STRONGYLOSIS IN EQUINES: A REVIEW.
We review strongylosis covering all aspects as it is one of the most important internal parasitic diseases of equines caused by nematodes of strongylidae family affecting more than 80% equids in the world. Majority of the work published has been focused on incidence epidemiology and control. Strongylus vulgaris one of the large strongyle is the most prevalent and pathogenic. Small strongyles exhibit mild symptoms of diarrhea and weight loss in the host whereas large strongyles show major pathogenesis. The pathogenesis encompasses severe enteropathy verminous arteritis damage of visceral organs embolisim / thrombosis leading to death and is mainly attributed to migrating larvae of parasites. In this report the scientific knowledge about development pathogenesis clinical findings diagnosis epidemiology treatment and control of strongylosis in equines has been reviewed for a period of last nine decades.
Inspite of substantial improvements in understanding the life cycle of strongyles adopting latest diagnostic techniques at molecular level and implementing the most modern control measures / treatment the disease is still prevalent and could not be eradicated from any part of the world. It is currently impossible to measure or detect the encysted larval load in living horse however there are exciting advancements in diagnosis of disease by means of molecular approaches such as cyathostomin gut-associated larval antigen-1 (Cy-GALA-1). The current strategy engaged in seasonal use of anthelmentics is the key to arrest the disease and overcome anthelmentic resistance. We conclude that there is still a need to thrash approaches / scientific knowledge towards understanding the problem to reduce economic / performance losses. The scientific studies for development of an effective vaccine are considered the need of the day.
Key words: strongylosis equines S. vulgaris arteritis faecal floatation migrating larvae. INTRODUCTION
Strongylosis has been reported from all parts of world and almost affects more than 90 % of horse population (Nielsen et al 2006). The small strongyles also called cyathostomins are among the most important intestinal nematodes of horses (Lyons et al. 1999) with almost 100% of horses infected with at least some species of small strongyles (Reinemeyer et al. 1984). Among the gastro-intestinal nematodes of horses large strongyle infections were diagnosed with an infection rate of 58.5% (Saeed et al. 2010). S. vulgaris has long been considered as one of the most common and pathogenic parasites of the horse (Claire and Masterson. 1987; Krecek et al. 1987; Tolliver et al. 1987; Peter and Waller. 1997; Gasser et al. 2004; Hubert et al. 2004; Martin et al. 2007; Toscan et al. 2012). It is estimated that 45 to 90 per cent of horses harbor S. vulgaris (McCraw and Slocombe 1976; Jubb et al. 1985).
The prevalence of infection is high and a few equines are likely to escape out of this disease by the end of their first year of life (0gbourne and Duncan 1985). Infestation with strongyles is complex and produces an inflammatory enteropathy which results in impaired intestinal motility and microcirculation (Love et al. 1999; Bechera et al. 2010; Neils et al. 2011; Pilo et al 2012). Clinically infection with small strongyles can cause mild disease symptoms such as weight loss loss of appetite poor hair coat intermittent diarrhoea lethargy deterioration of condition peripheral edema and disordered intestinal motility (McCraw and Slocombe 1976; Love and Duncan 1992; Love et al. 1992; Matthews and Morris 1995). After ingestion larvae travel through the digestive system to the large intestine
S. vulgaris migrate to the anterior mesenteric artery S. endentatus to the liver / flank area and S. equinus migrate to the liver and pancreas ( Owend and Slocombe. 1985; Monhan et al. 1995; Kuzmina et al 2012). Adult large strongyles live in the cecum and large intestine. Fourth (L4) and fifth (L5) stage larvae are responsible for arteritis necrosis and fibrosis of the cranial mesenteric artery and its branches (Patton and Drudge 1977; Duncan and Pirie 1985; Kuzmina et al. 2012). The occurrence of larvae in the cranial mesenteric artery was apparently known to the ancient Romans (McCraw and Slocombe 1974). Verminous arteritis in the cranial mesenteric artery and its branches has been reported as the cause of death in 10 to 33% of abdominal crises in the horse (Meads 1969; Proctor 1966) and in a few case studies of natural infections (Foster and Clark 1937; Ottaway and Bingham 1945; Deorani 1966).
Severe colic and death of horses is the consequence of thrombosis and embolism leading to infarction of the intestinal tract (Marinkovic et al. 2009). Many animals are not picked up early enough to interrupt the larval migration and are only diagnosed by faecal culture techniques indicating the presence of fertile adults (Morgan et al. 1994). Determining the number of strongyle eggs per gram of faeces (EPG) has been the most widely used method for diagnosing infection with adult strongyles (Herd 1995; Mathew and morris 1995; Kaplan 2010; Mahmood and Ashraf 2010). Molecular approaches have been developed that enable species identification of pre-adult stages of strongyle nematodes (Gasser et al. 2004; Matthews et al. 2004). Routine treatment for the removal of adult worms is essential as the large egg output of strongyles can cause contamination of a restricted area very rapidly.
Thiabendazole has been widely used and recently several other anthelmintics have been developed and approved for use in adult horses including benzimidazole compounds (Drudge et al 1975). The under dosing is the major cause of anthelmintic resistence (Toscan et al. 2012). Pasture management is considered a reliable control measure. The emergence of anthelmintic resistance demands review of old studies.
