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The larval rearing of gilthead seabream and seabass is a typical intensive rearing technology ideally involving complete control over the environmental parameters and fish population. Two different protocols are acknowledged. The first takes place in a lighted environment using rotifers as first feeding and adding microalgae in the gilthead seabream initial culture. The second protocol, frequently referred to as the “French technique”, is applied for seabass larval rearing and is characterised by a dark environment during the first days after hatching, and by the use of small newly hatched brine shrimp nauplii as first food.

Because it is more widely adopted and it is the only applicable technology for both species, the following description refers to the first protocol. It discusses in detail the working procedures concerning the management of the abiotic (water system, water chemistry and hygiene) as well as biotic factors (husbandry of fish larvae, live feeds and rearing environment). In this manual the term larval rearing refers to both the larval phase, i.e. when the yolk is the only energy source, and the post-larval phase, i.e. when feed is provided as external energy source.

Layout of the larval rearing system

The first four to six weeks of the life of young seabass and gilthead seabream are spent in a specific larval rearing unit of the hatchery. The most common equipment consists in a number of round fibreglass tanks of an individual capacity of 6-10 m3. Seawater is either recirculated through a biofilter in the case of a semi-open recirculation system, or is just pumped to the tank and discharged after use in the open systems.

Fig.46.01-2 Old and modern tanks for larval rearing units (photo STM Aquatrade)

As seabass and gilthead seabream reproduction takes place during the winter season, heating the water in the larval rearing tanks accelerates the growth rate. In some cases, where wells with higher temperature than seawater can be used, the heating requirements are reduced. A light hanged over each tank provides the necessary illumination to allow visual predation. Since live feed is mostly distributed by hand, automatic feeders are employed only during the last culture days before the transfer to the weaning section. Additional equipment include: water sterilizers, small insulated tanks for the short term stocking of live feed, tank cleaning devices (siphons, brushes, disinfection tanks for equipment, etc.) and manual or automatic devices to control main environmental parameters.

Preparing the larval rearing system

In advance of the onset of the reproduction season, the entire larval rearing system (piping, tanks, filters, sterilizers, air system, equipment, etc.) has to be properly set. Everything should be thoroughly cleaned with detergent and disinfected with a hypochlorite solution (500 ppm of active chlorine). A practical way to do it for tanks and piping is to fill them with the hypochlorite solution and keep the solution in circulation or standing (depending on the circuit layout) for a few days. It is also mandatory to perform these cleaning operations at the end of the rearing season, to avoid dirt drying and sticking.

During the production season, after a batch of fry has been transferred to the weaning sector, tanks and their outlet pipes should be similarly disinfected with hypochlorite. The outlet pipe system should be dismantled and kept in a hypochlorite solution overnight (further details are given in annex 16). If the system includes a biofilter, it has to start working at least 30 days before any larval rearing is started in order to allow a proper colonization of the filter medium by nitrifying bacteria. The disinfection of the biofilter components has to take place well in advance.

All consumables must be ordered well in advance and be at hand before production starts. A typical example of poor management is given by delays and mistakes in ordering items, which can quickly jeopardize the work of the hatchery. Some items such as special batches of brine shrimp cysts, food integrators and drugs may also run out of stock during the rearing season. The search for new suppliers is often a time-consuming and expensive activity. This recommendation also applies to having available an adequate stock of spare parts, in particular for pumps, sterilizers and piping, where a quick servicing can save the production of an entire season.

Well before the first stocking, the entire rearing system has to be assembled and tested running at full capacity. Any trouble with equipment must be solved either by fixing or by replacing it.

Environmental parameters for larval rearing

Since the larval and postlarval rearing conditions of gilthead seabream and seabass differ to some extent, the table below compares the two species for different parameters.

Fig.46.03 Hypoclorite disinfection tank (photo STM Aquatrade)

A comprehensive summary of the evolution of the main environmental parameters in the rearing system is also given in Annex 13. As a general rule, since fish mortality is highest during the first month, the greatest attention is required at that time.

Table 3.9 - Environmental parameters in the larval and post-larval rearing of seabass and gilthead seabream.

Gilthead seabream


Water temperature

Same as spawning temperature at incubation up to yolk sac resorption (-2 to +6 days). Slowly increasing (0.5° C/day) to reach 18-20° C, the choice depending on management considerations and period of the year. Particular care has to be given to maintaining water temperature during the first 25 days, when water renewal is nil or very limited. Fluctuation should never exceed 0.5° C in 24 h.

