Plates 22 & 23 (pp. 140 – 141) and Fig. 4 (p.142).
Species affected and geographic range
Khalil (1971) lists over 50 species of trematodes, from 15 families, occurring in a variety of freshwater fish in Africa. Of these, only the extraintestinal species are potentially harmful to fish; species of Sanguinicola (the blood fluke) infect Synodontis schall and Auchenoglanis occidentalis in the Sudan (Khalil, 1969) and Clarias lazera (Paperna, 1964b) and Oreochromis spp. in Israel. Callodistomid and opistorchid species infect the gall bladder and bile ducts of diverse fish species such as Polypterus bichir, Synodontis schall and Gymnarchus niloticus, while species of Phyllodistomum are found in the urinary bladder of siluroids, Ctenopoma kingsleye, Mastacembalus nigromarginatus and Gymnarchus niloticus. One representative of the Didimozoidae (parasites of fish tissues and internal cavities), Nemathobothrium labeonis, occurs (unencysted) in the eye orbits of Labeo spp. in the Sudan Nile.
Taxonomy, description and diagnosis
Trematodes or Digenea are flatworms (Platyhelminthes), heteroxenous (with a multiple host life cycle) and require (with only one exception) a mollusc as their first intermediate host.
Adult-stage digeneans usually have a dorso-ventrally flattened, oval body with a smooth, spiny or corrugated surface, a sucker around the antero-ventral mouth, and an additional ventral sucker or acetabulum. Both suckers are used for attachment and locomotion. The digestive system consists of a pharynx connected to the mouth opening, a short oesophagus and two blind intestinal caeca. Most trematodes are hermaphrodite, containing both male organs (testes, ducts and copulatory system) and female organs (ovary, vitelline glands, ducts and uterus). Some also contain a specialised copulatory organ (gonotyl in Heterophyes spp.) which is useful for differential diagnosis. Eggs are evacuated to the genital opening, and are usually oval and operculated (Schell, 1970).
Blood flukes (Sanguinicolidae) are slender, spiny, and lack anterior ventral suckers and pharynx. The intestinal caeca are short, X- or H-shaped. Eggs are thin-shelled and lack an operculum (Smith, 1972).
Didymozoidae are thread-like, with or without an expanded posterior region, and occur in pairs or small groups inside body cavities or within cysts or cyst-like formations in the tissues. Some are hermaphrodite, while others show variable degrees of separation into sexes (Dawes, 1946).
Differential diagnosis is difficult and requires experience with trematode taxonomy. Fixation, which allows further processing and adequate staining, should be done with Alcohol (70–95%) under moderate pressure of a glass slide or cover slip (depending on specimen size). Staining for demonstrating internal organs, if desired, may be done with either haematoxylin or carmine stains.
Life history and biology
The life histories of the trematodes which (at the adult stage) infect African fish have so far not been studied and their first molluscan host and other intermediate hosts remain unknown as yet. Data available on trematodes elsewhere (Dawes, 1946; Hoffman, 1967; Schell, 1970), may be summarised as follows:
Eggs of gut dwelling digeneans are released via defaecation, while eggs of those living in the gall-bladder are evacuated into the gut with bile. Eggs, produced by digeneans in the kidney or gonads, are evacuated from their host with the respective organ's products. If they are located in tissue or closed internal cavities they can only be liberated following death of the host or predation (Didymozoid eggs).
Eggs of blood flukes (Sanguinicolidae), containing a fully developed miracidium, accumulate in the terminal (distal) blood capillaries. Only those reaching the gill filament blood vessels release their miracidia, which then actively break through the gill tissue into the water (Davis et al., 1961; Smith, 1972). Eggs of Sanguinicola dentata in Clarias lazera (Paperna, 1964b) were accumulating in the kidney and seemed to evacuate via the urinary system.
Eggs, if laid undeveloped (Paramphistomatidae), begin their embryonic development only after evacuation from the host, apparently after being triggered by appropriate stimuli (the presence of oxygen and light). Eggs of many piscine digeneans, however, when laid contain a fully developed miracidium. Such eggs hatch immediately or soon after evacuation from the definitive host (Asymphilodora tincae - Van den Broek and de Jong, 1979). Fully embryonated eggs of Ophistorchiidae do not hatch, but infect snail hosts upon being swallowed.
Free-swimming miracidia are pyriform, and covered with cilia.
Both bivalve and gastropod molluscs serve as intermediate hosts for trematodes which reach the adult stage in fish. In fresh waters, both prosobranchs and pulmonates are involved. Trematodes demonstrate a high degree of specificity to their molluscan hosts. Bivalves are first intermediate hosts for Fellodistomatidae, Gorgoderidae and Allocreadiidae (Hoffman, 1967; Schell, 1970). Pulmonates are the molluscan hosts of blood flukes (Sanguinicola spp.), infecting freshwater fish (Lymnaea spp. of S. inermis of carp) (Smith, 1972), and of Plagiorchidae. Freshwater prosobranch snails are hosts to both Ophistorchiidae and Monorchidae.
