brown thrasher 75 12 28 (1–8) 0.37 (+/− 1.16)
Toxostoma rufum
Gray-cheeked thrush 5 1 1 (N/A) 0.20 (+/− 0.45)
Catharus minimus
Hermit thrush 37 4 7 (1–3) 0.19 (+/− 0.62)
Catharus guttatus
Ovenbird 8 2 5 (2–3) 0.50 (+/− 0.93)
Seiurus aurocapilla
Swamp sparrow 15 3 4 (1–2) 0.27 (+/− 0.59)
Melospiza georgiana
Winter wren 9 1 2 (N/A) 0.22 (+/− 0.67)
Troglodytes hiemalis
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Fig. 2.

Fig. 2.

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Percentage of birds with Ixodes ticks in southeastern Virginia by species. Number above each bar represents the number of individual birds surveyed for each bird species.

Two-hundred fifty-five Ixodes ticks, representing 4 Ixodes species, were collected during this study. Of the Ixodes collected, 84 were I. scapularis, 77 Ixodes brunneus, 64 I. affinis, and 30 Ixodes dentatus. Only three adult Ixodes ticks were detected on birds in this study, i.e., two female I. brunneus from an American robin (Turdus migratorius) and one I. brunneus female from a Carolina wren. Larvae from all four species were collected while nymphs were only collected for I. scapularis, I. brunneus, and I. affinis. Each Ixodes species was found parasitizing multiple bird species (Table 4), both resident and migratory. All four Ixodes species were found parasitizing Carolina wrens and northern cardinals (Cardinalis cardinalis), both of which are resident to the study area. American robins and brown thrashers, also resident bird species, were parasitized by three of the four Ixodes species. Migratory hosts were only parasitized by one or two of the four Ixodes species found.

Table 4.

Ixodes species collected from various avian hosts.

Tick Species Host Bird Species Migratory Status Individuals Infested (%) Mean number of Ixodes per bird (+/− SD)
Ixodes scapularis American robin resident 1.2 0.01 (+/− 0.11)
Brown thrasher resident 9.3 0.15 (+/− 0.51)
Carolina wren resident 11.8 0.35 (+/− 1.67)
Northern cardinal resident 0.3 0.00 (+/− 0.06)
Ovenbird migratory 25.0 0.50 (+/− 0.93)
Swamp sparrow migratory 6.7 0.07 (+/− 0.26)
Ixodes affinis American robin resident 0.6 0.01 (+/− 0.08)
Brown thrasher resident 2.7 0.05 (+/− 0.36)
Carolina wren resident 5.4 0.28 (+/− 2.14)
Common yellowthroat migratory 0.3 0.03 (+/− 0.17)
Dark-eyed junco migratory 4.8 0.05 (+/− 0.22)
Eastern towhee resident 5.6 0.06 (+/− 0.24)
Northern cardinal resident 0.3 0.00 (+/− 0.06)
White-throated sparrow migratory 0.6 0.01 (+/− 0.08)
Ixodes brunneus American robin resident 1.8 0.02 (+/− 0.13)
Brown thrasher resident 2.7 0.03 (+/− 0.16)
Carolina wren resident 4.3 0.25 (+/− 2.94)
Dark-eyed junco migratory 4.8 0.05 (+/− 0.22)
Hermit thrush migratory 10.8 0.19 (+/− 0.62)
Northern cardinal resident 0.3 0.00 (+/− 0.06)
Swamp sparrow migratory 13.3 0.20 (+/− 0.56)
Tufted titmouse resident 6.3 0.06 (+/− 0.25)
White-throated sparrow migratory 4.4 0.06 (+/− 0.33)
Ixodes dentatus Carolina wren resident 2.7 0.08 (+/− 0.75)
Brown thrasher resident 4.0 0.15 (+/− 0.95)
Gray-cheeked thrush migratory 20.0 0.20 (+/− 0.45)
Gray catbird resident 1.0 0.01 (+/− 0.10)
Northern cardinal resident 0.3 0.00 (+/− 0.06)
Winter wren migratory 11.1 0.22 (+/− 0.67)
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Phenologies of immature tick life stages (larvae and nymphs) on birds were determined during the 2-yr sampling period based on the number collected each calendar month for each Ixodes species. Ixodes scapularis larval feeding activity on bird hosts occurred from April through October with a peak in July; I. affinis larvae from June to December with peaks in August and October; I. dentatus larvae from July to November with the peak in October and November; and I. brunneus larvae in November and December with the peak in November (Figure 3). The number of Ixodes nymphs found was much lower than the number of larvae (55 nymphs versus 197 larvae). The peak feeding time for I. scapularis nymphs (representing the majority of nymphs collected) was during July and August, I. affinis nymphs had peaks in July and December, and I. brunneus nymphs had peaks in March and December (Figure 4); no I. dentatus nymphs were collected. Ixodes scapularis nymphal feeding overlapped with I. scapularis, I. affinis, and I. dentatus larval feeding from April to October.

