Ectoparasites of road-killed vertebrates in northwestern South Carolina, USA

Veterinary Parasitology 129 (2005) 313–322 www.elsevier.com/locate/vetpar

Ectoparasites of road-killed vertebrates in northwestern South Carolina, USA Mark P. Nelder *, Will K. Reeves 1 114 Long Hall, Department of Entomology, Soils, and Plant Sciences, Clemson University, Clemson, SC 29634-0315, USA Accepted 1 February 2005

Abstract Road-killed animals are overlooked as the source of ectoparasites for monitoring vectors of zoonotic pathogens. We demonstrate that by exclusively sampling road-killed animals, a wide spectrum of vertebrate hosts and ectoparasites can be collected. Fifty-one species of ectoparasites were recovered from 35 species of road-killed vertebrates in northwestern South Carolina. Approximately, 11% of the total known terrestrial vertebrate species in the region were examined, which included more than 25% of the known mammal species. Our sampling techniques produced new state and regional records for chewing lice, ticks, and parasitic mites. Most ectoparasites were alive when they were collected, which would allow them to be screened for zoonotic pathogens. # 2005 Elsevier B.V. All rights reserved. Keywords: Distribution; Ectoparasitic arthropods; Surveillance; Vector; Zoonoses

1. Introduction Ectoparasite surveys in South Carolina typically have been restricted to a single host taxon (e.g. rodents, Clark et al., 1998) or ectoparasite (e.g. ticks (Ixodida), (Williams et al., 1999)). Ticks (Williams et al., 1999; Reeves et al., 2002) and fleas (Siphonaptera) (Durden et al., 1999) are beginning * Corresponding author. Tel.: +1 864 656 5070; fax: +1 864 656 5069. E-mail address: [email protected] (M.P. Nelder). 1 Present address: Centers for Disease Control and Prevention, Viral and Rickettsial Zoonoses Branch, Mailstop G-13, 1600 Clifton Road, NE, Atlanta, GA 30333, USA.

to be thoroughly documented in South Carolina, while lice (Phthiraptera) and mites (Acari) remain understudied (Reeves et al., 2004). Our knowledge of potential arthropod vectors in South Carolina and reports of ectoparasite faunas encompassing a wide diversity of host species are limited. Ectoparasites may be vectors of zoonotic pathogens that cause diseases, such as Lyme disease, Powassan encephalitis, plague, Rocky Mountain spotted fever, and tularemia. In clinical cases, the source of infection is often unknown because arthropod vectors are overlooked. Ectoparasite-borne infectious diseases and parasites have caused significant morbidity and mortality in South Carolina (Adler and Wills, 2003).

0304-4017/$ – see front matter # 2005 Elsevier B.V. All rights reserved. doi:10.1016/j.vetpar.2004.02.029

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Some ectoparasitic arthropods are irritating pests of humans and domestic animals, regardless of their significance as vectors of disease. The specific hosts of ectoparasites can maintain enzootic pathogens or infect bridge vectors. While some ectoparasites, such as polyplacid lice or bird mites, are host specific and will not feed on humans, these arthropods can transmit pathogens to their natural hosts and maintain zoonotic foci. Other less host-specific ectoparasites, such as some fleas or ticks then become infected and transmit pathogens to humans. To fully understand and control zoonotic diseases, ectoparasitic arthropods and hosts should be accounted for. Literature searches, trapping hosts or ectoparasites, investigation of museum records or specimens, collections by veterinarians, and taxidermists’ animal skins are methods commonly used to determine the ectoparasite fauna of an area (e.g. Coyner et al., 1996; Clark et al., 1998; Durden et al., 1999; Williams et al., 1999; Reeves et al., 2002). The use of road-killed animals for determining ectoparasite faunas has been well documented but under-used. For instance, roadkilled cottontail rabbits (Wassel et al., 1980; Baird, 1983), coyotes (Foster et al., 2003), fox squirrels (Coyner et al., 1996), ground squirrels (Galloway and Christie, 1990), muskrats (Bauer and Whitaker, 1981), Virginia opossums (Hopkins, 1980), and various other mammals (Whitaker and Goff, 1979; Lavender and Oliver, 1996) have been used in conjunction with traditional methods to detect ectoparasites. Understanding the host associations and distributions of ectoparasites of animals is fundamental to increasing our knowledge of the ecology of arthropodborne pathogens. Here, we document new geographical and host records for ectoparasites of vertebrates of northwestern South Carolina by sampling road-killed animals.