Water temperature

Same as spawning temperature at incubation up to yolk sac resorption (-2 to+6 days). Slowly increasing (0.5° C/day) to reach 18° C within the complete swim- bladder inflation. Then increased to 20° C by the 15th day. Fluctuation should never exceed 0.5° C in 24 h.

Air temperature

Fluctuation within 1° C. That should preserve water temperature especially at night. Special attention should be paid when water renewal is nil or very limited.

Air temperature

Fluctuation within 1° C That should preserve water temperature especially at night. Special attention should be paid when water renewal is reduced.


Usually the same salinity at spawning (35-38 ppt = full seawater). A lower salinity down to 25-30 ppt during first feeding may enhance survival rate, but at a cost:

- at least two separate hydraulic circuits are needed
- live food settling is increased
- a change in salinity can increase stress.




16 hours light, 8 hours dark when temperature remains below 21° C. Above 21°C increase to 20h L/4h D.


16 hours light, 8 hours dark when temperature remains below 21° C. Above 21° C increase to 20h L/4h D.

Light intensity

1,000 to 3,000 lux at water surface till age 25 days, thereafter 500 to 1,000 lux until metamorphosis. During on/off operations use a 10-min twilight effect by means of a dimmer switch driven by a timer. Halogen lamps are advisable for light quality and cost effectiveness.

Light intensity

500 lux at water surface. The Alternative “French method” foresees complete darkness during a short initial period of 5-7 days.

Bottom aeration

A very slow fine bubbling of 0.1 l/min during first feeding. Gently, but constantly increased from 15th day on up to 0.6 l/min, related to larval activity, surface dirt and distribution of live feeds. If eggs are stocked directly into the larval tanks, a rather strong air flow keeps eggs suspended, and is reduced at hatching to be completely stopped at the end of hatching to allow debris settling and removal.

Bottom aeration

Same pattern, but twice as much flow due to the sturdy larval stage of bass.

Water renewal

None during first feeding light periods, moderate at dark (0.5 to 1 total tank renewal). Increasing steadily up to metamorphosis to 10 total tank renewals at night. Regular water quality monitoring should confirm or adjust renewal to the actual need. If eggs are stocked directly into the larval tanks, 1-2 renewals/day during incubation, and 3 to 6 during hatching (check water temperature).

Water renewal


Dissolved oxygen (DO)

DO saturation should remain between a minimum of80% and a maximum of 100%. Adjust aeration, water renewal, phytoplankton daily ration and bottom cleaning accordingly.

Dissolved oxygen


Total Ammonia Nitrogen (TAN)

It should not represent a major problem in the larval unit due to its scarce total biomass. In any case keep it below 0.5 ppm.

Total Ammonia Nitrogen (TAN)


Screen mesh

Use a 500 mm filter mesh when you want to discard uneaten rotifers and brine shrimp (as their nutritional value is lost if they remain there overnight). To keep them in the tank, if a water renewal is needed during feeding, replace with a 100 mm filter. If only artemia nauplii are fed, mesh size can be increased to 250 mm.

Change every time it is clogged or near to clogging, in particular just before dark. If eggs are stocked directly into the larval tanks, use a 400 µm filter during incubation and hatching.

Screen mesh



At a water temperature of 18°C, a photoperiod of 16h light - 8h darkness is provided to extend the predatory feeding and to increase larval growth. Higher temperatures require a longer light time to match increased larval activity and growth rate.

To automatically set light intensity and photoperiod, a dimmer switch controlled by a timer should be installed. The twilight effect should last 10 to 15 minutes to prevent stress induced by abrupt changes of light intensity at on/off.


For gilthead seabream in their early post-larval development stage, light intensity is critical to start a proper predatory activity. It should be at least 800 lux in the less lit areas of the tank surface, with an optimal range of 1 000 to 3 000 lux. In case of seabass, the light intensity at first feeding may be as low as 100 lux, with an optimal value of 500 lux.

In both cases direct solar light has to be avoided because it would be too strong and would create an interference with the artificial environment. Instead, incandescence bulb lamps or daylight halogen lamps are hung over the rearing tanks at a suitable distance from the water surface to provide the required light intensity.


Tank aeration maintains fish postlarvae and live feed in suspension and allows a proper mixing of the tank water. Aeration should be adjusted to avoid a stressful turbulence to which post-larvae are most sensitive in particular at two critical stages: during the first feeding, and during the formation of the swimbladder. Excessive water movement may prevent predatory activity of fish postlarvae, and may make more difficult the gulping of an air bubble at the water surface that is necessary to activate the inflation of their swimbladder.

