The early life stages of seabass and gilthead seabream are zooplankton-feeders, i.e. they prey on small free living planktonic animals. As no artificial larval diet can at present totally fulfil their nutritional requirements, their successful rearing still depends on an adequate supply of high quality live feeds, usually in the form of rotifers (fed on unicellular algae) and brine shrimp (Artemia spp).
This chapter describes the equipment and operation to mass produce these organisms, whose biology has been presented in Part 2. The design of the hatchery sections for production of live feed is described in the Design and Engineering part of this manual.
The technology for phytoplankton and zooplankton mass production has become very reliable and the production of live feeds is part of the standard working procedures in Mediterranean hatcheries. The efficiency of this part of the hatchery mainly depends on the implementation of standard procedures by well trained staff.
As live preys for first postlarval stages of seabass and gilthead seabream, two small animals are extensively used:
In farms dealing with seabass only, live feed production is often limited to the hatching of Artemia nauplii, which are obtained by incubating their dry resting eggs (cysts). The mouth of the seabass at first feeding is large enough to gulp brine shrimp nauplii and these larvae do not require the smaller rotifers as first feed, as it is in the case of gilthead seabream. The hatcheries working on both species, or on gilthead seabream alone, have to produce rotifers as well as microalgae.
As said in previous sections of the manual microalgae are now used not only for rotifer production (see below), but also to improve water quality in the larval tanks, creating the so called green water, which is used during the initial rearing phases.
In case of rotifer mass production, clear advantages of this organism are given by its fast reproductive rate and by the high densities that can be reached in the rearing facilities, up to 1,000 rotifers per ml and over. The daily increase of their populations ranges between 50 and 150%, depending on the production technique chosen and the nutritional value of their diet. Their main drawback is that to culture them microalgae are needed as food, at least in the initial steps. However, for their final production process in large volumes (see below) there are now good artificial feeds which can replace algae, whose mass production remains unavoidable at least in gilthead seabream production for the greenwater.
Fig. 23.00 A sort of simplified trophic chain is established in the hatchery.
On the other hand, the production of Artemia is greatly facilitated by the availability of dry resting eggs, which can be purchased from specialised suppliers. If properly canned and stored, brine shrimp cysts can remain viable for years.
As rotifers and nauplii are produced to fulfil the needs of the larval rearing unit, they have to be available at given times, in pre-set quantities and with their nutritional quality intact. To achieve this, the design and operation of the culture systems should pay special attention to the following points:
Mass production of phytoplankton for rotifers and green water in most Mediterranean hatcheries is limited to a few species such as: Chlorella sp, Isochrysis galbana, Pavlova lutheri, Nannochloropsis oculata and N. gaditana, Dunaliella tertiolecta and Tetraselmis suecica. These species have been selected on the basis of their size, nutritional value, culture easiness and absence of negative side effects, such as toxicity. Their nutritional value shows a great variability not only among different species, but also in genetically different populations of the same species (strains). For hatchery purposes, the species to be cultured should both fit well the local rearing conditions and have a high nutritional value for rotifers. The increasing availability of nutritional boosters as enrichment diets for both rotifers and brine shrimps, has made this choice easier.
Fig. 23.01 Mass culture of microalge (photo STM Aquatrade)
Microalgae population dynamics can be described by different phases:
It is advisable to harvest phytoplanktonic organisms during their log phase, since in the new culture they will grow more rapidly and will yield a more viable population.
For aquaculture purposes, microalgae are mass produced in three main ways: (i) batch (or discontinuous or multistep back-up system) culture, (ii) semi-continuous culture, and (iii) continuous culture.
In the batch culture a small axenic stock culture produces a series of cultures of increasing volume where the algal population of each culture vessel is entirely harvested at or near its peak density, i.e. while still conserving a good growth potential, to be used either as inoculum for other culture vessels, or to feed rotifers or be used in fish larval tanks. It typically makes use of small (few liters) to medium size (500 liters) containers, and it is kept indoor and under strictly controlled, if not properly axenic, conditions. It is considered by many authors the easiest and most reliable method of algal production, provided that the working protocol is strictly enforced. Algal quality is less erratic than in the semi-continuous method, even if the latter is more productive for any given volume.
In the semi-continuous system the algal population, when mature, is partially harvested at intervals. The harvested culture volume is replaced by fresh medium to keep growth going on. This culture is adopted to produce large amounts of algae and frequently uses large outdoor tanks. Their main drawbacks are: (i) the unpredictable duration, (ii) the risk of contamination by other organisms as competitors (other microalgal species), contaminants (bacteria) and predators (ciliate protozoa feeding on the algae), as well as (iii) the building up of metabolites, which can affect quality.
The continuous system is a steady-state continuous flow culture in which the rate of growth is governed by the rate of supply of the limiting factor. It is a balanced axenic system where the algal population is harvested and fertilised continuously. This method, though the most efficient over extended periods, produces limited amounts of high quality cells and requires complex equipment as well as advanced management. A relatively recent development of this system is represented by the photo-bioreactor, a continuous culture device that increases the density of cultured microalgae to very high levels under predictable environmental and microbiological conditions.
The microalgae produced can be concentrated to a dense liquid suspension by centrifugation, and can then be stored for more than one month in the refrigerator, still giving excellent viability when used. A new industry is now appearing, whose concentrated algal products can also fulfil the hatchery needs, saving the time-consuming and expensive production of microalgae in the hatchery.
The system described below is the batch culture, by far the most widely adopted method by Mediterranean hatcheries. Before its description, additional instructions are given concerning facilities, the preparation of the culture medium, and the equipment required.
Fig. 23.02 Old fashioned unit using artificial light for algae mass culture (photo M. Caggiano)
Algae are cultured in a dedicated sector of the live feeds production section, which is made of three working areas inside the hatchery building: a lab for duplicating small cultures, a conditioned room to maintain small culture vessels and pure strains and finally a large area for the mass cultures in PE bags or, less frequently, tanks. In the warmest Mediterranean areas, a light greenhouse can replace the latter.
Small volume cultures are kept in vessels ranging from 20-ml test tubes up to 18 l carboys. They can be made of borosilicate glass, polycarbonate, PET or any other material able to stand a sterilization process. These vessels are placed on glass shelves lightened by fluorescent tubes and equipped with a CO2 enriched air distribution system.
Hot-extruded tubular PE film is utilised for larger volumes bags. The film is usually 0.25 mm thick and its stretched width ranges from 45 to 95 cm. Two bag designs are widely adopted in Mediterranean hatcheries: the smaller suspended bag and the larger one placed within a steel wire cylindrical frame. The first type has a capacity of 60 I (single) to 150 I (double or U-shaped), whereas the latter, that stands on a saddle-like GRP base to improve circulation, can contain up to 450l. Their top is closed by a plastic cover to prevent contamination.
Fig 24.00 A typical scheme of a batch type production
All units are equipped with artificial lights, usually fluorescent tubes, an aeration system, often with an additional source of carbon dioxide, and stands for the culture vessels, i.e. light shelves for small volumes and metal racks or wired frames for PE bags.
The unit also stores the special equipment to process pre-treated seawater, such as fine filters and sterilizers, as well as a laboratory where nutrients and glassware are prepared and stored, and where the necessary monitoring operations are performed. Standard cleaning procedures have to be strictly followed to maintain proper hygienic conditions (see Annex 6).
In planktonic mass production, seawater is the medium of all culture vessels. The use of other mediums such as agar is limited to the preservation of pure algal strains. Ideally, seawater should be free from pathogens and pollutants. With this aim in mind, seawater is treated to remove suspended solids, contaminants and organisms and to improve its original parameters to fit the quality standards set for proper growth of microalgae. These methods are outlined below, while the description of related technical equipment can be found in the Engineering part of the manual.
The most common systems of seawater mechanical treatment (applicable to all culture volumes: strains, small volumes and mass cultures) for microalgae production (which also apply to rotifers and brine shrimps cultures), are described below.
Raw seawater is first pre-treated to remove the bulk of suspended solids and contaminant organisms. Different methods are followed, but the most commonly used is a combination of settling and sand filtration. If properly dimensioned, a settling tank is a useful device. Not only it improves the water quality at no cost by decanting its suspended matter, but also provides a reservoir of seawater to tackle unpredictable problems in seawater supply, such as a damaged main pump station or a temporary contamination of seawater (oil spill, river plume, exceptional storm). Settled water is then filtrated through a sand or bag filter that retains particles as small as 50,10 or 5 mm depending from the sector of destination. At this point filtered water can be used for most purposes in the hatchery, provided for some sectors like pytozooplankton it undergoes UV sterilisation before use. In the live feeds sector a higher degree of filtration is required and pre-treated water is further micro-filtered to a size of 5 mm for large culture volumes and down to 1 mm for small volumes and strain cultures For this finer filtering polyethylene wire cartridges, bag filters or diatomaceous earth filters are used. Such a fine filtration should even remove bacteria and other micro-organisms, but in reality the filtering capacity is not absolute and cannot totally guarantee such results, in particular under hatchery working conditions. It is therefore recommended to proceed with the final step, sterilization of filtered seawater.
Fig. 24.01 Mechanical sand filter for the water inlet (photo STM Aquatrade)
Different methods of water sterilization have been developed. The following description refers to the most common methods adopted for hatcheries. The choice is based on local availability of equipment and service and depends also on the amounts of water to sterilize, which are related to the size of the hatchery.