2. Materials and methods 2.1. Study area Our study area included portions of upstate South Carolina (Anderson, Oconee, and Pickens Counties). We collected in the upper piedmont ecoregion and did not collect in the Blue Ridge Mountain ecoregion. The piedmont consists of rolling hills and is a mosaic of

deciduous forests, grasslands, wetlands, and urban environments. The elevation ranges from approximately 180–370 m above sea level. The upstate of South Carolina, one of the most highly populated regions of the state, harbors approximately 314 species of terrestrial vertebrates: 38 amphibians, 170 birds, 62 mammals, and 44 reptiles (King, 1979; Bull and Farrrand, 1994; Whitaker, 1996). 2.2. Collection of ectoparasites Road-killed animal carcasses were examined for ectoparasites. Not all carcasses were appropriate for recovering ectoparasites; unidentifiable carcasses and those infested with maggots (Diptera: Calliphoridae, Sarcophagidae, or Muscidae) were not examined. Carcasses usually had maggots on them by 24–48 h after death. To restrict our study to road-killed animals as hosts, we examined only those carcasses showing signs of animal–vehicle collisions (i.e., crushed skulls or broken limbs). We collected carcasses between 1 April 2004 and 15 July 2004, but also included were eight prior collections. Carcasses were either reported to us by colleagues or spotted by the authors and retrieved immediately. While collecting carcasses and their ectoparasites, two pairs of nitrile gloves were worn to prevent possible infection by disease agents. Carcasses were placed in plastic garbage bags and transported to the laboratory for freezing (at 20 8C) or immediate inspection. Frozen specimens were allowed to thaw for approximately 30 min at room temperature (i.e., 22–24 8C) before inspection. Fleas were obtained by combing the fur with a flea comb, whereas other ectoparasites (e.g. chiggers, lice, ticks) were collected using forceps while separating the fur or feathers. Liberated fur was examined under a dissecting microscope for the presence of ectoparasites. Bags that contained carcasses were rinsed with 80% ethanol and the wash examined under a dissecting microscope for ectoparasites that might have emigrated or fallen off the carcass during transport or freezing. Birds were examined in the field and ectoparasites placed in labeled vials containing 80% ethanol. All ectoparasites were fixed in 80% ethanol or cleared in hot 85% lactic acid and slide mounted in Euparal or Hoyer’s medium. Occurrence of an ectoparasite is reported as prevalence for each host on which it was found. Prevalence is defined as the

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number of individuals of a host species parasitized by a particular ectoparasite divided by the total number of individuals of a host species examined. Identification of ectoparasites followed the keys provided by Price et al. (2003) (chewing lice); Gaud and Atyeo (1996) (feather mites); Benton (1983) (fleas); Maa (1966, 1969), Blanton and Wirth (1979), Sabrosky (1986) (flies); Wharton and Fuller (1952), Whitaker and Wilson (1974), Krantz (1978), Strandmann and Wharton (1958) (mites); Kim et al. (1986) (sucking lice); and Clifford et al. (1961) and Keirans and Litwak (1989) (ticks). Voucher specimens of ectoparasites were deposited in the Clemson University Arthropod Collection.