According to tank shape, water depth and inlet/outlet position, the aeration should create a slow up-welling current for proper vertical mixing of the entire water mass. In round tanks with conical bottom, this is obtained with a central fine airstone placed at 20 cm from the cone tip. In this way dirt and debris can settle and accumulate to facilitate cleaning.

Fig.47.01-2 Open and submerged water inlet for the larval rearing tank (photo STM Aquatrade)

A common device to inject air from a low-pressure aeration pipeline is the transparent PVC 6 mm-hose to which a standard plastic needle-valve is fitted. Diffusers range from the standard carborundum porous stone to textile porous hose..

Water flow

The water inlet has to be placed carefully in order to avoid dangerous strong currents (more than 10cm sec-1) that can trigger spinal deformities and impede the onset of predatory behaviour. As a general rule, the water inlet should be at the tank periphery and positioned in a way that would avoid tangential currents. A current velocity at the periphery of not more than 2cm sec-1 is suggested.

Dissolved oxygen

Oxygen is provided to fish through the water renewal and, to a limited extent at least in the larval rearing, through aeration. Since renewal rates during the first stages of larval culture are rather low, it is important to monitor dissolved oxygen content and to adjust it whenever it falls below 80% saturation.

The addition of microalgae at high density helps in stabilising DO concentrations during the period in which lights are on and water renewal is absent or reduced. As a rule, even if it becomes truly significant during weaning, the DO levels should be checked after feeding, when DO demand peaks, and after a prolonged period without water renewal. In presence of excessive amounts of microalgae coupled with absence of water renewal at night, a check of the DO level during and towards the end of the dark period is advisable, since algae are net oxygen consumers at night. Normal practice is to renew water in the larval tanks at night, so as to remove catabolites and to introduce oxygen when is most needed (final part of the light period).

A practical and effective way, though expensive, to ensure high DO levels is to add pure oxygen to the water in the tanks. This can be done either by injecting pure oxygen into the water piping system, or by bubbling it inside the tanks. The first way is more efficient since it prolongs the time in which oxygen and water are in contact before reaching the tanks, thus reducing oxygen losses. In a simpler but less efficient way, pure oxygen is directly injected into each tank through one or more fine air stones. Pure oxygen injected through additional diffusers must follow the same rule of avoiding potentially harmful water currents originated from the upward movement of oxygen bubbles.

In any case, it is recommended to install an emergency oxygen supply to overcome the risk of anoxic conditions that may occur unexpectedly. An oxygen supply can also be helpful when peaks of biomass are present during the last stages of fry rearing.

Fig.47.03 Stationary probe monitoring in the larval for O2 rearing tank (photo STM Aquatrade)

Outlet filters

The management of outlet removable screens follows the various feeding regimes. When water renewal takes place at dark, as fish larvae are visual predators they will not consume significant amounts of live preys at night during darkness. Rotifers and artemia nauplii lose nutritional value in a night and therefore can be eliminated during the water exchange. In this case a filter mesh of 500 µm is used to discharge them when water is renewed. When for any reason water has to be renewed also during the period in which lights are on, live feed has to be kept in the tanks for the fish which will be preying actively. In this case a 100 µm filter for rotifers and a 250 µm screen for artemia nauplii are used.

Outlet screens must be changed before they become completely clogged, and in any case should always be replaced with 500 µm screens just before dark. Check the filter material for dead larvae as this could be an early warning of possible troubles. It is recommended to have a set of outlet filters of different mesh sizes for every tank to save time during filter replacement.

After use screens should be cleaned by removing dirt with a water jet, then by a placing them for half-an-hour in a 500-ppm active chlorine bath followed by a thorough rinsing with freshwater. Screens should be stored in a place where they can dry out of the range of tank spray and should be at hand during replacement. If eggs are stocked directly into the larval rearing tanks, use a 400 µm filter during incubation and hatching.

Fig.48.01-2 Two different shapes of outlet screen for larval rearing tank (photo STM Aquatrade)

Fig.48.03-4-5-6 Larval rearing screen outlet changing, washing, disinfecting and stocking (photo STM Aquatrade)

Feeding seabass and gilthead seabream post-larvae

After being stocked in the tanks (150 to 250 larvae per liter should be a correct density), the larvae will continue their development during some days relying only on their yolk sac reserves. Depending on the rearing temperature they will start feeding on living micro-organisms in three to four days from hatching.

