Hatchery Management of Oyster, Mud crab and Mussel

Hatchery Management of Oyster, Mud crab and Mussel

IntroductionAny aquatic invertebrate animals having a cutaneous or calcareous shell surrounding there body and belonging to the phylum Mollusca, the class Crustacea (phylum Arthropoda), or phylum Echinodermata is known as shellfish. The term is often used for the edible species of the groups, especially those that are fished or raised commercially. The most commercially important shellfish are:

Mollusk: Oysters, mussels, scallops and clams Crustacean: Shrimp, prawn, lobster, crab and crayfish Echinoderm: sea urchins and sea cucumbers

Shellfish hatchery is a place where shellfish seeds are produced in a controlled way. Hatchery management is a branch of science which deals with the activities including from collection of brood shellfish to seed production. Culturing of shellfish has occurred since ancient times. Although controlled rearing of young shell has long existed, hatchery production is a more recent advancement. Producing seed under controlled conditions in a hatchery will disconnect its production from environmental factors and provide a reliable supply of seed. Oysters, mussels and mud crabs are the most important groups of shellfish after shrimp and prawn. These are popular among the western countries and becoming more popular all over the world. So hatchery management of oyster, mussel and crab is crucial.

Importance of oyster, mussel and mud crab hatchery management:

To produce required amount of healthy and high quality demanded oyster, mussel and mud crab seeds in proper time.

To produce high quality demanded hybrid shellfish. To avoid inbreeding problem. To avoid dependence on nature for seeds. To utilize manpower. It is a profitable business. To earn a lot of foreign currency by exporting them.

OBJECTIVES OF THE ASSIGNMENT

The main objective of the assignment is to have a good knowledge about hatchery management of oyster, mud crab and mussel to produce

required amount of healthy and high quality demanded oyster, mussel and mud crab seeds in proper time.

Hatchery management of oysterOyster hatcheries provide juvenile oysters for commercial production, restoration projects and research. Oyster hatchery techniques are well established and many publications describe how to produce oysters (see Additional Reading). This publication covers the basics of oyster hatchery production based on information from previous publications and our own experience. The information is generally applicable to oysters in the genus Crassostrea and is most applicable to the eastern oyster, Crassostrea virginica, in southern waters.

Oyster biologyAn oyster hatchery simply creates a controlled environment for the early portions of the oyster life cycle. Therefore, producers must understand oyster biology. Oysters occur naturally in dense aggregations, often called reefs or beds. Oysters thrive in estuarine waters with salinities of about 10 to 25 ppt, though they can tolerate lower and higher salinities. Oysters on natural reefs are stimulated to spawn when the water temperature rises in the spring. The release of sperm and eggs into the water further stimulates other oysters to spawn. This results in a mass release of reproductive products. Sperm fertilize eggs in the water column. Fertilized eggs develop and progress through a series of free-swimming larval `stages (Fig. 5.) over a period of 14 to 20 days, depending on water temperature. These stages are referred to as the trochophore, veliger and pediveliger. The trochophore larvae feed on very small algae as they move through the water column. Trochophore larvae quickly develop into more motile veliger larvae (Fig. 6). Toward the end of the larval cycle, pediveligers (Fig. 7) develop a foot that helps them find a suitable hard substrate on which to attach (set) and transform into small oysters. This stage is also called an “eyed larvae” because of the development of a pig-mented eye spot. Eyed pediveligers settle out of the water column when they are approximately 300 micrometers (μm) and may be stimulated to settle by the presence of adult oysters. Finding a hard substrate (cultch) is essential to their survival. The eyed larvae can move only very small distances, once they settle, in order to find a suitable spot. Once settled, they attach and transform into small oysters called spat. Spat soon begin feeding on algae by filtering water through their gills and a special structure (labial palps) located just in front of the mouth.

Site selectionThere are many critical elements to an oyster hatchery, but none is more important than location, or more specifically, location relative to water supply. Oyster hatcheries require large volumes of clean sea water with salinities in the range of 15 to 30 parts per thousand (ppt). Salinity is not a major issue in many areas, but some estuaries have periodic episodes of freshwater inflows that can reduce salinity below 10 ppt. Low salinity water is not conducive to spawning, larval development, or early growth of young oysters. Turbidity, potential pollutants, watershed development, boat traffic, and natural algae production are other aspects of water quality to consider. The local agency that oversees oyster harvesting areas should be consulted in regard to regulations that might affect the use of oysters produced from the proposed hatchery site.

Hatchery facilityOyster hatcheries vary in size and shape depending on budgets and pro-jected production. Most facilities have a separate pump station that brings saltwater from a nearby source to the hatchery. Having dual water lines and pumps provides a backup system and can also reduce the fouling of lines because one line can be allowed to go anoxic while the other is in use. Pumps and water lines are sized for the distance, height (head) and volume of water to be moved. Systems that can deliver 10 to 25 gallons (38 to 95 L) per minute simultaneously from multiple outlets will efficiently fill a number of large tanks and ensure adequate flow for growing oysters.

Figure 1. Large holding tanks for settling and water reserves.Before it enters the hatchery, water is often pumped to large holding tanks (Fig.1) where settling reduces turbidity. Or, the holding tanks can be bypassed and water pumped directly to the hatchery.

