Hostname: page-component-745bb68f8f-g4j75 Total loading time: 0 Render date: 2025-02-11T14:33:06.413Z Has data issue: false hasContentIssue false
  • English
  • Français

Semi-nested PCR for the specific detection of Habronema microstoma or Habronema muscae DNA in horse faeces

Published online by Cambridge University Press:  18 November 2004

D. TRAVERSA ,
A. GIANGASPERO ,
R. IORIO ,
D. OTRANTO ,
B. PAOLETTI  and
R. B. GASSER
D. TRAVERSA
Affiliation:
Department of Biomedical Comparative Sciences, Faculty of Veterinary Medicine, University of Teramo, Teramo, Italy
A. GIANGASPERO
Affiliation:
Department of Production Science, Engineering, Mechanics and Economy, Faculty of Agronomy, University of Foggia, Foggia, Italy
R. IORIO
Affiliation:
Department of Biomedical Comparative Sciences, Faculty of Veterinary Medicine, University of Teramo, Teramo, Italy
D. OTRANTO
Affiliation:
Department of Animal Health and Welfare, Faculty of Veterinary Medicine, University of Bari, Valenzano, Bari, Italy
B. PAOLETTI
Affiliation:
Department of Biomedical Comparative Sciences, Faculty of Veterinary Medicine, University of Teramo, Teramo, Italy
R. B. GASSER
Affiliation:
Department of Veterinary Science, The University of Melbourne, Werribee, Victoria 3030, Australia
Rights & Permissions [Opens in a new window]

Abstract

Habronema microstoma and Habronema muscae (Spirurida: Habronematidae) are parasitic nematodes which infect the stomach and/or skin of equids. The accurate diagnosis of gastric habronemosis is central to studying its epidemiology, but data on its distribution and prevalence are lacking, mainly due to the limitations of clinical and coprological diagnosis in live horses. To overcome this constraint, a two-step, semi-nested PCR-based assay was validated (utilizing genetic markers in the nuclear ribosomal DNA) for the specific amplification of H. microstoma or H. muscae DNA from the faeces from horses (n=46) whose gastrointestinal parasite status had been determined at autopsy and whose faeces were examined previously using a conventional parasitological approach. Of these horses examined at autopsy, some harboured adults of either H. microstoma (n=19) or H. muscae (n=4), and others (n=7) harboured both species. Most of them were also infected with other parasites, including strongylid nematodes (subfamilies Cyathostominae and Strongylinae), bots and/or cestodes; there was no evidence of metazoan parasites in 2 horses. Larvated spirurid eggs were detected in the faeces of 1 of the 30 horses (3·3%) shown to be infected with Habronema at autopsy. For this set of 46 samples, the PCR assay achieved a diagnostic specificity of 100% and a sensitivity of ~97% (being able to specifically detect as little as ~0·02 fg of Habronema DNA). The specificity of the assay was also tested using a panel of control DNA samples representing horse, the gastric spirurid Draschia megastoma and 26 other species of parasites from the alimentary tract of the horse. H. microstoma, H. muscae and D. megastoma could be readily differentiated from one another based on the sizes of their specific amplicons in the PCR. The results of this study showed that the performance of the PCR for the diagnosis of gastric habronemosis was similar to that of autopsy but substantially better than the traditional coprological examination procedure used. The ability to specifically diagnose gastric habronemosis in equids should have important implications for investigating the epidemiology and ecology of H. microstoma and H. muscae.