UV light sterilisation (applicable to all culture volumes)
UV light with a wave length of 265 nm (short wave UV or UV-C) has a strong germicidal effect based on its capacity to break the DNA helix. It is produced by special high or low pressure mercury vapour lamps whose germicidal capacity depends on several factors such as their power, seawater transmittance (transparency to UV), type and quantity of microorganisms to be destroyed, degree of purification required, water flow (contact time) and temperature. Seawater to be sterilized flows through one or more sealed chambers where it is irradiated by one or more lamps placed inside quartz tubes (transparent to UV light). The thickness of the water film inside the irradiation chamber should be such as to allow the maximum sterilization effect. If the power of UV lamps and chamber design are properly dimensioned, the contact time between water and quartz tubes is only of few seconds.
Fig. 24.02 Compact water treatment unit for zooplancton (photo STM Aquatrade)
For practical purposes, and under normal hatchery conditions, an intensity of at least 40 mJ/cm2, provided at the end of the life span of the lamps, removes 99% of most unwanted micro-organisms for fish farming from the treated water volume. With its auxiliary equipment (manual or automatic wipers, UV sensors and stabilizers, computer-aided control), this method is very effective and manageable and fully justifies its cost.
Chlorine sterilization (applicable to all culture volumes)
Active chlorine is a strong oxidizing agent, commercially available as liquid bleach (sodium hypochlorite or NaOCl) and as bleaching powder (CaOCl2). The percentage of active chlorine in these chemicals should always be checked in advance as it changes widely according to the producer: commercial grade NaOCl usually contains 5-15% active chlorine, while CaOCl2 contains 60-70%. Annex 7 describes how to prepare hypochlorite solutions, to assess their active chlorine content and the residual chlorine in treated water, as well as the methods to sterilize seawater and deactivate residual chlorine.
Independently from the method employed, a final dosage of 5 to 10 ppm of active chlorine is used to sterilize seawater. The contact time between water and chlorine should be at least one hour, after which any residual chlorine must be neutralised with sodium thiosulphate, Na2S2O3, (see Annex 7 for details). This technique is now widely used as final sterilisation step of water in larger vessel and of the culture equipment (air and oxygen tubing and diffusers, detritus traps, submersible water heaters)
Fig. 24.03 Chlorine container for laboratory use (photo STM Aquatrade)
Use of an autoclave, or wet vapour sterilization (applicable to small volume cultures)
With this method, applicable only up to 5-6 I volumes depending on the autoclave size, seawater is sterilised together with the culture vessels, usually made of Pyrex® glass, due to their resistance to heat. The autoclave should work at 120°C under a 2 atm pressure. Sterilization time ranges from 10 min (100-ml flasks) to 20 min (200-ml flasks) and 30 min (up to 5-6 I vessels). The neck of each container has to be covered with a loosened aluminium foil stopper to let vapour out during the sterilization.
Dry vapour sterilization (applicable to small volume cultures)
This method replicates the previous one, but the autoclave is replaced by an oven. As it works in dry vapour at ambient pressure, glass vessels filled with seawater are heated at 160-170°C for 2 to 3 hours. Dimensions and stoppers of vessels to be sterilized are the same ones used for wet vapour sterilization.
The exponential growth of the microalgal populations is regulated by four most important parameters: light, pH, turbulence and nutrients. Whereas the first three can be easily adjusted specific nutrients have to be added to the culture medium in proper quantities.
The main nutrients required are nitrogen (N) and phosphorus (P), followed by trace minerals, vitamins and chelating agents. Nutrient solutions are prepared in advance according the type of microalgae cultivated. With the exception of N and P solution, the other nutrients are stored as primary stock solutions, which are used to prepare the working solutions according to the day-by-day production schedule. Aseptic conditions have to be maintained in the preparation of the enrichment solutions. The vitamin solution cannot be sterilised because heating will deactivate the vitamins. The microalgae selected for the reproduction of seabass and gilthead seabream require the following fertilizers that refer to the enrichment medium Guillard f/2.
Trace elements and vitamins are first prepared as concentrated primary stock solutions: in this way, if properly stored, they may last several months. Trace elements are prepared as four different solutions, each stored, like vitamins, in a separate container. To prepare one litre of each stock solution, the following quantities (in grams) are required.
Trace element stock solutions
Solution A: ZnSO4 · H2O (30g) + CuSO4 · 5H2O (25g) + CoSO4 · 7H2O (30g) + MnSO4 · H2O (20g)
Solution B: FeCl3 · 6H2O (50g)
Solution C: Na2MoO4 · 2H2O (25g)
Solution D: EDTA · 2H2O (50g)
To prepare the solution put the components, according to the proportions indicated above, into one 1 -I graduated Pyrex® bottle and fill with distilled water (DW) to the mark. Deionized water can also be employed if distilled water is not available. When the components are fully dissolved, store at ambient temperature, avoiding exposure to direct light.
Vitamins stock solutions
B12 Cyanocobalamin (0.1 g)
B1 Thiamin (10g)
H Biotin (0.1g)
Place the indicated quantity of each vitamin into a sterilized 1 -I graduated Pyrex® bottle filled to the required volume with sterilized DW. When fully dissolved, store in refrigerator and keep away from light. The B12 solution should preferably be stored in a dark or aluminium wrapped bottle.
Warning: do not sterilize any vitamin solution.
The working solutions represent the way to add the nutrients, trace elements and vitamins directly to the seawater medium. Two working solutions are prepared by diluting the above mentioned stock solutions, whereas the third solution is prepared directly from industrial grade chemical salts of N, P and K.
Fig. 24.04 Working solutions ready to use (photo STM Aquatrade)
Mineral salts working solution
Mineral salts solution: NaNO3 (300g) + KH2PO4 (30g) +NH4Cl (20g)
Put the salts into one 1-I screw-capped oven-resistant glass bottle and fill with DW to the mark. If not available, deionized water can also be employed. When fully dissolved, sterilize either in autoclave or oven. Store at ambient temperature, avoiding direct light. Due to the comparatively higher requirements for the mineral salts stocking solution, quantities in excess of one liter are usually prepared at one time. Use 5 to 10-l glass autoclavable vessels, then store in 5- or 10-I plastic carboys with bottom tap.
Use: 1 ml per litre of seawater medium.
Trace elements working solution
One liter of trace element working solution is prepared according to the sequence and quantities oulined below:
Solution D (100 ml) + Solution A (10 ml) + Solution B (10 ml) + Solution C (10 ml).
Use only sterile glassware: a 100-ml cylinder and three 10-ml pipettes (one per solution). Mix the four solutions into a 1-I screw-capped oven-resistant glass bottle and fill with DW or deionized water and sterilise either in autoclave or oven. Store at ambient temperature, avoiding direct light.
Use: 1 ml per litre of seawater medium.
Vitamins working solution
Mix the following amounts of vitamins stock solutions: solution B12 (10 ml) + solution B1 (10 ml) + solution H (10 ml).
Use only sterile pipettes, one per vitamin. Dilute the mix to one litre volume with sterilized distilled water. Pour in a dark sterilized bottle (or wrapped in aluminium foil) and store in the refrigerator just before use.
Use: 1 ml per litre of seawater medium.
Warning: do not sterilize any vitamin solution
As for water medium and nutrients, also the equipment should be kept clean and disinfected to prevent contamination.
Small glassware such as volumetric pipettes, Petri dishes, Pasteur pipettes and beakers, are sterilized either in an autoclave or in an oven. Bigger glass containers as Erlenmayer flask, balloons and carboys are sterilized after being refilled with seawater. Pipettes are divided by volume capacity and stored into capped metal containers or wrapped in aluminium foil. Polyethylene (PE) bags are considered sterile as they are obtained by hot extrusion and do not need special treatments. Tanks, plastic jugs for nutrients and piping for pump transfer are disinfected with hypochlorite.
Fig. 24.05 Pipette container used with oven sterilization (photo STM Aquatrade)
All up-scaling tools and consumables that may be in contact with algae (aluminium foil, platinum needles, necks of tubes, flasks and balloons) are sterilized by flame using a Bunsen burner. On the glassware neck, flaming should lasts until all water drops have evaporated.
Sterilized hydrophobic cotton is commonly used as disposable stoppers for all glass containers. For its sterilization, it is packed in aluminium foil and sterilized in autoclave or oven. Cotton stoppers are disposed of after use. Hygiene and cleaning procedures in the live feed production sector are outlined in Annex 6.
This section describes how to proceed with the fertilization of the different culture vessels. All working nutrient solutions are diluted at 1 ml per litre of seawater medium. Pure strain cultures in test tubes, in marine agar and their initial small volumes are fertilized with half dosage (0.5 ml/l).
Fig. 24.06 Phytoplankton strain cultures (photo STM Aquatrade)
With small vessels, this operation should be carried out in a dedicated room kept clean and equipped with all the necessary tools, glassware and consumables. To reduce the risk of contamination, a UV-light ceiling lamp can be installed to provide germicidal irradiation when the room is not in use.