The miracidium, upon reaching the molluscan host, transforms into a “mother” sporocyst. Sporocysts yield a new generation of either sporocysts or rediae. Daughter stages migrate and settle in the molluscan hepato-pancreas. The sporocyst consists of only a tegumental sac, while the redia contains a muscular pharynx connected to a sac-like intestine, and a birth-pore located near the pharynx. In these, or their sporocyst or redia offspring, the cercariae are formed. Intramolluscan development of the Allocreadiidae, Haploporidae, and Monorchiidae (Dawes, 1946; Fares & Maillard, 1974; Van den Broek & De Jong, 1979) includes both sporocyst and redia stages. In Sanguinicolidae and Plagiorchidae (Hoffman, 1960; Smith, 1972), the cercariae are formed in the daughter sporocyst stage.
Cercariae already have the elements of mature digenean organisation, but with primordial genital organs. Cercariae may also have locomotive devices; a tail, in some forked, fins (the forked-tailed cercariae of sanguinicoliids also have a characteristic dorso-median fin fold), and a pair of eyes. The latter are lost when cercariae transform into metacercariae.
Of all piscine trematodes, only the blood flukes (Sanguinicolidae) and Transversotrematidae have cercariae which develop directly into adults in their definitive host. Cercariae of these flukes actively penetrate into their definitive piscine host (Sommerville & Iqbal, 1991; Rao & Ganapati, 1967). All other flukes which attain maturity in piscine hosts, reach their definitive host as waiting stage metacercariae. Cercariae transform into metacercariae when penetrating aquatic invertebrate or vertebrate (fish, tadpoles) organisms, or after encystment on plant material or other substrates in the water (example Haploporidae, Fares and Maillard, 1974). Transmission into the definitive hosts occurs when metacercariae are predated with their intermediate hosts, or browsed from the substrate by suitable fish hosts. Infection of fish by metacercarial stages is therefore closely linked to their food preferences.
Bucephalidae, and Acanthostomidae (in part) life histories involve fish as hosts for both metacercariae and adult stages. Tadpoles are second intermediate hosts for some Gorgoderidae (Schell, 1970).
Life histories involving molluscs as second intermediate hosts are found among members of very diverse digenean families; Monorchidae, Phyllodistomatidae, Azygiidae and Lepocreadiidae (Dawes, 1946; Schell, 1970). Monorchidae, developing in Bithynia or in bivalves, also exploit their first molluscan host for metacercarial encystment (Van den Broek and de Jong, 1979).
Other digeneans reach their definitive piscine host via planktonic or benthic organisms consumed as food. Common second intermediate hosts of digeneans infecting freshwater fish (such as Allocreadiidae), are larvae of aquatic insects; mayflies (Ephemeroptera), caddis-flies (Trichoptera), Dragon flies (Odonata) and Chironomidae and also various Crustacea, leeches, oligochaetes and planarians (see Dawes, 1946; Hoffman, 1967 and Schell, 1970).
Adult trematodes, infecting the digestive tract of fish, are considered harmless, even when their numbers are high. Extraintestinal trematode infections, on the other hand, are potentially pathogenic.
Thus far, only the pathological data on blood flukes (sanguinicolids which can cause considerable damage to the gills and impair respiration) are relevant to African fish. Adult worms and trapped eggs can physically obstruct the passage of blood, causing thrombosis and subsequent necrosis (Hoffman et al., 1985), while escape of miracidia through the gill epithelium causes loss of blood and may lead to anaemia (Evans, 1974; Davis et al., 1961). Proliferation of the arterial endothelium was reported in common carp infected with Sanguinicola inermis (Prost, M., Poland, in Lucky, 1964). Loss of blood was evident from the pale colour of the gills and the decline in packed cell volumes and oxyhaemoglobin levels (Evans, 1974a). Heavy infection compromises the host's ability to withstand stressful conditions, for example heavily infected cultured carp suffocated during transportation (Lucky, 1964; Smith 1972). In chronic infections, adult worms disperse and become stranded in the heart, kidneys and caudal vessels. Dispersed eggs become encapsulated, and may also become surrounded with a focal granuloma. Nodular foci have been demonstrated in the heart, head, kidney and spleen of carp (Lucky, 1964) and Oreochromis spp. In S. armata infected grass carp (Cteno-pharyngodon idella) and bighead (Aristichthys nobilis), tissue response to eggs, spread throughout the viscera, was negligible (Anderson and Shaharom-Harrison, 1986) and similarly to eggs of S. dentata infiltrating the kidney of Clarias lazera.