Fig. 3.

Fig. 3.

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The phenology of larval Ixodes spp. on birds in this study over a 2-yr period (2012–2014). Individual data points were fitted to a cubic spline curve. Symbols above the graph indicate the month when Borrelia burgdorferi sensu stricto-infected Ixodes ticks were collected. Infected I. scapularis larvae occurred at different points in the year. Inline graphic: infected I. scapularis larvae from Carolina wrens; Inline graphic: infected I. scapularis larvae from brown thrashers; Inline graphic: infected I. affinis larvae from Carolina wrens; and Inline graphic: infected I. brunneus larvae from swamp sparrows.

Fig. 4.

Fig. 4.

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The phenology of nymphal Ixodes spp. on birds in this study over a 2-yr period (2012–2014). Individual data points were fitted to a cubic spline curve. Symbols above the graph indicate the month when Borrelia burgdorferi sensu stricto-infected ticks were collected. Infected I. scapularis nymphs occurred at different points in the year. Inline graphic: infected I. scapularis nymphs from Carolina wrens; Inline graphic: infected I. scapularis nymphs from brown thrashers.

Tick-Borrelia Interactions

Borrelia spp. DNA was detected in 20 of the 255 immature Ixodes ticks screened by real-time PCR detecting the 23S rRNA region (Table 5); 14 were I. scapularis, 5 I. affinis, and 1 I. brunneus. Based on real-time PCR, only 5 ticks were positive for B. burgdorferi s.s. (Table 5), while no ticks tested positive for B. miyamotoi. The majority of infected Ixodes were larvae. All I. affinis and I. brunneus infected with Borrelia sp. were larvae. For I. scapularis, 7 larvae and 7 nymphs were infected.

Table 5.

Bird host and life stage associated with Ixodes ticks that tested positive for Borrelia burgdorferi sensu stricto.

Real-time PCR assay
(0-negative, 1-positive)		 Gene targeted sequencing
(Yes-able to amplify, No-did not amplify)		 Borrelia species detected
Bird Species Ixodes spp. Tick Life Stage Bb 23S Bb 16S Bb s.s. flaB ospC IGS
KP0 CARW I. scapularis Nymph 1 † † † No † B. burgdorferi s.l.
CH2 BRTH I. scapularis Larva 1 1 1 Yes Yes Yes B. burgdorferi s.s.
CH2 BRTH I. scapularis Nymph 1 1 0 Yes No No B. burgdorferi s.s.
CH2 CARW I. scapularis Larva 1 1 0 Yes No No B. burgdorferi s.s.
CH2 CARW* I. affinis Larva 1 1 0 Yes No Yes B. burgdorferi s.s.
CH2 CARW* I. affinis Larva 1 1 0 Yes No Yes B. burgdorferi s.s.
CH2 CARW* I. affinis Larva 1 0 0 Yes No No B. burgdorferi s.s.
CH2 CARW* I. affinis Larva 1 1 0 Yes No No B. burgdorferi s.s.
CH2 CARW* I. affinis Larva 1 0 0 Yes No No B. burgdorferi s.s.
CH1 SWSP I. brunneus Larva 1 0 0 Yes No No B. burgdorferi s.s.
PT2 CARW I. scapularis Nymph 1 † † Yes No † B. burgdorferi s.s.
PT2 CARW I. scapularis Nymph 1 † † Yes No † B. burgdorferi s.s.
PT2 CARW I. scapularis Nymph 1 0 0 Yes No No B. burgdorferi s.s.
SF0 CARW* I. scapularis Larva 1 1 1 Yes Yes Yes B. burgdorferi s.s.
SF0 CARW* I. scapularis Larva 1 1 1 Yes Yes Yes B. burgdorferi s.s.
SF0 CARW* I. scapularis Larva 1 1 1 Yes Yes Yes B. burgdorferi s.s.
SF0 CARW* I. scapularis Nymph 1 1 1 Yes Yes Yes B. burgdorferi s.s.
YRK1 CARW I. scapularis Larva 1 † † Yes No † B. burgdorferi s.s.
YRK1 CARW* I. scapularis Larva 1 † † Yes No Yes B. burgdorferi s.s.
YRK1 CARW* I. scapularis Nymph 1 † † Yes No † B. burgdorferi s.s.
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*

Same bird

†

Ran out of DNA

CARW = Carolina wren, BRTH = brown thrasher, SWSP = swamp sparrow

Eighteen of the twenty real-time positive samples were amplified and confirmed by sequencing of the flaB gene in both directions (Table 5); one was only sequenced in one direction for the flaB gene; the other ran out of template. All sequences were homologous with Borrelia burgdorferi s.s. with no other Borrelia species detected by sequence identity. Five Borrelia from I. scapularis were sequenced in at least one direction for the 3 genes tested and two Borrelia from I. affinis were sequenced in at least one direction for 2 of the 3 genes tested. The Borrelia from the single I. brunneus larva was only amplified and sequenced for the flaB gene. Sequencing indicated that these ticks carried B. burgdorferi s.s., with sequence identity over 570 bp for the ospC gene, 260 bp for flaB, and 308 bp for IGS.

