Plates 8–11, 12d and Fig. 1 (pp. 52–57)
Host specific species are associated with a wide range of fish species from most families. Ubiquitous or opportunistic species (Ichthyobodo necator, Chilodonella spp., and some species of Trichodina, Ambyphrya and Scopulata (Scyphidia) are particularly common in juvenile cichlids and carp.
The ubiquitous ectoprotozoans are cosmopolitan or trans-continentally dispersed via translocation of their cultured fish hosts (carp and tilapia in particular) (Ichthyobodo necator, Cryptobia branchialis, Chilodonella hexasticha, C. piscicola, Trichodina acuta, T. heterodentata and T. pediculus, T. reticulata, T. mutabilis, T. centrostrigata, Trichodinella epizootica, - Hoffman, 1978; Basson et al., 1983; Van As & Basson, 1987, 1989; Natividad et al., 1986; Shaharom-Harrison & Abdullah, 1988; Albaladejo & Arthur, 1989; Bondad-Reantaso & Arthur, 1989; Basson & Van As, 1993).
Distribution of the more specialised host-specific species follows that of their hosts, but may also be more restricted, sometimes to only one or a few watersheds. There is evidence for the presence of Ichthyobodo, Chilodonella, and in particular trichodinids and sessilians in a number of water systems in tropical Africa (Lake Volta and East African lake systems — Fryer, 1961; Paperna, 1968; Paperna & Thurston, 1968; Fryer & Iles, 1972), but taxonomic data are limited to only a few locations (Kazubski & El Tantawy, 1986; El Tantawy & Kazubski, 1986). The most comprehensive taxonomic data on trichodinids in African fish are from Southern Africa and the Zambezi river system. Data are also available from Israeli fish (Basson et al., 1983; Basson & Van As, 1987; Van As & Basson, 1989, 1992).
Description taxonomy and diagnosis
Integumental ectoprotozoan genera are readily differentiated (Kabata, 1985; Dykova & Lom, 1992), while diagnosis of species is difficult and often requires special staining.
Most ectoparasitic forms are readily detected in direct microscopic examination of skin and gill scrapings from live (or freshly killed) fish.
Flagellates may be further detected in air dried, methanol fixed, Giemsa stained smears.
Smears containing ciliates should be air dried, fixed in Bouin for 20 min., destained in 70% ethanols, brought to water, stained in a haematoxylin stain and mounted after dehydration.
Trichodinids for specific differentiation should be impregnated with silver. Air dried smears should be placed in 2% silver nitrate for 7–9 min. in the dark, rinsed in water and exposed to the sun or UV for 5–10 min.
Flagellates [Mastigophora, Kinetoplastida]:
|Cryptobia||free, spindle shaped, 10–30 × 3–5 μm in size (if C. branchialis), or pyriform when attached to the integument, with two flagellae, one wholly or partly adjunct to the body, kinetoplast rod-shaped or round.|
|Ichthyobodo||free, 13–26 × 2–7 μm in size, or attached to the integument; with four flagellae.|
|Chilodonella||rounded to oval, cytostome distinct, macronucleus round and cilia on the concave ventral surface are arranged in several concave parallel rows:|
|C. hexasticha||size 30–65 × 20–50 μm, with 6–8 ciliary lines on each side.|
|C. piscicola (syn: C. cyprini) size 33–100 × 24–60 μm, with more than 10 ciliary lines on each side.|
|Trichodina||cup shaped, 20–100 μm in diameter with concentric rows of cilia and a crown of denticles. The denticle shape is a distinct taxonomic feature; for differential specific diagnosis of African spp. see Basson et al., 1983; Basson & Van As, 1987; Van As, & Basson, 1989, 1992.|
Small trichodinids, predominantly from the gills, are bell shaped (Tripartiella and Paratrichodina) and often settle on the tips of the gill lamellae (see Basson & Van As, 1989, for generic division of Trichodinidae). In Trichodinella, the ray (the inner extension of the denticle) (Van As & Basson, 1989), is totally reduced, and in the other two genera is delicate or rudimentary (Kazubski & El Tantawy, 1986; Basson & Van As 1987).
Genera of sessile peritrichs are differentiated by their macronuclei and scopula (attachment leg) (Viljoen & Van As, 1983, 1985):
Scopulata (Scyphidia) round macronucleus and wide scopula;
Apiosoma (Glossatella) pyriform nucleus, small scopula;
Ambyphrya Ribbon shaped macronucleus and wide scopula.