Figure 2. Overhead water and air lines. Overhead water lines (Fig. 2) keep floor space cleaner and allow tanks to be filled directly. The plumbing is designed with a water filtration and treatment system consisting of some combination of rapid sand filters, cartridge filters, activated carbon, ultraviolet (UV) sterilization or pasteurization (Fig. 3).

Figure 3. Small cartridge and ultraviolet sterilizer filter system for treating seawater.

Treated sea-water is then suitable for larval and algal production.Tanks for larval production are circular, generally 250 gallons (946 L) or larger, and have center drains and sloping or conical bottoms. Drain pipes make it convenient to drain water and sieve larvae. An in-floor drain system (Fig. 4) that can handle the maximum expected water flow helps keep water off the floor.

Figure 4. In-floor water drains and nursery tanks.

Shallow rectangular tanks with drain pipes provide nursery space for juvenile oysters. Aeration throughout the hatchery is supplied by an appropriate size blower, overhead PVC piping, vinyl tubing, and good quality air stones. If algal cultures are to be produced, there must be a clean room with special lighting for starting cultures. Another larger space with special lighting, separate from the main hatchery, is needed to grow larger volumes of algae.

Basic hatchery proceduresThese are the basic procedures for spawning oysters, raising and setting larvae, and growing spat. No two hatcheries operate identically or in the same environment and there is no substitute for experience. Water treatmentGood water quality is essential to successful hatchery production (see Site Selection) but even high-quality water must be treated to remove unwanted organisms. Water used for spawning, mixing eggs and sperm, and growing larvae is typically filtered mechanically and treated with ultraviolet radiation. Large-scale operations and “low-tech” or “back yard” facilities may forgo UV treatment but will use some mechanical filtration. Mechanical filtration is usually done with a pressurized sand filter, cartridge filters for smaller volumes, or fine-mesh bags. Because of the rich array of organisms in Gulf of Mexico waters, mechanical filtration down to 1 μm with UV treatment can assist in successful spawning and larval production.

Hatchery operations, procedures and equipment:

Fig: General flow chart

A general flow chart of the hatchery system is shown in Fig.1. Only the major components broodstock maintenance and spawning, larval culture, setting and cultch preparation and algal culture—are shown. These four subsystems are essential to any oyster seed operation. The floor plan for the pilot hatchery is shown in Fig. 2. The figure merely indicates the location of the various subsystems in the pilot hatchery; it is not intended as a guide for hatchery construction. The characteristics of each hatchery site dictate its unique floor plan. At each location the components have to fit together; they shouldn't be forced. In practice it is not possible to separate the four hatchery subsystems. All subsystems—from broodstock maintenance and spawning to algal culture—must mesh together. The two major biological systems—larval growth and algae production— must be kept at their peaks at the same time. This synchronization is the key to a successful oyster seed hatchery.

WATER SUPPLYOyster seed hatcheries are similar to fish hatcheries in that water supply is of prime importance in site location. Water for the hatchery should have a salinity higher than 20 parts per thousand. Some water storage may be necessary to provide for short periods of low ambient salinity. Water temperatures should not exceed 20 C. Primary settling of solids is desirable and sand filtering may be necessary. Special water treatments required for various subsystems will be discussed later. The saltwater should be transferred and stored in non-toxic plastic containers. When in doubt, questionable materials should be tested against embryonic development of the oyster following accepted bioassay procedures.

Fig. 2. Floor plan

Brood stock Maintenance and Spawning

The brood stock maintenance and spawning subsystem is diagrammed in Fig. 3. It shows the role of the adult oyster in the hatchery system. In general terms, adults are taken from the field and returned to the field, less mortalities and those sacrificed for gametes.

SELECTION OF ADULTS In the Pacific Northwest, the Pacific oyster may be induced to spawn and viable gametes may be obtained at any time during the year. Year-round spawning is possible because the Pacific oyster normally does not completely spawn out in all Pacific Northwest estuarine areas, generally too cool to induce complete spawning. To insure a supply of spawners, 300 or 400 oysters should be moved to cool seawater (normally an area of direct ocean influence). Spawners should possess a suitable level of "fatness," or stored glycogen. During the four to eight weeks that the adults are in conditioning trays, they receive little if any food and must rely on stored reserves.

CONDITIONING Bi-monthly, 50 "fat" oysters are placed in the hatchery conditioning trays. These are flow-through aquaria using unfiltered seawater heated to a temperature of 18-20 C. During a four-week period the gametes in these adult oysters will mature and the adults will grow ready for induced spawning. These adults will be prime for induced spawning for the following two weeks. They may be retained an additional two weeks to assure a source of conditioned oysters in the event that problems occur with a succeeding group. Possible problems may include a failure of the oysters to reach spawning condition or a complete spawn-out during the initial four week conditioning period. It is always wise to have more

conditioned adults on hand than are actually needed to supply gametes for the hatchery. After two months in the hatchery, surviving adults may be returned to the oyster beds in the field. They should not be reused for at least six months. Spawned adults would not normally be reused at all for producing wild hatchery seed. In a breeding program, however, spawners should be marked and detailed records kept so that desirable brood stock can be re-spawned to improve the strain of hatchery produced seed. The records should include data about fecundity as well as larval survival and growth rates.