3. Results Fifty-one species of ectoparasites were found on 96 road-killed animals in northwestern South Carolina (Table 1). Ectoparasites were recovered from 68 of the 96 carcasses. Thirty-five species of road-killed vertebrates were examined, representing seven amphibians and reptiles, 12 birds, and 16 mammals. Roadkilled animals represented 11% of the total known terrestrial vertebrate diversity in the region. More than 25% of the area’s mammal species was represented in our samples. Some groups, such as salamanders and songbirds, are not prone to be near roads or were too small to be noticed during sampling even if they were killed. Domestic animals (e.g. cattle, horses, chickens, pigs), bats, some small mammals, and bears were not found as road-kills. Host species yielding a rich fauna of ectoparasites included the Virginia opossum, Didelphis virginiana Kerr (eight ectoparasite species); the raccoon, Procyon lotor (Linnaeus) (seven); the brown thrasher, Toxostoma rufum (Linnaeus) (seven); and the eastern gray squirrel, Sciurus carolinensis Gmelin (five) (Table 1). Tramp ectoparasites (i.e., defined here as those found on three or more hosts) and their number of host species were: Dermacentor variablilis (Say) (six); Orchopeas howardi (Baker) (four); Ctenocephalides felis (Bouche´ ) (three); Neotromicula whartoni (Ewing) (three); and Ornithonyssus sylviarum (Canestrini and Fanzago) (three). Reptiles and amphibians yielded few ectoparasites. The black rat snake, Elaphe obsoleta (Say), which

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yielded Microtrombicula trisetica (Loomis and Crossley), was the only reptile or amphibian found to harbor an ectoparasite (Table 1). Hosts on which we did not observe ectoparasites are footnoted in Table 1.

4. Discussion Our use of road-killed animals to detect ectoparasites yielded 51 species; however, further work is needed to assess emigration of ectoparasites from carcasses so that ectoparasite-infestation levels can be quantified. Using mostly road-killed cottontail rabbits (108 of 131 hosts), Wassel et al. (1980) found 21 species of ectoparasites. Their findings, coupled with ours, point to road-kills as a rich source of ectoparasites that should be used more often in ectoparasite surveys. One disadvantage is the emigration of ectoparasites from the carcass; mesostigmatid mites (Mesostigmatidae) and chiggers (Prostigmatidae) are the earliest to leave a carcass (<5 h) (Westrom and Yescott, 1975). Fleas also leave the carcass soon after host death (Cole and Koepke, 1947; Jameson, 1947; Gross and Bonnet, 1949; Stark and Kinney, 1962). The number of ectoparasites collected on a dead animal can be considered only a proportion of its living total (Colbo and MacLeod, 1976). Although fleas, mesostigmatid mites, and chiggers leave carcasses soon after host death, we found four, six, and five species of each, respectively. Polgenis gwyni (Fox) is commonly found on the cotton rat, Sigmodon hispidus Say and Ord (Durden et al., 2000), and other rodents (Clark and Durden, 2002). This finding is the first report of this flea infesting the Virginia opossum in South Carolina; however, P. gwyni infests Virginia opossums elsewhere in the southeastern U.S. (Lewis and Lewis, 1994). P. gwyni is a vector of Rickettsia typhi (Wolbach and Todd), the causative agent of murine typhus (Azad, 1990). We report C. felis for the first in time in South Carolina from the domestic cat, Felis silvestris Schreber, and the raccoon. C. felis is a vector or intermediate host of Acanthocheilonema reconditum (Grassi) (a causative agent of canine filariasis), Bartonella henselae Brenner, O’Connor, Winkler, and Steigerwalt (causative agent of cat scratch disease) (Chomel et al., 1996), other Bartonella spp. (Rolain et al., 2003), Dipylidium

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Table 1 Ectoparasites of road-killed animals from northwestern South Carolina (between 16 December 1994 and 15 July 2004) Ectoparasite Flies (Diptera) Biting midges (Ceratopogonidae) Culicoides snowi Wirth and Jones

Bot flies (Oestridae) Cuterebra sp. (abdominalis or buccata)

Keds (Hippoboscidae) Lipotena mazamae Rondani

Ornithoica vicina Walker

Fleas (Siphonaptera) Ctenocephalides felis (Bouche´ )

Host (ectoparasite prevalence)

Collection Dataa,b

Sciurus carolinensis Gmelin (1/15)

