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A molecular assessment of Ostertagia leptospicularis and Spiculopteragia asymmetrica among wild fallow deer in Northern Ireland and implications for false detection of livestock-associated species

Parasites & Vectors volume 17, Article number: 141 (2024) Cite this article

Abstract

Background

Wild deer populations utilizing livestock grazing areas risk cross-species transmission of gastrointestinal nematode parasites (GINs), including GINs with anthelmintic resistance (AR) traits. Wild deer have been shown to carry problematic GIN species such as Haemonchus contortus and Trichostrongylus species in the UK, but the presence of livestock GINs in Northern Ireland deer populations is unknown. Also, is it not known whether AR traits exist among GINs of deer such as Ostertagia leptospicularis and Spiculopteragia asymmetrica in pastureland where anthelmintics are heavily used.

Methods

Adult-stage GIN samples were retrieved from Northern Irish wild fallow deer abomasa. Individual specimens were subject to a species-specific PCR analysis for common sheep and cattle GIN species with ITS-2 sequence analysis to validate species identities. In addition, the beta-tubulin gene was subject to sequencing to identify benzimidazole (BZ) resistance markers.

Results

ITS-2 sequencing revealed O. leptospicularis and S. asymmetrica, but species-specific PCR yielded false-positive hits for H. contortus, Teladorsagia circimcincta, Trichostrongylus axei, T. colubriformis, T. vitrinus and Ostertagia ostertagi. For beta-tubulin, O. leptospicularis and S. asymmetrica yielded species-specific sequences at the E198 codon, but no resistance markers were identified in either species at positions 167, 198 or 200 of the coding region.

Discussion

From this report, no GIN species of significance in livestock were identified among Northern Ireland fallow deer. However, false-positive PCR hits for sheep and cattle-associated GINs is concerning as the presence of deer species in livestock areas could impact both deer and livestock diagnostics and lead to overestimation of both GIN burden in deer and the role as of deer as drivers of these pathogens. ITS-2 sequences from both O. leptospicularis and S. asymmetrica show minor sequence variations to geographically distinct isolates. AR has been noted among GINs of deer but molecular analyses are lacking for GINs of wildlife. In producing the first beta-tubulin sequences for both O. leptospicularis and S. asymmetrica, we report no BZ resistance in this cohort.

Conclusions

This work contributes to genetic resources for wildlife species and considers the implications of such species when performing livestock GIN diagnostics.

Graphical Abstract

Background

Across the northern hemisphere, including Great Britain and Northern Ireland (NI), wild deer populations are increasing [1,2,3], resulting in deer encroachment onto livestock pastures and increased contact with livestock [4,5,6]. This raises questions about the impact of deer on livestock farming, especially given that deer species are susceptible to a number of economically significant livestock parasites, principally gastrointestinal nematodes (GINs) [5,6,7,8,9,10,11].

GIN infections of ruminants are complex, with many species capable of infecting a range of hosts across livestock and wildlife barriers [5, 7, 11]. For instance, pathogenic species of livestock such as Teladorsagia circimcincta [11] and Haemonchus contortus have been reported in natural infections of wild cervid species [8, 12,13,14]. Conversely, wildlife GIN species can cause infections in livestock such as Ostertagia leptospicularis in sheep [15,16,17,18,19] and cattle [17].

In the UK and Republic of Ireland alone, GINs cause an estimated €280 million in production losses annually, with a further €42 million spent on anthelmintic control measures [20]. Use of anthelmintics is by far the most common GIN control measure utilised in commercial livestock settings, but long-term reliance on a few classes of compounds has led to the emergence and spread of anthelmintic resistance among livestock GIN populations [21,22,23,24,25,26,27,28,29]. As such, AR is a major problem within the livestock industry and threatens closure of farms in areas where no anthelmintic control approaches are effective [30,31,32]. However, there are limited analyses of AR traits among GIN species of wild deer.

AR GIN populations are known to exist amongst farmed deer, for instance benzimidazole-resistant H. contortus have been reported [14]. Reduced anthelmintic efficacy has also been demonstrated in more cervid-restricted species, including in O. leptospicularis, Spiculopteragia spiculoptera and S. asymmetrica, for which no molecular markers for resistance are known [33,34,35,36,37,38]. Unlike farmed deer, wild cervids are not routinely treated with anthelmintics; therefore, carriage of resistant genotypes may be seen as indicative of interspecies contact with livestock or farmed deer [5, 7, 38,39,40]. However, AR traits are also known to occur naturally in free living nematodes such as Caenorhabditis elegans [41]. Additionally, wildlife may be exposed to low levels of anthelmintics in the environment such as in wastewater or on pasture [42, 43]. Therefore, GINs of cervids may also have developed resistance traits and warrants investigation.

In the face of widespread AR-GINs in livestock farming systems, approaches to improve diagnostics have been made partly to enable sustainable use of anthelmintics and to better our knowledge of GIN infections spatiotemporally [44, 45]. It is also critical to differentiate GIN species to provide an appropriate anthelmintic treatment [46]. However, morphological diagnostic approaches require expert knowledge and time-consuming methods to accurately diagnose specific species [47]. Currently, a number of DNA-based diagnostic approaches are in use or under development to detect and enumerate the presence of different GIN species based on species-specific markers that can detect individual GIN parasites by PCR [48,49,50,51,52,53,54,55], loop-mediated isothermal amplification (LAMP) [52, 56] or ITS-2 meta-barcoding [57, 58]. Methods have also been developed for the detection of deer and other wildlife GINs in both wildlife and livestock contexts [39, 46, 59, 60]. However, as these emerging technologies evolve, DNA databases need to keep pace with accurate and comprehensive data to ensure clarity of speciation [46]. A lack of knowledge remains pertaining to the accuracy and utility of DNA diagnostic tools in the presence of wildlife GIN species.

In NI, deer are the most prominent wild ruminants with fallow (Dama dama), red (Cervus elaphus) and Japanese sika (Cervus nippon) present, all of which are susceptible to GINs of livestock [61, 62]. However, GIN infections and AR in wild deer have gone largely under-investigated at both local and international scales. In this study, abomasa samples were obtained from three populations of Northern Ireland’s most abundant deer species, fallow deer, for nematode speciation and subsequent genetic analysis for evidence of resistance to widely used benzimidazole (BZ) anthelmintics. In addition, we cross-validate known PCR diagnostic primer sets against deer-specific species identified in this study to determine the potentiality of wildlife species yielding false-positive PCR results for livestock diagnostics.

