Prevalence of multiple resistant Haemonchus and Ostertagia species in goats and cattle in Machakos, Eastern Kenya

E O Mungube, L W Wamae¹, G A Omondi and G Mwangi¹

Kenya Agricultural and Livestock Research Organization (KALRO), Veterinary Research Centre, Muguga, P.O. Box 32-00902 Kikuyu, Kenya
wamateka@hotmail.com
¹ KALRO Headquarters Nairobi, P.O. Box 57811-02100 Nairobi; Kenya

Abstract

Anthelmintic resistance has seriously undermined helminth control in ruminants. Faecal egg count reduction tests (FECRT) conducted between 5^th and 27^th January 2015 established prevalence of anthelmintic resistant nematodes in a goat and a cattle herd. Study animals were identified through a pre-treatment faecal egg counts (FECs) where 116 goats and 60 cattle with FECs ≥150 were selected. The animals were allocated into four treatment groups namely the control (untreated), valbazen, Nilzan plus and ivermectin using a complete randomised design. Valbazen and Nilzan plus were administered per os at 10 mg/kg and 7.5mg/kg, respectively to the experimental goats and cattle. Ivermectin was injected subcutaneously around the neck at 200mcg/kg.

Fourteen days post-treatment, experimental animals in each of the 4 groups were faecal sampled for FECs and persisting nematode species. Goats treated with valbazen, Nilzan plus and ivermectin had FEC reduction of 16%, 90% and 98%. This showed that Valbazen and Nilzan plus were ineffective against nematodes of goats. Haemonchus species persisted amongst goats treated with the 3 drugs. Valbazen, Nilzan plus and Noromectin treatment of cattle resulted in 98%, 95% and 99% reduction in FECs. Although all the 3 drugs were effective against nematodes in cattle, Ostergia species persisted amongst cattle treated with the 3 drugs. Molecular tools are needed to confirm prevalence of multiple resistant Haemonchus and Ostertagia species in the semi-arid region of Kenya.

Keywords: anthelmintic resistance, Haemonchus, cattle, goats, Ostertagia, refugia

Introduction

Helminthosis is a major cause of mortality and sub-optimal productivity in pastoral farming systems across sub-Sahara Africa (SSA). Compared to the temperate climatic zones, the impact of gastro-intestinal helminths in the SSA could be higher since the tropical climate is associated with a wider range of agro-ecological factors suitable for the survival of diversified hosts and helminth species (Sissay et al 2006). Heavy helminth infections if left untreated could be fatal to calves and small ruminants in addition to causing sub-clinical disease in adult cattle associated with lowered productivity, premature culling and making animals unsuitable as replacement breeding stock (Murphy et al 2006).

Although grazing management and treatment of animals using anthelmintic drugs (ADs) are the common strategies used in the control of helminths in livestock, ADs are by far the most favoured method by livestock keepers (Coles et al 2006). Despite the continued use of ADs, production losses still remain high. This could be due to the poor quality of ADs (Monteiro et al 1998) and the increasing threat of anthelmintic resistance (AR) that has been reported in many parts of the world (Kaplan 2004).

In most small ruminant production systems, anthelmintic resistant nematodes are a problem of major economic concern to farmers (Kaplan 2004). Several reports on AR especially those associated with benzimidazoles (BZ) in the small ruminant production systems have been reported. In Kenya notable BZ resistant nematodes in the sheep and goat production systems reports have been documented (Wanyangu et al 1996; Maingi et al 1998; Waruiru et al 1998; Gatongi et al 2003). The same has been reported in Tanzania (Bjorn et al 1990; Ngomuo et al 1990; Keyyu et al 2002), South Africa (Van Wyk et al 1999; Vatta et al 2001; Bakunzi et al 2003), Zambia (Gabrie et al 2001), Nigeria (Mbah et al 1992), Cameroon (Ndamukong and Sewell 1992), Zimbabwe (Mukaratiwa et al 1997), Ethiopia (Sissay et al 2006). On the other hand, AR is scantly reported in cattle possibly because of the infrequent use of anthelmintics on this class of animals (Kaplan 2004). The few reports on AR in bovine mostly originate from regions where cattle are kept under grazing management systems like in New Zealand (Coles 2002), South America (Soutello et al 2007; Suarez and Cristel 2007) and Europe (Coles 2002; 2004; Demeler et al 2009).