There were many instances of co-feeding on birds with Borrelia-infected ticks. The Carolina wren from site KP0 was found parasitized by three I. scapularis nymphs in August; one of which was positive for Borrelia sp. by real-time PCR; we were unable to determine the species of Borrelia. At site CH2, four birds were parasitized by Borrelia-infected ticks (one each in April, May, July, and August). Two birds from site CH2 had co-feeding ticks; one bird had two I. scapularis larvae (one of which was positive for B. burgdorferi s.s.); the other bird had one I. scapularis nymph (negative for Borrelia spp.) and 27 I. affinis larvae (five of which were positive for B. burgdorferi s.s.). At site CH1, the swamp sparrow was parasitized by two I. brunneus larvae in December, but only one was positive for B. burgdorferi s.s. At site PT2, three birds were parasitized by Borrelia-infected I. scapularis nymphs in July, August, and September. Two birds had other Ixodes ticks co-feeding with the infected I. scapularis nymphs (one bird with a second I. scapularis larva; one bird with one I. scapularis nymph and one I. affinis larva), but none of these co-feeding ticks were positive for B. burgdorferi s.s. At site SF0, one Carolina wren was parasitized by three Borrelia-infected I. scapularis larvae and one nymph in June. This bird was also parasitized by three other I. scapularis larvae and one I. brunneus nymph. At site YRK1, two Carolina wrens were parasitized by I. scapularis larvae and nymphs in June. One bird had 11 larvae and one nymph but only one larva tested positive for B. burgdorferi s.s.; the other bird had 12 larvae and five nymphs with one larva and one nymph testing positive for B. burgdorferi s.s.

Discussion

Few Ixodes ticks were found parasitizing passerine birds over the 2-yr period of this study in southeastern Virginia, with a subset of ticks being positive for B. burgdorferi s.s. Four Ixodes species were found parasitizing birds: I. scapularis was the most common species detected, a species considered medically important as it is the only human-biting vector of B. burgdorferi s.s. in Virginia (Piesman and Sinsky, 1988). Ixodes affinis and I. brunneus have not been observed to bite humans (Nadolny et al., 2011; Merten and Durden, 2000; Luttrell et al., 1996), while I. dentatus has rarely been reported to feed on humans (Kollars and Oliver, 2003). Despite I. affinis and I. brunneus not feeding on humans and the rarity with which I. dentatus feeds on humans, these three tick species contribute to the sylvatic cycle of B. burgdorferi s.s. in animals, including avian hosts (Heller et al., 2016). Ixodes affinis, in particular, serves as a vector and plays an important role in the sylvatic cycle of B. burgdorferi s.s. involving small mammals (Nadolny et al., 2011).

The seasonal activity (phenology) of immature I. scapulars varies geographically, with debate over whether spring active nymphs and late summer active larvae (as occurs in the northeastern United States) or synchronous phenology of nymphs and larvae based on computer models is better for transmission of B. burgdorferi (Ogden et al., 2018). A reversed pattern, with larvae active earlier in the year and nymphs in the autumn, seen in the southeastern United States may explain the lower Lyme disease incidence in this part of the country (Ogden et al., 2018). Additionally, there is an East-West gradient (northeast to upper Midwest) that results in a bimodal larval phenology in the Midwest. Two states in which data were collected by Ogden et al. (2018) closest to Virginia showed differences in immature I. scapularis phenologies; Tennessee having nymphs feeding before larvae, while North Carolina had larvae before nymphs. Stromdahl et al. (2014), using information from I. scapularis collected from humans over a 15-yr period from 1997 to 2012, reported that in Pennsylvania the nymphal peak was in June, while in Minnesota both nymphs and adult I. scapularis peaked in June. Phenologies of the immatures of most Ixodes species in Virginia have not been fully characterized. Our study provides important insight into the timing of host-seeking activity for immature I. scapularis, I. affinis, I. brunneus, and I. dentatus on avian hosts in this region. Host-seeking activity of I. affinis in Virginia has been previously examined (Nadolny et al., 2018); on small mammals, I. affinis larval feeding activity peaks in winter, but on birds, we found I. affinis larval feeding activity peaked later in August. We observed this same trend for nymphs, where peak feeding activity is in April on small mammals but much later, July to September, on birds. Despite these differences, phenological data for all immature Ixodes ticks from any host are useful in understanding the disease ecology of an area. Phenological data for immature Ixodes ticks collected from birds is particularly important for determining dispersal of potentially infected ticks into novel areas. The phenologies we observed do not match those reported for immature I. scapularis from nearby states (Stromdahl et al., 2014; Ogden et al., 2018); therefore, determining the phenology of Ixodes spp. in southeastern Virginia is important in understanding the role of these ticks in the prevalence of pathogenic B. burgdorferi and the risk of human infection.