Stalked sessile peritrichs — Heteropolaria with elongate body and curled macronucleus (Foissner et al., 1985); Epistilis cup-shaped with horseshoe-shaped macronucleus (Viljoen & Van As, 1983). Some Apiosoma also develop on stalks.
Suctoria: (Trichophyra and other genera) — cilia lacking, variable numbers of tentacles arise from the rounded body.
Life cycle and biology
Most ectoprotozoans, flagellates as well as ciliates have simple life histories. Species of Cryptobia are ectoparasites as well as intestinal and vascular parasites. It has been shown that an ectoparasitic phase occurs in two vascular species (Woo, 1987). Both ectoparasitic flagellates, I. necator and Cryptobia spp., occur either free swimming or attached to the integument, the former through a cytoplasmic protrusion (Schubert, 1968) and the latter by attachment with the flagellum (Lom, 1980).
Reproduction is usually by binary fission. Conjugation is sometimes observed in ciliates. Sessile species also bud and give birth to a free swimming mobile generation, reminiscent of mobile peritrichs, which settle on suitable substrates (fish).
The sessile suctorians reproduce by internal and external budding, the detached buds are ciliated. As the buds become attached to a new location on the piscine integument, cilia are shed and tentacles appear (Hoffman, 1978).
Spores or other forms of waiting stages are unknown; the suggested existence of waiting stages, such as encysted forms of Chilodonella hexasticha in the gills (Rowland et al., 1991) or free cysts (Bauer et al., 1969), has to be confirmed.
Water temperatures do not seem to be an important parameter, in spite of reports of low temperatures being more optimal for reproduction of Chilodonella piscicola and some trichodinids (Bauer et al., 1969). Massive infections with I. necator, both species of Chilodonella and the ubiquitous trichodinids and sessile species, occur in low (12–17°C) and high (25–30°C) ambient temperatures in southern Africa and Israel. Most freshwater ectoparasitic protozoans disappear in ponds with increased salinities (above 2000 ppm chlorinity), only I. necator and some Cryptobia are tolerant and become the predominant parasites in fish of such ponds. There are also halophilic species of Ambyphrya and Scyphidia which infect fish (grey mullet) in estuaries.
A number of ciliates (species of Tetrahymena, Ophryoglena, Glaucoma, Colpidium and others — see Hoffman, 1978) are facultative parasites, or opportunists which will colonise fish in special circumstances, most often when fish are stressed or traumatised (Hoffman, 1978). All others mentioned above are obligatory parasites which will apparently survive for only a limited time outside their hosts. Non-parasitic sessile peritrichs are different species from those colonising living organisms. Trichodinids and sessile species found on aquatic invertebrates comprise different species from those infecting fish (Van As & Basson, 1987; Viljoen & Van As, 1983, 1985). There are, however, a few documented exceptions: T. pediculus being reported from both hydra and fish, and T. diaptomi a parasite of a calanoid copepod, which temporarily invaded hatchery grown fry of Clarias gariepinus (Basson et al., 1983; Basson & Van As, 1991).
There are several degrees of adaptation of trichodinids to their piscine hosts: ubiquitous species, of an opportunistic nature, which are always found on the fish skin but never on the gills (T. pediculus and T. acuta); other ubiquitous species occur both on gills and skin (T. heterodentata); additional, seemingly ubiquitous, widespread species appear to have a variable degree of predilection for one fish family or another (cichlids or cyprinids).
Among the latter, trichodinids with seemingly related morphological characteristics (e.g. pediculus-like, acuta-like and nigra-like), in different geographical regions, demonstrate definite affinities to a particular group of hosts and may in fact comprise diverse species (Van As & Basson, 1989). Host specific trichodinids, are all, with only a few exceptions, gill parasites: T. centrostrigata and great numbers of small trichodinids mainly species of Tripartiella, are associated with Cichlidae; T. reticulata occurs mainly in goldfish, T. kazubski has been found in South African Barbus spp. and T. nobilis and T. kupermani mainly in asian carp (Basson et al., 1983; Van As & Basson, 1987, 1989; Abaladejo & Arthur, 1989).