Methods

Wild fallow deer specimen retrieval and DNA extraction

Seven fallow deer (Dama dama) were culled by trained stalkers, in adherence with Northern Ireland legislation, across three locations, Co. Antrim (Randalstown Forest), Co. Down (Tollymore Forest) and Co. Londonderry (Ardkill), during the legal open season (November 2019). Randalstown Forest is a 172 ha mixed conifer and broadleaf woodland situated at the northern edge of Lough Neagh. Tollymore Forest is a 630 ha mixed conifer and broadleaf woodland at the foot of the Mourne Mountains in the southeast of the country and Ardkill has several small forested areas situated in the north west of the country. Each of these woodland areas is surrounded by cattle and sheep pasture.

Adult nematodes were washed from abomasum tissue, cleaned in ddH2O and then stored in 70% EtOH at − 20 °C. Individual worms were washed thrice in 0.5 ml lysis buffer [100 mM KCl, 20 mM Tris pH 8, 2.5 mM MgCl2, 0.9% IGEPAL® CA-630 (Sigma-Aldrich, St. Louis, MO, USA), 0.9% Tween-20 (Sigma-Aldrich) and 0.02% gelatin, see nemabiome.ca website for details] and then transferred by pipette to an ethanol-cleaned microscope slide under a stereomicroscope and bisected in individual droplets of lysis buffer by bleach cleaned scalpel to break the cuticle. Worm pieces were transferred to individual tubes by bleach-cleaned no. 5 forceps with one sample taken per worm. DNA lysis was adapted from the Nemabiome protocol [57] with each sample digested in 100 µl lysis buffer and 4 µl proteinase K (0.8 mg/ml final concentration, NEB, Ipswich, MA, USA), which was vortexed, centrifuged and then digested overnight or at 56 °C in a water bath. Proteinase K was inactivated at 95 °C for 20 min and samples were stored at − 20 °C (20 mg/ml solution).

Species-specific PCR screening

PCR reactions were performed with GoTaq® G2 Hot Start Taq Polymerase (Promega, Madison, WI, USA) at recommended concentrations (2.5 mM MgCl2, 0.2 mM dNTPs (PCRbio, London, UK) and 0.4 µM of each primer). Previously published primers and the recommended annealing temperatures used are listed in Additional file 2: Table S1 [49, 63,64,65,66,67,68,69,70]. For all reactions, 25 μl volumes were used with 1 μl of DNA lysate per reaction or nuclease-free water for no template controls. For all PCR reactions, a 95 °C/2 min initial denaturation step and a final extension at 72 °C for 5 min was performed. For Bisset et al. primers [49], touchdown PCR conditions were used including: 12 cycles of 95 °C denaturation for 15 s, 60 °C annealing (with 0.5 °C decline per cycle), 72 °C extension for 30 s, followed by 25 cycles of 95 °C for 15 s, 54 °C for 15 s and 72 °C for 30 s as previously described. For other primers, 35 cycles of 95 °C denaturation for 15 s, 52–58 °C annealing for 15 s and 72 °C extension for 30 s were used, with as previously described. See Additional file 2: Table S1 for specific details. PCR products were visualized by precast 1 × concentration SYBRSafe (Thermo Fisher, Waltham, MA, USA) on 1.3% agarose gel electrophoresis in TAE buffer.

Sanger sequencing

PCR was performed with Q5® High-Fidelity DNA Polymerase (New England Biolabs) per manufacturer’s instructions. For speciation, ITS-2 NC1-forward (5′-ATTGCGCCATCGGGTTCATTCC-3′) and NC2-reverse (5′-TTAGTTTCTTTTCCTCCGCT-3′) primers were used [67]; 1 μl of DNA lysate was used per 25 μl reaction. Cycling conditions included an initial denaturation (98 °C/30 s), 35 cycles (98 °C/15 s, 52 °C/30 s, 72 °C/30 s) and final extension (72 °C/2 min). For beta-tubulin sequences, previously designed forward (5′-NNNACGCACTCTTTGGGAGGAGG-3′) and reverse (5′-NNNTGTGAGTTTTAGTGTGCGGAAG-3′) primers were used that span exons 4 and 5 and intron 4 of the beta-tubulin isotype 1 gene [71]; 1 μl of DNA lysate was used per 25 μl reaction. Cycling conditions included an initial denaturation (98 °C/30 s), 35 cycles (98 °C/15 s, 55 °C/30 s, 72 °C/35 s) and final extension (72 °C/2 min). PCR products were assessed by 1.3% agarose gel electrophoresis with single bands purified by Wizard® SV Gel and PCR Clean-Up System (Promega) and assessed by NanoDrop® ND-1000 (Thermo Fisher). Purified PCR products were sequenced by Eurofins TubeSeq service and assessed for quality and base-calling using Bioedit 7.2.5 software (University of North Texas).

Phylogenetic analyses of ITS-2 sequences

Individual ITS-2 sequences were speciated in Geneious Prime® (2023.1.1 Build 2023-04-03) using the ‘identify organism’ tool to BLAST against the nemabiome ITS-2 database [72]. Sequences and hits with the highest overall sequence query coverage and identity are shown in Additional file 3: Table S2. A single consensus sequence was produced from MUSCLE (PPP) aligned sequences and submitted GenBank for O. leptospicularis (Genbank accession no. OR284984) and S. asymmetrica (Genbank accession no. OR284985).

Phylogenetic trees were produced using previously documented protocols for GIN ITS-2 phylogenetics [71] using Geneious Prime®. Briefly, ingroup and outgroup taxa were selected and aligned by MUSCLE (PPP) with trees produced using Jukes-Cantor model computed with 10,000 bootstrap replicates and rooted on T. axei. Comparison species were selected from known species in wild ruminants [11] alongside other representative Haemonchidae with random representative sequences from the nemabiome ITS-2 database [72]. To compare individual samples, the same method was used except consensus sequences from ingroup taxa were selected to minimize tree size while still comparing sample variance.