It is commonly accepted that resistant genes exist as rare alleles in natural populations (Michel et al 1982; Prichard 1990; Jackson 1993) and that anthelmintic resistance develops when selection pressure is high. The two main factors which select for resistance and accelerate resistance development are treatment frequency and a high number of anthelmintic treatments with the same anthelmintic family for years (Sykes et al 1992). At the KALRO Katumani goat herd, albendazole has been used as a preferred anthelmintic uninterrupted for over 20 years raising suspicion for possible resistance by the common gastro-intestinal nematodes. It is reported that although sheep, cattle and goats may harbour similar nematode parasites, the BZ dose needed for treating goats should be a slightly higher (Chartier et al 1999; Hennessy and Alvineriem 2002; Sangster et al 1993) than that for sheep and cattle. In many farms, similar BZ doses are used for treating the animals hastening the risk of BZ resistant nematodes development in goats.

There exists empirical evidence to suggest that the risk for AR increases with under-dosing, lack of anthelmintic class rotation and high drench frequency (Domke et al 2011), justifying the need for regular evaluation of the efficacy of common anthelmintic drugs used in worm control regimes. This continuous analysis of the treatment response of the marketed anthelmintic drugs will ensure that farmers are adequately advised on the appropriate drugs to use as dewormers hence assuring them of value for their money. The aim of this study was to investigate the prevalence of AR for the common anthelmintics used for controlling worms in cattle and goat flocks and to also identify the resistant nematode genera in selected large-scale farms around Machakos, Eastern Kenya.

Methodology

The study was carried out in two large-scale farms: a goat herd at KALRO Katumani and a cattle herd at New Astra ranch. Both herds are located in the Machakos County, Kenya (Fig 1).

The two herds are located 20 km apart within agro-ecological zone lower midland 4 (LM 4) (Jaetzold and Schmidt 2006) where bimodal rainfall pattern is experienced. The long rains start in mid-March to early May while the short rains start in mid-October to mid-December (Rao et al 2011). The total rainfall for the two seasons is approximately 450 mm and is not only erratic but also poorly distributed in time and space. 

In this zone, agro-pastoralism is practiced and farmers commonly grow cowpeas, pigeon peas green grams, beans and to some limited extent cassava. They also rear cattle which include the boran, zebu and recently introductions of Friesian and zebu crosses. The small East African goats (SEAG) and Galla goats and Toggenburg and the Galla/SEAG crosses are reared. Although sheep is still a minority animal, the few who have sheep, keep the red Maasai sheep and dorper. The cattle and goats within LM 4 are extensively grazed on common pastures with minimal supplementation.

Experimental animals

A total of 116 experimental goats were purposively selected from a goat herd of 230 at KALRO Katumani on the basis of their faecal egg counts (FECs). These goats were dual purpose goats; a composite breed consisting of Galla and the SEAGs. The experimental goats were identified after screening all goats aged at least 3 months for their FECs with those whose FECs ≥150 qualifying as experimental animals. Before the final list of goats was compiled, those which for one reason or the other had received anthelmintic treatment in the last three months were dropped from the list of experimental animals.

At the New Astra cattle flock, 60 experimental cattle were also purposively selected from a population of 155 cattle aged between 3 months and 12 months from the general herd and ear-tagged. A FEC screening exercise was undertaken to identify those with FECs of ≥150 which were included as experimental cattle. Like for the experimental goats, any cattle which had been treated with anthelmintics in the last three months were also dropped from the list of experimental animals. The 60 eligible cattle were ear tagged and a list prepared for ease of follow-up.

Experimental design and treatments

The experiment was undertaken in three stages namely the pre-treatment FEC screening stage, treatment of selected animals and post-treatment FECs between 5^th January 2015 and 22^nd February 2015. A complete randomised design was used during this study where the experimental goats and cattle were randomly allocated into four groups namely; the control (untreated) group, valbazen® (albendazole 10%) treatment group, Nilzan plus® (Levamisole and oxyclozanide) treatment group and Noromectin® (ivermectin) treatment group (Table 1).

Both goats and cattle within the valbazen treatment group received 10 mg/kg of albendazole 10% (valbazen, Ultravetis EA Ltd) instead of the 7.5mg/kg per os to accommodate the higher albendazole dose for goats. The goats and cattle on Levamisole 1.5%, and Oxyclozanide 0.382% (Nilzan plus, Cooper Kenya Ltd) received 7.5mg/kg per os. The ivermectin treated goats and cattle received 200mcg/kg of ivermectin 1% (Noromectin, Norbrook Pharmaceuticals Ltd, Schering, UK) which was injected subcutaneously around the neck region.  The treatments were calculated on the basis of body weight estimated using a weighing band.

