Regular issues
Special Issues

Southeastern Naturalist
    SENA Home
    Range and Scope
    Board of Editors
    Editorial Workflow
    Publication Charges

Other EH Journals
    Northeastern Naturalist
    Caribbean Naturalist
    Urban Naturalist
    Eastern Paleontologist
    Eastern Biologist
    Journal of the North Atlantic

EH Natural History Home

Helminth Parasites of Pit Vipers from North Carolina
Elijah Davis, Jeffrey C. Beane, and James R. Flowers

Southeastern Naturalist, Volume 15, Issue 4 (2016): 729–741

Full-text pdf (Accessible only to subscribers.To subscribe click here.)


Site by Bennett Web & Design Co.
Southeastern Naturalist 729 E. Davis, J.C. Beane, and J.R. Flowers 22001166 SOUTHEASTERN NATURALIST 1V5o(4l.) :1752,9 N–7o4. 14 Helminth Parasites of Pit Vipers from North Carolina Elijah Davis1, Jeffrey C. Beane2, and James R. Flowers1,* Abstract – We surveyed for helminth parasites salvaged specimens of 27 Agkistrodon contortrix (Copperhead), 4 Agkistrodon piscivorus (Cottonmouth), and 7 Crotalus horridus (Timber Rattlesnake) collected between 2003 and 2010 from various locations in North Carolina. We detected 10 previously described helminths (2 trematodes: Ochetosoma kansensis, Travtrema stenocotyle; 1 cestode: Proteocephalus sp.; 6 nematodes: Kalicephalus inermis coronellae, Kalicephalus costatus parvus, Physalopterid larvae, Physaloptera squamatae, Capillaria colubra, Strongyloides serpentis; and 1 acanthocephalan: Macracanthorhynchid cystacanths). Herein, we report 7 new host records and 7 new geographic-distribution records with notes on host–parasite biology. Introduction North Carolina contains 4 physiographic provinces (Mountains, Piedmont, Sandhills, Coastal Plains) that support at least 71 reptile species, including 37 snake species (Palmer and Braswell 1995). Yet, helminths from only 5 snake species from North Carolina have been reported: Nerodia sipedon (L.) (Northern Water Snake), Nerodia erythrogaster (Forster in Bossu) (Plain-bellied Water Snake), Nerodia taxispilota (Holbrook) (Brown Water Snake), Agkistrodon piscivorus (Lacépède) (Cottonmouth), and Coluber constrictor L. (Racer). Primary reports include a single helminth survey by Collins (1968, 1969) of 4 North Carolina snakes and 3 additional studies, each listing a single ophidian helminth species from North Carolina (Brooks 1979, Richardson and Nickol 1995, Sprent 1988). We queried the databases of the US National Parasite Collection (USNPC 2015), and the Harold W. Manter Laboratory of Parasitology (HWML 2016) for deposited North Carolina ophidian helminths. Of the 33 lots of North Carolina ophidian helminths deposited in the USNPC, only 2 (#11829 “Cestode” from “Water Moccasin” from Wilmington, and #97949 Ochetosoma kansense from Cottonmouth from Bertie County) were from viperids, and none of the 5 lots of North Carolina snake helminths deposited in the HWML were from viperids. In an effort to increase understanding of the helminths of North Carolina snakes, we have conducted helminth surveys of salvage snakes. To that end, we have examined 27 Agkistrodon contortrix (L.) (Copperheads), 4 Cottonmouths, and 7 Crotalus horridus L. (Timber Rattlesnakes) from various counties in North Carolina for helminths and report their helminth fauna herein. Notes on the life cycle and host–parasite relationship of each parasite are included. 1Department of Population Health and Pathobiology, College of Veterinary Medicine, North Carolina State University, 1060 William Moore Drive, Raleigh, NC 27607. 2North Carolina State Museum of Natural Sciences, 1626 Mail Service Center, Raleigh, NC 27699-1626. *Corresponding author - Manuscript Editor: Jeff Lauren Southeastern Naturalist E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 730 Methods During general faunal surveys for the North Carolina State Museum of Natural Sciences, J.C. Beane has utilized salvaged snakes (mostly road-killed) to provide morphological, biological, ecological, and locale data for North Carolina herpetofauna. Since 2003, many of these salvaged specimens have been collected, stored, and donated to the parasitology group of the College of Veterinary Medicine at North Carolina State University, Raleigh, NC. Host specimens were stored frozen until just prior to necropsy. At necropsy, we isolated, opened, and examined various organs (mouth, esophagus, stomach, intestine, liver, pancreas, lungs, and kidneys) from the carcass and thoroughly washed the tissues in 0.9% saline. Using a dissecting microscope, we collected helminths directly from the organs or from the sediment of saline washings. We employed standard parasitological procedures (Pritchard and Kruse 1982) to fix and process helminth specimens. We used Semichon’s carmine to stain trematodes, cestodes, and acanthocephalans, and cleared nematodes with glycerin. We deposited representative parasites in the Harold W. Manter Laboratory of Parasitology (HWML), University of Nebraska-Lincoln, Lincoln, NE. Taxonomy and common names of snake hosts follow Conant and Collins (1998) and Palmer and Braswell (1995); species authorities follow the Integrated Taxonomic Information System (2016). Results and Discussion Twenty-one of 38 (55%) viperids examined harbored helminths, including 14 of 27 (52%) Copperheads, 4 of 4 (100%) Cottonmouths, and 3 of 7 (43%) Timber Rattlesnakes. We report 10 previously described helminths (2 trematodes, 1 cestode, 6 nematodes, and 1 acanthocephalan) and include the mean intensity (Bush et al. 1997) and range of each helminth infection, the number of snakes infected, and the counties of host collection (Tables 1, 2). In addition to the specimens included in the tables, 1 Cottonmouth from Moore County and 3 Timber Rattlesnakes (1 each from Bertie, Hoke, and Moore counties) were collected but were too damaged to necropsy. Trematoda Ochetosoma kansensis (Crow) Skrjabin and Antipin. (HWML #101991, #101993). A female Copperhead collected on 16 June 2004 from Wake County harbored 6 O. kansensis in its mouth and esophagus. We found 1 O. kansensis in the esophagus of a male Copperhead collected on 21 August 2003 from Randolph County. This mouth fluke is a common parasite of snakes and has been reported previously from at least 15 colubrid and viperid species, including Copperhead, Cottonmouth, and Timber Rattlesnake; its geographic range in the US is listed as Arkansas, Florida, Georgia, Illinois, Kansas, Louisiana Missouri, Oklahoma, Tennessee, and Texas (Ernst and Ernst 2006). Although this is the first published report of O. kansensis from North Carolina, specimens (USNPC #97949) from another Southeastern Naturalist 731 E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 Cottonmouth collected from Bertie County, NC are deposited in the USNPC. A male Cottonmouth collected on 01 August 2009 from Moore County harbored 2 Ochetosoma sp.; however, the specimens were too distorted to determine specific identifications. Identification of Ochetosoma species follows the key of Dubois and Mahon (1959). During studies of O. kansensis and Ochetosoma laterotrema, Sogandares-Bernal and Grenier (1971) found these digeneans to exhibit a snail–amphibian–snake life cycle. They experimentally infected the aquatic snail Physella anatina (Haldeman) as first intermediate host, tadpoles of the Rana pipiens (Schreber) (Northern Leopard Frog), as second intermediate hosts, and Cottonmouths as definitive hosts. Such a life cycle is supported by Palmer and Braswell (1995), who have listed the amphibians Ambystoma opacum (Gravenhorst) (Marbled Salamander) and Plethodon sp. (slimy salamander) as food records for North Carolina Copperheads, and the amphibians Pseudotriton montanus Baird (Mud Salamander), Marbled Salamder, Hyla cinerea (Schneider) (Green Treefrog), Rana utricularia (Cope) (Southern Leopard Frog), and Rana catesbeiana Shaw (Bullfrog) for Cottonmouths. Another ophidian mouth fluke, Ochetosoma aniarum, also utilizes a similar snail–amphibian– snake life cycle (Byrd 1935). Table 1. Helminths from 14 of 27 Agkistrodon contortrix (Copperhead) collected from North Carolina. Counties where snake host was collected: Co = Columbus, Da = Dare, Mn = Montgomery, Mo = Moore, Ra = Randolph, Ri = Richmond, Sa = Sampson, Sc = Scotland, Wi = Wilkes, Wk = Wake, Ws = Washington. †denotes new host record and ‡ denotes new locality record. § = this digenean specimen too distorted to identify. Number Intensity Helminth species HWML # of snakes Mean Range Counties Trematoda Ochetosoma kansensis (Crow)‡ 101991, 101993 2 3.5 1–6 Ra, Wk Travtrema stenocotyle (Cohn)†‡ 101994 1 1.0 - Ra Unidentified digenean§ 1 1.0 - Mo Cestoda Proteocephalus sp.‡ 102069 2 1.5 1–2 Ra Nematoda Kalicephalus inermis coronellae (Ortlepp) 96262, 96263, 4 1.8 1–3 Mn, Sc 96265 Strongyloides serpentis Little ‡ 96264, 96266 2 2.5 2–3 Mo, Ri Capillaria colubra Pence† 1 3.0 - Da Physaloptera squamatae Harwood‡ 1 2.0 - Sa Physalopterid larvae† 1 1.0 - Sc Acanthocephalan Macracathorhynchid cytacanths†‡ 101992 2 1.0 - Ra, Ri No helminths 13 Co, Mn, Mo, Ra, Ri, Sc, Wi, Ws Southeastern Naturalist E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 732 Travtrema stenocotyle (Cohn) Pereira. (HWML #101994). The small intestine of a male Copperhead, collected on 21 August 2003 from Randolph County, was infected with a single Travtrema stenocotyle. This digenean was first described in North America by McIntosh (1939) as Leptophyllum tamiamiensis from “three lots of material” with “A. piscivorus” listed as the only host species for all three lots. The geographic locations for the hosts were Washington DC, National Zoological Park (USNPC #14523.02, #59988), New York Zoological Park (USNPC #44035), and Florida (USNPC #44033, #44034, #44033, #98842, #98843). Later that same year, Leptophyllum ovalis (USNPC #9312, #9313, #80681, #80682, and HWML #31092) was described from the small intestine of Brown Water Snakes that had been purchased from Florida and kept in Iowa for 6 months (Byrd and Roudabush 1939, Platt and Prestwood 1990). Also, 1500 specimens of L. tamiamiensis from a Florida “cotton-mouth” (Byrd and Roudabush 1939) were later used to describe the excretory system of this parasite (Byrd et al. 1940). Schad (1953) reported this fluke from “Elaphe quadrivittata deckerti” (probably the Yellow Rat Snake) without providing a geographic location, and considered L. ovalis to be a synonym of L. tamiamiensis. Goodman (1958), who reported this trematode from Farancia abacura abacura (Holbrook) (Florida Eastern Mud Snake), moved the species to the genus Travtrema Pereira 1929. Because the generic