Tilapia

Diet

Allspice © Jonathunder

Baby's breath © PiPi

Tilapia may survive on algae or duckweed alone, but combined feeding results in higher growth rates. Superior growth is achieved by partial replacement with feed from animal origin, such as fishfeed or insects and worms.

Protein

Protein requirement for maximum performance of tilapia larva is 35 - >50%, decreasing with increasing fish size.[1][2][3] Juvenile tilapia need 30-40% protein, while adult tilapia need 20-30% dietary protein for optimum performance. Tilapia broodstock need 35-40% protein for optimum reproduction, spawning efficiency, and larval growth and survival.[4][5][6].

Carbohydrates

Tilapia is relatively well equiped (due to alpha-Amylases[7] produced by Aeromonas, Bacteroidaceae and Clostridium strains[8]) for utilizing carbohydrates.[9] Tilapia can utilize up to 46% dietary starch without growth retardation.[10] Tilapias do better on starch than on glucose (even when the glucose diet is supplemented with chromic oxide) [11] In tilapia, polysaccharides have antibacterial properties, increasing resistance to infections.[12]

Starch consists of amylose and amylopectin. Amylopectin is more susceptible to enzymatic cleavage. A high-dietary amylose-amylopectin ratio (> 0.24; as in corn starch) decreases starch digestibility and postprandial peak blood glucose and triglycerides, and inhibits feed efficiency and growth in tilapia.[13] Amylopectin accounts for about 75% of the starch weight in reserve starch (in storage organs) [14] and typically more than 90% in transitory starch (in photosynthetic organs)[15]. Spirodela polyrrhiza amylose content was 21%[16], compared to 25% in potatoes[17], 7-44% in barley[18] and 25–28 in wheat.[19]

Feeds with smaller granule size have higher starch digestibility than those with bigger granules. Spirodela polyrrhiza granules range from 1 to 8 μ [20], compared to 2-55 μ in wheat starch [21], 7 to 22 µ in corn starch and 8 to 110 µ in potatoes [22].

Fat

Tilapia do well on fat versus carbohydrates as a source of energy.[23]

Oxidative stress

The use of tuna by-product meal as a total replacement for fish meal into tilapia diets increases oxidative stress.[24] Dietary duckweed (Spirulina) incorporation increases antioxidant activity in tilapia.[25]

Chito-oligosaccharides

In tilapia stomach, intestine, and serum, chitinases are present. [26] Chitinases are enzymes required to enzymatically decompose chitin. Chitin is an amino polysaccharide; a long chain of many monosaccharide (N-acetylglucosamine) units. A chito-oligosaccharide is a short chain of just a few linked units of N-acetylglucosamine. Chitin may be split into chito-oligosaccharides. Chitin plus other composite materials (eg glycoproteins) forms the exoskeletons of insects and the flexible body wall of larva / caterpillars. The peritrophic matrix of the tobacco hornworm (Manduca sexta) may contain 40% chitin.[27] (Peritrophic matrices usually contain 3 to 13% chitin[28]) Chitin is also an ingredient of cell walls in algae.[29][30] Dietary intake of chito-oligosaccharides can improve intestinal health, and improve tilapia resistance to infection.[31] Dietary intake of chito-oligosaccharides at up to 4 g/kg may considerably improve growth, survival and immune response.[32] Dietary intake of chito-oligosaccharides at 3 to 5 g/kg for juvenile GIFT tilapia may increase growth performance, nonspecific immunity and promote blood lipid metabolism.[33]

Supplements

• Saponin-rich plants such as Quillaja saponaria (Soap bark tree), yucca and Sapindus saponaria (wingleaf soapberry, western soapberry, jaboncillo) may increase growth of (and influence male to female ratio in) tilapia.[34] Gypsophila paniculata (baby's breath) and Quillaja saponaria (both high in saponins) may induce release of LH (Luteinizing hormone)[35] Supplementing Tilapia fry with 700 mg/kg extracted Quillaja saponin extract resulted in significantly increased growth, higher number of males, higher LH release and higher serum cholesterol.[36] Saponin supplementation diminishes egg production by females.[37] Intestinal magnesium uptake in tilapia is stimulated by chloride and inhibited by saponins (which may induce intravesicular magnesium accumulation.[38] In tilapia, saponins selectively permeabilize the plasma membrane.[39]
• Allspice Pimenta dioica (10 g/kg fish) acts as a growth promoter to improve feed utilization and weight gain in Mozambique Tilapia fry and acts an antimicrobial agent to enhance disease resistance during first feeding of fry.[40] Allspice (aka Jamaica pepper, myrtle pepper, pimenta, pimento, English pepper, newspice) is dried unripe berries of Pimenta dioica.
• Honey bee pollen (2.5% (w/v)) increases growth performance parameters, feed efficiency ratio and immunological, hematological and biochemical parameters in young Nile tilapia.[41]
• A supplemental mixture (at 0.5 to 2% w/w) composed of astragalus, angelica, hawthorn, Licorice root and honeysuckle may increase lysozyme, superoxide dismutase and peroxidase activity, decrease malondialdehyde levels and reduce mortality in Aeromonas hydrophila-challenged tilapia.[42]

.

