Martin et al. Parasites & Vectors (2019) 12:39 https://doi.org/10.1186/s13071-018-3283-9 RESEARCH Open Access Culicoides species community composition and infection status with parasites in an urban environment of east central Texas, USA Estelle Martin1*, Elaine Chu1, Phillip Shults1, Andrew Golnar1, Dustin A. Swanson2, Jamie Benn3, Dongmin Kim1, Peter Schneider3, Samantha Pena4, Cassie Culver4, Matthew C. I. Medeiros5, Sarah A. Hamer6 and Gabriel L. Hamer1* Abstract Background: Despite their importance as vectors of zoonotic parasites that can impact human and animal health, Culicoides species distribution across different habitat types is largely unknown. Here we document the community composition of Culicoides found in an urban environment including developed and natural sites in east central Texas, a region of high vector diversity due to subtropical climates, and report their infection status with haemoparasites. Results: A total of 251 individual Culicoides were collected from May to June 2016 representing ten Culicoides species, dominated by C. neopulicaris followed by C. crepuscularis. We deposited 63 sequences to GenBank among which 25 were the first deposition representative for six Culicoides species: C. arboricola (n =1);C. nanus (n =4);C. debilipalpis (n =2);C. haematopotus (n =14);C. edeni (n =3);andC. hinmani (n = 1). We also record for the first time the presence of C. edeni in Texas, a species previously known to occur in the Bahamas, Florida and South Carolina. The urban environments with natural area (sites 2 and 4) had higher species richness than sites more densely populated or in a parking lot (sites 1 and 3) although a rarefaction analysis suggested at least two of these sites were not sampled sufficiently to characterize species richness. We detected a single C. crepuscularis positive for Onchocercidae gen. sp. DNA and another individual of thesamespeciespositiveforHaemoproteus sacharovi DNA, yielding a 2.08% prevalence (n = 251) for both parasites in this species. Conclusions: We extend the knowledge of the Culicoides spp. community in an urban environment of Texas, USA, and contribute to novel sequence data for these species. Additionally, the presence of parasite DNA (Onchocercidae gen. sp. and H. sacharovi)fromC. crepuscularis suggests the potential for this species to be a vector of these parasites. Keywords: Biting midges, Avian hosts, Onchocercidae, Haemosporida, Vector-parasite association, Integrative taxonomy * Correspondence: [email protected]; [email protected] 1Department of Entomology, Texas A&M University, College Station, Texas, USA Full list of author information is available at the end of the article © The Author(s). 2019 Open Access This article is distributed under the terms of the Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/), which permits unrestricted use, distribution, and reproduction in any medium, provided you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons license, and indicate if changes were made. The Creative Commons Public Domain Dedication waiver (http://creativecommons.org/publicdomain/zero/1.0/) applies to the data made available in this article, unless otherwise stated. Martin et al. Parasites & Vectors (2019) 12:39 Page 2 of 10 Background with a 6 volt battery, USA) baited with 1.5 kg of dry ice The family Ceratopogonidae (Diptera) includes the were run from approximately 18:00 h to 8:00 h. Traps genus Culicoides, commonly known as biting midges or were placed in 4 localities: a grocery store parking lot “no-see-ums”. Culicoides spp. are hematophagous pests (site 1), research park natural area (site 2), and two pri- and vectors of viruses, protozoans and filarial worms vate homes in residential neighborhoods (site 3 and site [1–3]. A major focus of research on Culicoides spp. is re- 4) (Fig. 1). While site 1 and 3 are located in urban devel- lated to their roles as vectors of epizootic hemorrhagic oped areas, sites 2 and 4 are located in urban natural en- disease virus (EHDV), bluetongue disease virus (BTV) vironments with more vegetation in the surrounding and African horse sickness virus (AHSV) [3]. Beside landscape. In site 4, Culicoides traps were set in three lo- their impact on livestock, Culicoides also transmit para- cations, including one location next to a chicken coop. sites to wildlife such as the avian trypanosome Trypano- After collection, trap contents were chilled on ice for soma bennetti [4] and Haemoproteus parasites [5, 6]; transport to the laboratory where specimens were identi- however, distributional patterns and infection preva- fied to the species level based on wing pattern [17, 18] lences of these parasites remain largely unknown. and stored at -20 °C until further processing. One neglected area in the study of Culicoides spp. is We used an integrative taxonomic approach to identify their ecology in urban environments, where there is an Culicoides. Thirteen specimens were mounted on per- interface with human