Finding larval ticks infected with B. burgdorferi s.s. is suggestive that some birds were infected with B. burgdorferi s.s. Host reservoir competence for B. burgdorferi s.s. is variable among avian taxa, as well as within individuals of a given species (Richter et al., 2000; Ginsberg et al., 2005). Within the United States, up to 20% of Ixodes ticks, and up to 50% of nymphs, collected from birds have been reported to be infected with B. burgdorferi s.l. (Smith et al., 1996; Hamer et al., 2012a, 2012b; Dingier et al., 2014; Newman et al., 2015; Heller et al., 2016), indicating that birds may serve as reservoirs of Borrelia burgdorferi s.s. Avian reservoir competency studies found American robins, song sparrows (Melospiza melodia,) northern cardinals, and gray catbirds (Dumetella carolinensis) are reservoir-competent for B. burgdorferi s.s. (Ginsberg et al., 2005). We found Ixodes species parasitizing American robins, gray catbirds, and northern cardinals, but none of these ticks were infected with B. burgdorferi s.s. or any other Borrelia species; however, it is noted that our sample size of Ixodes from these birds was relatively low.

As an alternative to a systemic host infection, the Ixodes larvae in our study could have become infected with B. burgdorferi s.s. on hosts by concomitant feeding with other infected ticks through a localized infection (Patrican, 1997b). Two of the infected ticks were from different life stages feeding on the same Carolina wren host; in both incidences, we found an I. scapularis nymph and larva (both infected) feeding. The wren may have served as a host in which a localized infection of B. burgdorferi s.s. could have occurred following the bite from an infected nymph that then allowed the larvae to acquire the pathogen. The phenological data obtained in this study demonstrated that I. scapularis nymphs actively seek hosts and feed from April to October, overlapping with I. scapularis, I. affinis, and I. dentatus larval feeding. These findings suggest the other infected I. scapularis and I. affinis larvae in our study may have encountered birds parasitized by infected I. scapularis nymphs during their blood meals, even if co-feeding was not documented at the time of collection.

The proportion of birds parasitized by Ixodes ticks in general, but B. burgdorferi s.s.-infected Ixodes ticks in particular, was low in comparison to that seen with small mammal sampling in other regions of southeastern Virginia (Anderson et al., 1985, 1987). Although small mammals may not have the capacity for long-distance dispersal, they are host to Ixodes ticks and amplify reservoirs in the enzootic cycle of B. burgdorferi s.s. (Levine et al., 1985). For brown thrashers and Carolina wrens, prevalence rates of individuals parasitized by infected ticks varied from 2.7 to 4.8%, respectively. The brown thrasher and the Carolina wren are both ground foragers; however, the lower prevalence in the brown thrasher is not unexpected since this species is known to be reservoir-incompetent for B. burgdorferi s.s. (Ginsberg et al., 2005). Swamp sparrows had the highest prevalence of individuals parasitized by B. burgdorferi s.s.-infected ticks (6.7%, 1 of 15 birds surveyed) albeit based on a small sample size. Systemic B. burgdorferi s.s. infection (i.e. reservoir competency) in Carolina wrens and swamp sparrows has not been studied to date.

We did not find evidence that one bird species was more likely to harbor B. burgdorferi s.s.-infected Ixodes ticks than another. The mean infestation number of ticks per host species was low for all bird species parasitized by Ixodes juveniles, with numbers typically <1 Ixodes tick per host (Table 3). However, the mean infestation number of infected ticks per host species may have been skewed because of small sample sizes for some bird species. For instance, 25% (2 of 8) of ovenbirds were parasitized by Ixodes ticks, but none were positive for B. burgdorferi s.s The risk of encountering a B. burgdorferi s.s.-infected tick may change with a larger host sample size, but for our study region, the low number of captures per host species likely represents the relative abundance and presence of hosts in this region and does provide an assessment of host roles and parasitism by infected Ixodes ticks.

It is important to note the locations on the bird where the Ixodes ticks were attached. We predominantly surveyed the head and vent region, where ticks regularly congregate; however, a recent paper by Roselli et al. (2020) noted other body regions that were parasitized by non-Ixodes ticks including the flank, thigh, and wing. These areas were not routinely checked for ticks in our study, which may have contributed to a lower number of ticks detected per bird and/or the overall number of ticks detected on certain bird species.

