Plates 12 and 13 (pp. 67 & 68), Fig. 1G (p. 57)
Species affected and geographical range
Parasitic dinoflagellids, the marine Amyloodinium ocellatum and the freshwater
Piscinoodinium pillulare and P. limneticum, are not discriminatory in their choice of
piscine hosts and have been implicated in mass mortalities of tropical marine and
freshwater aquarium fish (Jacobs, 1946; Schaeperclaus, 1954; Paperna, 1980). A.
ocellatum has infected sea water acclimatised Oreochromis mossambicus and Aphanius
dispar in inland salt pans; some strains of this parasite survive in salinities as low as 10
ppt. Schaperclaus (1954) reports P. pillulare infection in 14 species of tropical
ornamental freshwater fish of diverse families as well as in carp and crucian carp.
Epizootic infections and mortalities were recently reported in farmed cyprinid fish in
Malaysia, including grass carp (Ctenopharyngodon idella), bighead (Aristichthys nobilis),
Leptobarbus hoevenii and Puntius gonionotus (Shaharom-Harrison et al., 1990). The
presence of P. pillulare has never been established in Africa, but this ubiquitous parasite
may eventually be found. If introduced with culture seed, it is likely to become
established.
Diagnosis
Trophonts, when reaching the final stage of growth, are visible to the naked eye (80–100
μm diameter) as white spots (similar to that seen in ichthyophthiriasis) and turn dark blue
when exposed to Lugol's-iodine. They are oval with a smooth wall and with inner
aggregates of globules. In Malaysian fish, clinical signs of P. pillulare infection comprise
both a rust-coloured appearance of the skin, indicating the presence of the parasite
trophonts (20–75 × 14–50 μm), and a dense covering of mucus (Shaharom- Harrison et
al., 1990).
Life cycle and biology
The life cycle of the dinoflagellid fish parasite is comprised of a parasitic feeding stage
(trophont) which attaches to integumentary epithelial cells, and an encysted dividing
stage (tomont) which is detached from the host. The trophonts of P. pillulare derive an
essential part of their nutrition from photosynthesis. Trophonts dislodged at any time
during their trophic stage will transform into a dividing tomont. Divisions yield a motile
infective stage (dinospore) which attaches to a new host. There are several detailed
studies of A. ocellatum (Paperna, 1984a), but comparable detailed data on the
freshwater fish dinoflagellids are lacking. Data on P. limneticum growth and division
(Jacobs, 1946), suggests that parasites reach a “maturation” prior to detachment. P.
pillulare trophonts on the gills, at 23–25°C, develop from dinospore to detached tomont
in three to four days. The tomont then completes division to the dinospore stage within
50–70 hours. At 15–17°C, the process of division is lengthened to 11 days
(Schaperclaus, 1954).
Pathology and epizootiology
In larval fish, infections were limited to the skin, whereas in large fish the highest parasite
densities occurred on the gill filaments and in the buccal-pharyngeal integument. Fish
recovering from the epizootic infestation through a gradual decrease in infection, could
not be reinfected (Paperna, 1980).
A. ocellatum is attached to and feeds from the host epithelial cell by means of rhizoids, which penetrate the host cell. The consumed cell gradually degenerates and collapses. Damage to infected cells leads to focal erosion of the epithelium. Prolonged infection exhausts a generation of mucus cells and leads to accelerated desquamation. Proliferation of the epithelium causes obliteration of the gill lamellae, while the inner strata of the epithelium become spongious and in some cases undergo complete lysis (Paperna, 1980). Attachment and penetration organelles of P. pillulare differ from those seen in A. ocellatum, in that the host cell is penetrated by nail-like extensions. However, damage to the host cell is similar (Lom & Schubert, 1983). Significant histopathological changes are only seen in the gills, where most of the infection occurs, namely a massive proliferation of the branchial epithelium which causes fusion of the lamellae by a confluent cellular mass (Shaharom-Harrison et al., 1990).
Piscinoodinium infection in Malaysia initially occurred among ornamental fish, but it spread eventually to pond farmed local and exotic cyprinids, causing mortality in fry of Puntius gonionotus in particular, although clinical signs were also apparent in a wider range of cyprinid fish species (Shaharom-Harrison et al., 1990).