First feeding

At hatching, fish larvae are not yet completely formed and, among other things, lack functional eyes and mouth. Moreover they do not have an active swimming behaviour. In the first three to six days after hatching, and depending on water temperature, the fish larva therefore relies only on its yolk sac reserves as food source. At the end of this period the young fish has developed functional eyes, which are recognisable by their dark colour, its mouth has opened and the digestive tract, though still primitive, can now assimilate food. Then, its swimming behaviour becomes active and the animal is thus able to keep a horizontal position. At this stage the post-larval stage begins and the young fish starts feeding on live preys, such as rotifers or brine shrimp nauplii (the latter only a first feed in the case of seabass). These live feeds are supplied on time to the larval tank (see annex 17 and 18 for feeding regime of seabass and gilthead seabream).

The onset of the first larval feeding is a crucial step in the young fish life: if something goes wrong, starvation quickly kills a weakened animal. Starvation is actually a major cause of larval mortality. Therefore it is important to help them to overcome this delicate and key phase. The main actions to be taken are the following:

Fig.49.01-2 Gilthead seabream larvae: before first feeding and with functional digestive tract (photo STM Aquatrade)

Fig.50.01 Manual live feed distribution (photo STM Aquatrade)

Fig.50.02 Larval rearing tank during Artemia nauplii feeding (photo STM Aquatrade)

Table 3.10 - Repletion rate and number of ingested preys per larva at first feeding of gilthead seabream and seabass larvae.


Repletition rate

Actual ingested rotifer/ARTEMIA


Gilthead seabream


3rd day




4th-5th day




6th-8th day




9th day


up to 50

up to 100


Transition from live feed to artificial food

Feeding on live prey usually lasts 40 to 50 days, according to water temperature, species, rearing protocol and the opinion of the larval unit manager. The sooner the fish move to an artificial diet, the better in terms of savings on labour, overall costs and time. With this aim in mind, the timing of first dry food supply has been continuously anticipated in recent years, thanks also to new more elaborated artificial diets, including vitamins and immunostimulants, which fit better the larval requirements in terms of composition, size, buoyancy, and flavour. Special processing techniques can now blend high quality ingredients into micro-particles showing excellent stability in water, a slow sinking rate and attracting fish larvae better.

Before these advanced diets were available, fish were weaned on freshly prepared wet diets, formulated according to the nutritional requirements of other fish species. Special attention had to be given to the quality of the raw materials and integrators, as well as to their processing. A detailed description of such a wet diet (moist feed) and an example of feeding rates are presented in Annex 24.

The main difficulty encountered when trying to feed an artificial diet for the first time is how to stimulate fish to accept it, which is a time-consuming task since they are used to live preys. This delicate transition is indeed one of the main sources of larval mortality. In practice, at the beginning inert feed is distributed daily in very small quantities to accustom fish to a new flavour. To incentivate its consumption, it is advisable to start the distribution of inert feed in the morning, after the night starvation, and well before live feed is offered. Once dry food is accepted by most fish, it can be distributed by automatic feeders. To induce feed ingestion by the rest of the fish population, brine shrimp can be dropped close to the automatic feeder. The slow water movement will attract fish just below the feeder. When feeders are employed, distribution can be automatically set with a timer. In any case it is recommended to watch closely the ingestion rate. Artificial feed leftovers have to be avoided not only because they represent a loss of money, but mainly because they can severely pollute the water in the tank.

Fig.51.01 Belt feeder and the Artemia nauplii dispenser during gilthead sea bream weaning (photo STM Aquatrade)

From a management point of view, feeding fish with an artificial diet involves a series of changes in the rearing environment (see Annex 13):

In larval rearing cannibalism is not yet a major problem as it is in the weaning unit, but towards the end of the production cycle it may require attention, in particular when fish of different sizes and stage of metamorphosis coexist. The problem tends to be more serious with seabass. In presence of cannibalism, the supply of both artificial and live feeds should be better calibrated and its frequency should be increased. Another possible solution is to dilute the population in two or more tanks or introduce microalgae again, which would reduce the visibility and in consequence the more aggressive behaviour. This subject is extensively treated in the description of the weaning sector below.

Feeding protocol

As already indicated, the diet of seabass and gilthead seabream post-larvae in their first weeks of life is represented by small animals, rotifers and brine shrimp larval stages, whose biology, culture methods and nutritional value have been previously discussed in the manual. Microalgae are also provided to feed rotifers and improve the overall quality of the rearing environment. This section will deal with their handling and distribution to fish.

