Review Article

Amphistome infections in domestic and wild ruminants in East and Southern Africa: A review

Davies M. Pfukenyi, Samson Mukaratirwa

Received: 02 Nov. 2017; Accepted: 11 Sept. 2018; Published: 18 Oct. 2018

Copyright: © 2018. The Author(s). Licensee: AOSIS.
This is an Open Access article distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.


In this article, the main amphistome species infecting domestic and wild ruminants in East and Southern Africa, their snail intermediate hosts and epidemiological features are reviewed and discussed. Twenty-six amphistome species belonging to nine genera from three families occur in domestic and wild ruminants in the region under review and over 70% of them belong to the genera Calicophoron, Carmyerius and Cotylophoron. Of the amphistome species, 76.9% are shared between domestic and wild ruminant hosts – an important observation when considering the different options for control. Seven freshwater snail species belonging to four genera from two families act as intermediate hosts of the identified amphistome species, with the genus Bulinus contributing 57% of the snail species. Some of the snails are intermediate hosts of amphistome species belonging to the same genus or to different genera; a phenomenon not yet fully elucidated as some snails are reported to be naturally infected with amphistome cercariae of unidentified species. Only nine (34.6%, 9/26) of the amphistome species have known snail intermediate hosts, while most (65.4%, 17/26) have unknown hosts. Species of intermediate hosts and the potential of the flukes to infect these hosts, the biological potential of the snail hosts, the definitive hosts management systems and their grazing habits are considered to be the main factors influencing the epidemiology of amphistomosis. Based on the epidemiological features of amphistome infections, various practical control options are discussed. Further research is necessary to determine amphistome–snail associations, develop diagnostic tests that can detect prepatent infections in the definitive host, determine the burden and economic importance of amphistomosis in domestic and wild ruminants and the efficacy of different anthelmintics in the treatment of patent infections.


Amphistomosis is a disease of domestic and wild ruminants caused by digenetic trematodes of the superfamily Paramphistomoidea Fischoeder, 1901 (Lotfy et al. 2010). The superfamily has a cosmopolitan distribution and is composed of hundreds of species belonging to 12 families (Jones 2005). Given their ubiquity and their abundance within hosts, it seems likely that the importance of these flukes is underestimated globally (Lotfy et al. 2010). Various species of the different paramphistomoid families, especially members of Paramphistomidae and Gastrothylacidae, cause amphistomosis among ruminants. The disease is caused by a severe infection with immature flukes in the small intestines of immunologically incompetent hosts. The amphistomes are responsible for lower nutrition conversion and result in a loss of weight and/or a decrease in milk production, causing great economic losses (Horak 1971). However, most reports on the disease do not quote the responsible amphistome species as they are difficult to identify from a systematic point of view (Horak 1971). Calicophoron microbothrium is probably the biggest cause of this disease in Africa (Dinnik 1964a). Knowledge of the different amphistome species infecting domestic and wild ruminants facilitates a better understanding of the amphistome–host associations and the epidemiology of the disease.

A wide range of gastropods belonging to the genera Bulinus Müller 1781, Biomphalaria Preston 1910, Ceratophallus Brown and Mandahl-Barth 1973 and Galba Müller 1774 act as the intermediate hosts of amphistomes in Africa (Dinnik 1961, 1965; Dinnik & Dinnik 1954; Southgate et al. 1989; Wright, Southgate & Howard 1979). The prevalence of snail-borne diseases such as amphistomosis is influenced by both the abundance of infected definitive hosts and the abundance and efficiency of the snail intermediate hosts. Hence, the epidemiology and seasonal patterns of infection with amphistomes is determined to a large extent by the availability of the snail intermediate hosts and the grazing habits of the definitive hosts (Horak 1971; Rolfe et al. 1991). Information on the snail hosts of different amphistome species is essential as knowledge of the amphistome–snail associations has an influence on amphistomosis epidemiology and control.

In this review, to avoid confusion, genera of parasites and snail hosts have been abbreviated using the first three letters of the genus name and these include: for amphistomes – Bilatorchis (Bil.), Calicophoron (Cal.), Carmyerius (Car.), Choerocotyloides (Cho.), Cotylophoron (Cot.), Gastrothylax (Gas.), Gigantocotyle (Gig.), Orthocoelium (Ort.) and Stephanopharynx (Ste.) and for snail hosts – Biomphalaria (Bio.), Bulinus (Bul.) and Ceratophallus (Cer.). The authorities of the digenean families and species and that of the snail species referred to in this review can be found in Table 1.

TABLE 1: Checklist of amphistome species and their ruminant and snail intermediate hosts reported in east and southern African countries.

In this paper, we review the information available to date on amphistome species infecting domestic and wild ruminants in east and southern African countries, the snail intermediate hosts, as well as the epidemiology of amphistomosis and available control options.

Amphistome species infecting ruminants in East and Southern Africa

Reported amphistome species and their respective domestic and wild ruminant hosts in east and southern African countries are shown in Table 1. Data show that the documented species belong to four families: Choerocotyloididae, Gastrothylacidae, Paramphistomidae and Stephanopharyngidae, and nine genera: one from Choerocotyloididae (Choerocotyloides Prudhoe, Yeh & Khalil 1964), two from Gastrothylacidae (Carmyerius Stiles & Goldberger 1910 and Gastrothylax Poirier 1883), five from Paramphistomidae (Bilatorchis Fischoeder, 1901, Calicophoron Näsmark 1937, Cotylophoron Stiles & Goldberger 1910, Gigantocotyle Näsmark 1937 and Orthocoelium [Stiles & Goldberger 1910] Price & McIntosh 1953) and one from Stephanopharyngidae (Stephanopharynx Fischoeder 1901).

Twenty-six species occur in domestic and wild ruminants in the area under review. Seventy-seven per cent of them (20/26) belong to Calicophoron, Carmyerius and Cotylophoron with the genus Calicophoron accounting for approximately 35% of the species, followed by Carmyerius (27%) and Cotylophoron (15%). Seventy-five per cent (9/12) of the known Calicophoron species and more than 40% of Carmyerius (43.8%, 7/16) and Cotylophoron (57.1%, 4/7) species occur in the area under review. However, less than 40% of Gastrothylax (33.3%, 1/3), Gigantocotyle (25%, 1/4) and Orthocoelium (9.1%, 1/11) known species occur in ruminants in east and southern Africa. Most of the Calicophoron species have a wider distribution with respect to countries where reported compared with species of the other genera. Calicophoron microbothrium has the widest distribution followed by Cot. cotylophorum and Cal. raja. Calicophoroncalicophorum, Cal. phillerouxi, Cal. sukari, Cal. fuelleborni and Cal. spatiosus also have a wider distribution compared with the rest of the other species. Seven species; Bil. papillogenitalis, Cal. daubneyi, Car. bubalis, Car. dollfusi, Cho. Onotragi, Cot. macrosphinctris and Ort. scoliocoelium had the narrowest distribution, being reported in only one country each.

The majority of the species (76.9%) are shared between domestic and wild ruminant hosts and approximately 12% of them have not been documented in domestic ruminants as yet (Bil. papillogenitalis, Car. bubalis and Cot. macrosphinctris), while another 12% (3/26) have not been reported in wild ruminants (Cal. daubneyi, Car. dollfusi and Ort. scoliocoelium). Approximately 85% (22/26) are found in domestic and 88% (23/26) in wild ruminants. All the species recorded in domestic ruminants occur in cattle with Cal. daubneyi, Car. dollfusi and Ort. Scoliocoelium documented in this ruminant host only. Sheep are hosts to 55% (12/22), while goats are hosts to 50% of the species reported in domestic ruminants. Half (11/22) of the species have been reported in all the domestic ruminants with most (63.6%, 7/11) of them being Calicophoron species. The range of wild ruminant hosts varies for the different species. Cotylophoron cotylophorum has the highest wild ruminant host range, 19 host species belonging to 10 genera followed by Cal. raja, 18 host species belonging to 12 genera, Cal. microbothrium, 17 host species belonging to 11 genera and Car. spatiosus, 10 host species belonging to 10 genera. Calicophoron sukari has the lowest wild ruminant host range, with only one wild ruminant host, that is, the buffalo. Of the wild ruminant hosts, buffaloes are hosts to 78% (18/23) of the amphistome species recorded in wild ruminants followed by the waterbuck (52.2%, 12/23), the Kafue lechwe (47.8%, 11/23), the roan antelope (47.8%, 11/23) and the hartebeest (43.4%, 10/23). The blue wildebeest, bushbuck, eland, impala, kudu and sable antelope are also hosts to more than 25% of the amphistome species documented in wild ruminant hosts.

Mixed farming systems of cattle and game, particularly antelope, have become an important agricultural activity in most east and southern African countries. In addition, there has been the creation of Transfrontier Conservation Areas (TFCAs) involving many African countries, particularly in Southern Africa, resulting in increased livestock–wildlife interface areas. Therefore, domestic and wild animals are coming into ever more intimate contact in many interface areas, particularly in rural areas at the edges of the TFCAs and in farms practising mixed cattle and game farming, thus promoting the possibility of parasite exchange. These observations are important when considering the different options for their control. For instance, Phiri et al. (2011) observed that the host range of many helminths found in the Kafue lechwe is broad and they could serve as a potentially stable source of infection to domestic animals such as goats and cattle. Hence, issues concerning livestock management and conservation may arise.