Development and Life Cycle: All species of equine strongyles have direct life cycles (Bucknell et al. 1995; Osterman 2005; Kuzmina 2006; Martin et al. 2007). There are significant differences in number of eggs in female uteri in various strongyle species where the egg number differs 50100 times between species (Kuzmina et al. 2012). Eggs are laid by adult female strongyles and passed in the faeces into the external environment where they hatch to the first stage larvae (L1s) at 12 39 OC with adequate moisture. The minimum temperature for eggs to hatch is 78 OC (Ogbourne 1975). From the egg on the ground / pasture L1 emerges which grows and molts to L2 and then to the L3 stage. The L3 is the infective stage and under optimal summer conditions it requires about ten days to two weeks to develop from the time the egg is passed. L3s retain an outer protective sheath and are more resistant to chilling and dehydration than the L1s and L2s.
Dehydration can prevent L3s from leaving the faeces and gaining contact to herbage. Most larvae climb no higher than 10 cm from the soil surface can move 15 cm horizontally and during rain L3s migrate from the faeces to the surrounding herbage most efficiently. (Bucknell et al. 1995; Edward 2007). After ingestion of L3 by the host they pass to the small intestine remove external covering and initiate the internal phase of development. Removal of protective covering depends upon the stimuli from physiological / biochemical conditions in the gut of the host. Larvae of large strongyles emerge from the sheath through an anterior cap whereas larvae of small strongyle escape via a longitudinal slit in the region of the oesophagus (Kuzmina et al. 2012). Removel of outer covering at 38 OC within 3 hours using an artificial intestinal fluid comprising trypsin pancreatin sodium bicarbonate and sodium dithionite has been achieved experimentally (Kuzmina et al. 2006).
Internal phase of large strongyles larval development encompasses a somatic migration whereas those of small strongyles burrow into the glands in the caecum and colon and become encysted with no further migration. (Eysker et al. 1986; Gasser et al 2004). In the submucosa next molting occurs ie L4 on about day 4 or 5 PI. Working against the flow of blood the L4s gradually move up the arterial system of the intestine. By eight days PI larvae have reached the cecal and ventral colic arteries. When these larger arteries are achieved the route of migration is marked by a twisty thread of fibrin on the intima and by day 14 larvae may be found in mural thrombi. The ileo-ceco-colic and cranial mesenteric arteries are reached between 11 and 14 days PI.
The traveling advance attains its climax by the 19th day at which time larvae may be found in almost any part of the arterial system but are always most abundant in arteries close to the origin of the cranial mesenteric artery (McCraw and Slocombe 1976; Hopfer et al. 1984; Osreman 2005). The molt to the fifth stage (L5) occurs as early as 9 days PI and by 120 days. At this stage most larvae are preadults measuring up to 18 mm long. S. vulgaris larvae tend to remain in the arterial site until they molt to the fifth stage though many fourth stage larvae are apparently swept away before the last molt occurs. Larval size and the thrust from the flow of blood are important factors in the separation of larvae from arterial lesions. The preadult larvae reach the small arteries on the serosal surface of the large intestine and terminal small intestine. Unable to migrate further in arteries the young one of large strongyle become encased in pea-sized nodules.
These nodules are numerous four months after infection; after their escape from nodules into the lumen of the intestine S. vulgaris require another six to eight weeks before reaching sexual maturity. The prepatent period (the time from ingestion of L3s to the excretion of eggs in the faeces) is 57 months (McCraw and Slocombe 1976; Kuzmina et al. 2006; Martin et al. 2007; Niels et al 2011). The prepatent period among the species varies from 6 months to 12 months (Urquhart et al. 1996; Osterman 2005). Prenatal infection lacks any evidence (Andersen 2013). Deviant larvae can sometimes migrate to kidneys thoracic cavity and testis. The pattern of larval migration among large and small strongyles differs; (1). Large strongyles larvae migrate widely within the host through extra-alimentary tissues with a minimum of 6 month prepatent period (Kaplan 2004). (2)
Small strongyles larvae invade the lining of the cecum and ventral colon where they grow within fibrous cysts in the mucosa or submucosa and can reside as long as 2.5 years (Reinemeyer 2009). Now it is understandable that the ecological requirements for larval development and insistence are matching for both groups as inferred from above mentioned studies and described by Hutchinson et al. 1989 too however no dependable information is available on the environmental conditions required for the development of L1s and L2s. For small strongyles the life cycles of individual species have also not yet been determined. The question on influence of fecundity on proportion of species in strongyles community needs further studies.