The didimozoids (N. labeonis) recovered from the orbits were all young and unencysted and were never observed penetrating the eyeball, and thus did not cause any detectable pathological effect (Khalil, 1969).
All trematodes are host specific and transmission may, at most, involve species of the same or very close genera. The presence of suitable vector snails in the habitat is essential for transmission (see the chapter on the epizootiology of metacercarial infection in relation to the environmental aspects of snail distribution - p. 130).
Sanguinicola infection is fairly common (quantitative data are lacking) in Oreochromis aureus of L. Kinneret, Israel. Data on blood fluke species, in the other African piscine hosts, are scarce. Blood flukes were reported in 6% of examined Auchenoglanis occidentalis, in the Sudan Nile (Khalil, 1969). The cichlid blood fluke recently became established in a fish farm, in Israel, in concrete holding tanks fed by open canals of surface water. By the time infection was detected, only Physa snails were present, which may not have been the vector snails.
American species of Sanguinicola have been implicated in massive mortalities of hatchery reared salmonids after their vector snails became established in the culture system (Davis et al., 1961; Evans, 1974; Hoffman et al., 1985). S. inermis (transmitted by Lymnaea spp.), at times severely infects common carp in extensive eastern European ponds which allow propagation of Lymnaea (Lucky, 1964). Anderson and Shaharom-Harrison (1986) reported the introduction of S. armata with infected bighead carp (Aristichthys nobilis) and grass carp (Ctenopharyngodon idella) into fish farms in Malaysia.
Of Labeo species (4 spp.) examined in the Nile, 53% harboured in their orbits 3 to 10 N. labeonis (Didymozoidae); in 41 out of 49 fish both eyes were infected (Khalil, 1969). Trematodes of the digestive tract are often very common, and also numerous (Khalil, 1969, Ukoli, 1969), in some samples of Bagrus spp. all fish were infected by Phyllodistomum, some with up to 150 worms. Infections of Phyllodistomum bavuri in Clarias gariepinus in Kruger National Park (northern Transvaal) occurred throughout the year with no evident seasonal fluctuation in incidence of infection or worm burden (Boomker, 1984).
Control (see in Metacercaria section below).
Most freshwater and estuarine fish are potential hosts, but juvenile fish, bottom dwellers and shallow water inhabitants are most vulnerable.
Metacercarial infections were found in fish in all studied inland water bodies in Africa and the Near East (Fahmy & Selim, 1959; Paperna, 1964; Williams & Chaytor, 1966; Khalil, 1969, 1971; Paperna & Thurston, 1968; Van As & Basson, 1984).
Piscivorous birds are the definitive hosts for many of the metacercariae found in fish. Consequently, bird migration along the eastern (over the Syro-African rift) and the western routes from Eurasia to Africa is the greatest contributory factor for dispersal of metacercarial infections. The other equally important factor is the presence of suitable molluscan intermediate hosts. It has also been suggested that aquatic birds help in the dispersal of aquatic snails. Water bodies from the Jordan system throughout the Nile to the Rift Valley lakes share common snails (Bulinus truncatus, same species group Lymnaea and Melanoides tuberculata), and similar fish (cichlids, Clarias and Barbus), which become infected by the same metacercariae (“black spot” Neascus, Clinostomum tilapiae, C. complanatum, Eculinostomum heterostomum, Centrocestus spp. Phagicola spp. and Haplorchis spp.). All of these have been demonstrated to have herons, cormorants and pelicans as definitive hosts. Where species can be determined (in Clinostomatidae), data suggests transcontinental distribution in both East and West Africa (Ukoli, 1966a, b).
A different pattern of distribution characterises heterophyiids utilising the lagoon dwelling euryhaline snail Pirenella conica as a molluscan host. These parasites, which develop in both avian and mammalian definitive hosts, occur in fish of estuarine and lagoon habitats of the Mediterranean as well as the Red Sea and Indian Ocean coasts (Balozet & Callot, 1938; Taraschewski, 1984; Taraschewski & Paperna, 1981), but are apparently absent in the remaining coastal regions of the continent.
Taxonomy, description and diagnosis
Members of some families or even certain genera may be recognised through characteristic structural affinities, aided by additional features such as the type and location of encystment. In other instances even family affinities cannot be determined. To a limited extent mature trematodes may be obtained from metacercariae through experimental infection of known or suspected definitive hosts; herons and pelicans in the case of Strigeoidea or Clinostomatidae (Ukoli, 1966a,b; Williams, 1967; Donges, 1974), dogs and cats with Heterophyiidae (Witenberg, 1929), or laboratory mice, rats, chicks or ducklings when the trematode is non-fastidious in its choice of definitive host (Khalil, 1963; Williams & Chaytor, 1966, and particularly heterophyiids - Sommerville, 1982a, Taraschewski, 1984).