name Leptophyllum (Verhoeff), was previously occupied by a Myriapoda, Goodman (1958) transferred all previously described species (Leptophyllum stenocotyle Cohn, Travtrema travtrema Pereira, L. tamiamiensis McIntosh, and L. ovalis Byrd and Roudabush ) in Table 2. Helminths from 4 of 4 Agkistrodon piscivorus (Cottonmouth) and 3 of 7 Crotalus horridus (Timber Rattlesnake) collected from North Carolina. County where snake host was collected: Br = Brunswick, Bt = Bertie, Mo = Moore, On = Onslow, Ri = Richmond, Sc = Scotland, and Wi = Wilkes. †denotes new host record and ‡ denotes new locality record. § = only strobila fragments (3–15 per snake) were collected; thus, a definitive count of tapeworms could not be made. Of the 4 Cottonmouths from which tapeworms were collected, 0, 0, 1, 12 scolices were found. Number Intensity Snake species/Helminth species HWML # of snakes Mean Range Counties Agkistrodon piscivorus Trematoda Ochetosoma sp. 1 2.0 - Mo Cestoda Proteocephalus sp. 99989, 99990, 4 ? 3–15§ Br, Mo, 99991, 102070, Ri, Sc 102071, 102072 Nematoda Capillaria colubra Pence† 96267, 96268 2 1.5 1–2 Br, Mo Crotalus horridus Nematoda Kalicephalus costatus parvus (Ortlepp)†‡ 96270 1 1.0 - Bt Capillaria colubra Pence† 96269 1 1.0 - On Unidentified nematode larvae 99988 1 4.0 - Wi No Helminths 4 Mo Southeastern Naturalist 733 E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 synonymy under T. stenocotyle (Cohn) Pereira. It is unclear if Tkach (2008) agrees with this synonymy. The Copperhead and North Carolina are new host and locality records, respectively, for T. stenocotyle. Hamann and Gonzalez (2009) and Hamann et al. (2012) have reported the larval stage of T. stenocotyle from Argentinian amphibians; indicating that, like the mouth flukes of the genus Ochetosoma, T. stenocotyle also utilizes a snail–amphibian– snake life cycle. Cestoda Proteocephalus sp. Weinland. (HWML #99989, #99990, #99991, #102069, #102070, #102071, #102072). The same male Copperhead that was infected with T. stenocotyle, also harbored 2 Proteocephalus sp. tapeworms, but no scolices were found. We found a single Proteocephalus sp. scolex from a second male Copperhead collected from Randolph County. All 4 Cottonmouths of this study harbored from 3 to 15 fragments of Proteocephalus sp. strobilae. We recorded very few scolices (0, 0, 1, 12) from these Cottonmouths, probably due to the salvage nature of the host specimens. Although species-level identifications were not possible, the tapeworms were most likely Proteocephalus marenzelleri (Barrois) or Proteocephalus perspicua (LaRue). Collins (1968, 1969) reported 2 Proteocephalid tapeworms (P. marenzelleri and P. perspicua [as Ophiotaenia marenzelleri and Ophiotaenia perspicua, respectively]) from North Carolina Cottonmouths; however, the current survey is the first report of a Proteocephalus sp. from Copperheads. We used Khalil et al. (1994) for Cestode identifications. Earlier workers provided details of the life cycle of the ophidian tapeworm P. perspicua, which is likely one of the unidentified species of the tapeworms found in the present study. Herde (1938) reported that tapeworm eggs collected from P. perspicua from Oklahoma Nerodia rhombifer (Hallowell) (Diamondback Water Snakes) hatched in tap water. The newly hatched larval tapeworm (onchosphere) penetrates the copepod hosts, Cyclops viridis (Jurine) and Microcyclops varicans (Sars), and develops to the next larval stage (procercoid). Thomas (1941) utilized P. perspicua adult worms from a Texas Diamondback Water Snake and specimens of Michigan Northern Water Snake to conduct his life-cycle studies. Cyclops vulgaris and C. viridis were used as first intermediate hosts. After ingesting infected crustacean hosts, tadpoles of the Rana clamitans (Latreille) (Green Frog) and Northern Leopard Frog became infected with the second tapeworm larval stage (pleurocercoid). Adult frogs that had been infected as tadpoles retained the infection after metamorphosis. Laboratory-reared Thamnophis sirtalis (L.) (Garter Snakes) and Northern Water Snakes became infected with juvenile and adult tapeworms after being fed infected tadpoles or adult frogs. Ulmer and James (1976) found pleurocercoid larvae, which they believed to be P. perspicua, from 36 Northern Leopard Frogs and 1 Bufo americanus Holbrook (American Toad) from Iowa. It would seem that a Randolph County, NC, Copperhead, in the present study, had a predilection for amphibians because this snake was infected with a mouth fluke (O. kansensis.), an intestinal fluke (T. stenocotyle), and 2 Proteocephalus sp. tapeworms, all of which utilize amphibians as their second intermediate hosts. Southeastern Naturalist E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 734 Nematoda Kalicephalus inermis coronellae (Ortlepp) Schad. (HWML #96262, #96263, #96265). Four Copperheads collected in 2 counties (Montgomery and Scotland) of the North Carolina Sandhills were infected with the ophidian hookworm, K. i. coronellae. A total of 7 K. i. coronellae were collected from the snakes’ esophagus or intestine. Ernst and Ernst (2006) listed this hookworm from 25 snake species, including Copperheads, from Colorado, Florida, Georgia, Massachusetts, New Mexico, North Carolina, and Texas. Kalicephalus costatus parvus (Ortlepp) Schad. (HWML #96270). A Timber Rattlesnake collected in 2007 from Bertie County harbored a single K. c. parvus. However the damage to the salvaged host was so extensive that the nematode’s host organ of residence was not discernable. This is the first report of K. c. parvus from a North Carolina Timber Rattlesnake. Species identification and synonymy for Kalicephalus species follow that of Schad (1962). While investigating the life cycles of kalicephalid nematodes, Schad (1956) was able to experimentally infect snakes (Pituophis spp. Holbrook [Bullsnakes]), Garter Snakes, and Storeria dekayi [Holbrook] [Brown Snakes]) with Kalicephalus parvus, Kalicephalus agkistrodontis, and Kalicephalus rectiphilus. He considered the most likely routes of infection in nature to be “ingestion” of infective larvae introduced into the mouth on the snake’s tongue during sensory reception and/or through larval skin penetration. Schad (1956) suggested that kalicephalid larvae might be more likely to skin-penetrate recently fed snakes, which likely have higher than ambient temperatures during prey digestion. Like the ancylostomatid nematodes of mammals, kalicephalids appear to have direct life cycles, with potential for the use of prey species as paratenic (transport) hosts. And like the hookworm larvae that infect mammals, kalicephalid infective larvae require warm, moist, and shaded habitats to develop and extend survival within the environment. Physalopterid larvae. A single physalopterid larva was collected from the stomach of a Scotland County Copperhead. As suggested by Goldberg and Bursey (2001), such larvae are likely temporary residents associated with the inclusion of insects in a snake’s diet. The Copperhead provides a new host record; although physalopterid larvae have been previously reported from North Carolina amphibians (Dyer and Brandon 1973, Mann 1932, Rankin 1937, Walton 1935). Physaloptera squamatae Harwood. A Copperhead from Sampson County harbored 2 specimens of P. squamatae. The present study is the first report of P. squamatae from North Carolina. We utilized Anderson et al. (2009) and Chabaud (1956) to determine the nematodes’ generic group; Bursey and Brooks (2011) and Chabaud (1956) were used for the specific identity of physalopte rids. Ophidian physalopterids are relatively large nematode parasites that often gain the attention of herpetologists (McCauley 1945, Ortenburger 1928) as well as parasitologists. In the US, there have been 4 nominal species reported from snakes: Physaloptera abjecta Leidy, Physaloptera obtusissima Molin, P. squamatae Harwood, and Physaloptera variegata Reiber, Byrd, and Parker. The first to be reported from a US snake was P. abjecta by Leidy (1856); since then there has been much published regarding the validity of the physalopterid fauna of snakes. Southeastern Naturalist 735 E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 Without mentioning the South American species, P. obtusissima Molin or Physaloptera monodens Molin, Harwood (1932) erected P. squamatae as a new species from Agkistrodon mokasen (Texas Copperhead) and a Scincella lateralis (Say in James as Leiolopisma laterale) (Ground Skink). Although Morgan (1941a, b; 1943) and McAllister et al. (2010) have considered P. squamatae to be a synonym under P. obtusissima, many others (Baker 1987; Bowie 1974; Brooks 1963, 1972; Bursey and Brooks 2011; Chabaud 1956; Goldberg and Bursey 2000; Goldberg et al. 1994; Goldberg et al. 2002; McAllister and Bursey 2007; McAllister et al. 2014; Ortlepp 1937; Price and Underwood 1984; Reiber et al. 1940; and Telford and Bursey 2003) recognize P. squamatae as a valid species. In their text-key of American reptilian physalopterids, Bursey and Brooks (2011) separated the 2 species, P. obtusissima and P. squamatae, by the location of the 3rd pair of sessile papillae on the male’s caudal end. It is noteworthy that during his extensive work on the Physalopterids, Morgan (1940; 1941a, b, c; 1943) initiated a host-record error for P. squamatae that was propagated by later authors. In error, Morgan reported that Harwood’s (1932) snake host (Agkistrodon mokasen) was a “water moccasin”, not the correct host, “copperhead snakes (Agkistrodon mokasen)” (see Harwood 1932:20). McAllister and Bursey (2007) eventually corrected this host error. The terrestrial versus semiaquatic habitats of Copperheads and Cottonmouths, respectively, makes this an important biological correction. Physalopterid nematodes utilize insects as intermediate hosts. Although Palmer and Braswell (1995) list various insects as food items for North Carolina Copperheads, it is interesting that since Harwood’s (1932) original report of P. squamatae from a Texas Copperhead and a Ground Skink, this worm has only been reported from lizards (McAllister and Bursey 2007, McAllister et al. 2014). Also, all 22 museum lots (19 from USNPC and 3 from HWML) listed as P. squamatae are from lizards. This finding suggests that reports of P. squamatae from Copperheads (Harwood 1932, present study), may be “a by-product of diet and not parasites sensu stricto” Bursey and Brooks (2011). Further evidence of P. squamatae potentially being an incidental parasite in Copperheads is that Palmer and Braswell (1995) listed Grounds Skinks as food items for North