Pathogenesis: Naturally infected horses usually carry a mixed load of large and small strongyles in the intestine (Owend and Slocombe 1985). The pathogenesis of strongylus has been studied based on elucidation of experimental mono specific infections (Drudge et al 1966; McCraw and Slocombe 1974; Malan et al. 1982; Alam et al 1999). The damage caused by large and small strongyles is attributed to larval stages. Small strongyles have small buccal capsules and feed superficially on the mucosa (Ogbourne and Duncan 1985). Large strongyles have large buccal capsules which they attach to the intestinal mucosa pull out a plug of tissue absorb the host cells crack the blood vessels and suck blood feed on the mucosa and consume blood (Levine 1980). Hemorrhage occurs subsequent to feeding at the injured site which eventually is marked by a scar. The larvae cause minimal inflammatory response as long as they remain encysted however their synchronous emergence of large number results in diffuse
Inflammation of the cecum and ventral colon (Love et al. 1999). In all the cases a normocytic normochromic anaemia is observed in affected equines (Ogbourne 1985). The L4s and L5s migrate through the arterial system and cause verminous arteritis with marked intima thickening infiltrated with inflammatory cells. They can cause mechanical damage and inflammation in the liver pancreas and peritoneal cavity. A considerable reduction of lesions in the cranial mesenteric artery was found approximately nine months after infection with S. vulgaris larvae (Duncan and Pirie 1985). Incidence of lesions is 86% in the cranial mesenteric artery followed by 62.5% in the cecal and colic arteries (Poynter 1969). Depending on larval burden infected horses show clinical signs of pyrexia anorexia and colic. Colic has long been considered related to thrombosis or embolism in these vessels (Bueno et al 1979).
In older horses the cranial mesenteric and ileo-ceco-colic arteries are often encased in a large nodular mass. Occlusion of the right coronary artery was observed due to S. vulgaris larvae and in the kidney too (Cronin and Leader 1982). S. vulgaris was also suspected as an important cause of cerebrospinal nematodiasis (Little et al. 1974). Several studies have reported alterations in blood parameters and blood chemistry as a consequence of S. vulgaris infection including a decrease in RBC PCV total serum proteins and an increase in WBC. Intestinal haemorrhages lead to reduced RBC survival loss of albumin in intestine leads to increased albumin catabolism (Drudge et al. 1966; Duncan and Pirie 1985; McCraw and Slocombe 1976; Patton and Drudge 1977; Peter and Waller 1997; Nielsen 2012). In ponies with a natural infection a reduced level of ileo-ceco-colic motility has been demonstrated with electromyographic techniques (Bueno et al. 1979). At necropsy the crater-like ulcers caused by large strongyles are often more numerous than the worms suggesting that they move periodically to new sites of attachment (Andersen et al. 2013).
Due to the difficulty in differentiating the effect of species in naturally acquired mixed infections there is lack of detailed information on the pathological effects of individual species of small strongyles (Ogbourne 1976; Reinemeyer et al. 1984; Lyons et al. 1997). Several studies have shown a marked decrease of S. vulgaris infection worldwide but on the other hand many studies showed that this parasite continues to exert its pathogenic role even when its detection rate is quite low within the strongyle population infecting horses (Pilo et al. 2012).
Clinical Signs: The acute signs related with large strongyles are due to migrating larvae and are seen during the first few weeks after infection. The severity of signs is related to the number of larvae ingested the age and previous occurrence of the host. (McCraw and Slocombe 1985). Older horses are often observed to have arterial lesions without a history of specific signs although signs detected in field cases can be correlated with findings at necropsy (Gasser et al 2004). Anemia emaciation poor coat and poor performance are frequently attributed to large strongyles while in the intestine. Diaarrhoea is more common sign in small strongyle infection than with large strongyles (McCraw and Slocombe 1976). The main clinical sign in small strongyle infection is weight loss (Love et al. 1999). Other typical clinical signs are profuse / sudden onset of diarrhoea loss of body condition debility with normal appetite and subcutaneous oedema of the limbs / ventral abdomen.
Death is relatively common with mortality rate of greater than 50% (Love et al. 1999). The encysted larvae of small strongyles can emerge synchronously from intestinal wall leading to the clinical disease called larval cyathostominosis' which is associated with clinical signs of oedema diarrhoea pyrexia weight loss colic and can be fatal in up to 50% of cases (Gasser et al 2004). Fever in S. vulgaris infection is attributed to tissue damage or a toxic substance elaborated by larvae. The most steady change in early S. vulgaris infection would result a rapid increment in total white cell (WBC) counts. These values rise sharply during the first three weeks to levels of 17000 to 22700/mm3. Eosinophils values will increase after the second week and demonstrate little change in acute infection. Increments in serum total protein and globulin fractions occur as early as the first week following infection.
Thrombus formation can block arteries causing infarction of intestinal walls and/or intermittent lameness and is commonly associated with clinical signs of marked pyrexia anorexia severe colic and death (Pilo et al. 2012). Under natural conditions severe symptoms are rarely seen because foals may tolerate large numbers of larvae ingested in small doses over a long period (Urquhart et al 1996). Number of adult strongyles in the intestine required to provoke clinical signs are not yet known and need elaboration / further studies.
Diagnosis: Signs and symptoms are not valuable for diagnosis. It has usually relied on the use of the method of faecal flotation (Duncan and Pirie. 1985; Nautrup et al 2003; Gasser et al. 2004; Kaplan and Nielsen. 2010; Andersen et al. 2013). Faecal egg counts are useful for comparing the efficacy of anthelmintic compounds detecting drug resistance and determining the correct gap between anthelmintic treatments (Herd 1992; Warnick 1992). Since it is not possible to distinguish strongyle eggs of different species morphologically faecal samples are cultured to allow the development to L3s which may be collected for study. The Baermann technique is also a successful method for the recovery of small strongyles immature larvae in the faeces of clinically diseased horses. In case of larval strongylosis fecal egg count technique is assumed to be of no value. It is impossible currently to measure or even detect the encysted larval load while the horse is still alive (Osterman 2005).
