Trichostrongylus axei: Abomasal Hairworm of Cattle and Sheep (Gastric Parasitism)

Taxonomic Classification and Morphology

Trichostrongylus axei (Cobbold, 1879) is a slender, hair-like nematode belonging to the family Trichostrongylidae, order Strongylida. It is a common abomasal parasite of cattle, sheep, and goats, and occasionally infects horses, pigs, and deer [1, 2]. The adult worms are small, measuring 3-5 mm in length (males) and 4-6 mm (females), with a filariform appearance that gives rise to the common name "hairworm" [3]. Male worms possess a well-developed copulatory bursa with two large lateral lobes and a small dorsal lobe, supported by characteristic ray patterns used for species identification [4]. The spicules are short, stout, and curved, often with a distinctive barb at the tip [5].

The cuticle bears fine longitudinal ridges (synlophe) that vary among trichostrongylid species but are less pronounced than in Ostertagia or Haemonchus [6]. Eggs of T. axei are thin-shelled, morulated, and measure approximately 75-95 µm by 35-46 µm, indistinguishable from those of other trichostrongylids by standard flotation [7]. Larvae must be cultured to the third stage (L3) for species-level identification using morphological keys [8, 9].

Host Range and Geographic Distribution

Trichostrongylus axei has a broad host range, primarily infecting ruminants. In cattle, it is frequently found co-infecting the abomasum along with Ostertagia ostertagi and Haemonchus placei [10, 11]. In sheep, it commonly coexists with Teladorsagia circumcincta and Trichostrongylus colubriformis [12, 13]. The parasite has a worldwide distribution, with prevalence reported in temperate and subtropical regions. In Europe, surveys have recorded T. axei in grazing dairy cattle in Germany and Austria [2], in dairy heifers in England and Wales [14], and in Austrian beef cattle [15]. In North America, it is prevalent in cattle from the United States and Canada [16, 17]. In Japan, it is part of the abomasal nematode complex in dairy cattle in Hokkaido [18]. In tropical regions such as The Gambia, T. axei has been identified in N'Dama cattle, often in a state of hypobiosis during dry seasons [19].

Life Cycle and Epidemiology

The life cycle of T. axei is direct, similar to other trichostrongylids. Adult worms reside in the abomasum, attached to the mucosal surface or lying free in the mucus [3]. Eggs pass out in feces, and under favorable field conditions (15-30°C, adequate moisture), they hatch to release first-stage larvae (L1) that feed on bacteria and develop through two molts to become infective third-stage larvae (L3) in 7-14 days [1, 20]. The L3 migrate onto herbage and are ingested by grazing animals. The exsheathment of L3 occurs in the rumen or abomasum, triggered by carbon dioxide and pH changes [1, 21]. After exsheathment, the larvae penetrate the abomasal mucosa and undergo two more molts to reach the adult stage. The prepatent period is approximately 18-21 days in cattle and sheep [3, 17].

Environmental factors profoundly influence transmission. Cool, moist conditions favor survival of L3 on pasture, while hot, dry conditions reduce larval longevity [2, 22]. In temperate regions, transmission peaks in spring and autumn. Hypobiosis (arrested larval development) is an important epidemiological feature observed in T. axei, though it is less pronounced than in O. ostertagi [11, 19]. In the Gambia, hypobiotic larvae have been recovered during the dry season, enabling the parasite to persist through unfavorable periods [19]. In northern Japan, seasonal fluctuations in worm burdens have been documented, with peaks in late autumn [18]. Co-infection with O. ostertagi can modulate the hypobiotic response; experimental studies have shown that concurrent infection with both species increases the proportion of inhibited O. ostertagi larvae, likely due to host immune interactions [11].

The following Mermaid diagram summarizes the life cycle of Trichostrongylus axei in cattle and sheep:

graph TD
 A[Adult worms in abomasum] --> B[Eggs in feces]
 B --> C[L1 in dung pat]
 C --> D[L2 in dung pat]
 D --> E[L3 infective on herbage]
 E --> F[Ingested by host]
 F --> G[Exsheathment in rumen/abomasum]
 G --> H[L4 penetrate mucosa]
 H --> I[L5 develop in lumen]
 I --> A
 H --> J[Hypobiosis arrested L4]
 J --> I

Pathogenesis and Clinical Signs

Trichostrongylus axei is a blood-feeding nematode, though its pathogenic impact is generally less severe than that of Haemonchus contortus or Ostertagia ostertagi [3]. The adult worms irritate the abomasal mucosa, causing a catarrhal inflammation with increased mucus production [11, 23]. Histologically, there is hyperplasia of gastric glands, infiltration of eosinophils and lymphocytes, and occasional sloughing of surface epithelium [11, 24]. The damage leads to a protein-losing gastroenteropathy, reduced digestive efficiency, and impaired growth.

