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5 The post-spawning process


The post-spawning process includes facility maintenance, water quality management; broodstock handling; washing, selection, holding and transport of nauplii; postlarval rearing, maintenance, health management, assessment of condition, selection and risk assessment for stocking, shipping and transfer; and documentation and record keeping.

5.1 Facility maintenance

To achieve consistent production of high quality larvae, the production facilities must be maintained in optimal condition

Facilities must be maintained so as to optimize the conditions for the growth, survival and health of the shrimp broodstock, larvae and PL, minimizing the risks of disease outbreaks. In order to facilitate this, a set of protocols must be drawn up by the hatchery management as part of the Standard Operating Procedures (SOPs) and followed strictly by all staff members at all times. The hatchery's SOPs should include procedures for a sanitary dry out following each cultivation cycle (for larval rearing), or at least every three to four months (for maturation facilities), with a minimum dry period following cleaning of seven days. This will help prevent the transmission of disease agents from one cycle to the next.

All tanks and equipment should be thoroughly cleaned on a regular basis, cleaned and disinfected after use, and cleaned and disinfected again before starting a new production cycle

Tanks used for broodstock spawning, egg hatching, and holding of nauplii and postlarvae should be thoroughly cleaned after each use. The procedures used for cleaning and disinfection are basically the same for all tanks and equipment. They include scrubbing with clean water and detergent to loosen all dirt and debris, disinfecting with hypochlorite solution (20-30 ppm active ingredient) and/or a 10% solution of muriatic[8] acid (pH 2-3), rinsing with abundant clean water to remove all traces of chlorine and/or acid, and then drying. The walls of tanks may also be wiped down with muriatic acid; outdoor tanks and small tanks can be sterilized by sun drying.

The following points should be considered:

· Tanks should be washed and disinfected at the end of every production cycle.

· All hatchery equipment should be regularly cleaned and disinfected.

· Concrete tanks painted with marine epoxy or plastic-lined tanks are easier to clean and maintain than bare cement tanks.

· After harvesting the larvae from a larval rearing tank, the tank and all of its equipment should be disinfected. Similarly, once all of the tanks in a room have been harvested, the room itself and all its equipment should be disinfected.

· Tanks can be filled to the maximum level and hypochlorite solution added to achieve a minimum concentration of 20-30 ppm active ingredient. After 48 hours, the tanks can be drained and should be kept dry until the next cycle starts.

· All equipment and other material used in the room (filters, hoses, beakers, water and air lines etc.) can be placed in one of the tanks containing hypochlorite solution after first cleaning with a 10% muriatic acid solution.

· Broodstock maturation tanks and all associated equipment should be cleaned and disinfected following a typical operation period of three to four months.

· Water pipes, air lines, air stones etc. should be washed on a monthly basis (or during dry-out) with the same chlorine concentration and/or a 10% solution of muriatic acid (pH 2-3) by pumping from a central tank.

· All hatchery buildings (floors and walls) should be periodically (once per cycle is recommended) disinfected.

· All other equipment should be thoroughly cleaned between cycles.

· Before stocking tanks for a new cycle, they should once again be washed with detergent, rinsed with clean water, wiped down with 10% muriatic acid and once more rinsed with treated water before filling.

· Disinfection procedures may require adjustment according to the special needs of the facility.

· Appropriate safety measures must be taken when handling the chemicals used for disinfection. Procedures regarding chemical usage and storage, wearing of protective gear etc. should be included in the hatchery's Standard Operating Procedures (SOPs).

Recommended products, concentrations and frequencies for the disinfection of various hatchery items are also given in OIE (2003).

5.2 Water quality management

The hatchery infrastructure should allow appropriate cleaning and disinfection of the incoming water

Incoming water should be cleaned and disinfected through chlorination and filtration before being distributed to different working areas (hatchery, algal culture, Artemia etc.). Distribution should be designed to avoid the risk of cross-contamination. Water and air distribution systems should be designed to allow for pumping of disinfecting solutions through the system and to permit complete drying during dry-outs.

Ideally, maturation and larval rearing facilities should be built to take advantage of a supply of oceanic water

However, it is possible to use seawater brought into the facility from elsewhere or suboptimal seawater with the appropriate filtration and disinfection techniques. In general, a closed recirculating system is more biosecure than an open water system, but requires additional biological and mechanical filtration and disinfection to maintain optimal water quality.

"Sub-sand beach wells" should be used for primary water filtration

The most common system for filtration of raw seawater entering the hatchery from the sea is through the use of "sub-sand beach wells". These consist of a series of filter galleries, well-points, wells, tips etc. that allow primary filtration before entering the hatchery. They also limit fouling organisms, pathogen hosts, red tides and some pathogens, which direct intake systems do not.

Water carrying a high sediment load should be passed through sedimentation tanks to remove suspended solids

A minimum storage capacity of 50% of the total capacity is required when the reservoirs can be refilled twice daily.

Incoming water should be disinfected to destroy any remaining pathogens and any heavy metals present removed by chelation

Calcium (or sodium) hypochlorite (10 ppm active ingredient for not less than 30 min), and/or ozone, or UV light should be used to disinfect the incoming water after initial filtration and sedimentation. After treatment with chlorine, the water in the reservoir must be checked by ortho-toluidine (3 drops in 5 mL of water sample) to ensure no chlorine residual remains (indicated by a yellow colour) before the water is used. A chart or whiteboard must be provided giving the date and time of treatments and the results of these tests signed by the person who is responsible for the water treatment. Once the chlorine has dissipated or been neutralized with sodium thiosulfate (1 ppm for every 1 ppm of chlorine remaining), EDTA can be applied to chelate any heavy metals present (quantities depending on concentrations of heavy metals and use).

The temperature of the water should be adjusted before it enters the production units

A boiler and heat exchange system, typically located between the reservoir and production units, may be required to adjust water temperatures to within the range required (generally between 28 and 32 °C depending on area and stage, see Table 4).

The water filtration system following the reservoirs should consist of sand filters, activated carbon, and other filtering elements such as cartridge filters or membrane filters for water uses requiring fine filtration.

Sand filters must be properly maintained

Sand filters must be backwashed at least two times per day (or as required based on the suspended solids loading of the incoming water) for a sufficient length of time to assure the cleaning of the filter.

Being able to open the filters to check for channelling and thorough backwashing is an advantage. At the beginning of each production cycle, the sand must be replaced by clean sand that has been previously washed with sodium hypochlorite solution at 20 ppm active ingredient or 10% muriatic acid solution (pH 2-3). Activated carbon should be replaced at least once every hatchery cycle to maintain efficiency.