A method for detecting mucosal larval stages would be valuable in the diagnosis of larval strongylosis. Recently cyathostomin (small strongyles) gut-associated larval antigen-1 (Cy-GALA-1) has been identified which is a target of serum IgG (T) responses in experimentally and naturally infected horse populations (McWilliam et al. 2010). Within the past two decades molecular approaches have been developed that enable species identification of pre-adult stages of strongyle nematodes like characterization of strongyle nematode ribosomal DNA sequences (Gasser et al. 2004). The first and second internal transcribed spacers (ITS-1 and 2) and the intergenic spacer (IGS) have been used as genetic markers for species identification. From the ITS sequences species-specific oligonucleotide primers for some of the most common species (S. vulgaris
C. catinatum C. nassatus C. longibursatus and C. goldi) have been designed and used in a PCR system thus allowing species-specific amplification of parasite DNA in eggs and larvae (Hung et al. 2000). IGS oligoporbes are used to study the effect of anthelmintic treatment at the species level (Matthews et al. 2004). IGS oligoprobes have been used in a PCR-ELISA (hybridisation assay) for the detection of PCR products from L4s collected from horses suffered from diarrhoea (Nielsen 2012). A microchip-based capillary electrophoresis technology has been employed successfully for species differentiation of closely related cyathostomins (Posedi et al. 2004). A copro antigen ELISA has shown promise with moderate to good diagnostic sensitivity and specificity as well as a positive correlation with worm numbers (Kania and Reinemeyer 2005; Skotarek et al. 2010). Change in the blood picture associated with S. vulgaris is not unlike that seen in bacterial infections (Drudge et al. 1984).
Alterations in blood biochemical and haematological parameters can be detected in a proportion of infected horses. Hypoalbuminaemia is a common finding in naturally infected horses which is probably due to the increased permeability of the intestines. A rise of b-globulin in serum has also been reported in natural infections. A marked reduction of serum fructose amines (glycated serum proteins) has been reported for horses with experimental cyathostomin infection (Dowdall et al. 2004)
Foregoing in view it is evident that recent advancements in molecular approaches lack specific diagnosis because of immunological cross-reactivity among species. The need of a reliable diagnostic assay to detect larval cyathostomins is still obligatory. There is a big need for specific criteria in the diagnosis of verminous arteritis in horses. Thus at the present time PCR cannot be recommended as an efficient and reliable means of surveillance. Cy-GALA-1 identification in small strongyles infestation is an exciting advancement but a semi quantitative assay has to be developed for authentication.
Epidemiology and Prevalence: Strongyles infestation can involve millions of nematodes covering a large range of species (Ogbourne 1975; Ogbourne 1976; Reinemeyer et al. 1984; Bucknell et al. 1995; Osterman 2005). In a single host 17 different species have been recorded (Bucknell et al. 1995). More than 50 species of equine strongyles have been reported (Lichtenfels et al. 2002). Small strongyles involve 80% of the total parasite population in a horse. The highest incidence of infection in yearlings (nearly 90%) and the lowest (46.6%) in foals has been recorded. Egg production varies seasonally and it has been demonstrated that least egg production occurs in winter rising during the spring with maximal production during August / September (Poynter 1954). In Ontario it was found that egg counts were high in August but during May to July were lesser for thoroughbred standardbred and show horses than for pleasure or commercial animals. (Slocombe and McCraw 1973).
The percentage of large strongyles viable eggs was lowest in winter but maximum in May and remained high throughout the summer (Ogbourne 1975). Higher egg excretion has been recorded in spring and summer (Herd 1990 Saeed et al. 2010). Season has no impact on the prevalence of strongyle infections but shedding intensity of strongyle eggs is affected by season and significantly higher egg excretion was recorded in spring and summer (Nielsen 2012). Higher EPG were recorded in young horse (= 3 year) as compared with lder horses no difference in the prevalence of strongyle infections as influenced by sex and excretion of eggs was also not affected by the sex of the animals (Saeed et al. 2010). L3s survival on pasture was estimated at 2-4 weeks in the summer wet season and 8-12 weeks in the autumn-winter dry season (April- August). Hot dry spring weather (pre-wet season) is the most unfavourable for larval development migration and survival (Hutchinson et al. 1989).
L3s were recovered from herbage samples from plots 3-4 weeks after the faeces had been deposited during May-October (Ramsey et al. 2004). The proportion of larvae successfully surviving during winter appeared to be maximal in faecal deposited on pasture in September of the previous year (up to 42.0% of the initial number of larvae). Larvae were observed surviving winter in soil beneath the faecal pats (Kuzmina et al. 2006). L3s are capable of surviving severe cold especially under a protective snow cover which somewhat stabilizes the climatic variations (Urquhart et al. 1996). Usually 90% of the adult worms are distributed throughout the dorsal and ventral colon and the remaining 10% are found in the caecum (Reinemeyer et al 1984; Gawor 1995; Collobert-Laugier et al. 2002). Prevalence of strongylosis is not affected by age or sex (Saeed et al. 2010).