Enrichment procedure for small vessels (up to 20l capacity):
1 prepare on the bench the required vessels containing sterilized water, nutrient working solution and the necessary tools;
2 carefully loosen their aluminium foil caps;
3 heat each flask neck and the inner side of the aluminium foil caps with a burner; close again;
4 heat opening and screw cap of each bottle containing stock solution in the same way;
5 take a sterile pipette from its sealed container (if in a metal case, open and close it at the flame);
6 with the pipette place one nutrient solution at a time in each flask, using a new pipette for each solution;
7 repeat the flaming of necks and aluminium caps on the enriched flasks.
Enrichment procedure for polyethylene bags (up to 500l capacity):
1 The bags are filled with the same treated and UV-light sterilized seawater utilised for small V vessels, the only difference being that the bags are not sterilized. Due to the larger amount of nutrients involved, use clean and sterilized graduated cylinders to fertilize bags.
2 Cut a 20 cm-long slit on the upper rim (empty part) of the bag;
3 add 1 ml per liter of each solution.
When cutting the bag, bear in mind the final culture level, i.e. after the addition of the inoculum.
Mass culture of phytoplankton starts from pure strains of selected species and proceeds by upscaling from small (0.5 I) up to large volumes in PE bags (450I) or tanks (1 000 l and up). In finfish breeding, where large amounts of microalgae are not required, the last upscaling step is the PE bags. See annex 8 for a typical example of microalgae upscaling protocol, and annex 9 for the daily workplan and culture file for microalgae.
Fig. 24.07-8 Flask sterilization and enrichment (photo STM Aquatrade)
The culture of pure strains of the selected microalgal species is the starting point of the mass production process. Strain quality is therefore essential for any successful production process. A good selection of pure strains of different algae should always be kept in a dedicated facility in the hatchery. This practice also allows the selection of the most suitable strains under local conditions and at a given time.
Pure guaranteed strains of algae, as well as of rotifers, are normally available from a few laboratories and institutions in the Mediterranean region and Europe. It is strongly recommended to regularly renew their lines, not only in case of culture crashes, but also to control the often unavoidable contamination or decline in quality. Strains from other hatcheries should better be avoided because of their possible decline in quality and the associated risk of introducing contaminated strains.
The algal pure strains are kept under standard controlled environment in conditioned rooms or in especially designed incubators, in which routine work can be performed under strict hygienic control. The pure strain cultures are usually kept in small glass containers, such as 10 to 25 ml test tubes or 100-ml glass Erlenmeyer flasks, closed by a sterile stopper (screw cap or a folded aluminium foil).
Pure-strains cultures should be maintained at a steady or resting stage, i.e. under environmental conditions which allow them to reproduce, but not to increase exponentially in number. In this way, their sexual reproduction is fostered, thus increasing their genetic variability, and the growth of unwanted organisms such as other algal species, bacteria and ciliate protozoa is prevented. Culture parameters are therefore kept below the values adopted for mass production. In particular, only half dose of nutrients is used, water temperature is kept at around 14-16°C, light intensity ranges from 300 (test tubes) to 1 000 lux (flasks) and no aeration and carbon dioxide are provided.
Under routine conditions, strain cultures are usually renewed every month. In the replication process, an inoculum of 0.1-0.2 ml (from test tubes) or 0.5 to 1 ml (from Erlenmeyer flasks) is taken from the best old culture which is free of contamination, to inoculate three new vessels of the same size to start a new strain. The old culture is then either utilised for upscaling, or is discarded. Strain culture vessels should be stirred at least once a day by hand, paying attention not to stir bottom debris up.
Warning: in the management of pure algal strains quality is essential: get rid of any tube or flask which is found to be contaminated by bacteria, fungi, ciliates, nematodes or different algal species.
The steps to duplicate algal strains at test tube level are described in the following paragraphs. All required equipment and consumables have to be well cleaned and sterilised in advance.
Fig. 24.09 Pure strains kept under controlled conditions (photo STM Aquatrade)
Choose the test tubes that show the best algal populations at naked eye and check under the microscope a sample taken from each of them for contaminant organisms (use a new sterile pipette for each sampling). Then keep only the uncontaminated cultures and put them on a stand avoiding any stirring.
For each selected test tube:
1 prepare four sterilized test tubes with cap on a stand;
2 prepare at least 50 ml of sterilized seawater medium, enriched with half dose of the standard nutrients mix (see above);
3 heat necks and caps of all test tubes (new and old vessels) by means of a manually operated burner and let them cool;
4 fill each new test tube with 10 ml of seawater medium taken with a graduated pipette;
5 with a sterile 1 ml-pipette take 0.5 ml of mature culture from near the surface of the selected test tube;
6 Inoculate 0.1 ml of the old culture into each new test tube; be careful to make the drops fall freely into the culture medium without touching the tube walls; with one hand open and close the tube cap; with the other hand handle the pipette; do not mix or agitate the tubes;
7 place the used pipettes into the rinser cylinder;
8 after inoculation, heat the upper part of each new test tube thoroughly and cover with its cap, previously flamed; as an alternative, use sterilized hydrofobic cotton or flamed aluminium foil as stopper;
9 discard the old culture, if hot needed for other replicas, and clean the empty tube following the usual cleaning routine for glassware (annex 6);
10 write date and algal species on all new test tubes with a waterproof marker;
11 place the newly inoculated test tubes on a rack in a shelf reserved to pure strain cultures.
Even under axenic conditions, pure strains can be contaminated by other algal species or micro-organisms: in such cases before up-scaling the cultures the microalgal populations have to be purified before being used as inoculum.
Two methods to purify contaminated algal cultures are of common use: successive dilutions of the original contaminated culture, and picking up of single cells from the original culture. Both techniques are also applied when new algal species are isolated from the wild.
A simple technique to purify a contaminated strain is to proceed with repeated sub-cultures obtained by progressive dilutions of the original sample. Dilution rates can be very high, depending on the number of the sub-cultures. In the procedure described below a dilution rate of 10-10 is reached. The same nutrient quality and environmental condition of the initial culture are used.
Example of dilution:
1. prepare 10 sterile test tubes with caps on a tube rack;
2. sterilize 500 m I of seawater in an Erlenmeyer flask, then fertilize it with half dose of usual mix of nutrients when cool;
3. using a sterile 10-ml pipette add 9 ml of enriched seawater to each tube, close loosely with flamed caps and number them 1 to 10;
4. put the test tube containing the strain to be diluted in the rack, remove the cap and flame its neck;
5. using a 1-ml sterile pipette take 1 ml and add it to the tube No. 1, then stir gently;
6. using a new sterile 1 ml-pipette repeat the previous step by taking 1-ml inoculum from tube No. 1 and inoculate it into tube No. 2;
7. repeat the same procedure with the remaining tubes, each time pipetting 1 ml from the previous tube (gently stirred) into the next one; flame necks and caps and let them coo);
8. keep under controlled environmental conditions.
When cell growth reappears, check samples of the tubes under the microscope and get rid of the tubes that are still contaminated, typically the initial ones, and keep only the purified cultures, usually in the more diluted tubes. Repeat the process using the last dilutions if necessary, and in any case at least every three months to always have a safe amount of purified cultures ready at hand.
Picking up method
Two systems which produce single cells are described below: the agar plate method and the capillary method:
Agar plate method
A solid medium can grant more stable conditions for the growth of the desired species. This technique needs some sterile equipment like Petri dish and platinum hooks.
1. prepare the solid medium by adding 1.5 g of agar powder to 100 ml of sterile fertilised seawater;
2. heat the medium using a Bunsen burner, stirring with a sterile glass rod to dissolve the agar, and pour it on 3 to 5 sterilised Petri dishes;
3. allow to cool and solidify;
4. using a sterilized 1 ml-pipette take 0.1 ml from the initial contaminated strain culture and drop it into each Petri dish;
5. spread the sample over the agar with a sterile platinum hook and incubate at desired environmental conditions placing the dish upside down, so that water drops will not form on the lid and then fall on the culture;
6. once algal colonies are observed, take a sample by means of a sterile platinum hook and check under the microscope
7. monospecific colonies should be kept in agar for successive replication or might be up-scaled by replication (i.e. a small portion of the colony together with some agar, using a sterile hook) on a 50-ml or 500-ml Erlenmeyer flask to continue in the liquid medium.
This technique follows the dilution method, but the inoculum is obtained by selecting single cells of the desired species by means of a capillary pipette handled under a microscope.
The isolation or purification of cultured strains should be repeated as many times as required to produce contaminant-free cultures.
As indicated above the upscaling of microalgae production starts from small containers (0.5 ml) and proceeds through various steps up to the mass production in PE bags (up to 450l). Each step involves an increase of the culture volume: when mature (i.e.: in log phase), the algal population of a smaller volume is sacrificed to replicate the same vessel and to inoculate larger vessels.
Small-medium size (0.5 to 10l) cultures of microalgae are usually kept in borosilicate-glass containers with large necks, such as Erlenmeyer flasks or other flat bottom round flasks (balloons) and carboys. The flasks are usually closed with a stopper made of sterilised cotton or of plastic which can stand sterilization in autoclave.
Fig. 24.10 Petri dish with culture solid medium (photo STM Aquatrade)
All these cultures are provided with a proper medium to support algal growth (treated and fertilized seawater), good aeration supplemented with carbon dioxide (CO2) as an additional carbon source, and a strong light. In this way it is possible to reach quickly the log-phase growth. Aseptic conditions should be maintained to avoid contamination and culture crashes.