Ectoparasitic protozoa are variable in their effect on their hosts. Pathological effects are density dependent, when both the size of the parasite population and the nature of the tissue responses are modulated by the physiological (clinical) condition of the fish. Hostile environments (stressful conditions) compromise the fishes' capacity to counteract infection.
Ichthyobodo necator attaches itself to epithelial cells and through an inserted protrusion consumes their contents (Schubert, 1968), whereas Chilodonella spp. browse the epithelial surface (Paperna & Van As, 1983). Histopathological changes in the integument following infection by Chilodonella spp. and I. necator are an outcome of two counteracting cellular processes — hyperplasia of the epithelial cells, including mucus cells and chloride cells, versus a progressive cellular destruction. Cellular destruction primarily occurs due to direct action of the parasites, and later by enhanced abrasion of the peripheral cells after the depletion of mucus forming cells. The production of mucus cells is limited. Accelerated mucus cell production stimulated by the infection apparently exhausts resources for mucus production, and the infected fish become “dry”. Some parasites seem to yield cytotoxins or proteolytic enzymes which could be the cause of spongiosis, which affects both the proliferated and unchanged epithelial layer (Robertson et al., 1981; Paperna & Van As, 1983). Secondary cellular damage due to degeneration, necrosis and desquamation results in the degradation and disintegration of the epithelial layer.
Cryptobia attachment through the flagellum does not induce any pathological or even ultrastructural cellular damage (Lom, 1980), contrary to reports of morbidities associated with this parasite (Woo, 1987).
Although there are a number of reports on poor condition and mortalities, particularly of fry, coinciding with massive infestation of trichodinids, Trichodinella epizootica in particular (Lom, 1973), and the sessilians Apiosoma, Ambyphrya and Scopulata (Fijan, 1961; Meyer, 1970; Paperna et al., 1984; Lightner et al., 1988; Paperna 1991), histopathological changes in events of massive infections by these ectoprotozoans are hardly evident, if occurring at all (Fitzgerald et al., 1982; Paperna, unpublished 1985). Trichodinella epizootica in carp (Lom, 1973) and Tripartiella cichlidarum in cichlids (Paperna, 1991) cause some erosion of the gill epithelium. However, food vacuoles of trichodinids revealed bacteria rather than sloughed cells (Paperna, unpublished). Ultrastructural observation on attached Apiosoma did not reveal any interference with the host cell serving as substrate (Lom & Corliss, 1968; Lom, 1973; Fitzgerald et al., 1982) or peripheral tissue response. Thus, mortalities following massive colonisation of gills by sessilians (Fijan, 1961) could result from the dense cover of sessilians disrupting gas exchange through the respiratory epithelium. The only exception among these infections are the colonies of the stalked sessilia Heteropolaria (Epistilis) which cause lesions (“red sore”) at the stalk attachment to the fish skin, these inflamed haemorrhagic lesions are also contaminated with the bacterium Aeromonas hydrophila (Esch et al., 1976; Miller & Chapman, 1976). Reported localised infection above the opercular bone (in cultured tilapia in Israel) resulted in aggravation of the lesion into a wide (6 mm in diam.) perforation of the bone (Paperna, 1991).
Suctorians (Trichophyra spp.) in certain instances cause cytological damage to the gill lamellae cells in direct contact with the parasites and subsequent hyperplasia and haemorrhages of the gill tissue (Heckmann & Caroll, 1985).
The course of infection by ectoparasitic protozoans is determined either individually or by the interaction of the following factors:
mobility of the fish.
the fish's capacity to activate its defence systems.
Reduced mobility facilitates parasite colonisation as well as moderating loss through detachment and drift from the integumental surface.
Defence mechanisms other than epithelial hyperplasia, and specific immune responses to integumental ectoparasites have not yet been studied (except in I. multifiliis, see pp. 61–62), although spontaneous recovery from infection has been frequently observed. Juvenile fish and fish under stress (and at below optimum ambient temperature) have both limited mobility and apparently immunological incompatibility, being either naive or immunosuppressed (Sniezko, 1964; Avtalion, 1981).
Heavy infections by ectoparasitic protozoans are mainly found in young fish (less than one year old) when overcrowded and confined to restricted habitats, and under stress conditions. In these circumstances opportunistic and ubiquitous species are involved. Infections otherwise, in grown-up fish, are very low and host-specific species predominate.
