HOW TO RUN A PRODUCTION OF LIVE FEEDS (E-book)

Introduction

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:

all-female (amictic) populations of the rotifer Brachionus plicatilis (60 - 350 mm in length);
larval stages (nauplii and metanauplii) of a small crustacean, the brine shrimp Artemia spp. (length: 400 - 800mm).

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:

adequate dimensioning of the production facilities, including additional space for back-up cultures as a precaution against culture collapses;
proper daily renewal and up-scaling of the cultures (standard operating procedures);
maintenance of strict hygienic conditions in the culture environment (cleaning procedures);
close control of culture conditions (monitoring procedures).

Mass culture of microalgae

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)

Population dynamics

Microalgae population dynamics can be described by different phases:

the lag-phase, where, just after the inoculum, the cells increase in size, but not in number, and begin to absorb the nutriens supplied with the culture medium;
the log-phase (or esponential phase), where cells reproduce very fast and population growth is exponential;
the transitional phase (or declining growth phase), where growth rate slows down;
the stationary phase, where cells remains constant in number and reproduction is balanced by death;
the decline phase, where cell number decreases since death rate exceeds growth.

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.

Mass production systems

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)

Mass culture facilities for microalgae

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).

Preparation of the culture medium

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.

Mechanical filtration

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)

Sterilization

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.

Enrichment

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.

Primary stock solutions

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 B₁₂solution should preferably be stored in a dark or aluminium wrapped bottle.
Warning: do not sterilize any vitamin solution.

Working solutions

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

Culture equipment sterilization

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.

Enrichment of culture vessels

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.

Batch culture of microalgae

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)

Pure strain culture

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.

Protocol for test tubes replication

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.

Protocol for purification of algal strains

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.
Dilution method
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.
Example:

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.

Capillary method:
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.

Upscaling culture conditions

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:

temperature	20 ± 2°C
salinity	25 to 30 ppt
light intensity	4000 to 8 000 lux
aeration	50 to 100% of the culture volume per minute
carbon dioxide	2 % of air volume

Fig. 25.01-2 Algae flask replication (photo STM Aquatrade)

Temperature
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.
Salinity
In the selected species salinity does not represent a limiting factor within a range of 15 to 40 ppt.
Light
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 hatchery’s 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
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.
Carbon dioxide
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.).

Scaling up protocol

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)

Monitoring algal populations

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:

to determine the growth curve for each species of algae under local conditions;
to devise criteria for quick identification of possible troubles (eg.; presence of foam, sedimentation pattern, changes in colour, etc.);
to determine optimal utilization time, i.e. the age at which the algal population reaches the peak of the log-phase;
to adjust environmental conditions to maximise production;
to control possible contaminants and try countermeasures.

Counting microalgae

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.

Mass culture of rotifers

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.)