1.AN,15.IV.04,MPN,A 2.AN,1.V.04,MPN,A 3.PI,7.V.04,MPN,A 4.AN,10.V.04,WKR,A 5 PI,13.V.04,MAM,A 6.AN,15.V.04,MPN,A 7.PI,16.V.04,WKR,A 8.PI,16.V.04,WKR,P 9.OC,21.V.04,WKR,A 10.AN,16.VI.04,MPN,A 11.OC,18.VI.04,WKR,A 12.AN,22.VI.04,MPN,A 13.PI,22.VI.04,WKR,A 14.AN,30.VI.04,MPN,A 15.PI,8.VII.04,WKR,A

Sylvilagus floridianus (Allen) (1/3)

1.OC,2.VI.04,WKR,A 2.PI,21.IX.03,WKR,P 3.AN,14.VII.04,MPN,A

Odocoileus virginianus (Zimmermann) (2/3)

1.AB,15.IV.04,KDC,A 2.AN,1.VI.03,WKR,P 3.PI,8.VI.04,MPN/WKR,P 1.PI,25.VI.04,WKR/MPN,P 1.PI,13.VI.04,WKR,P 2.PI,19.VI.04,WKR,A 3.PI,2.VII.04,MPN,A

Strix varia Barton (1/1) Toxostoma rufum (Linnaeus) (1/3)

Didelphis virginiana Kerr (5/7)

Felis silvestris Schreber (2/3) Procyon lotor (Linnaeus) (3/10)

1.PI,29.II.03,WKR,P 2.PI,26.V.04,MPN,P 3.OC,27.V.04,WKR,A 4.PI,9.VI.04,WKR,P 5.PI,13.VI.04,MPN,P 6.PI,22.VI.04,MPN,P 7.PI.13.VII.04,MPN,A 1.PI,15.IX.03,MAM,P 2.PI,13.VI.04,WKR,P 3.AN,30.VI.04,MPN,A 1.AN,27.IV.04,MPN,A 2.OC,21.V.04,MPN,A 3.OC,27.V.04,WKR,A 4.OC,8.VI.04,MPN,P 5.PI,11.VI.04,MPN,A 6.AN,2.VII.04,MPN,P 7.PI.5.VII.04,SKK,A 8.PI,5.VII.04,SKK,A 9.OC,8.VII.04,WKR,A 10.OC.15.VII.04,WKR,P

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Table 1 (Continued ) Ectoparasite Orchopeas howardi (Baker)

Host (ectoparasite prevalence)

Collection Dataa,b

Didelphis virginiana (1/7) Tamias striatus (Linnaeus) (1/4)

1.A 2.P 3.A 4.A 5.A 6.A 7.A 1.AN,8.II.03,WKR,P 2.OC,9.V.04,WKR,A 3.PI.2.VII.04,WKR,A 4.PI.15.VI.04,HEC,A 1.P 2.A 3.A 4.A 5.A 6.P 7.A 8.P 9.A 10.A 1.P 2.P 3.P 4.P 5.P 6.A 7.P 8.A 9.P 10.P 11.A 12.A 13.A 14.A 15.A 1.A 2.A 3.P 4.A 5.A 6.A 7.A 1.PI,16.VI.04,MPN,P

Procyon lotor (3/10) Sciurus carolinensis (8/15) Polygenis gwyni (Fox) Pulex simulans Baker Lice (Phthiraptera) Sucking lice (Anoplura) Enderleinellus longiceps Kellogg and Ferris Enderleinellus marmotae Ferris

Didelphis virginiana (1/7) Vulpes vulpes (Desmarest) (1/1)

Sciurus carolinensis (1/15) Marmota monax (Linnaeus) (2/7)

Hoplopleura sciuricola Ferris

Sciurus carolinensis (2/15)

Neohaematopinus sciuri Jancke

Sciurus carolinensis (6/15)

Chewing lice (Mallophaga) Brueelia dorsale Williams Chelopistes meleagridis (L.) Columbicola columbae (L.) Degeeriella fulva (Geibel) Menacanthus eurysternus (Burm)