The feeding protocols for gilthead seabream and seabass are given in Annexes 17 and 18 respectively. In both cases, the protocol is based on the following assumptions:

Gilthead seabream aged 3 to 7 days receive a daily amount of 20 million rotifers per tank together with 40 liters of mature algal culture (at 12x106 cell/ml). From day 8 to day 12 the amount of rotifers is increased by 20% to 24 millions and to 28 millions from day 13 to 16, whereas the algal supplement remains at 40 litres.

From day 17 the first brine shrimp nauplii are fed to the postlarvae: they should be of a particularly small strain in order to facilitate their gulping by the still small-mouthed fish. The Artemia AF cyst are an example of such strains. The amount used ranges from 0.1 to 0.5 millions. At the beginning, frequent controls on fish are recommended to check is they are accepting the new food item. Together with the first nauplii, the rotifer ration is increased to 32 millions, whereas microalgal supplements are progressively reduced to 20 litres. At this time the first artificial feed (of a very small size, 80-200 µm), is also distributed. The quantity offered is limited to 1-3 g, but its function at this stage is mainly to start getting postlarvae accustomed to this new taste.

From day 20, algae are further reduced to 10 litres, rotifers quantities begin to decrease (to 20 millions), being replaced by an increased amount of artemia AF (0.5-1 millions) and, for the first time, also artificially enriched artemia metanauplii, produced with cheaper artemia strains (0.3-0.6 millions are offered). Inert feed is also gradually increased to 10 g.

From day 24 to day 27 algae are progressively eliminated, rotifers decrease to 10 millions, whereas artemia AF increases to 1.5 millions and artemia EG or RH to 3 millions. Artificial feed is also increased to 15 g.

From day 28 the distribution of algae, rotifers and artemia AF nauplii is suspended and only artemia EG or RH (10 millions) and inert feed (15-20 g) are fed to fish, whose average weight should now be about 5 mg.

From day 34 to day 39 fish are given more EG or RH artemia (12 millions) and 20 g of inert feed of 80-200 µm size, plus 10 g of the larger 150-300 µm size.

From day 40 to day 43, when metamorphosis from post-larval to juvenile shape (fry) has started, the distribution of the 80-200 µm inert feed ceases, and it is replaced by more EG or RH artemia (up to 16 millions) and an additional 20 g of 150-300 µm feed. From this point the fish are ready to be moved to the weaning sector.

In seabass, whose post-larvae are much larger than in the case of gilthead seabream, feeding with brine shrimp starts two weeks earlier. Fish aged 3 to 7 days receive a daily amount of 20 millions rotifers, 2 millions small size brine shrimp nauplii (cysts Artemia AF or BE) and 40 liters of mature algal culture, the latter one decreasing gradually till day 23.

From day 8 to 12 the amount of rotifers is increased to 25 millions and AF or BE artemia nauplii to 3 millions.

From day 13 to 16, rotifers decrease to 15 millions, and AF or BE artemia nauplii ration is increased to 4 millions.

Fig.52.01 High concentration of gilthead seabream around Artemia nauplii dispenser (photo STM Aquatrade)

From day 17 a few grams (between 1-3 g) of artificial feed of very small size (80-200 µm) are distributed together with 10 million rotifers for the smaller part of the fish population with 6 million of AF or BE artemia nauplii plus the first 2 millions of EG or RH nauplii for the larger fish.

From day 20, the enriched brine shrimps of large size(EG or RH) offered are 14 million, rotifers decrease to 5 millions and inert feed is gradually increased to 10 g.

From day 24 microalgae and rotifer distribution ceases, the EG or RH artemia nauplii ration increases to 16 million and the inert feed to 10 -15 g.

From day 28 the distribution of artemia is increased to 20 millions and inert feed stays at 10-15 g, although but 10 g of 150-300 µm feed are added.

From day 34 the fish get the same previous amount of artemia metanauplii. Distribution of the 80-200 µm inert feed particles ceases and 20 g of 150-300 µm are fed.

From day 40, when metamorphosis from post-larval to juvenile (fry stage) is almost completed, the artemia metanauplii ration is decreased (down to 16 millions) and the 150-300 µm inert feed is increased to 20 g. From now on fish will be ready to move to the weaning sector.