Fig: Broodstock sub-system

INITIATING SPAWNINGSpawning is generally induced in conditioned oysters by manipulating water temperature. Conditioned adults are placed in aquaria with recirculating seawater. Water temperature is raised first to 25 C and then to 30 C over a half hour period. The temperature is then allowed to fluctuate between 25 C and 30 C. This temperature manipulation may initiate spawning of one or both sexes. If spawning does not occur, the aquaria may be drained, left empty for a few minutes, and then refilled with heated seawater. Although the biological basis for it is not clear, this technique seems to stimulate pumping and occasionally spawning by the oysters. If temperature manipulation does not initiate spawning, sex products (either eggs or sperm) should be added to the water. There appears to be a hormone released by spawning oysters that stimulates other adults to spawn. This hormone has not been identified, but it is the basis for stimulating spawning with gametes. The sex products used to stimulate spawning can be obtained in any one of several ways. If one or more oysters spawn by temperature stimulation, they are isolated in small aquaria to collect sex products. These gametes may then be used to induce other adults to spawn. If no adults spawn in this fashion, a

conditioned adult may be sacrificed to obtain gametes to stimulate other conditioned adults. Alternatively, excess gametes may be frozen and stored for future use as a spawning inducement. If the adult oysters have been properly conditioned, these techniques nearly always induce spawning. On rare occasions when spawning does not occur, the procedure should be repeated the following day.

FERTILIZATION Unless mating specific individuals, eggs from at least two females should be mixed. Fertilization takes place by mixing sperm and eggs in the ratio of 2-4 ml of dense sperm suspension to 4 liters of egg suspension (approximately one million eggs). Care should be taken to avoid adding too much sperm to the egg suspension. The presence of excess sperm can result in a condition known as polyspermy, which leads to abnormal embryonic development and poor survival. The fertilized eggs should be passed through an 80 micron screen to remove excess debris. The eggs are afterwards diluted with a known volume of saltwater and a sample of known volume is withdrawn with a pipette. The eggs are counted and an estimate is made of the total egg count. An example may make the sampling procedure clearer: An unknown number of eggs are diluted to 10 liters. After agitation to insure equal distribution of the eggs, a 1 ml sample is withdrawn and further diluted to 100 ml. This subsample is agitated and a 1 ml sample is withdrawn and put in a small dish. The eggs in this sample are counted using a dissecting microscope. Three or more samples are counted and averaged. The average count is multiplied by 1 x 106 to obtain the total number of eggs or by 100 to find the number of eggs per milliliter. Oyster larvae may also be counted by this method. The second dilution may not be necessary. If not, multiply by 1 x 10 4 to obtain the total population. After counting, the fertilized eggs should be diluted to not more than 200 eggs per milliliter and allowed to develop for 24 hours at 25C. After enough gametes have been collected and fertilized, the adult oysters should be placed in cold running seawater. This will usually end the spawning response, although some individuals will occasionally continue spawning until completely spent.

Figure 8. Fertilized egg with first polar body.

Larval CultureTwenty four hours after fertilization, when held at a temperature of 25 C, the fertilized eggs will have developed into swimming, straight-hinged

veligers ready to feed. These larvae now enter the larval rearing subsystem (Fig. 4) and must be provided with cultured algae. The tanks commonly used for rearing the larvae through the free swimming stage are relatively large, at least 500 liters. Exact tank size will be dictated by the individual goals of each hatchery.

Fig. 4. Larval rearing subsystem

Larval careTanks are cleaned, disinfected with sodium hypochlorite (bleach), and filled with treated sea water before they are stocked with fertilized eggs. Tanks should be gently aerated so that eggs and subsequent larvae are mixed throughout the tank. From this point until larvae are ready to set, larval care consists of feeding algae, draining tanks every 2 days (daily as larvae near setting), sieving and counting larvae, cleaning and refilling tanks, and restocking larvae at the appropriat density. Table 1 outlines a schedule for draining, the suggested mesh size for sieving, the larval density, and the food density. Most fertilized eggs develop into trochophore larvae within 12 to 20 hours. These become veliger larvae (also called straight-hinge or D-shaped larvae) within 20 to 48 hours. The first draining and sieving (Fig. 9) is done at about 48 hours. Water is drained slowly through the appropriate size sieve (Table 1) and the retained larvae are placed in a known volume of treated seawater (e.g., 10 L). Several 1-ml samples are taken, the larvae are counted in a Sedgewick-Rafter cell, and the average number is used to calculate the total number of larvae, as in the egg count. Larvae are restocked in a cleaned and disinfected tank filled with treated seawater at the recommended density, five per ml or about 20,000 per gallon. This process is repeated every 2 days (daily as larvae near setting) with appropriate reductions in larval density (Table 1) until larvae are ready to set.