Toxostoma rufum (Linnaeus) (2/3) Meleagris gallopavo Linnaeus (1/1) Columba livia Gmelin (1/1) Buteo jamaciensis (Gmelin) (1/1) Turdus migratorius Linnaeus (1/2)

Neotrichodectes mephiditis (Packard)

Mephitis mephitis (Schreber) (2/2)

Oxylipeurus corpulentus Clay Penenirmus auritus (Scopoli) Picicola marginatulus (Harrison) Stachiella octomaculatus (Paine) Stachiella larseni Emerson

Meleagris gallopavo (1/1) Dryocopus pileatus (Linnaeus) (1/1) Dryocopus pileatus (1/1) Procyon lotor (9/10) Mustela vison Schreber (2/2)

Mites (Acari) Astigmata Asiochirus blarinae Fain and Hyland Falculifer rostratus (Buchholz) Gycyphagus hypudaei Koch Listophorus dozieri Radford Schizocarpus indianensis Fain, Whitaker, and Smith

Analgidae sp. 1 Analgidae sp. 2 Syringobiidae sp. Vexillaridae sp.

1.A 2.P 3.A 4.A 5.A 6.A 7.A 8.A 9.A 10.A 11.A 12.A 13.A 14.A 15.A 1.PI,4.V.04,MPN,A 2.OC,5.V.04,WKR/MPN,A 3.PI,11.V.04,MPN,P 4.PI,19.V.04,MPN,A 5.AN,11.VI.04,MPN,P 6.AN,17.VI.04,MPN,A 7.PI,20.VI.04,MPN,A 1.A 2.A 3.A 4.A 5.A 6.A 7.A 8.A 9.A 10.A 11.P 12.P 13.A 14.A 15.A 1.A 2.P 3.P 4.P 5.P 6.P 7.A 8.A 9.P 10.A 11.A 12.A 13.A 14.A 15.A 1.A 2.Pc 3.P 1.OC,9.V.04,WKR,Pc 1.PI,28.VI.04,WKR,Pc 1.AN,12.II.03,SMM,Pc 1.PI,21.VI.04,WKR,A 2.PI,19.VI.04,WKR,P 1.AN,20.II.03,WKR,P 2.PI,22.V.04,WKR,P 1.Pc 1.AN,3.V.04.MPN/WKR,Pc 1.Pc 1.Pc 2.P 3.P 4.P 5.P 6.P 7.P 8.P 9.P 10.A 1.OC,16.VI.04,MPN,Pc 2.OC,16.VI.04,WKR,P

Blarina brevicauda (Say) (1/1) Columba livia (1/1) Didelphis virginiana (1/7) Didelphis virginiana (2/7) Castor canadensis Kuhl (1/3)

1.PI,14.XII.03,KDC,P 1.P 1.A 2.P 3.A 4.A 5.A 6.A 7.A 1.A 2.P 3.A 4.A 5.A 6.P 7.A 1.PI,3.I.03,WKR,A

Mimus polyglottos (Linnaeus) (1/1) Toxostoma rufum (1/3) Charadrius vociferous Linnaeus (1/1) Parus carolinensis Audubon (1/1)

2.PI,10.II.04,WKR,P 3.PI,17.VI.04,WKR/MPN,A 1.AN,11.VI.04,MPN,P 1.P 2.A 3.A 1.PI,29.VI.04,WKR/MPN,P 1.OC,6.VI.04,WKR,P

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Table 1 (Continued ) Ectoparasite Mesostigmata Androlaelaps fahrenholzi (Berlese)

Dermanyssus gallinae (DeGeer) Haemolaelaps glasgowi (Ewing) Ornithonyssus bacoti (Hirst) Ornithonyssus sylviarum (Canestrini and Fanzago)

Ornithonyssus wernecki (Fonseca) Prostigmata Archaemyobia inexpectatus Jameson Euschoengastia blarinae (Ewing) Microtrombicula trisetica (Loomis and Crossley)

Miyatrombicula cynos (Ewing) Neotrombicula whartoni (Ewing)

Ticks (Ixodida) Amblyomma americanum (L.)