Analysis of beta-tubulin sequences

Sanger sequencing was performed as described above for a sub-set of individual specimens. Individual sequences were assessed for the presence of four known markers of BZ resistance, F167Y (TTC/TAC), E198A (GAR/GCA) or E198L (GAR/TTA) and F200Y (TTC/TAC) [58, 71, 73, 74]. Aligned and consensus sequences are shown in Additional file 4: File S1. Consensus sequences were submitted to GenBank for O. leptopiscularis (PP077401) and S. asymmetrica (PP077402).

Data presentation

Unless otherwise stated, original data were tabulated in Microsoft Excel with figure layouts generated in Adobe® Illustrator 2023 (Adobe Inc).

Results

Assessment of gastrointestinal nematodes from fallow deer necropsies

To determine whether deer in Northern Ireland harbour sheep- or cattle-associated GINs, adult worms were collected from abomasa and subjected to Sanger sequencing molecular speciation using NC1/NC2 pan-nematode primers (Additional file 2: Table S1). In abomasa O. leptospicularis and S. asymmetrica coinfections were identified in 4/7 of individuals, with more S. asymmetrica identified overall (Table 1). No other GIN species were identified.

Table 1 Tally of gastrointestinal nematodes identified from deer necropsies
Full size table

Inter-sample sequence variation was minimal for O. leptospicularis (96.43–100% identity, median = 100%) and S. asymmetrica (94.62–100% identity, median = 100%) and all individual samples clustered when compared to other Haemonchidae (Additional file 1: Fig. S1). Consensus sequences for each species yielded mildly distinct variations from known isolates (Fig. 1) and may represent Ireland-specific variants for both O. leptospicularis and S. asymmetrica.

Fig. 1
figure 1

ITS-2 phylogenetic neighbour-joining tree from isolated samples compared to parasites of wild ruminants and similar Haemonchidae species. Random representative sequences selected from the Nemabiome ITS-2 database for reference species. Sequences aligned by MUSCLE (PPP) and organised by the Jukes-Cantor distance model (10,000 bootstrap replicates) with Trichostrongylus axei used as an outgroup. Tree displays species and accession number used with consensus sequences from in-study specimens colour coded. Bootstrap values > 70% shown

Full size image

Cross-validation of sheep and cattle GIN species-specific PCR primers against deer parasites

An array of species-specific primers have previously been designed to detect sheep and cattle GIN species. To determine whether O. leptospicularis or S. asymmetrica yield cross-contamination of sheep and cattle GINs, we performed a PCR screen from published primers [49, 66]. We found that primers for H. contortus, T. circimcincta, T. axei, T. colubriformis and T. vitrinus cross-reacted with both O. leptospicularis and S. asymmetrica, while a primer set for O. ostertagi cross reacted with O. leptospicularis (Table 2, Additional file 2: Table S1, Fig. 2). Overall, the majority of primer sets worked as expected against these wildlife species (Table 2), but false-positive reactions produced single PCR product bands that were indistinguishable in size from positive controls (Fig. 2). One of the primer sets tested for T. circimcincta (‘TeciFd3’) has previously documented cross-reactivity to O. leptospicularis [49].

Table 2 False-positive PCR hits from species-specific primer sets
Full size table
Fig. 2
figure 2

Species-specific PCR screen with false-positive hits against Ostertagia leptospicularis and Spiculopteragia asymmetrica specimen pools. Primer sets shown as outlined in Additional file 2: Table S1 and results summarised in Table 2, with target species and expected PCR product sizes (in brackets) tested against O = O. leptospicularis, S = S. asymmetrica, (+) = positive control DNA, and (−) = negative no template control. Ladder = 100 bp with double-intensity 500-bp band

Full size image

Beta-tubulin sequences from O. leptospicularis and S. asymmetrica

A total of 30 sequences from O. leptospicularis (n = 15) and S. asymmetrica (n = 15) were performed using previously published primers [75]. All samples tested yielded susceptible sequences at positions 167 (TTC), 198 (GAG/GAA) and 200 (TTC) (Additional file 4: File S1). For all O. leptospicularis GAG sequences were found at position 198, while for all S. asymmetrica GAA was found at position 198. Sequences had high identities across individuals with 203/226 matching bases for O. leptospicularis (91.11–100% pairwise identity). For S. asymmetrica 219/230 bases matched (96.52–100% range in pairwise identity) (Additional file 1: File S1).

Discussion

Little is known relating to populations of GINs among Northern Irish fallow deer and whether these GINs harbour AR traits. To elucidate the status of GIN presence among wild fallow deer in Northern Ireland, we investigated deer carcasses from individuals covering three counties inhabited by over two-thirds of the total NI sheep population (67%) and over half the cattle population (53%) [76]. In this study we identified two species commonly associated with cervids, O. leptospicularis and S. asymmetrica among abomasa of seven deer. Both of these species are frequently reported in cervids and occasionally in sheep and cattle, but molecular resources for these species are lacking. To add to available resources, we provide consensus ITS-2 Sanger sequences from a total of 22 O. leptospicularis (OR284984) and 53 S. asymmetrica (OR284985), with individual sequences presented in Additional file 3: Table S2. In addition, we provide the first reported beta-tubulin sequences for both species (PP077401-PP077402) to our knowledge.

In improving genetic resources for these species, we note the importance of including wildlife and less common species in development of molecular diagnostic assays, such as for monitoring the presence of livestock GINs. Interest in molecular diagnostics (particularly PCR based diagnostics) of GINs and other helminths is broad and growing. Methods for detection vary and rely on standardization with available parasite materials. In principle, PCR diagnostics utilise conserved regions of parasite genomes, such as the ITS-2 region of rDNA with species- or genus-specific primers or probes designed at variable sites [49,50,51,52,53,54,55]. PCR-based amplification yields fluorescence from specific probes in loop-mediated isothermal amplification (LAMP) [52, 56], Q-PCR [54, 68, 77] and ddPCR [78], size-specific products in endpoint PCR and semi-quantitative PCR [48, 49] or products with specific melt curves in high-resolution melt curve analyses [48, 55]. Molecular monitoring of GINs can utilize a number of genetic sources such as: (1) adult worms from necropsies [55, 79], (2) faecal DNA [48, 79], (3) isolated eggs from faeces [77], (4) cultured larvae [53, 57], (5) pasture larvae [80] and (6) environmental DNA [81]. However, validation of probe specificity has typically not taken wildlife species into consideration, although multiple studies have been mindful to detect wildlife species [39, 46, 59, 60]. In this study, we test O. leptospicularis and S. asymmetrica DNA from fallow deer in NI against a number of livestock GIN specific primers using endpoint PCR with size specific primers.