Snail intermediate hosts

Table 1 shows the reported intermediate snail hosts of different amphistome species recorded in the study areas under review. Data show that seven snail species – Bio. pfeifferi, Bul. forskalii, Bul. globosus, Bul. nasutus, Bul. tropicus and Cer. natalensis all Planorbidae Rafinesque 1815 and Galba truncatula belonging to Lymnaeidae Rafinesque 1815 – are so far confirmed intermediate hosts of identified amphistome species. The genus Bulinus contributes 57% (4/7) of the confirmed snail intermediate hosts, while the remaining genera contribute one species each. The data also show that some snail species are intermediate hosts of amphistome species belonging to the same genus, for example, Bul. forskalii (Cal. microbothrium and Cal. phillerouxi), Bul. tropicus (Cal. calicophorum and Cal. microbothrium) and Cer. natalensis (Car. exoporus and Car. mancupatus) or amphistome species belonging to different genera, for example, Bul. globosus (Cal. microbothrium and Car. parvipapillatus) and Cer. natalensis (Car. exoporus, Car. mancupatus and Ort. scoliocoelium). The capacity of the various snail species to act as intermediate hosts for paramphistomoids has not been fully elucidated yet, as some of the snail hosts have been reported to be naturally infected with amphistome cercariae of unidentified species (Chingwena et al. 2002a; Dinnik 1961; Loker, Moyo & Gardner 1981; Lotfy et al. 2010; Mukaratirwa et al. 1998; Pfukenyi et al. 2005a; Wright et al. 1979). Adult amphistomes are difficult to identify using their anatomical and morphological features as they have thick robust bodies in which the internal organs are difficult to characterise (Jones 1990). As amphistomes in snail hosts are in their larval stages, species identification is made even more difficult. Hence, the technical difficulties in making precise species identifications impact greatly on the better understanding of amphistome–snail associations. However, Lotfy et al. (2010) demonstrated ITS2 as a good molecular marker for amphistome identification, which can be used to identify both adult amphistome and cercariae to species level.

The current review shows that only nine (34.6%, 9/26) amphistome species (Cal. calicophorum, Cal. daubneyi, Cal. microbothrium, Cal. phillerouxi, Cal. sukari, Car. exoporus, Car. mancupatus, Car. parvipapillatus and Ort. scoliocoelium) have known snail hosts. Except for Cal. microbothrium, presently with four known snail hosts (Bul. forskalii, Bul. globosus, Bul. nasutus and Bul. tropicus) in East and Southern Africa, all the other species have one known snail host each. In addition, Bio. pfeifferi and Melanoides tuberculata are known experimentally to serve as snail hosts of this parasite (Chingwena et al. 2002b). Calicophoron microbothrium is widely distributed in the areas under review and the wide range of its snail hosts probably supports its reported broad geographical distribution.

Data under review show that most of the known amphistome species have unknown snail hosts (65.4%, 17/26). The snail hosts of four Calicophoron species (Cal.bothriophoron, Cal. clavula, Cal. raja and Cal. sukumum) are currently not known. In Tanzania, amphistome cercariae of unidentified Calicophoron species were recorded in Bul. forskalii (Lotfy et al. 2010). Lotfy et al. (2010) suggested that besides Cal. phillerouxi, which is known from Bul. forskalii, two other Calicophoron species known in Tanzania, Cal. bothriophoron and Cal. sukumum with unknown snail hosts, cannot be ruled out. In the East African region, Bul. abyssinicus is reported as the intermediate host of Cal. clavula in Somalia (Sobrero 1962). Dinnik and Hammond (1968) suggested Bul. globosus as a likely snail host of Cal. raja as it is experimentally proven to be susceptible to infection. Four Carmyerius species with presently unknown snail hosts are Car. bubalis, Car. dollfusi, Car. gregarious and Car. spatiosus. Amphistome cercariae belonging to an unidentified species of the family Gastrothylacidae were recorded from Cer. natalensis in Kenya (Lotfy et al. 2010). The genus Carmyerius is one of four genera belonging to the family Gastrothylacidae. Hence, besides Car. Mancupatus and Car. exoporus already known from this snail host, one other Carmyerius species known in Kenya, Car. spatiosus with unknown snail hosts, cannot be ruled out (Lotfy et al. 2010). Besides Cer. natalensis, Wright et al. (1979) also suggested Bul. forskalii as likely snail hosts of Car. spatiosus in Zambia. To date, all four Cotylophoron species have unknown snail hosts. The other species with unknown snail hosts are Bil. papillogenitalis, Cho. onotragi, Gas. crumenifer, Gig. symmeri and Ste. compactus. However, even though not yet confirmed, Bul. forskalii has been speculated to act as the intermediate host of Ste. compactus (Dinnik 1965).

Epidemiological features of amphistome infections in ruminants in East and Southern Africa

The epidemiology and prevalence of amphistomosis depend on several factors. These include the species of definitive and intermediate hosts (Rolfe et al. 1991), the potential of the flukes to infect these hosts (Dinnik 1964a; Dinnik & Dinnik 1954; Horak 1967), the topography and biological potential of the snail hosts (Dinnik 1964a; Horak 1971; Rolfe et al. 1991; Swart & Reinecke 1962a, 1962b), the definitive hosts’ management systems and their grazing habits as well as climate (Rolfe et al. 1991).

Data on amphistome infection prevalence are scarce for the reviewed countries and are currently only available from six countries (Table 2). The prevalence data are based on coprology and fluke counts with most studies having conducted in cattle. Because of difficulties in amphistome species identification, specific species prevalence data are lacking. Prevalence studies are limited for goats, sheep and wild ruminants. The available data show a high prevalence in the Kafue lechwe, but low prevalence rate in goats and sheep. In cattle, the coprological prevalence varies from 23.7% to 86.5%, while it varies from 25.5% to 96% on fluke counts, the high prevalence perhaps being explained by the fact that amphistome infection in ruminants is commonly because of several species. In the highlands of Kenya, Cal. microbothrium, Cal. daubneyi and Cal. jacksoni were recovered from a single animal and in few cases Cal. sukari, Car. exoporus and Car. mancupatus were present as well (Dinnik 1964a). Another combination of six species (Cal. microbothrium, Cal. phillerouxi, Cal. raja, Cot. cotylophorum, Car. parvipapillatus and Ste. compactus) was found in an ox in Zambia (Dinnik 1964a). Amphistomes recovered in slaughtered cattle were a combination of Cal. microbothrium and Cot. jacksoni in Tanzania (Keyyu et al. 2006). Infections with different amphistome species are also reported in Zimbabwe in cattle (Dube et al. 2004; Dube & Tizauone 2014) and in sheep and goats (Dube, Masanganise & Dube 2010). In addition, most amphistome species (85%) are shared between domestic and wild ruminant hosts (Table 1), providing a potentially stable source of infection among the ruminant animals. Furthermore, the availability of a wide range of the snail hosts (Table 1) with high biological potential also increases the successful propagation of amphistomes in the environment leading to increased infection exposure in ruminants. Limited routine anthelmintic treatment, particularly in rural communities who practice communal grazing, and the lack of effective drugs against amphistomes are also possible explanations for the high prevalence in domestic ruminants. An increase in the prevalence of amphistome infections has been reported in Western Europe (Foster et al. 2008; Mage et al. 2002; Murphy et al. 2008; Toolan et al. 2015). Besides an improvement in quality of diagnosis, the increase has also been attributed to the absence of an effective anthelmintic against amphistome infections (Mage et al. 2002).

TABLE 2: Prevalence of amphistomes in ruminants in east and southern African countries based on faecal egg and fluke counts.

Studies on animal-breed predisposition to amphistome infection are limited. Indigenous cattle breeds were observed to have a significantly higher prevalence and intensity than the exotic breeds and crosses in Kenya and Uganda (Howell 2011; Kanyari, Kagira & Mhoma 2010). In Tanzania, the Maasai Zebu cattle had a significantly higher prevalence than the Iringa Red cattle; however, the numbers of animals involved were too small for any meaningful interpretations to be made (Nzalawahe et al. 2015). Literature reports a variable effect of sex in domestic ruminants. Despite females tending to record higher prevalences than males, the associations were not significant (Kanyari et al. 2009, 2010; Keyyu et al. 2006; Phiri, Chota & Phiri 2007a; Phiri, Phiri & Monrad 2006). However, Pfukenyi et al. (2005a) observed significantly higher prevalences in pregnant and lactating cows compared with bulls, oxen and dry cows. Similarly, Howell (2011) reported a significantly higher prevalence in female cattle compared with males. The differences between sexes could probably be related to grazing patterns, sex hormones and treatment regimes.