Our results indicate that passerine birds harbor both larval and nymphal life stages of B. burgdorferi s.s.-infected Ixodes ticks, albeit in low numbers and on a small number of hosts. Host species that were resident year-round were parasitized by more B. burgdorferi s.s.-infected Ixodes ticks than migratory species, suggesting long-distance dispersal of infected ticks initially parasitized in the southeastern region of Virginia is not common. Resident bird species parasitized by Ixodes ticks (e.g. Carolina wrens) may explain movement of B. burgdorferi-infected ticks within localized regions and may help in assessing the human risk of encountering infected ticks. We acknowledge that several of the species we categorized as “resident” (e.g. American robin, brown thrasher, Eastern towhee, and gray catbird) do exhibit some form of seasonal movement throughout parts of their range (Cavitt and Haas, 2020; Greenlaw, 2020; Smith et al., 2020; Vanderhoff et al., 2020); however based on our recapture data indicating some individuals did not exhibit such seasonal movements we classified them as resident. Moreover, given the timing of parasitism, we suspect the source of the feeding ticks was locally obtained, rather than being a function of seasonal movement. However, we acknowledge that some individuals sampled during the winter could be temperate migrants wintering at our field sites. Recognizing that many of the resident host species in this study are found associated with urban areas (Heller et al., 2019), we cannot determine if Borrelia burgdorferi s.s. infections in our region can be explained by the dispersal of infected ticks on passerine birds. We also do not know if our findings scale up to explain patterns exhibited at much larger continental scales. This, due in part, to the particular community of tick species found in our study area; in particular, the relative abundance of I. affinis to I. scapularis.

Supplementary Material

1

Acknowledgements

This work was funded in part by National Institutes of Health grant 1R01AI136035 as part of the joint National Institutes of Health-National Science Foundation-United States Department of Agriculture Ecology and Evolution of Infectious Diseases program and in part by grant no. K25AI067791 (to HDG) from the National Institute of Allergy and Infectious Diseases. This study was also supported through start-up funds to ELW provided by Old Dominion University (ODU), ODU’s Research for Undergraduates in Math and Sciences, and the Virginia Academy of Science.

We thank all of our volunteer field assistants that assisted with data collection and entry with particular thanks to: Tyler Chavers, Amy Johnson, Ally Lahey, Ashley Morris, Dorothy Paine, Annie Sabo, and Alexis White. Also special thanks to Chelsea Wright for her contributions to pathogen testing.

Research compliances included: Old Dominion University IACUC Protocol # 13-018, US Department of the Interior Federal Bird Banding Permit #23803, Virginia Department of Game and Inland Fisheries Wildlife Salvage Permit # 044737, Virginia Department of Game and Inland Fisheries Scientific Collection Permit #044735, Commonwealth of Virginia Department of Conservation and Recreation Research and Collecting Permit #FLKPYR-RCP-030512, and the United States Fish and Wildlife Service Permit #MG71673A-0.