Faecal egg counts reduction test

Faecal egg counts from experimental animals were analysed for helminth eggs during the pre- and post-treatment phases. The pre-treatment faecal sampling and FECs were undertaken between 5^th and 7^th January 2015 while treatment of experimental animals was conducted on the 12^th January 2015. Post-treatment faecal sampling and FECs were conducted between 3^rd and 26^th and 27^th January 2015, 14 days after treatment. This was to allow for comparison in FECs for experimental animals for the four experimental groups before and after treatment to measure the effect of drugs on helminth egg shedding.

Faecal samples for each experimental goats and cattle belonging to each of the four treatment groups were collected per rectum and appropriately labelled with animal tag number, breed, sex, date and herd identity. The samples were packed in separate bags and zipped before they were delivered to the helminthology laboratory at the Veterinary Research Institute, KALRO Muguga in an ice-packed cool boxes.

The FECs were done using modified McMaster technique (Kaufmann 1996). Briefly, 4 g of faeces were weighed and put into a petri dish. Some saturated sodium chloride solution was added and thoroughly mixed and stirred using two wooden spatulas. The suspension was transferred into a measuring cylinder by straining through a sieve before more saturated salt solution was added to make it 60 ml. The suspension was then transferred into a sealable container and properly mixed by gentle shaking. Both chambers of the McMaster slide (Chalex, Wallowa, USA) were carefully loaded with the suspension to avoid transferring air bubbles in the counting chambers and eggs counted using a compound microscope at × 40 magnification. The eggs per gram faeces (EPG) or faecal egg counts (FECs) was calculated as the number of eggs counted multiplied by a correction factor of 50.

Copro-cultures

Identification of the prevailing GINs in the study area was done to show which species were sensitive and which ones were resistant to the three test anthelmintic drugs. Copro-cultures were done using the method described by Kaufmann (1996) between 5^th and 18^th January 2015 for the pre-treatment phase and 8^th and 22^nd February 2015 for the post-treatment. Briefly, about 50g of faecal samples from the experimental goats and cattle belonging to the albendazole treatment group, Nilzan plus treatment group, ivermectin treatment group and untreated control group were separately pooled by treatment group. The 8 (4 for goats and another 4 for cattle) pooled faecal samples were broken down into fine particles using a wooden spatula and mixed with vermiculite (Rajapack^®, Birkenfeld, Germany) and tap water, ensuring they remained moist and crumbly but not really wet. The mixture was then filled into 8 separate plastic culture dishes identified by treatment group and animal before partially covering them and incubating at 27ºC for 7 days. Larvae 3 were recovered using the Baermann technique and their identification done as described by Kaufmann 1996. Morphological features used in the identification were the shape of the anterior part and the number of gastro-intestinal cells of the 3^rd stage larvae.

Data analysis

The arithmetic mean, percentage reduction and 95% confidence interval of the FECs for the 4 treatment groups were calculated by the method described by Coles et al (1992). The percentage reduction is 100 (1 - x̄t/x̄c) where x̄ is the arithmetic mean of the FEC, t, treatment groups (valbazen®, Nilzan plus®, Noromectin®) at day 14 post-treatment and c is the control group FEC at 14 day. The 95% confidence interval was calculated by applying the formula 100[1 - x̄t/x̄c exp (- 2.048√Y2)] for the lower limit and 100[1 - x̄t/x̄c exp (+ 2.048√Y²)] for the upper limit. Y², the variance of reduction was calculated as:

 Y²=Variance of treatment group / (n, for treatment group* x̄²t) + Variance of control group/(n, for control group* x̄²c)

The interpretation of the FECR test results (Coles et al 1992) was as follows: resistance was declared where the percentage reduction in FECs was < 95% and where the lower 95% confidence interval was < 90%. Where only one of the above conditions was accomplished, emerging resistance was suspected.