Adult domestic ruminants are reported to have a significantly higher prevalence compared with young animals (Howell 2011; Kanyari et al. 2009, 2010; Keyyu et al. 2005, 2006; Nzalawahe et al. 2014; Pfukenyi et al. 2005a; Phiri et al. 2007a; Vassilev 1999). This is attributed to a long exposure time in adults leading to immunity against the pathogenic effects of immature amphistomes but still having the mature ones maintaining their high egg production capacity (Horak 1971). The resistance to amphistome re-infection in cattle was demonstrated clinically (Horak 1967) with no simultaneous studies on the cellular effector systems that characterise the acquired resistance. Mavenyengwa et al. (2008) showed that the resistance to Cal. microbothrium re-infection in cattle involves eosinophils and mast cells that are targeted at immature flukes. Epidemiologically, adults act as a constant source of infection, but clinical amphistomosis remains a problem for young animals, with adults grazing the same pastures exhibiting no clinical effects despite continued egg production (Boray 1959, 1969; Butler & Yeoman 1962; Rolfe et al. 1991). Amphistomes may survive for up to two years in the definitive hosts, providing a virtually constant source of infection for successive generations of snail hosts. In rural areas, the grazing management of young and adult animals may differ, where young animals graze around farms or homesteads while adults are trekked long distances to valleys, flood plains or swampy areas where they are exposed to high metacercariae-contaminated pastures (Keyyu et al. 2005, 2006).

Animal grazing area and/or habitat is significantly associated with prevalence and intensity of amphistomes in domestic ruminants (Howell 2011; Kanyari et al. 2009, 2010; Keyyu et al. 2005, 2006; Nzalawahe et al. 2014, 2015; Pfukenyi et al. 2005a; Phiri et al. 2006). The prevalence is highest in animals grazing in areas characterised by wetlands or swampy or marshy grazing areas where the distribution of suitable snail habitats is widespread. For instance, the highveld region in Zimbabwe, characterised by wet/swampy grazing areas where distribution of snail habitats is widespread, is associated with a higher prevalence compared with the lowveld which is characterised by dry land grazing with a focal distribution of snail habitats (Pfukenyi et al. 2005a, 2005b). Similarly, the presence of wetlands and high livestock density in the cattle grazing areas of the Zambian western and southern provinces is associated with an increased risk of acquiring amphistome infections (Phiri et al. 2006) and the same observations have been reported in Kenya (Kanyari et al. 2009, 2010), Tanzania (Keyyu et al. 2005, 2006; Nzalawahe et al. 2015) and Uganda (Howell 2011). In Tanzania, traditional communal grazing areas exhibited the highest prevalence of amphistomes compared with other sectors and this is attributed to heavy contamination of the habitats with eggs where intermediate host snails breed, because of high stocking densities with subsequent heavy metacercarial density on vegetation grazed by the animals especially during the dry season (Keyyu et al. 2005, 2006). Villages practising irrigation of crops are associated with high amphistome infection rates in Tanzania (Nzalawahe et al. 2014) as this provides favourable ecological conditions for growth of snail hosts and development of trematode larval stages.

The prevalence in domestic ruminants as measured by coprology follows a seasonal pattern with an increase towards the end of the dry season and a peak during the wet months of the year (Keyyu et al. 2005; Pfukenyi et al. 2005a; Phiri et al. 2007a; Reinecke 1983; Vatta & Krecek 2002). Outbreaks of acute clinical amphistomosis because of immature flukes are usually confined to the drier months of the year (Boray 1969; Butler & Yeoman 1962; Dinnik 1964a; Horak 1967, 1971; Rolfe et al. 1991; Vassilev 1999). Towards the end of the rainy season and onset of the dry season, conditions in permanent water sources become favourable for an increase in the number of the snail intermediate hosts, reaching their peak during the mid-to-end of the dry season (Chingwena et al. 2002a; Pfukenyi et al. 2005a, 2005b; Phiri et al. 2007b). As the snail hosts are extremely adaptable and prolific breeders, this ensures their widespread availability as well as heavy shedding of cercariae which encyst on vegetation surrounding the habitats (Dinnik 1964a). The proportion of infected snails increases from the end of the rainy season into the dry season (Chingwena et al. 2002a; Pfukenyi et al. 2005a). A combination of high snail numbers, asexual multiplication of the fluke in infected snails and survival of snails in suitable environments for several months may result in shedding of large numbers of cercariae. During this period, the infective metacercariae are spread over pastures surrounding permanent water sources where they can survive for several months. This coincides with pasture areas being narrowed around permanent water sources or wetland environments where animal concentration becomes high (Pfukenyi et al. 2005b). Contamination rates of these areas are increased, resulting in more snails being infected and high numbers of metacercariae on surrounding herbage, leading in turn to acute infections of animals with amphistomes. Thus, a build-up of immature flukes occurs, accounting for clinical amphistomosis outbreaks and low amphistome prevalence as measured by coprology. The outbreaks are common in ruminants that graze in marshy or swampy areas and are usually confined to the dry season. However, on irrigated pastures, moisture is often adequate for the survival of snail hosts and metacercariae, and hence outbreaks can occur throughout the year. In Tanzania, some villages practising year-round zero-grazing had high levels of amphistome infections attributed to the acquisition of cattle fodder from irrigation canals and swamps contaminated with metacercariae (Nzalawahe et al. 2014).

Development of amphistomes into adults takes 5–9 months (Dinnik & Dinnik 1962) and the prepatent period is 56–89 days (Dinnik & Dinnik 1962; Horak 1971). Five to 9 months after infection, the immature flukes become fully mature and this would lead to high faecal egg production and thus, account for the high prevalence during the rainy season as measured by coprology. During this period, abundant grazing and alternative water sources are available. Hence, drinking from and grazing around infected permanent water sources is greatly reduced. Furthermore, snail habitats and pastures are constantly flooded, and thus snails and the parasitic free-living stages are regularly flushed (Pfukenyi et al. 2005b). In summary, the intermediate and definitive hosts acquire most of the infection during the beginning and/or middle of the dry season. This results in immature fluke infections and clinical amphistomosis during the dry season and patent (mature fluke) infections during the wet months and at the end of the dry season. However, the timing may vary depending on location, length of the rainy season and the grazing habits of the animals (Pfukenyi et al. 2005b).

Impact on production

Adult flukes are not associated with clinical amphistomosis (Mavenyengwa, Mukaratirwa & Monrad 2010). However, in heavy infections they have been hypothesised to cause weakness, recurrent ruminal tympany, ruminal atony, weight loss, anaemia and production losses (Anuracpreeda, Wanichanon & Sobhon 2008). They are also reported to be associated with inflammation of the mucosa and mucoid diarrhoea (Rolfe & Boray 1993). Based on coprology, poor body condition is reported to be significantly associated with high amphistome prevalence in cattle (Kanyari et al. 2010). A similar observation was noted in small ruminants (Kanyari et al. 2009), but the association was not significant. Cattle infected with more than 500 adult amphistomes had a significant reduction in final carcass mass when compared with controls (Marchand 1984; Dube & Tizauone 2014). The concurrent infection of amphistomes with other parasites known to depress growth rate such as strongyles (Kanyari et al. 2010), Fasciola species (Kanyari et al. 2010; Keyyu et al. 2006; Nzalawahe et al. 2014; Phiri et al. 2006; Yabe et al. 2008) and Moniezia species (Kanyari et al. 2010) is a likely explanation of the significant association between poor body condition and amphistome infections. However, further studies on the effect of adult amphistomes on production are required.

Clinical amphistomosis is caused by the immature flukes that lodge in the first 3 m of the small intestine (Mavenyengwa et al. 2010). The occurrence of clinical amphistomosis and subsequent clinical pathology in ruminants is dependent on the dose, pathogenicity of the species and the level of establishment of the metacercariae in the host’s small intestine (Horak 1967; Mavenyengwa et al. 2010). In ruminants, the disease is characterised by anorexia, anaemia, submandibular oedema, and hypoproteinemia, foul-smelling fetid diarrhoea, general weakness, polydipsia, and a reduction in feed conversion, weight and milk production and mortality in young animals (Boray 1969; Horak 1966; Mavenyengwa et al. 2010; Mohan 2011; Pillai & Alikutty 1995; Rolfe, Boray & Collins 1994; Spencer, Fraser & Chang 1996). Together with gastrointestinal nematodes, amphistome infection in cows can reduce milk production by approximately 0.4 L/day – 3 L/day (Mohan 2011; Spencer et al. 1996). Anthelmintic treatment of dairy cows infected with gastrointestinal nematodes (oxfendazole) and amphistomes (oxyclozanide) resulted in a significant increase in milk production, averaging 0.4 L/day (Spencer et al. 1996). The reduction in milk yield during clinical amphistomosis is associated with fetid diarrhoea (Mohan 2011). Mohan (2011) also reported anoestrus during clinical amphistomosis, while a functional obstruction or paralytic ileus of the intestine because of severe amphistomosis was reported in a cow (Yogeshpriya et al. 2011). Despite their ubiquity and abundance, as well as an increase in their prevalence in domestic ruminants, the economic importance of amphistome infections is not yet fully known and is likely to be underestimated in eastern and southern Africa – an area which requires further studies.