Heavy worm loads may be found in individuals of all ages and horses may contribute to infected pastures throughout their lives (Osterman 2005). The occurrence of S. vulgaris has greatly decreased in recent decades (Herd 1990). In herds where anthelmintics have not been used to control parasites the prevalence has remained high (Gawor 1995).
We believe that the available studies indicate major climatic and seasonal differences in rates of development and persistence of free-living stages specifically the infective stage L3. This enables us to focus anthelmintic treatments at times when adequate refugia are present. In broad terms it is therefore necessary to avoid treatments of strongylosis during the winter months in cold temperate climates and during summer months in warm/hot climates in order to retard the development of anthelmintic resistance. Since understanding of larval ecology is prerequisite for development of rational anthelmintic control programs therefore more research is needed for a thorough quantitative description of strongyle larval bionomics and a better understanding of their basic epidemiology.
Control and Treatment: Usually equines are treated with anthelmintic drugs to eliminate adult strongyles from the large intestines to prevent excessive contamination of pastures with eggs and L3s. Thiabendazole has been widely used and several other drugs have been developed or approved for use in adult horses including benzimidazole tetrahydropyrimidines and organic phosphorus compounds (Drudge et al 1975; Tolliver 1987; Drudge et al. 1984; Saeed et al. 2008). Thiabendazole at the rate of 250 mg/kg body weight given through stomach tube on two consecutive days is useful (Coffman and Carlson 1971). Currently there are three main classes of commonly-used drugs categorised by their mode of action: the benzimidazoles (e.g. thiabendazole cambendazole fenbendazole and oxibendazole) pyrantel and the macrocyclic lactones e.g. ivermectin and moxidectin (Gasser et al. 2004). In the 1990's treatment intervals practiced for adult horses were 8 weeks for ivermectin and 46 weeks for other anthelmintics. Many combinations of macrocyclic lactones (abamectin ivermectin moxidectin) including ivermectin combined with pyrantel (tetrahydropyrimidine) and ivermectin combined with praziquantel (pyrazinoisoquinolin derivative) a pharmaceutically formed generic paste containing ivermectin 4% were tested for their effectiveness to control gastrointestinal nematodes of horses (Toscan et al. 2012).
Alike formulations of ivermectins had different efficacies calculated by reduction of EPG (Mariana et al. 2010). Stages of efficacy of the tested drugs varied against S. edentatus S. equinus and S. vulgaris. The generic paste (ivermectin 4%) was less effective than the conservative drugs. The efficacy of Oxafex Ivomec and Farbenda has been established as 94.7 98 and 81% respectively on day l4-post medication. On day 28th post medication it was 100% 96% and 86% respectively (Saeed et al. 2008). Now a days it is recommended to reduce the treatment intensity significantly to holdup further development of anthelmintic resistance (Kaplan and Nielsen 2010). In severe enteropathy the administration of non steroidal anti inflammatory agent is also required. Single IV dose of 0.6 mg.kg-1body weight meloxicam once a day is recommended for horses (Mahmood and Ashraf 2010).
There is general agreement that the traditional treatment at frequent intervals should be abandoned and that parasite control be maintained with far fewer anthelmintics (Nielsen 2012). Prevention by routine deworming of horses is unnecessary in all regions during the 6 month period that comprises the unfavorable season for strongyle transmission. During this interval environmental conditions largely prevent new parasites from developing. Even if horses have high egg counts during that period relatively few of those eggs can develop into adults. Therefore the goals of parasite control are being accomplished by the climate and compound treatment is not required (Reinemeyer 2009). Biological control especially the predacious fungi have established good potential as an adjunct for strongyle control and such a product could easily have a market in equine establishments.
There is a rising market for nematophagous fungal spores and the biotech industry must be encouraged to finalize and market a product for equine usage. Commercial products are being promoted in Australia (Edward 2007). Since small numbers of infective larvae may have grave effects on foals thus mares in foal or newly foaled should be regularly examined for strongyle eggs and treated with suitable anthelmintics. Climatic influences cannot effectively clean pastures from one grazing season to the next (Martin et al. 2007). The use of herbal compounds as anthelmentics against strongylosis is yet to be explored.
Alam S.S. A.N Anwar and M.N. Khan (1999). Studies on strongylosis in equines with special emphasis on haematology and chemotherapy. Pak J. Biological Sciences. 2: 634-636. Andersen U. V. I. T. Hakansson T. Roust M. Rhod K .
E. Baptiste and M. K Nielsen (2013). Developmental stage of strongyle eggs affects the outcome variations of real-time PCR analysis. Vet Parasitol. 191: 191 196. Bechera A.M. M. Mahlingc M.K. Nielsend K. Pfisterb and Salzburg (2010). Selective anthelmintic therapy of horses in the federal states of Bavaria Germany. An investigation into strongyle egg shedding consistency. Vet. Parasitol. 171: 116 122.
Bucknell D. G. R. B. Gasser and I. Beveridge (1995). The prevalence and epidemiology of gastrointestinal parasites of horses in Victoria Australia. Int. J. Parasitol. 25: 711724. Bueno L. Y. Ruckeluesch and P. Dorchies (1979).