New born cichlids, as soon as they were weaned from parental care, and sometimes before, became heavily infected by trichodinids and sessilians of the genera Ambyphrya and Scopulata. Infestation reached its climax level in fish 10–12 mm long. Such infections occur in natural habitats (lakes), man-made impoundments, as well as in hatchery installations (Fryer, 1961; Paperna et al., 1984; Paperna, unpublished report 1985). Heavy infections, however, were not found in all the breeding habitats of the investigated lake system. Conditions for infestation varied with habitat and ambient conditions and were positively related to the abundance of fry schools.
Level of infection in the fry sharply declined as fish gained in size (Paperna, unpublished report). The decline in infection also coincided with changes in parasite species composition, the ubiquitous, generalists and opportunists (T. pediculus [=? T. migala], T. acuta [=?T. compacta, see Basson & Van As, 1989], T. heterodentata, and species of Ambyphrya and Scopularia) being gradually replaced by species specific to cichlids (Tripartiella spp. Trichodina centrostrigata and species of Apiosoma) (Basson et al., 1983; Kazubski & El Tantawy, 1986; El Tantawy & Kazubski, 1986; Basson & Van As, 1987; Van As & Basson, 1989).
Heavy infections with ubiquitous trichodinids (T. pediculus) and sessile peritrichs (mainly Scopulata spp.) also occur in carp fry in hatcheries and nursery ponds, and likewise as fish grow, are replaced by more specialised species (such as T. nigra, T. mutabilis and Apiosoma spp.) (Basson et al., 1983; Shaharom-Harrison & Abdullah, 1988; Albaladejo & Arthur, 1989).
Heavy infections (by trichodinids and sessile species) accumulate in fish - small spp. of Barbus, Alestes, cyprinodontids and juvenile cichlids and Clarias spp. crowded into residual pools in rivers drying-out during the dry season. In larger water bodies in Africa, infections with both trichodines and sessilians in fish other than fry may be common but low (Paperna, 1968; Viljoen & Van As, 1985; Kazubski, 1986).
Low temperature stress plays an important role in epizootic outbreaks of ectoprotozoan infections in cichlid fish outside the limits of the tropical environment and of populations introduced to non-tropical countries such as the southern USA.
Heavy infections by skin and gill protozoa, predominantly of Chilodonella spp., are a frequent occurrence in overwintering stocks of cultured tilapia hybrids (Oreochromis aureus x niloticus) in Israel, and O. mossambicus in ponds and dam reservoirs in southern Africa (Du Plessis, 1952; Oldewage & Van As, 1987; Paperna et al. 1983; Paperna, 1984).
In small ponds (1 hectare) fish are not spared even in relatively mild winters, with minimum temperatures above 13°C. Fish in lakes and large reservoirs on the other hand, become severely affected only in extremely cold winters, with temperatures declining to 10°C and below. Mortalities often occur from the cumulative effects of ectoprotozoans, dermal saprolegniases and systemic bacterial diseases, all mediated by the stress of low temperature.
In addition to temperature stress, overwintering tilapia in ponds are often stressed by overcrowded stocking and inadequate feeding. Intermittence of higher and lower ambient temperatures, characteristic of the Mediterranean type winters, increases the unpredictability of food demand by fish and thus complicates feeding schedules.
Infestation levels rise by late fall, with increased abundance of trichodinids and C. hexasticha. Fish succumbing in early winter were predominantly hyperinfected by C. hexasticha. Late winter and early spring mass mortalities (even when temperatures were already rising above 15°C) were associated with C. piscicola hyperinfections. C. piscicola is abundant in carp in some ponds already by early winter, however, it will only infect tilapia at the end of the cold season when they become compromised by prolonged stress.
Ichthyobodo necator hyperinfections are morbid to cichlids as well as to fish of other families. Mortalities occur in fish overcrowded in holding tanks, ponds and in both warm and cold water conditions. Natural infection was also revealed in Aplocheilichthys gambiae from a pool in Ghana. In Israel Cryptobia spp. occasionally swarm the gills of tilapia, goldfish and silver carp and also, in the latter, in low saline waters (8–10 ppt salt) but data from Africa are lacking.
O. mossambicus appears to be more tolerant to low temperatures in water of higher salinities, and also where most ectoparasites are excluded (except l. necator). Members of the genus Tilapia, in Israel (T. zillii) and in southern Africa (T. rendalli & T. sparmanii) are also less affected in freshwaters by low temperatures and are rarely heavily parasitised.