Cartridge filters must exchanged daily

For cartridge filters, two sets of filtering elements must be available and these sets should be exchanged every day. Used filters are washed and disinfected in a solution of calcium (sodium) hypochlorite at 10 ppm active ingredient or 10% muriatic acid solution for one hour. Some filter materials are sensitive to muriatic acid and thus care must be taken when this disinfectant is used.

The filters are then rinsed with abundant treated water and dipped in a recipient containing a solution of 10 ppm sodium thiosulfate to neutralize chlorine (if used). Two or more new sets of filters should be used for each hatchery cycle, depending upon the suspended solids loading of the seawater.

The recommended final size of filtration depends on the uses of the water as shown in Table 4.

Table 4. Recommended water filtration standards and water temperatures for different hatchery needs.

WATER USE

FILTER SIZE
(mm)

TEMPERATURE
(ºC)

Maturation

15

28 to 29

Hatchery

5

28 to 32

Spawning and hatching

0.5-1.0

29 to 32

Algae culture (indoor/pure)

0.5

18 to 24


If recirculation systems are employed, additional biological filtration should be used

To prevent cross-contamination between different areas of the hatchery, separate recirculation systems should be used for each area requiring them Water recirculation systems are the most efficient systems for broodstock maturation, as they reduce the need for water replacement and residual water discharge. Recirculation systems help maintain stable physical and chemical parameters in the water and also help concentrate mating hormones in maturation, as well as providing better biosecurity.

If recirculation of seawater is required for any area of the hatchery, additional biological filtration will be required to remove dissolved organic material. There are many types of biofilters, all of which incorporate living elements (denitrifying bacteria) that must be cultivated or "spiked" (additional biological material added to the filter) prior to use, so that their effects are optimized at all stages of the cycle. They also require periodic cleaning in a way that does not kill their beneficial bacterial inhabitants.

The water to be used in the spawning and hatching tanks and pure algal culture facilities must be the same quality

The spawning and hatching tanks and pure algal culture facilities must receive water of the same quality and treated in the same way as the water used in the maturation and larval rearing units (i.e. with the addition of UV light sterilization and filtration to 0.5 or 1 µm). Additionally, for hatching and spawning, EDTA is often required at up to 20-40 ppm to ensure heavy metals are chelated and made unavailable, and Treflan at 0.05-0.1 ppm is usually used to combat fungi.

Water distribution should be designed so that each area of the hatchery can be disinfected separately

Water distribution from the reservoir to the various areas of the hatchery should be designed so that each area can be disinfected without compromising the other areas. In this way, regularly scheduled disinfections can be accomplished at times appropriate to each area and cross-contamination between areas can be avoided. Temperature and salinity regulation may vary between different sectors and is facilitated by well-designed distribution systems. In addition, each area has specific filtration requirements, which can be established prior to point of use, appropriate to each area of the hatchery. Pumps, pipes and filtration equipment should be sized so that maximum expected water exchange rates can be maintained to ensure optimal conditions are met at all times.

5.3 Broodstock disinfection

After removing the spent females from the spawning tanks, they should be immersed in iodine-PVP (20 ppm/15 sec) before returning them to the tank of origin.

5.4 Washing of nauplii

Harvested nauplii at stage 4 can be treated by bath immersion in Treflan (0.05-0.1 ppm) to prevent fungal contamination, followed by a thorough wash in filtered and sterilized water and a dip in an iodine-PVP solution (50-100 ppm for 1-3 min) or chloramine-T solution (60 ppm for 1 min), followed immediately by further washes with clean seawater.

Other washing steps have also been described using formalin and iodine-PVP. Chen et al. (1992) and Brock and Main (1994) described a method in which the nauplii are given a 30 second dip in both formalin (300 ppm) and iodophore (100 ppm) before rinsing with filtered sterile seawater for three minutes prior to stocking. This can be effective in removing debris and fouling organisms such as bacteria and protozoa, and may minimize the transmission of viral diseases.

5.5 Selection of nauplii

Nauplii should be harvested using a light to attract them to the water surface

As the nauplii display strong positive phototaxis, healthy nauplii can be harvested using a light to attract them to the water surface. Those that remain at the bottom of the tank are discarded, reducing the percentage of weak and deformed nauplii. After harvesting, the number of good nauplii is counted to provide the hatching rate. In good batches, the hatching rate should be >70%. If lower rates are encountered, a decision is made as to whether the whole batch should be discarded and investigations initiated to discover the cause of the problem.

The activity and colour of the nauplii should be evaluated and the percentage of deformities estimated

A deformity rate of <5% is generally considered acceptable. An estimate is made of the naupliar condition using the extent of the positive phototaxis. To carry out this test, a sample of larvae is placed in a translucent container next to a light source and the displacement of the animals is observed. If 95% or more of the larvae move strongly towards the light, the batch is good; it is intermediate if 70% or more respond, and poor if less than 70% move towards the light. Poor batches may be discarded, depending upon the selection criteria of each hatchery.

5.6 Holding of nauplii

Harvested nauplii must be held under optimal conditions until they are stocked

The harvested nauplii can be held at a density of 20 000-40 000/litre, with continuous light, clean water and aeration until they are ready to be stocked in hatchery tanks. As with eggs, nauplii (stage 4) can be treated with Treflan and/or disinfected. The equipment and material used to harvest the nauplii must be washed daily with a calcium (sodium) hypochlorite solution (30 ppm active ingredient) to prevent contamination of subsequent batches.

5.7 Transportation of nauplii

The nauplii should be transported at densities of 15 000 - 30 000 nauplii/litre, depending on distance or time to the hatchery

Transportation is normally done in double plastic bags containing 10-15 litres of water and filled with pure oxygen. The bags are then packed in cardboard and/or styrofoam boxes, although plastic buckets and tanks are sometimes used. The temperature of the packing and shipping water is adjusted from 28-30 oC down to between 18 and 25 oC (and sometimes not at all), according to the travel time and distance to the receiving hatchery. Salinity is maintained at 32-35 ppt. Upon arrival at the purchasing hatchery, the nauplii should again be disinfected.

If possible, the transport vehicle should first be disinfected before entering the hatchery facilities. After unpacking the nauplii, the packing material must be incinerated.

5.8 Larval rearing and maintenance

Larval rearing should produce the best quality, high-health postlarvae possible

In many cases, where hatcheries and farms form distinct economic units, larval quality is often sacrificed for economy. However, in reality the most economic strategy is to produce postlarvae that will grow quickly, are free from disease and that will give a high survival and production rate in the grow-out facilities. In order to achieve this, all areas involved in larval rearing must be designed for optimal efficiency, cleanliness and productivity.