Keywords

coprological diagnosisepidemiologyhorseHabronema microstomaHabronema muscaesecond internal transcribed spacer of ribosomal DNAsemi-nested PCR
Type
Research Article
Information
Parasitology , Volume 129 , Issue 6 , December 2004 , pp. 733 - 739
Copyright
© 2004 Cambridge University Press

INTRODUCTION

The parasitic nematodes Habronema microstoma Creplin, 1849 and Habronema muscae Carter, 1865 (Spirurida: Habronematidae) cause gastric, cutaneous and/or muco-cutaneous habronemosis in equids (i.e., horses, donkeys, mules and zebras) (Owen & Slocombe, 1985; Anderson, 2000). These dioecious parasites mate in the stomach of the equid (definitive) host. After mating, the females produce embryonated eggs which are released via the faeces into the environment. In the faeces, the embryonated eggs or developing nematode larvae (during their dung-dwelling phase) are ingested by an intermediate host, a muscid fly, such as Musca domestica or Stomoxys calcitrans (see Roubaud & Descauzeaux, 1922; de Jesus, 1963; Dunn, 1969; Anderson, 2000). In the fly, the nematode undergoes larval development from the first (L1) to the third stage (L3), which is synchronous with the development of the fly. The infective L3s are deposited by the fly imago on to the moist skin (e.g., around nostrils, mouth, skin lesions, eyes and/or genitalia) of the definitive host when the imago feeds (Roubaud & Descauzeaux, 1922). When deposited around the nostrils and/or mouth, the L3s are swallowed, after which they develop into adults in the stomach (hence, gastric habronemosis). When deposited at other cutaneous sites or muco-cutaneous transitions, the parasites remain as larvae and do not reach sexual maturity. These larvae cause local inflammatory responses that relate to cutaneous and/or muco-cutaneous habronemosis.

Habronemosis caused by H. microstoma and/or H. muscae is considered to have a global distribution (Lyons et al. 1983; Pandey, Ohuelli & Verhulst, 1992; Borgsteede & van Beek, 1998; cf. Traversa et al. 2004). However, there are still significant knowledge gaps regarding the ecology and epidemiology of these parasites, mainly due to diagnostic limitations. Clinical diagnosis of both gastric and cutaneous forms of habronemosis is unreliable, because symptoms associated with gastric infection may ‘overlap’ with other types of colic (cf. Euzéby, 1961; Soulsby, 1982), and the cutaneous granulomatous lesions are similar to those of other diseases (see Inzana & Carter, 1990; Pascoe, 1990; Chaffin, Schumacher & McMullan, 1995). Traditionally, surveys of habronemosis have relied on the post-mortem examination of equids for adult worms in the stomach (e.g., Hass, 1979; Lyons et al. 1983, 1987, 1994; Pandey et al. 1992; Bucknell, Gasser & Beveridge, 1995) because diagnosis in live equids has major limitations. While adults of the spirurid Draschia megastoma (which usually induce large, fibrotic nodules in the stomach of equids) can be readily identified based on a funnel-shaped buccal capsule, the morphological identification and differentiation of species of Habronema, based on the measurements of the cylindrical buccal capsule, is very challenging. Coprological diagnosis of gastric habronemosis, based on the detection of larvated eggs by flotation or the detection of larvae employing the Baermann technique or coproculture, has a very low sensitivity, even for horses with a high intensity of infection (unpublished findings), which is considered to relate largely to the low biotic potential of species of Habronema (cf. Euzéby, 1961; Dunn, 1969; Soulsby, 1982). Even when eggs or larvae are detected, they cannot be identified to species. Another method is the gastric lavage (cf. Euzéby, 1961; Dunn, 1969; Soulsby, 1982) followed by the microscopical examination of sedimented washings for larval or adult stages, but this approach requires sedation and restraint of the horse, is invasive (and thus risky) and time consuming. Hence, given these limitations, there is significant merit in developing improved diagnostic tools.

DNA methods based on the polymerase chain reaction (PCR), employing species-specific genetic markers, have proven particularly applicable to the copro-diagnosis of a number of different parasitic nematode infections (e.g., Gasser, 1999; Hung et al. 1999; Verweij et al. 2000, 2001). Extending from previous work, we recently demonstrated that the sequences of the second internal transcribed spacer (ITS-2) of nuclear ribosomal DNA allowed the accurate identification of H. microstoma and H. muscae, enabling the construction of species-specific primers (within the ITS-2) for use in the PCR (Traversa et al. 2004). In the present study, we established and assessed a semi-nested PCR assay using these primers for the detection of H. microstoma and/or H. muscae DNA in equine faeces, as a molecular epidemiological and ecological tool.