Metacercariae may be released from their cysts for better examination either by teasing or applying pressure, or with digest solutions (in Pepsin, 5% in 0.1N/HCI and then 1% Trypsine with 0.5% Sodium-taurocholate in 1% NaHCO3).
Strigeoid metacercariae (Strigeidae, Diplostomatidae and Cyathocotylidae) encyst in a variety of organs, including the inside of the eye ball. Some diplostomatids remain temporarily or ultimately (in the eye lens and retina) unencysted. The mature metacercaria is divided into a cup-shaped forebody carrying the suckers, and a cylindrical hindbody containing the rudiments of the reproductive organs. The function of the ventral sucker is taken over by a new holdfast (tribocytic) organ (Hoffman, 1960). The cysts of some (of the larval genus Neascus) occurring in the skin accumulate melanophores, or other skin chromophores (“Black spot”).
Clinostomatid cysts and worms are the largest (up to 5 mm in diameter and 10 × 3 mm in size) and the worm's intestine is loaded with a yellow to orange substance.
Heterophyiids are covered by spines. The male genital pore of Heterophyes spp., Stictodora and others is accompanied by spines, arranged on a special round structure (gonotyle) or otherwise. Some have one or two rows of oral spines (Parascocotyle or Phagicola and Ascocotyle) and also encyst within a cartilaginous capsule on the gill filaments (Centrocestus) (Witenberg, 1929; Paperna, 1964a; Farstey, 1986). Oral spines of a different pattern occur in metacercariae of Echinostomatidae.
Life history and biology
The general pattern of trematode life history and the development of each of its larval stages has been outlined previously (13.1, in the description of adult trematode life histories).
The most common definitive hosts of Diplostomatidae (and other Strigeoidea), Clinostomatidae and Heterophyiidae encysting as metacercariae in fish are piscivorous birds. Mammalian hosts, including dogs, play an important part in dissemination of Heterophyiidae and the stregioid Prohemistomum vivax (Witenberg, 1929; Fahmy & Selim, 1959). Heterophyiidae, notably Heterophyes heterophyes, are very versatile in their choice of definitive hosts and will develop to maturity in both mammals and birds. Crocodiles (and possibly Nile monitors) are definitive hosts to metacercariae of the clinostomatid Nephrocephalus (Dollfus, 1930), and Pseudoneodislostomum thomasi (Fischthal & Thomas, 1970) which infect Bagrus and Clarias spp.
Herons are the common definitive hosts of Diplostomatidae and natural infection of B. levantinus has been found in Ardea purpurea. Eggs of diplostomatids are shed undeveloped; light and oxygen trigger the onset of embryonic development. Data on incubation schedules for African species are lacking. Eggs of D. spathaceum, incubated at 29°C, hatched after 9–11 days, while infected snails (Lymnaea peregra) commenced shedding within 6–9 weeks (Whyte et al., 1988). Cercariae of all diplostomatids are fork-tailed (furcocercariae). Bulinus truncatus, the snail host of Bolbophorus levantinus was found shedding 7 weeks after being placed with freshly laid eggs (at an ambient temperature of 22–24°C (Paperna & Lengy, 1963). B. truncatus from the fringes of L. Kinneret also shed furcocercariae which developed, in juvenile cichlids, into blackspot (Neascus). The vector of Neascus causing blackspot in L. Victoria cichlids is the local bulinid, B. ugandae. Blackspot metacercariae occurring on non cichlid fish might well be a different species. Metacercariae of B. levantinus developed only in species of Oreochromis. In Bolbophorus levantinus, metacercariae were shown to develop from a ‘distome’ to a strigeoid form, their posterior half distending while becoming densely filled with vesicular cells (reserve bladder - Hoffman, 1960). These are released into the cyst lumen at the end of the metacercarial development and the posterior end becomes the genital segment (Paperna & Lengy, 1963; Yekutiel, 1985). The same process apparently occurs in metacercariae of Ornithodiplostomum and Postdiplostomum, in which the posterior segment is comprised of a “reserve bladder” (Hoffman, 1960).
Hyperparasitism, i.e. a cyst within another cyst of an apparently different species of Diplostomatidae has been revealed in Clarias gariepinus muscles in Israel and in Uganda.
Definitive hosts for species of Clinostomum and Euclinostomum are herons, pelicans, cormorants and darters (Anhinga rufa). In all of these the adult trematodes become attached to the wall of the posterior pharynx and in the laryngeal zone. Some species, however, may restrict their choice of hosts; C. complanatum fails to become established in pelicans (Finkelman, 1988). Eggs, shed by worms, are either washed directly to the water habitat, or swallowed and defaecated.