Entrance to the larval rearing area(s) should be restricted

Entrance to the larval rearing area(s) should be restricted to the personnel that work in these areas. Sanitary mats or footbaths containing a disinfectant solution (e.g. calcium or sodium hypochlorite solution, >50 ppm active ingredient) must be placed at the entrance of each room of the hatchery. The disinfectant solution must be replaced as necessary. At each entrance to the larval rearing room(s), container(s) with iodine-PVP (20 ppm) and/or 70% alcohol should be available and all personnel must wash their hands in the disinfection solution(s) on entry to, and exit from, the rooms.

Each room should have a complete complement of materials for routine operation.

Each room should have a complete complement of materials for routine operation (filters, meshes, buckets etc.). A tank (500-600 litres) containing disinfectant (hypochlorite solution, 20 ppm active ingredient) should be provided to disinfect hoses, buckets, etc. Common-use equipment can be placed in this disinfecting tank at the end of every day and rinsed before re-use the following day. The disinfectant in this tank should be replaced daily or as required.

All materials and equipment should be for the exclusive use in each room, and should not leave the room or be used elsewhere

Additionally, beakers, nets etc. used for each tank should be maintained in a bucket filled with sodium hypochlorite solution (20 ppm active ingredient) and dedicated to that one tank to prevent cross-contamination between tanks within the same unit.

Larvae and postlarvae should be routinely checked for quality

Samples of larvae and postlarvae for routine checking should be taken in disposable plastic containers (paper cups or 300 mL plastic beakers) that are disposed of once used. After the daily check is complete, the larvae or postlarvae should be discarded into a plastic container with sodium hypochlorite (20 ppm active ingredient) or another suitable disinfectant. Larvae and postlarvae used in the daily checks must never be returned to the larval rearing rooms or larval tanks.

The infrastructure for larval culture should include of one or more units of conical or "V"-shaped larval rearing tanks

The infrastructure for larval culture consists principally of one or more units of conical or "V"-shaped larval rearing tanks (the tanks are sometimes in two phases: one from nauplius to PL4-5, then larger, flat bottomed tanks or raceways for postlarvae or nursery culture). Supporting infrastructure (discussed in more detail elsewhere) includes a water storage, treatment, heating and distribution system; an aeration system; live feed production facilities for algae and Artemia (and others); laboratories for health checks, bacteriology and feed preparation; offices and an area for packing and shipping postlarvae.

5.9 Larval nutrition and feed management

High standards of feed preparation must be maintained

All feed preparation, especially of live feeds (algae, Artemia and others), is a critical control point (CCP), because feed can be contaminated through inappropriate handling. All sources of live, fresh or frozen food should be considered from the point of view of pathogen risk. The source, treatment, storage and use of feed items should be reviewed and steps taken to ensure that they are safe and properly handled.

Entry to the algal culture and Artemia culture rooms must be restricted to authorised personnel

Staff from these areas should not be able to enter other production areas. At the entrance of each room a footbath containing a disinfecting solution (calcium (sodium) hypochlorite, >50 ppm active ingredient) should be placed. This solution must be replaced as often as necessary. As in other areas, container(s) of disinfectant solution (20 ppm of iodine-PVP and/or 70% alcohol) should be placed at the doors and all staff should wash their hands on entering and leaving the room.

It is beyond the scope of this manual to detail exact feeding protocols for larval rearing. The feeding regimes should be based on the specific requirements of the various larval stages, backed by frequent and detailed examinations of the feeding activity of the larvae in each tank. Indications are given in this section of significant points to bear in mind.

Algae

An extremely high standard of hygiene must be maintained for microalgal cultures

Microalgal culture requires extreme hygiene in the laboratory phases, including thorough disinfection and filtration (to <0.5 mm) of all water and air supplies, through the use of sterilizers for all equipment and water, to the use of pure laboratory-grade fertilizing chemicals and air-conditioning to maintain temperatures between 18-24 oC.

Pure cultures of algae must be maintained using appropriate sanitary and microbiological procedures

Single-celled algae such as Chaetoceros, Thalassiosira, Tetraselmis, Isochrysis and Dunaliela are most commonly used. Pure cultures of all the algal species used should be maintained and cultured and subcultured on site, at all stages (from agar plates and tubes/bottles in the laboratory to massive on-growing outside). Appropriate sanitary and microbiological procedures should be used to ensure the quality of the culture. Contamination with protozoans that feed on algae, other species of algae, and bacteria (in particular harmful Vibrio spp.) should be avoided. Alternatively, pure starter cultures can be purchased from reputable algal culture laboratories and be on-grown in the hatchery's massive tanks using sanitary procedures. The procedure of buying one lot of pure algal culture and continuously subculturing it throughout each larval culture cycle is not recommended, as it can easily lead to contamination of the algae and eventually, of the larvae themselves.

All algal culture tanks must be washed and disinfected after each harvesting

Following disinfection of the algal culture tanks with calcium (sodium) hypochlorite solution (10 ppm active ingredient), they should be rinsed with clean, treated water and washed with a 10% muriatic acid before being left to dry.

Planktonic microalgae are usually offered to the larvae from the last naupliar stages, so that upon metamorphosis to the first feeding stage (zoea 1), the larvae will be able to begin feeding immediately. Concentrations are usually maintained at 80 000 -130 000 cells/mL throughout the zoea l and mysis stages, and then decline through the postlarval stages as the larvae become more carnivorous. During postlarval or nursery culture, benthic algae are often used, as the postlarvae will begin grazing algae from the walls of the tanks.

Artemia

Measures should be taken to ensure that Artemia do not pose a risk of disease introduction

Certification may be requested for freedom from TSV, WSSV and YHV viruses by PCR analysis for all Artemia cysts purchased.

Artemia should be decapsulated

Although Artemia cysts may not carry major viral pathogens (Sahul Hameed et al. 2002), they are certainly significant sources of bacterial, fungal and protozoan diseases. Therefore. decapsulation of the cysts is recommended to avoid contamination of the Artemia culture water and the possibility of resulting contamination of larval rearing water.

Decapsulation is carried out using 40 g of caustic soda (NaOH) and 4 litre of chlorine liquid (8-10% active ingredient) in 4 litres of seawater per 1 kg of Artemia cysts to be decapsulated. During decapsulation, the decapsulation mixture must be maintained below 20 oC using ice to prevent damage to the cysts.

As soon as the cysts begin to turn orange (indicating successful decapsulation), they are rinsed with 100 g of sodium thiosulphate added to the water containing decapsulated Artemia to stop the chlorination. The decapsulated cysts can then be washed with clean fresh water and stored in a super-saturated brine solution until needed for hatching.