Carolina Copperheads. Also McAllister et al. (2014) found P. squamatae to be the most common helminth of Ground Skinks from Oklahoma and Arkansas, and “in some lizards, represented massive infections (up to 64 worms)”. Capillaria colubra Pence. (HWML #96267, #96268, #96269). We found all species of examined viperids (Copperhead, Cottonmouth, and Timber Rattlesnake) to be infected with C. colubra. After Pence’s (1970) original species description of C. colubra from a Louisiana Coluber constrictor priapus Dunn and Wood (Southern Black Racer), Collins (1973) corrected the identifications of his 1968 study, from Capillaria heterodontis Harwood to C. colubra. Thus, C. colubra has been previously reported from North Carolina in 3 species of water snakes (Northern Water Snake, Nerodia fasciata (L.) [Banded Water Snake], and N. e. erythrogaster (Forster in Bossu) Southeastern Naturalist E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 736 [Redbelly Water Snake]) (Collins 1968, 1969, 1973). Finally, Collins (1968, 1969) reported Capillaria sp. from 18.8% of his Cottonmouth hosts, but did not provide a species determination. The larger morphometrics (body length and egg size) and the striated spicules of male specimens distinguish our specimens as C. colubra. Although previous reports of this worm were from the hosts’ oviducts, 2 of our snake hosts (a Copperhead and a Cottonmouth) were male. The salvage nature of our host specimens prevents definitive localization of the worms within the snakes. The current study is the first to report C. colubra from the Copperhead, Cottonmouth, and Timber Rattlesnake. We utilized Biserkov et al. (1994), Harwood (1932), and Pence (1970) to determine species identification. Nematodes of the Family Capillariidae display either direct life cycles in which infective eggs are ingested by the definitive host or indirect life cycles in which infective larvae within a prey host is ingested by the predatory definitive host (Moravec et al. 1987). Unfortunately the life cycles of ophidian capillarids have not been investigated. However, one can speculate that snakes may become infected via either route—direct ingestion of an infective egg from a contaminated environment or ingestion of infective capillarid larvae within an annelid or arthropod prey. Strongyloides serpentis Little. (HWML #96264, #96266). Two Copperheads, 1 from Moore County and 1 from Richmond County, were infected with the ophidian threadworm, S. serpentis. Mati and Melo (2014) and Santos et al. (2010) have confirmed that Strongyloides ophidiae Pereira of South American snakes is a distinct species from S. serpentis and Strongyloides gulae Little of North American snakes. Little (1966) originally described S. serpentis from 9 species of Louisiana snakes, including Copperheads and Cottonmouths; however, this parasite has only been reported by Fontenot and Font (1996) from Louisiana Nerodia cyclopion (Duméril, Bibron, and Duméril) (Green Water Snake). The current survey is the first report of S. serpentis from North Carolina. Descriptions by Little (1966) were utilized for species identification. Although the life cycle of S. serpentis has not been reported; a closely related species, Strongyloides ophidiae Pereira, has been studied by Mati and Melo (2014). These researchers found that snakes are infected via cutaneous penetration of infective larvae from a contaminated environment. Warm, moist, shaded habitats promote the development and survival of Strongyloides spp. These nematode parasites also have a unique life cycle in which, under such favorable conditions, multiple generations of male and female free-living worms can produce large numbers of infective larvae within a short period, thus making this an important parasite for captive herpetological collections (Mati and Melo 2014). Unidentified nematode larvae. (HWML #99988). A Timber Rattlesnake from Wilkes County was found to have 4 nematode larvae in its esophagus. The larvae were ~1.84 mm long, with a 0.215-mm strongyliform esophagus, a 0.075-mm-long tail (anus to tip) and transverse cuticular striations. The stomal region and body were indistinct, but the tip of the tail formed a dorsally directed crescent-shaped Southeastern Naturalist 737 E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 hook. Although we were unable to identify these larvae, they possess a similar size and appearance as the larvae of Baylisascaris procyonis (Stefanski and Zarnowski) (Raccoon Ascarid), as described and illustrated by Bowman (1987). Acanthocephala Macracanthorhynchid cystacanths. (HWML #101992). Acanthocephalan larvae were collected from 2 Copperheads. Collins (1968, 1969) reported the centrorhynchid cystacanth, Centrorhynchus conspectus Van Cleave and Pratt, from North Carolina snakes, including 3 species of water snakes and the Cottonmouth. A spherical proboscis with relatively few large hooks distinguished our specimens as macracanthorhynchids; centrorhynchids have a cylindrical proboscis with many small hooks. This report is the first of macracanthorhynchid cystacanths from Copperheads of North Carolina. We used Amin (2013) and Petrochenko (1956, 1958) for acanthocephalan identifications. Macracanthorhynchids utilize a terrestrial indirect life cycle. The mammalian definitive host releases