Fig.52.02-3 UV lamp and titanium plate exchanger are frequently used for temperature and bacterial growth stabilisation of the whole larval rearing unit. (photo STM Aquatrade)

Daily distribution of live feed

Rotifers and brine shrimp are distributed by hand into the areas of the tank surface where larval density is lower. Feed is distributed three times per day, starting as soon as the lights have been switched on in the morning until four hours before the artificial sunset, in late evening. A quick distribution of the first ration in the morning is recommended to stop the forced starvation, which takes place during darkness.

The daily ration should be distributed every 6 hours in the following way (time is indicative):

A prey density check is highly recommended before the second and the third distributions to adjust the concentration and thus avoid situations of over or under feeding. Use a 1-ml pipette to take two to three samples at different places in the tank and count the number of rotifers or artemia nauplii at naked eye or with the help of a portable lens.

From an operating point of view, the head of the larval sector prepares a feeding schedule for the day with the quantities to be distributed to each tank for the three meals. Rotifers and brine shrimp are quoted in millions: the worker has therefore to convert this figure in liters of stocked culture, according to the density marked on each stocking tank.

The live feeds should be distributed by keeping the jar above the water surface at the centre of the tank, and gently pouring its content over the rising air bubbles, to obtain an optimal dispersion. The operator has to take the highest care to avoid any splashing, waves or current the could induce stress to larvae.

First feeding with rotifers has already been discussed. Shifting from rotifers to brine shrimp is done progressively, feeding artemia nauplii from small size strains to gradually adapt the growing larva to the new food. The worker should check the ingestion of Artemia by regularly sampling larvae with a transparent beaker and looking at their digestive tract, which assumes a pale orange coloration when nauplii are ingested. See Annexes 21 and 22 for examples of record-keeping forms to help daily and weekly management of live food production and distribution.

When rotifers are fed to the fish larvae, microalgae are added daily to the rearing tanks to obtain a final density of 500 000 to 800 000 cells/litre. Because of their high PUFA content, preferable species are Nannochloropsis oculata, N. gaditana and Isochrysis galbana (Tahitian strain), the latter being particularly rich in the essential DHA.

Apart from maintaining the rotifer high nutritional value, their other positive effects in the intensive rearing environment are thought to be a certain bacteriostatic capacity and a shading effect that reduces the larval aggressive behaviour. Larval culture in clear water is also feasible, but it gives lower average results in terms of survival and size homogeneity.

In large hatcheries, where algal volumes to be distributed are important, a more practical way is given by the use of centrifugal reversible pumps connected to a stocking tank placed above the larval tanks. The required daily volume of algal culture is pumped into the tank and then easily distributed to the rearing tanks by gravity.

Notice: before distributing the microalgae, the tank cleaning must have been completed in all tanks as water turbidity prevents from seeing the bottom.

Fig.52.04 Wine reversible pump used to transfer live food (photo STM Aquatrade)

Daily storage of live feed

As soon as produced, live feed is stored in the daily stocking tanks of the larval rearing unit, to be available for distribution during the hours when the lights are on. The most practical tanks are PRF rounded containers with a conical bottom equipped with a drainage ball valve. Their capacity ranges from 100 to 500 litres although larger tanks are also used, according to the larval unit size. A central strong aeration obtained with a coarse air stone maintains live feed in suspension and keeps oxygen levels within safe margins. It is in any case advisable to install an emergency supply of pure oxygen as not only the nutritional value of dead rotifers and brine shrimps is low, but also the risk that they can spoil the rearing environment is very high.

To prevent the loss of their nutritional value, live feed is kept at low temperatures (5-10°C) which reduce the metabolic rate. For this purpose, a complete heat insulation is applied to the tank sides, bottom and removable top. Styrofoam or polyurethane mats and bubble plastic foils are commonly used. Sealed ice bags or blue ice packs are then introduced in the tank and regularly replaced when melted. To avoid contamination, sealed ice bags should be discarded after use, whereas blue ice packs should be disinfected in a 500 ppm hypochlorite solution before reuse. Remember to control the presence of punctures and to thoroughly rinse them with freshwater before freezing.

An alternative and more appropriate option is to stock the concentrated cultures in buckets into a big freezer, whose thermostat has been set to a temperature range of 0-10 °C. The lid of the freezer is drilled to fit the air or pure oxygen hose and supply the highly dense populations with the necessary oxygenation and water circulation. As already mentioned the best cold storage is obtained using a refrigerated tank.

Maximum daily stocking density for live feed is as follows:

Remember to clearly mark the stocking density on each tank, otherwise the staff in the larval rearing unit will not be able to distribute the live feed properly.