Footnotes

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References

  1. Anderson JF, Johnson RC, Magnarelli LA, Hyde FW, 1985. Identification of endemic foci of Lyme disease: isolation of Borrelia burgdorferi from feral rodents and ticks (Dermacentor variabilis). J. Clin. Microbiol 22, 36–38. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Anderson JF, Johnson RC, Magnarelli LA, 1987. Seasonal prevalence of Borrelia burgdorferi in natural populations of white-footed mice (Peromyscus leucopus). J. Clin. Microbiol 25, 1564–1566. [DOI] [PMC free article] [PubMed] [Google Scholar]
  3. Barbour AG, Bunikis J, Travinsky B, Hoen AG, Diuk-Wasser MA, Fish D, Tsao JI, 2009. Niche partitioning of Borrelia burgdorferi and Borrelia miyamotoi in the same tick vector and mammalian reservoir species. Am. J. Trop. Med. Hyg 81, 1120–1131. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Brinkerhoff J, Gilliam WF, Gaines D, 2014. Lyme disease, Virginia, USA, 2000–2011. Emerg. Infect. Dis 20, 1661–1668. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Bunikis J, Garpmo U, Tsao J, Berglund J, Fish D, Barbour AG, 2004. Sequence typing reveals extensive strain diversity of the Lyme borreliosis agents Borrelia burgdorferi in North America and Borrelia afzelii in Europe. Microbiology 150, 1741–1755. [DOI] [PubMed] [Google Scholar]
  6. Cavitt JF, Haas CA, 2020. Brown Thrasher (Toxostoma rufum), version 1.0. In Birds of the World (Poole AF, Editor). Cornell Lab of Ornithology, Ithaca, NY, USA. 10.2173/bow.brnthr.01 Accessed 4 December 2020. [DOI] [Google Scholar]
  7. CDC (Centers for Disease Control and Prevention)., 2015. Relapsing Fever, https://www.cdc.gov/relapsing-fever/. Accessed 4 December 2020.
  8. CDC (Centers for Disease Control and Prevention)., 2018. Lyme Disease: Recent Surveillance Data. https://www.cdc.gov/lyme/datasurveillance/recent-surveillance-data.html. Accessed 4 December 2020.
  9. Courtney JW, Kostelnik LM, Zeidner NS, Massung RF, 2004. Multiplex real-time PCR for detection of Anapiasma phagocytophilum and Borrelia burgdorferi. J. Clin. Microbiol 42, 3164–3168. [DOI] [PMC free article] [PubMed] [Google Scholar]
  10. de la Fuente J, Van Den Bussche RA, Kocan KM, 2001. Molecular phylogeny and biogeography of North American isolates of Anapiasma marginale (Rickettsiaceae: Ehrlichieae). Vet. Parasitol 97, 65–76. [DOI] [PubMed] [Google Scholar]
  11. Dingier RJ, Wright SA, Donohue AM, Macedo PA, Foley JE, 2014. Surveillance for Ixodes pacificus and tick-borne pathogens Anaplasma phagocytophilum and Borrelia burgdorferi in birds from California’s inner coast range. Ticks Tick Borne Dis. 5, 436–45. [DOI] [PubMed] [Google Scholar]
  12. Eggeman DR, Johnson FA, 1989. Variation in effort and methodology for midwinter waterfowl inventory in the Atlantic flyway. Wildl. Soc. Bull 17, 227–233. [Google Scholar]
  13. Gatewood AG, Liebman KA, Vourc’h G, Bunikis J, Hamer SA, Cortinas R, Melton F, Cislo P, Kitron U, Tsao J, Barbour AG, Fish D Diuk-Wasser MA., 2009. Climate and tick seasonality are predictors of Borrelia burgdorferi genotype distribution. Appl. Environ. Microbiol 75, 2476–2483. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. Gauthier-Clerc M, Clerqin MY, Handrich Y, 1998. Hyperinfestation by ticks Ixodes uriae: a possible cause of death in adult King Penguins, a long-lived seabird. Colon. Waterbird. 21, 229–233. [Google Scholar]
  15. Ginsberg HS, Buckley PA, Balmforth MG, Zhioua E, Mitra S, Buckley FG, 2005. Reservoir competence of native North American birds for the Lyme disease spirochete, Borrelia burgdorferi. J. Med. Entomol 42, 445–449. [DOI] [PubMed] [Google Scholar]
  16. Graham CB, Maes SE, Hojgaard A, Fleshman AC, Sheldon SW, Eisen RJ, 2018. A molecular algorithm to detect and differentiate human pathogens infecting Ixodes scapularis and Ixodes pacificus (Acari: Ixodidae). Ticks Tick Borne Dis. 9, 390–403. [DOI] [PMC free article] [PubMed] [Google Scholar]
  17. Greenlaw JS, 2020. Eastern Towhee (Pipilo erythrophthalmus), version 1.0. In Birds of the World (Rodewald PG, Editor). Cornell Lab of Ornithology, Ithaca, NY, USA. 10.2173/bow.eastow.01 Accessed 4 December 2020. [DOI] [Google Scholar]
  18. Hamer SA, Goldberg TL, Kitron UD, Brawn JD, Anderson TK, Loss SR, Walker ED, Hamer GL 2012a. Wild birds and urban ecology of ticks and tick-borne pathogens, Chicago, Illinois, USA, 2005 – 2010. Emerg. Infect. Dis 18, 1589–1595. [DOI] [PMC free article] [PubMed] [Google Scholar]
  19. Hamer SA, Hickling GJ, Keith R, Sidge JL, Walker ED, Tsao JI, 2012b. Associations of passerine birds, rabbits, and ticks with Borrelia miyamotoi and Borrelia andersonii in Michigan, USA. Parasit. Vectors 5, 1–11. [DOI] [PMC free article] [PubMed] [Google Scholar]