While the majority of primers tested produced no cross-reactivity against either species, we generated false-positive hits at expected product sizes for H. contortus (1/5 primers), O. ostertagi (1/4 primers), T. circimcincta (1/5 primers), T. axei (1/3 primers), T. colubriformis (1/3 primers) and T. vitrinus (2/2 primers). We should note that the T. circimcincta primer set (TeciFd3) is known to cross-react to O. leptospicularis [49] and so this finding is not surprising. However, cross-reactivity with both O. leptospicularis and S. asymmetrica is concerning as it raises questions about potential other wildlife species that could contaminate molecular diagnostic approaches. While closely related species such as O. ostertagi might be expected to yield false-positive hits, the variety of species detected is concerning since there is potential for O. leptospicularis and S. asymmetrica DNA to be present in any number of sources.

With deer co-grazing on pasture becoming increasingly prominent, deer GINs may be increasingly deposited on pastures where the eggs or larvae can contaminate faeces collected from grass for faecal egg counts and molecular diagnostics, contaminate pasture larvae counts or contaminate environmental DNA collections. In addition, as O. leptospicularis can cause infections in both cattle and sheep, molecular misidentifications can also be made from worms collected at necropsy. PCR diagnostics have previously been performed on adult worms collected from roe deer, but such studies are robust and have included additional checks such as ITS-2 sequencing to confirm species identity [13]. However, whether PCR alone is sufficient to identify livestock parasites remains to be determined.

Cervid species such as O. leptospicularis may also pose a threat to the performance of cattle [17] and sheep [15, 19] and are capable of altering intestinal pH [16]. Interestingly, distinct O. leptospicularis populations in cattle have been found and possibly adapt to local hosts [10]. Spiculopteragia asymmetrica is also identified in some farmed ruminants such as mouflons [59] but may have capacity to overspill into more livestock species in areas where wildlife are declining or where shared grazing is increasing [5, 82]. Infections of O. leptospicularis in livestock may also produce sterile hybrid offspring with O. ostertagi [18]. This in turn can result in false-positive detection of sheep or cattle parasites by PCR from pasture larvae or pasture collected faecal samples since deer faeces will contaminate the pasture. Furthermore, sterile hybrid infections will also go undetected in faecal egg counts as these have reduced or ablated egg production [18]. However, both O. leptospicularis and S. asymmetrica cross-reacted with the same off-target primer sets, with the exception of O. ostertagi which did not yield a false-positive hit for S. asymmetrica. As such, it is probable that primers can be effectively designed even in the presence of off-target species from wildlife contamination.

Beyond ITS-2 markers we also provide the first beta-tubulin sequences for both O. leptospicularis and S. asymmetrica. Monitoring resistance markers in wildlife is crucial to determine the extent of overlap between anthropogenic and wild habitats and has been applied for species with wild and farmed populations such as bison [75]. A number of single-nucleotide polymorphism (SNP) sites that confer resistance to BZ are known for other GIN species [83], but have yet to be identified in wildlife restricted species. Here we find that beta-tubulin sequences vary in intronic areas but predicted protein coding regions match with predicted and known other nematode protein sequences. No AR SNPs were detected for any samples indicating that despite the small sample size collected, there is no evidence for AR among these species in wild cervids in Northern Ireland currently. While we did not find any evidence of BZ resistance in our sample cohort, we demonstrate that effective monitoring of BZ resistance sites by sequencing is possible as has been reported for livestock species [71]. These sequences also provide a means to develop PCR monitoring for BZ SNPs as previously developed for livestock species [31].

Despite the lack of resistance markers present in the current study, monitoring for AR among wildlife is critical to determine potential toxicological impacts of livestock–wildlife interfaces. For instance, widespread use of anthelmintics has led to ecological contamination in a wide variety of substrates including water [84, 85], soil [42, 86] and plants [42, 86]. However, most common anthelmintics such as BZ, macrocyclic lactones and levamisole are photolabile and will degrade in sunshine, and they may not pose a significant direct effect in sunny summer conditions [87]. Nevertheless, build-up of anthelmintic residues on pasture can lead to auto-dosing of untreated animals [42]. In this case, sheep dosed with albendazole were allowed to excrete on fodder plants. Later feeding of fodder plants from contaminated pasture led to albendazole detected in untreated sheep [42]. As such, it is entirely possible for wildlife species to be similarly impacted if co-grazing in dirty pasture.

Overall, this dataset provides a baseline to monitor BZ resistance markers in O. leptospicularis and S. asymmetrica as well as details of ITS-2 sequences for improved species-specific molecular diagnostics of livestock GINs.

Conclusions

We provide a case report of abomasal worms from seven fallow deer culled in regions with relatively high sheep ownership. Individual adult worms were isolated and speciated, revealing O. leptospicularis and S. asymmetrica. To our surprise, both species yielded cross-reactivity to a number of livestock GIN species using previously published species-specific PCR primers, a result that highlights the need to consider wildlife species when designing molecular diagnostics. Since coinfections of livestock and wildlife are diverse and often overlap, it is critical to build upon molecular resources for wildlife species which are a neglected part of the ecology of parasite dispersal in shared environments.

Availability of data and materials

Baseline data from nematode species counts, PCR results and DNA sequences are presented in the manuscript and supplemental materials.

Abbreviations

AR:

Anthelmintic resistance

GIN:

Gastrointestinal nematode parasite

ITS-2:

Ribosomal internal transcribed spacer 2 region

NI:

Northern Ireland

PCR:

Polymerase chain reaction

References

  1. Apollonio M, Andersen R, Putman R. European ungulates and their management in the 21st century. Cambridge: Cambridge University Press; 2010.

    Google Scholar 

  2. Carden RF, Carlin CM, Marnell F, McElholm D, Hetherington J, Gammell MP. Distribution and range expansion of deer in Ireland. Mamm Rev. 2011;41:313–25.

    Article  Google Scholar 

  3. Ward AI. Expanding ranges of wild and feral deer in Great Britain. Mamm Rev. 2005;35:165–73.

    Article  Google Scholar 

  4. Böhm M, White PC, Chambers J, Smith L, Hutchings MR. Wild deer as a source of infection for livestock and humans in the UK. Vet J. 2007;174:260–76.