Artemia should be hatched at 1-2 kg cysts/mt of seawater under constant light and vigorous aeration for 24 hours or until fully hatched

Artemia nauplii are then harvested, disinfected with a 20 ppm sodium hypochlorite solution, or better, chloramine-T at 60 ppm for 3 min, and washed with fresh water. They can then be fed live, or frozen and fed when needed, or placed into separate tanks for enrichment (for 3-12 hours), or for on-growing for feeding to postlarval stages.

After harvesting the Artemia, the hatching tanks should be thoroughly cleaned

After harvest, the tanks used to hatch Artemia must be washed with detergent and water, and then disinfected using a sponge dipped in sodium hypochlorite solution (20 ppm active ingredient), rinsed with abundant treated (filtered and sterilized) water and washed again with a 10% solution of muriatic acid.

Frozen Artemia nauplii or adults should be stored in a separate, exclusive freezer. Basic hygienic protocols (SOPs) must be implemented at all times.

Artificial feeds

Although artificial feeds generally do not pose a health risk, they must be properly used and stored

Many kinds of artificial or formulated feeds are available for use during larval rearing. These types of feeds generally do not pose the same health risks as live feeds, because they can be maintained free from contamination quite easily.

Artificial feeds include dried algae, liquid feeds, microencapsulated feeds, flakes and crumbled pellets, and mineral and vitamin supplements and enrichments. These are used in various sizes according to the stage of larval development and in various combinations, depending upon hatchery preferences, water quality and nutritional requirements. However, they are usually used primarily as supplements to live feeds.

Generally, as long as high quality feeds are selected and they are stored correctly in cool, dry conditions, used promptly once the container is opened, and not used excessively (as this can lead to water quality issues), they should not present any health-related difficulties.

5.10 Larval health management

If good numbers of high quality larvae are to be produced, tight control must been maintained on the many factors involved in managing larval health in the hatchery

There are many factors involved in managing larval health in the hatchery. A tight control must be maintained on all of these factors throughout the larval rearing cycle if good numbers of high quality larvae are to be produced. Some of the more common factors affecting larval health during the larval culture cycle (assuming that high quality nauplii have been stocked according to the methods outlined earlier in this section) are shown in Table 5.

Table 5. Some factors affecting shrimp larval health and possible control measures.

FACTOR

EFFECTS

CONTROL
MEASURES

STANDARD

Excessive Stocking Density

Stress
Cannibalism
Poor water quality

Reduce stocking density

100 to 250 nauplii/litre

Poor Water Quality

· Sea water (A)
· Tank water (B)

Mortalities
Late moulting
Deformities

Improve water quality by filtration, chlorination and/or sterilization (A)
Increase water exchange (B)

Filter < 5 mm
Activated carbon
Chlorination (10 ppm) followed by nutralization
Ozone and UV 20-100% water exchange per day

Long Stocking Period

Increased infection rates of later stocked larvae

Limit number of days in stocking hatchery

3-4 days per unit

Poor Feeding (Quality and/or Frequency)

Cannibalism
Malnutrition
Epibiont fouling
Poor water quality

Appropriate feeding programme,
Frequent checks on feed consumption and water quality

Feed every 2 to 4 hours to satiation with high quality feeds

Quality and/or Quantity of Algae

Mortality in zoeal stages
Fouling of larvae

Routine counts and quality checks

Chaetoceros or
Thalassiosira 80 000 to 130 000 cells/mL

Artemia Nauplii

Source of bacteria leading to mortality

Disinfection of Artemia nauplii

hypochlorite at 20 ppm active ingredient

Stocking density

The density at which nauplii are stocked should not be excessive

Overstocking can result in stress and in later stages, and may lead to cannibalism and reductions in water quality, especially when survival rates are high. In general, stocking rates for nauplii should be in the range of 100-250 nauplii/litre (100 000 - 250 000 per mt) of water. Lower stocking densities are typically used where larvae are grown to harvest size in a single tank, while higher densities can be used where a two-tank system is used. In the latter system, the larvae are typically cultured in a conical or "V" or "U"-bottomed tank at high density until PL4-5 and then transferred to flat-bottomed tanks for the later, benthic stages at reduced densities of up to 100 PL/litre.

Poor survival may reduce the density of larvae in a larval rearing tank to a level below where it is cost-effective to feed (because larval rearing tanks are generally fed according to volume of water rather than number of larvae).

Water quality

Good water quality must be maintained

Water quality has a major impact on the health and performance of larval batches. Poor water quality can lead to poor growth, low survival, late moulting/staging, increased epibiont fouling and deformities. Water for larval rearing should be filtered to around 5 µm and disinfected with chlorine, ozone and/or UV. The temperature should be maintained between 28 and 32 °C and salinity above 30 ppt, at least until postlarval stages are reached. Dissolved oxygen levels should be maintained as close to saturation (6.2 ppm at 30 oC) as possible, but at least above 5 ppm. A pH of around 8 should be maintained. Overfeeding is one of the major causes of water quality deterioration and should be avoided.

Water quality is also maintained through aeration that is sufficient to prevent uneaten food and faeces from settling on the tank bottom (normally necessitating tanks with angled bottoms) and regular siphoning of the tank to prevent a build up of anaerobic sludge on the bottom.

Water exchange should be carefully handled

Generally, no water is exchanged until the mysis stage is reached, although water levels are generally increased throughout the zoeal stages, because nauplii are usually stocked in tanks that are only half full. After the mysis stage, typically 20-100% of the water is exchanged daily, depending upon the stocking density and water quality parameters. Care should be taken to make sure that the water used to top-up the tanks and all exchange water is at a similar temperature, salinity and pH to the water in the tank, and free from chlorine to avoid unduly stressing the larvae.

Hatcheries should also consider the use of probiotics and bacterial enzymes

In an effort to maintain water quality, prevent bacterial blooms and reduce or eliminate the requirement for antibiotics during larval culture, hatcheries are turning increasingly towards the use of probiotic powders or solutions of beneficial bacteria and bacterial enzymes. As for all these products and supplements, care must be taken in selecting those which do not pose their own health risks and which are efficacious and cost-effective.

Stocking period

Each separate unit of larval rearing tanks within a hatchery or, preferably, the whole hatchery should be stocked with nauplii in as short a time period as possible

Each separate unit of larval rearing tanks within a hatchery or, preferably, the whole hatchery should be stocked with nauplii in as short a time period as possible, usually limited to three to four days. Prolonging this stocking period often results in increased incidence of disease for the later-stocked larvae, presumably through bacterial contamination from the older to the younger tanks.