Few instances of mortalities coincided with heavy infections, concomitantly or exclusively, by trichodinids, sessilians (Apiosoma), Chilodonella spp. and I. necator, in overwintering carp, but occurred more often in relation to other stress factors such as high levels of overcrowding or high nitrite concentrations (Fijan, 1961; Sarig, 1971; Paperna, unpublished 1985).
Heavy infections by Chilodonella spp. seems to have an excluding effect on other integumental protozoans. Otherwise, skin and gill ectoparasites coexist, and are even synergistic, with metazoan ectoparasites (Gyrodactylus and Argulus) and skin lesions (epithelioma), (Paperna & Kohn, 1964; Sarig, 1971; Paperna, unpublished 1985). Mass mortalities of farmed Clarias gariepinus (in the Central African Republic) were associated with mass infestation by Chilodonella hexasticha.
Epistilis infections, including red sore and opercular perforations only occur sporadically with no particular link to overwintering.
Treatment with formalin is still the only effective means to control massive ectoparasitic infections in all warm water cultured fish species. In Israeli fish farms, ponds are sprayed with formalin up to concentrations of 25 or 40 ppm (of the 37% commercial product) (Sarig, 1970; Lahav & Sarig, 1972). Efficacy of formalin treatments is affected by ambient temperatures, water quality, including salinities and parasites treated. Product quality is variable, and is particularly affected by storage, resulting in accumulation of polymerised (paraformaldehyde) sediment. Trichodinids were readily eradicated with treatment by 25 ppm, while elimination of Chilodonella was achieved after treatment with 40–50 ppm. Van As et al. (1984) also demonstrated differential efficacy with the type of fish treated, e.g. 25 ppm per 24h was effective in cleaning infected carp, while with tilapia fry it has been achieved with 45 ppm per 24h.
Albaladejo, J.D. & Arthur, J.R., 1989. Some trichodinids (Protozoa: Ciliophora: Pertrichida) from freshwater fishes imported into the Philippines. Asian Fisheries Science, 3: 1–25.
Avtalion, R., 1981. Environmental control of the immune response in fish. CRC Crit. Rev. Environ. Control., 11: 163–188.
Basson, L. & Van As, J.G., 1987. Trichodinid (Ciliophora; Peritricha) gill parasites of freshwater fish in South Africa. Sys. Parasitol., 9, 143–151.
Basson, L. & Van As, J.G., 1989. Differential diagnosis of the genera in the family Trichodinidae (Ciliophora: Peritrichida) with description of a new genus ectoparasitic on freshwater fish from southern Africa. Sys. Parasitol., 13: 153–160.
Basson, L. & Van As J.G., 1991. Trichodinids (Ciliophora: Peritrichia) from a calanoid copepod and catfish from South Africa and notes on host specificity. Sys. Parasitol., 18: 147–158.
Basson, L. & Van As, J.G., 1993. First record of European trichodinids (Ciliophora: Peritrichida), Trichodina acuta Lom, 1961 and T. reticulata Hirschmann et Partsch, 1955 in South Africa. Acta Protozool. 32: 101–105.
Basson, L., Van As, J.G. & Paperna, I., 1983. Trichodinid parasites of cichlids and cyprinid fishes of South Africa and Israel. Sys. Parasitol., 5: 245–257.
Bauer, O.N., Musselius, V.A. & Strelkov, Yu. A., 1969. Diseases of Pond Fishes. Publisher “Kolos” Moskva. In English: Israel Program for Scientific Translations, Jerusalem, 1973.
Bondad-Reantaso, M.G. & Arthur, J.R., 1989. Trichodines (Protozoa: Ciliophora: Peritrichida) of Nile tilapia (Oreochromis niloticus) in the Philippines. Asian Fisheries Science, 3: 27–44.
Du Plessis, S.S., 1952. Fish diseases in Transvaal. C.S.A. Symp. Hydrobiol. and Inland Fish., Entebbe 37: 128–130.
El-Tantawy, S.A.M. & Kazubski, S.L., 1986. The trichodinid ciliates from fish, Tilapia nilotica from the nile delta. Acta Protozool., 25: 439–444.