    Article  PubMed  Google Scholar 

  5. Brown TL, Airs PM, Porter S, Caplat P, Morgan ER. Understanding the role of wild ruminants in anthelmintic resistance in livestock. Biol Lett. 2022;18:20220057.

    Article  PubMed  PubMed Central  Google Scholar 

  6. Csivincsik Á, Nagy G, Halász T, Zsolnai A. Shared pastures and anthelmintic resistance in wildlife and livestock. Agric Conspec Sci. 2017;82:189–91.

    Google Scholar 

  7. Barone CD, Wit J, Hoberg EP, Gilleard JS, Zarlenga DS. Wild ruminants as reservoirs of domestic livestock gastrointestinal nematodes. Vet Parasitol. 2020;279:109041.

    Article  CAS  PubMed  Google Scholar 

  8. Chintoan-Uta C, Morgan ER, Skuce PJ, Coles GC. Wild deer as potential vectors of anthelmintic-resistant abomasal nematodes between cattle and sheep farms. Proc Biol Sci. 2014;281:20132985.

    CAS  PubMed  PubMed Central  Google Scholar 

  9. Walker JG, Morgan ER. Generalists at the interface: nematode transmission between wild and domestic ungulates. Int J Parasitol Parasit Wildl. 2014;3:242–50.

    Article  Google Scholar 

  10. Wyrobisz-Papiewska A, Kowal J, Łopieńska-Biernat E, Nosal P, Polak I, Paukszto Ł, et al. Morphometric and molecular analyses of Ostertagia leptospicularis Assadov, 1953 from ruminants: species diversity or host influence? Animals. 2021;11:182.

    Article  PubMed  PubMed Central  Google Scholar 

  11. Wyrobisz-Papiewska A, Kowal J, Nosal P, Chovancová G, Rehbein S. Host specificity and species diversity of the Ostertagiinae Lopez-Neyra, 1947 in ruminants: a European perspective. Parasit Vectors. 2018;11:369.

    Article  PubMed  PubMed Central  Google Scholar 

  12. Davidson W, McGhee M, Nettles V, Chappell L. Haemonchosis in white-tailed deer in the southeastern United States. J Wildl Dis. 1980;16:499–508.

    Article  CAS  PubMed  Google Scholar 

  13. Lehrter V, Jouet D, Liénard E, Decors A, Patrelle C. Ashworthius sidemi Schulz, 1933 and Haemonchus contortus (Rudolphi, 1803) in cervids in France: integrative approach for species identification. Infect Genet Evol. 2016;46:94–101.

    Article  PubMed  Google Scholar 

  14. Nagy G, Csivincsik Á, Zsolnai A, Sugár L. Benzimidazole resistance in Haemonchus contortus recovered from farmed red deer. Parasitol Res. 2016;115:3643–7.

    Article  PubMed  Google Scholar 

  15. Eysker M, Jansen J. Population build up of gastrointestinal nematode infections in ewes and lambs on pasture grazed by calves in the previous year. Res Vet Sci. 1982;32:203–5.

    Article  CAS  PubMed  Google Scholar 

  16. Hertzberg H, Guscetti F, Lischer C, Kohler L, Neiger R, Eckert J. Evidence for a parasite-mediated inhibition of abomasal acid secretion in sheep infected with Ostertagia leptospicularis. Vet J. 2000;159:238–51.

    Article  CAS  PubMed  Google Scholar 

  17. Mckenna PB. Checklist of helminth parasites of terrestrial mammals in New Zealand. N Z J Zool. 1997;24:277–90.

    Article  Google Scholar 

  18. Suarez V, Durette-Desset M, Cabaret J. Description of Ostertagia ostertagi and Ostertagia leptospicularis hybrids in experimentally infected sheep. J Parasitol. 1993;79:874–8.

    Article  CAS  PubMed  Google Scholar 

  19. Vlassoff A, McKenna PB. Nematode parasites of economic importance in sheep in New Zealand. N Z J Zool. 1994;21:1–8.

    Article  Google Scholar 

  20. Charlier J, Rinaldi L, Musella V, Ploeger HW, Chartier C, Vineer HR, et al. Initial assessment of the economic burden of major parasitic helminth infections to the ruminant livestock industry in Europe. Prev Vet Med. 2020;182:105103.

    Article  CAS  PubMed  Google Scholar 

  21. Baiak BHB, Lehnen CR, da Rocha RA. Anthelmintic resistance in cattle: a systematic review and meta-analysis. Livest Sci. 2018;217:127–35.

    Article  Google Scholar 

  22. Charlier J, Bartley D, Sotiraki S, Martinez-Valladares M, Claerebout E, Von Samson-Himmelstjerna G, et al. Anthelmintic resistance in ruminants: challenges and solutions. Adv Parasitol. 2022;115:171–227.

    Article  CAS  PubMed  Google Scholar 

  23. Coles G. Anthelmintic resistance in sheep. Vet Clin N Am Food Anim Pract. 1986;2:423–32.

    Article  CAS  Google Scholar 

  24. Morgan ER, Charlier J, Hendrickx G, Biggeri A, Catalan D, Von Samson-Himmelstjerna G, et al. Global change and helminth infections in grazing ruminants in Europe: impacts, trends and sustainable solutions. Agriculture. 2013;3:484–502.

    Article  Google Scholar 

  25. Mphahlele M, Molefe N, Tsotetsi-Khambule A, Oriel T. Anthelmintic resistance in livestock. Helminthiasis. IntechOpen. (2019), https://doi.org/10.5772/intechopen.87124.

  26. Sankar M, Yashica K, Ghosh S. Strategies to reduce anthelmintic resistance in livestock. Indian J Comp Microbiol Immunol Infect Dis. 2021;42:32–40.

    Google Scholar 

  27. Sutherland IA, Leathwick DM. Anthelmintic resistance in nematode parasites of cattle: a global issue? Trends Parasitol. 2011;27:176–81.

    Article  PubMed  Google Scholar 

  28. Vineer HR, Morgan ER, Hertzberg H, Bartley DJ, Bosco A, Charlier J, et al. Increasing importance of anthelmintic resistance in European livestock: creation and meta-analysis of an open database. Parasite. 2020;27:69.