Control
A. ocellatum is controlled by continuous application of copper sulphate, 0.75 ppm into
infected tanks. A further option is a mixture of 5-hydrate copper sulphate with citric acid
monohydrate, to yield 0.15 ppm copper ion concentration in the water (Hignette, 1981;
Kabata, 1985). The same methodology will apparently effectively control Piscinoodinium
infection, although concentrations should be adjusted to the freshwater medium and the
fish targeted for treatment. In freshwater with a pH below 7.0 (in tropical aquaculture),
concentrations above 0.3 ppm may be lethal to fish (e.g. Puntius gonionotus fry).
Species affected and geographical range
Most species of freshwater fish are susceptible, although some may be more so than
others. The world wide distribution of I. multifiliis (Hoffman, 1970) has apparently been
facilitated by the widespread translocation of cultured and ornamental fish. The presence
of this parasite in autochthonous fish, in remote areas of the world including, southern
Venezuela (Ventura & Paperna, 1985) and Northern Transvaal in South Africa (Paperna,
unpublished) may suggest, however, that many populations, particularly those in the
tropics, are comprised of a mix of autochthonous and introduced parasites. Data
available from Africa are limited to Southern Africa (in cichlids, carp, Barbus spp., trout
and eels - Du Plessis, 1952; Lombard, 1968; Jackson, 1968; Van As & Basson, 1984)
and Uganda (on native Barbus amphigramma and exotic Lebistes reticulatus from small
streams at Kajansi - Paperna 1972). It is very common in Israel, in both farmed (Sarig,
1971; Hines & Spira, 1973a) and wild fish including cichlids (Ventura & Paperna, 1985).
Description and diagnosis
Gross signs - white spots on the skin and the gills, which under microscopic examination
reveal (in skin and gill scrapings) uniformly ciliated organisms with a small cytostome,
which may reach up to 1 mm in diameter. Staining with either haematoxylins or Giemsa
(after adequate fixation, in such as Bouin) reveals a large crescent-shaped
macronucleus and small micronucleus. Ichthyophthirius multifiliis is a monotypic genus
of hymenostomatid ciliates.
Life history and biology
Trophonts (feeding stages) develop within the integumentary epithelium, always above
the basal membrane (Ventura & Paperna, 1985; Ewing & Kocan, 1992). By maturity,
which is reached in 2 days at ambient temperatures of 25–28°C (3–4 days at 21–24°C),
the parasite evacuates the host tissue and settles within 2–6 hours on a substrate in the
water to form a cyst-encapsulated tomont (dividing stage). Parasites evicted from the
tissue before the scheduled time for their spontaneous departure, fail to develop into
tomonts and eventually die (Ewing & Kocan, 1992; Paperna, unpublished observations).
Within the cyst, tomonts undergo successive binary fissions with a resulting yield of
250–2000 tomites (infective, free swimming stages), which after release will seek a
suitable host. The division of tomonts into tomites, in ambient temperatures of 25–28°C,
is completed within 15–20 hours (Bauer, 1959; Meyer, 1969; Paperna, unpublished
observations; 7–8 hours according to Hoffman, 1978).
Invasion of tomites (teronts) (30–45 μm long) into the host integument is facilitated by the excretion of a sticky substance from subpellicular crystalline organelles named mucocysts. Active penetration causes focal necrosis of the epithelial cells. It has been suggested that hyaluronidase and other enzymes may be produced by the penetrating parasite (Ewing et al., 1985). In the absence of a suitable host, tomites will lose their infective potential within 24 hours at 24–28°C (Ewing & Kocan, 1992). Higher temperatures hasten trophont maturation and tomont division, but at lower temperatures, slower development allows the growth of larger trophonts (0.8–1.0 mm in 5–10°C vs 0.5–0.7 mm in 20–24°C), yielding tomonts with higher numbers of tomite progeny (Ewing et al., 1986). In lower ambient temperatures the survival of the tomites is prolonged, thus, allowing more time to locate a host. Low temperatures do not interrupt propagation, a full cycle is completed at 20°C in 3–5 days, at 15°C in 7–14 days and at 10°C in 21–35 days (Bauer, 1959; Meyer, 1969). Data on the effect of other environmental parameters is less conclusive, although it has been suggested that dissolved oxygen levels below 1 mg/l affect parasite reproduction (Bauer, 1959).