For practical purposes the main parameters for the cultures are usually the same for the different species of algae mentioned in the sections above:
20 ± 2°C
25 to 30 ppt
4000 to 8 000 lux
50 to 100% of the culture volume per minute
2 % of air volume
Fig. 25.01-2 Algae flask replication (photo STM Aquatrade)
For mass culturing purposes, the optimal temperature for the above mentioned algal species ranges between 20 and 24°C. Generally speaking, temperatures lower than 16°C and above 27°C will slow down growth rates, whereas those above 30°C are normally lethal. That makes room conditioning necessary as artificial lights and insufflated air from the aeration system can raise temperature dangerously.
Low temperatures are used for pure strains only, where growth should be kept as slow as possible. As low temperatures also affect bacterial growth, non-axenic cultures should be maintained at the lowest possible temperature consistent with a good growth, to prevent bacterial growth.
In the selected species salinity does not represent a limiting factor within a range of 15 to 40 ppt.
Light is the source of energy for photosynthesis and therefore in mass cultures algae are usually kept in continuous light. Fluorescent tubes are the commonest choice for providing light due to their low power consumption, low installation cost and limited heat production compared to bulb lamps. Spectral quality suitable for algal growth is provided by Cool white and Daylight models, often installed in equal numbers. The commonest tube size is 42 with waterproof contacts. See Part 4 Engineering for their installation in the hatcherys algal unit.
Even if each algae species has its own preferred light intensity for best growth, for practical purposes intensity in mass cultures is kept in the range of 2 500-8 000 lux, while higher intensity is used on the small volumes shelves (5 000-8 000 lux) because of their higher cell density. Large volumes (bag cultures) can also be illuminated by natural daylight, entering from sufficiently large windows or from a continuous glass wall, provided that the cultures are not directly exposed to external contamination.
Fig. 25.03 Lighted table used for mid size culture up scaling (photo STM Aquatrade)
Aeration is used to maintain the culture in turbulent state, preventing settling of cells and exposing all cells to light. It also supplies carbon dioxide, which is fixed by the algae during photosynthesis, and provides essential pH stabilization.
Transparent PVC tubing, of 6 mm internal diameter, are commonly utilized to deliver air to the culture vessels. In small vessels this tubing is connected to glass pipes which fit the stopper and reach the bottom of the vessel, while in PE bags they are simply forced into a small hole near their bottom (the water pressure keep the holes sealed), whereas in tanks a weight keep them submersed.
Aeration should be moderate in the first two days of culture, then should be increased adjusting it according to culture growth. For this purpose screw clamps or cheaper plastic needle aquarium valves are required to adjust the air flow. If aeration produces foam, it is an early warning of culture troubles.
As its normal content in air is low (approx. 0.03%), carbon dioxide is often supplied at 2% by volume to optimize culture growth. Commercial grade CO2 bottles are utilised, and the gas is injected into the main air pipe via a dispensing vessel in which gas bubbling reveals the gas flow. Since carbon dioxide is heavier than air, to prevent stratification the pipe makes some ups and downs after the point where it is injected.
Depending on the algal species, the CO2 supply, and the volume of inoculum, working cultures normally reach their log-phase in 5 to 7 days. At this point, cultures can be utilized either to start new cultures, to be fed to rotifers or to be used as green water in fish tanks..
The volume of the algal inoculum is usually 15 to 20% of the new volume. Smaller or larger inocula could be used to decrease or increase growth rate. In culture up-scaling, the new vessels have to be inoculated with a sufficiently high microalgae density in order to ensure a rapid growth and to limit the risk of contamination with different algae or other micro-organisms (protozoa, nematodes, fungi, etc.).
The following section describes in detail the operation to replicate different volumes.
Inoculation of flasks from test tube
Follow the same procedure as previously described for pure strains. Each 0.1-1 flask will receive 50 ml of enriched medium and 0.5 ml of inoculum. At this stage, no aeration is required. When mature, each small flask will inoculate a new 2-l flask.
Fig. 25.04 Alternative use of olives containers for algae culture (photo Ittica Mediterranea)
Fig. 25.05 Very small PE bags of phytoplankton at Ittica Ugento (photo STM Aquatrade)
Fig. 25.06 Medium and large size bags (photo STM Aquatrade)
Inoculation of 2 and 6-l flasks from another 2 or 6-l flask:
1. prepare the necessary amount of new flasks filled with sterilised seawater, as well as all equipment required for the operation (pipettes, nutrients, cotton stoppers, aluminium foil, glass tubing, etc.);
2. select the mature culture that will be used as inoculum, checking a sample under the microscope for contaminants;
3. remove its cap and flame its neck; then close with a flamed aluminium foil stopper and let it cool;
4. in the meanwhile, add the fertilizing working solutions to each new flask, at a rate of 1 ml/l; using a new sterile pipette for each solution;
5. flame their necks and aluminium caps thoroughly;
6. when cool, remove the stopper and pour some algal culture of the old flask into the new vessels at a rate of about 10% of the receiving volume, avoiding to wet their neck with the inoculum, then gently shake flasks to mix the new culture;
7. flame thoroughly their necks, introduce the sterilized glass tubing for aeration and close tightly with sterilised cotton stopper (or any other type of sterile cap);
8. write date and algal species on the new flask;
9. place the flasks on the lighted shelf and connect to the air delivery system, adjusting its flow to a gentle bubbling;
10. after on hour, check all new vessels for a proper air bubbling.
Use only the upper layer of the old culture, leaving dead cells and debris in the flask used to inoculate the new ones. The size of the inoculum for small volumes is only 10% of the new volume because of the high cell density. Remember to get rid of every contaminated flask. The above mentioned procedure applies to the upscaling of the other medium size vessels.
Fig. 25.07-08 PE bags inoculation with the help of an self-priming pump (photo STM Aquatrade)
Inoculation of a PE bag from a flask or from another bag
1. prepare the bag, either suspended to a rack or placing it inside a cylindrical frame made of wire;
2. fill the bag with sterilized water; leaving enough free space for the volume to be inoculated; wait a couple of hours to check for possible leaks; if found, seal them or replace the bag;
3. introduce nutrients and inoculum;
4. fit two PVC tubes for aeration to the bottom of the bag, connect them to an air line and adjust the air flow to a gentle bubbling;
5. add the nutrients using a graduated cylinder at the usual rate of 1 ml/l of each working solution;
6. select a mature culture that will be used as inoculum, either from a large flask or from another bag, and check a sample under the microscope for contaminants;
7. in case a flask is used as inoculum, remove its cap and flame its neck; then close with a flamed aluminium foil stopper and let it cool;
8. then, remove the stopper and pour the algal culture from the flask into the bag. The inoculum should be about 10% of the receiving volume;
9. if the inoculum comes from another bag, with a self-priming plastic pump transfer the inoculum from the old bag into the new one. The volume to be transferred should be about 15-20% of the receiving volume (a bigger inoculum is necessary to compensate for a less dense culture);
10. on each bag mark date, algal species, origin of the inoculum (bag or flask) and the bag serial number.
To reduce the risk of contamination, smaller bags are usually inoculated from flasks, whereas larger bags are inoculated from smaller bags.
Fig. 25.09-10-11 Large volume cultures (photo STM Aquatrade)
A regular check of microalgae cultures is essential to prevent crashes and to keep high quality standards. The main parameters to be monitored are: colour, density, pH and contaminant levels. As an example, a change in colour to opaque grey and a pH level lower than 7.5 may indicate a high degree of bacterial contamination. A lighter colour than normal may reveal insufficient nutrients or poor lighting. However, during the peak production season in the hatcheries a close monitoring of algal cultures can hardly be assured, due to the chronic lack of staff and time. Mass cultures are normally checked at naked eye by experienced staff and strict controls are usually restricted to pure strains and small vessels.
To partly by-pass this problem, the staff can do a good preliminary job outside the peak production season, when they are not so absorbed by production and there is some more time. Then test culture cycles can be closely monitored, the algal culture growth can be followed daily by counting the number of cells per ml with a haemocytometer (see below), and their average growth curves can be plotted against values obtained with a colorimeter. During more busy days, a colorimetric monitoring (comparing values against the test curves) gives a fast and reliable indication of algal cultures growth.
These test cultures are useful for a number of other reasons as it is possible:
A Neubauer haemocytometer can be used for counting microalgae cells with diameters ranging from 2 to 20 mm and at densities up to 500 million cells/ml.
This device consists of a thick rectangular slide with an H-shaped trough delimiting two counting areas. With its special cover slip in place, each area forms a chamber 0.1 mm deep. The total area of each chamber is 9 mm2. Each chamber is divided into 1 mm2 squares: the four corner squares are subdivided into 16 smaller squares, whereas the central one is subdivided into 25 smaller squares, 0.2 x 0.2 mm, each with an area of 0.04 mm2.