This phenomenon is often associated with the so-called "zoea-2 syndrome", where late zoea 1 and early zoea 2 stage larvae refuse to eat and suffer high mortality with associated bacterial problems. This problem may be controlled through restricting the time of stocking to less than four days, using probiotics and maintaining good cleanliness in all areas of the hatchery at all times.

Nutrition and feeding

The quantity, quality and management of feed should be closed monitored

The quantity, quality and management of feed can have an important impact on larval health and survival. Failure to provide sufficient feed of the right quality can lead to stress, poor growth, mortality, increases in cannibalistic behaviour, deformity and increased levels of epibiont fouling. This is especially true when a large proportion of the feed used is formulated diets. When using formulated diets as a supplement to live feeds, it important is to feed small amounts of high quality, appropriately sized, nonpolluting diets frequently. As a guide, particle sizes should be 10-50 mm for zoea, 100-200 mm for mysis, and 200-300 mm for early postlarval stages. A feeding frequency of every two to four hours is generally regarded as sufficient.

For the majority of the larval feed requirements, reliance should still be placed on high quality live feeds, including algae and Artemia

However, insufficient or poor quality algae can also have severe consequences for larval health. Heavy mortality in the zoea stage, for example, has been linked to algal quality, and insufficient algae during this stage (<80 000 - 130 000 cells/mL) can result in the larvae having insufficient reserves to complete the stressful moult to mysis. Algal concentrations and quality should be regularly monitored to ensure that they are sufficient for the stage being fed.

5.11 General assessment of larval condition

Assessment of larval condition should be one of the main activities carried out in the hatchery

Assessment of larval condition is usually done in the morning, and decisions on water exchange, feeding and other management activities made so that action can be taken in the afternoon. The larvae in each tank should be inspected two to four times each day. Initially, a visual inspection of the larvae, the condition of the water in the rearing tank and the feed is made. A sample of larvae can be taken with a beaker and inspected with the naked eye. Observations are made on the larval stage, health, activity, behaviour and abundance of feed and faeces in the water. Records may also be taken of water quality parameters, and the amount of food in the tank.

The same, or a separate sample of larvae, should also be taken to the laboratory for a more detailed microscopic examination. This will provide information on the stage, condition, feeding and digestion and presence of any disease or physical deformity.

Samples may also be sent once or twice during the cycle for analysis in a PCR laboratory for screening for viral diseases.

The type of observations that are made can be categorized into three levels, based on the health assessment levels described in Table 2.

Level 1 Observations

Level 1 observations are based on simple visual features of the larvae and water condition that can be easily seen with the naked eye in a glass beaker of animals taken from the tank. Special attention is paid to the behaviour or activity of the larvae, their swimming behaviour (according to the larval stage), water quality, presence of feed and faeces and later on, size disparity and homogeneity. These observations and the scoring system used are summarized in Table 6.

Table 6. Summary of Level 1 assessments of larval health.

CRITERIA

SCORE

STAGE

OBSERVATION

Swimming activity

10

All stages

Daily (2-4x) observations

Active (> 95%)

Intermediate (70-95%)

5

Weak (on bottom) (< 70%)

0

Phototaxism

10

Zoea

Daily (2-4x) observations

Positive (>95%)

Intermediate (70-95%)

5

Negative (< 70%)

0

Faecal string (cord)

10

Zoea

Daily (2-4x) observations

Present (90-100%)

Intermediate (70-90%)

5

Absent (<70%)

0

Luminescence

 

Mysis

Night observation of the tank

Absent

10

Present (<10%)

5

Abundant (>10%)

0

Homogenous stage

10

All Stages

Daily (2-4x) observation

High (80-100%)

Intermediate (70-80%)

5

Low (< 70%)

0

Intestinal contents

10

Mysis

Daily (2-4x) observation

Full (100%)

Half full (50%)

5

Empty (<20%)

0

Swimming activity

The swimming activity of the larvae changes dramatically but characteristically through the larval cycle. Zoea l stages will swim rapidly and consistently forwards, usually in circles, filter feeding on phytoplankton. Mysis, by comparison, swim backwards with intermittent flicks of their tails, maintaining themselves in the water column and feeding visually on phyto- and zooplankton. PL, again turn to swimming rapidly and consistently forward, initially planktonically, but at least from PL4-5 onwards, benthically, searching for food, unless maintained in the water column by strong aeration. Within these distinct modes of swimming, if >95% of the larvae are observed to be swimming actively, they are given a score of 10; if 70-95% are active, they are given a score of 5; and if <70% are active, they are given a score of 0.

Phototaxis

Zoea stage larvae should retain a strong positive phototaxis and move towards light. To test this, a sample of larvae is placed in a translucent container next to a light source and the displacement of the animals is observed. If 95% or more of the larvae move strongly towards the light, the larvae are good and given a 10; if 70-95% respond, they are acceptable and given a 5; and if less than 70% move towards the light, they are considered weak and given a score of 0.

Faecal string (cord)

During the zoea l stages, when the zoea are feeding almost exclusively on algae, long faecal strings can be seen projecting from the anus and loose in the water column. When 90-100% of the larvae have these long, continuous strings all along the digestive tube, through their bodies and continuing outside, they are considered well fed and given a score of 10. When 70-90% have these strings, or they are short or discontinuous, they are given a score of 5; and when <70% of the larvae have these strings, the larvae are not eating and they are given a score of 0.

Luminescence

This factor is observed directly in the larval rearing tank in absolute darkness. Larval luminescence is generally due to the presence of luminescent bacteria such as Vibrio harveyi. If no luminescence is observed, a score of 10 is given; if the observed luminescence appears low (up to 10% of the population), the score is 5; and if above 10% of the population are luminescent, the score is zero.

Stage homogeneity

This indicates the uniformity of larval stages in a tank. If 80% or more of the population is in the same stage, a score of 10 is given; if between 70 and 80% are at the same stage, the score is 5; and if less than 70% are in the same stage, the score is zero.

It should be noted that when larval shrimp moult, it is normal to see a decrease in the stage homogeneity, so the time at which the stage homogeneity is determined has to be taken into consideration. This is also true for postlarvae when they are moulting.

Intestinal contents

The intestinal contents can be observed in older larval stages. The intestine is visible as a dark line from the hepatopancreas in the larva's head region that is easily observed in larvae held in a clear container, such as a glass beaker. This is useful as a guide to larval feeding and feed availability. If most of the larvae observed are full, a score of 10 is given; if half of the larvae have food in the intestine, a score of 5 is given; and if <20% of the larvae have food in the intestine, the score is zero.