Feeding ManagementOyster larvae feed by filtering small, single-cell algae from the water. They must be supplied with the right size food at a density that makes the food easy to encounter. There are several methods for supplying algae to larvae. The simplest is to coarsely filter (10 to 25 μm) natural waters to keep out zooplankton and large algae and then provide the water directly to the larvae. A second method involves filtering natural water in the same way and then fertilizing it to stimulate algae growth and reproduction. After a significant amount of algae is produced, it is fed to the oysters. Both of these methods have worked for hatcheries but the results can vary considerably; and, the water can be contaminated by unwanted zooplankton or the wrong kinds of algae.A third method is to separately culture several species of algae from pure cultures of each desired species. Algae species that have been used to grow oyster larvae include Chaetocerus gracilis, Isochrysis galbana, Pavlova spp., and Nannochloropsis spp. Several studies have shown that a mix of algae species results in better growth. Culturing algae can be labor intensive, requiring repeated sterilization of glassware as the algae is moved through a series of larger containers. Several continuous culture methods have been developed that can reduce labor and provide larger volumes. See the Additional Reading section for sources of more detailed information on culturing algae.A fourth method is to purchase concentrated algae from commercial producers. While often expensive, commercially produced algae may be cost effective depending on the size of the oyster hatchery.However it is obtained, algae must be added daily to the larval culture tanks at concentrations that result in the densities listed in Table 1. Intensively cultured algae are very dense and often a diluted subsample must be counted. To do this, a drop of diluted culture water is placed on a hemacytometer (a special microscope slide with finely etched squares to aid counting) and the cells within several 1-mm-square areas are counted. The cell count is divided by the number of 1-mm-square areas counted and then multiplied by 10,000 to get the cells per ml. This number is then multiplied by the dilution factor.The volume of culture water needed to achieve the desired density in the larval tanks is determined from the calculated density of algae. For example, if the hemacytometer count shows 100 cells in four 1-mm-square areas, the number of cells per 1-mm-square area is 25. Multiply by 10,000 to get 250,000 cells per ml. If the sample was originally diluted by a factor of 10, multiply by 10 to get 2,500,000 cells per ml in the original culture. The desired density of algae at the beginning of larval culture is 25,000 cells per ml. Suppose the larval tank is 250 gallons (946 L). Multiply the larval

tank volume (946,000 ml) by the desired algae density (25,000 cells/ml) and divide by the density of cells in the algae culture (2,500,000 cells/ml) to get 9,460 ml of plankton culture to be added to the larval tank.

Record keeping and evaluation methodAn oyster hatchery, like any other business, should be constantly evaluated for best use of effort, equipment and space. The method to be described should provide a means of optimizing biological reliability and procedural efficiency in hatchery operation and management. The key to optimization lies in periodically reviewing the biological and procedural data, identifying problem areas and making selected changes in operating routines to solve the problems. After operating according to the altered routine for a period of time, the resulting data should be reviewed to judge the effectiveness of the change. This evaluation procedure should be an integral part of the hatchery routine. Continual improvements must be made to optimize productivity.

HATCHERY ROUTINESThe evaluation operates in a cyclic manner. Periodic review encourages implementation of carefully considered changes in the operating routine (Fig. 11). The routine should be established with a daily schedule of duties accompanied by a thorough description of each task. Operation of the hatchery is broken down into units of work, or tasks. Each task should be defined in such a way that time expended on it can be identified exclusively with that task. A daily record is kept of times spent on each task. Table 3 shows a hypothetical breakdown of the proportion of time expended on various tasks. The table is meant only as a guide to show how a technician's time may be recorded; actual times will vary among workers and hatcheries. If excessive time is indicated, techniques should be streamlined.SignificanceOysters are an important component of seafood production and provide widely appreciated ecological services. Oyster hatcheries can produce oysters for commercial culture operations, restoration projects, and a variety of basic and applied research projects. Oyster hatcheries have played an important role in breeding disease-resistant oysters, triploid and tetraploid oysters, and faster growing oysters.

Hatchery management of Mud Crab

The mud crab (Scylla serrata) is a highly regarded and valued table food item in both Australia and Asia. Mud crab aquaculture is not currently undertaken in the NT but it has been carried out in a number of Asian countries for many years. Mud crab farming in these areas is generally based on catching juveniles from the wild and using them to stock into mangrove enclosures, pens or ponds for grow out. Aquaculture can supply

crabs to a range of markets, such as the regular hard shelled crabs and niche markets for females with mature ovaries, and soft shelled crabs.

Site selectionIt is rare that a hatchery is sited in an optimal location. More commonly, it is acompromise based on land availability, cost, existing infrastructure and proximity orlogistical connections to grow-out areas. The basic attributes required for a mud crab hatchery site include:• An unpolluted source of marine seawater and freshwater;• Ability to discharge treated hatchery wastewater streams;• A site with land suitable for construction of hatchery buildings;• Access to reasonable transport arrangements for staff and products.For a mud crab farming venture (or its component parts) to be viable, it is essentialThat logistics are such that they do not impinge on its ongoing operation. Factors to beConsidered include:• transport (air, sea and road);• Availability of staff;• Accommodation;• Political stability;• Supplies;• Services available;• Power and water supply;• Proximity to markets;• Potential for flooding or other natural disasters to affect operations.The cost of establishing and operating a mud crab farm can vary significantly depending on where it is sited; typically, the more remote the location chosen, the more expensive it is. These costs may be offset by other factors such as cheap labor, outstanding growing conditions or other special circumstances. The preparation of a detailed business plan that takes these factors into consideration is strongly recommended in order to ensure the underlying viability of a business is not compromised by the logistics of its operation.