Diagnosis of amphistomes in live animals is still dependent on faecal detection of eggs (Rieu et al. 2007) and this method only detects the presence of adult rumen fluke infection (Malrait et al. 2015). The filtration technique with sieves and sedimentation is the most accurate method to identify eggs in faeces (Horak 1971). Using contrast stains such as methylene blue or methyl green to distinguish amphistome eggs from Fasciola ova is advisable. One drawback of this diagnostic method is that, in acute infections, it is highly probable not to find eggs or only very few as this is usually associated with massive infection with immature flukes (Horak 1971). The agreement between a modified McMaster method and necroscopic diagnosis of amphistome infection is reported to be high (Rieu et al. 2007) with no significant differences being observed between the two methods. The modified McMaster method showed a significant association between eggs per gram (epg) counts and parasite burden; more than 100 epg indicated the presence of more than 100 adult amphistomes in the rumen and/or reticulum (Rieu et al. 2007). Similarly, the mini-FLOTAC is a reliable method of assessing the presence of adult amphistome infection with both sensitivity and specificity being above 0.9 (Malrait et al. 2015). A good correlation was found between faecal egg count (FEC) and estimated rumen fluke burden with a FEC > 200, indicating the presence of more than 200 adult rumen flukes in the rumen and/or reticulum (Malrait et al. 2015). The adult worms are difficult to identify to species level because most have thick robust bodies in which the internal organs are difficult to see. Even by using histological techniques, species identification is still problematic (Lotfy et al. 2010). As the flukes responsible for disease are sexually immature, specific identification is made even more difficult and the diagnosis has to rely on the dubious procedure of identifying a few adult worms, which may be present in the rumen of the animal (Horak 1971). Because of these problems, PCR-based techniques providing rDNA ITS2 sequences have proven to be reliable tools to identify amphistome species and to determine their phylogenetic relationships (Itagaki et al. 2003; Rinaldi et al. 2005). Using cercariae and rediae from snail hosts and adult flukes obtained from slaughterhouses, Lotfy et al. (2010) confirmed ITS2 as a good molecular marker for amphistome identification that can also be used to determine phylogenetic and amphistome–snail associations.

The clinical diagnosis of amphistomosis remains challenging as immunological techniques are usually not conclusive (Horak 1967, 1971). Croposcopic examination cannot be used for the early diagnosis of clinical amphistomosis which is vital for prompt treatment before considerable damages and economic losses are incurred. For the identification of immature flukes, the recommended method is to mix approximately 10 g of faeces with 100 mL – 200 mL of water (Horak 1971). The mixture is allowed to stand for 5 min, followed by decanting any supernatant fluid and then repeating the procedure four to five times. Young flukes, resembling small white or pink rice grains, will be seen after pouring the sediment on a black surface for examination (Horak 1971). In dead animals, postmortem, pathological and clinical pathological findings combined with the presence of immature flukes in the affected intestines would be confirmative. The gross pathological, histopathological and clinical pathological findings are as described in the literature (Horak 1966, 1967, 1971; Horak & Clarke 1963; Mavenyengwa et al. 2005, 2008, 2010; Pillai & Alikutty 1995). An indirect ELISA performed to detect coproantigens in faecal supernatants of 100 cattle known to be infected with Gas. crumenifer had a sensitivity of 74% (Kandasamy & Devada 2011). Generally, the sensitivity of the indirect ELISA ranges from 74% to 86% and its specificity from 79% to 90% (Hassan et al. 2005; Kandasamy & Devada 2011; Salib et al. 2015; Sanchis et al. 2012; Shivjot et al. 2009). However, Shivjot et al. (2009) reported a very low specificity of 23.7%. The ELISA was shown to be more specific and accurate but less sensitive than Western blotting for the diagnosis of amphistome infections in cattle and buffaloes (Salib et al. 2015). Results indicate the feasibility of ELISA for the detection of coproantigens of amphistome infections, especially for the diagnosis of immature amphistomosis where faecal examination may not reveal eggs.


The available epidemiological information on amphistomes of ruminants in the area under review can be used to design appropriate integrated control measures. Options available for the control of amphistome infections are mainly based on chemical treatment, non-chemical management practices and immunological control.

Chemical treatment

Chemical control involves treatment with a product that is effective against both adult and immature flukes. Oxyclozanide given twice, three days apart, has a high efficacy against both adult and juvenile amphistomes (Rolfe & Boray 1987) and a high anthelmintic performance in cattle (Arias et al. 2013; Rolfe & Boray 1987; Spencer et al. 1996) and small ruminants (Paraud et al. 2009; Rolfe & Boray 1988; Sanabria et al. 2014). Studies in Tanzania showed a reduced efficacy of levamisole–oxyclozanide combination against amphistomes in cattle (Keyyu et al. 2008) and this is of great concern as they are the commonly available drugs in the country. However, levamisole is widely used to treat nematode infections in livestock and it is not intended as treatment against trematodes. When given orally at a higher dosage (10 mg/kg), closantel has a high efficacy against mature flukes (Arias et al. 2013). However, treatment of mature flukes with intra-ruminally (Rolfe & Boray 1993) or subcutaneously administered (Malrait et al. 2015) closantel is not effective. In countries where oxyclozanide is unavailable, the use of closantel to treat against mature flukes is recommended. When administered at high doses (50 mg/kg and 100 mg/kg), niclosamide has 94% – 99% efficacy against immature amphistomes (Rolfe & Boray 1987).

Even though it is not of direct benefit to the animal, treatment against mature amphistomes will prevent egg laying and thus reduce pasture contamination (Horak 1971), while treatment against the immature flukes will reduce the impact of the disease. During the rainy season, mature amphistomes are expected and anthelmintic treatment with drugs effective against adult flukes is indicated. The strategic anthelmintic treatment against mature amphistomes should be given in adult animals at the end of the rainy season (Pfukenyi et al. 2005a, 2005b) or beginning of the dry season (Keyyu et al. 2005) to reduce the opportunity for snail infections. The timing of this treatment is dependent on local factors, length of the rainy season and the grazing habits of the animals. Where possible, adult animals targeted for treatment should have high levels of infection based on coprology. Depending on availability, oxyclozanide or closantel can be administered during this period to treat against mature amphistomes.

Disease epidemiology indicates that large burdens of immature amphistomes are expected during the dry season. As adult animals are resistant to the pathogenic effects of the migrating immature amphistomes, the target for treatment would be young animals being exposed to the infection for the first time (Pfukenyi et al. 2005a). Hence, the first anthelmintic treatment can be administered in young animals during the mid-dry season period when maximum migration of immature amphistomes starting 3–4 weeks after infection in the early dry season is expected. To remove potentially high burdens of immature amphistomes acquired later in the dry season, a second treatment could be given towards the end of the dry season (Pfukenyi et al. 2005a). Oxyclozanide or niclosamide can be administered during this period to treat against immature amphistomes. In communal areas, animals are communally grazed and for optimum benefits, the recommended anthelmintic treatments should be well organised and preferably done at the same time within a village. Where cattle are dipped for the control of ticks, dip tank facilities where all animals are gathered during dipping sessions could be used for organised fluke control (Pfukenyi et al. 2005a, 2005b).

The efficacy of medicinal plant extracts against amphistomes has recently been evaluated. The ethanol extract of Punica granatum L. (Lythraceae), commonly known as pomegranate, is highly effective against amphistomes in naturally infected sheep (Lalhmingchhuanmawii, Veerakumari & Raman 2014). The authors concluded that the plant extract could be successfully used as an anthelmintic to treat amphistomes in domestic ruminants. Similarly, an aqueous extract of Acacia concinna (Willd.) DC. (Fabaceae) significantly reduced egg counts of amphistomes in naturally infected sheep and also restored the haemato-biochemical profile to normal in extract-treated sheep (Priya, Veerakumari & Raman 2013). However, efficacy of the P. granatum and A. concinna extracts was not established in immature amphistomes. Other studies have also shown medicinal plants extracts to be effective against amphistomes (Elango & Rahuman 2011; Kamaraj et al. 2010).

Chemical control of the snail hosts through application of molluscicides such as niclosamide may also be done. To achieve cost-effective control, this type of control should be done during the peak transmission period to reduce numbers of infected snails and cercarial shedding. Thus, the application could be done during the mid-dry and towards the end of the dry season (Pfukenyi et al. 2005b). The application is practical and economical in areas where snail habitats are focal and not widespread, but regular application may be necessary because of the rapid recovery of the snail populations during brief periods of favourable conditions. However, molluscicide application causes environmental pollution and also kills non-targeted aquatic organisms (Roberts & Suhardono 1996).