Esch, G.H., Hazen, T.C., Dimock, R.V. & Gibbons, J.W., 1976. Thermal effluent and the epizootiology of the ciliate Epistilis and the bacterium Aeromonas in association with centrachid fish. Trans. Am. Microsc. Soc., 95: 687–693.
Fijan, N., 1961. Massive invasion of carp fry (Cyprinus carpio L.) by protozoan of the genus Glossatella (Croatian text, English summary). Vet. Arth. Zagreb, 32: 30–33.
Fitzgerald, M.E., Simco, B.A. & Coons, L.B., 1982. Ultrastructure of the peritrich ciliate Ambyphrya ameiuri and its attachment to the gills of the catfish Ictalurus punctatus. J. Protozool., 29: 213–217.
Fryer, G., 1961. Observation on the biology of the cichlid fish Tilapia variabilis Boulenger in the northern waters of Lake Victoria and the Victoria Nile. Rev. Zool. Bot. Afr., 64: 1–33.
Fryer, G. & Iles, T.D., 1972. The cichlid fishes of the great lakes of Africa. T.F.H. Publications, Neptune City, N.J.
Foissner, W., Hoffman, G.L. & Mitchell, A.J., 1985. Heteropolaria colisarum Foissner & Schubert, 1977 (Protozoa: Epistylididae) of North American freshwater fishes. J. Fish Dis., 8: 145–160.
Heckmann, R.A. & Carroll, T., 1985. Host-parasite studies of Trichophyra infesting cutthroat trout (Salmo clarki) and longnose suckers (Catastomus catastomus) from Yellowstone lake Wyoming. Great Basin Nat., 45: 255–265.
Hoffman, G.L., 1978. Ciliates of freshwater fishes. In: Kreier, J.P. (ed.) Parasitic Protozoa, II. pp. 584–632.
Kabata, Z., 1985. Parasites and diseases of fish cultured in the tropics. Taylor & Francis. London and Philadelphia.
Kazubski, S.L., 1986. The trichodinid ciliates from fish, Tilapia sp. from Lake Victoria (Kenya) and description of Trichodina equatorialis nom. nov. Acta Protozool., 25: 445–448.
Kazubski, S.L. & El-Tantawy, S.A.M., 1986. The ciliate Paratrichodina africana sp. n. (Peritricha, Trichodinidae) from Tilapia fish (Cichlidae) from Africa. Acta Protozoologica, 25: 433–438.
Lahav, M. & Sarig, S., 1972. Control of unicellular parasite fauna. Acta Parasitol. Pol., 22: 49–66.
Lightner, D., Redman, R., Mahoney, L., Dickerson, G. & Fitzsimmons, K., 1988. Major diseases encountered in controlled environment culture of tilapias in fresh water and brackish water over a three year period in Arizona. In: Pullin, R.S.V., Bhukaswan, T., Tonguthai, K. & Maclean J.L. (eds.) The Second International Symposium on Tilapia in Aquaculture. ICLARM Conference Proceedings 15, Dep. Fish. Bangkok, Thailand, ICLARM Manila, Philippines, pp. 111–116.
Lom, J., 1973. The mode of attachment and relationship to the host of Apiosoma piscicola Blanchard and Epistilis Iwoffi Faure-Fremiet, ectocommensals of freshwater fish. Folia Parasitol. (Praha), 20: 105–112.
Lom, J., 1980. Cryptobia branchialis Nie from fish gills: ultrastructural evidence of ectocommensal function. J. Fish Dis., 3: 427–436.
Lom J. & Corliss, J.D., 1968. Observation on the fine structure of two species of the peritrich ciliate genus Scyphidia and on their mode of attachment to their host. Trans. Am. Microsc. Soc., 87: 493–509.
Miller, R.W. & Chapman, W. R., 1976. Epistilis and Aeromonas hydrophila infections in fishes from North Carolina reservoirs. Prog. Fish Cult. 38, 165–168.
Meyer, F. P., 1970. Seasonal fluctuations in the incidence of diseases on fish farms. Spec. Publ. Am. Fish. Soc., 5: 125–128.
Natividad, J.M., Bondad-Reantaso, M.G. & Arthur, J.R., 1986. Parasites of Nile tilapia (Oreochromis niloticus) in the Philippines. In: Maclean, L.J., Dizon, L.B. & Hosillos, L.V. (eds.) The First Asian Fisheries Forum. Asian Fisheries Society, Manila, Philippines. pp. 225–259.