MATERIALS AND METHODS

From February to December 2003, the gastro-intestinal tracts of 46 native horses (both sexes; 1–9 years of age), slaughtered in accordance with the animal welfare and ethics laws in Italy in the abattoirs of Pescara, Chieti and L'Aquila (Abruzzo region, central Italy), were examined for the presence of metazoan parasites. Macroscopic examination did not reveal any metazoan parasites in sites (including lung, liver and peritoneal cavity) external to the gastro-intestinal tract. The nematodes collected from the gastro-intestinal tracts were cleared in lactophenol and identified morphologically, according to existing descriptions and keys (Euzéby, 1961, 1981; Lichtenfels, 1975; Tolliver, 2000). Other parasites, such as bots (species of Gasterophilus) and cestodes were identified according to their site of predilection in the host and characteristic morphological features (Zumpt, 1965; Euzéby, 1981; Schmidt, 1986). Of the 46 horses, 30 (nos 1–30) were infected with H. microstoma and/or H. muscae (3 to 148 adults per horse) (Table 1). The vast majority (97%) of them also harboured bots in the stomach and/or strongylid nematodes (subfamilies Cyathostominae and/or Strongylinae) in the large intestine. There was no evidence of habronemosis in 14 horses (nos 31–44), of which 1 had Trichostrongylus axei in the stomach, 3 had Anoplocephala perfoliata in the ileo-caecal junction or caecum, 1 had Anoplocephala magna in the small intestine and 1 had Oxyuris equi in the large intestine. No metazoan parasites were detected in 2 horses (nos 45 and 46) (Table 1).

A faecal sample (~300 g) was collected (post-mortem) from the rectum from each of the 46 horses, and divided into a 100 g aliquot for coprological examinations (within 24 h) and a 200 g aliquot for molecular analysis (stored at −20 °C until use). Individual faecal samples were subjected to flotations (Euzéby, 1981) using saturated solutions of each NaCl, ZnSO4 and HgI2/KI, with specific gravities of 1·2, 1·35 and 1·45, respectively. For this method, ~23 g of faeces were added to 60 ml of flotation solution, centrifuged at 600 g for 5 min, and an aliquot (~100 μl) of supernatant was aspirated (with a Pasteur pipette), transferred to a glass slide and examined using a light microscope (Axioscope 40, Zeiss, Oberkochen, Germany) at a magnification of 200 times. Of the 46 samples examined, larvated spirurid eggs were detected in one sample (no. 22) using the HgI2/KI flotation solution (Table 1).

The 46 faecal samples obtained from horses (Table 1) were then processed for molecular analysis. For each, 10~20 g aliquots were subjected to the flotation procedure in 15 ml tubes using ZnSO4 solution, the rationale being to enrich eggs, larvae and/or DNA from the faecal samples. For each flotation, 40 μl of supernatant were aspirated using a micropipette, and all ten 40 μl aliquots pooled into an Eppendorf tube. Each tube (containing the 400 μl) was exposed to 3 freeze-thaw cycles (3 min at −196 °C and 80 °C, respectively). Subsequently, genomic DNA was extracted from individual samples using the QIAamp® DNA Stool Mini Kit (QIAgen, Hilden, Germany) and stored at −20 °C. In addition to the 46 DNA samples extracted from the faecal samples, a panel of 48 control DNA samples representing the equine host (n=2) and representing 27 different species of parasites, including D. megastoma, T. axei (Nematoda), Gasterophilus intestinalis (Arthropoda), Fasciola hepatica (Trematoda), A. perfoliata, A. magna, Paranoplocephala mammillana (Cestoda), Parascaris equorum, O. equi and a range of species of Strongylinae and Cyathostominae (Nematoda), including Strongylus vulgaris, Strongylus edentatus, Strongylus equinus, Triodontophorus brevicauda, Triodontophorus serratus, Triodontophorus tenuicollis, Coronocyclus coronatus, Cyathostomum catinatum, Cyathostomum pateratum, Cylicocyclus insignis, Cylicocyclus leptostomus, Cylicocyclus nassatus, Cylicocyclus brevicapsulatus, Cylicodontophorus euproctus, Cylicostephanus calicatus, Cylicostephanus goldi, Cylicostephanus longibursatus and Cylicostephanus poculatus (see Hung et al. 1999; Traversa et al. 2004), were included to test the specificity of the two-step, semi-nested PCR established. The absence of PCR inhibition (by faecal contaminants) in individual samples was verified using an approach described previously (see Traversa et al. 2004).