Amblyomma maculatum Koch Dermacentor variabilis (Say)

Haemaphysalis leporispalustris (Packard) Ixodes affinis Neumann Ixodes brunneus Koch Ixodes cookei Packard

Ixodes texanus Banks a

Host (ectoparasite prevalence)

Collection Dataa,b

Castor canadensis (1/2) Sciurus carolinensis (1/15) Columba livia (1/1) Toxostoma rufum (1/3) Tamias striatus (1/4) Mus musculus Linnaeus (1/1) Didelphis virginiana (1/7) Turdus migratorius (1/2)

1.P 2.A 3.A 1.A 2.A 3.A 4.A 5.P 6.A 7.A 8.A 9.A 10.A 11.A 12.A 13.A 14.A 15.A 1.P 1.P 2.A 3.P 1.A 2.P 3.A 4.A 1.OC.15.VII.04,CEB,P 1.A 2.A 3.A 4.A 5.A 6.A 7.P 1.P 2.A

Strix varia (1/1) Toxostoma rufum (1/3) Didelphis virginiana (2/7)

1.P 1.P 2.A 3.A 1.A 2.P 3.A 4.A 5.P 6.A 7.A

Didelphis virginiana (2/7) Blarina brevicauda (Say) (1/1) Elaphe obsoleta (Erdnatter) (1/4)

1.A 2.P 3.A 4.A 5.A 6.P 7.A 1.PI,14.XII.03,KDC,P 1.AN,28.V.04,MPN,A 2.PI,8.VI.04,MPN,Pc 3.OC,10.VI.04,WKR,A 4.AN,25.VI.04,WKR,A 1.OC,16.XII.94,RAM,P 1.P 2.A 3.A 1.A 2.A 3.A 4.P 5.A 6.A 7.A 8.A 9.A 10.A 1.P 2.A 3.A

Glaucomys volans (Linnaeus) (1/1) Castor canadensis (1/3) Procyon lotor (1/10) Sylvilagus floridianus (1/3) Meleagris gallopavo (1/1) Odocoileus virginianus (1/3) Procyon lotor (1/10) Canis latrans Say (1/1) Canis lupus Linnaeus (1/2) Canis latrans Say (1/1) Didelphis virginiana (4/7) Marmota monax (2/7) Procyon lotor (6/10) Vulpes vulpes (1/1) Toxostoma rufum (1/3) Sylvilagus floridianus (1/3) Corvus brachyrhynchos Brehm (1/1) Mephitis mephitis (1/2) Procyon lotor (2/10) Vulpes vulpes (1/1) Procyon lotor (6/10)

1.P 1.P 2.A 3.A 1.A 2.A 3.A 4.A 5.A 6.A 7.A 8.A 9.P 10.A 1.AN,5.VII.04,MPN/WKR,P 1.AN,30.VI.04,WKR,P 2.PI.14.VII.04,MPN 1.P 1.A 2.P 3.P 4.P 5.P 6.A 7.A 1.A 2.A 3.A 4.A 5.P 6.P 7.A 1.P 2.A 3.P 4.A 5.P 6.P 7.A 8.P 9.P 10.A 1.P 1.P 2.A 3.A 1.P 2.A 3.A 1.PI,1.III.04,CSD,P 1.P 2.A 1.P 2.A 3.A 4.A 5.A 6.A 7.A 8.A 9.P 10.A 1.P 1.P 2.P 3.A 4.A 5.P 6.P 7.A 8.P 9.P 10.A