Results

The mean pre-treatment FECs for the experimental goats were 1989 for the control group, 2261 for the valbazen treatment group, 1942 for the Nilzan plus treatment group and 2112 for the Noromectin treatment group (Table 2). A notable reduction in FECs between the pre-treatment and post-treatment phases of for the goats belonging to the control group was observed. This notwithstanding, the FECRT results displayed in Table 2 shows that valbazen treated goats only had 16% reduction indicative of presence of albendazole resistant GIN nematodes within the goat herd (Table 2). Nilzan plus (Levamisole 1.5%/Oxyclonide 0.382%) treatments in goats resulted in a 90% reduction in the FEC output indicating likely presence of levamisole resistant GINs in the goat herd. Noromectin (Ivermectin) treatment resulted in a higher reduction percentage of 98% meaning the drug was effective against majority GINs in the studied goat herd.

Of the 116 goats selected to participate on the study, 8 (1 each from the Valbazen^® and Nilzan plus^® could not be traced at the time of post-treatment faecal sampling. Some of the goats had died shortly after being selected.

Copro-culture of pooled faecal samples from goats during the pre-treatment phase established that a number of GINs were prevalent in the goat herd at Katumani. Haemonchus species was the most prevalent GIN species followed by Trichostrongylus, Oesophagostomum and Telardorsagia (Ostertagia) in that order (Figure 2).

However, copro-culture results for the day 14 post-treatment sampling had Cooperia species manifesting itself in addition to Haemonchus, Telardosargia (Ostertagia), Trichostrongylus and Oesophagostumum species (Table 3).

Haemonchus species was the most dominant GIN species which persisted in all the three anthelmintic treatment groups. It’s prevalence among albendazole treated goats was 31%, 28% among Nilzan plus treatment group and 100 % among the ivermectin treated goats (Table 3). On the contrary, 28% of Telardosargia (Ostertagia) and 56% Trichostrongylus species persisted among the valbazen and Nilzan plus treated goats. The prevalence of Cooperia species was 27% and 16%, respectively for the valbazen and Nilzan plus treated goats.

The mean pre-treatment FECs for the study cattle were 475 for the control group, 514 for the valbazen treatment group, 604 for the Nilzan plus treatment group and 443 for the ivermectin treatment group (Table 4). Like was the case for goats, there was a general reduction in FECs for the control group from a mean FEC of 475 to 342 comparing the pre-treatment and post-treatment phases. Valbazen treated cattle had reduction in FECs of 98% meaning the majority of GINs in the study cattle were still sensitive to valbazen (Table 4). Nilzan plus treatment resulted in a 95% reduction in the FEC output also a sign that GINs in the study cattle were still sensitive to levamisole. Ivermectin treatment resulted in the highest reduction percentage of 99% meaning the drug was effective against majority GINs in the study cattle.

Cultured pooled pre-treatment faecal samples collected from cattle resulted showed prevalence of Haemonchus species as the most common followed by Trichostrongylus, Oesophagostomum and Ostertagia (Figure 3).

The post-treatment faecal sample culturing indicated the prevalence of Haemonchus, Trichostrongylus, Ostertagia, Oesophagostomum and Cooperia species in the treated cattle (Table 5).

Multiple drug resistant Ostertagia species persisting in valbazen with a prevalence of 50%, 100% in Nilzan plus and Noromectin treatments, respectively (Table 5). The prevalence of Haemonchus species resistant to albendazole was 50%.

Discussion

The findings by this study on the three test drugs; Valbazen^® (albendazole 10%) Nilzan^® plus (levamisole 1.5% and oxyclozanide 3%) and Noromectin^® (ivermectin 1%) on their effect on shedding of helminth eggs showed that ivermectin, a member of the macrocyclic lactones (MLs) was efficacious against GIN parasites in both cattle and goats treated with this drug. There is hence a ray of hope for farmers who rear livestock extensively to still rear healthy and productive livestock free of infectious helminths provided that ivermectin is used rationally. Although, the overall treatment response of ivermectin in reducing FECs of GINs by this study agreed well with those reported in other studies (Wanyangu et al 1996; Maingi et al 1998; Waruiru et al 1998; Sissay et al 2006), the presence of Haemonchus contortus populations amongst ivermectin treated goats is cause for worry since failure to eliminate this GIN species, one of the most pathogenic nematodes of ruminants is likely to result in serious economic losses in terms of reduced productivity and in severe cases mortalities of affected animals (Uhlinger et al 1992).