    Article  Google Scholar 

  29. Waller P. The development of anthelmintic resistance in ruminant livestock. Acta Trop. 1994;56:233–43.

    Article  CAS  PubMed  Google Scholar 

  30. Erez MS, Kozan E. Anthelmintic resistance in farm animals. Kocatepe Vet J. 2018;11:322–30.

    Google Scholar 

  31. Geary TG. Ivermectin 20 years on: maturation of a wonder drug. Trends Parasitol. 2005;21:530–2.

    Article  CAS  PubMed  Google Scholar 

  32. Kaplan RM. Drug resistance in nematodes of veterinary importance: a status report. Trends Parasitol. 2004;20:477–81.

    Article  CAS  PubMed  Google Scholar 

  33. Hoskin S, Pomroy W, Wilson P, Ondris M, Mason P. The efficacy of oral ivermectin, pour-on ivermectin and pour-on moxidectin against naturally acquired infections of lungworm and gastrointestinal parasites in young farmed deer. Deer Branch Course. 2005;22:21.

    Google Scholar 

  34. Lawrence D. Cervine anthelmintics-the bubble has burst. Proc Deer Branch NZVA. 2011;28:87–92.

    Google Scholar 

  35. Lawrence D, Macgibbon J, Mason P. Moxidectin pharmacokinetics and resistance in deer. Proc Deer Branch NZVA. 2012;29:41–5.

    Google Scholar 

  36. Lawrence D, Macgibbon J, Mason P. Efficacy of levamisole, moxidectin oral, moxidectin injectable and monepantel against ostertagia-type nematodes in deer. In: Proceedings of the NZVA conference; 2013. p. 233–240.

  37. Leathwick DM, Mason PC, Fraser K, Miller CM, Green P. Route of administration affects the efficacy of moxidectin against Ostertagiinae nematodes in farmed red deer (Cervus elaphus). Vet Parasitol. 2021;298:109525.

    Article  CAS  PubMed  Google Scholar 

  38. Pyziel-Serafin AM, Vetter W, Klich D, Anusz K. Exchanged communities of abomasal nematodes in cervids with a first report on in red deer. J Vet Res. 2023;67:87–92.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  39. Beaumelle C, Redman E, Verheyden H, Jacquiet P, Bégoc N, Veyssière F, et al. Generalist nematodes dominate the nemabiome of roe deer in sympatry with sheep at a regional level. Int J Parasitol. 2022;52:751–61.

    Article  CAS  PubMed  Google Scholar 

  40. Verheyden H, Richomme C, Sevila J, Merlet J, Lourtet B, Chaval Y, et al. Relationship between the excretion of eggs of parasitic helminths in roe deer and local livestock density. J Helminthol. 2020;94:e159.

    Article  CAS  PubMed  Google Scholar 

  41. Wit J, Rodriguez BC, Andersen EC. Natural variation in Caenorhabditis elegans responses to the anthelmintic emodepside. Int J Parasitol Drugs Drug Resist. 2021;16:1–8.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  42. Navrátilová M, Raisová Stuchlíková L, Matoušková P, Ambrož M, Lamka J, Vokřál I, et al. Proof of the environmental circulation of veterinary drug albendazole in real farm conditions. Environ Pollut. 2021;286:117590.

    Article  PubMed  Google Scholar 

  43. Prasse C, Löffler D, Ternes TA. Environmental fate of the anthelmintic ivermectin in an aerobic sediment/water system. Chemosphere. 2009;77:1321–5.

    Article  ADS  CAS  PubMed  Google Scholar 

  44. Van Wyk JA. Refugia—overlooked as perhaps the most potent factor concerning the development of anthelmintic resistance. Onderstepoort J Vet Res. 2001;68:55–67.

    PubMed  Google Scholar 

  45. Vande Velde F, Charlier J, Claerebout E. Farmer behavior and gastrointestinal nematodes in ruminant livestock—uptake of sustainable control approaches. Front Vet Sci. 2018;5:255.

    Article  PubMed  PubMed Central  Google Scholar 

  46. Beaumelle C, Redman EM, de Rijke J, Wit J, Benabed S, Debias F, et al. Metabarcoding in two isolated populations of wild roe deer (Capreolus capreolus) reveals variation in gastrointestinal nematode community composition between regions and among age classes. Parasit Vectors. 2021;14:594.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  47. van Wyk JA, Mayhew E. Morphological identification of parasitic nematode infective larvae of small ruminants and cattle: a practical lab guide. Onderstepoort J Vet Res. 2013;80:1–14.

    Google Scholar 

  48. Airs PM, Ventura-Cordero J, Mvula W, Takahashi T, van Wyk J, Nalivata P, et al. Low-cost molecular methods to characterise gastrointestinal nematode co-infections of goats in Africa. Parasit Vectors. 2023;16:1–15.

    Article  Google Scholar 

  49. Bisset S, Knight J, Bouchet C. A multiplex PCR-based method to identify strongylid parasite larvae recovered from ovine faecal cultures and/or pasture samples. Vet Parasitol. 2014;200:117–27.

    Article  CAS  PubMed  Google Scholar 

  50. Francis EK, McKay-Demeler J, Calvani NED, McDonell D, Šlapeta J. Which larvae are they? Use of single larva for the molecular confirmation of Cooperia pectinata and Cooperia punctata in Australian cattle. Vet Parasitol. 2020;278:109033.

    Article  CAS  PubMed  Google Scholar 

  51. Högberg N, Lidfors L, Hessle A, Segerkvist KA, Herlin A, Höglund J. Effects of nematode parasitism on activity patterns in first-season grazing cattle. Vet Parasitol. 2019;276:100011.

    Article  Google Scholar 

  52. Ljungström S, Melville L, Skuce PJ, Höglund J. Comparison of four diagnostic methods for detection and relative quantification of Haemonchus contortus eggs in feces samples. Front Vet Sci. 2018;4:239.

    Article  PubMed  PubMed Central  Google Scholar 

  53. Redman E, Queiroz C, Bartley DJ, Levy M, Avramenko RW, Gilleard JS. Validation of ITS-2 rDNA nemabiome sequencing for ovine gastrointestinal nematodes and its application to a large scale survey of UK sheep farms. Vet Parasitol. 2019;275:108933.