Immunological control

Hafeez and Rao (1981) showed that the lifespan and pathogenicity of amphistomes developing from gamma irradiated (2 or 3 krad) metacercariae were greatly reduced with the higher irradiation dose resulting in the complete absence of the flukes in infected animals. Single vaccination of kids and lambs with 3000 irradiated (2 or 3 krad) metacercariae stimulated a significant degree of resistance against challenge and the resistance was more pronounced in the group vaccinated with a higher irradiation dose (Hafeez & Rao 1981). Earlier, Horak (1967) successfully immunied sheep, goats and cattle against massive artificial infections with Cal. microbothrium. The animals were given immunising infections with at least 40 000 metacercariae and later challenged with larger doses of metacercariae (Horak 1967). Cattle were the most suitable subjects for immunisation with the immunity being effective for at least a year post-immunisation (Horak 1967, 1971). Mavenyengwa et al. (2008) demonstrated that cattle acquire resistance to amphistome infection. This resistance is targeted at immature amphistomes and it involves eosinophils and mast cells. However, despite promising immunisation results, the mass production of snail hosts and metacercariae remains a challenge and a major limiting factor (Horak 1967, 1971; Mavenyengwa et al. 2006; Swart & Reinecke 1962a, 1962b). Thus, the success of a large-scale immunisation program is dependent on a viable metacercariae mass production system.

Non-chemical control

The best preventive method against amphistome infections is to keep domestic ruminants from infected pastures (Pfukenyi et al. 2005b). Fencing-off or drainage of wetlands or marshy/swampy areas and provision of clean pastures and cercariae-free water in troughs are advised (Roberts & Suhardono 1996). Similarly, habitat management through vegetation clearance is also effective in controlling the snails (Woolhouse & Chandiwana 1990). However, habitat management and complete separation of stock from snail habitats are only practical and economical where the snail habitats are focal and not widespread (Pfukenyi et al. 2005b). These control methods are not feasible in communal grazing areas. It is also important to repair any leaks in dams and water troughs as they can create an ideal habitat for the snail hosts.


Twenty-six amphistome species belonging to nine genera from three families occur in domestic and wild ruminants in the area under review and seven snail species belonging to four genera from two families act as their intermediate hosts. Eighty-five per cent of the amphistome species are shared between domestic and wild ruminant hosts. Some snails are intermediate hosts of amphistome species belonging to the same genus or to different genera – a phenomenon not yet fully elucidated. Only nine (34.6%) of the amphistome species have known snail intermediate hosts, while most (65.4%) have unknown hosts. The epidemiology of amphistomosis depends on the species of definitive and intermediate hosts and the potential of the flukes to infect these hosts, the topography and biological potential of the snail hosts, the management systems of the definitive host and their grazing habits and climatic factors. Based on current epidemiological information, the strategic anthelmintic treatment against mature amphistomes should be given in adult animals at the end of the rainy or early dry season. The anthelmintic treatment in young animals against immature amphistomes should be administered during the mid-dry and towards the end of the dry season. Further research is necessary to determine the economic importance of amphistomosis, amphistome–snail associations, efficacy of different anthelmintics and to develop diagnostic tests that can detect prepatent infections in the definitive host.


The authors would like to thank the anonymous reviewers whose suggestions improved the manuscript.

Competing interests

The authors declare that they have no financial or personal relationships that may have inappropriately influenced them in writing this article.

Authors’ contributions

Both authors contributed equally in the writing of this manuscript.


Anderson, I.G., 1983, ‘The prevalence of helminths in impala, Aepyceros melampus (Lichteinstein, 1812) under game ranching conditions’, South African Journal of Wildlife Research 13, 55–70.

Anuracpreeda, P., Wanichanon, C. & Sobhon, P., 2008, ‘Paramphistomum cervi: Antigenic profile of adults as recognized by infected cattle sera’, Experimental Parasitology 118, 203–207.

Arias, M., Sanchis, J., Francisco, I., Francisco, R., Pineiro, P., Cazapal-Monteiro, C. et al., 2013, ‘The efficacy of four anthelmintics against Calicophoron daubneyi in naturally infected dairy cattle’, Veterinary Parasitology 197, 126–129.

Boray, J.C., 1959, ‘Studies on intestinal amphistomosis in cattle’, Australian Veterinary Journal 35, 282–287.

Boray, J.C., 1969, ‘Studies on intestinal amphistomosis in sheep due to Paramphistomum ichikawai, Fukui 1922’, Revue de Medecine Veterinaire 4, 290–308.

Butler, R.W. & Yeoman, G.H., 1962, ‘Acute intestinal paramphistomiasis in Zebu cattle in Tanganyika’, Veterinary Record 74, 227–231.

Bwangamoi, O., 1968, ‘Helminth parasites of domestic and wild animals in Uganda’, Bulletin of Epizootic Diseases in Africa 16, 429–454.

Caeiro, V.M.P., 1961, ‘Acerca de alguns “Paramphistominae” nao assinalados em territorios Portugueses’, Revista Ciȇncias Veterinárias Lisbon 56, 68–106.

Chingwena, G., Mukaratirwa, S., Kristensen, T.K. & Chimbari, M., 2002a, ‘Larval trematode infections in freshwater snails from the highveld and lowveld areas of Zimbabwe’, Journal of Helminthology 6, 283–293.

Chingwena, G., Mukaratirwa, S., Kristensen, T.K. & Chimbari, M., 2002b, ‘Susceptibility of freshwater snails to the amphistome Calicophoron microbothrium and the influence of the species on susceptibility of Bulinus tropicus to Schistosoma haematobium and Schistosoma mattheei infections’, Journal of Parasitology 88, 880–883.[0880:SOFSTT]2.0.CO;2

Cruz e Silva, J.A., 1971, ‘Contribuição para o estudo das helmintoses das espécies pecuárias do Sul do Save’, Veterinária Moçambicana Lourenço Marques 4, 33–42.

Dinnik, J.A., 1951, ‘An intermediate host of the common stomach fluke, Paramphistomum cervi (Schrank), in Kenya’, East Africa Agricultural Journal 16, 124–125.

Dinnik, J.A., 1954, ‘Paramphistomum sukari n.sp. from Kenya cattle and its intermediate host’, Parasitology 44, 414–421.

Dinnik, J.A., 1956, ‘On Ceylonocotyle scoliocoelium (Fischoeder, 1904) and its intermediate host in Kenya, East Africa’, Journal of Helminthology 30, 149–156.

Dinnik, J.A., 1961, ‘Paramphistomum phillerouxi sp. nov. (Trematoda: Paramphistomidae) and its development in Bulinus forskalii’, Journal of Helminthology 35, 69–90.

Dinnik, J.A., 1962, ‘Paramphistomum daubneyi sp. nov. from cattle and its snail host in the Kenya highlands’, Parasitology 52, 143–151.

Dinnik, J.A., 1964a, ‘Intestinal paramphistomiasis and P. microbothrium Fischoeder in Africa’, Bulletin of Epizootic Diseases in Africa 12, 439–454.

Dinnik, J.A., 1964b, ‘Paramphistomum sukumum sp. nov. from cattle in the Sukumaland area of the lake Region, Tanganyika’, Parasitology 54, 201–209.

Dinnik, J.A., 1965, ‘The snail hosts of certain Paramphistomidae and Gastrothylacidae (Trematode) discovered by the late Dr. P. L. LeRoux in Africa’, Journal of Helminthology 39, 141–150.

Dinnik, J.A. & Dinnik, N.N., 1954, ‘The life cycle of Paramphistomum microbothrium Fischoeder, 1901 (Trematoda: Paramphistomidae)’, Parasitology 44, 285–299.

Dinnik, J.A. & Dinnik, N.N., 1955, ‘Stomach flukes (Trematoda: Paramphistomidae) found in cattle, sheep and goats in the Highlands of Kenya’, Annual Report of the East African Veterinary Research Organization 1954/55, 84.

Dinnik, J.A. & Dinnik, N.N., 1957, ‘Development of Paramphistomum sukari Dinnik, 1954 (Trematoda: Paramphistomidae) in a snail host’, Parasitology 47, 209–216.

Dinnik, J.A. & Dinnik, N.N., 1960, ‘Development of Carmyerius exoporus Maplestone (Trematoda: Gastrothylacidae) in a snail host’, Parasitology 50, 469–480.

Dinnik, J.A. & Dinnik, N.N., 1962, ‘The growth of Paramphistomum microbothrium Fischoeder to maturity and its longevity in cattle’, Bulletin of Epizootic Diseases in Africa 10, 27–31.

Dinnik, J.A. & Hammond, J.A., 1968, ‘Division of helminth diseases’, Annual Report of the East African Veterinary Research Organization 1967, 47.

Dinnik, J.A., Walker, J.B., Barnett, S.F. & Brocklesby, D.W., 1962, ‘Some parasites obtained from game animals in western Uganda’, Bulletin of Epizootic Diseases in Africa 11, 37–44.

Dube, S., Dlamini, N.R., Masanganise, K.E. & Dube, C., 2004, ‘Abattoir studies on paramphistomes recovered from cattle in Masvingo and Manicaland Provinces of Zimbabwe’, Folia Veterinaria 48, 123–129.

Dube, S., Masanganise, K.E. & Dube, C., 2010, ‘Studies on paramphistomes infecting goats and sheep from Gwanda District in Zimbabwe’, Zimbabwe Journal of Science & Technology 5, 55–64.