Regions of ITS-2 were amplified from these DNA samples using a two-step, semi-nested PCR protocol. The PCR conditions were optimized by titration of MgCl2 and dNTP concentrations, and varying annealing temperatures, cycle numbers and times. PCR (in 50 μl) was performed in 10 mM Tris–HCl, pH 8·4; 50 mM KCl; 2·5 mM MgCl2; 0·5 μg bovine serum albumin, 250 μM of each dNTP; 50 or 100 pmol of each primer with 1.25 U AmpliTaqGold (Applied Biosystems, Foster, CA, USA). Primer set D (5′-GAGTCGATGAAGAACGCAG-3′) – B1 (5′-GAATCCTGGTTAGTTTCTTTTCCT-3′) was used in a first round of PCR, followed by a second round of PCR using primer set Hmi (5′-GATCGCAATATGTGTAACAC-3′) – B1 or Hmu (5-CTGGTAAAGCATCAATGCATCAGGTATG-3) – B1 (Traversa et al. 2004); the specificity of these primer sets in a single-step PCR was reported previously (Traversa et al. 2004). Cycling was performed as follows: initial denaturation at 94 °C for 12 min (required for Taq Gold activation), followed by 30 cycles (first round) or 35 (second round) of 94 °C for 30 s (denaturation), 58 °C for 45 s (annealing) and 72 °C for 45 s (extension), followed by a final extension at 72 °C for 7 min. In the first round of PCR, 2 μl of DNA template were included in the reaction. In the second round, an aliquot of each amplicon (cf. results) was subjected under the same cycling conditions. Samples without DNA template (no-DNA controls) and a known positive control (DNA extracted from H. microstoma or H. muscae specimens) were included in each amplification run. PCR products were subjected to electrophoresis in 1·6% agarose gels using TBE (65 mM Tris–HCl, 27 mM boric acid, 1 mM EDTA, pH 9; Bio-Rad, Richmond, CA, USA) as the buffer and GeneRuler 100 bp SM 0321 (Fermentas, Hanover, MD, USA) as a size marker, and then stained with ethidium bromide and detected upon ultraviolet transillumination.

Second-round amplicons produced using each primer set Hmi-B1 or Hmu-B1 were purified over mini-columns (Ultrafree-DA, Millipore, Bedford, MA, USA) and subjected to automated sequencing (version 2; Applied Biosystems in a ABI-PRISM 377). Sequences were determined in both orientations (using the same primers) and electro-pherograms verified by eye. The sequences were aligned using the ClustalX program (see Thompson et al. 1997), and compared with one another and with those of H. microstoma (Accession number: AY251023) and H. muscae (Accession number: AY251024) determined previously (Traversa et al. 2004) and available in the GenBank™ database. The smallest amount of H. microstoma or H. muscae DNA yielding detectable amplicons by the semi-nested PCR was estimated to be 0·02 fg by multiple serial titrations into faecal samples collected from helminth-free foals.