Basic infrastructure

Water ManagementWater sources utilized should be free of significant pollution and within the pH range 7.5–8.5. This pH recommendation is based on the requirements of marine shrimp, as little work has been undertaken on the effect of pH on mud crab growth and survival. For pond farms, both a brackish to marine source of water and a separate freshwater source are ideal to manage water salinity at the preferred level. The daily requirements for a farm requiring pumped water need to be calculated, and potential pump sites examined to ensure that sufficient quantities of water will be available for the size of the farm being planned. Factors such as the availability of water for pumping at different phases of the tide will need to be included in the calculations. Similarly, the availability of

freshwater resources, which vary throughout the year in response to local rainfall patterns, should be examined. Freshwater for salinity control is most likely to be required in the driest times of the year. As mud crabs often live in areas of turbid coastal waters, high turbidity is not a major issue, with the exception of water required in hatcheries. However, the use of sand or other filtration methods can reduce highly turbid water to water suitable for hatchery and live feed production. While mud crabs can survive a wide salinity range in culture (5–40 ppt), optimalgrowth appears to be in the range of 10–25 ppt for S. serrata, although research has not been undertaken for all species, for the entire size range of each species and certainly not from all countries where they are grown. conditions. In northern Australia, optimal growth for S. serrata was at a temperature of 30 oC, with good growth from 25 to 35 oC.

Hatchery operations, procedures and equipment

QUARANTINEAny potential broodstock for a mud crab hatchery should be examined carefully before they are placed in holding tanks. Only crabs that are in good condition, have no missing limbs, no necrotic spots on their shell and are not carrying any fouling or parasitic organisms should be used. Female crabs carrying an egg mass should preferably not be brought into a broodstock facility. This is because the egg mass of these crabs was spawned in the wild and, as such, its disease status would be unknown, as would be the nutritional status of the female carrying the eggs. However, berried mud crabs (females carrying an egg mass) can be utilized. If berried, those crabs carrying egg masses with brown, grey or black eggs are closest to hatching and will not have to be kept in hatching tanks for too long. To mitigate against the risk of transferring exogenous pathogens into the hatchery, all new broodstock should be bathed in an appropriate disinfectant before placement in a broodstock tank. While formalin (40 percent formaldehyde) at 150–200 ppm for 30–60 minutes is the most common treatment, other chemicals such as potassium permanganate, malachite green and methylene blue have been used, with regular treatment every 2–3 days for 15 minutes recommended. Such treatment has been found to have no detrimental effect on either the mud crabs, their eggs or larvae.

BROODSTOCK SELECTIONBroodstock can be sourced from the wild, from pond-reared animals or from domesticated improved broodstock. Although wild broodstock currently have better reproductive performance than pond-reared broodstock, those reared in ponds are still of significant commercial value. This highlights the need for improved broodstock nutrition for pond-reared broodstock, so that they can match the performance of wild broodstock. While significant research has been undertaken on the genetics of mud crabs, little work has been done on domesticating stock and rearing for improved characteristics. From an operational perspective, on entering the hatchery, broodstock from any source should be treated the same.

Mud crab broodstock can be held in broodstock tanks at densities of 1–5/m2, depending on crab size. When mature female crabs, as assessed by the appearance of the abdominal flap are sourced from the wild or from ponds, they are generally already mated and fertilized. There is, therefore, usually no need to hold male broodstock. However, when immature females are used, they will readily mate with any male provided when they moult to maturity.

INCUBATION AND HATCHINGOnce mud crabs are carrying an egg mass, they no longer need to be fed. It has been found that a 2 °C difference in broodstock water temperature can lead to problems with zoeal viability. While mud crab broodstock and larvae can be successfully kept at temperatures of 25–32 °C, it is recommended that the water for all tanks be kept within as small a temperature range as possible. Such an approach has also been found to produce improved results in shrimp hatcheries, where operators typically try to maintain temperatures within ±1 °C. The embryonic development of Scylla spp. has been described with a 5, 9 or 10-point scale; however, from a practical perspective, it is critical that hatchery staff have live feed and tanks prepared in time to look after the larvae as soon as eggs hatch. To that extent, it is critical that hatchery staff are familiar with the typical time from spawning to hatching, at any given temperature, and in particular can recognize the pre-hatching phase of development. From a practical perspective, simple systems are best, so the most useful scale to use is probably the five-point scale of Thach (2009) (Table 5.1). Egg health and development can be assessed by quick observation of the egg mass and excising several small bunches of eggs from different areas of the egg mass for observation under a low-power microscope.