populations. Prior studies have fo- manent slides to observe anatomic characteristics useful cused on specific species, such as pestiferous salt-marsh for species identification and were in parallel processed species [7] or species from specialized habitats such as by a non-destructive DNA extraction method modified zoos [8], and the vector-host-pathogen interactions in from the Gentra Puregene Kit (#D-5500A) (Gentra Sys- urban environments, could be of potential interest tems, Inc., Minneapolis, USA). Whole specimens were highlighting their potential as vectors of pathogens. Sev- added to individual Eppendorf tubes containing 100 μl eral species of Culicoides are known vectors of Haemo- of Cell Lysis Solution and 1 μl of Proteinase K and incu- sporida [9, 10], parasitic protozoans of amphibians, bated overnight at 55 °C. Specimens were not crushed to reptiles, birds and mammals [10]. These parasites can preserve morphological features. The specimens were cause acute epizootic outbreaks that have severely chilled at 0 °C for 20–30 min, after which 35 μl of 8.0 M altered avian communities by affecting long-term demo- ammonium acetate were added to each tube. The sam- graphic processes such as reproductive rate and sur- ples spun at 2200× rpm for 7 min in an Eppendorf 5424 vivorship [10–12]. Culicoides are also involved in the Centrifuge. The supernatant was pipetted out of the tube transmission of filarial nematodes in the family Oncho- and used to complete the DNA extraction protocol. Ap- cercidae with infection report in the California quail, the proximately 100 μl of 95% ethanol were added to the American crow and the great-tailed grackle [13–15]. tube containing the exoskeleton to prevent further Recently, a high prevalence of haematozoan parasites breakdown. The exoskeletons were separated into body was documented in a population of great-tailed grackles regions (head, thorax, abdomen and wings) and trans- (Quiscalus mexicanus) in College Station, Texas, includ- ferred to 15.0% acetic acid for 10 min, 2-propanol for 10 ing Heamoproteus, Plasmodium, trypanosomes and filar- min, and then 100.0% clove oil. Each body region of a ial worms (Golnar et al., unpublished data). Previous single specimen was slide mounted in Canada balsam work suggests that mosquitoes are not the vectors for under its own coverslip. Slides were kept at 40 °C for 8– these parasites and instead Culicoides are the more likely 10 h and air dried for 72 h [19]. Slides were deposited in vectors [16]. Here, we document the Culicoides spp. the Texas A&M University Entomology Museum under inhabiting the urban environment of College Station, accession number 735. This method allowed for identifi- Texas, USA, and screen individuals for blood-borne cation of specimens both molecularly and morphologic- parasites known to occur in local birds. The ally. Sequences of the cytochrome c subunit 1 (cox1) characterization of Culicoides species in this area where gene were obtained using the protocol described below. a high burden of avian parasites circulates provides an All the remaining specimens (n = 238) were homoge- ideal opportunity to evaluate their potential role as vec- nized in a Mini-Beadbeater-96 (Bio Spec Products Inc., tors of haematozoan parasites. Bartlesville, OK, USA) with three 4 mm PYREX Solid Beads (Sigma-Aldrich, Inc., St. Louis, MO, USA) in 200 Methods μl of Hank’s Buffer Salt Solution (ThermoFisher Scien- Culicoides collection and identification tific, Waltham, MA). DNA was extracted from 100 μlof We collected Culicoides from College Station, Texas (30° the homogenized material (MagMAX CORE Nucleic 37'40.7"N, 96°20'3.8"W) during the months of May and Acid Purification Kit from Applied Biosystems, Foster June, 2016. Centers for Disease Control and Prevention City, CA). A PCR assay targeting a 710 bp region of the miniature light traps (CDC-LT, BioQuip model 2836BQ, cox1 gene of invertebrates [20] was used for molecular Martin et al. Parasites & Vectors (2019) 12:39 Page 3 of 10 Fig. 1 Trapping sites across College Station, Texas, USA. Blue boxes and site numbers are locations where traps were set identification of Culicoides species (Additional file 1: KT794162.1; KT794164.1; KT794165.1; KT794167.1; Table S1). The PCR consisted of 1.5 μl of genomic DNA, KT794171.1) as well as 17 previously unpublished se- 0.5 μM of each primer, 12.5 μl of 1× Premix from the quences (GenBank: MH751220, MH751222, MH751223, Epicentre Failsafe PCR purification kit (Epicentre Bio- MH751224, MH751226, MH751227, MH751228, technologies, Madison, WI, USA), and 1 unit of enzyme MH751235, MH751243, MH751244, MH751246, mix (total volume 25 μl). The thermal cycling profile MH751248, MH751252, MH751258, MH751267, consisted of denaturation at 95 °C for 3 min, followed by MH751273, MH751280) from specimens collected in 35 cycles of 94 °C for 1 min, 45 °C for 1.5 min, 72 °C for Wisconsin, Wyoming and South Carolina and identified 2 min, and a final extension step at 72 °C for 5 min.