RESULTS AND DISCUSSION

In a first step, the PCR assay described recently (Traversa et al. 2004) was assessed for its capacity to amplify H. microstoma or H. muscae DNA from all of the 30 faecal samples from horses known to be infected with Habronema (see Table 1), but it did not achieve effective amplification from them. To overcome this problem, the assay was modified to become a two-step, semi-nested PCR. Amplification was first conducted for 30 cycles using the primer set D-B1. Then, 4 μl of a 1/40 dilution of each D-B1 amplicon (determined to be optimal) were transferred to a fresh tube containing the same PCR reaction buffer (46 μl) with either primer set Hmi-B1 or Hmu-B1, and amplified for 35 cycles. The rationale was that the primer set D-B1 in the first round of PCR would amplify ITS-2 from both species of Habronema (undetectable on agarose gels) but not from DNA derived from host or other metazoan parasites (cf. Traversa et al. 2004). Upon amplification in the second round of PCR, the primer sets Hmi-B1 and Hmu-B1 would specifically amplify products of 196 bp and 395 bp from samples containing H. microstoma and H. muscae DNA, respectively, but not from host or other parasites (cf. Traversa et al. 2004). This two-step PCR was validated, and its specificity and sensitivity established using the panel of 46 test samples. Its specificity was also assessed using a panel of 48 control DNA samples from horse and from parasites, including T. axei and D. megastoma, G. intestinalis, F. hepatica, A. perfoliata, A. magna, P. mammillana, P. equorum, O. equi and a range of members of the Strongylinae and Cyathostominae.

In the first round of PCR, no amplicons were produced for any of the 46 test samples. In the second round, 18 amplicons of ~200 bp and 4 amplicons of ~400 bp were produced from samples from horses infected with either H. microstoma (n=19) or H. muscae (n=4). Amplicons of ~200 bp and ~400 bp were both amplified from samples from horses (n=7) infected with both H. microstoma and H. muscae (see Table 1; Fig. 1). Using the two-step PCR, no amplicons were produced for any of the 14 samples from horses infected with other parasites, including T. axei, bots, cestodes, strongylids and O. equi (Table 1). There was no evidence of PCR inhibition in any of these 14 samples, or in sample no. 1 which did not yield amplicons after 2 rounds of PCR but which was from a horse known to be infected with H. microstoma (Table 1). No amplicons were produced from any of the control DNA samples from horse or from non-spirurid parasites. While no amplicon was produced from any of the DNA samples (n=3) representing the spirurid D. megastoma using the primer set Hmu-B1, a product of ~160 bp was amplified from all of them employing primer set Hmi-B1; this amplicon was shown by sequencing to represent partial ITS-2 of D. megastoma and was thus specific (results not shown).

Fig. 1. Example of an agarose gel showing PCR products amplified from horse faecal samples by the two-step, semi-nested PCR. Lane M: size marker; Lanes 1–6: samples 2, 10, 12, 19, 28 and 29 (Hmi-B1); Lanes 7–9: samples 3, 9, 18 (Hmu-B1); Lane 10: Hmu-B1 positive control (Habronema muscae DNA); Lanes 11 and 12: samples 34 and 40; Lane 13: negative control (no-DNA sample); Lane 14: Hmi-B1 positive control (Habronema microstoma DNA). Bands of <100 bp (lanes 6, 7, 9 and 11–14) represent primer dimers.

All ~200 bp and ~400 bp amplicons (n=36) produced in the two-step PCR using the primer sets Hmi-B1 and Hmu-B1 were sequenced and demonstrated to represent appropriate regions of ITS-2 of H. microstoma and H. muscae, respectively. For each species, the sequences obtained (available from the corresponding author) were ~99% (mean value) similar to their respective sequence in the GenBank™ (see Traversa et al. 2004). Sequence variation among all Hmi-B1 (n=25) and all Hmu-B1 (n=11) amplicons was ~0·4% and ~2% (mean percentages), respectively. This magnitude of variability was consistent with that recorded in a previous study and relates mainly to microsatellite variation (cf. Traversa et al. 2004). While not the subject of the present study, such variation may be useful for investigating, using a mutation scanning approach, the genetic make-up of each species of Habronema from different geographical regions (see Gasser et al. 2001).