Dube, S., Sibula, M.S. & Dhlamini, Z., 2015, ‘Molecular analysis of selected paramphistomes isolates from cattle in southern Africa’, Journal of Helminthology 90, 784–788.

Dube, S., Siwela, A.H., Masanganise, K.E. & Dube, C., 2002, ‘Prevalence of paramphistomes in Mashonaland West, Central and East, and Midlands Provinces, Zimbabwe’, Acta Zoologica Taiwanica 13, 39–52.

Dube, S. & Tizauone, M., 2014, ‘Paramphistomes in Matebeleland South Province Zimbabwe and their effect on aspects of blood plasma composition in infected cattle’, IOSR Journal of Agriculture & Veterinary Science 7, 133–138.

Eduardo, S.L., 1980, ‘Bilatorchis papillogenitalis n.g., n.sp. (Paramphistomidae: Orthocoeliinae), a parasite of the red leche (Kobus leche Gray, 1850) from Zambia’, Systematic Parasitology 1, 203–210.

Eduardo, S.L., 1983, ‘The taxonomy of the family Paramphistomidae Fischoeder, 1901 with special reference to the morphology of species occurring in ruminants. III. Revision of the genus Calicophoron Nasmark, 1937’, Systematic Parasitology 5, 25–79.

Eduardo, S.L., 1984, ‘The taxonomy of the family Paramphistomidae Fischoeder, 1901 with special reference to the morphology of species occurring in ruminants. IV. Revision of the genus Gigantocotyle Nasmark, 1937 and the elevation of the subgenus Explanatum Fukui, 1929 to full generic status’, Systematic Parasitology 6, 3–32.

Eduardo, S.L., 1985a, ‘The taxonomy of the family Paramphistomidae Fischoeder, 1901 with special reference to the morphology of species occurring in ruminants. V. Revision of the genus Cotylophoron Stiles et Goldberger, 1910’, Systematic Parasitology 7, 3–26.

Eduardo, S.L., 1985b, ‘The taxonomy of the family Paramphistomidae Fischoeder, 1901 with special reference to the morphology of species occurring in ruminants. VI. Revision of the genus Orthocoelium (Stiles and Goldberger, 1910) Price & McIntosh, 1935’, Systematic Parasitology 7, 125–158.

Eduardo, S.L., 1986, ‘The taxonomy of the family Paramphistomidae Fischoeder, 1901 with special reference to the morphology of species occurring in ruminants. VIII. The genera Stephanopharynx Fischoeder, 1901, and Balanorchis Fischoeder, 1901’, Systematic Parasitology 8, 57–69.

Eduardo, S.L., 1987, ‘Zoogeographical affinities of paramphistomids of ruminants’, Transactions of the National Academy of Science & Technology (Phils.) 9, 229–242.

Elango, G. & Rahuman, A.A., 2011, ‘Evaluation of medicinal plants extracts against ticks and fluke’, Parasitology Research 108, 513–519.

Fitzsimmons, W.M., 1964, ‘A host check of helminth parasites from domestic animals in Nyasaland’, British Veterinary Journal 120, 186–190.

Fitzsimmons, W.M., 1971, Report on the helminthiasis research project, Lesotho, 1967–1970, Agricultural Information Service, Maseru, Lesotho.

Foster, A.P., Otter, A., O’Sullivan, T., Cranwell, M.P., Twomey, D.F., Millar, M.F. et al., 2008, ‘Rumen fluke (paramphistomosis) in British cattle’, Veterinary Record 162, 528.

Hafeez, Md. & Rao, B.V., 1981, ‘Studies on amphistomiasis in Andhra Pradesh (India) VI. Immunization of lambs and kids with gamma irradiated metacercariae of Cercariae indicae XXVI’, Journal of Helminthology 55, 29–32.

Hassan, S.S., Kaur, K., Joshi, K. & Juyal, P.D., 2005, ‘Epidemiology of paramphistomosis in domestic ruminants in different districts of Punjab and other adjoining areas’, Journal of Veterinary Parasitology 19, 43–46.

Horak, I.G., 1966, ‘Studies on paramphistomiasis. VIII. The pathogenesis and symptoms of the disease in sheep’, Journal of the South African Veterinary Association 37, 428–430.

Horak, I.G., 1967, ‘Host-parasite relationships of Paramphistomum microbothrium Fischoeder, 1901, in experimentally infected ruminants, with particular reference to sheep’, Onderstepoort Journal of Veterinary Research 34, 451–540.

Horak, I.G., 1971, ‘Paramphistomiasis in domestic ruminants’, Advances in Parasitology 9, 33–72.

Horak, I.G. & Clarke, R., 1963, ‘Studies on paramphistomiasis. V. The pathological physiology of the acute disease in sheep’, Onderstepoort Journal of Veterinary Research 30, 45–160.

Howell, A., 2011, ‘Snail-borne diseases in bovids at high and low altitude in eastern Uganda: Integratedparasitological and malacological mapping’, MSc dissertation, Liverpool School of Tropical Medicine.

Itagaki, T., Tsumagari, N., Tsutsumi, K. & Chinone, S., 2003, ‘Discrimination of three amphistome species by PCR-RFLP based on rDNA ITS2 markers’, Journal of Veterinary Medical Sciences 65, 931–933.

Jones, A., 1990, ‘Techniques for hand-sectioning thick- bodied Platyhelminths’, Systematic Parasitology 15, 211–218.

Jones, A., 2005, ‘Superfamily Paramphistomoidea Fischoeder, 1901’, in A. Jones, R.A. Boray & D.I. Gibson (eds.), Keys to the Trematoda, pp. 221–327, CABI Publishing and the Natural History Museum, New York.

Jooste, R., 1987, ‘Internal parasites of wild-life in Zimbabwe: Impala (Aepyceros melampus)’, Zimbabwe Veterinary Journal 18, 44–55.

Jooste, R., 1989, ‘A checklist of the helminth parasites from the larger domestic and wild mammals of Zimbabwe’, Transactions of the Zimbabwe Science Association 64, 15–32.

Kamaraj, C., Rahumann, A.A., Bagavan, A., Elango, G., Rajakumar, G., Zahir, A.A. et al., 2010, ‘Evaluation of medicinal plant extracts against blood-sucking parasites’, Parasitology Research 106, 1403–1412.

Kandasamy, A. & Devada, K., 2011, ‘Feasibility of coproantigen detection in amphistomosis’, Journal of Veterinary & Animal Sciences 42, 5–7.

Kanyari, P.W.N., Kagira, J.M. & Mhoma, J.R.L., 2009, ‘Prevalence and intensity of endoparasites in small ruminants kept by farmers in Kisumu Municipality, Kenya’, Livestock Research & Rural Development 21, 11.

Kanyari, P.W.N., Kagira, J.M. & Mhoma, J.R.L., 2010, ‘Prevalence and intensity of endoparasites in cattle within urban and peri-urban areas of Lake Victoria Basin, Kenya with special reference to zoonotic potential’, Scientia Parasitologica 11, 171–178.

Keyyu, J.D., Kassuku, A.A., Kyvsgaard, N.C. & Monrad, J., 2008, ‘Comparative efficacy of anthelmintics against Fasciola gigantica and amphistomes in naturally infected cattle in Kilolo District, Tanzania’, Tanzania Veterinary Journal 25, 40–47.

Keyyu, J.D., Kassuku, A.A., Msalilwa, L.P., Monrad, J. & Kyvsgaard, N.C., 2006, ‘Cross-sectional prevalence of helminth infections in cattle on traditional, small-scale and large-scale dairy farms in Iringa District, Tanzania’, Veterinary Research Communications 30, 45–55.

Keyyu, J.D., Monrad, J., Kyvsgaard, N.C. & Kassuku, A.A., 2005, ‘Epidemiology of Fasciola gigantica and amphistomes in cattle on traditional, small-scale dairy and large-scale dairy farms in southern highlands of Kenya’, Tropical Animal Health & Production 37, 303–314.

Kock, N.D., Kampamba, G., Mukaratirwa, S. & Du Toit, J., 2002, ‘Disease investigation into free-ranging Kafue lechwe (Kobus leche kafuensis) on the Kafue Flats in Zambia’, Veterinary Record 151, 482–484.

Laidemitt, M.R., Zawadzki, E.T., Brant, S.V., Mutuku, M.W., Mikoji, G.M. & Loker, E.S., 2017, ‘Loads of trematodes: Discovering hidden diversity of paramphistomoids in Kenyan ruminants’, Parasitology 144, 131–147.

Lalhmingchhuanmawii, K., Veerakumari, L. & Raman, M., 2014, ‘Anthelmintic activity of Punica granatum ethanol extract against paramphistomes in infected sheep’, Journal of Research in Animal Science 2, 079–086.

Le Roux, P.L., 1930b, ‘A preliminary communication on the life cycle of Cotylophoron cotylophoron and its pathogenicity for sheep and cattle’, in 16th Report of the Director of Veterinary Services and Animal Industries, Union of South Africa, pp. 243–254.