To count cells in a sample, proceed as follows:
1. take a 5-ml sample from each culture and place each sample in a separate test tube. Add one drop of Lugol solution and mix well;
2. prepare clean Neubauer slides and covers;
3. put one drop of well mixed algal suspension on each Neubauer chamber and cover with its cover slip;
4. check under low magnification that the algal cells are evenly distributed: avoid the presence of air bubbles, over flowing, underfilling and uneven distribution of cells. Allow the cells to settle for 5 minutes before counting;
5. start counting at the top left square and count only those cells which lie within or touch the boundary lines chosen according one of the two possibilities: either left and bottom boundary lines or right and top boundary lines;
6. for cells larger than 6 mm and not too dense, make a total count in each of the four corner squares and in the central square; then repeat this count in the second chamber;
7. for minute cells and denser populations, count the cells in 5 smaller squares in the larger central square, then repeat this count in the second chamber;
8. calculate average cell density as follows: if all cells are counted in individual blocks each with an area of 1 mm2 and a volume of 0.1 mm3, the average cell density expressed as cells/ml is given by the total count divided by the number of blocks and multiplied by 10 000;
9. if all cells in ten (five in upper chamber + five in lower chamber) smaller squares (volume = 0.004 mm3)in the central block are counted, the average cell density is the total count multiplied by 4 000 000 and divided by 10, i.e. the total count multiplied by 400 000;
10. record the count for each sample and introduce it in the algae population growth curve prepared for each culture container.
For the breeding of many marine finfish species the rotifer Brachionus plicatilis is, up to now, the only live feed that can be used in their very early larval stages. While not a compulsory choice in seabass feeding, its mass production is required for the successful breeding of gilthead seabream (and of other Sparids, grouper and grey mullet), whose small mouth cannot accept larger preys at the onset of larval feeding. Among other valuable characteristics as live feed for fish, B. plicatilis was also chosen due to the relative easiness to culture it in large scale.
Two main strains are used. The so-called small strain (S-type) and a large strain (L-type), 50% bigger in dry weight than the S-type. The average length of the lorica in the adult S-type rotifers is 130 µm, whereas it is 240 µm in the L-type. The two strains also show different temperature and salinity tolerances. In the last years mainly S-type rotifers are reared in the hatcheries.
Fig. 26 Brachionus rotundiformis and Brachionus plicatilis (modified from Fu et al.)
Rotifers can reproduce both sexually (mictic reproduction) and asexually (amictic reproduction), according to the environmental conditions. Usually amictic, rotifers may turn to sexual reproduction when sudden changes in salinity or temperature take place. Then, they produce large resting eggs, similar to brine shrimp cysts. However, in hatcheries is the asexual reproduction that provides the large amounts of animals required for the early feeding of fish larvae. Rotifer population dynamics under mass rearing conditions follow different phases, mimicking those of microalgae:
The quality of the rotifer population to start new cultures is even more important than in the case of microalgae. To be used as inoculum, the rotifer population must still be in the middle of its log-phase with at least 20% fertility rate (measured as percentage of eggs over total rotifers. Populations in their last declining phase, characterised by limited motility, scarce repletion and absence of egg-bearing animals, should always be discarded. With a proper inoculum and under optimal rearing conditions, a rotifer population should reach its harvesting density within 4 to 5 days.
Under hatchery conditions, rotifer populations can reach the following densities:
in flasks and bags, after 5 to 7 days:
- S-type rotifers, 500 to 700 ind/ml
- L-type rotifers, 150 to 250 ind/ml
in tanks, after 4 to 6 days:
- S-type rotifers, 1000 and more ind/ml
- L-type rotifers, 400 ind/ml
As this microscopic animal is a filter feeder, its nutritional value strictly depends on its food. In hatcheries, the species is first cultured on microalgae, following the same scale-up protocol described for microalgae, then its final mass production is achieved in large tanks where artificial diets are fed to rapidly increasing numbers, improving at the same time their nutritional value.
The rotifer B. plicatilis is a rather sturdy species able to tolerate a wide range of salinity, temperature and ammonia levels. It can also use several food sources, provided that particle size remains within a 2-20 µm range. Obviously, the highest growth rate is achieved under more restricted environmental parameters, and is closely related to the selected rotifer strain and feeding provided,. In particular, good yields are obtained with high dissolved oxygen levels, temperature at 25°C, pH at 7.5-8.5, salinity in a 20-30 ppt range, less than 1 mg/l free ammonia (NH3), and moderate turbulence. Light is required only when rotifers are fed microalgae.
Dissolved oxygen levels
During the upscaling phase in algal vessels, the oxygen requirements of rotifers are fulfilled by microalgae photosynthetic activity. In mass culture, artificial diets, enrichment boosters, high rotifer densities and metabolic products, contribute to deplete oxygen in the culture medium. To keep its levels within safe margins, i.e. above 80% saturation, a strong aeration is supplied, linked to an emergency oxygen delivery system. See section below for technical details.
In mass culture, a temperature as high as 30°C is acceptable, but as bacterial and other contaminants would also increase at these levels, a more manageable 20-25°C range is commonly adopted.
Fig. 27 Medium and large volumes cultivated in the same room (photo STM Aquatrade)
An acceptable range is pH 5-9, but the buffer capacity of seawater keeps it within a narrower range. A low pH also influences the balance between the toxic un-ionized ammonia and the ionized form (NH4+). The lower pH also influences the balance between the toxic un-ionized ammonia and the ionized form (NH4 reduces the fraction of toxic NH3.
The acceptable salinity range for this rotifer species is quite broad: 1-60 ppt. However there are differences between the two strains, the optimal salinity for the S-type strain being 18-20 ppt, whereas L-type grows better at 30 ppt.
A moderate to strong turbulence is required to keep food particles and rotifers in suspension. The aeration system should provide enough water circulation, but the position of the air diffusers should be adjusted in such a way to avoid stirring and re-suspension of bottom sediments: airstones are hung 15 cm above the bottom, both along the periphery and in the centre of the tanks.
Inside the hatchery building, phytoplankton shares the same culture facilities with rotifers, which have a specific section with large tanks for their final mass production. In Mediterranean hatcheries, these tanks have a volume ranging from 1 to 10 m3. These tanks are made of FRP, PE or PVC-lined concrete. As water circulation and particles sedimentation are important in rotifer culture (see below), a widely adopted design is the round tank with conical bottom. This section for mass production of rotifers is equipped with an air distribution system, air conditioning and seawater heaters to keep high water temperatures. Special filtering devices to harvest and rinse rotifers are also available.
As mass culture of rotifers does not require light (except for the phases in which algae are used as food), a normal lighting system is installed. Treated seawater (filtered and sterilized) is provided by the same source that feeds the algal section.
As in microalgae production, hygiene is important and requires the strict implementation of standard cleaning procedures (see Annex 6).
The same procedures and precautions described in the algal production section apply to rotifer culture. The only enrichment added to the rotifer culture medium, be it either a log-phase algal culture or treated seawater plus artificial diet, is represented (cyanocobalamin) as a fertility by the addition of vitamin B12 booster for the rotifers. Its dosage is usually 100 ml of B12 stock solution per m3 of rotifer culture in tanks, whereas small vessels and bags are fertilized at the rate of 1 ml/litre. In both cases the vitamin is added together with the inoculum.
To prepare the stock solution put 0.1 g of vitamin B12 into a sterilised 1-l graduated Pyrex bottle and fill with sterilised DW to the mark. When fully dissolved, store in the refrigerator. Warning: never sterilise vitamin solutions.
Fig. 28 Large rotifers tanks for mass culture (photo STM Aquatrade)
The first phases of rotifer cultures (strains and small volume cultures) are performed in the same conditioned room where microalgae are kept, but on separate stands to avoid the risk of possible contamination. The same propagation techniques described for microalgae strains are used for rotifer strains. As for microalgae, culture conditions discourage exponential growth. Rotifer strain inoculums are kept at 1 or 2 individuals per ml in a low-density algal culture and no vitamin is added. Both test tubes and small flasks are used to maintain rotifer strain cultures.
A clean culture of egg-rich rotifers from a 0.5-l flask is usually inoculated directly into a 5 to 10-l flask, bypassing the 2-l flask stage. The inoculum should provide an initial density of 10 to 20 rotifers per ml. A larger inoculum results in a faster pace of population growth. Rotifers should always be inoculated in algae cultures which have not yet reached their peak growth (that is their log-phase). Environmental conditions should be the same ones at which algae are kept. However, aeration can be reduced to diminish the amount of foam and bottom sediments produced by the metabolic activity of rotifers.
Vitamin B12 is routinely added to all new vessels, at 1 ml of stock solution per litre of algal culture. Mature rotifer cultures from small vessels (5 liters or larger) are poured into bags with an algal population that has not yet entered its full log phase. Five to ten % inoculum is used at this stage, i.e. one 5-liter vessel is used to inoculate one 100-l bag, following the rule of the new culture starting at 30-50 rotifers/ml. Using a sterile volumetric cylinder, add 1 ml of the Vitamin B12 per litre of culture; mark the date on the bag surface, together with the origin of the inoculum and the serial number to facilitate its handling and record keeping.
If the population is growing normally and remains free from contaminants, the horizontal culture method is also frequently used for rotifer culture upscaling. In this case, a good bag makes it possible to inoculate a group of new bags with a initial density of at least 30-50 rotifers/ml.
In the same way as it happens in microalgae cultures, also rotifer cultures are subject to occasional failures (culture crashes). A culture crashes either when rotifers do not multiply as foreseen, or when the entire culture dies abruptly. The possible causes are described below in the Monitoring section. As a precaution against these problems, always maintain a certain number of culture bags in excess of those required by the production schedule. They could be useful to replace quickly a crashed culture at any stage.