Obvious suspicion of resistance to both albendazole and levamisole was evident among the studied goats. Although this study pioneered investigation on anthelmintic resistance in livestock under extensive system of management, the findings were in accordance with resistance situation in small ruminants under intensive production systems (Wanyangu et al 1996; Maingi et al 1998; Waruiru et al 1998). Similar albendazole and levamisole resistant nematodes has been reported in Ethiopia (Sissay et al 2006) in Tanzania (Bjorn et al 1990; Ngomuo et al 1990; Keyyu et al 2002) in South Africa (Van Wyk et al 1999; Vatta et al 2001; Bakunzi et al 2003) among other places. Despite the evidently clear prevalence of albendazole resistant GINs in goats, evidence for GINs resistant to levamisole was borderline. Moreover, this finding was quite unexpected and surprising since levamisole has not been used as an anthelmintic in Katumani before. However, because the efficacy of levamisole in treated goats was 90 % it suggests existence of a window of opportunity for slowing the resistance as suggested by Bakunzi (2003).

Although many reports on GIN resistance in sheep and goat farms in SSA (Bjorn et al 1990; Wanyangu et al 1996; Mukaratiwa et al 1997; Maingi et al 1998; Waruiru et al 1998; Gatongi et al 2003; Sissay et al 2006) have been made, AR in cattle GINs is a still a rare occurrence. This is perhaps because of the low frequency of anthelmintic treatments in cattle. One of the first reports on anthelmintic resistance in cattle in Africa was made in Mali where one study suspected presence of albendazole resistant GIN populations in cattle (Mungube et al 2013). Contrary to the findings by Mungube et al (2013), the present study has shown that albendazole is still efficacious against GINs within the cattle flock studied. However, levamisole, an imadathiazole tested in this study failed to produce an appreciable treatment response (FECRT was <95%) amongst treated cattle. This increased suspicion about possible presence of levamisole resistant GIN populations in the cattle at New Astra ranch. It is not clear why this was the case since levamisole is rarely used as an anthelmintic although it was strongly linked to the inflows into the ranch of breeding stock sourced from anthelmintic resistance hotspot areas within the central highlands and Rift Valley of Kenya (Maingi et al 1998; Waruiru et al 1998).

The magnitude of anthelmintic resistance involving levamisole was less severe in the New Astra ranch compared to the KALRO Katumani flock. This is could be because of the differences in the husbandry practices and management levels in the two herds. At the New Astra ranch, anthelmintic treatment is selectively done (treatment of the calves and sick animals) compared to the mass treatments done at Katumani. The infrequent use of anthelmintic drugs reduces the risk of resistance development since a reservoir of susceptible worms remains within the population which helps to dilute the resistant alleles (Hoste et al 2002). At the New Astra ranch, close interaction between cattle and wild herbivores like giraffes, wildebeests, antelopes and zebras is beneficial in maintaining refugia. The interfacing of cattle with wildlife ensures that the GINs in cattle after they are exposed to anthelmintic treatments are constantly diluted or replaced by the GIN nematodes from wildlife which are unexposed to drugs. This helps to delay resistance development in agreement with the consideration by most parasitologists that levels of refugia is the single most important factor contributing to selection for anthelmintic resistant parasites (Van Wyk 2002). Maintaining parasites in refugia is a key point in controlling and delaying the development of resistance, because the susceptible (anthelmintic sensitive) worms genes are preserved (Waghorn et al 2008).

Nematodes in goats are likely to develop resistance against common BZ formulations faster than those in sheep and cattle (Chartier et al 1999; Hennessy and Alvinerie 2002; Sangster et al 1993). This is because goats metabolise anthelmintics more rapidly than other domestic ruminants which may also accelerate the rate of selection for resistance in nematode parasites. Moreover, at KALRO Katumani, goats are kept intensively at high stocking rates which sometimes forces them to graze instead of browsing hence predisposing them to severe problems of GINs as observed by Sissay et al (2006). Under the intensified rearing conditions, very frequent treatments with anthelmintic drugs are practiced in response to the high helminth infection risk precipitating in the earliest and worst cases of anthelmintic resistance recorded in ruminant livestock (Jackson et al 1992; Sykes et al 1992). Anthelmintic resistance can also be selected at lower treatment frequencies, especially when the same drug is used over many years. This actually confirms that the high GIN prevalence of albendazole resistant nematode populations in goats could be that this drug has continuously been used in the control helminths at Katumani for well over 20 years. Coles (1995) reported the development of AR even when only two or three treatments were given annually.