    Article  CAS  PubMed  Google Scholar 

  54. Roeber F, Morrison A, Casaert S, Smith L, Claerebout E, Skuce P. Multiplexed-tandem PCR for the specific diagnosis of gastrointestinal nematode infections in sheep: an European validation study. Parasit Vectors. 2017;10:1–11.

    Article  Google Scholar 

  55. Skorpikova L, Reslova N, Magdalek J, Vadlejch J, Kasny M. The use of high resolution melting analysis of ITS-1 for rapid differentiation of parasitic nematodes Haemonchus contortus and Ashworthius sidemi. Sci Rep. 2020;10:15984.

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  56. Melville L, Kenyon F, Javed S, McElarney I, Demeler J, Skuce P. Development of a loop-mediated isothermal amplification (LAMP) assay for the sensitive detection of Haemonchus contortus eggs in ovine faecal samples. Vet Parasitol. 2014;206:308–12.

    Article  CAS  PubMed  Google Scholar 

  57. Avramenko RW, Redman EM, Lewis R, Yazwinski TA, Wasmuth JD, Gilleard JS. Exploring the gastrointestinal “Nemabiome”: deep amplicon sequencing to quantify the species composition of parasitic nematode communities. PLoS ONE. 2015;10:e0143559.

    Article  PubMed  PubMed Central  Google Scholar 

  58. Redman E, Whitelaw F, Tait A, Burgess C, Bartley Y, Skuce PJ, et al. The emergence of resistance to the benzimidazole anthlemintics in parasitic nematodes of livestock is characterised by multiple independent hard and soft selective sweeps. PLoS Negl Trop Dis. 2015;9:e0003494.

    Article  PubMed  PubMed Central  Google Scholar 

  59. Halvarsson P, Baltrušis P, Kjellander P, Höglund J. Parasitic strongyle nemabiome communities in wild ruminants in Sweden. Parasit Vectors. 2022;15:341.

    Article  PubMed  PubMed Central  Google Scholar 

  60. Halvarsson P, Höglund J. Sheep nemabiome diversity and its response to anthelmintic treatment in Swedish sheep herds. Parasit Vectors. 2021;14:114.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  61. Laca-Megyesi Š, Königová A, Babják M, Molnár L, Rajský M, Szestáková E, et al. Wild ruminants as a potential risk factor for transmission of drug resistance in the abomasal nematode Haemonchus contortus. Eur J Wildl Res. 2020;66:1–6.

    Article  Google Scholar 

  62. Móónica S-D. Abomasal parasites in wild sympatric cervids, red deer, cervus elaphus and fallow deer, dama dama, from three localities across central and western spain: relationship to host density and park management. J Parasitol. 2004;90:1378–86.

    Article  Google Scholar 

  63. Amarante MRV, Bassetto CC, Neves JH, Amarante AFT. Species-specific PCR for the identification of Cooperia curticei (Nematoda: Trichostrongylidae) in sheep. J Helminthol. 2014;88:447–52.

    Article  CAS  PubMed  Google Scholar 

  64. Amarante MRV, Santos MC, Bassetto CC, Amarante AFT. PCR primers for straightforward differentiation of Haemonchus contortus, Haemonchus placei and their hybrids. J Helminthol. 2017;91:757–61.

    Article  CAS  PubMed  Google Scholar 

  65. Bandyopadhyay S, Naskar S, Devi P, Bera AK, De S, Pan D, et al. Multiplex PCR for identification of parasitic eggs of Oesophagostomum spp. Isolated from sheep and goat. Proc Zool Soc. 2009;62:125–9.

    Article  Google Scholar 

  66. Burgess CGS, Bartley Y, Redman E, Skuce PJ, Nath M, Whitelaw F, et al. A survey of the trichostrongylid nematode species present on UK sheep farms and associated anthelmintic control practices. Vet Parasitol. 2012;189:299–307.

    Article  CAS  PubMed  Google Scholar 

  67. Gasser RB, Chilton NB, Hoste H, Beveridge I. Rapid sequencing of rDNA from single worms and eggs of parasitic helminths. Nucleic Acids Res. 1993;21:2525–6.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  68. Höglund J, Engström A, Von Samson-Himmelstjerna G, Demeler J, Tydén E. Real-time PCR detection for quantification of infection levels with Ostertagia ostertagi and Cooperia oncophora in cattle faeces. Vet Parasitol. 2013;197:251–7.

    Article  PubMed  Google Scholar 

  69. Wimmer B, Craig BH, Pilkington JG, Pemberton JM. Non-invasive assessment of parasitic nematode species diversity in wild Soay sheep using molecular markers. Int J Parasitol. 2004;34:625–31.

    Article  CAS  PubMed  Google Scholar 

  70. Zarlenga DS, Barry Chute M, Gasbarre LC, Boyd PC. A multiplex PCR assay for differentiating economically important gastrointestinal nematodes of cattle. Vet Parasitol. 2001;97:201–11.

    Article  Google Scholar 

  71. Avramenko RW, Redman EM, Melville L, Bartley Y, Wit J, Queiroz C, et al. Deep amplicon sequencing as a powerful new tool to screen for sequence polymorphisms associated with anthelmintic resistance in parasitic nematode populations. Int J Parasitol. 2019;49:13–26.

    Article  CAS  PubMed  Google Scholar 

  72. Workentine ML, Chen R, Zhu S, Gavriliuc S, Shaw N, Rijke JD, et al. A database for ITS2 sequences from nematodes. BMC Genet. 2020;21:74.

    Article  PubMed  PubMed Central  Google Scholar 

  73. Sargison ND, Macleay M, Morrison AA, Bartley DJ, Evans M, Chaudhry U. Development of amplicon sequencing for the analysis of benzimidazole resistance allele frequencies in field populations of gastrointestinal nematodes. Int J Parasitol Drugs Drug Resist. 2019;10:92–100.

    Article  PubMed  PubMed Central  Google Scholar 

  74. Von Samson-Himmelstjerna G, Blackhall WJ, McCarthy JS, Skuce PJ. Single nucleotide polymorphism (SNP) markers for benzimidazole resistance in veterinary nematodes. Parasitology. 2007;134:1077–86.

    Article  Google Scholar 

  75. Avramenko RW, Redman EM, Windeyer C, Gilleard JS. Assessing anthelmintic resistance risk in the post-genomic era: a proof-of-concept study assessing the potential for widespread benzimidazole-resistant gastrointestinal nematodes in North American cattle and bison. Parasitology. 2020;147:897–906.