The two-step PCR assay assessed herein achieved a diagnostic sensitivity of 96·7% and a specificity of 100% using the present test sample set. These values are similar to those (~95% sensitivity and 100% specificity) achieved recently by Verweij et al. (2001) using a similar PCR specific for the detection of DNA from Oesophagostomum bifurcum or Necator americanus in the faecal samples from humans shown to be infected by coproculture (which has a substantially higher diagnostic sensitivity for these strongylid nematodes than it does for gastric spirurids of equids). These findings demonstrate that the present PCR is a useful tool for the diagnosis of gastric habronemosis in horses. As there is but one (since unconfirmed) report of a single specimen of D. megastoma from the large intestine (rather than the stomach) of a donkey in Italy (Ricci & Sabatini, 1992), the present study focused on H. microstoma and H. muscae in this country. Nonetheless, this molecular tool could be readily applied in geographical regions where D. megastoma also occurs in equids, because the three species of spirurid could be readily differentiated based on the sizes (i.e. ~200 bp, ~400 bp and ~160 bp) of the specific amplicons produced. Clearly, this method overcomes the majority of limitations of traditional diagnostic techniques, and thus provides a powerful approach for epidemiological, drug efficacy and ecological studies.

The PCR tool established can now be used to conduct ante-mortem surveys of the prevalence of H. microstoma and H. muscae infections in equids, which is likely to have been underestimated to date due to the inherent limitations of previous copro-diagnostic approaches. Given the controversy surrounding gastric habronemosis as a clinical entity (cf. Euzéby, 1961; Soulsby, 1982), it may also provide a means of assessing whether species of Habronema represent a risk factor in equine colic. The PCR method may also be of use to evaluate (indirectly) the efficacy of compounds against species of Habronema via the monitoring of the decline or absence of parasite-specific DNA in faeces after treatment in live horses with gastric habronemosis, thus circumventing the need to sacrifice them. This latter statement is supported by the fact that DNA was amplified from the faeces of horses with a very low number of Habronema specimens (e.g., 2 females and 1 male; sample no. 2; Table 1) in the stomach and by the minimum amount (~0·02 fg) of Habronema DNA required for effective amplification. This approach should be particularly applicable to H. microstoma, for which no detailed drug efficacy data are yet available. The PCR assay could also assist in the diagnosis of cutaneous habronemosis, which is sometimes difficult to differentiate from other dermopathies of equids (such as botryomycosis, pythiosis, onchocercosis, equine sarcoid and squamous cell carcinoma) with similar clinical signs (see Inzana & Carter, 1990; Pascoe, 1990; Chaffin et al. 1995), via the specific amplification of Habronema DNA from skin biopsy samples. Moreover, the present assay could also be employed for the (indirect) identification of the key arthropod intermediate hosts for species of Habronema via the specific detection of their DNA in muscid flies, because these have not yet been identified for a range of countries (due to the major constraints in specifically identifying larvae in them).

This project was supported financially through the Italian Ministry of the Scientific and Technological Research (COFIN 40% grant). Other support from the Australian Academy of Science and the Australian Research Council (to R.B.G.) is acknowledged.