In cattle, high burdens (e.g., >50,000 worms) can cause diarrhea, weight loss, submandibular edema ("bottle jaw"), and reduced milk yield in dairy cows [2, 25]. In young calves, experimental infections have shown reduced appetite and growth rates [17]. In sheep, the clinical syndrome is similar to that caused by Teladorsagia circumcincta, with diarrhea, poor fleece quality, and loss of condition [12]. The parasite can contribute to the "parasitic gastroenteritis" complex, especially when co-occurring with other species.

In contrast to Ostertagia ostertagi, T. axei does not typically cause the raised "Morocco leather" lesions characteristic of ostertagiosis type I, but rather diffuse inflammatory changes [11, 24]. Experimental single-species infections in calves have demonstrated that T. axei induces an eosinophilic infiltration and mucosal thickening, but without the severe nodular hyperplasia seen with O. ostertagi [11].

Diagnosis

Diagnosis of T. axei infection relies on detection of eggs in feces using standard flotation techniques (e.g., saturated sodium chloride, specific gravity 1.20) [26]. Because eggs are morphologically indistinguishable from those of other trichostrongylids, identification to species requires larval culture and differentiation of third-stage larvae (L3) using morphological keys [8, 27]. The Baermann technique is not required for abomasal nematodes, as larvae are not pulmonary stages. However, for differential diagnosis, fecal egg count reduction tests (FECRT) can be used to monitor anthelmintic efficacy, and identification of cultured larvae to species can improve interpretation of resistance patterns [27, 28].

Necropsy and abomasal worm counts remain the gold standard for quantifying burdens [3, 5]. The abomasum is opened, washed, and the contents and digest of the mucosa are examined for adult worms. Speciation of adult males relies on spicule morphology and bursal ray patterns [5]. For field surveys, composite sedimentation techniques are used [14, 15].

Molecular diagnostics are increasingly employed for species-specific detection and quantification. PCR-based nemabiome sequencing targeting the ITS-2 rDNA region can identify T. axei from fecal samples with high specificity [29, 30]. This nemabiome approach allows simultaneous profiling of the entire gastrointestinal nematode community and is particularly useful for mixed infections.

Treatment and Control

Anthelmintic treatment for T. axei follows the same guidelines as for other abomasal nematodes. Macrocyclic lactones (e.g., ivermectin, eprinomectin), benzimidazoles (e.g., fenbendazole, albendazole), and imidazothiazoles (e.g., levamisole) are effective [5, 7, 31, 32, 33]. Ivermectin and eprinomectin have demonstrated high efficacy against adult and larval stages [5, 31]. A sustained-release bolus of ivermectin has shown efficacy against T. axei in cattle [33]. However, resistance to benzimidazoles and macrocyclic lactones is emerging in trichostrongylid populations globally, and FECRT should be performed to confirm product efficacy in a herd or flock [27, 34].

Control strategies integrate pasture management, stocking density reduction, and strategic anthelmintic treatments to minimize pasture contamination. Rotational grazing can reduce exposure to infective larvae, though the effect may be less pronounced for T. axei than for other species due to its shorter survival on pasture [35]. In sheep, combining grazing with cattle or horses can take advantage of host specificity; T. axei is a generalist, but cross-grazing with resistant species may still reduce overall larval burden [36]. Biological control using nematophagous fungi (e.g., Duddingtonia flagrans) has been evaluated for T. axei and other trichostrongylids, with some success in reducing larval numbers on pasture [37, 38]. Immunological control is not yet available, but vaccine development against Trichostrongylus colubriformis has shown some cross-protection in experimental models.

The following table summarizes recommended diagnostic and control measures:

Drug Resistance

Anthelmintic resistance in T. axei has been reported, although less extensively than in Haemonchus contortus or Teladorsagia circumcincta. Resistance to benzimidazoles has been documented in sheep flocks in several countries, and macrocyclic lactone resistance is suspected but not as well characterized [27, 34]. The FECRT, with a reduction threshold of <95% (mean) and lower 95% confidence interval <90%, is the field standard for detecting resistance [27]. When resistance is suspected, larval culture and differentiation are essential to attribute resistance to a particular species, as mixed infections are common [27].

Public Health Considerations

Trichostrongylus axei is primarily a veterinary parasite and rarely infects humans. Accidental human infections have been reported following ingestion of contaminated raw vegetables or water, but such cases are sporadic and of minor clinical significance. No specific public health control measures are required beyond standard food hygiene. This article focuses strictly on veterinary aspects.

Conclusion

Trichostrongylus axei is a widely distributed abomasal nematode of cattle and sheep that contributes to production losses through impaired growth, reduced milk yield, and predisposition to parasitic gastroenteritis. Its small size and morphological similarity to other trichostrongylids necessitate careful larval differentiation and, increasingly, molecular nemabiome approaches for species-level diagnosis. Anthelmintic resistance is emerging, making integrated control strategies and regular efficacy testing imperative.

References

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