Level 2 Observations

Level 2 observations are based on microscopic examination and squash mounts, if necessary, of a randomly taken sample of at least 20 larvae per tank (more for larger tanks). Special attention is paid to the state of the hepatopancreas and intestinal contents, necrosis and deformity of limbs, fouling organisms and the presence of baculovirus in the faeces or hepatopancreas of older larvae. These observations and the scoring system used are summarized in Table 7.

Table 7. Summary of Level 2 assessments of larval health.

CRITERIA

SCORE

STAGE

OBSERVATION

Hepatopancreas (lipid vacuoles)

10

All stages

Daily (2-4x) observations

High (>90%)

Moderate (70-90%)

5

Low (< 70%)

0

Intestinal content

10

All stages

Daily (2-4x) observations

Full (>95%)

Moderate (70-95%)

5

Empty (< 70%)

0

Necrosis

10

All stages

Daily (2-4x) observations

Absent (0%)

Moderate (<15%)

5

Severe (>15%)

0

Deformities

10

All stages

Daily (2-4x) observations

Absent (0%)

Moderate (<10%)

5

Severe (>10%)

0

Epibionts

10

All stages

Daily (2-4x) observations

Absent (0%)

Moderate (<15%)

5

Severe (>15%)

0

"Bolitas

10

All stages

Daily (2-4x) observations

None

1 to 3

5

>3

0

Baculovirus

10

Mysis

Daily (2-4x) observations

Absent (0%)

Moderate (<10%)

5

Severe (>10%)

0

· Sloughed cells of hepatopancreas and intestine expressed as number of "bolitas" in the digestive tract)

Condition of the hepatopancreas and gut contents

The condition of the hepatopancreas gives an indication of larval feeding and digestion. It is observed using a wet mount of a sample of larvae on a microscope slide at a magnification of 40X. In healthy larvae showing active feeding and digestion, the hepatopancreas and midgut will be full of small, easily observed bubbles (digestive or "lipid" vacuoles) and strong peristalsis will be seen in the intestine. If 90% or more of the animals sampled show abundant lipid vacuoles and/or a full gut, a score of 10 is given; if the sample shows 70 to 90% of individuals with lipid vacuoles and/or a moderately full gut, a score of 5 is given; and if it is less than 70% and/or the intestine is empty, the score is zero.

Necrosis

Necrosis of the larval body and limbs, which is an indication of cannibalism or possible bacterial infection, can be observed by light microscope under low power. If necrosis is absent, a score of 10 is given; where <15% of the animals show some necrosis, a score of 5 is given; and where >15% show necrosis, indicating a severe infection is present, a score of 0 is given.

Deformities

Deformities may indicate poor quality nauplii, if in the early stages, and bacterial infections or mishandling and stress later on. Typically, the fine setae on the limbs of the larvae and/or their rostrums may appear bent, broken or missing; the tail may appear bent; or the gut may terminate before the anus. Typically, no remedies exist for these problems (unless due to rough handling), and such deformed larvae will die. In severe cases, it may be preferable to discard the whole tank as soon as possible to prevent infection of other tanks. Where deformities are absent, a score of 10 is given; ff <10% have deformities, a score of 5 is given; and if >10% present deformities, a score of 0 is given.

Epibiont fouling

The larvae may become host to a range of fouling organisms ranging from bacteria and fungi through to protozoans of many species. These will typically attach to the exoskeleton on the head and body, and particularly around the gills of the larvae. Where the infections are slight, the next moult may remove the fouling without further problems, but in severe cases, the fouling will persist or reoccur in the next stage, indicating poor water quality and necessitating action. Where fouling is absent, a score of 10 is given; if <15% have temporary or permanent fouling, a score of 5 is given; and if >15% are fouled continuously, a score of 0 is given.

Baculovirus

Baculoviruses can usually be detected in whole or squashed (stained with malachite green for Monodon baculovirus) preparations of hepatopancreas or faecal strands from larger-sized larvae, using a high powered light microscope to spot the characteristic viral occlusion bodies (which, in the case of MBV, are dark coloured and tetrahedral). The expression of bacculoviruses is often mediated by stress, and if seen, reductions in levels of stress can often reduce prevalence and the associated problems of growth depression. Where baculoviruses are absent, a score of 10 is given; if <10% have baculovirus, a score of 5 is given; and if >10% are infected, a score of 0 is given.

"Bolitas"

"Bolitas" is the Spanish name given to a syndrome involving the detachment of epithelial cells from the intestine and hepatopancreas, which appear as small spheres within the digestive tract. It is believed to be caused by bacteria and can be fatal. Some success in preventing "bolitas" condition has been achieved by rapid stocking of the hatchery (within three to four days), use of probiotics, and good health and feeding management practices.

The value of Level 1 and 2 scoring

When all of these level 1 and 2 observations are made and recorded for each tank of larvae at each stage and the appropriate scores given in each case, an overall picture of larval health can be derived, with higher numbers relating to healthier larvae and vice versa. With experience, it becomes easy to judge the overall health of each tank of larvae and to recommend courses of action to combat the problems encountered, depending on the scores obtained.

Level 3 Observations

Level 3 observations utilizing molecular techniques and immunodiagnostics are not normally required until the postlarvae are ready to be transferred to on-growing facilities. PCR and/or dot-blot techniques are commonly used to test for major viral pathogens. However, PCR is recommended as it is relatively more sensitive than dot-blot.

5.12 Selection of postlarvae for stocking

Good hatchery management should be practised to ensure high postlarval quality

Many factors affect the quality of PL. Feed quality and quantity, moulting, water quality (temperature, salinity, ammonia, suspended solids, faeces), use of antibiotics, diseases and bad management practices can all have an impact on the quality of postlarvae produced by a hatchery. These factors can be regulated through the use of good hatchery management practices, and this will have a major impact on the quality of the postlarvae produced.

As mentioned previously, the larval production plan should be aimed at producing the highest quality animals possible, because subsequent on-growing performance is directly related to postlarval quality. It is thus in the on-growing phase where postlarval quality is of most importance.

There are many indicators of health and quality that can be used to determine postlarval selection. These indicators fall under the previously mentioned (see Table 2) three categories or levels and are detailed in Tables 8, 9 and 10.

Moulting

The postlarvae should be checked that they are moulting easily, but that they are not moulting when intended for harvest and transport, because this will reduce survival rate during this critical time. Also check that there are no moults stuck to the heads of the PL, resulting in bent antennae and impediment of feeding, and ultimately starvation and death. This is usually caused by inadequate feeding, poor food quality and/or bacterial disease usually related to poor water quality. Thus, increased water exchange and revision of feeding protocols can be used to combat this problem.

