Fig. 7 (p. 175)
Species affected and geographical range
Present in representatives of diverse African fish families (Golvan, 1957, 1965; Khalil, 1971). The geographic range of these parasites is sometimes narrower than that of their specific hosts, for example, the cichlid parasite Acanthogyrus tilapiae is widespread in tropical Africa including Madagascar (introduced), but it has not yet been found in the Near East, the Sudan Nile or South African cichlids (Paperna, 1964; Khalil, 1969; Van As & Basson, 1983).
Description taxonomy and diagnosis
Acanthocephala are readily recognised by their evaginable proboscis crowned with several rows of recurved hooks. In the encased larval stage, in tissues, the spiny proboscis is retracted. The worms are sac-like, containing lemnisci connected to the proboscis and genital organs opening posteriorly. The sexes are separate and the male opening is within a membranous bursa. An alimentary canal is absent.
The number and arrangement of the hooks on the proboscis are the main criteria for differentiation of species. A wider range of anatomical details are considered for determination of higher taxa (Kabata, 1985).
Incubation in tap water helps to extract the proboscis prior to fixation in hot or cold alcohol 70%, formol saline 4%, or AFA.
Life history and biology
All acanthocephalans develop via one or more intermediate hosts (heteroxenous). Adult acanthocephalans are all gut parasites. Eggs are laid into the intestinal lumen and evacuated with faeces. First intermediate hosts of piscine acanthocephala are amphipods, isopods, copepods or ostracods. The first larvae, the acanthella (acanthor), hatch from eggs after being swallowed by a suitable invertebrate host. Some species will develop to the adult stage when their larvae in the invertebrate host are ingested by the definitive vertebrate host (George & Nadakal, 1973; Schmidt, 1985). Fish can also serve as intermediate hosts, harbouring a second larval stage (the acanthor or cystacanth). Definitive hosts of such acanthocephalans are either predatory fish or piscivorous birds (Hassan & Qasim, 1960; Hoffman, 1967).
Life histories and intermediate hosts of acanthocephala of African fish are at present unknown.
Pathogenic effects of acanthocephalans are due to attachment of the adult parasite in the digestive tract and also to the encapsulation of larval stages in the tissues. In low to moderate infections, pathological effects are localised around the attachment of the adult worm. The extent of damage is proportional to the depth of penetration of the proboscis. It is negligible when parasites are attached to the epithelial mucosa only (Acanthogyrus and Acanthocephalus spp.), and becomes extreme, with extensive granuloma and subsequent fibrosis, when the worm's proboscis is anchored in the muscle layer or entirely perforates the intestinal wall (Pomphorhynchus spp.) (Paperna & Zwerner, 1976; McDonough & Gleason, 1981; Kabata, 1985). The depth of penetration of some species, may vary in different host fish (Taraschewski, 1988). Extensive inflammation, peritonitis due to perforation of the gut and systemic clinical changes (anurhersia) will occur only in massive infections, most often in farmed fish (Bullock, 1963; Bauer, 1959). In juvenile fish (cichlids <60 mm long) a single attached specimen of Acanthogyrus tilapiae obstructed the digestive tube, apparently with no clinical implications (Douellou, 1992 a,b). Low to moderate infections with larval stages (cystacanths) in visceral organs (liver, spleen) caused only local changes while heavy infection, in juvenile fish in particular, led to extensive granuloma, fibrosis and ultimately atrophy through fibrosis of either a portion of or the entire organ (Paperna & Zwerner, 1976). Information on infection among fish in Africa is very limited and none of the conditions described above have ever been reported.
Host specificity of acanthocephalans is variable and may be evaluated only where sufficient data are available, which is not the case for most African fish species. Acanthogyrus tilapiae is specific to Cichlidae, while other species have been found in Cyprinidae, Paragorgorhynchus albertianum is indiscrminate in its choice of hosts (Khalil, 1971). Epizootiological data are limited to natural infections: In the Sudan White and Blue Nile, 5–27 Tenuisentis niloticus occurred in 93% of Heterotis niloticus, 6–43 Neoechinorhynchus sp. in 26% of Citharinus citharus and 2–5 unidentified acanthocephala in 60% of Synodontis batensoda (Khalil, 1969). Similarly abundant infections were found in the same fish in L. Chad (Troncy, 1974, 1977; Troncy & Vassilides, 1973). Acanthogyrus tilapiae is a fairly common parasite of cichlid fish in tropical Africa, in Lake Kariba, 63% of the Tilapia rendalli, and all four Oreochromis mortimeri examined harboured worms, of which, one specimen had over 100 worms (Douellou, 1992a,b).
To control infections in coldwater fish farms, medicated feed with Bithionol (2.2-thio bis (4,6-dichlorophenol), is recommended, at a dose of 0.2 g/kg fish (Hoffman, 1983). Feeds medicated with Di-N-butyl tin oxide are also potentially effective (see 14.1).
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ILLUSTRATIONS - Fig. 7 page 175 with legend
Fig.7. Acanthocephala and various other parasites of fish. A. Paragorgorhynchus chariensis, male, 10–11 mm long. B,C. Pallisentis tetraodontis, female, 4.5 mm long, proboscis and whole view (A–C, after Troncy, 1977). D. Larval stages of acanthocephalans: 1. egg (of Neoechinorhynchus, 60×25 μm); 2. Acanthella from Gammarus amphipod, 1–4 mm long; 3. Acanthella from ostracods (2–4 mm long). 4. Cystacanthus (Acanthor) from fish (3–6 mm long). E. Piscicolid leeches (Hirudinea) (80–100 mm). F. Pentastomid larva. G. Parasitic larva of mutelid bivalve (after Fryer, 1970). H. Unionid glochidium embedded in the gill tissue.