At the end of the day any live feed leftover can be concentrated by filtering, then packed in sealed plastic bags, named and dated and deep frozen. These leftovers can be used as an emergency supply in case of a production failure or delay, bearing in mind their generally reduced nutritional value.

Hygiene in the larval rearing environment

Due to the relatively high water temperature and density of living organisms, hygienic conditions in the larval rearing tanks can deteriorate rapidly. Moreover the quantity of organic matter creates a favourable environment for bacterial growth, often harmful to the delicate larval fish. A complete control of the hatchery hygienic conditions should therefore be contemplated and duly enforced. The health status of marine fish larvae depends mainly on a well-planned prevention including the treatment of the raw seawater and the compliance with strict cleaning routines, as detailed in Annex 16. Table 11 below gives a minimum daily planning for larval rearing controls during the day. The procedures check list is given in Annex 23.

Table 3.11- Daily schedule of controls



control 1/

Control 2/

Biological control

Various works





Rotifer/Artemia quality

Light switched on,
Change screens


Larval behaviour

Clean make-up
cartridges 3/
Clean UV/by-pass
Siphon tank bottom




Larval behaviour


Rotifer/Artemia quality









Rotifer/Artemia quality

Siphon tank bottom




Rotifer/Artemia quality





Larval behaviour





Larval behaviour

Change screens


Switch off light

1/ T-DO = Temperature + Dissolved Oxygen measurements
2/ WAF = Water renewal + Aeration + Filters overall controls
3/ In case a semi-closed system is present

Monitoring and controls

As soon as the larval population develops an active behaviour and starts feeding, the following controls should be performed to monitor its health status:

Quantitative evaluation of feeding performance

Presence and quantity of ingested food gives such a clear indication of fish health status to become one of the most important controls to be carried out during the first ten days of larval rearing. To correctly estimate the preying activity, a sample containing 30 to 50 fish from each tank population should be examined under the microscope every day. This figure can be doubled in the very first days of feeding. See Table 11 above for feeding patterns of seabass and gilthead seabream post-larvae.

Larval feeding examination of each tank:

Try not to damage the larvae with the pipette. Remember that an excessive pressure on the cover slide may result in the expulsion of eaten rotifers through the anus of the larvae. Morphological condition assess should be made as quickly as possible since heat irradiated by microscope lamp makes the larval body to shrink in a minute.

The presence of ingested rotifers can be easily recognized by the presence of their masticatory parts (mastax), left undigested in the larval gut. Under a 100x magnification they clearly appear amongst rotifer’s eggs and other debris. In practice, only the number of mastax found is recorded to estimate the total number of rotifers eaten. The number of preys per larva ranges from 2-3 (early feeding) to over 50.

To check the ingestion rate of brine shrimp nauplii, it is sufficient to visually estimate the percentage of repleted larvae (carefully sampled in a 100-ml transparent beaker and checked for their reddish-coloured digestive tracts as the deep orange nauplii are visible through the transparent larval skin).

Observations on prey quantity should be integrated with an assessment of fish behaviour, which can be done easily and in a quicker way (see below the section on evaluation of stress).

Qualitative evaluation of stress

Stress in fish larvae induces both morphological and behavioural changes that can be detected by the hatchery operator in order to improve culture conditions or to replace as soon as possible a poor larval batch. The main criteria for stressed fish larvae are:

Starvation is the more obvious indication that something has gone wrong. It is a general response to stress and therefore it is impossible to link this deadly condition to a single possible cause. With the sole exception of acute intoxication, all rearing parameters, alone or more probably in association, may stop larval feeding. As a starved fish will not survive for long, it is also important to monitor the onset of first feeding (see above).

There is some scientific evidence that a direct correlation exists between environment-induced stress and the appearance of calculi in the urinary system of gilthead seabream and seabass larvae. Although there is no confirmed correlation between calculi and death, they are often associated to starvation and consequently are present at a higher rate in dead larvae. The early appearance of calculi in a larval population is therefore considered as a stress indicator.