Growth

Tilapia nilotica fingerlings grow optimally on diets containing 30-40% protein, 12-15% lipids, and 30-40% digestible carbohydrate.[43]

Temperature

Feed efficiency and protein efficiency ratio are better and juvenile fish grow better at 28°C than at 22°C and 34°C. Liver gluconeogenesis and lipogenesis are increased by low temperature (22°C).[44]

Masculinization

Estrogen synthesis is crucial for ovarian differentiation, and transcription of the aromatase gene can be proposed as a key step in that process in fish. Treatments with an aromatase inhibitor (ATD, 1,4,6- androstatriene-3-17-dione) result in 75.3% masculinization of an all-female (XX) population in tilapia (dosage 150 mg/kg of food). The effectiveness of the aromatase inhibition by ATD is demonstrated by the marked decrease of the gonadal aromatase activity in treated animals versus control.[45]

Aromatase activity as a key factor in sexual differentiation in Oreochromis niloticus.[46] The most sensitive time to aromatase inhibitors lies in the first week (between 7 and 14 days post hatch). Treatment with the aromatase inhibitor Fadrozole (nonsteroidal) showed a dose-dependent increase in the percentage of males from 0 to 200 mg per kg. At higher doses (200 to 500 mg / kg), the percentage of males remained more or less constant (92.5-96.0%).[47]

The masculinizing actions of 17alpha-methyltestosterone (MT) are most potent at up to day 20 of age.[48]

Treating female tilapia Oreochromis niloticus with methyltestosterone (at a dose of 50 mcg/g diet) resulted in 100% masculinization.[49]

Treatment with tamoxifen and letrozole (200mg/kg feed) to fingerlings of O. niloticus for 60 days brought about 98.5% masculinization. Treatment with 17α methyltestosterone (35 mg/kg feed) to fingerlings of O. mossambicus after 8 days post hatch for 60 days obtained 100% sex reversed males with excellent growth.[50]

During the restricted developmental period temperature is of great influence. The critical period for elevated-temperature-induced masculinization lays between days 10 and 15 post-hatch.[51] Higher temperatures (during 5 days) before they are 5 days old induces deformities. Masculinization is induced at elevated temperatures (28 to 32C) during 5 days after 10 days old.[52]

Feminization

The critical period for low-temperature-induced feminization lays between days 5 and 10 post-hatch.[53] The period of maximal feminizing action of 17beta-estradiol (E(2)) upon sex ratio is before 10 days posthatching in tilapia (Oreochromis mossambicus).[54]

During the restricted developmental period temperature is of great influence. Higher temperatures (during 5 days) before they are 5 days old induces deformities. Gonadal feminization is induced at lower temperatures (20C) for 5 days before 10 days old.[55]

Parasites

Tilapia may be infected with:

• Nematodes / roundworms, such as Contracaecum sp.[56] (eg Contracaecum multipapillatum [57]), Paracamallanus cyathopharynx, Procamallanus laevionchus [58], Rhabdochona esseniae [59], Rhabdochona paski [60], Gnathostoma spinigerum, Gnathostoma doloresi and Gnathostoma hispidum.[61] Human gnathostomiasis infection is related to ingestion of "ceviche".[62]
• Cestodes (aka tapeworms), such as Amirthalingamia sp. (eg Amirthalingamia macracantha; higher prevalence and intensity in wet season [63]), and Cyclustera sp.[64] (eg Cyclustera magna [65])
• Trematodes / flukes (encysted metacercariae)[66], such as Clonorchis sinensis (human liver trematode)[67], Tylodelphys sp.[68], Stictodora tridactyla (in brackish-water fish)[69], Ribeiroia marini [70], Clinostomum sp.[71] ((eg Clinostomum Complanatum, Clinostomum tilapiae, Euclinostomum hetereostomum [72]), Diplostomum sp.[73], Neascus sp, Acanthostomum sp.[74], Bolbophorus sp.[75], Haplorchis yokogawai [76], Haplorchis pumilio, Haplorchis taichui, Centrocestus formosanus, Stellantchas musfalcatus, Echinochasmus japonicus [77], Heterophyes heterophyes, Heterophyes aequalis, Pygidiopsis genata, Stictodora sp., Phagicola sp.[78] (eg Phagicola ascolonga[79]) and Prohemostomumn vivax.[80] These trematodes loose their viability after 48 hours at -5°C.[81] The conditions of gas supersaturation (gas pressure > 111.2% saturation; N supersaturation, O2 undersaturation) may result in heavy monogenetic trematode infections in tilapia.[82] In Thailand, fish-borne zoonotic trematode (FZT) metacercarial infections were found in Nile tilapia from cage (2.5%) and pond aquaculture systems (10%) and in wild caught fish.(53%) [83] In China, tilapia from nursery and grow-out ponds were sampled from monoculture, polyculture and integrated aquaculture systems, revealing a 1.5% prevalence of FZT infections (Heterophyidae and Echinostomatidae); lower than in wild caught fish. Integrated systems using animal manure and latrine wastes as fertilizer did not show a higher prevalence.[84] In Vietnam, the overall FZT prevalence in tilapia from wastewater ponds was 2.0%, but much higher in tilapia from farm ponds. [85]
• Monogenea (very small flatworms), such as Dactylogyrus minutus [86], Neobenedenia melleni [87][88], Enterogyrus cichlidarum (associated with overcrowding [89] and infection intensity increases with (tilapia) host size[90]), Gyrodactylus malalai (ectoparasite)[91], Gyrodactylus ergensi [92], Scutogyrus longicornis [93] and of the genus Cichlidogyrus [94], such as Cichlidogyrus tilapiae, Cichlidogyrus sclerosus [95], Cichlidogyrus dossoui [96], Cichlidogyrus halli [[97], Cichlidogyrus thurstonae [98], Cichlidogyrus berradae, Cichlidogyrus revesati, Cichlidogyrus legendrei and Cichlidogyrus lemoallei.[99] The highest parasite number was recorded in reservoir-dwelling, and lowest in stream-dwelling tilapia.[100] Gyrodactylus niloticus infection may particularly become lethal (in tilapia) when followed by exposure to the bacterial pathogen Streptococcus iniae.[101] Low stocking density and low water temperature may result in lower monogenea parasitism rate.[102]
• Acanthocephala (thorny-headed worms), such as Acanthogyrus (Acanthosentis) tilapiae.[103] and Polyacanthorhynchus kenyensis [104] In Kenya, infection rates in wild Tilapia zillii ranged from 4.1 to 77.7%.[105]
• Ciliatea ('small flagella'), such as Epistylis sp.[106], Paratrichodina africana [107], Trichodina compacta [108], Trichodina magna [[109] and Ichthyophthirius multifiliis.[110] Ichthyophthirius multifiliis load increases susceptibility and mortality of tilapia to Streptococcus iniae (Gram-positive bacteria) infection.[111]
• Dinophyceae (Dinoflagellata; "red tide"), such as Piscinoodinium pillulare.[112] (a dominant parasite in Brasilian tilapia ponds)[113]
• Myxosporea, such as Henneguya suprabranchiae [114], Myxobolus heterosporus [115] (induces total destruction of the intestine structure [116]), Myxobolus nounensis [117], Myxobolus dossoui (host size effect), Myxobolus zillii (seasonal pattern) [118], Myxobolus microcapsularis [119], Myxobolus agolus, Myxobolus brachysporus, Myxobolus clarii, Myxobolus cichlidarum, Myxobolus tilapiae and Myxobolus camerounensis.[120]
• Conoidasida (unicellular), such as Goussia cichlidarum.[121]
• Phyllopharyngea (single-cell parasites), such as Chilodonella hexasticha.[122]
• Trhypochthoniellus longisetus longisetus, a mite.[123]
• Lamproglena sp. (Lernaeidae; small crustaceans) [124]
• Pentasomida (tongue worms; tiny crustaceans), such as Leiperia cincinnalis [125], Subtriquetra wedli [126] and Subtriquetra riley.[127]
• Anodontites trapesialis (the larval stage of this mussel).[128]

No parasite vaccines exist.[129] Tilapia are relatively resistant to parasitic infections.[130] The prevalence of heterophyid encysted metacercariae infection decreases as fish size increases.[131] When dissolved oxygen, temperature, pH, nitrite and ammonia are within normal rate parasites may cause little harm to tilapia.[132] Parasitic infection by Acanthogyrus tilapiae provokes an aggressive host response.[133] Tilapia may produce an induced humoral immune response against Cichlidogyrus sp.[134] Oreochromis mossambicus initiated a strong immune protection by direct exposure with even a small number of parasites.[135] Gyrodactylus numbers fluctuate independently of temperature. In anticipation of immune defenses reaching the fish surface through mucus, Gyrodactylids (Monogenea) worms progressively move away from fins with high mucus cell density to those with low density.[136] Tilapia may also develop resistance to Neobenedenia sp. (Monogenea) infection.[137]

Freshwater mollusks in tilapia ponds, such as Melania tuberculata, Melanoides turricula, Pomacea flagellata, Haitia cubensis and Anodontiles luteola, may host various parasites that are harmful to tilapia.[138] The Biomphalaria sp. snails are often found in tilapia ponds (fresh water reservoirs are their natural habitat[139]) and are intermediates for Schistosoma mansoni.[140] Biomphalaria cf. havanensis is an intermediate host for Diplostomum ompactum, causing higher levels of infections in cultured tilapia than wild tilapia.[141] The snail Bulinus truncatus is an intermediate host for Clinostomum tilapiae, Euclinostomum heterostomum, Bolbophorus levantinus and Neascus-type metacercariae. The snail Melanoides tuberculata is an intermediate host for Centrocestus sp. and Haplorchis sp. The snail Melanopsis costata is an intermediate host for Pygidiopsis genata, Phagicola longa.[142] The snail Thiara granifera is commonly infected with Haplorchis pumilio.[143] Dogs, cats and pigs may also be intermediate hosts for metacercariae.[144]