Postlarval quality assessment using Level 1 procedures

Swimming activity

The vigour of swimming activity should be assessed as a general guideline of postlarval health using the techniques described for larvae in Section 7.12.1. The larvae can also be put into a bowl and the water swirled with a finger. Healthy postlarvae should orient themselves facing the current and not fall into a pile at the bottom of the bowl, being unable to resist the current. They should also respond to tapping the side of the bowl by jumping.

Table 8. Summary of postlarval quality assessment using Level 1 procedures.

Criteria

Observations

Qualitative Assessment

Score

Moulting

Moults in the water

< 5%

10

Moults not sticking to head of PL

5-10%

5

>10%

0

Swimming Activity

Activity level of postlarval swimming behaviour

Active

10

Intermediate

5

Low

0

Direct Observation of Luminescence

Night-time observation of the tank

<5%

10

5-10%

5

>10%

0

Survival Rate and Clinical History of Tank

Estimation of survival rate in each tank

>70%

10

40-70%

5

<40%

0

Luminescence

The prevalence of luminescence as an indication of potentially pathogenic Vibrio spp. infections should be determined using the techniques described for larvae in Section 7.12.1 or using Level 2 techniques described below. Presence of luminescence requires immediate treatment (probiotic use can sometimes be successful) in order to prevent more severe infections.

Survival rate

The survival rate of postlarvae in each tank should be estimated as an indication of the general state of health, clinical history and lack of problems during the cycle.

Each of these Level 1 postlarval quality assessments are carried out visually on randomly taken samples of >20 animals (where appropriate) and the scoring system detailed in Table 8 applied.

Postlarval quality assessment using Level 2 procedures.

Level 2 assessments are carried out on a randomly selected sample of >20 postlarvae per tank which are examined using low- and high-power light microscopy. The scoring system detailed in Table 9 is then used to score the quality of each batch of postlarvae produced.

Muscle opaqueness

An examination should be made of the body of the PL, concentrating on the bend of the tail around the 4th-5th abdominal segments. The normally transparent muscles turn opaque due to various reasons, including bacterial infection. This problem can be quite serious and potentially fatal if left untreated.

Deformities

Postlarvae should be examined for various deformities such as bent rostrum, enlarged head due to moulting problems, or missing or damaged limbs due to bacterial infections, to estimate general health. Some deformities are fatal.

Size variation

To determine the size variation, measure individually the length of at least 50 postlarve and calculate the mean length and the standard deviation. The coefficient of variation (CV) is obtained by dividing the standard deviation by the mean. If the CV is equal to or less than 15%, the size variation is considered low (score 10); if the CV is between 15% and 25%, the size variation is moderate (score 5); and if it is greater than 25% the size variation is high (score 0).

When postlarvae moult, it is normal that the CV will increase, so the time at which the CV is determined has to be taken into consideration. If the CV is found to be high, the test should be repeated after a day to give time for the whole population to complete the moult.

Gut content

Examinations of the intestinal tract for its contents and appearance (not just the colour) should be made to assess the PL's feeding level according to the criteria shown in Table 9. The presence of empty guts may be the first sign of disease, or may just be due to inadequate feeding. In either case, it should be investigated immediately. It is important to examine postlarvae immediately following sampling.

Colour of the hepatopancreas

The hepatopancreas should not be transparent and should have a good coloration. Typically, it should be dark yellow ferrous or ochre in colour, however, the colour of the hepatopancreas can be greatly influenced by the quality and colour of the diets fed and tanks used. A darker coloured hepatopancreas generally indicates better health. Care must be taken when using some flake feeds, as these may contain dyes that stain the hepatopancreas almost black, without necessarily contributing to the animals' health.

Condition of the hepatopancreas

The hepatopancreas of the postlarvae should be examined for its general condition, which is primarily indicated by the number of lipid vacuoles and its overall size. The presence of a relatively large hepatopancreas with a large number of lipid vacuoles is considered a sign of good health. Postlarvae with a small hepatopancreas containing few lipid vacuoles is a sign of under feeding, and improved feeding prior to harvest may be required in order to enhance their quality.

Epibiont fouling

Postlarvae should be examined for any epibiont or organic matter fouling on the exoskeleton or gills (usually consisting of protozoans such as Zoothamnium, Vorticella, Epistylis or Acineta, filamentous bacteria or dirt and organic matter). Fouling can normally be moulted off or treated with formalin at up to 20-30 ppm for one hour (with full aeration).

Melanization

Postlarvae should be examined for melanization, which often occurs where limbs have been cannibalized or where bacterial infections have occurred. Excessive melanization is a cause for concern and requires treatment through water quality and feeding regime enhancement, and sometimes reductions in stocking density, to prevent cannibalism and reduce bacterial loads.

Gill development

The state of gill development should be examined, as it gives a good idea of when the postlarvae are able to tolerate salinity changes, which often occur when the shrimp are transferred to the on-growing facilities. When the gill lamellae have become branched like Christmas trees, approximately around PL9-10, they are generally able to tolerate fairly rapid changes in salinity (up to 1 ppt/hr down to 5 ppt, or 0.1 ppt/hr below 5 ppt) and can easily be acclimated to on-growing conditions. Where the gill lamellae remain unbranched, the shrimp should not be subjected to major or rapid salinity changes and should not be considered ready for transfer from the postlarval tanks.

Intestinal peristalsis

A high-power microscopic examination of the intestinal tract of the postlarvae should be conducted in order to ascertain the peristaltic activity of the intestinal muscles. Strong gut peristalsis, in combination with a full gut, is an indication of good health and high nutritional status.

Baculovirus

Refer to page 43.

Muscle to gut ratio

A microscopic examination of the relative thickness of the ventral abdominal muscle and the gut in the 6th abdominal segment of the tail of the postlarvae should be conducted to determine the muscle to gut ratio. This gives a useful indication of the nutritional status of the animal. High muscle to gut ratios are preferable (see Table 9).

Table 9. Summary of postlarval quality assessment using Level 2 procedures.