It is known that the commonest GINs of small ruminants include Haemonchus contortus, Telodorsagia circumcinta, Trichostrongylus axei, Nematodirus spp, and Cooperia spp (Radostits et al 2002). Haemonchus contortus and T. circumcincta represent most of the parasite burdens seen in small ruminants, with H. contortus being present in highest numbers. Anthelmintic resistance is present in all of these parasites, but the prevalence is highest for H. contortus, making it the most economically important GIN of sheep and goats (Uhlinger et al 1992). This assertion was confirmed by the results of this study. With anaemia as the principal pathologic effect from infection with H. contortus (Radostits et al 2002), FAMACHA (van Wyk et al 2002; Kaplan et al 2004) can be an effective tool for identifying animals that require treatment (but only for H. contortus) as a substitute to the blanket mass worming of animals after every 3 months irrespective of their anaemic status. Using the FAMACHA will ensure that the number of anthelmintic treatments administered are greatly reduced which in turn significantly diminishes selection pressure for resistance and concomitant reduction in drug costs (Fleming et al 2006).

The results by this study showed a general reduction in faecal egg outputs comparing the FECs of the control group at pre-treatment and at post-treatment. This seemed to suggest that season influences helminth egg shedding in agreement with observations by Mungube et al (2013). It has also been reported that there is suppression of nematode egg output during the dry season, attributed to the different survival mechanisms of the various nematodes (Kaufmann and Pfister 1990). Some nematode species like Cooperia, Bunostomum and Oesophagostomum survive in their hosts as adults during the dry season compared to Haemonchus species which are hypobiosed (Kaufmann and Pfister 1990). This was actually confirmed by this study since eggs of Cooperia species which were not detected during pre-treatment phase (rainy season) were among those detected during the post-treatment phase sample collection which occurred during the dry season.

Conclusions

• Prolonged use of albendazole is likely to accelerate development of albendazole resistant GINs.

  
  

• The use of albendazole in treating GINs in goats in Machakos should be reconsidered owing to the prevalence of albendazole resistant nematodes.

  
  

• Although ivermectin is still largely effective against GINs in both cattle and goats, it is not effective against Haemonchus species in goats and Ostertagia species in cattle

Recommendations

• Discontinue the use of albendazole 10% for worm control in goats in and around KALRO Katumani.

  
  

• Despite the prevalence of multiple resistant Ostertagia species in cattle, albendazole 10% still remains an effective anthelmintic in the control of GINs in cattle.

  
  

• Since Haemonchus species were prevalent amongst both cattle and goats, adoption of FAMACHA for making treatment decisions will rationalize and prolong the use of anthelmintics especially ivermectin.

  
  

• Adpotion of integrated worm control practices like pasture management, improved nutrition among others may be of help in the face of multiple resistant Haemonchus and Ostertagia species

  
  

• Further studies using more robust molecular tools are needed to confirm prevalence of multiple resistant Haemonchus and Ostertagia species in the semi-arid region of Kenya.

  
  

• Further studies to establish anthelmintic handling and use practices as well as quantify active ingredients in marketed albendazole 10% and Nilzan plus should be conducted.

Acknowledgements

The authors acknowledge financial support from the Kenya Agricultural Productivity and Agribusiness Project (KAPAP). We also sincerely thank the livestock managers at KALRO Katumani Research Centre and the New Astra ranch for their cooperation.

References

Bakunzi F R 2003 Anthelmintic resistance of nematodes in communally grazed goats in a semi-arid area of South Africa. Journal South Africa Veterinary Association 74: 82-83.

Bjorn J, Monrad J, Kassuku A A and Nansen P 1990 Resistance to benzimidazole in Haemonchus contortus of sheep in Tanzania. Acta Tropica 48: 59-67.

Chartier C, Etter E, Pors I and Alvinerie M 1999 Activity of eprinomectin in goats against experimental infections with Haemonchus contortus, Teladorsagia circumcincta and Trichostrongylus colubriformis. Veterinary Record 144: 99–100.

Coles G C, Bauer C, Borgsteede F H M, Geerts S., Klei T R, Taylor M A and Waller P J 1992 World Association for the Advancement of Veterinary Parasi­tology (W.A.A.V.P) methods for the detection of anthelmintics resistance in nema­todes of Veterinary importance, Veterinary Parasitology 44: 35 – 44

Coles G C, Papadopoulos E and Himonas C A 1995 Tubulin, resistance and worms. Parasitology Today 11: 183–185

Coles G C 2002 Cattle nematodes resistant to anthelmintics: why so few cases? Veterinary Research 33: 481-489.