parasite eggs in their feces. The eggs are then ingested by an arthropod, most often a coleopteran, in which a larval acanthocephalan develops. The life cycle is completed when the mammalian host ingests the arthropod host. However, if the arthropod host is ingested by a snake or other reptile, the larval acanthocephalan will encyst as a cystacanth within the tissues of the reptile host. The life cycle is then completed if the snake becomes prey to a mammalian host such as Procyon lotor (L.) (Raccoon), Canis latrans Say (Coyote), fox, Canis lupus familiaris L. (Domestic Dog), Felis catus L. (Domestic Cat), skunk, etc. (Petrochenko 1956, 1958). We report herein 7 new host records and 7 new geographic distribution records for various helminths from 3 North Carolina pit vipers. Previous to our study, only a small percentage (5 of 37 species or 13.5%) of North Carolina snakes had been surveyed for helminth parasites. We documented and thereby add 2 more hosts to that list; however, additional surveys on those species that have not yet been examined in the state could reveal additional host and geographic records. Nelder and Reeves (2005) discussed the advantages of utilizing salvaged hosts for parasitic studies, including the simplification of host collection, as well as the avoidance of euthanasia procedures along with associated regulatory issues. Our study has identified disadvantages of using salvaged hosts. Some hosts may be so damaged or deteriorated that confident determination of the helminths’ natural habitat within the host is impossible. Helminths may become distorted or deteriorated to the point of rendering specific identification impossible. Based on our inability to determine the intensity of infection or the specific identifications of the tapeworms, it would seem that tapeworms are the first to deteriorate to such a point. However, despite these disadvantages, the parasitological study of salvaged herpetofauna is still herpetologically and helminthologically fruitful because such records yield information about the extent of the host–parasite interactions across the landscape. Southeastern Naturalist E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 738 Acknowledgments We thank herpetological collectors that collected and donated snakes to the North Carolina State Museum of Natural Sciences. Salvage permits were provided by the North Carolina Wildlife Resources Commission. We are grateful to Dr. Gabor Racz and Dr. Scott Gardner of the Harold W. Manter Laboratory of Parasitology and Pat Pilitt and Dr. Eric Hoberg of the US National Parasite Collection for assistance with museum specimens and data. Funding for this project was provided by the Merial Veterinary Scholars Program in cooperation with the NCSU-CVM Summer Research Internship Program. Literature Cited Amin, O.M. 2013. Classification of the Acanthocephla. Folia Parasitologica 60:273–305. Anderson, R.C., A.G. Chabaud, and S. Willmott. 2009. Keys to the Nematode Parasites of Vertebrates. Archival Volume. CAB International, Oxfordshire, UK. 463 pp. Baker, M.R. 1987. Synopsis of the Nematoda parasitic in amphibians and reptiles. Memorial University of Newfoundland Occasional Papers in Biology 11:1–325. Biserkov, V.Y., F. Mészáros, and N. Chipev. 1994. On the validity of the species considered synonyms of Paracapillaria sonsinoi (Parona, 1897) (Nematoda: Capillariidae). Parasitologia Hungarica 27:53–56. Bowie, L.A. 1974. Comparative study of the gastrointestinal nematodes of two sceloporid lizards in Florida. M.Sc. Thesis. University of Florida, Gainesville, FL. 47 pp. Bowman, D.D. 1987. Diagnostic morphology of four larval Ascaridoid nematodes that may cause visceral larva migrans: Toxascaris leonina, Baylisascaris procyonis, Lagochilascaris sprenti, and Hexametra leidyi. Journal of Parasitology 73:1198–1215. Brooks, D.R. 1979. New records for amphibian and reptile trematodes. Proceedings of the Helminthological Society of Washington 46:286–289. Brooks, G.R. 1963. Intestinal helminths of the Ground Skink (Lygosoma laterale). Virginia Journal of Science 14:198. Brooks, G.R. 1972. Intestinal helminths of the Ground Skink (Lygosoma laterale). Quarterly Journal of the Florida Academy of Sciences 35:8–14. Bursey, C.R., and D.R. Brooks. 2011. Nematode parasites of Costa Rican snakes (Serpentes) with description of a new species of Abbreviata (Physalopteridae). Comparative Parasitology 78:333–358. Bush, A.O., K.D. Lafferty, J.M. Lotz, and A.W. Shostak. 1997. Parasitology meets ecology on its own terms: Margolis et al. revisited. Journal of Parasitology 83:575–583. Byrd, E.E. 1935. Life-history studies of Reniferinae (Trematoda, Digenea) parasitic in Reptilia of the New Orleans area. Transactions of the American Microscopical Society 54:196–225. Byrd, E.E., and R.L. Roudabush. 1939. Leptophyllum ovalis n. sp., a trematode from the Brown Watersnake. Journal of Parasitology 25:471–473. Byrd, E.E., M.V. Parker, and R.J. Reiber. 