Eggs are shed undeveloped, and like those of diplostomatids, require oxygen and light for development. Miracidia of C. tilapiae hatched following 10 days incubation at 25–30°C (under constant illumination) and those of C. marginatum after 11–13 days (Finkelman, 1988).
In Israel, B. truncatus was shown to be the intermediate host for Clinostomum tilapiae (Finkelman, 1988). Elsewhere in Africa, where B. truncatus is absent, C. tilapiae is likely to be transmitted by other bulinids (in South Africa). Another bulinid, Bulinus (Physopsis) globosus, is the vector of Euclinostomum heterostomum (Donges, 1974). Clinostomum complanatum develops through species of Lymnaea (Radix) (Lo et al., 1982; Finkelman 1988).
C. tilapiae infected snails start to shed after 40 days and C. complanatum infected snails after 30 days (Finkelman, 1988). Cercariae are fork tailed with a dorso-median fin fold (similar to that seen in sanguinicolid cercariae).
Shed eggs of heterophyiids, contain a miracidium which hatches and commences development when ingested by the vector snail (Khalil, 1937). Heterophyiid snail hosts are prosobranch snails; Melanoides tuberculata (host to Centrocestus spp., Haplorchis spp. and Stellantchasmus falcatus) in freshwater inland habitats (Khalifa et al. 1977, Sommerville, 1982, Farstey, 1986) and Pirenella conica and species of Hydrobia in euryhaline waters (notably Heterophyes spp. and Stictodora spp.) (Khalil, 1937; Martin, 1959; Taraschewski & Paperna, 1981). Heterophyiid cercariae have an undivided tail (Pleurolophocercous).
Clinical effects of infection are often not obvious. The presence of metacercariae in supposedly sensitive organs such as the brain, cranial nerves or spinal cord [Diplostomum mashonense and D. tregenna, in Clarias spp. (Beverly-Burton, 1963; Khalil, 1963)], does not necessarily imply a debilitating impact on the fish, even at relatively high infection loads, and despite visible structural damage. Sudden, massive outbreaks of infection can be fatal. Cercariae penetrate via the skin and gills (Hoglund, 1991). Exposure to massive numbers of cercariae may kill fry within a few hours (cichlids infected by Haplorchis pumilio - Sommerville, 1982a), but such exposures are not representative of naturally occurring infections. Cercariae penetrated and encysted deeper in the tissues of small fish and the large cysts interfered with organ function. The large (0.5–0.8 mm in diam.) and numerous (over 50) cysts of B. levantinus, established in muscles of young cichlids (<50 mm long), induce severe body deformities (Paperna & Lengi, 1963; Yekutiel, 1985). Metacercariae form massive infections in juvenile (O-class) fish and have, therefore, been implicated as an important cause of natural mortalities at this stage of their lives (Centrocestus spp. in gills and Bolbophorus levantinus in muscles of cichlid fish - Yekutiel, 1985; Farstey, 1986; Paperna, 1991). Population studies and field observations suggest that fish heavily infected by metacercariae are selectively removed from the host population (Chubb, 1979). Heavy gill infection appears to lower respiratory efficiency. During 3hrs of transport, all young cichlids (Sarotherodon galilaeus) with heavily Centrocestus-infected gills (116±48 per fish), succumbed, while all lightly infected (same size, with 15±15 per fish) survived (Farstey, 1986).
The pathological impact of cardiac infections by Phagicola and Ascocotyle in cichlids (and also grey mullets) was not evaluated. Trout infection with Apatemon gracilis resulted in fibrogranulomatosis of the epicardium and failure in in-vitro pumping performance (Tort et al., 1987; Watson et al., 1992).
Pronounced inflammatory response and focal haemorrhages accompanies penetration and early migration (in muscles penetrated by H. pumilio - Sommerville, 1982a). The inflammatory reaction, predominated by infiltrating macrophages, is particularly intense around unencysted migrating metacercariae and preceded the eventual enclosure in a fibrous capsule of the encapsulating metacercaria (B. levantinus - Yekutiel, 1985). The fibrous capsules produced by the host, are superimposed on the acellular wall secreted by the encysting cercaria.
Cysts consolidating around certain skin metacercariae may incorporate dermal melanophores and exceptionally, other chromophores. Such metacercariae, termed “black spot”, are formed in infections by the strigeoid larval genus Neascus (species of Crassophialia, Ornithodiplostomum and Uvulifer - Hoffman, 1960) and many others whose adult stages are unknown.
Centrocestus metacercariae on gills become encysted in a cartilaginous capsule, which is comprised of a cartilaginous extension of the filament's ray. Proliferation of the gill epithelium around the forming capsule, with the resulting obliteration of the lamellar structure, is apparently the cause of the observed respiratory malfunction in the infected fish (Farstey, 1986).