For a given road-kill, data are reported as follows: Specimen Number; location: AB = Abbeville County, AN = Anderson Co., PI = Pickens Co., and OC = Oconee Co.; date: day, month, year (‘‘0’’s refer to the first decade of the 21st century); collector; presence (P) or absence (A) of ectoparasite on specimen. Collection data are given only the first time a host taxon is mentioned in the table. b Road-kills with no ectoparasites were Butroides striatus (L.) (1.PI.15.VI.04,WKR), Elaphe guttata (L.) (1.PI,14.V.04,MPN), Eumeces fasciatus (L.) (1.PI,30.VI.04,WKR), Rana catesbeina Shaw (1.AN,22.VI.04,WKR/MAM), Rana sphenocephala Cope (1.PI,20.VI.04,MPN,2– 5.PI, 5.VII.04,MPN), Stenotherus odoratus (Latreille) (1.PI,16.VI.04,WKR), and Terrepene carolina (L.) (1.PI,14.VI.04,LWB). c State record of this ectoparasite.

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caninum (Linnaeus) (double-pored tapeworm infection) (Durden and Traub, 2002), Rickettsia felis La Scola, Meconi, Fenollar, Rolain, Roux, and Raoult (murine typhus-like disease), and R. typhi (Azad et al., 1997). We report O. howardi from the chipmunk, Tamias straitus (Linnaeus), Virginia opossum, and raccoon for the first time in South Carolina. O. howardi is a common flea of the eastern grey squirrel and other arboreal mammals (Wilson, 1978). This flea is a laboratory and natural vector of Rickettsia prowazekii da Rocha-Lima, the causative agent of sylvatic epidemic typhus (Sonenshine et al., 1978; McDade, 1987). Pulex simulans Baker is reported for the first time on red fox, Vulpes vulpes (Linnaeus), from South Carolina; it is a common flea of carnivores throughout the southeastern U.S. (Durden and Kollars, 1997). Because the latter three species of fleas were found on the Virginia opossum, this host might be capable of maintaining pathogens, acting as a potential reservoir of zoonotic pathogens. The Virginia opossum could be a bridge reservoir of zoonoses between alternative hosts of these fleas. Our discovery of Ixodes texanus Banks on raccoons is significant because this tick was previously reported only from the coastal plain of South Carolina (Durden et al., 1999). This tick is a vector of Rickettsia rickettsii (Wolbach), the causative agent of Rocky Mountain spotted fever (Sonenshine et al., 2002). We extend the known range of Ixodes affinis Neumann from the coastal plains and sandhills of South Carolina (Durden and Keirans, 1996; Durden et al., 1999) to include the piedmont ecoregion. The eastern cottontail rabbit, Sylvilagus floridianus (Allen), was a new host record for this tick. I. affinis harbors Borrelia burgdorferi Johnson, Schmid, Hyde, Steigerwalt, and Brenner sensu lato, the causative agent of Lyme disease (Clark et al., 1998); however, its competence as a vector of this spirochete is unknown. This tick was found on rodents from the coastal plain of South Carolina (Clark et al., 1998). Amblyomma americanum (Linnaeus) was recorded from the raccoon, turkey, and white-tailed deer during our study. A. americanum is a vector of Ehrlichia chaffeensis Anderson, Dawson, Jones, and Wilson (etiologic agent of human monocytic ehrlichiosis) (Lockhart et al., 1995), Ehrlichia ewingi Anderson, Greene, Jones, and Dawson (canine ehrlichiosis), Francisella tularensis McCoy and Chapin (tularemia), and R. rickettsii