Fig. 29 Mature algae bags ready to be inoculate with rotifers (photo STM Aquatrade)
Rotifer mass culture is carried out in large tanks as mentioned above. Because of the very high density achieved (up to 1.000 individuals per ml or more), rearing procedures and protocols to maintain strict hygienic conditions (see Annex 6) have to be rigorously applied. Such routine procedures are described below.
There are basically two main mass rearing methods for rotifers: the older technique that is based on algae and bakers yeast as food for the rotifers, and the more recent one that uses an artificial diet, the Culture Selco® produced by INVE SA of Belgium (or similar). Both are described below.
Before starting a new production cycle, normally after the harvest of the previous rotifer culture, prepare the tank as follows:
1. rinse with tap water to eliminate the bulk of organic debris;
2. wash it thoroughly with brush and detergent and rinse it again;
3. wash or spray the tank walls with 500 ppm active chlorine solution (Annex 6); after a couple of hours, drain the tank and rinse it well until the chlorine smell is gone;
4. let the tank dry and fill it with sterilized heated water only when needed; as an alternative possibility, fill the tank with seawater and sterilize with hypochlorite, then neutralize the residual chlorine with sodium thiosulphate (see method in Annex 7).
Repeat the procedure for the equipment to be used in the tanks: aeration tubing, drain valves and suspended traps (see below). A practical procedure is to assemble all small equipment in the new tank, fill with SW and sterilize with hypochlorite: the equipment will be disinfected as a consequence.
To be used as inoculum, the rotifer population must still be in the middle of its log-phase with at least 20% fertility rate (measured as percentage of eggs over total rotifers). Never use rotifers which have already reached the last phase characterised by limited motility, scarce repletion and no egg presence.
In algae/yeast fed tanks, the initial density of inoculum, one of the most important factors in rotifer culture, should be kept at least at 100 animals/ml, with an optimal density of 150-200 animals/ml. High density cultures with artificial feeding need up to 500 rotifers per ml (see below). With an initial density of 200 animals/ml, rotifer density should reach its peak within four to six days at 25°C.
Fig. 30.00 The classic cylindroconical rotifers tank (photo STM Aquatrade)
Tank inoculum may come from large rotifer bags (vertical upscaling), or from other tanks (horizontal upscaling). One day before being harvested to be used as inoculum, rotifers in the tanks should be fed a fertility booster such as Protein Selco® by INVE SA. In order to reduce contamination and culture crash risks, horizontal upscaling should be limited to 7-8 cycles, after which the inoculum should be again taken from large bags.
As previously indicated two feeding methods are most widely adopted: 1) a combination of algae and bakers yeast and 2) a totally artificial diet. For its reliability and higher output, the latter is progressively replacing the first method.
Mass culture with algae/yeast as food
The initial method of mass culturing rotifers in Mediterranean hatcheries, makes use of a common and easily available food staple, the bakers yeast Saccaromyces cerevisiae. It is a labour and cost sparing food, which has no nutritional value for rotifers that feed on bacteria associated with the yeast.
Compared to the artificial diets, this method has a lower yield and requires more time, typically one extra day. Density at harvest rarely exceeds 450 rotifers/ml with an average daily increase ranging from 19 to 33%. In addition, rotifers should be enriched with high levels of (n-3) HUFA and vitamins. A major constraint of this method is the absolute necessity to improve the otherwise very poor nutritional quality of yeast-fed rotifers before their distribution to fish larvae. The enrichment procedure, which takes place the day before harvesting the rotifers is explained below.
Fig. 30.01 Large containers with hypochlorite solution are used to keep cleaned small equipment (photo STM Aquatrade)
1. fill the tank with sterilized sea water diluted with tap water to obtain a 20 ppt salinity; check the chlorine content of tap water and, if present, neutralize it with an excess of sodium thiosulphate (see Annex 7). Take care to leave enough space for the algal cultures to be supplied as food (about 30% of the tank volume).
2. place the air diffusers and switch on the aeration;
3. place the traps for ciliates and impurities;
4. select the most suitable rotifer bags to be used for inoculum, checking for contaminants and using only clean batches; 5. filter the selected batch or batches (see below for procedure);
6. inoculate the tank to achieve an initial density of 150-200 rotifers/ml. This is considered day 0;
7. add algal culture as 20% of tank volume to provide rotifers with their initial food; as usual, the algae should be in their log-phase and from non contaminated cultures, even if of different species.
8. the next day (day 1) fill the remaining 10% volume with algal culture;
9. on the tank file, record all information on the culture growth, food distributed and environmental parameters monitored (see below for details and Annex 9 for a file sample);
10. from day 1 on, feed with bakers yeast at the following rates, according to the recorded rotifer density.
Daily feeding rate
less than 50
more than 100
The daily amount of fresh yeast is divided into 4 equal rations fed at 2 and 8 am, 2 and 10 pm (the last two rations are given by the night watchman). Each yeast ration is taken from weighed out of the yeast cake kept in the refrigerator and placed in a plastic bucket filled with tap water at a concentration of 100 g/l. A kitchen or better an industrial blender helps to separate yeast cells and keep them suspended in water. Feed immediately and discharge any leftover. Prepare fresh for every meal.
Mass culture with Culture Selco® as food
A different technique based on a compound feed has been developed a few years ago by the Belgian Company INVE SA. The product, named Culture Selco® (CS) is a dry and complete rotifer diet that does not require algae and is also effective as enrichment medium. Particle size (5 to 7 µm) and physical characteristics ensure an optimal uptake by rotifers. The feed composition includes proteins (>35%), lipids (>15%, of which 23% are PUFA), carbohydrates (30%), carotenoids and other micronutrients as minerals and vitamins A, D3, F and C.
The average daily production of rotifers fed on CS ranges consistently from 45 to 60% of the initial rotifer density. In addition rotifers are enriched with high levels of the essential (n-3) PUFA and vitamins.
Fig. 30.02-03 Rotifers mass culture performed in very large heated tanks at Ittica Mediterranea (photo STM Aquatrade)
This diet has made rotifer mass culturing reliable and predictable, and has consistently reduced the need for algal cultures and their associated labour and facilities requirements. New rotifer cultures can be easily started from old ones, thanks to their enhanced fertility.
The following schedule gives the CS daily feeding rate (DFR), expressed as grams of CS per million rotifers. The day before harvest rotifers are enriched with other artificial diets made by INVE, Protein Selco® or DHA Protein Selco® (see Enrichment below for details).
a/Replace CS with Protein or DHA Protein Selco®
Fig. 30.04 An industrial blender for yeast or Culture Selco® suspension (photo STM Aquatrade)
To prepare CS suspend the amount required for a single meal in tap water, up to 50 g CS/l, and mix vigorously for 3 minutes (use a kitchen or better an industrial blender). Mixing or shaking by hand or using a magnetic stirrer is not sufficient to separate the CS cells. Remember that cell agglomerates left in the feed suspension cannot be ingested by rotifers because of their large size. Take care not to overfeed as uneaten feed can also quickly spoil water quality. Feed the daily amount in four to six meals evenly distributed over the 24-h period. In case it would be necessary, the feed suspension can be stored at a temperatures below 8°C, and the amount needed for the whole day can be prepared at one time. The feeding ration can thus be distributed from the stored suspension at each meal.
Fig. 30.05-6 CS preparation and distribution to the tanks (photo STM Aquatrade)
Before transferring rotifers to a new tank, place about 25% of the first days food ration in the tank, so that the feed would be already evenly spread when the rotifers enter the medium. Add a few drops of silicon based antifoam agent, such as Rhodorsil Antimousse AM 70414 by Caldic Belgium NV.
Aerating the rotifer culture is essential to provide oxygen, to keep rotifers and food cells suspended and to optimise tank cleanliness. The aeration rate and the air diffusers positioning should be carefully adjusted. A reasonably strong aeration is detected by an evenly spread water turbulence at water surface without large air bubbles.
Number and positioning of air diffusers depend on tank shape and volume. In all cases, however, they should be suspended at about 15 cm above the tank bottom to prevent re-suspension of sediments. As to the number of diffusers, a reliable rule of thumb is to keep a distance of approximately 60 cm among them.
Fig. 30.07 CS distribution by gravity to the tanks at Ittica Ugento (photo STM Aquatrade)
In round tanks with conical bottom, bottom sediments are removed by letting them first settle for 10 15 minutes in absence of aeration (take care to check DO levels during the operation). Then the bottom valve is opened for a few seconds and it is closed when the outflowing water is again clean. Repeat the operation twice a day, in the morning and evening.
Complete the cleaning procedure by removing any greasy layer that may be formed at the water meniscus with a sponge or a paper cloth. Never dip your hands in water.
The rotifer tank water can be rapidly polluted by particulate matter, faeces and flocks of uneaten food. Their removal may prevent an excessive bacterial development and will increase the oxygen available for rotifers. As the continuous aeration of the water volume prevents in part the settlement of particles, their removal is achieved by means of particle traps. These devices consist of floating mats of coarse sponge-like material, such as the housing polishing mat Scotch Briteä. In the case of a 3,000-l tank, three pieces measuring 15 x 100 cm are hung vertically in the water (by means for example of wooden stick laid across the tank top rim), and are kept vertical by a terminal weight. The water circulation pushes particulate matter against the trap, where they adhere and clog the mat. These mats are also an ideal substrate for Vorticella, a sessile ciliate that competes with rotifers for food. To be effective, the traps must be cleaned at least twice a day.