Coles G C, Jackson F, Pomroy W E, Prichard R K, von Samsom-Himmelstjerna G, Silverstre A, Taylor M A and Vercruysse J 2006 The detection of anthelmintic resistance in nematodes of veterinary importance. Veterinary Parasitology 136: 167-185. 

Demeler J, Van Zeveren A M J, Kleinschimdt N, Vercruysse J, Höglund J, Koopmann R, Cabaret J,  Claerebout E, Areskog M and von Samson-Himmelstjerna G 2009  Monitoring the efficacy of ivermectin and albendazole against gastro intestinal nematodes of cattle in Northern Europe. Veterinary Parasitology 160: 109-115.

Domke A, Chartier C, Gjerde B, Leine N, Vatn S, Østerås O and Stuen S 2012 Worm control practice against gastro-intestinal parasites in Norwegian sheep and goat flocks. Acta Veterinary Scandinavian 53: 29. doi: 10.1186/1751-0147-53-29

Fleming S A, Craig T, Kaplan R M, Miller J E, Navarre C and Rings M 2006. Anthelmintic resistance of gastrointestinal parasites in small ruminants. Journal of Veterinary International Medicine 20: 435-444

Gabrie S, Phiri I K and Vercruysse J A 2001. A survey on anthelmintic resistance in nematode parasites of sheep in Lusaka, Zambia. Onderstepoort Journal Veterinary Research 68: 271-274.

Gatongi P M, Njoroge J M, Scott M E, Ranjan S, Gathuma J M, Munyua W K, Cheruiyot H and Prichard R K 2003 Susceptibility to IVM in a field strain of Haemonchus contortus subjected to four treatments in a closed sheep-goat flock in Kenya. Veterinary Parasitology 110 (2): 235–240.

Hennessy D R and Alvinerie M 2002 Pharmacokinetics of the macrocyclic lactones: conventional wisdom and new paradigms In: Vercruysse J., Rew R.S. (eds) Macrocyclic lactones and antiparasitic activity. CAB International pp 97-123

Hoste H, Chartier C, Lefrileux Y, Goudeau C, Broqua C, Pors I, Bergeaud J P, Dorchies P H 2002 Targeted application of anthelmintics to control trichostrongylosis in dairy goats: result from a 2-year survey in farms. Veterinary Parasitology 110: 101–108.

Jackson F 1993 Anthelmintic resistance – the state of play, British Veterinary Journal 149: 123-138.

Jaetzold R, Schmidt H, Hornetz B and Shisanya C 2006 Farm Management Handbook of Kenya Vol II: Natural Conditions and Farm Management Information 2^nd Edition Part C, East Kenya, Subpart C1 Eastern Province, pp. 573.

Kaplan J 2004 Drug resistance in nematodes of veterinary importance: a status report. Trends in Parasitology 20: 477-481.

Kaufmann J 1996 Parasitic Infections of Domestic Animals: A Diagnostic Manual. Birkhäuser Verlag, Basel. pp 29-54.

Kaufmann J and Pfister K 1990 The seasonal epidemiology of gastrointestinal nematodes in N'Dama cattle in The Gambia. Veterinary Parasitology 37: 45-54.

Keyyu J D, Mahingika H M, Magwisha H B, Kassuku A A 2002 Efficacy of albendazole and levamisole against gastrointestinal nematodes of sheep and goats in Morogoro, Tanzania. Tropical Animal Health and Production 34 (2): 115-120.

Maingi N, Bjorn H, Gichohi V M, Munyua W K and Gathuma J M 1998 Resistance to benzimidazoles and levamisole in nematode parasites of sheep in Nyandarua District of Kenya. Acta Tropica 69: 31–40.

Mbah A H, Ogunrinade A F and Dina O A 1992 Benzimidazole resistance in strongyles of sheep in Ibadan, Nigeria. African Livestock Research 1: 16-17.

Michel  J F, Cawthorne R J G, Anderson R M, Armour J, Clarkson M J, Thomas R J 1982 Resistance to anthelmintics in Britain: husbandry practices and selective pressure, Facts and reflections. IV. Resistance of parasites to anthelmintics. A workshop in the Commission of European Communities animal pathology programme held at the Central Veterinary Institute, Lelystad, The Netherlands. pp. 41-50.

Monteiro A M, Wanyangu S W, Kariuki D P, Bain R, Jackson F, McKellar  Q A 1998 Pharmaceutical quality of anthelmintics sold in Kenya. Veterinary Record 142: 396-398.