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(Burgdorfer et al., 1975). Amblyomma maculatum was removed from a coyote in this study. This tick is a vector of the protozoan Hepatozoon americanum Vincent-Johnson, Macintire, Lindsay, Lenz, Baneth and Shkap, which can kill domestic dogs (Mathew et al., 1998; Ewing et al., 2002). Of all ticks collected from road-killed animals, D. variabilis was the most numerous found on several canine species, raccoons, Virginia opossums, and groundhogs. In the eastern U.S., D. variabilis is the primary vector of R. rickettsii (Strickland et al., 1976). Road-killed animals, especially the raccoons in our study, appeared to attract questing D. variabilis. Both B. burgdorferi and St. Louis encephalitis virus have been isolated from D. variabilis (McLean et al., 1985); however, the tick’s role in the transmission of these disease agents is not known. Haemaphysalis leporispalustris (Packard), found on a brown thrasher, might be involved with the enzootic cycling of Rocky Mountain spotted fever, tularemia, and Q fever (Strickland et al., 1976). Ixodes brunneus Koch was found on an American crow, Corvus brachyrhynchos Brehm, and is known to harbor R. rickettsii (Clifford et al., 1969). Ixodes cookei Packard is reported here for the first time for South Carolina from the skunk, Mephitis mephitis (Schreber), and red fox. I. cookei is a vector of Powassan encephalitis virus (Berge, 1975). Although the vector potentials of the flies, lice, and mites found during our study are unknown, many are noteworthy. The bird mesostigmatid-mites Dermanyssus gallinae (DeGeer) and O. sylviarum were found on several species of birds. These mites are known to harbor and mechanically transmit St. Louis encephalitis virus and western equine encephalitis virus (Mullen and O’Connor, 2002). These two mites are possible bridge vectors because they parasitize a wide range of birds and will bite humans. We suggest that West Nile virus should be sought in these mites, as they might maintain this arbovirus in wild animals. We collected the tropical rat mite, Ornithonyssus bacoti (Hirst) (Macronyssidae), a potentially significant ectoparasite because it will bite humans and infests other domestic animals, including rodents, dogs, cats, and birds. O. bacoti is a vector of filarial nematodes to rodents and Hantann virus, and is a vector of Rickettsia akari Huebner, Jellison, and Pomerantz (Rickettsialpox) and Yersinia pestis (plague) in the laboratory (Yunker, 1964; Mullen and O’Connor, 2002).

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We report the presence of the chigger M. trisetica for the first time in South Carolina on a black rat snake, E. obsoleta. M. trisetica was previously not reported in South Carolina (Reeves et al., 2004). Chiggers are more common on animals in cooler months (Whitaker and Loomis, 1978); this might explain the low species richness of this group during our warm-weather study. Most of the chewing lice of birds reported here are state records for South Carolina because little work has been done on this group. Except for scattered reports (Peters, 1933, 1936), these results are the first report of bird lice for northwestern South Carolina. The majority of ectoparasites collected during this study were collected alive. We propose the use of such ectoparasites to screen for animal and human pathogens. For example ticks might be screened for pathogens such as Rickettsia spp. and Borrelia spp. The collection of ectoparasites and screening for pathogens could lead to the discovery of emerging infectious agents of both animals and humans.

5. Conclusion Our use of road-killed animals has several advantages over conventional methods (e.g. live trapping) of host assays for ectoparasites. First, the method is less time consuming because it does not require deployment of traps and their routine inspection. Second, animals do not have to be euthanized or anesthetized, preventing the chance bite or scratch from a wild and possibly infectious animal. Third, traditional methods usually only target a restricted group of animals (e.g. Sherman1 traps for small rodents). In contrast, our method allows for a wide variety of hosts to be inspected. Research involving a specific pathogen or ectoparasite might best be served by a targeted trapping of certain host species. Taken together, the advantages make this method economically and logistically feasible. Because only a small proportion of the total potential host species (11%) was inspected over a brief period (summer), 51 species of ectoparasites for the area is only a fraction of the total that exists. Further use of conventional methods of ectoparasite surveillance, accompanied by road-kill surveys, should provide a better understanding of the ectoparasitic arthropods in this and other regions of the world.

Acknowledgements We thank the following people for giving us the location of carcasses: Charles Beard, Karen Burton, Layla Burgess, Hugh Conway, Kristin Cobb, Candis Duncan, Sam-Kyu Kim, James Korecki, Monica MacCarroll, Stanlee Miller, Robert Moss, and Eric Paysen. We also thank Lance A. Durden and William J. Wrenn for help in the identification of some of our ectoparasites. We thank Peter H. Adler and Alfred G. Wheeler, Jr. for reviewing an earlier version of the manuscript. This is technical contribution number 5040 of the South Carolina Agriculture and Forestry Research System, Clemson University.

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