  20. Heller EL, Wright CL, Nadolny RM, Hynes WL, Gaff HD, Walters EL, 2016. New records of Ixodes affinis (Acari: Ixodidae) parasitizing avian hosts in southeastern Virginia. J. Med. Entomol 53, 441–445. [DOI] [PMC free article] [PubMed] [Google Scholar]
  21. Heller EL, Gaff HD, Brinkerhoff RJ, Walters EL, 2019. Urbanization and tick parasitism in birds of coastal southeastern Virginia. J Wildl. Manag 83, 975–984. [Google Scholar]
  22. Hinshaw VS, Wood JM, Webster RG, Deibel R, Turner B, 1985. Circulation of influence viruses and paramyxoviruses in waterfowl originating from two different areas of North America. Bull. World Health Organ 63, 711–719. [PMC free article] [PubMed] [Google Scholar]
  23. Hojgaard A, Lukacik G, Piesman J, 2014. Detection of Borrelia burgdorferi, Anaplasma phagocytophilum and Babesia microti, with two different multiplex PCR assays. Ticks Tick Borne Dis. 5, 349–351. [DOI] [PubMed] [Google Scholar]
  24. Johnson BJB, Happ CM, Mayer LW, Piesman J, 1992. Detection of Borrelia burgdorferi in ticks by species-specific amplification of the flagellin gene. Am. J. Trap. Med. Hyg 47, 730–741. [DOI] [PubMed] [Google Scholar]
  25. Keirans JE, Clifford CM, 1978. The genus Ixodes in the United States: a scanning electron microscope study and key to the adults. J. Med. Entomol Suppl. 2, 1–149. [DOI] [PubMed] [Google Scholar]
  26. Keirans JE, Hutcheson HJ, Durden LA, Klompen JSH, 1996. Ixodes (Ixodes) scapularis (Acari:Ixodidae): redescription of all active stages, distribution, hosts, geographical variation, and medical and veterinary importance. J. Med. Entomol 33, 297–318. [DOI] [PubMed] [Google Scholar]
  27. Kingry LC, Anacker M, Pritt B, Bjork J, Respicio-Kingry L, Liu G, Sheldon S, Boxrud D, Strain A, Oatman S, Berry J, Sloan L, Mead P, Neitzel D, Kugeler KJ, Petersen JM, 2018.Surveillance for and discovery of Borrelia species in US patients suspected of tickborne illness. Clin. Infect. Dis 66, 1864–1871. [DOI] [PMC free article] [PubMed] [Google Scholar]
  28. Kollars TM, Oliver JH, 2003. Host associations and seasonal occurrence of Haemaphysalis leporispalustris, Ixodes brunneus, I. cookei, I. dentatus, and I. texanus (Acari: Ixodidae) in southeastern Missouri. J. Med. Entomol 40, 103–107. [DOI] [PubMed] [Google Scholar]
  29. Kugeler KJ, Farley GM, Forrester JD, Mead PS, 2015. Geographic distribution and expansion of human Lyme disease, United States. Emerg. Infect. Dis 21, 1455–1457. [DOI] [PMC free article] [PubMed] [Google Scholar]
  30. Levine JF, Wilson ML, Spielman A, 1985. Mice as reservoirs of the Lyme disease spirochete. Am. J. Trap. Med. Hyg 34, 355–360. [DOI] [PubMed] [Google Scholar]
  31. Levy S, 2013. The Lyme disease debate host biodiversity and human disease risk. Environ. Health Perspect 121, a120–a125. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Loss SR, Noden BH, Hamer GL, Hamer SA, 2016. A quantitative synthesis of the role of birds in carrying ticks and tick-borne pathogens in North America. Oecologia 182, 947–959. [DOI] [PubMed] [Google Scholar]
  33. Luttrell MP, Creekmore LH, Mertins JW, 1996. Avian tick paralysis caused by Ixodes brunneus in the southeastern United States. J. Wildl. Dis 32, 133–136. [DOI] [PubMed] [Google Scholar]
  34. McLain DK, Wesson DM, Oliver JH Jr, Collins FH, 1995. Variation in ribosomal DNA internal transcribed spacers 1 among eastern populations of Ixodes scapularis (Acari: Ixodidae). J. Med. Entomol 32, 353–360. [DOI] [PubMed] [Google Scholar]
  35. Merten H, Durden L, 2000. A state-by-state survey of ticks recorded from humans in the United States. J. Vector. Ecol 25, 2–13. [PubMed] [Google Scholar]
  36. Nadolny RM, Gaff HD, 2018. Natural history of Ixodes affinis in Virginia. Ticks Tick Borne Dis. 9, 109–119. [DOI] [PubMed] [Google Scholar]
  37. Nadolny RM, Wright CL, Hynes WL, Sonenshine DE, Gaff HD, 2011. Ixodes affinis (Acari: Ixodidae) in southeastern Virginia and implications for the spread of Borrelia burgdorferi, the agent of Lyme disease. J. Vector Ecol 36, 464–467. [DOI] [PubMed] [Google Scholar]
  38. Newman EA, Eisen L, Eisen RJ, Fedorova N, Hasty JM, Vaughn C, Lane RS 2015. Borrelia burgdorferi sensu lato spirochetes in wild birds in northwestern California: associations with ecological factors, bird behavior and tick infestation. PLoS One 10, e0118146. [DOI] [PMC free article] [PubMed] [Google Scholar]