Mukaratirwa R, Charakupa R and Hove T 1997 A survey of anthelmintic resistance on ten sheep farms in Mashonaland East Province, Zimbabwe. Journal of South Africa Veterinary Association 68: 140-143.

Mungube E O, Rexa F, Hinney B, Fritz S, Sanogo Y, Rösel K, Diall O, Randolph T and Clausen P H 2013 Detection of albendazole resistant gastro-intestinal nematodes in village cattle of south-eastern Mali. East African Agricultural and Forestry Journal 78 (1:, 113-118.

Ndamukong K J N and Sewell M M H 1992 Resistance to benzimidazole anthelmintics by trichostrongyles in sheep and goats in north-west Cameroon. Veterinary Parasitology 41: 335–339.

Ngomuo A J, Kassuku A A and Ruheta M R 1990 Critical controlled test to evaluate resistance of field strains of Haemochus contortus to thiophanate. Veterinary Parasitology 36: 21-26.

Prichard R K 1990 Biochemistry of anthelmintic resistance, Resistance of parasites to antiparasitic drugs: Round Table Conference, VII International Congress of Parasitology, Paris, Texas, United States.

Radostits O M, Blood D C and Gay C C 2002 Veterinary Medicine. A text book of the diseases of cattle, sheep, pigs, goats and horses. 13^th edn., Bailliere Tindall, USA.

Rao K P C, Ndegwa W G, Kizito K and Oyoo A 2011 Climate Variability and Change: Farmer Perceptions and Understanding of Intra-Seasonal Variability in Rainfall and Associated Risk in Semi-Arid Kenya. Experimental Agriculture 47: 267-291.

Sangster N C, Steel J W, Collins G H and Hennessy D R 1993 Comparative kinetic disposition of oxfendazole in sheep and goats before and during infection with Haemonchus contortus and Trichostrongylus colubriformis. Journal of Veterinary Pharmacology and Therapeutics 16: 245–253. doi: 10.1111/j.1365-2885

Sissay M M, Asefa A, Uggla A and Waller P J 2006 Anthelmintic resistance of nematode parasites of small ruminants in eastern Ethiopia: exploitation of refugia to restore anthelmintic efficacy. Veterinary Parasitology 135 (3-4: 337-346.

Suarez H V and Cristel S L 2007 Anthelmintic resistance in cattle nematode in the western Pampeana Region of Argentina. Veterinary Parasitology 144 (1-2): 111-117.

Sykes A R, McFarlane R G and Familton A S 1992 Parasites, immunity and anthelmintic resistance. in: Progress in sheep and goat research, CAB International, Wallingford, UK, , pp. 179-191.

Taylor M A, Hunt K R and Goodyear K L 2002 Anthelmintic resistance detection methods: a review. Veterinary Parasitology 103: 183–194

Uhlinger C and Kristula M 1992 Effects of alternation of drug classes on the development of oxybendazole resistance in a herd of horses. Journal of American Veterinary Medicine Association 201 51-55.

Van Wyk J A 2002 Refugia-overlooked as perhaps the most potent factor concerning the development of anthelmintic resistance. Onderstepoort Journal Veterinary Research 68: 55–67

Van Wyk J A, Stenson M O, Van der Merwe J S, Vorster R J and Viljoen P G 1999 Anthelmintic resistance in South Africa: surveys indicate an extremely serious situation in sheep and goat farming. Onderstepoort Journal Veterinary Research 66: 273-284.

Vatta A F, Letty B A, Van Der Linde M J, Van Wyk E F, Hansen J W and Kreck R C 2001 Testing for clinical anaemia caused by Haemonchus species in goats farmed under resource-poor conditions in South Africa using an eye colour chart developed for sheep. Veterinary Parasitology 99: 1-14.

Waghorn T S, Leathwick D M, Miller CM and Atkinson D S 2008 Testing the concept that leaving susceptible parasites in refugia will slow the development of anthelmintic resistance. New Zealand Veterinary Journal 56 (4): 158-163.

Wanyangu S W, Bain R K, Rugutt M K, Nginyi J M and Mugambi J M 1996 Anthelmintic resistance amongst sheep and goats in Kenya. Preventive Veterinary Medicine 25: 285–290.

Waruiru R M, Kogi J K, Weda E H and Ngotho J W 1998 Multiple anthelmintic resistance on a goat farm in Kenya. Veterinary Parasitology 175: 191-197.

Received 3 November 2015; Accepted 9 November 2015; Published 1 December 2015

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