1940. A new genus and two new species of digenetic trematodes, with a discussion on the systematics of these and certain related forms. Journal of Parasitology 26:111–122. Chabaud, A.G. 1956. Essai de révision des Physaloptères parasites de reptiles. Annales de Parasitologie Humaine et Comparee 31:29–52. Collins, R.F. 1968. The helminths of Natrix spp. and Agkistrodon piscivorus piscivorus (Reptilia: Ophidia) in eastern North Carolina. M.A. Thesis. Wake Forest University, Winston Salem, NC. 30 pp. Southeastern Naturalist 739 E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 Collins, R.F. 1969. The helminths of Natrix spp. and Agkistrodon piscivorus piscivorus (Reptilia: Ophidia) in eastern North Carolina. Journal of the Elisha Mitchell Scientific Society 85:141–144. Collins, R.F. 1973. New host and locality records for Capillaria colubra Pence, 1970. Journal of Parasitology 59:1020. Conant, R., and J.T. Collins. 1998. A Field Guide to Reptiles and Amphibians: Eastern and Central North America. 3rd Edition. Houghton Mifflin, Boston, MA. 616 pp. Dubois, G., and J. Mahon. 1959. Étude de quelques trématodes Nord-Americains (avec note sur la position systematique de Parorchis Nicoll 1907) suivie d’une revision des genres Galactosomum Looss 1899 et Ochetosoma Braun 1901. Bulletin de la Société Neuchâteloise des Sciences Naturelles 82:191–229. Dyer, W.G., and R.A. Brandon. 1973. Helminths of three sympatric species of cave-dwelling salamanders in southern Illinois. Transaction of the Illinois Academy of Science 66:23–29. Ernst, C.H., and E.M. Ernst. 2006. Synopsis of helminths endoparasitic in snakes of the United States and Canada. Society for the Study of Amphibians and Reptiles Herpetological Circular 34. 86 pp. Fontenot, L.W., and W.F. Font. 1996. Helminth parasites of four species of aquatic snakes from two habitats in southeastern Louisiana. Journal of the Helminthological Society of Washington 63:66–75. Goldberg, S.R., and C.R. Bursey. 2000. Transport of helminths to Hawaii via the Brown Anole, Anolis sagrei (Polychrotidae). Journal of Parasitology 86:750–755. Goldberg, S.R., and C.R. Bursey. 2001. Helminths of six species of colubrid snakes from southern California. Bulletin of the Southern California Academy of Sciences 92:43–51. Goldberg, S.R., C.R. Bursey, and F. Kraus. 1994. Helminth parasites of the Bark Anole, Anolis distichus and the Brown Anole, Anolis sagrei (Polychrotidae), from Florida and the Bahamas. Caribbean Journal of Science 30:275–277. Goldberg, S.R., C.R. Bursey, and R. Tawil. 2002. Seasonal variation in the helminth community of the Brown Anole, Anolis sagrei (Polychrotidae), from Oahu, Hawaii. American Midland Naturalist 148:409–415. Goodman, J.D. 1958. Travtrematinae nom. nov. for Leptophyllinae Byrd, Parker, and Reiber, 1940 (Plagiorchiidae: Trematoda). Journal of Parasitology 44:106–109. Hamann, M.I., and C.E. González. 2009. Larval digenetic trematodes in tadpoles of six amphibian species from northeastern Argentina. Journal of Parasitology 95:623–628. Hamann, M.I., A.I. Kehr, and C.E. González. 2012. Community structure of helminth parasites of Leptodactylus bufonis (Anura: Leptodactylidae) from northeastern Argentina. Zoological Studies. 51:1454–1463. Harold W. Manter Laboratory of Parasitology (HWML). 2016. HWML database. Available online at Accessed 5 April 2016. Harwood, P.D. 1932. The helminths parasitic in the Amphibia and Reptilia of Houston, Texas and vicinity. Proceedings of the United States National Museum 81:1–71. Herde, K.E. 1938. Early development of Ophiotaenia perspicua La Rue. Transactions of the American Microscopical Society 57:282–291. Integrated Taxonomic Information System (ITIS). 2016. Taxonomic Information Database. Available online at Accessed 1 April 2016. Khalil, L.F., A. Jones, and R.A. Bray. 1994. Keys to the Cestode Parasites of Vertebrates. CAB International, UK. 751 pp. Leidy, J. 1856. A synopsis of entozoa and some of their ecto-congeners observed by the author. Proceedings of the Academy of Natural Sciences of Philadelphia 8:5–59. Southeastern Naturalist E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 740 Little, M.D. 1966. Seven new species of Strongyloides (Nematoda) from Louisiana. Journal of Parasitology 52:85–97. Mann, D.R. 1932. The ecology of some North Carolina salamanders with special reference to their parasites. M.A. Thesis. Duke University, Durham, NC. 51 pp. Mati, V.L.T., and A.L. Melo. 2014. Some aspects of the life history and morphology of Strongyloides ophidiae Pereira, 1929 (Rhabditida: Sytongyloididae) in Liophis miliaris (Squamata: Dipsadide). Neotropical Helminthology 8:203–216. McAllister, C.T., and C.R. Bursey. 2007. First report of the nematode, Physaloptera squamatae (Spirurida: Physalopteridae) in Oklahoma, with a summary of hosts. Proceedings of the Oklahoma Academy of Science 87:65–67. McAllister, C.T., C.R. Bursey, and P.S. Freed. 2010. Helminth parasites of herpetofauna from the Rupunini District, Southwestern Guyana. Comparative Parasitology 77:184–201. McAllister, C.T., C.R. Bursey, M.B. Connior, L.A. Durden, and H.W. Robison. 2014. Helminth and arthropod parasites of the Ground Skink, Scincella lateralis (Sauria: Scincidae), from Arkansas and Oklahoma, USA. Comparative Parasitology 81:210–219. McCauley, R.H., Jr. 1945. The Reptiles of Maryland and the District of Columbia. R.H. McCauley Jr. Hagerstown, MD. 194 pp. McIntosh, A. 1939. Description of a plagiorchioid trematode, Leptophyllum tamiamiensis, n. sp., from a poisonous snake. Proceedings of the Helminthological Society of Washington 6:92–94. Moravec F., J. Prokopic, and A.V. Shlikas. 