To clean particle traps proceed as follows:
1. take them out of the tank carefully, avoiding dripping of trapped material back into the water;
2. remove all particles with a high pressure jet of tap water;
3. dip the trap in a 500 ppm hypochlorite solution for one hour to disinfect;
4. rinse well with tap water, dip in a thiosulphate solution to neutralize residual chlorine and place again in the tank.
Fig. 31.01-2 Scotch Briteä after one night of work and during cleaning procedures (photo STM Aquatrade)
At harvest, rotifers are filtered and rinsed before being fed to fish or utilized as inoculum for new tanks. For this purpose, a double submerged filter is used. The inner filter has a mesh size of 250-300 µm to retain larger particles, flocks of agglomerated food particles and ciliates which would rapidly clog the finer filter. The outer filter has a 50 µm filter mesh. Its capacity should be large enough to keep safely the whole rotifer population for the time needed to complete harvest and rinsing. Both filters are placed inside a large wheeled container full of water to avoid pressure build-up from the outgoing water that would smear rotifers against the net. A gentle air bubbling along the inner side of the filter helps to keep the filter free from clogging.
Harvesting and rinsing protocol for round-conical tanks with central drain:
1. prepare the harvesting device, always clean and disinfected with hypochlorite;
2. inject pure oxygen into the tank to be harvested for 10 to 15 minutes to have a supersaturated medium (at 10 ppm DO) in which rotifers could safely stand filtering operation;
3. switch off aeration for 10 - 15 minutes and purge the bottom sediments;
4. into the central drain fit a PVC pipe with holes at 15 cm from the drain; wait 5 minutes;
5. fix a flex hose to the bottom drainage valve and place the other end into the harvesting device;
6. open the valve and start filtering;
7. regulate the water flow so that the filter does not clog and the culture water does not overflow; do not exceed 100 l/minute;
8. stop water flow and clean the clogged pre-filter whenever necessary to avoid overflow of rotifer concentrated water;
9. while harvesting, check for possible loss of rotifers through the net by sampling some filtered water with a beaker or a Petri dish;
10. at the end of filtration, close the valve and rinse for 10 to 15 minutes the rotifers with filtered sterilized seawater at the same temperature as the tank of origin.
Fig. 32.01-2-3-4 Harvesting procedures of rotifers mass cultures (photo STM Aquatrade)
Recent advances in mass culture technique yield higher rotifer densities. As an example, the procedure described below shows how to produce one billion rotifers daily with a battery of five 1-m3 round tanks with conical bottom.
Inoculum: 500 rotifers/ml, harvesting at 1 000 rotifers/ml in four days. Prepare five tanks following the above mentioned procedure.
Cleaning operations (three times a day):
1. remove traps, air tubing and diffusers and wash carefully with tap water;
2. stop aeration for 10 minutes and purge the tank cone through its bottom valve;
3. put back in place clean and disinfected traps, air tubing and diffusers.
Feeding schedule (distributed at 2 and 8 am, 2 and 10 pm)
a/Of which 50% Protein Selco® and 50% Culture Selco®: Calculate the daily amount, distribute before the inoculum 100 g of it (100 ppm in a 1-m3 tank), then divide the remaining quantity into three rations, using the same proportion for the whole day 0, at 4 and 10 pm and 2 am.
b/Replace Culture Selco® by Protein or DHA Protein Selco®.
c/1/3 of the harvest is used to inoculate other tanks, and 2/3 for fish larvae feeding (direct or after 12 hours-enrichment in a new tank).
Rotifers have a limited nutritional value for marine finfish larvae. In the past their nutritional value was upgraded by an enriching process before their harvest through feeding them with microalgae rich in PUFA and vitamins such as Chlorella, Nannochloropsis and Isochrysis.
At present, the enrichment is provided by specially formulated artificial diets like the above mentioned Selcoâ products. This oil emulsion gives excellent results in terms of high levels of EPA, DHA and vitamin C, which was not possible with the only use of algae. Moreover, labour, time, investment and running costs are spared. Rotifers can be enriched either in their mass culture tanks or after harvesting by placing them in dedicated enrichment tanks.
The first method produces an enrichment of the tissues, as it is continuous along the entire culture period. The acquired fatty acids reserves are more stable and are less exposed to a rapid decrease in nutritional value during starvation. This method also saves time and reduces handling losses.
The second system is a short term enrichment or, rather, a gut enrichment. It implies the harvesting and rinsing of the rotifers and the preparation of a separate enrichment tank.
Enrichment with algae:
Enrichment with Selco® products or other similar products:
Fig. 33.00 Enriched rotifer (photo INVE Aquaculture)
The content of nutrients decreases rapidly in rotifers that are not immediately consumed by fish larvae. In starving rotifers the total PUFA loss reaches 60% after 6h at 18°C. Even in green water, i.e. with microalgae, this loss remains important (about 40% after 6 h). To prevent this degradation in nutritional quality, enriched rotifers not immediately fed to fish should be stored in containers at low temperature as follows:
Check all rotifer cultures daily for both quantitative and qualitative evaluations. From each vessel, flask, bag and tank, take a 1 ml sample and observe under the stereoscopic microscope.
Measure the following parameters: quantitative parameters:
Fig. 33.01-2 Rotifers populations are daily checked and counted (photo STM Aquatrade)
In addition the following qualitative parameters of the culture should be checked:
All these information must be recorded in the individual file of each culture. All daily monitoring procedures for the rotifers sector are given in Annex 10.
Fig. 34.01-2 Micropipette and microscope used to evaluate population parameters (photo STM Aquatrade)
Having a larger size than rotifers, the larval stages of a small crustacean, the brine shrimp Artemia salina are used as the second (after rotifers), and last live food fed to fish larvae before their weaning on artificial feed. Strictly speaking, Artemia is not cultivated in the hatchery as is the case for algae and rotifers, but their larval stages are obtained by incubating and hatching their resting eggs, available as dry storable cysts, which can be purchased from specialised suppliers. The first Artemia larval forms, the nauplii, which are also the smallest and richest in yolk, are followed by a larger metanauplius, whose nutritional value has to be boosted by feeding them special enrichment diets 12 to 24 hours before being offered to fish.
Fig. 35.01-2-3-4 Various Artemia stages (photo INVE Aquaculture)
Many types of brine shrimp cysts from different locations in the world are marketed: they differ in naupliar size, important in gilthead seabream first feeding, in nutritional value and in hatching characteristics. For practical purposes they can be categorized as follows.
Table 3.6 - Biometrical analysis parameters of some Artemia nauplii strains
Two additional parameters characterize the Artemia batches: the number of cyst per gram and their hatching rate (the number of nauplii produced per gram of cysts). The best strains can give about 290 000 - 300 000 nauplii per gram of cyst hatched, with a hatching rate close to 95%. In a hatchery the use of good quality cysts allows a synchronization of the production cycle on a 24-h period, with the harvest of freshly hatched nauplii coinciding with the start of the incubation of new batches. Basic information on Artemia biology and life history is given in Part 2 of this manual.
Artemia cyst shells are usually contaminated with bacteria, spores of fungi and other microorganisms. Fish larvae can be infected when untreated empty shells, unhatched cysts or cyst hatching medium residues are introduced into the larval rearing tank.
Before incubation cysts should therefore be disinfected. This process also improves hatching by reducing the bacterial load of the hatching medium. Disinfection is done by keeping the cysts for a few minutes in a hypochlorite solution at a maximum density of 50 g/litre. This product is easily available as commercial grade bleach. The duration of the treatment varies according to the active chlorine concentration of the disinfecting solution. Typical duration is:
As in commercial bleach the chlorine content may range from 5 to 15%, it is mandatory to check the actual chlorine concentration in the bleach that is going to be used. This can be done either by titration or by determination of the refractive index. The titration method is explained in Annex 7.
The following example shows how to disinfect one kg of cysts in a 200 ppm hypochlorite solution obtained from a household bleach with 5% active chlorine:
1 one kg of cysts needs 20 l of fresh water for the disinfecting solution;
2 if this solution is going to be used for a 20 minutes bath you will need 20 l x 200 mg/l = 4 000 mg = 4 g active chlorine;
3 the quantity of 5% bleach required to give 4 g active chlorine is: (1 000/50) x 4 = 80 ml
4 pour 80 ml of 5% bleach in 20 l of fresh water;
5 add one kg of cysts; place an airstone for continuous aeration to keep cysts in suspension, and keep the cyst in the solution for 20 minutes;
6 harvest cysts on a sieve (125 µm mesh size) and rinse thoroughly with plenty of tap water;
7 transfer the rinsed cysts to the incubation tank.
A more effective way to obtain completely contaminant-free cysts is decapsulation, which implies the elimination of the cysts thick external layer, the chorion, by chemical oxidation. This process, which requires greater attention, has additional advantages. As they spend less energy to hatch after the removal of the chorion, the hatching nauplii have better nutritional value. Moreover, fish do not risk to suffocate by gulping empty or unhatched cysts offered together with the nauplii, as it may happen when using disinfected cysts.
The decapsulation process consists in four steps: hydration, treatment in a chlorine solution, washing and deactivation of the residual chlorine. The example described below refers to the decapsulation procedure of one kg of cysts.