Oldewage, W.H. & Van As, J.G., 1987. Parasites and winter mortalities of Oreochromis mossambica. S. Afr. J. Wildl. Res., 17: 7–12.
Overstreet, R.M. & Howse, H.D., 1977. Some parasites and diseases of estuarine fishes in polluted habitats of Mississippi. Ann. N.Y. Acad. Sci., 298: 427–462.
Paperna, I., 1968. Ectoparasitic infections of fish of Volta lake, Ghana. Bull. Wildl. Dis. Ass., 4: 135–137.
Paperna, I., 1984. Winter diseases of cultured tilapia. In: Agrigrup (ed.) Fish Diseases. Fourth COPRAQ session. Editora ATP Madrid (Espanha) pp. 139–147.
Paperna, I., 1985. Infectious diseases of fish in extreme environment. Final report to the Ministry for Science and Art of Lower Saxony, Germany - unpublished.
Paperna, I. & Kohn, A., 1964. Studies on the host-parasite relations between carp and populations of protozoa and monogenetic trematodes in mixed infestations. Rev. Brazil. Biol., 24: 269–276.
Paperna, I. & Thurston, J.P., 1968. Report on ectoparasitic infections of freshwater fish in Africa. Bull. Off. int. Epizoot., 69: 1192–1206.
Paperna, I. & Van As J.G., 1983. Epizootiology and pathology of Chilodonella hexasticha (Kiernik, 1909) (Protozoa, Ciliata) infections in cultured cichlid fishes. J. Fish Biol., 23:441–450.
Paperna, I., Van As J.G. & Basson L., 1984. Review of diseases affecting cultured cichlids. In: Fishelson, L. & Yaron, Z. (ed.) International Symposium on Tilapia in Aquaculture, Proceedings. Tel Aviv University Press. pp. 174–184.
Robertson, D.A., Roberts, R.J., & Bullock, A.M., 1981. Pathogenesis and autoradiographic studies of the epidermis of salmonids infested with Ichthyobodo nacator (Henneguy, 1883). J. Fish Dis., 4: 113–125.
Rowland, S.J., Ingram, B.A. & Prokop, F.B., 1991. Suspected cysts of the protozoan parasite Chilodonella hexasticha. Bull. Eur. Ass. Fish Pathol., 11: 159–161.
Sarig, S., 1971. The prevention and treatment of diseases of warmwater fish under subtropical conditions, with special emphasis on intensive fish farming. T.F.H Publications Inc., Jersey City, N.J. 127 p.
Schubert, G., 1968. The injurious effects of Costia necatrix. Bull. Off. int. Epiz., 69: 1171–1178.
Shaharom-Harrison, F. & Abdullah, S.Z., 1988. Study of trichodinid ectoparasites from bighead carp Aristichthys nobilis, grass carp Ctenopharyngodon idella and lampam jawa Puntius gonionotus in peninsular Malaya. Trop. Biomed., 5: 131–137.
Sniezko, S.F., 1964. The effect of environmental stress on outbreaks of infectious diseases of fishes. J. Fish. Biol., 6: 197–208.
Van As, J.G. & Basson, L., 1987. Host specificity of trichodinid ectoparasites of freshwater fish. Parasitol. today, 3: 88–90.
Van As, J.G. & Basson, L., 1989. A further contribution to the taxonomy of the trichodinidae (Ciliophora: Peritricha) and a review of the taxonomic status of some fish ectoparasitic trichodines. Sys. Parasitol., 14: 157–179.
Van As, J.G. & Basson, L., 1992. Trichodinid ectoparasites (Ciliophora: Peritrichida) of freshwater fishes of the Zambesi River system, with a reappraisal of host specificity. Sys. Parasitol., 22: 81–109.
Van As, J.G., Basson, L. & Theron, J., 1984. An experimental evaluation of the use of formalin to control trichodiniasis and other ectoparasitic protozoans on fry of Cyprinus carpio L. and Oreochromis mossambicus (Peters). S.Afr.J.Wildl.Res., 14: 42–48.
Viljoen, S. & Van As J. G., 1983. A taxonomic study of sessile peritrichians of a small impoundment with notes on their substrate preferences. J. Limnol.Soc.Sth.Afr., 9: 33–42.