1987. The biology of nematodes of the family Capillariidae Neveu-Lemaire, 1936. Folia Parasitologica 34:39–56. Morgan, B.B. 1940. The Physalopterinae (Nematoda) of North America. Journal of Parasitology 26(8):40. Morgan, B.B. 1941a. A summary of the Physalopterinae (Nematoda) of North America. Proceedings of the Helminthological Society of Washington 8:28–30. Morgan, B.B. 1941b. The Physalopterinae (Nematoda) of North American vertebrates. Ph.D. Dissertation. University of Wisconsin, Madison, WI. 102 pp. + 2 Plates. Morgan, B.B. 1941c. Additional notes on North American Physalopterinae (Nematoda). Proceedings of the Helminthological Society of Washington 8:63–64. Morgan, B.B. 1943. The Physaloptera (Nematoda) of Reptiles. Le Naturaliste Canadien 70:179–185. Nelder. M.P., and W.K. Reeves. 2005. Ectoparasites of road-killed vertebrates in northwestern South Carolina, USA. Veterinary Parasitology 129:313–322. Ortenburger. A.I. 1928. The Whip Snakes and Racers: Genera Masticophis and Coluber. Memoirs of the University of Michigan Museums Vol. 1. University of Michigan. Ann Arbor, MI. 247 pp. Ortlepp, R.J. 1937. Some undescribed species of the nematode genus Physaloptera Rud., together with a key to sufficiently known forms. Onderstepoort Journal of Veterinary Science and Animal Industry 9:71–84. Palmer, W.M., and A.L. Braswell. 1995. Reptiles of North Carolina. The University of North Carolina Press, Chapel Hill, NC. 412 pp. Pence, D.B. 1970. Capillaria colubra sp. n. from the oviducts of Coluber constrictor priapus. Journal of Parasitology 56:261–264. Petrochenko, V.I. 1956. Acanthocephala of Domestic and Wild Animals. Vol. 1. Moscow: Izdatel’stvo Akad. Nauk SSSR (In Russian). 1971. English translation. Israel Program for Scientific Translations, Ltd., Jerusalem, Israel. 465 pp. Southeastern Naturalist 741 E. Davis, J.C. Beane, and J.R. Flowers 2016 Vol. 15, No. 4 Petrochenko, V.I. 1958. Acanthocephala of Domestic and Wild Animals. Vol. 2. Moscow: Izdatel’stvo Akad. Nauk SSSR . (In Russian). 1971. English translation. Israel Program for Scientific Translations, Ltd., Jerusalem, Israel. 478 pp. Platt, T.R., and A.K. Prestwood. 1990. Deposition of type and voucher material from the helminthological collection of Elon E. Byrd. Systematic Parasitology 16:27–34. Price, W.W., and H. Underwood. 1984. Intestinal helminths of the Cuban Anole, Anolis sagrei sagrei, from Tampa, Florida. Florida Scientist 47:205–207. Pritchard, M.H., and G.O. Kruse. 1982. The Collection and Preservation of Animal Parasites. Technical Bulletin No. 1. The Harold W. Manter Laboratory, University of Nebraska Press, Lincoln, NE. 141 pp. Rankin, J.S. 1937. An ecological study of parasites of some North Carolina salamanders. Ecological Monographs 7:169–269. Reiber, R.J., E.E. Byrd, and M.V. Parker. 1940. Certain new and already known nematodes from Amphibia and Reptilia. Lloydia 3:125–144. Richardson, D.J., and B.B. Nickol. 1995. The genus Centrorhynchus (Acanthocephala) in North America with description of Centrorhynchus robustus n. sp., redescription of Centrorhynchus conspectus, and a key to species. Journal of Parasitology 81:767–772. Santos, K.R. dos, B.C. Carlos, K.S. Paduan, S.M. Kadri, T.H. Barrella, M.R.V. Amarante, P.E.M. Ribolla, and R.J. da Silva. 2010. Morphological and molecular characterization of Strongyloides ophidiae (Nematoda, Strongyloididae). Journal of Helminthology 84:136–142. Schad, G.A. 1953. Leptophyllum ovalis Byrd and Roudabush, a synonym of Leptophyllum tamiamiensis McIntosh. Journal of Parasitology 39:673. Schad, G.A. 1956. Studies on the genus Kalicephalus (Nematoda: Diaphanocephalidae). I. On the life histories of the North American species K. parvus, K. agkistrodontis, and K. rectiphilus. Canadian Journal of Zoology 34:425–452. Schad, G.A. 1962. Studies on the genus Kalicephalus (Nematoda: Diaphanocephalidae). II. A taxonomic revision of the genus Kalicephalus Molin, 1861. Canadian Journal of Zoology 40:1035–1165 + plates I–IV. Sogandares-Bernal, F., and H. Grenier. 1971. Life cycle and host-specificity of the plagiorchiid trematodes Ochetosoma kansensis (Crow, 1913) and O. laterotrema (Byrd and Denton, 1938). Journal of Parasitology 57:297. Sprent, J.F.A. 1988. Ascaridoid nematodes of amphibians and reptiles: Ophidascaris Baylis, 1920. Systematic Parasitology 11:165–213. Telford, S.R., Jr., and C.R. Bursey. 2003. Comparative parasitology of squamate reptiles endemic to scrub and sandhills communities of north-central Florida, USA. Comparative Parasitology 70:172–181. Thomas, L.J. 1941. The life cycle of Ophiotaenia perspicua La Rue, a cestode of snakes. Revista de Medicina Tropical y Parasitologia, Bacteriologia, Clinica y Laboratorio 7:74–78. Tkach, V.V. 2008. Family Plagiorchiidae Lühe, 1901. Pp. 295–325, In D.I. Gibson, A. Jones and R.A. Bray (Eds.). Keys to the Trematoda. Vol. 3. CAB International and Natural History Museum London, UK. 824 pp. Ulmer, M.J., and H.A. James. 1976. Studies on the helminth fauna of Iowa II. Cestodes of amphibians. Proceedings of the Helminthological Society of Washington 43:191–200. United States National Parasite Collection (USNPC). 2015. USNPC database. Available online at xlsx. Accessed 23 June 2015. Walton, A.C. 1935. The Nematoda as parasites of Amphibia. II. Journal of Parasitology 21:27–50.