Le Roux, P.L., 1930a, ‘Helminthiasis of domestic stock in the Union of the South Africa’, Journal of the South African Veterinary Medical Association 1, 43–66.

Le Roux, P.L., 1932, List of helminths collected from mammals and birds in the Mazabuka area, Northern Rhodesia, Annual Report (1931), Department of Animal Health, Northern Rhodesia, Appendix B., p. 31–34.

Le Roux, P.L., 1934, Report of the Assistant Veterinary Research Officer, Annual Report (1934), Department of Animal Health, Northern Rhodesia, p. 28.

Loker, E.S., Moyo, H.G. & Gardner, S.L., 1981, ‘Trematode-gastropod associations in nine non-lacustrine habitats in the Mwanza region of Tanzania’, Parasitology 83, 381–399.

Lotfy, W.M., Brant, S.V., Ashmawy, K.I., Devkota, R., Mkoji, G.M. & Loker, E.S., 2010, ‘A molecular approach for identification of paramphistomes from Africa and Asia’, Veterinary Parasitology 174, 234–240.

Madzingira, O., Mukaratirwa, S., Pandey, V.S. & Dorny, P., 2002, ‘Helminth parasites of cattle and antelope in a mixed farming system in Zimbabwe’, in The Tenth International Congress of Parasitology, Vancouver, Canada, August 04–09, 2002, pp. 627–637.

Mage, C., Bourgne, H., Toullien, J.M., Rondelaud, D. & Dreyfuss, G., 2002, ‘Fasciola hepatica and Paramphistomum daubneyi: Changes in prevalences of natural infections in cattle and in Lymnaea truncatula from central France over 12 years’, Veterinary Research 33, 439–447.

Malrait, K., Verschave, S., Skuce, P., Loo, H.V., Vercruysse, J. & Charlier, J., 2015, ‘Novel insights into the pathogenic, diagnosis and treatment of the rumen fluke (Calicophoron daubneyi) in cattle’, Veterinary Parasitology 207, 134–139.

Marchand, A., 1984, ‘Economic effects of the main parasitosis of cattle’, Revue Medecine Veterinaire 135, 299–3302.

Mavenyengwa, M., Mukaratirwa, S. & Monrad, J., 2010, ‘Influence of Calicophoron microbothrium amphistomosis on the biochemical and blood cell counts of cattle’, Journal of Helminthology 84, 355–361.

Mavenyengwa, M., Mukaratirwa, S., Obwolo, M. & Monrad, J., 2005, ‘A macro- and light microscopical study of the pathology of Calicophoron microbothrium infection in experimentally infected cattle’, Onderstepoort Journal of Veterinary Research 72, 321–332.

Mavenyengwa, M., Mukaratirwa, S., Obwolo, M. & Monrad, J., 2006, ‘Observations on mass production of Calicophoron microbothrium metacercariae from experimentally and naturally infected Bulinus tropicus’, Onderstepoort Journal of Veterinary Research 73, 95–100.

Mavenyengwa, M., Mukaratirwa, S., Obwolo, M. & Monrad, J., 2008, ‘Bovine intestinal cellular responses following primary and challenge infections with Calicophoron microbothrium metacercariae’, Onderstepoort Journal of Veterinary Research 74, 109–120.

Mettam, R.W.M., 1932, Identification list of helminths from Departmental Collection 1920–1931, Annual Report (1931) Veterinary Department, Uganda, Appendix IB, p. 20.

Mettrick, D.F., 1962, ‘Some trematodes and cestodes from mammals of Central Africa’, Revista Biologica Lisbon 3 (2/4), 149–170.

Mohan, S.K., 2011, ‘Amphistomosis – An unnoticeable threat’, Journal of the Indian Veterinary Association 9, 60–61.

Mukaratirwa, S., Kristensen, T.K., Siegismund, H.R. & Chandiwana, S.K., 1998, ‘Genetic and morphological variations of populations belonging to the Bulinus tropicus/truncatus complex (Gastropoda: Planorbidae) in south western Zimbabwe’, Journal of Molluscan Studies 64, 435–446.

Munang’andu, H.M., Siamudaala, V.M., Munyeme, M. & Nalubamaba, K.S., 2012, ‘Detection of parasites and parasitic infections of free-ranging wildlife on a Game ranch in Zambia: A challenge for disease control’, Journal of Parasitology Research 2012, Art. # 296475, 1–8.

Munyeme, M., Munang’andu, H.M., Muma, J.B., Nambota, A.M., Biffa, D. & Siamudaala, V.M., 2010, ‘Investigating effects of parasite infection on body condition of the Kafue lechwe (Kobus leche kafuensis) in the Kafue Basin’, BMC Research Notes 3, 346.

Murphy, T.M., Power, E.P., Sanchez-Miguel, C., Casey, M.J., Toolan, D.P. & Fagan, J.G., 2008, ‘Paramphistomosis in Irish cattle’, Veterinary Record 162, 831.

Nzalawahe, J., Kassuku, A.A., Stothard, R.J., Coles, C.G. & Eisler, M.C., 2014, ‘Trematodes infection in cattle in Arumeru District, Tanzania are associated with irrigation’, Parasites & Vectors 7, 107.

Nzalawahe, J., Kassuku, A.A., Stothard, R.J., Coles, C.G. & Eisler, M.C., 2015, ‘Associations between trematode infections in cattle and freshwater snails in highland and lowland areas of Iringa Rural District, Tanzania’, Parasitology 142, 1–10.

Ortlepp, R.J., 1961, ‘N’oorsig van Suid-Afrikaanse helminte veral met verwysing na die wat in ons wild herkouers voorkom’, Tydskrif vir Natuurwetenskap 1, 203–212.

Paraud, C., Gaudin, C., Pors, L. & Chartier, C., 2009, ‘Efficacy of oxyclozanide against the rumen fluke Calicophoron daubneyi in experimentally infected goats’, Veterinary Parasitology 180, 265–267.

Pfukenyi, D.M., Monrad, J. & Mukaratirwa, S. 2005b, ‘Epidemiology and control of trematode infections in cattle in Zimbabwe: A review’, Journal of the South African Veterinary Association 76, 9–17.

Pfukenyi, D.M., Mukaratirwa, S., Willingham, A.L. & Monrad, J., 2005a, ‘Epidemiological studies of amphistomes infections in cattle in the highveld and lowveld communal grazing areas of Zimbabwe’, Onderstepoort Journal of Veterinary Research 72, 67–86.

Phiri, A.M., Chota, A., Muma, J.B., Munyeme, M. & Sikasunge, C.S., 2011, ‘Helminth parasites of the Kafue lechwe antelope (Kobus leche kafuensis): A potential source of infection to domestic animals in the Kafue wetlands of Zambia’, Journal of Helminthology 85, 20–27.

Phiri, A.M., Chota, A. & Phiri, I.K., 2007a, ‘Seasonal pattern of bovine amphistomosis in traditionally reared cattle in the Kafue and Zambezi catchment areas of Zambia’, Tropical Animal Health & Production 39, 97–102.

Phiri, A.M., Phiri, I.K., Chota, A. & Monrad, J., 2007b, ‘Trematode infections in freshwater snails and cattle from the Kafue wetlands of Zambia during a period of highest cattle-water contact’, Journal of Helminthology 81, 85–92.

Phiri, A.M., Phiri, I.K. & Monrad, J., 2006, ‘Prevalence of amphistomiasis and its association with Fasciola gigantica infections in Zambian cattle from communal grazing areas’, Journal of Helminthology 80, 65–68.

Pike, A.W. & Condy, J.B., 1966, ‘Fasciola tragelaphi sp. nov. from the sitatunga, Tragelaphus spekei Rothschild, with a note on the prepharyngeal pouch in the genus Fasciola L.’, Parasitology 56, 511–520.

Pillai, U.N. & Alikutty, K.M., 1995, ‘Clinico-hematological observation in bovine amphistomiasis’, Indian Journal of Veterinary & Medicine 15, 38–39.

Porter, A., 1921, ‘The history of some trematodes occurring in South Africa’, South African Journal of Science 18, 156–163.

Porter, A., 1938, ‘The larval trematode found in certain South African Mollusca, with special reference to schistosomiasis (bilharziasis)’, Publications of the South African Institute for Medical Research 8, 1–492.

Priya, P., Veerakumari, L. & Raman, M., 2013, ‘Anthelmintic efficacy of Acacia concinna against paramphistomes in naturally infected sheep’, Journal of Applied Animal Research 41, 183–188.

Prudhoe, S., 1957, ‘Trematoda-Exploration du Parc National de l’Upemba’, in Mission G.F. de Witte 1946–49), vol. 1, p. 48, Royal Belgian Institute of Natural Sciences, Brussels.

Prudhoe, S., Yeh, L.S. & Khalil, L.F., 1964, ‘A new amphistome trematode from ruminants in Northern Rhodesia’, Journal of Helminthology 38, 57–62.

Reinecke, R.K., 1983, Veterinary helminthology, Butterworths, Durban.

Rieu, E., Recca, A., Benet, J.J., Saana, M., Dorchies, P. & Guillot, P., 2007, ‘Reliability of coprological diagnosis of Paramphistomum sp. infection in cows’, Veterinary Parasitology 146, 249–253.