Disturbances of digestive motility in horses associated with strongyle infection. Vet. Parasitol. 5: 253-260. Claire Nichol. I and W.J. Masterson (1987). Molecular and Biochemical Characterisation of surface antigens of Strongylus vulgaris of potential immunodiagnostic importance. Parasitology. 25: 29-38 Coffman J. R. and K. L. Carlson (1971). Verminous arteritis in horses. J. Am. vet. med. Ass. 158: 1358-1360.
Collobert-Laugier C. H. Hoste C.Sevin and P. Dorchies (2002). Prevalence abundance and site distribution of equine small strongyles in Normandy France. Vet Parasitol. 110: 77-83. Cronin M. T. I and G. H. Leader (1952). Coronary occlusion in a thoroughbred colt. Vet. Rec. 64: 8.
Deorani V. P. S. (1966). Studies on pathological lesions due to Strongylus vulgaris infection in an Indian pony. Indian Vet. J. 43: 865-867. Dowdall S. M .J. C. J. Proudman T. R. Klei T. Mair and J. B. Matthews (2004). Characterisation of IgG(T) serum antibody responses to two larval antigen complexes in horses naturally- or experimentally-infected with cyathostomins. Int. J. Parasitol. 34: 101108.
Drudge J. H. Lyons E. T. Szanto J. (1966). Pathogenesis of migrating stages of helminths with special reference to Strongylus vulgaris. In: Soulsby EJL editor. Biology of Parasites. New York: 199214. Drudge J. H. E. T. Lyons and S. C Tolliver. (1975). Critical tests of suspension paste and pellet formulations of cambendazole in the horse. Am. J. Vet. Res. 36: 435-439.
Drudge J. H. Lyons E. T. Tolliver S. C. (1984). Critical tests of morantel-trichlorfon paste formulation against external parasites of the horse. Vet. Parasitol. 14: 55-64. Duncan J.L. (1975). Immunity to Strongylus vulgaris in the horse. Equine Vet. J. 7: 4-11. Duncan J.L. and H. M. Pirie (1985). The pathogenesis of single experimental infections with Strongylus vulgaris in foals. Research in Vet. Sci. 18: 82-93.
Edward C. (2007). Integrated pest management for the horse farm. Publication No. 07/090 Rural Industries Research and Development Corporation Barton Australia. Eysker. M. J. Jansen F.N.J. Kooyman and M.H. Mirck. (1986). Control of strongylosis in horses by alternative grazing .Vet Parasitology. 19: 103- 115.
Eysker. M. J. Jansen F.N.J. Kooyman M.H. Mirck and T.H. Wensing (1988). Comparison of two systems for cythostome infections in the horse and further aspects of epidemiology of these infections. Vet. Parasitology. 22: 105-112. Foster A. O. and H. C. Clark (1937). Verminous aneurysm in equines of Panama. American J. Tropical Medical Hygiene. 17: 85-89. Gasser R. B. G. C. Hung N. B. Chilton and I. Beveridge (2004). Advances in developing molecular- diagnostic tools for strongyloid nematodes of equids: fundamental and applied implications. Molecular and Cellular Probes 18: 3-16.
Gawor J. J. (1995). The prevalence and abundance of internal parasites in working horses autopsied in Poland. Vet. Parasitol. 58: 99-108. Herd R. P. (1990). The changing world of worms: The rise of the cyathostomes and the decline of Strongylus vulgaris. The Compendium on Continuing Education for the Practicing Veterinarian. 12: 732-736. Herd R. P (1992). Performing equine fecal egg counts.
Vet. Med. 87: 240-244. Hopfer S.M. H.J. Van Kruiningen B.V. Amsterdam and W.H. Daniels (1984). The elimination of equine strongyles and haematological and pathological consequences following larvicidal dose of thiabendazole. Vet. Parasitol. 14: 21-32.
Hubert J.D. T. L. Seahorn T. R. Klei G. Hosgood D. W. Horohov and R. M. Moore (2004). Clinical signs and hematologic cytokine and plasma nitric oxide alterations in response to Strongylus vulgaris infection in helminth-naAve ponies. Can. J. Vet. Res. 68: 193200. Hutchinson G.W. S.A. Abba and M.W. Mfitilodze (1989). Seasonal translation of equine strongyle infective larvae to herbage in tropical Australia. Vet Parasitol. 33: 251-263.
Hung G. C. N. B. Chilton I. Beveridge and R. B. Gasser (2000). Molecular systematic framework for equine strongyles based on DNA sequence data. Int J. of Parasitology. 30: 95-103. Jubb K. V. F. P. C. Kennedy and N Palmer (1985). In: Pathology of Domestic Animals 3rd edit. 3. Academic Press Orlando: 50.
Kaplan Ray M. R. Thomas klel T. Eugene Lyons Guy Luster H. Charles Courtney D. Dennis French C. Sharon Tolliver N. Anand Vidyashanka and Ying Zhao (2004). Prevalence of anthelmentic resistant cyathostomes on horse farm. JAVMA 6: 225. Kaplan R. M. and M. K. Nielsen (2010). An evidence based approach to equine parasite control. Equine Vet Educ. 22: 306-16. Kania S. A. and C. R. Reinemeyer (2005). Anoplocehala perfoliata coproantigen detection. a preliminary study. Vet. Parasitol. 127: 115119. Krecek R.C. F. S. Malan R. K. Reinecke and V. Vos (1987). Nematode parasites from Burchell's zebras in South Africa. J. Wildl. Dis. 23: 404 411.