Criteria

Observations

Qualitative Assessment

Score

Muscle Opaqueness

Opaque muscle in tail of PL

<5%

10

5-10%

5

>10%

0

Deformities

Deformities in limbs and head

<5%

10

5-10%

5

>10%

0

Size variation (CV)

Calculation of CV of postlarval size

<15%

10

15-25%

5

>25%

0

Gut content

Degree of fullness of digestive tract

Full

10

Moderate

5

Empty

0

Colour of the Hepatopancreas

Relative coloration of hepatopancreas

Dark

10

Pale

5

Transparent

0

Condition of the Hepatopancreas

Relative quantity of lipid vacuoles

Abundant

10

Moderate

5

Epibiont Fouling

Degree of fouling by epibionts

<5%

10

5-10%

5

>10%

0

Melanization

Melanization of body or limbs

<5%

10

5-10%

5

>10%

0

None

0

Gill Development

Degree of branching of gill lamellae

Complete

10

Intermediate

5

Slight

0

Intestinal Peristalsis

Movement of gut muscle

High

10

Low

5

Baculovirus

Daily (2-4x) observation of Mysis

Absent (0%)

10

Moderate (<10%)

5

Severe (>10%)

0

Muscle to Gut Ratio

Comparison of ratio between muscle and gut thickness

>3:1

10

1-3:1

5

<1:1

0

"Bolitas" (sloughed cells of hepatopanceas and intestine)

Number of bolitas in digestive tract

None

10

1 to 3

5

>3

0

Stress Test

If < 75%, re-testing is recommended

>75%

10

"Bolitas" (sloughed cells of hepatopancreas and intestine)

Refer to page 43.

Stress test

At harvest, or once the postlarvae reach PL10, a stress test can be carried out. There are several stress tests, and the most common method is to place a randomly selected sample of about 300 animals in a beaker with water at 0 ppt salinity, leave them for 30 minutes and then return them to 35 ppt (or ambient) water for another 30 minutes. Following this, the survivors are counted and the percentage of resistant individuals calculated. Stress tests should not be carried out when the postlarvae are moulting, as they are unduly stressed at this time. Some hatcheries have used 100 ppm of formalin for 30 minutes as the stress test, with similar success.

Postlarval quality assessment using Level 3 procedures.

Level 3 assessments should be carried out on a statistically determined number of postlarvae (usually 150 for a population > 10,000) from each tank (in order to provide a 95% confidence level at 2% prevalence in the result) using PCR techniques for the detection of important viral pathogens. This testing must be done according to standard protocols by a competent health laboratory, following all the rules for sampling, preservation and transport of the samples. For a detailed discussion of sampling for disease detection, see OIE (2003).

The only acceptable result for any of these viral pathogens is a negative result (which scores 10 points - see Table 10), where both negative and positive controls have simultaneously given their corresponding expected results. All batches testing positive should be destroyed.

Table 10. Summary of postlarval quality assessment using Level 3 procedures.

Analysis

Observations

Qualitative Determination

Score

PCR

WSSV/YHV

Negative

10

IHHNV

Negative

10

TSV

Negative

10

5.13 Risk assessment for stocking

A table summarising the three levels of postlarval quality and the points system should be used to determine the fate of the PL

As with larval quality assessment, a summary table should be made of these three levels of postlarval quality and the points system employed (using some or all of the above indicators, depending on circumstances). This table then is used to determine which tanks of postlarvae are selected for on-growing, which may require treatment before selection, and which will be rejected. As before, experience will guide the manager in his selection of indicators to use and of a cut-off point for points scored, below which the postlarvae batch will be treated or rejected.

The risk of stocking a given batch of postlarvae must be carefully assessed.

The decision to stock or not to stock a batch of postlarvae is ultimately an assessment of risk. No fixed guidelines or standards can be provided, as this generally comes from experience, but the following guide can be used to reduce the risk of experiencing mortalities or poor growth in pond culture of Penaeus vannamei. In this risk analysis, the order of importance of assessment is Level 3 > Level 2 > Level 1.

The following criteria can be used:

· Postlarvae must pass Level 3 assessment.

- Postlarvae must be PCR or dot-blot negative for YHV, IHHNV, WSSV and TSV.

· Provided that postlarvae passed Level 3 assessment, the following guide can be used for Level 2:

- A score greater than 100 represents a low risk of severe disease problems, therefore recommended

- A score of 65-100 represents a moderate risk of severe disease problems.

- A score less than 65 represents a high risk of severe disease problems, therefore not recommended.

· Provided that animals pass Level 2 assessment, the following guide can be used for Level 1:

- A score greater than 30 represents a low risk of severe disease problems, therefore acceptable.

- A score of 20-30 represents moderate risk of severe disease problems.

- A score less than 20 represents a high risk of severe disease problems, therefore not recommended.

5.14 Shipping and transfer of postlarvae

Postlarvae must be carefully and appropriately packed for shipping to grow-out facilities

Postlarvae can be transported in large tanks or in boxes with plastic bags and at densities that may vary from 500 to 1200 PL/litre, depending upon duration and method. Two plastic bags (one inside the other) of 25-30 litre capacity are commonly used, filling with 10-15 litres of filtered water, adding the desired quantity of postlarvae and then filling with pure oxygen, bubbled into the water. As food source, live Artemia nauplii are typically added at about 15-20 nauplii per postlarvae for every four hours of transport. A few granules of washed, new, activated carbon may also be added to each bag to assist in maintaining low ammonia levels during long transportation times. The bags are then sealed with elastic bands and placed into sealed cardboard cartons for short distances and/or polystyrene for added insulation over long distances.

The temperature used and the stocking densities employed during transportation will vary according to the travel time and distance to be shipped. Typically, no temperature reduction is needed if the hatchery is close to the farm site, but temperatures will be reduced to 25-28 oC for transportation times of one to three hours, to 23-25oC over transportation times of 3-12 hours and 18-23 oC for over 12 hours. Such temperature reduction is used to lower the metabolic rate of the larvae so that they will use less oxygen, excrete less waste and remain calm during transportation. The salinity of the water should be that to which the postlarvae have been acclimated, which should be similar to that expected in the grow-out facility.

Strict biosecurity measures should be followed

All shipping containers and equipment (nets, air stones, air lines etc.) should be disinfected before and after use (see appropriate sections in this document for procedures). If plastic bags are used, they should be incinerated after use; they should not be re-used for shipping postlarvae or broodstock shrimp.

The vehicles that deliver the postlarvae are a potential source of contamination, as they may visit several farms and hatcheries in the course of making deliveries. If possible, postlarval packing should take place at a point isolated from the production facilities, and the transport trucks (at least the wheels and tires) should be disinfected before entry to the hatchery.

5.15 Documentation and record keeping

A comprehensive and up-to-date system of documentation and record keeping should be established

Good documentation and record keeping are fundamental to any system of good management practice. It is important that a detailed set of written standard operating procedures (SOPs) is developed and a system of regular employee training programmes be established. It is also important that the SOPs are reviewed regularly and kept up to date.


[8] In the past, muriatic acid was referred to 3:1 HCl and HNO3, but currently it is referred to as 34-37% HCl.

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