MonitoringA monitoring programmed needs to be established and maintained throughout each larval rearing period. Parameters to be monitored and recorded for each larval rearing tank include:• Water temperature• Water salinity• Water treatments• Tank cleaning• Larval stage• Condition/appearance of larvae• Behavior of larvae• Density of larvae• In-tank feed densities• Addition of feed• Chemical treatments

Maintaining larval water qualityWater treatment for mud crab larval rearing varies with location and availability of filtration equipment. Where the water quality of incoming

seawater available to hatcheries is suboptimal, treatment by chlorination can be used, followed either by chemical de-chlorination, or aeration for 2–3 days to remove residues. Other water treatment options for mud crab larval rearing water include ozone treatment (followed by carbon filtration), UV sterilization, microfiltration and microbial conditioning. Over the duration of a mud crab larval rearing run, waste feed and metabolites will build up in the tanks. If the tanks and water are not adequately maintained, water quality can deteriorate and bacterial levels increase, which can affect larval survival. During mud crab larval rearing, draining down tanks and replacing water with fresh high-quality water is routine. This can be undertaken as frequently as every day, to once every five days. The percentage of water changed varies from 30–70 percent of the tanks’ total water volume. The frequency and volume of water exchange may be linked to the monitoring of water parameters or the level of particular pathogens, e.g. luminescent bacteria. During drain-down, tank walls can be cleaned with sterile wipes or sponges, although the ease with which this can be undertaken depends on the size of larval rearing tanks used. In addition, dead zoea, excess feeds and any wastesthat settle within tanks can be removed by siphoning. Another approach to maintain water quality in larval rearing tanks is to establish a recirculating system, which screens and treats water in tanks. Such systems need to be able to screen mud crab larvae and feed so that they remain in the tank. Such recirculating systems are not common in mud crab aquaculture at present, but as they can provide enhanced water quality compared with other systems, and mud crab larvae are sensitive to water quality, this may change in the future. An approach intermediate to water exchange or recirculation is a system set-up where water is flowed through larval rearing tanks, constantly refreshing water quality in the tanks, limiting the buildup of metabolites and the concentration of potential pathogens. As biofilms, which can contain bacterial pathogens, can build up on the walls and floors of larval rearing tanks, it is advisable to keep larvae dispersed in the water column. To do this, gentle, non-turbulent in-tank aeration, and directional flow of water within tanks should be established. Simple airlifts, which keep larvae off the bottom of tanks, are a useful tool in this regard. While nitrite has been shown to be toxic to mud crab larvae, the levels at which it is toxic (4–7 mg/liter for different larval stages) are approximately an order of magnitude higher than nitrite levels commonly found in mud crab larval systems (<0.5 mg/litre) and, therefore, it is of little concern to commercial hatchery operators. Similarly, mud crab larvae are not affected by the levels of ammonia routinely found in mud crab larviculture systems, which are also well below toxic levels.

DISEASE MANAGEMENT AND TREATMENT IN MUD CRAB FARMING

A limited number of treatments have been developed to assist mud crab farming operations. Much of the work in disease management has been directed to hatchery operations to improve mud crab larval survival, where the control of both bacterial and fungal infections has been critical.

Improved pond management is the other area where substantial improvements can be made. While prophylactic treatments, such as antibiotics for bacteria, or fungicides for fungi, have been used successfully to improve survival in mud crab larval systems, more progress has been made by the development of improved culture systems that reduce the risk of such infections in the first place. The use of any chemical to treat a disease must be within government regulations controlling their use and under the supervision of trained staff.

Hatchery management of MusselMussel is the common name used for members of several families of clams or bivalvia mollusca, from saltwater and freshwater habitats. These groups have in common a shell whose outline is elongated and asymmetrical compared with other edible clams, which are often more or less rounded or oval. The word "mussel" is most frequently used to mean the edible bivalves of the marine family Mytilidae, most of which live on exposed shores in the intertidal zone, attached by means of their strong byssal threads ("beard") to a firm substrate. A few species (in the genus Bathymodiolus) have colonised hydrothermal vents associated with deep ocean ridges. In most marine mussels the shell is longer than it is wide, being wedge-shaped or asymmetrical. The external colour of the shell is often dark blue, blackish, or brown, while the interior is silvery and somewhat nacreous. The word "mussel" is also used for many freshwater bivalves, including the freshwater pearl mussels. Freshwater mussel species inhabit lakes, ponds, rivers, creeks, canals, grouped in a different subclass, despite some very superficial similarities in appearance. Freshwater Zebra mussels and their relatives in the family Dreissenidae are not related to previously mentioned groups, even though they resemble many Mytilus species in shape, and live attached to rocks and other hard surfaces in a similar manner, using a byssus. They are classified with the Heterodonta, the taxonomic group which includes most of the bivalves commonly referred to as "clams".

Site selectionThere are many critical elements to a mussel hatchery, but none is more important than location, or more specifically, location relative to water supply. Oyster hatcheries require large volumes of clean sea water with salinities in the range of 15 to 30 parts per thousand (ppt). Salinity is not a major issue in many areas, but some estuaries have periodic episodes of freshwater inflows that can reduce salinity below 10 ppt. Low salinity water is not conducive to spawning, larval development, or early growth of young oysters. Turbidity, potential pollutants, watershed development, boat traffic, and natural algae production are other aspects of water quality to consider. The local agency that oversees oyster harvesting areas should be consulted in regard to regulations that might affect the use of oysters produced from the proposed hatchery site.