The hydration, a necessary step as the complete removal of the chorion may only happen when cysts are spherical, proceeds as follows:
1. water volume required: around 6 l per kg (maximum amount: 200 g/l); both fresh and sea water can be used; water temperature should be between 20 and 25°C;
2. duration: 45 minutes;
3. aeration: sufficiently strong to keep cysts in constant suspension; use an open end pipe in a 10 l bucket;
4. collect the hydrated cysts on a sieve and treat them immediately with the decapsulation solution.
The decapsulation solution requires a source of hypochlorite, usually liquid bleach (NaOCl), and an alkaline product to increase pH level of the decapsulation solution above pH10. Usually technical grade caustic soda (sodium hydroxide NaOH) is utilized. The first product is added at 0.5 g active chlorine per gram of cysts, and the second as 0.15 g of sodium hydroxide per gram of cysts. For hydrated cysts the procedure is as follows (figures refers to one Kg of cysts):
1. prepare 0.5 g Cl x 1 000 g cysts = 500 g of active chlorine, equal to 3 33 l of a 15% bleach;
2. prepare 0.15 g NaOH x 1 000 g cysts = 150 g of NaOH, equal to 0.375 l of a 40% NaOH solution;
3. put the bleach and NaOH in a suitable container (e.g.: a 20 l plastic bucket) and fill with seawater to 14 litres (14 - 3.33 - 0.375 = about 10.3 l of seawater)
4. provide a strong aeration and eventually if available add antifoam;
5. place the hydrated cysts in the bucket;
6. control the temperature: it should remain within 25°-30°C. In case of higher temperatures, add ice to prevent that it reaches 40°C which are lethal for the cysts;
7. verify cyst colour changes. The change in cyst colour confirms that decapsulation is in progress: the cyst colour shifts from dark brown to grey and finally to orange, which is the colour of the nauplius body seen by transparency through its outer cuticular membrane, left exposed by the dissolution of the chorion. The process usually lasts 5 to 15 minutes.
8. using a pipette or a graduated cylinder, check floatability: non decapsulated cysts will float and decapsulated cysts will sink; as soon as all cysts have turned orange, stop the process by harvesting them on a sieve and rinse thoroughly with plenty of tap water and rinse well until no more chlorine smell is noticed;
9. the residual hypochlorite adsorbed by the decapsulated cysts has to be neutralised by dipping them in a 0.1% solution of sodium thiosulfate (Na2S2O3. 5H2O) for 5 minutes;
10. then, after a final rinsing, they are transferred to the incubation tank.
Decapsulation is an exothermic reaction: during the process the temperature will raise by approximately 12°C, from 20 to about 32°C.
The chemicals used for this process are toxic to humans and must be handled properly. Wear gloves and protective eyeglasses.
Check if the local legislation requires any special permission for (or bans) the use of any of these products.
Always check the chlorine content of the bleach used, to optimize the entire procedure and give constant results. See Annex 7 for detailed description of procedure and reagents
Fig. 36.01-2-3-4 Artemia industrial decapsulation (photo INVE Aquaculture)
Besides incubation, the decapsulated cysts can be either stored or, unless not commonly, be fed directly to fish. For short-term storage, up to one week, keep in the refrigerator at a temperature of 0 - 4°C. Decapsulated cysts that have to be stored for a longer period have to be dehydrated in a saturated brine solution as follows:
As the water content of cysts should be about 16-20% after this process, maximum hatchability is guaranteed for a storage period of only a few months.
In an emergency, they can be directly offered to fish larvae, provided that water circulation and aeration can keep them in suspension for a sufficient time to be ingested by fish. As the decapsulated cysts are 50% smaller than newly hatched nauplii, they may be used as initial food for small fish larvae.
As soon as the dry cysts are placed in water to be hydrated, the embryos inside the cyst start their metabolic activity. The free swimming nauplius, called instar I, hatches after about 20 to 24 hours if incubated under optimal conditions as follows:
- tank design: round with conical bottom, white gel-coated inside with a semi-transparent window near the cone tip for harvesting; a drain with a valve is installed in the cone tip;
- filtered and sterilized seawater, 35 ppt salinity;
- water temperature 28-30°C;
- strong aeration to provide a vigorous water agitation, to keep cysts in suspension: air is provided through the open end of a 1/2 inch PVC pipe placed close to the tank bottom;
- dissolved oxygen level above 4 ppm;
- pH over 8; if needed, add sodium bicarbonate (NaHCO3) at the rate of about one gram (previously dissolved) per litre;
- a strong illumination of 2 000 lux at the water surface during the first incubation hours; light can be provided by two neon m3 (photo STM Aquatrade) tubes (2 x 58 W) placed just above the tank rim;
-cyst density for incubation: 2.5 g per litre
Fig. 37.01 Artemia incubation tanks of 1 m3 (photo STM Aquatrade)
Artemia should be harvested when at the energy-rich instar I larval stage, just after hatching. This occurs in about 22 h at 28°C. To assess the proper time, sample the incubation medium with a 5-ml glass pipette and check for nauplii and umbrella stages (embrios still attached to their cyst) under the stereomicroscope.
To harvest nauplii, proceed in the following way:
1 Purge the bottom tip of unhatched cysts that has sunk by opening the drain valve for a few seconds;
2 Fit a flexible hose at the drain valve and place its open end into a container full of seawater equipped with a central filter.
3 Stop aeration, switch off the light above the tank, cover its top with a light-proof lid and place a light source at the window on the tank bottom: empty cyst shells will float, unhatched cysts will sink, whereas freshly hatched nauplii will concentrate at the tip of the conical bottom, attracted by light because of their positive phototactism. To attract the nauplii use a 150 W bulb lamp placed in a waterproof holder;
4 Wait 10 minutes, the time the nauplii will separate from their empty shells and start to harvest the tank, either fully or partially, by draining it into the filter. After another 10 minutes, a second harvest may be carried out if all nauplii were not harvested earlier. Do not drain the tank completely to keep floating empty shells in it. Adjust the water flow to avoid clogging the screen/water overflowing. Suggested maximum water flow is 100 l/min. If the water level in the filter stands higher than the water level in the tank, it would also be possible to harvest nauplii using a small submersible pump. When filtering, check with a glass beaker for possible losses of nauplii that may escape the filter.
5 During the settling time monitor the oxygen level, which should not drop below 2 ppm, as well as excessive crowding nauplii above the tip of the conical bottom. If necessary prior to harvest inject pure oxygen into the culture tank for 10 to 15 minutes to raise the dissolved oxygen content up to 10 ppm.
6 Rinse the nauplii thoroughly (for about 15 minutes), preferably with fresh water, to wash out the hatching debris.
7 Put the rinsed nauplii in a temporary container filled with a known volume of filtered and sterilised seawater and count them (see the procedure below).
8 Transfer nauplii to the storage tank filled with sterilised sea water and adjust the water volume to give a maximum density of 4 million nauplii per litre; provide a moderate aeration from the cone tip and add blue ice (or sealed ice bags ice) to keep water temperature at 5 to 10°C or better use a cold storage refrigerated tank or a freezer when available. At higher temperatures their nutritional value decreases rapidly.
9 If the nauplii have to be enriched, transfer them to a previously prepared enrichment tank.
Fig. 38.01-2-3-4 Artemia harvesting (photo STM Aquatrade)
To assess the hatching results and to feed the larval rearing tanks at the established densities you have to count the Artemia nauplii.
Three methods are described below, one for high nauplii densities, such as after harvesting and in a cold storage tank, one more common for counting the nauplii when they are in the incubation tank and the other for low nauplii densities, similar to those which can be found in fish tanks.
Counting high density nauplii samples:
Counting nauplii samples from the incubation tank:
Counting low densities nauplii samples:
The two main criteria to evaluate hatching results are:
Older larval stages of brine shrimp, the metanauplii, are used typically as feed for growing fish post-larvae. However, their poor nutritional value has to be boosted by dedicated enrichment diets that are rich in essential n-3 HUFA. Such diets can only be given when Artemia feeds actively, which is during its larval stages of instar II and instar III. First feeding in brine shrimps actually coincides with their moult into the second Instar stage.
The best results are obtained when the hatching time and the moulting pace are exactly known. The beginning of the enrichment can easily be determined by observation. After the first 18 hours small samples are taken every hour and checked under the microscope. The appearance of second instar stage is easily detected since it is larger than the first instar and presents a gastrointestinal tract. Enrichment will begin as soon as the first instar II stages appear.
The duration of the enrichment process as well as the type of product to be used depend on the HUFA content that is desired. Full enrichment takes 24 hours and two doses of enrichment emulsion, at time zero and 12 h later. Short-term enrichment takes 12 h and only the initial dose.
Fig. 39.01 Enriched Artemia (photo INVE Aquaculture)
Optimal enrichment conditions
Prepare the enrichment meal as specified by the producer, and make sure to prepare a new enrichment emulsion for each meal. At the end of the enrichment time harvest the metanauplii as usual, rinsing them thoroughly with seawater until no oily emulsion is noticed in the outflowing water.
Fig.40.01-2 Artemia cold storage tank (photo STM Aquatrade)
Cold storage of enriched Artemia nauplii
Enriched metanauplii rapidly loose their nutritional value at room temperature, as it happens to rotifers, unless they are stored in cold seawater (4 to 10°C). Keep density below 4 millions per litre. Equipment and procedure are the same described for cold storage of nauplii.
Annex 11 shows examples of daily procedures and recording files for Artemia sector.