Viljoen, S. & Van As J. G., 1985. Sessile peritrichs (Ciliophora: Peritricha) from freshwater fish in the Transvaal, South Africa. S.Afr.J. Zool., 20: 79–96.
Woo, P.T.K., 1987. Cryptobia and cryptobiosis in fishes. Advan.Parasitol., 26: 199–237.
ILLUSTRATIONS page 52.
ILLUSTRATIONS (pages 52 – 57)
Plate 8. Ectoparasitic Protozoa: a. Ichthyobodo necator on gill arch of Oreochromis aureus, Israel. b. l. necator on mouth lining of wild goldfish, Israel. c–e. Cryptobia infections, Israel: c, on farmed goldfish; d. on Hypophthalmichthys molitrix (silver carp) gill rackers. f–m. Chilodonellosis: f. O. aureus, Israel; g. live Chilodonella hexasticha, O. mossambicus, S. Africa; h. C. hexasticha, silver impregnated, O. mossambicus, S. Africa; i. C. piscicola (=cyprini), O. mossambicus, S. Africa (h,i. by courtesy of L. Basson); k–m. Gill damage in O. aureus x niloticus, Israel: k,l, abrasion and desquamation (arrows - proliferating chloride cells) and m, epithelial hyperplasia.
Plate 9. Ectoparasitic Protozoa continued: a. Scanning electron microscopic view of severe gill chilodonellosis in O. aureus x niloticus Israel. b. Ambyphrya sp. from O. mossambicus S. Africa (haematoxylin stained). c. same Ambyphrya sp. live. d. Apiosoma sp. live ex O. mossambicus. e. Stalked Apiosoma sp. O. mossambicus. f. Scanning electron microscopic view of Scopulata sp. O. aureus x niloticus fry, Israel. g. Scopulata constricta from O. mossambicus, S. Africa (haematoxylin stained)(courtesy of L. Basson). h. Enlarged view of live, stalked Apiosoma, O. mossambicus, S. Africa. i. Scopulata sp. live, from O. mossambicus, S. Africa. k. Apiosoma piscicola from carp skin (haematoxylin stained).
Plate 10. Ectoparasitic Protozoa: trichodinids: a,b. Trichodina heterodentata: a, live, carp, Israel; silver impregnated, O. aureus x niloticus, Israel. c. Scanning electron microscopic (SEM) view of T. reticulata of goldfish. d,e. SEM view of O. aureus x niloticus gill infection with Tripartiella cichlidarum; f. same as d, in histology.
Plate 11. Ectoparasitic Protozoa: trichodinids continued: Silver impregnated trichodinids: a. Trichodina compacta (prev. acuta), O. mossambicus, S. Africa. b. T. migala (prev. pediculus), O. mossambicus, S. Africa. c. T. mutabilis, Barbus paludinosus, S. Africa. d. T. centrostrigata, O. mossambicus, S. Africa. e. T. reticulata, farmed goldfish, Israel. f. T. minuta, O. mossambicus, S. Africa. g. Trichodinella epizootica, Carp, S. Africa. h. Tripartiella cichlidarum, O. aurea, Israel. i. Hemitrichodina robusta, Marcusenius macrolepidotus (by courtesy of L. Basson).
Fig. 1. Ectoparasitic protozoa (page 57 with legend)
Plate 8. Ectoparasitic Protozoa (legend p. 52).
Plate 9. Ectoparasitic Protozoa continued (legend p. 52).
Plate 10. Ectoparasitic Protozoa: trichodinids (legend p. 52).
Plate 11. Ectoparasitic Protozoa: trichodinids continued (legend p. 52).
Fig. I. Ectoparasitic and intestinal Protozoa: A. Ichthyobodo necator free (left) and attached (10–15 μm long axis). B. Ichthyobodo sp. from Aplocheilichthys gambianus from South Ghana (10 μm long axis). C. Cryptobia (length 6–8 μm). D. Hexamita sp. from tilapia hybrid gut (7–12 μm). E. Thecamoeba (40 μm diam.) F. Entameoba (15 μm diam.). G. Life cycle of Ichthyophthirius multifiliis: 1. Trophont; 2. Dividing tomont; 3. End of division - tomites (theronts) escape from the cyst residues; 4. Tomite (theront). H. Protoopalina (150–350 μm long).