DEVELOPMENT OF INFRASTRUCTURE AND FACILITIES

Although the existing hatchery and the support system for live organisms in Kantian are well developed, improvement is needed for the efficient implementation of the finfish programme, especially the mass seed production of marine finfish in the future.

Recommendations for improvement are outlined below:

a) Brood stock development tank

Modifying one 660 m outdoor concrete tank at the seafront for holding the broodstock in addition to the existing cages.

Dividing the tank into 4 equal compartments with polyethylene netting and wooden frame.

Installing one unit of 4-inch diameter pump at the edge of the tank to facilitate daily water changes at 50–60% per day.

Providing the tank with 1 unit of 2-inch air blower for water circulation and aeration.

b) Hatchery and nursery complex

The existing holding tanks in the hatchery are large (20–40 m3) and deep (2 m), and are suitable for brood stock conditioning, gonadal maturation trials and spawning tanks. Outdoor concrete tanks are suitable for live food organism production. It is essential to develop the 150 m unused land adjoining the existing hatchery into a larvae-rearing and nursing complex, which should consist of 20 units of 1 m3, 20 units of 2 m3 and 10 units of 4 m3 fiberglass tanks for larval-rearing purposes.

c) Water intake system

Since the hatchery needs a large quantity of seawater, it should have its own pumping system.Two types of water-intake systems are recommended:

Pumping seawater directly from the sea. Pumping seawater through sump pit.

d) Water supply system

Although the seawater in front of the hatchery is very clear, filtered seawater is necessary for larviculture. The layout of the water system is

shown in Figure 2. Unfiltered seawater will be used for the maturation and brood stock tanks and filtered seawater for larval-rearing.

LARVAL FEED DEVELOPMENT

One of the key factors to ensure success in marine finfish hatchery operations is the timely supply of the necessary food organisms in sufficient quantities. Ways of assuming continuous sources and mass production techniques are discussed below:.

CRITERIA FOR LARVAL FEED SELECTION

Feeds suitable for fish larvae are characterized as follows:

a. They should be accepted by the fish.b. The feed should be of a size which can be eaten easily by the

larvae.c. The feed should have high dietary value especially in highly

unsaturated fatty acids (HUFA), essential to the growth and survival of the larvae.

d. The feed should be easy to mass-produce in large quantities.

BROODSTOCK

A sufficient supply of broodfish is essential for a successful induced breeding operation or artificial propagation. There are two sources of finfish broodstock: wild stock and those from ponds or cages. The disadvantages of wild stock is the uncertainty of capturing them, while the advantage of pond or cage reared broodstock is that they are already accustomed to culture conditions and consequently easier to develop into suitable broodfish.

SELECTION OF SUITABLE BROODSTOCK

Fish selected for broodstock should be fast-growing, active, and among the largest and strongest individuals of their age group, and free of parasites and disease.

BROODSTOCK MANAGEMENT

Gonad development is affected by nutrition (food) and environmental factors indicated below:

Nutrition

There is paucity of information on the nutritional requirement of broodstock and suitable practical diets. Standard practices for feeding broodstock are not well documented. At present, broodstock is fed following traditional or empirical lines. The formulated feed used are

generally those commercially available as feed for rearing fish to marketable size.

Data accumulated to date indicate that poor nutrition can result in poor or negative reproductive performance and that lack of a vitamin supplement can affect sperm quality. Reliance on natural food may also lead to poor or variable reproductive performance. It has been shown that fatty acids, especially in the case of ovarian lipids, tend to utilize the highly unsaturated fatty acids.

Environment

- Photoperiod

One of the factors considered of great importance to the inducement of sexual matuation and spawning is photoperiod. Photoperiod manipulation is now being employed to alter the normal production of a cultured fish species, for example, mullet, rabbitfish, rainbow trout, tilapia, carp and catfish. The greatest advantage of altering the spawning time of the cultured species is the availability of fry for stocking in ponds, pens and cages throughout the year.

- Temperature

Water temperature is another important factor which influences the maturation and spawning of fish. Data accumulated to date show that the functional maturity in some species of fish is directly controlled by temperature; in others, the time of spawning is regulated by the day-length cycle, and occurs at the time when temperature is optimum for survival and food supply is adequate.

- Salinity

Some species of fish, e.g., salmon, migrate from the marine to the freshwater environment in order to spawn, while others, such as eels, migrate from freshwater to the marine environment to complete their reproductive cycle. This definitely shows that salinity is related to maturation and spawning. Salinity may influence gametogenesis but probably does not function as a synchronizer for the timing of maturation.

- Other environmental factors

Aside from photoperiod, temperature and salinity, other less obvious factors may affect the maturation and spawning of broodstock, such as rainfall, stress, sex ratios, stocking density, isolation from human disturbance, dissolved oxygen, social behaviour of fish, heavy metals, pesticides, and irradiation.

SPAWNING

At present, two major techniques are employed in the mass-production of marine finfish fry in Southeast Asian countries: artificial fertilization and induced spawning.

Artificial fertilization

Spawners are caught in natural spawning grounds near the mouth of rivers or in saltwater lakes. The degree of maturity of the collected spawners is immediately checked. The dry method of fertilization is normally used. The eggs are stripped directly from the female into a dry and clean container where the milt is added. A feather is used to mix the milt and eggs for about 5 minutes. Filtered seawater is added to the mixture while stirring, and it is then allowed to stand undisturbed for 5 minutes.