    Article  PubMed  PubMed Central  Google Scholar 

  76. DAERA. Agricultural Census in Northern Ireland 2022_Data Tables-1. In: DEPARTMENT FOR AGRICULTURE, E. A. R. A., editor. NI Direct: Gov.UK; 2022.

  77. McNally J, Callan D, Andronicos N, Bott N, Hunt PW. DNA-based methodology for the quantification of gastrointestinal nematode eggs in sheep faeces. Vet Parasitol. 2013;198:325–35.

    Article  CAS  PubMed  Google Scholar 

  78. Elmahalawy ST, Halvarsson P, Skarin M, Höglund J. Droplet digital polymerase chain reaction (ddPCR) as a novel method for absolute quantification of major gastrointestinal nematodes in sheep. Vet Parasitol. 2018;261:1–8.

    Article  CAS  PubMed  Google Scholar 

  79. Reslova N, Skorpikova L, Kyrianova IA, Vadlejch J, Höglund J, Skuce P, et al. The identification and semi-quantitative assessment of gastrointestinal nematodes in faecal samples using multiplex real-time PCR assays. Parasit Vectors. 2021;14:391.

    Article  CAS  PubMed  PubMed Central  Google Scholar 

  80. Mcfarland C, Rose Vineer H, Chesney L, Henry N, Brown C, Airs P, et al. Tracking gastrointestinal nematode risk on cattle farms through pasture contamination mapping. Int J Parasitol. 2022;52:691–703.

    Article  PubMed  Google Scholar 

  81. Chan AHE, Saralamba N, Saralamba S, Ruangsittichai J, Chaisiri K, Limpanont Y, et al. Sensitive and accurate DNA metabarcoding of parasitic helminth mock communities using the mitochondrial rRNA genes. Sci Rep. 2022;12:9947.

    Article  ADS  CAS  PubMed  PubMed Central  Google Scholar 

  82. Morellet N, van Moorter B, Cargnelutti B, Angibault J-M, Lourtet B, Merlet J, et al. Landscape composition influences roe deer habitat selection at both home range and landscape scales. Landsc Ecol. 2011;26:999–1010.

    Article  Google Scholar 

  83. Evans MJ, Chaudhry UN, Costa-Júnior LM, Hamer K, Leeson SR, Sargison ND. A 4 year observation of gastrointestinal nematode egg counts, nemabiomes and the benzimidazole resistance genotypes of Teladorsagia circumcincta on a Scottish sheep farm. Int J Parasitol. 2021;51:393–403.

    Article  CAS  PubMed  Google Scholar 

  84. Crane M, Boxall ABA, Barrett K, Environmet S. P. W. O. V. M. I. T. Veterinary medicines in the environment: from the SETAC Pellston workshop on veterinary medicines in the environment, Pensacola, Florida, USA, 12–16 February 2006, SETAC. CRC Press; 2009.

  85. Sim WJ, Kim HY, Choi SD, Kwon JH, Oh JE. Evaluation of pharmaceuticals and personal care products with emphasis on anthelmintics in human sanitary waste, sewage, hospital wastewater, livestock wastewater and receiving water. J Hazard Mater. 2013;248–249:219–27.

    Article  PubMed  Google Scholar 

  86. Iglesias LE, Saumell C, Sagüés F, Sallovitz JM, Lifschitz AL. Ivermectin dissipation and movement from feces to soil under field conditions. J Environ Sci Health B. 2018;53:42–8.

    Article  CAS  PubMed  Google Scholar 

  87. Horvat AJM, Babić S, Pavlović DM, Ašperger D, Pelko S, Kaštelan-Macan M, et al. Analysis, occurrence and fate of anthelmintics and their transformation products in the environment. Trends Anal Chem: TrAC. 2012;31:61–84.

    Article  CAS  Google Scholar 

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Acknowledgements

The authors would like to acknowledge the Forestry Service Northern Ireland and Donald Whiteside for the kind provision of sample materials and AFBINI for their support.

Funding

This work was supported by DAERA’s annual postgraduate studentship (M Lyons & T Brown) and UKRI grant BB/S014748/1. For the purpose of open access, the author has applied a Creative Commons Attribution (CC BY) licence to any Author Accepted Manuscript version arising.

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Authors and Affiliations

  1. School of Biological Sciences, Queen’s University Belfast, 19 Chlorine Gardens, Belfast, BT9 5DL, UK

    Maggie Lyons, Tony L. Brown, Eric. R. Morgan & Paul M. Airs

  2. Agri-Food and Biosciences Institute Northern Ireland, 12 Stoney Road, Belfast, Co Antrim, BT4 3SD, UK

    Maggie Lyons & Angela Lahuerta-Marin

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  1. Maggie Lyons
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Contributions

ML—Data curation, Formal Analysis, Investigation, Writing—original draft, Writing—review & editing, TLB—Data curation, Investigation, Methodology, Parasite extraction, Writing—review & editing, AL-M—Supervision, Writing—review & editing. ERM—Conceptualization, Funding acquisition, Investigation, Methodology, Project administration, Supervision, Writing—original draft, Writing—review & editing, PMA—Conceptualization, Data curation, Formal Analysis, Investigation, Methodology, Software, Visualization, Writing—original draft, Writing—review & editing.

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Correspondence to Paul M. Airs.

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The manuscript does not contain clinical studies, patient data or individual data and is an observational study of parasites from waste tissue following culling of fallow deer by trained and licenced stalkers in Northern Ireland, in accordance with legislation and within the designated open season.

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Supplementary Information

Additional file 1. File S1

-  Multiple sequence alignments and identity matrices for Beta tubulin isoform 1 sequences.

Additional file 2. Table S1

- Primers and cycling conditions used for PCR.

Additional file 3. Table S2

- ITS-2 Sanger sequences for each individual analysed.

Additional file 4. Supplementary Materials

- Supplementary Figures and Table headings.

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Lyons, M., Brown, T.L., Lahuerta-Marin, A. et al. A molecular assessment of Ostertagia leptospicularis and Spiculopteragia asymmetrica among wild fallow deer in Northern Ireland and implications for false detection of livestock-associated species. Parasites Vectors 17, 141 (2024). https://doi.org/10.1186/s13071-024-06147-2

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Parasites & Vectors

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