Rinaldi, L., Perugini, A.G., Capuano, F., Fenizia, D., Musella, V., Veneziano, V. et al., 2005, ‘Characterization of the second internal transcribed spacer of ribosomal DNA of Calicophoron daubneyi from various hosts and locations in southern Italy’, Veterinary Parasitology 131, 247–253.

Roach, R.W. & Lopes, V., 1966, ‘Mortality in adult ewes resulting from intestinal infestation with immature paramphistomes complicated by severe fascioliasis’, Bulletin of Epizootic Diseases in Africa 14, 317–323.

Roberts, J.A. & Suhardono, 1996, ‘Approaches to the control of fasciolosis in ruminants’, International Journal for Parasitology 26, 971–981.

Rolfe, F., Boray, J.C. & Collins, G.H., 1994, ‘Pathology of infection with Paramphistomum ichikawai in sheep’, International Journal for Parasitology 24, 995–1004.

Rolfe, P.F. & Boray, J.C., 1987, ‘Chemotherapy of paramphistomosis in cattle’, Australian Veterinary Journal 64, 328–332.

Rolfe, P.F. & Boray, J.C., 1988, ‘Chemotherapy of paramphistomosis in sheep’, Australian Veterinary Journal 65, 148–150.

Rolfe, P.F. & Boray, J.C., 1993, ‘Comparative efficacy of moxidectin, an ivermectin/clorsulon combination and closantel against immature paramphistomes in cattle’, Australian Veterinary Journal 70, 265–267.

Rolfe, P.F., Boray, J.C., Nichols, P. & Collins, G.H., 1991, ‘Epidemiology of amphistomosis in cattle’, International Journal for Parasitology 21, 813–819.

Sachs, R. & Sachs, S., 1968, ‘A survey of parasitic infections of wild herbivores in the Serengeti region northern Tanzania and the Lake Rukwa region southern Tanzania’, Bulletin of Epizootic Diseases in Africa 16, 455–472.

Salib, F.A., Halium, M.M.A., Mousa, W.M. & Massieh, E.S.A., 2015, ‘Evaluation of indirect ELISA and Western blotting for the diagnosis of amphistomes infection in cattle and buffaloes’, International Journal of Livestock Research 5, 71–81.

Sanabria, R., Moreno, L., Alvarez, L., Lanusse, C. & Romero, J., 2014, ‘Efficacy of oxyclozanide against adult Paramphistomum leydeni in naturally infected sheep’, Veterinary Parasitology 206, 277–281.

Sanchis, J., Sanchez-Andrade, R., Macchi, M.I., Pineiro, P., Suarez, J.L.C., Cazapal-Monteiro, J.L. et al., 2012, ‘Infection by Paramphistomidae trematodes in cattle from two agricultural regions in NW Uruguay and NW Spain’, Veterinary Parasitology 191, 165–171.

Sey, O., 1982, ‘Revision of the genus Cotylophoron Stiles et Goldberger, 1910 (Trematoda: Paramphistomata)’, Helminthologia 19, 11–24.

Sey, O., 1983, ‘Revision of the family Gastrothylacidae Stiles et Goldberger, 1910 (Trematoda, Paramphistomata)’, Acta Zoologica Academiae Scientiarum Hungaricae 29, 223–252.

Sey, O., 2000, Handbook of the zoology of amphistomes, CRC Press, Boca Raton, FL.

Sey, O. & Graber, M., 1979a, ‘Examinations of amphistomes (Trematoda: Paramphistomata) of some African mammals’, Revue d’Élevage et de Médecine Vétérinaire des Pays Tropicaux 32, 161–167.

Sey, O. & Graber, M., 1979b, ‘Cotylophoron macrosphinctris sp. n. (Trematoda: Paramphistomata) from the African buffalo, Bubalus (Syncerus) caffer Sparrman’, Annales de Parasitologie Humaine et Comparee 45, 297–302.

Shivjot, K., Singla, L.D., Hassan, S.S. & Juyal, P.D., 2009, ‘Standardization and application of indirect plate ELISA for immunodiagnosis of paramphistomosis in ruminants’, Journal of Parasitic. Diseases 33, 70–76.

Sibula, M.S., Dhlamini, Z. & Dube, S., 2014, ‘Molecular characterization of paramphistomes from cattle from Matabeleland region (Zimbabwe) using random amplified polymorphic DNA (RAPDs) and amplified ribosomal DNA restriction analysis (ARDRA)’, Advanced Biomedical Research 5, 92–99.

Sobrero, R., 1962, ‘Ricostruzione del ciclo di vita di Paramphistomum clavula (Näsmark, 1937), parassita dei ruminanti in Somalia’, Parasitologia 4, 165–167.

Southgate, V.R., Brown, D.S., Warlow, A., Knowles, R.J. & Jones, A., 1989, ‘The influence of Calicophoron microbothrium on the susceptibility of Bulinus tropicus to Schistosoma bovis’, Parasitology Research 75, 381–391.

Spencer, S.A., Fraser, G.C. & Chang, S., 1996, ‘Responses in milk production to control of gastrointestinal nematode and paramphistome parasites in dairy cattle’, Australian Veterinary Journal 74, 456–459.

Swart, P.J., 1954, ‘The identity of so-called Paramphistomum cervi and P. explanatum, two common species of ruminant trematodes in South Africa’, Journal of the South African Veterinary Medical Association 26, 463–473.

Swart, P.J. & Reinecke, R.K., 1962a, ‘Studies on paramphistomiasis. I. The propagation of Bulinus tropicus Krauss, 1848’, Onderstepoort Journal of Veterinary Research 29, 183–187.

Swart, P.J. & Reinecke, R.K., 1962b, ‘Studies on paramphistomiasis. II. The mass production of metacercariae of Paramphistomum microbothrium Fischoeder 1901’, Onderstepoort Journal of Veterinary Research 29, 189–195.

Toolan, D.P., Mitchell, G., Searle, K., Sheehan, M., Skuce, P.J. & Zadoks, R.N., 2015, ‘Bovine and ovine rumen fluke in Ireland – Prevalence, risk factors and species identity based on passive veterinary surveillance and abattoir findings’, Veterinary Parasitology 212, 168–174.

Vassilev, G.D., 1994, ‘Prevalence and seasonality of internal parasite infections detectable by faecal examination of cattle in Chiweshe communal farming area of Zimbabwe’, Zimbabwe Veterinary Journal 25, 41–63.

Vassilev, G.D., 1999, ‘Prevalence of internal parasite infections of cattle in the communal farming areas of Mashonaland East Province, Zimbabwe’, Zimbabwe Veterinary Journal 30, 1–17.

Vatta, A.F. & Krecek, R.C., 2002, ‘Amphistome infection of goats farmed under resource-poor conditions in South Africa’, Onderstepoort Journal of Veterinary Research 69, 327–329.

Von Roth, H.H. & Dalchow, W., 1967, ‘Untersuchungen über den Wurmbefall von Antilopen in Rhodesien’, Zeitschrift für Angewandte Zoologie 54, 203–226.

Wandera, J.G., 1969, ‘Reflection on sheep diseases in Kenya’, Bulletin of Epizootic Diseases in Africa 17, 121–126.

Waruiru, R.M., Mbuthia, P.G. & Kimoro, C.O., 1993, ‘Prevalence of gastrointestinal parasites and liver flukes in calves in Mathira Division of Nyeri District, Kenya’, Bulletin of Animal Health & Production in Africa 41, 291–296.

Woolhouse, M.E.J. & Chandiwana, S.K., 1990, ‘The epidemiology of schistosome infections of snails: Taking the theory into the field’, Parasitology Today 6, 65–70.

Wright, C.A., Southgate, V.R. & Howard, G.W., 1979, ‘A note on the life-cycles of some amphistome flukes in Zambia’, Journal of Helminthology 53, 251–252.

Yabe, J., Phiri, I.K., Phiri, A.M., Chembensofu, M., Dorny, P. & Vercruysse, J., 2008, ‘Concurrent infections of Fasciola, Schistosoma and Amphistomum spp. in cattle from Kafue and Zambezi river basins of Zambia’, Journal of Helminthology 82, 373–376.

Yeh, S.L., 1957, ‘On a new paramphistome trematode Gigantocotyle leuroxi sp. n. from the stomach of the red leche, Onotragus leche from Northern Rhodesia’, Parasitology 47, 432–434.

Yogeshpriya, S., Ajithkumar, S., Dhanesh, V., Kutty, R.R. & Alex, P.C., 2011, ‘Paralytic ileus due to amphistomosis in a cow’, Journal of the Indian Veterinary Association 9, 48.


Crossref Citations

1. SEM and molecular approaches to identify Calicophoron clavula in Saudi Arabia
O. Soliman, M. M. Montaser, A. A. Ashour, M. I. Soliman, A. H. Nigm
Journal of Parasitic Diseases  vol: 44  issue: 1  first page: 239  year: 2020  
doi: 10.1007/s12639-019-01187-3