INTRODUCTION
⌅Parasitic gastrointestinal nematode infections remain a leading factor militating against the productivity and profitability of livestock business, especially the small ruminant enterprise, all over the world. In the tropics, these infections are mainly caused by the trichostrongylid nematodes, particularly, Haemonchus contortus, a blood-sucking nematode (Saddiqi et al., 2011). Infections are often associated with significant economic losses ranging from insidious loss of body conditions to outright mortality. The infection by this parasite is often widespread and occurs all year round in the tropics (Chiejina, 1986; Sowemimo et al., 2012). High H. contortus burden may lead to death especially in young animals. Under field conditions, co-infection with other species of the nematode sometimes occurs (Squire et al., 2019).
The control methods available to farmers in the tropics involve reducing worm burden in the animal through anthelmintic drug treatment in combination with controlled grazing, which consequently reduces the contamination of pastures (Barger, 1999). However, communal ownership of farmlands in most communities in the tropics limits their use for controlled grazing (Githiori et al., 2003). Consequently, anthelmintic intervention remains the most effective means of controlling the disease, but their continuous use and efficacy are limited by the unaffordability of the drugs, their uncertain availability, and the emergence of worm strains that are resistant to the available drugs (Epe & Kaminsky, 2013). In addition, there has been an increasing concern over chemical residues in edible animal products associated with the use of anthelmintic drugs in livestock (Waller, 1997).
Anthelmintic control of helminths involves routine treatment with synthetic drugs belonging to different anthelmintic families, namely, benzimidazoles, imidazothiazoles, and macrocyclic lactones. This has inevitably led to the selection of resistant strains of gastrointestinal nematode parasites, and particularly H. contortus, which resulted in partial or total inefficacy of most anthelmintic classes (Roeber et al., 2013). Given the lack of prospect of developing a vaccine against these parasites and the economic impacts caused by the increasing resistance to anthelmintics, it is important to either discover novel molecules or compounds able to control multi-resistant nematodes or seek viable alternatives that are effective, affordable, safe, and less selective for resistant worms.
Medicinal plants have served as a constant source of remedies for a variety of diseases over centuries. Plants have been a rich source of antimicrobial and anthelmintic agents and their products are used medicinally in different parts of the world as sources of many potent and safe drugs, including anthelmintics (Lai et al., 2005; Abdul-Ghani et al., 2011; Harvey et al., 2015; Irum et al., 2015). This study was therefore designed to investigate the anthelmintic efficacies of three medicinal plants, namely, Annona senegalensis, Cochlospermum planchonii, and Sarcocephalus latifolius against H. contortus.
MATERIAL AND METHODS
⌅Plant collection
⌅The plants used in this study were selected through a structured questionnaire administered to livestock farmers in Northern Nigeria. The questionnaire elicited information on herbs used in the treatment of animal diseases, of which the three plants were among those used by farmers to treat gastroenteritis, including those due to nematode parasitism in ruminants. The plants were collected within the vicinities of the University of Agriculture, Makurdi, Benue State. They were identified by a plant taxonomist in the Department of Botany, University of Agriculture, Makurdi, where voucher specimens were deposited in an herbarium. The plant materials, which include the root of CP and leaves of AS and SL were air-dried at room temperature (22-370C) and relative humidity of 39-45.6% for 6-8 days. Thereafter, the dried samples were ground to a fine powder with a hammer mill and then stored in an air-tight container at room temperature until needed.
Plant extraction
⌅Acetone and aqueous (water) extraction were performed on each of the three plant materials. Acetone was used because of its ability to extract compounds with a wide range of polarities based on its superiority as an extractant, and on several parameters, as described in several studies (Eloff, 1998; Kotze & Eloff, 2002; Eloff et al., 2005). One gram of each of the plant materials was separately extracted with 10 mL of acetone (>99% technical grade, Merck) in polyester centrifuge tubes. Aqueous extraction of the plants was chosen, because water is the solvent used by the natives. The tubes were vigorously shaken on an orbital shaker for 30 minutes, and then centrifuged at 4000 × g for 10 minutes. Thereafter, the supernatant of each extraction was filtered using Whatman No.1 filter paper into pre-weighed glass containers. The solvents were allowed to evaporate under a stream of air in a fume hood at room temperature to obtain the dried extract. The extracts were stored at 4°C until required and reconstituted in 5% dimethyl sulfoxide (DMSO) when needed for the assay.
Recovery and preparation of H. contortus eggs for egg hatch assay
⌅H. contortus eggs used in the assay were obtained from faeces collected per rectum from goats carrying mono-specific infections of H. contortus. Approximately 3 g of the faecal sample were crushed and made relatively liquid (slurry) by adding 42 mL of saturated sodium chloride solution, and the suspension passed through a tea strainer. The filtrate was placed in 15 mL test tubes on the bench undisturbed for 15 minutes to allow the worm eggs to float to the top of the tube. Thereafter, the top portions of the fluid containing the eggs were poured into another tube which was then resuspended with water to dilute and wash out the salt solution. The tubes were centrifuged at ×1000 g for 5 minutes after which the supernatant was decanted and the sediment resuspended with water. This was repeated three times after which the ‘cleaned’ eggs were resuspended with deionized water and placed in a universal bottle. For use in the assay, 1 mL of the egg suspension was further diluted to contain approximately 100 eggs in 200 µL. Egg count was carried out using the modified McMaster egg counting technique as described in Fakae et al. (1999).
Egg Hatch Assay (EHA)
⌅Egg hatch assay (EHA) was conducted following the guidelines of the World Association for the Advancement of Veterinary Parasitology (WAAVP) (Coles et al., 1992). Approximately 100 H. contortus egg suspension in 200 µL of deionized water was incubated with different concentrations (0.625, 1.25, 2.5, 5.0, 10.0 and 20 mg/mL) of each plant extract in 5% DMSO in a 48 –flat-bottomed microtitre plate to obtain a final tested concentration of 0.3125 to 10 mg/mL in 2.5% DMSO. Albendazole served as a positive control and was dissolved in 5% DMSO in de-ionized water to obtain different concentrations (0.01 to 25 µg/mL), while 5% DMSO served as the negative control. The setup was incubated in triplicate for each extract at 270C for 48 hours. At the end of 48 hours, a drop of Lugol’s iodine solution was added to each well and the number of larvae vs unhatched eggs (including larvated ones) was counted.
Thereafter, Probit analysis (Finney, 1971) was conducted to determine the lethal concentration (LC50) of the extracts and albendazole. The percentage inhibition of egg hatching was calculated using the formula by Cala et al. (2012):
where E = % inhibition of egg hatching, and L1 = Number of larvae in a particular well. All experiments were undertaken in triplicate on three separate occasions.