Pathology
Ichthyophthiriasis is fatal to fish of all sizes. Chronic infection will cause serious damage
to the skin, fin and gills; corneal infection impairs vision (Hines & Spira, 1973a, 1974a).
The infective stage invades the integumentary epithelium and becomes established in
the basal layer of the epithelium just above the basal membrane. Cellular damage in low
to moderate infections remains restricted to the infected site. In addition to the damage
caused to epithelial cells by the feeding and expanding parasites, in heavy infections
mass exodus of parasites from the epithelial layer, having completed their scheduled
growth, causes its erosion and detachment from the basal membrane. In some
infections, parasites cause widespread lysis of the inner layer of the epithelium.
Prolonged infection also induces epithelial proliferation and haemorrhagic inflammation,
causing the integument to become severely disintegrated (Hines & Spira, 1974a;
Ventura & Paperna, 1985; Ewing et al., 1986). Hines and Spira's (1973b, 1974a,b)
haematological and clinical data from heavily infected fish reveal evident physiological
dysfunction resulting apparently from both direct pathological damage induced by the
parasite and as a by-product of the stress response.
Epizootiology
Ichthyophthirius multifiliis is one of the most common, troublesome and difficult to control
of fish pathogens. Epizootic infections have been reported in cold water salmonid farms
(also in Africa, Du Plessis, 1952; Lombard, 1968) and warm water farmed carp, eels,
Clarias gariepinus and Ictalurus punctatus (channel catfish) (Meyer, 1970; Sarig, 1971;
Hines & Spira, 1973a; Hine, 1975; Jackson, 1978; Khalifa et al. 1983; Paperna,
unpublished). Fish may maintain low, subclinical (enzootic) infection (in preimmunity),
while encysted tomonts may persist in the habitat. Enzootic infections in native fish have
been found in Lebistes reticulatus in Uganda (Paperna, 1972), in cichlids and cyprinids
native to the Lake Kinneret system in Israel (Paperna, unpublished), in glass eels,
cyprinids and cichlids in native habitats of South Africa (Jackson, 1978; Van As &
Basson, 1984), and in a variety of native fish in the southern United States (Allison &
Kelly, 1963). Transition from nonclinical enzootic to epizootic clinical infection is usually
stress-mediated, prompted by adverse growth conditions such as overcrowding, poor
feeding and excess nitrogenous waste. Epizootic infection, however, never occurs in
overwintering tilapia or Clarias gariepinus in Israel, or southern Africa, but rather,
coincides (also in southern USA - Meyer, 1970) with the warming of the water in early
spring when fish are still kept in overcrowded conditions after winter storage. In South
Africa, 6.4% of wild glass eels in the Southern Cape are infected. Via these, fish infection
has been introduced into eel nurseries where elvers, especially those not completely
acclimated, succumbed to severe infestations (Hine, 1975; Jackson, 1978).
Spontaneous recovery has been observed in both natural infections in natural habitats and in holding facilities, and even in experimental infections in aquaria (Paperna, 1972; Lahav & Sarig, 1973). The potential for spontaneous recovery varied with fish species. Infection in scaled fish, notably cichlids, regressed faster than in smooth skinned fish (eels, Clarias spp. and other siluriforms, mirror and leather carp) (Paperna, 1972 and unpublished observations). After recovery, fish were refractory to reinfection or retained a merely subclinical chronic infection (Hines & Spira, 1974c; Wahli & Meier, 1985; Paperna, unpublished observations). The observed interspecific variation in susceptibility to infection could, however, also result from differential compatibility of various fish species to man-made habitats and variable vulnerability to stress.
Spontaneous recovery from infection and resistance to reinfection of recovered fish indicate that fish are capable of developing defence mechanisms against I. multifiliis (Hines & Spira, 1974c). Spontaneous recovery observed in carp at temperatures as low as 10°C (Lahav & Sarig, 1973) implies some protective responses other than via humoral antibody production, which becomes suppressed in carp below 12°C (Avtalion, 1981).
Hines & Spira (1974c) demonstrated immobilisation of free swimming tomites with sera taken from carp after their recovery from infection. The infective stages were also shown to be unable to penetrate the skin of resistant carp. Immobilisation tests with trophonts showed that in infected trout anti-parasitic activity of the mucus increases quickly after infection and decreases soon after the infection has disappeared. The anti-parasitic activity of the serum, in the same fish, increases slowly but remains at a higher level for at least 7 months (Wahli & Meier, 1985). Fish immunised with Tetrahymena spp. developed a resistance to a challenge of lchthyophthirius infection (Goven et al., 1981).