Fig.53.01-2 Gilthead seabream calculi (photo STM Aquatrade)

Calculosis can be easily detected by examining the lower part of the larval urinary duct (urethra and urinary bladder) under a microscope at 100 magnifications. This condition becomes evident by the appearance of a single stone-like corpuscle or a chain of smaller ones, reddish or grey in colour. Sometimes, they completely obstruct the urethra. The count of renal calculi may be done when the repletion rate is being evaluated and can be recorded in the same sheet. When a large calculosis, say in more than 40% of the fish examined, is observed, it can be interpreted as a sign of poor rearing conditions which typically will result in a low survival rate. In this case environmental and feeding parameters such as the following should be checked:

Another stress revealing parameter is represented by the presence of a large amount of post-larval fish that do not actively swim and attack preys. At the mercy of water currents, their passive behaviour prevents an efficient hunting of live prey. Such abnormal conditions can be easily detected because the artificial lights reflects in their eyes in a typical and specific way when these animals turn passively upside-down: small glimmering points appear throughout the rearing tank. This condition may lead to the loss of a consistent part of the population. An unhealthy rearing environment, together with a possible congenital factor, are considered as the most probable causes of this distress.

By comparison, a healthy fish displays the following normal preying behaviour:

Larval schooling starts at an age of 5 days and ends approximately between the 10th and the 15th day. It is believed that schooling indicates a healthy population. The so-formed larval shoals result from:

In each tank there are one or more shoals, typically in the calmest places of the tank, slowly moving around. Samples to control the state of the population are taken inside these shoals.

A particular type of behaviour, an erratic swimming at the water meniscus, should also be considered as a possible response to stress. This syndrome is characterized by a frenzied activity of larvae, which seem to be attracted by the water meniscus where they get stuck, beating the tank wall head-on or being shaken by spasmodic head-up movements. This affects their feeding rate and consequently their survival.

Apparently only a small part of the population shows these symptoms for a long time, thus suggesting a possible chronic stress or unsuitable rearing conditions. Therefore, massive mortality will occur only in populations that exhibit the meniscus-stress syndrome to a large extent. Usually, this syndrome accounts for part of the “normal” larvae mortality when their age is between 10 and 30 days.

If measures cannot be taken to counter the stress inducing factors, the larval section manager should consider the opportunity to eliminate the compromised population and quickly start a new batch.

Control of swim bladder development

At 18°C, the swim bladder formation starts between day three and four. The first evidence of swim bladder inflation is noticed between days five and seven, looking as a small air bubble inside a tissue vesicle that becomes clearly visible under the microscope (at 40x magnification). A few days later, a second bubble develops and joins the first one to form a almost spherical body that will gradually expand to become an elongated vesicle.

Many authors agree that the initial activation of the swim bladder relies on the gulping of air at the water surface. A temporary connection between the swim bladder area and the mouth (typical of physoclist teleost fish) makes this process possible. This active air swallowing is apparently crucial for a proper swim bladder development: if this air gulping cannot take place, the swim bladder will not form, as the conduct remains open only for a few days. The absence or the incomplete filling of an active swim bladder has severe consequences for the fish, causing a deformed backbone. This, linked to limited or negative buoyancy and an abnormal swimming behaviour, reduces the feeding rate and slows growth. Even if the deformed fish reaches marketable size, its price will be well below that of normal fish. Therefore an early and correct determination of the percentage of swim bladder inflation is vital to decide how to proceed with the rearing programme.

Amongst the factors interfering with normal swim bladder activation the following are considered to be important:

Any obstacle at the air/water interface that prevents larval fish to gulp air is responsible for a poor rate of swim bladder inflation. The presence of an oily layer originating from rotifers being fed an enrichment diet is considered particularly dangerous.

Fig.54.01-2 Surface skimmers (photo STM Aquatrade)

The introduction of “surface skimmers” has overcome this problem. These floating devices are traps that continuously remove any floating debris and grease by blowing air at low pressure tangentially to the water surface. Skimmers are periodically cleaned (at least three times per day) with either a beaker or a using a soft paper foil.

Fig.55.01 Heavy loaded skimmer before cleaning (photo STM Aquatrade)

In the early larval stages the swim bladder can be easily observed under a microscope at 20 to 40 magnifications, thanks to the transparency of the larval body and the brightness of the gas bubble, whose surface reflects light as a mirror. Monitoring should be made after complete filling of the swim bladder on samples aged 15 to 20 days.

The procedure for swim bladder inflation control in each tank is as follows:

1. Sample 30 to 50 larvae per tank or use the sample taken for checking first feeding;

2. Pipette the larvae on a slide with as less water as possible. Group them together.

3. Cover with the glass lid, remove excess water with filter paper and observe them at 20 and 40 magnifications

4. Look for the presence of the swim bladder.

5. Record all findings on a dedicated form (Annex 19).

Fig.56.01-2 Cleaning procedure for surface skimmers (photo STM Aquatrade)

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