  39. Norte AC, Margos G, Becker NS, Albino Ramos J, Núncio MS, Fingerle V, Araújo PM, Adamík P, Alivizatos H, Barba E, Barrientos R, 2020. Host dispersal shapes the population structure of a tick-borne bacterial pathogen. Molec. Ecol 29, 485–501. [DOI] [PubMed] [Google Scholar]
  40. Ogden NH, Pang G, Ginsberg HS, Hickling GJ, Burke RL, Beati L, Tsao JI, 2018. Evidence for geographic variation in life-cycle processes affecting phenology of the Lyme disease vector Ixodes scapularis (Acari: Ixodidae) in the United States, J. Med. Entomol 55, 1386–1401. [DOI] [PubMed] [Google Scholar]
  41. Patrican LA, 1997a. Absence of Lyme disease spirochetes in larval progeny of naturally infected Ixodes scapularis (Acari: Ixodidae) fed on dogs. J. Med. Entomol 34, 52–55. [DOI] [PubMed] [Google Scholar]
  42. Patrican LA, 1997b. Acquisition of Lyme disease spirochetes by cofeeding Ixodes scapularis ticks. Am. J. Trap. Med. Hyg 57, 589–593. [DOI] [PubMed] [Google Scholar]
  43. Piesman J, Sinsky RJ, 1988. Ability of Ixodes scapularis, Dermacentor variabilis, and Amblyomma americanum (Acari: Ixodidae) to acquire, maintain, and transmit Lyme disease spirochetes (Borrelia burgdorferi). J. Med. Entomol 25, 336–339. [DOI] [PubMed] [Google Scholar]
  44. Reiter M, Schötta A-M, Müller A, Stockinger H, Stanek G, 2015. A newly established real-time PCR for detection of Borrelia miyamotoi in Ixodes ricinus ticks. Ticks Tick Borne Dis. 6, 303–308. [DOI] [PubMed] [Google Scholar]
  45. Richter D, Spielman A, Komar N, Matuschka FR, 2000. Competence of American robins as reservoir hosts for Lyme disease spirochetes. Emerg. Infect. Dis 6, 133–138. [DOI] [PMC free article] [PubMed] [Google Scholar]
  46. Roselli MA, Cady SM, Lao S, Noden BH, Loss SR, 2020. Variation in tick load among bird body parts: implications for studying the role of birds in the ecology and epidemiology of tick-borne diseases. J. Med. Entomol 57, 845–851. [DOI] [PubMed] [Google Scholar]
  47. Scott JD, Fernando K, Banerjee SN, Durden LA, Byrne SK, Banerjee M, Mann RB, Morshed MG, 2001. Birds disperse ixodid (Acari: Ixodidae) and Borrelia burgdorferi-infected ticks in Canada. J. Med. Entomol 38, 493–500. [DOI] [PubMed] [Google Scholar]
  48. Smith RJ, Hatch MI, Cimprich DA, Moore FR, 2020. Gray Catbird (Dumetella carolinensis), version 1.0. In Birds of the World (Poole AF, Editor). Cornell Lab of Ornithology, Ithaca, NY, USA. 10.2173/bow.grycat.01 Accessed 4 December 2020. [DOI] [Google Scholar]
  49. Smith RP Jr., Rand PW, Lacombe EH, Morris SR, Holmes DW, Caporal DA, 1996. Role of bird migration in the long-distance dispersal of Ixodes dammini, the vector of Lyme disease. J. Infect. Dis 174, 221–224. [DOI] [PubMed] [Google Scholar]
  50. States SL, Huang CI, Davis S, Tufts DM, Diuk-Wasser MA, 2017. Co-feeding transmission facilitates strain coexistence in Borrelia burgdorferi, the Lyme disease agent. Epidemics 19, 33–42. [DOI] [PMC free article] [PubMed] [Google Scholar]
  51. Stromdahl E, Hamer S, Jenkins S, Sloan L, Williamson P, Foster E, Nadolny R, Elkins C, Vince M, Pritt B, 2014. Comparison of phenology and pathogen prevalence, including infection with the Ehrlichia muris-like (EML) agent, of Ixodes scapularis removed from soldiers in the midwestern and the northeastern United States over a 15 year period (1997–2012). Parasit Vectors 7:553. [DOI] [PMC free article] [PubMed] [Google Scholar]
  52. Tsao K, Bent SJ, Fish D, 2013. Identification of Borrelia burgdorferi ospC genotypes in host tissue and feeding ticks by terminal restriction fragment length polymorphisms. Appl. Environ. Microbiol 79, 958–964 [DOI] [PMC free article] [PubMed] [Google Scholar]
  53. Vanderhoff N, Pyle P, Patten MA, Sallabanks R, James FC, 2020. American Robin (Turdus migratorius), version 1.0. In Birds of the World (Rodewald PG, Editor). Cornell Lab of Ornithology, Ithaca, NY, USA. 10.2173/bow.amerob.01 Accessed 4 December 2020. [DOI] [Google Scholar]
  54. Williamson PC, Billingsley PM, Teltow GJ, Seals JP, Turnbough MA, Atkinson SF, 2010. Borrelia, Ehrlichia, and Rickettsia spp. in ticks removed from persons, Texas, USA. Emerg. Infect. Dis 16, 441–446. [DOI] [PMC free article] [PubMed] [Google Scholar]
  55. Wright CL, Hynes WL, White BT, Marshall MN, Gaff HD, Gauthier DT, 2014. Single-tube real-time PCR assay for differentiation of Ixodes affinis and Ixodes scapularis. Ticks Tick Borne Dis. 5, 48–52. [DOI] [PMC free article] [PubMed] [Google Scholar]
  56. Zahler M, Gothe R, Rinder H, 1995. Genetic evidence against a morphologically suggestive conspecificity of Dermacentor reticulatus and D. marginatus (Acari: Ixodidae). Int. J. Parasitol 25, 1413–1419. [DOI] [PubMed] [Google Scholar]

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