Kuzmina T. A. Y. I. Kuzmin and V. A. Kharchenko (2006). Field study on the survival migration and overwintering of infective larvae of horse strongyles on pasture in central Ukraine Vet. Parasitol. 141: 264272. Kuzmina T. A. E. T. Lyons S. C. Tolliver I. I. Dzeverin and V. A. Kharchenko (2012). Fecundity of various species of strongylids (Nematoda: Strongylidae) parasites of domestic horses. Parasitol Res. 111: 22652271. Lamb J. D. Gayle. Hallowel and M. Hany (2012).
Lipodemic analysis of serum from horses with strongyle infection. J. Vet Sci Technol. 3: 6 Lendala S. M.M. Larsena H. BjArnb J. Cravenc M. Chrieala and S.N. Olsend (1998). A questionnaire survey on nematode control practices on horse farms in Denmark and the existence of risk factors for the development of anthelmintic resistance.Vet Parasitol.78: 49-63. Levine N.D. (1980). Nematode parasites of domestic animals and of man. (2ndedition). Burgess Publishing Company Minneapolis Minnesota. Little P. B. L. Sein and P. Fretz (1974). Verminous encephalitis of horses: Experimental induction with Strongylus vulgaris larvae. Am. J. vet. Res. 35: 1501-1510.
Love S. and J. L. Duncan. (1992). The development of naturally acquired cyathostome infection in ponies. Vet. Parasitol. 44: 127142. Love S. D. Murphy and D. Mellor (1999). Pathogenicity of cyathostome infections. Vet. Parasitol. 85: 113122. Lyons E. T. S. C. Tolliver S. S. Collins J. H. Drudge and D. E. Granstrom (1997). Transmission of some species of internal parasites in horses born in 1993 1994 and 1995 on the same pasture on a farm in central Kentucky. Vet Parasitol. 70: 22540.
Lyons E.T. S.C. Tolliver J.H. Drudge (1999). Historical perspective of cyathostomes: prevalence treatment and control programs. Vet. Parasitol. 85: 97112. Mahmood K.T. and M. Ashraf (2010). Pharmacokinetics of ecofriendly meloxicam in healthy horses. Pakistan J. Sci. 62 (3): 198-201.
Malan F. S. V. de Vos R. K. Reineke and J. M. Pletcher (1982). Studies on Strongylus asini. I. Experimental infestation of equines. Onderstepoort J. Vet. Res. 49: 1513. Mariana Ionita K. Daniel Howeb T. Eugene Lyons Sharon C. Tolliver M. Ray Kaplan Ioan Liviu Mitrea and Michelle Yeargan. (2010). Use of a reverse line blot assay to survey small strongyle (Strongylida: Cyathostominae) populations in horses before and after treatment with ivermectin. Vet. Parasitol. 168: 332337.
Marinkovic D. A. K. Sanja V. Krstic and K. Milijana (2009). Morphological findings in the cranial mesenteric artery of horses with verminous arteritis. Acta Vet. 59: 231241. Martin. K. Nielsen Ray M. Kaplan M. Thamsborg Jesper Monrad Susanne and N. Olsen. (2007). Climatic influences on development and survival of free-living stages of equine strongyles. The Vet. J. 174: 2332. Martin K. Nielsen S. David Peterson Jesper M.Thamsborg Susanne N. Olsen M. Ray and Kaplan. (2008). Detection and semi- quantification of Strongylus vulgaris DNA in equine faeces by real-time quantitative PCR. Parasitology. 38: 443453.
Matthews A.G. and J. R. Morris (1995). Cyathostomiasis in horses. Vet. Rec. 136: 52 Matthews J. B. J. Hodgkinson S. M. J. Dowdall and C. J. Proudman (2004). Recent developments in research into the Cyathostominae and Anoplocephala perfoliata. Veterinary Research 35: 371-381.
McCraw B. M. and J. O. D. Slocombe (1974). Early development of and pathology associated with Strongylus edentatus. Can. J. comp. Med. 38: 124-138. McCraw B. M. and J. O. D. Slocombe (1976). Strongylus vulgaris in the horse: a review. Canadian Vet. J. 17: 150-157.
McCraw B.M. and J. O. D. Slocombe (1985). Strongylus equinus: development and pathological effects in the equine host. Can J Comp Med. 49: 37283. McWilliam H.E.G. A. J. Nisbet S. M. J. Dowdall J. E. Hodgkinson and J. B. Matthews (2010). Identification and characterisation of an immunodiagnostic marker for cyathostomin developing stage larvae. Int. J. Parasitol. 40: 265275.
Nautrup O. S. Schumann T. Pedersen and A. L. Eriksen (2003). Recovery of live immature cyathostome larvae from faeces of horses by Baermann technique. Vet Parasitol. 116: 259-263. Niels C. J. Kyvsgaarda L. Jenny l. Line Andreasena A. Luz L. Olivaresb K. Martin Nielsenc and M. Jesper (2011). Prevalence of strongyles and efficacy of fenbendazole and ivermectin in working horses in El Sauce Nicaragua. Vet Parasitology. 181: 248 254. Nielsen M. K. J. Monard and S.N. Olsen (2006). Prescription -only anthelmintics- a questionnaire surveillance and control of equine strongyles in Denmark. Vet Parasitol. 135: 47 55.