In spite of the large size (3–7 mm) of the clinostomatid cysts, neither skin infection nor muscle and visceral infection induces severe histopathology or gross pathological effects in fully grown or even juvenile fish. Heterotis niloticus tolerates infections as high as 130 Nephrocephalus metacercariae and up to 70 metacercariae of Clinostomum sp. were counted in muscles of individual Synodontis membranaceus (Ukoli, 1969) and of C. complanatum, in Tor (Barbus) canis (Finkelman, 1988). Seemingly healthy looking cichlids (Tristramella simonis, in L. Kinneret) are occasionally found virtually covered by cutaneous cysts of Clinostomum spp (Paperna, 1964a,b). Very young fish (O. mossambicus, 40–60 mm long), however, succumbed to infection by 3–5 cysts of Euclinostomum heterostomum in the viscera. Donges (1974) reports kills of experimentally hyperinfected O. mossambicus fingerlings by Euclinostomum heterostomum, 30–35 and 62 days post infection with loads of 75–81 worms.
Damage to the eyes of fish is caused by metacercariae with a predilection, or even site-specificity to that organ or as a non-specific side-effect, for example, corneal infection by integument-encysting metacercariae, which impairs eye vision. This condition is aggravated when metacercariae are accompanied by melanophores (black spot). The specific lens parasite Diplostomum spathaceum is unknown from African waters, but several infections by other diplostomatid metacercariae have been reported, usually invading the anterior or vitreous humor rather than the lens. Infected eyes often contain up to 4 metacercariae, 2.5 mm in size (with aggregates of up to 200) free, and encysted. Mashego (1982) reports cysts containing both juvenile metacercariae as well as some enveloped, more advanced metacercariae of the same or different species. Severe infection leads to exophthalmos, cataracts, and even complete collapse of the eye. Blindness can be uni- or bilateral (Thurston, 1965; Lombard, 1968; Douellou, 1992).
Prevalence data and host records hint that infestation by metacercariae occurs only, or predominantly in shallow waters where most vector snails live. In lakes, Melanoides tuberculata and Pirenella conica are often found at high densities on the fringe of the shore line. Nonetheless, in large lakes such as Lake Victoria, shallow lakes such as L. George or very small, but relatively deep lakes such as L. Kinneret, infections of skin with black spot, gills with Centrocestus and inner tissues with Bolbophorus levanticus (seen only in L. Kinneret) and Haplorchis spp., are common and high only among young fish, or species confined to shallow water (T. zillii and some species of Haplochromis).
Older fish in offshore water are only sparsely or exceptionally found infected. The only exception are some species of offshore Haplochromis where infection is also retained in the older age classes (which apparently spend part of their time inshore).
Daily cercarial production in heterophyiids is around 300–500 and may last for over a year. Daily cercarial output in pulmonate snails is often similar (Wright, 1971; Paperna, unpublished), or even higher (in Bolbophorus levantinus 2000–3500 per 24 hr period -Paperna & Lengy, 1963) but overall production time is restricted, as diplostomatids developing in pulmonates have only sporocyst stages. This can explain the prevalent and often extremely heavy infections which are often observed in fish of inshore waters.
Studies in Lake Kinneret, have shown (Farstey, 1986) that the highly prevalent metacercarial infections are sustained by a relatively sparse infection in the snails: 0.6–9% (with one instance of 32%) of M. tuberculata were shedding Haplorchis cercariae and 2–10% were shedding Centrocestus.
The distribution pattern of infection with all these metacercariae was very over dispersed, for example, with an average of less than 40 gill metacercaria per fish in L. Victoria cichlids, some were showing infection by 120 and up to 800. Statistically significant overdispersion parameters (best fit to negative binomials and others) were also demonstrated in B. levantinus infections of juvenile cichlids (Yekutiel, 1985).
In the Nile Delta lakes, Oreochromis niloticus as well as the grey mullets are important hosts of heterophyiids (including the zoonotic human pathogen H. heterophyes -Taraschewski, 1984) and Prohemistomum vivax metacercaria, with dogs acting as the main source for eggs to infect the vector snail (Wells & Randall, 1956; Fahmi & Selim, 1959; Taraschewski, 1984). Infection accumulates with time and in large fish approximates 100%. Metacercariae loads in O. nilotica were not reported but in grey mullets 300 to 3000 Heterophyes spp. were counted per 1g of fish flesh (Paperna & Overstreet, 1981).
Eye infections by Diplostomatidae were reported in 85% of examined T. rendalli and O. mortimeri in Lake Kariba (Douellou, 1992), the majority had bilateral infection. Eye infections are prevalent in less than 30% of L. Victoria Haplochromis (Thurston, 1965). In South Africa, in some dams (in Transvaal, Lombard, 1968), virtually all fish were found to be affected (100% of Barbus paludinosus with 5–20 metacercariae per fish). Outbreaks of ocular infection also occurred in farmed tilapia as well as in the introduced trout and large-mouth bass (Micropterus salmoides) (Lombard, 1968).