References

REFERENCES

ANDERSON, R. C. ( 2000). The superfamily Habronematoidea. In Nematode Parasites of Vertebrates. Their Development and Transmission, 2nd Edn, pp. 428–430. CABI Publishing, Guilford, UK.
BORGSTEEDE, F. H. & VAN BEEK, G. ( 1998). Parasites of stomach and small intestine of 70 horses slaughtered in The Netherlands. Veterinary Quarterly 20, 31–34.CrossRefGoogle Scholar
BUCKNELL, D. G., GASSER, R. B. & BEVERIDGE, I. ( 1995). The prevalence and epidemiology of gastrointestinal parasites of horses in Victoria, Australia. International Journal for Parasitology 25, 711–724.CrossRefGoogle Scholar
CHAFFIN, M. K., SCHUMACHER, J. & McMULLAN, W. C. ( 1995). Cutaneous pythiosis in the horse. Veterinary Clinics of North America: Equine Practice 11, 91–103.CrossRefGoogle Scholar
DE JESUS, Z. ( 1963). Observations of habronemiasis in horses. Philippines Veterinary Medicine 2, 133–152.Google Scholar
DUNN, A. M. ( 1969). Habronemiasis and Drascheiasis (sic. Draschiasis). In Veterinary Helminthology, pp. 199–200. W. Heinemann Medical Books Ltd, London, UK.
EUZEBY, J. ( 1961). Les Maladies Vermineuses des Animaux Domestiques et leurs Incidences sur la Pathologie Humaine. Maladies dues aux Nemathelminthes. Fascicule Premier. Vigot Frères, Paris, France.
EUZEBY, J. ( 1981). Diagnostic expérimental des helminthoses animales. Travaux pratiques d'helminthologie vétérinaire. Livre 1. Edition “Informations Techniques des Services Vétérinaires”, Paris, France.
GASSER, R. B. ( 1999). PCR-based technology in veterinary parasitology. Veterinary Parasitology 84, 229–258.CrossRefGoogle Scholar
GASSER, R. B., ZHU, X. Q., BEVERIDGE, I. & CHILTON, N. B. ( 2001). Mutation scanning analysis of sequence heterogeneity in the second internal transcribed spacer (rDNA) within some members of the Hypodontus macropi (Nematoda: Strongyloidea) complex. Electrophoresis 22, 1076–1085.3.0.CO;2-8>CrossRefGoogle Scholar
HASS, K. ( 1979). Equine parasitism. Veterinary Medicine and Small Animal Clinician 74, 980–988.Google Scholar
HUNG, G.-C., GASSER, R. B., BEVERIDGE, I. & CHILTON, N. B. ( 1999). Species-specific amplification by PCR of ribosomal DNA from some equine strongyles. Parasitology 119, 69–80.CrossRefGoogle Scholar
INZANA, T. J. & CARTER, G. R. ( 1990). Fungi causing systemic or deep infections. In Diagnostic Procedures in Veterinary Bacteriology and Mycology, 5th Edn (ed. Carter, G. R. & Cole, J. R.), pp. 433–455. Academic Press Inc., San Diego, CA, USA.CrossRef
LICHTENFELS, J. R. ( 1975). Helminths of domestic equids. Proceedings of the Helminthological Society of Washington 42, 1–92.Google Scholar
LYONS, E. T., TOLLIVER, S. C., DRUDGE, J. H., SWERCZEK, T. W. & CROWE, M. W. ( 1983). Parasites in Kentucky thoroughbreds at necropsy: emphasis on stomach worms and tapeworms. American Journal of Veterinary Research 44, 839–844.Google Scholar
LYONS, E. T., TOLLIVER, S. C., DRUDGE, J. H., SWERCZEK, T. W. & CROWE, M. W. ( 1987). Common internal parasites found in the stomach, large intestine and cranial mesenteric artery of thoroughbreds in Kentucky at necropsy (1985 to 1986). American Journal of Veterinary Research 48, 268–273.Google Scholar
LYONS, E. T., TOLLIVER, S. C., STAMPER, S., DRUDGE, J. H., GRANSTROM, D. E. & COLLINS, S. S. ( 1994). Transmission of some species of internal parasites in horses born in 1990, 1991 and 1992 in the same pasture on a farm in central Kentucky. Veterinary Parasitology 52, 257–269.CrossRefGoogle Scholar