Carp were immunised following controlled exposure to the infective tomite (teront) stage, and survived challenges with high infective doses, but lost protection after being immunosupressed by the administration of corticosteroids. These results could have simulated a stress mediated situation. Levels of humoral antibodies in immunosuppressed fish, however, remained the same as in the immunised group, which further confirms the involvement of other than humoral type immune systems in the protection processes against l. multifiliis infections (Houghton & Matthews, 1990).
Immunisation with killed vaccines gave less satisfactory results, although better protection was obtained through intraperitoneal inoculation of live teronts (Burkart et al., 1990). Oreochromis aureus mothers vaccinated through the latter method, passively transferred a protective immunity to their fry (Subasinghe & Sommerville, 1989). Additionally to immunity passed from mothers via eggs, demonstrable by antibodies in the soluble extracts of fry tissues, a protective immunity was acquired directly from the parent mouth during the brooding period (Sin et al., 1994).
Control
Both trophonts, localised beneath the epithelial layer of the integument, and the encysted
tomonts, attached to substrates in the aquatic habitat, are resistant to practically all
externally applied usable antiparasitic agents.
Infection can be effectively controlled only by destruction or elimination of the free dividing tomonts or the tomites they release. In warm water systems (24–28°C), three to four daily transfers of fish to clean tanks will effectively reduce infection, while enabling the fish to develop tolerance to reinfections. Tomonts can be effectively removed from large circulating tanks by repeated brushing with vacuum suction. Spontaneous recovery and transition into a refractory state will be further promoted by management techniques which alleviate stressing conditions (improving water flow, accelerating aeration and reducing stocking densities). Chemical parasiticides will be effective only through continuous or repeated daily application. Of the many listed (Meyer, 1969), the only cost-effective remedy for large scale farming systems is Malachite green at a dose of 0.05 ppm for continuous application (3–4 days) or up to 0.15 ppm (depending on the specific fish tolerance which varies with species - siluriforms are particularly susceptible). Formalin will dislodge some of the trophonts and is often applied mixed with Malachite green (50 ppm with 0.05 ppm) (Sarig, 1971).
Systemic therapy seems to be the only means of effective control. Elimination of tissue trophozoites was reported in several species of ornamental aquarium fish fed medicated food (Tetra, MA 100/50) containing Malachite green in a non-water soluble formulation for 4 days (neither drug concentration in the food, nor daily rations are given; Schmahl et al., 1992). For use in commercial food-fish culture for human consumption, the cost efficiency of Malachite medicated feed formulations and their toxicity to humans must be considered.
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ILLUSTRATIONS (pages 66 – 68)
ILLUSTRATIONS
Plate 12. Dinoflagellids and Ichthyophthyriasis: a. Piscinoodinium sp. on Colisa lisa fry (Photo A. Diamant). b. Enlarged view of a. c. Amyloodinium ocellatum on farmed Sparus aurata fry, Red Sea. d. Trichophyra sp. (Suctoria) from Morone saxatilis, USA. e–i. Ichthyophthirius multifiliis: e, live trophont, carp, Israel (by courtesy of S. Sarig); f, benign infection on eel's (Anguilla anguilla) skin, live; g, Silver impregnated, Oreochromis mossambicus, SA; h, histology of same infection as f; i, benign infection carp.
Plate 13. Ichthyophthiriasis continued: a. heavily infected Barbus amphigramma, from Uganda; b. Early invasion of the pharyngeal mucosa of young eel, experimental. c. Heavy infection in skin of Clarias lazera fry, Israel; d, Gill infection of Oreochromis mossambicus, Transvaal, South Africa. e. Parasites displacing gill lamellae of carp, Transvaal, South Africa. f. Cytolysis in gill infection of Haplochromis flavii-josephi, Lake Kinneret, Israel. g. Infection resulting in proliferation of the gill epithelium of Apistogramma sp. from South Venezuela.
Plate 12. Dinoflagellids and Ichthyophthiriasis (legend p. 66).
Plate 13. Ichthyophthiriasis continued (legend p. 66).