Of the clinostomatids, C. tilapiae and Euclinostomum heterostomum infections are widespread (Lombard, 1960), although prevalence in some habitats (in dams in Transvaal - Britz et al., 1985) may reach 76%, numbers of recovered worms only exceptionally exceed 10. Worm load per fish of C. complanatum in Tor (Barbus) canis in Lake Kinneret is considerably higher (up to 70, mean 33). Large numbers of metacercariae are found in Gnathonemous macrolepidotus infected with C. vanderhorsti (Ortlepp, 1935) in southern Africa.
Intensively utilised earth ponds, with their heavy organic and nitrogenous load and muddy (eutrophic) bottoms are unfavourable habitats for all snails. Omnivorous fish, such as carp and siluroid catfish, eat thin-shelled snails and their spawn. Metacercarial infections in intensive earth pond systems, therefore, occur only sporadically, as episodes restricted to a single growing season, and are eliminated when ponds are returned to routine intensive cultivation (Paperna, 1980). Snails can only proliferate in mesotrophic ponds with a solid substrate (earth or gravel), holding a low fish biomass, for instance a pond used for spawning, as a nursery or for holding broodstock. Extensive systems; dam reservoirs and similar large water bodies holding lower fish biomass, or smaller units, ponds or pools with frequent or continuous water exchange fringed by trailing and floating weeds; offer better conditions for vector snails and are attractive to piscivorous birds (De Bont and De Bont-Hers, 1952; Lombard, 1968; Paperna, 1968). At times, indoor circulation systems, raceways and hatcheries become heavily populated with snails (Stables & Chappell, 1986), but transmission in these systems is often limited to sanguinicolids (Hoffman et al., 1985). Metacercarial infections are usually prevented where piscivorous birds can be excluded by an efficient netting system. In cultured cichlids in Israel, and tropical and southern Africa, the following massive metacercarial infections, sometimes resulting in mortalities, have been recorded: gill infections of Centrocestus, and subcutaneous Haplorchis transmitted by M. tuberculata (Sommerville, 1982, 1982a; Paperna, 1991), skin Neascus (“black-spot”), muscle infection with Bolbophorus levantinus (Paperna, 1991) and visceral infections of Clinostomum tilapiae and Euclinostomum heterostomum (Lombard, 1968; Britz et al., 1985) transmitted by B. truncatus (Finkelman, 1988). Gill infection by Centrocestus formosanus resulting in mass mortality has been reported from farmed eels (Anguilla japonica) in Japan (Yanohara & Kagei, 1983). Lymnaea (Radix) transmitted C. complanatum, heavily infected farmed loach (Misgurnus anguillicaudatus) and ayu (Plecoglossus altivelis) in Taiwan, causing growth retardation and lower rates of survival (Liu, 1979; Lo et al., 1981). Additional data on species of clinostomids and heterophyiids troubling farmed fish in Southeast Asia are provided by Kabata (1985).
The most practicable preventative method of controlling digenean infection in farmed fish is elimination of the vector snail. Available measures include use of chemical molluscicides, environmental manipulation and use of molluscophagous fish.
Extensive literature exists on the control of snails which are vectors of schistosomes and Fasciola (McCullough & Mott, 1983; Madsen, 1990). Of all the molluscicides developed to control these snails, only copper sulphate is of any practical use in fish ponds and circulation systems. Molluscicidal concentrations of niclosamide (=Bayluscide, Beyer 73) and N- tritylmorpholin (=Frescon, WL 8008, Shell product) currently recommended for snail control are toxic to fish (Cowper, 1971). Copper sulphate (5-hydrate) molluscicide concentrations are tolerated by most fish (although some species, and younger fish may be more susceptible). It is an inexpensive compound, widely used in fish ponds as an algicide (Sarig, 1971), and it can be safely applied at a dose of 3.5 ppm to brackish-water ponds and at 2 ppm to neutral and hard freshwater ponds. However, in acid and soft freshwaters (pH 6.8, calcium ions >12 ppm) the same or even lower concentrations become toxic to fish. Copper salt may be applied by continuous dosing at a lower concentration (1 ppm), or as a low-soluble formulation (as copper carbonate or copper oxide) to produce long term residual effects. The safety of this compound to fish has been demonstrated under these conditions (Hoffman, 1970). Treatment of drained ponds or raceways by copper sulphate, prior to stocking, delayed but did not prevent repopulation by snails (Stables and Chappell, 1986).