The fertilized eggs are then transported to the hatchery for subsequent hatching.

Figure 4. Flow of operational processes in a bivalve hatchery.

Induced spawning by hormone injection

In induced spawning, the hormones used include the following:

SPH - acetone dried pituitary gland homogenate of coho salmon prepared by the British Columbia Research Council, Vancouver, Canada; 1 g powder contains 17.6 mg gonadotropin.

HCG - human chorionic gonadotropin, manufactured by Ayerst Laboratories, New York.

Before injection, HCG is dissolved in 3 ml of its accompanying diluent. The solution is then used to homogenize the acetone dried pituitary gland of salmon to be used for induced spawning.

Fig: Seed production of mussel.

FERTILIZATION AND INCUBATION

The fish that are induced to spawn by hormone injection will be ready to spawn within 9–12 hours after the final injection. The schedule of injections for subsequent spawning must be synchronized with the natural spawning time of the fish which occurs in late evening between 18.00 and 24.00 h. On the other hand, with the stripping method, it is still necessary to extract the eggs from gonads by cannulation and examine them under microscope. The fish has spawned only if at least 40% of the eggs are transparent. Stripping is always done by gently pressing the abdomen with the thumb and forefingers, beginning to apply pressure just foreward of the genital pore. The eggs are fertilized immediately after stripping, using the dry method, the milt being hand-stripped from the hormone-treated male. The eggs and milt are mixed gently but thoroughly using turkey feathers. After at least 3 minutes, seawater (34 ppt) is added to the mixture while stirring it. After another 3 minutes, the fertilized eggs are transferred in a scoop net (mesh size = 500 micron) and washed thoroughly with seawater isohaline in the incubation tanks. The incubators are strongly aerated to prevent the eggs from sticking together. The eggs are incubated at ambient temperature ranging from 25° to 30°C and at a salinity of 34 ppt. Six hours after the start of incubation, dead eggs are removed from time to time by stopping the aeration for about 5 minutes. Fertilized eggs float in seawater with a salinity of at least 34 ppt while unfertilized eggs sink.

LARVAL Management

Larvae of red seabream and black seabream were used to demonstrate the larval-rearing technique at the Centre. The rearing tanks are made in plastic in a circular shape. Volume ranges from 1 to 10 m. The tanks are usually protected from sunshine and heavy rain. Five hours before hatching, the developing eggs are transferred to larval-rearing tanks. The tanks are gently aerated. The larvae start to hatch 16–25 hours after fertilization, depending on the temperature and species. The usual stocking density of developing eggs is 100–200 eggs/litre.

Rearing environment

Good quality seawater at 30–31 ppt is required for larval-rearing. Water temperature is also important and should range from 26° to 28°C to promote fast growth of larvae. Larva, tanks are prepared one or two days prior to the transfer of newly-hatched larvae. Filtered seawater is added to the tanks and very mild aeration is provided. After stocking, unicellular algae (Tetraselmis sp. or Chlorella spp.) are added to the tank and maintained at a density of 8–10 × 10 or 3–4 × 10 per ml for Tetraselmis sp. and Chlorella spp. respectively. These algae serve a dual purpose: as a direct food to the larvae and other rotifers and as a water conditioner in the rearing tank. The day following stocking, the bottom of the larvae-rearing tank should be cleaned, and every day thereafter. This is done by siphoning off unfertilized eggs, faeces, dead larvae and uneaten food accumulating on the bottom of the tank. About 20% of the tank water is changed daily for the first 25 days of the rearing period, then increased to 40–60% per day for the remaining culture period. Since seabass can also be cultured in freshwater, it is recommended to reduce the salinity of the rearing water when the larva is still in the hatchery, before it is transferred to the freshwater environment. Beginning with the twentieth day, salinity can be reduced gradually until a freshwater condition is reached on the twenty-fifth day.

RECOMMENDATIONS

However, for the benefit of the Government, and to fully utilize the equipment and manpower, the project should include studies on induced breeding of other economically important finfish species, since the spawning seasons differ from one species to auother. The assignment of an international expert to work full time in the field with the local staff would be useful. Existing facilities, especially the hatchery and nursery system, should be further improved in order to facilitate the Centre's mass production of marine fish fry. The 150 m2 unused land adjoining the existing hatchery should be developed into the nursing area to accommodate the fibreglass nursery tanks which have been ordered by FAO/UNDP. The existing hatchery facilities will be used for maturation and spawning trials. The 660 m2 outdoor concrete tank at the seafront should be modified to maintain the broodstock including division of the tank into

four equal compartments of polyethylene netting with wooden frame. The advantage of this tank is that the fish can be reared the whole year round without risking damage by the typhoons. Broodstock of economically important species is needed to stock the tank, one species per compartment. An inter-disciplinary and team approach should be adopted to implement the project activities instead of having one person in charge of one species. Since there are many interdependent factors, a number of parameters should be simultaneously studied, in order to obtain a valid result and to fully utilize the available facilities and manpower.

Reference

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