OWEN, J. & SLOCOMBE, D. ( 1985). Pathogenesis of helminths in equines. Veterinary Parasitology 18, 139–153.CrossRefGoogle Scholar
PANDEY, V. S., OUHELLI, H. & VERHULST, A. ( 1992). Epidemiological observations on stomach worms of donkeys in Morocco. Veterinary Research Communications 16, 273–279.CrossRefGoogle Scholar
PASCOE, R. R. ( 1990). A Colour Atlas of Equine Dermatology. Wolfe Publishing Ltd, Ipswich, UK.
RICCI, M. & SABATINI, A. ( 1992). Notizie sugli elminti parassiti del cieco e del colon degli equidi in Italia. Parassitologia 34, 53–60.Google Scholar
ROUBAUD, E. & DESCAUZEAUX, J. ( 1922). Evolution de l'Habronema muscae Carter chez la mouche domestique et de l'H. microstomum Schneider chez le stomoxe (Note préliminaire) comme Agent Vecteur des Habronèmosis d'Équides. Bulletin de la Société de Pathologie Exotique 15, 572–574.Google Scholar
SCHMIDT, G. D. ( 1986). Handbook of Tapeworm Identification. CRC Press, Boca Raton, Florida, USA.
SOULSBY, E. J. L. ( 1982). Helminths, Arthropods and Protozoa of Domesticated Animals, 7th Edn. Baillière, Tindall and Cassell, Eastbourne, UK.
THOMPSON, J. D., GIBSON, T. J., PLEWNIAK, F., JEANMOUGIN, F. & HIGGINS, D. G. ( 1997). The ClustalX windows interface: flexible strategies for multiple sequence alignment aided by quality analysis tools. Nucleic Acids Research 24, 4876–4882.CrossRefGoogle Scholar
TOLLIVER, S. G. ( 2000). A Practical Method of Identification of the North American Cyathostomes (Small Strongyles) in Equids in Kentucky. Published by the Kentucky Agricultural Experiment Station-College of Agriculture, University of Kentucky, Lexington, Kentucky, USA.
TRAVERSA, D., GIANGASPERO, A., GALLI, P., PAOLETTI, B., OTRANTO, D. & GASSER, R. B. ( 2004). Specific identification of Habronema microstoma and Habronema muscae (Spirurida, Habronematidae) by PCR using markers in ribosomal DNA. Molecular and Cellular Probes 18, 215–221.CrossRefGoogle Scholar
VERWEIJ, J. J., PIT, D. S., VAN LIESHOUT, L., BAETA, S. M., DERY, G. D., GASSER, R. B. & POLDERMAN, A. M. ( 2001). Determining the prevalence of Oesophagostomum bifurcum and Necator americanus infections using specific PCR amplification of DNA from faecal samples. Tropical Medicine and International Health 6, 726–731.CrossRefGoogle Scholar
VERWEIJ, J. J., POLDERMAN, A. M., WIMMENHOVE, M. C. & GASSER, R. B. ( 2000). PCR assay for the specific amplification of Oesophagostomum bifurcum DNA from human faeces. International Journal for Parasitology 30, 137–142.CrossRefGoogle Scholar
ZUMPT, F. ( 1965). Myiasis in Man and Animals in the Old World. Butterworths, London, UK.
Figure 0

Table 1. Horses determined to be infected (+) or not infected (−) with Habronema microstoma (Hmi), Habronema muscae (Hmu), Trichostrongylus axei (Ta), Gasterophilus intestinalis (Gi), cestodes (Ce), Oxyuris equi (Oe), Cyathostominae (Cy) and Strongylinae (St) at autopsy, and comparison with faecal examination results (for larvated spirurid eggs) obtained using different flotation solutions and those achieved employing the two-step, semi-nested PCR

Figure 1

Fig. 1. Example of an agarose gel showing PCR products amplified from horse faecal samples by the two-step, semi-nested PCR. Lane M: size marker; Lanes 1–6: samples 2, 10, 12, 19, 28 and 29 (Hmi-B1); Lanes 7–9: samples 3, 9, 18 (Hmu-B1); Lane 10: Hmu-B1 positive control (Habronema muscae DNA); Lanes 11 and 12: samples 34 and 40; Lane 13: negative control (no-DNA sample); Lane 14: Hmi-B1 positive control (Habronema microstoma DNA). Bands of <100 bp (lanes 6, 7, 9 and 11–14) represent primer dimers.