The environmental limits imposed on snail survival in fish farm systems are discussed above. Regular weed control, performed manually, or with herbicides (Paperna, 1980) can decimate snail populations. Of all listed and recommended molluscophagous fish (DeBont and DeBont Hers, 1952; Carothers & Allison, 1968), only black carp (Myelopharyngodon piceus) was routinely employed (with mixed success) in water supply lakes (Leventer, 1979). Experience with commercial fish farms is still insufficient.
Worm infection control:
Praziquantel (Biltricide[R], Bayer AG, Germany) has been shown to be effective against digeneans and cestodes of men and animals, as well as being safe (Andrews et al., 1983). Preliminary trials demonstrated praziquantel's parasiticidal effect on Diplostomum spathaceum metacercariae in rainbow trout fed on medicated feed (Bylund & Sumari, 1981). This was followed by Szekely and Molnar's (1991) report on the elimination of all D. spathaceum metacercariae from herbivorous carp. Recommended application is by feeding a single dose of 300 mg kg-1 body mass. Three sequential lower doses of 35–100 mg kg-1 yielded 88–100% efficacy, and bath treatments of 1 mg l-1 for over 9hrs to 10 mg l-1 for 1hr showed 100% and 93–94% efficiency, respectively. Mr N. Kraus, manager of Kibbutz Hamaapil, Israel, fish farm, used a veterinary formulation of praziquantel (Droncit) to kill off metacercariae of Centrocestus, Haplorchis and Bolbophorus levantinus in juvenile tilapia (70 mm in length). Dissolved praziquantel in dip tanks was found to retain its therapeutic efficiency and may therefore be reused for over a month. In spite of its promising therapeutic qualities, praziquantel's use in fish farms is uneconomic due to its high price, except in very special circumstances such as high-priced ornamental fish, breeders or valuable genetic stock.
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Plate 22 Trematoda: a,b. Sanguinicola infection-eggs a. in gills of Mugil cephalus, S. Africa; b. in heart of Oreochromis aureus. c–e: Juvenile cichlids (Tristramella sacra and Tilapia zillii) naturally infected with Neascus (Black spot) (fine arrows) and Bolbophorus levantinus, Israel; e, skinned fish to demonstrate B. levantinus infection in muscles (thick arrows). f–i: infection with B. levantinus in T. sacra; f, early and g, late metacercariae, live; h. histological section showing inflammatory process on the periphery of the encysted metacercaria in the muscles and i, monocytes -macrophages infiltrating around the unencysted metacercaria. j,k. Neascus (blackspot) metacercaria (a Postdiplostomum) on skin of O. aureus x niloticus, Israel.
Plate 23. Trematoda continued: a. Tilapia zillii from Israel, with Clinostomum cutaneum encysted under scales. b. Centrocestus infection in gills of T. zillii, Israel. c. Pygidiopsis metacercariae on the gut wall (Killifish, USA). d. Histological view of Centrocestus metacercariae on gills of O. aureus, Israel. e. Melanoides tuberculata, L. Kinneret, Israel. f. Physopsis globosus, Ghana; g. Bulinus truncatus, Ghana. h. Ascocotyle metacercaria from heart of Liza ramada, Israel. i. Pygidiopsis metacercaria on the gut wall (see c) k. Heterophyes heterophyes in muscles of L. ramada, Sinai lagoons. l. An heterophyiid metacercaria in the liver of L. aurata, Israel.
Fig. 4. Trematoda: A. Allocreadium ghanensis, adult (3 mm long) from the intestine of Synodontis sp. (After Fischthal & Thomas, 1972). B. Sanguinicola dentata, adult, from kidney circulatory system of Clarias lazera; d, vas deferens; i, intestine; l, vagina (non functional); o, ovary; od, oviduct; ot, ootyp; t, testes; u, uterus; v, vitellaria. C. Metacercaria of Pygidiopsis genata. D. Met. of Phagicola longa. E. Met. of Heterophyes heterophyes. F. Gonotyle of H. aequalis. G. Gonotyle of H. heterophyes. H. Met. of Stictodira. I. Gonotyle of S. sawakiensis. J. Redia of heterophyiid. K. Cercaria of heterophyiid. L. Head of Phagicola italica. M. Head of Centrocestus spp. N. Head of echinostomatid metacercaria. O. Sporocyst of diplostomatids. P. Furcocercaria of diplostomatids. Q. Diplostomulum (Diplostomum spp. met.). R,S. Young and developed Neascus (Met. of Postdiplostomum and Ornithodiplostomum spp.). T. Met. of Clinostomum tilapiae (length 4–8 mm). U. Met. of Euclinostomum heterostomum (length 6–8 mm). V. Met. of Clinostomum sp. (“cutaneum”) beneath the scales of cichlid fish (see Pl. 23a) (length 6 mm).
Plate 22. Trematoda (legend p. 139).
Plate 23. Trematoda continued (legend p. 139).
Fig. 4. Trematoda (legend p. 139).