Nielsen M. K. (2012). Sustainable equine parasite control: Perspectives and research needs Vet Parasitol. 185: 32 44. Ogbourne C. P. (1975). Epidemiological studies on horses infected with nematodes of the family Trichonematidae . Int J Parasitol 5: 667720. Ogbourne C. P. (1976). The prevalence relative adundance and site distribution of nematodes of the subfamily Cyathostominae in horses killed in Britain. J. Helminthol. 50: 203214.
Ogbourne C.P. and J. H. Duncan (1985). Strongylus vulgaris in the horse: its biology and veterinary importance (2nd edition). Commonwealth Inst. Parasitol. Misc. Publ. No. 4 Commonwealth Agricultural Bureaux Farnham Royal United Kingdom: 68. Osterman Lind E. (2005). Prevalence and control of strongyle nematode infections of horses in Sweden. Doctoral thesis. ISSN 1652-6880 ISBN 91-576-7028- 5. Ottaway C. W. and M. L. Bingham (1945). Further observations on the incidence of parasitic aneurysm in the horse. Veterinary Record. 58: 155-159.
Owend J. and Slocombe. (1985). Pathogenesis of helminths in equines. Vet Parasitol. 18 : 139- 153. Patton S. and J. H. Drudge (1977). Clinical response of pony foals experimentally infected with Strongylus vulgaris. American J. Vet. Res. 38: 2059-2066.
Peter J and Waller. (1997). Anthelmintic resistance. Vet Parasitol. 72: (1997) 391-412. Pereira J.R. and S.S.S. Vianna. (2006). Gastrointestinal parasitic worms in equines in the Paraiba Valley State of Sao Paulo Brazil. Vet Parasitol. 140: 289295 Pilo C. A. Alteaa S. Pirinob P. Nicolussic A.
Varcasiaa M. Genchid and A. Scalaa (2012) Strongylus vulgaris (Looss 1900) in horses in Italy: Is it still a problem Vet Parasitol. 184: 161167. Posedi J. M. DrogemA1/4ller T. Schnieder J. Hoglund J. R. Lichtenfels and G. V. Samson- Himmelstjerna (2004). Microchip capillary electrophoresis-based comparison of closely related cyathostomin nematode parasites of horses using randomly amplified polymorphic DNA polymerase chain reaction. Parasitology Research. 92: 421-429.
Poynter D. (1954). Seasonal fluctuation in the number of strongyle eggs passed by horses. Vet. Rec. 66: 74-78. Poynter D. (1969). Some observations on the nematode parasites of horses. In Proceedings of the Second International Conference on Equine Infectious Diseases. Paris 2: 269-289. Ramsey Y. H. R. M. Christley J. B. Matthews J. E. odgkinson J. McGoldrick and S. Love (2004). Seasonal development of Cyathostominae larvae on pasture in a northern temperate region of the United Kingdom. Vet Parasitol. 119: 307-318.
Reinemeyer C.R. S. A. Smith A. A. Gabel and R. P. Herd (1984). The prevalence and intensity of internal parasites of horses in the USA. Vet. Parasitol. 15: 7583. Reinemeyer C. R. (2009). Controlling Strongyle Parasites of Horses. A Mandate for Change 352 Special Lectures. 55. AAEP Proceedings.
Saeed K. Z. Qadir S. A. Khan K. Ashraf and S. Nazir (2008). Evaluation of some broad spectrum antiparasitic drugs against natural strongyle infections in horses. J. Anim. Pl. Sci. 18: 64-66. Saeed K. Z. Qadir K. Ashraf and N. Ahmad (2010). Role of intrinsic and extrinsic epidemiological factors on strongylosis in horses. J. Anim. Pl. Sci. 20 (4): 277-280. Skotarek S.L. D. D. Colwell and C. P. Goater (2010). valuationof diagnostic techniques for Anoplocephala perfoliata in horses from Alberta Canada. Vet. Parasitology. 172: 249 255.
Tolliver S.C. E. T. Lyons and J. H. Drudge (1987). Prevalence of internal parasites in horses in critical tests of activity of parasiticides over 28- year period (19561983) in Kentucky. Vet. Parasitol. 23: 273284. Toscan G. A. S. Cezar R. C. F. Pereir G. B. Silva L. A. Sangioni L.S.S. Oliveir and F.S.F. Vogel (2012). Comparative performance of macrocyclic lactones against large strongyles in horses. Parasitology. 61: 550553. Urquhart G. M. J. Armour J. l. Duncan A. M. Dunn and F. W. Jennings (1996). Vet Parasitology. 2nd edn. Blackwell Science Ltd. Oxford. pp 4- 10: 42-47. Warnick L. D. (1992). Daily variability of equine faecal strongyle egg counts. Cornell Veterinarian. 82: 453-46.
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|Publication:||Journal of Animal and Plant Sciences|
|Date:||Feb 28, 2015|
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