Daily operations

Daniels etal. (1992) and Valenti et al. (1998) supplied step-by-step guides for daily tasks associated with freshwater prawn recirculation systems. Morning duties include monitoring both the system and the larvae. Water temperature, flow and level of water should be checked and screens cleaned or changed. Water losses by evaporation should be replaced with freshwater. Care should be taken to ensure that water replacement does not shock either the larvae or the biofilter. Mechanical filters should also be monitored and cleaned as needed. Larvae are fed ( Artemia or prepared diet) and then checked following the steps described in section 5.4.3. Artemia nauplii are harvested daily with excess quantities frozen for future use and new cysts are prepared for hatching.

In the afternoon, aeration and water flow should be temporarily turned off and visible waste feed and faeces siphoned (Fig. 5.10) in order to avoid increases in organic matter and subsequent increases in heterotrophic bacteria. These are potentially pathogenic bacteria, which compete with nitrifying bacteria and contribute to additional inorganic nitrogen in the water by decomposition. Prior to siphoning, the bottom and sides of the tank may also be scrubbed to remove all algae and organic accumulations; however, during the cleaning, aeration must be maintained to prevent larvae from being trapped between the mop and sides orbottom ofthe tank. Daniels etal. (1992) proposeda

Fig. 5.10 Siphon cleaning system.

daily cleaning routine beginning by day 3 and ending after the appearance of PL. Siphoned waste can be used to estimate larval mortality and feeding rates (see section 5.4.5). From day 18 to 20 onwards, siphoning provides an opportunity to observe the presence of PL. Larvae or PL that are removed during the cleaning process are returned to the same tank. Once cleaning and siphoning are done, aeration is restarted and, if water exchange is to be done that day, it normally occurs at this time. Then, larvae are fed again.

Water quality and larval health are critical for achieving good growth and survival. Failure to be attentive to the needs of the larvae can cause either chronic or acute larval stress. Chronic stress can delay larval development, cause poor larval health, and subsequent poor survival, whereas acute stress causes immediate mass mortality. Any of these can result in decreased production and increased costs. In closed systems, daily control of the majority of the water quality parameters is not essential because conditions are usually stable. However, the nitrification process should be rigorously monitored. Nitrite and ammonia levels should be measured daily until the system is stable, then both should be monitored several times weekly. The amount of monitoring is dependent upon the experience ofthe operational management, its confidence in the system and the amount of water exchanged daily. Newer systems should be monitored more often to ensure the stability of the system.

Normally, rearing tanks need some replacement of water daily. In recirculating systems, replacement of the amount removed during the cleaning (siphoning) process is relatively simple. However, in certain cases where water quality is observed to be very poor (e.g. due to an error in feeding) a major water exchange may be necessary. In flow-through systems, water exchange is essential, especially when inert feeding is commenced. In some hatcheries, little or no water exchange occurs during the first 5 to 7 days when only live food is provided. After that, the tank water is changed every day, or every second day, depending on water quality. Some hatchery operators favour a greater water exchange every other day, since this stimulates moulting. Water exchange at an average of 20 to 30%/day continues until day 18 to 20. Then, when the first PL are observed, daily water exchange is increased to 50 to 60%/day. Again, water exchange is observed to stimulate moulting and, at this moment, metamorphosis to PL. Thus, increasing the rate of water exchange may shorten the period when metamorphosis of the batch occurs. This observation is yet to be scientifically confirmed.

Possible diseases (Chapter 14) should be avoided, especially bacterial populations. Daniels et al. (1992) suggested that bacterial concentrations should be monitored several times a week using a dilution method or some other type of testing. However, they emphasised that decreasing food consumption by larvae is an indirect indicator of bacterial or water quality problems. An experienced manager can use this indicator to identify potential problems and act before a major loss of larvae occurs. Bacteria settled on the surface of all submerged materials, including tank walls, form biofilms that are more resistant to antibiotics and sanitis-ers. Carvalho & Mathias (1998) recommended periodically collecting the scrapings from submerged filter and tank walls for microscopic analysis. Pseudomonas, Aeromonas, Bacillus and Vibrio were the most common genera found in M. rosenbergii hatcheries in India (Sahul Hameed et al. 2003; Jayaprakash et al. 2006; Kennedy et al. 2006; Dass et al. 2007), whereas Chryseomonas, Vibrio, Cellulomonas, Aeromonas and Pasteurella dominated the water, debris, larvae and tank walls in a hatchery in Saudi Arabia (Al-Harbi & Uddin 2004). Most pathogenic bacteria in the water column or biofilms are autochthonous to the hatchery system and may be opportunistic pathogens. Prevention of disease through rigorous water management and sanitation is the best methodology forthe control ofpathogens. Many ofthe chemicals used to treat the larvae cause deformities and/or their use may be cost prohibitive.

Health management depends upon cleanliness and minimising the introduction of disease organisms. Massive blooms of Zoothamnium, Epistylis and hydroids, which are harmful to larvae, occur if hatchery hygiene is inadequate or if the incoming water is poorly treated. Every effort should be taken to keep equipment, supplies and food clean at all times. Precautions should be taken to prevent the introduction of disease organisms or chemical contaminants by workers and visitors entering the hatchery. Chemical footbaths and washing of hands prior to and when returning to work are recommended (Carvalho & Mathias 1998; Valenti et al. 1998). Carvalho & Mathias (1998) recommended cleaning rearing tanks immediately after emptying, disinfecting with 50 mg/L formaldehyde solution, rinsing with clean freshwater, and then drying. Regular disinfection of all equipment with a 100 mg/L sodium hypochlorite solution, followed by sun-drying, is recommended. The transfer of water and equipment from one larval tank to another without disinfection must be avoided. All utensils, such as beakers, porous stones, hoses and buckets should be disinfected every other day using 2.5 mg/L active chlorine, rinsed with freshwater and dried prior to storing. Good personal hygiene is also advisable and the use of tobacco in or around the hatchery should be discouraged. New (1990) noted that the use of antibiotics, and sometimes sulpha drugs, in larval rearing was commonplace. This practice is not normally publicised, and cannot be recommended. Hsieh et al. (1989) described the use of the antibiotics chlo-ramphenicol and tetracycline, as well as furazolidone, formalin and iodine, in larval rearing; their indiscriminate use was reported to be a problem in Taiwanese hatcheries at that time.

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