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Commercial hatchery production of blacklip and Mabe (giant black-winged) pearl oysters and following nursery culture to maturity has been successfully carried out for more than two decades in Japan where the stock of the mother-of-pearl shells are very limited (Shokita 1990). However, technical information has remained within property of private. At present, existing pearling industry in Oceania is mainly based on the collecting wild stock of mother-of-pearl shells or spat, and the declining wild stock and fluctuation of natural spat fall are major concerns. Recent hatchery development for the blacklip pearl oyster in Oceania region (Ito et al., 1995; Ito, 1996) is a promising outlook for establishing new cultured pearl industry by rebuilding depleted stocks or sustaining stocks without heavy pressure on natural stocks.

Hatchery technique for producing Mabe (the giant black-winged pearl oyster), Pteria penguin, has also been exercised for almost three decades by the Japanese companies, following the several years of research by the prefectural government organizations in late 1950s (Shinmura and Toyoda, 1956 & 1957; Shinmura et al., 1959; Matsui and Yamaguchi, 1958) and 1960s (Kozuka et al., 1961; Yamaguchi and Kozuka, 1961). Based on field observations, spawning season in the southern Japan is in summer and autumn, from early June to late October, where ambient water temperature ranges 25 – 29°C and particularly, gonad is most matured in August. In Vava'u, the Kingdom of Tonga, spawning season ranges from November to February, with gonad conditions being acceptable for hatchery operation between September and April

The size at which they reach to the settlement stage ranges from 240 to 280μ (average 250μ ) in shell length. Minimum water temperature for survival seems to be around 16°C. Optimum water temperature is around 25°C and physiological function deteriorates markedly at 23°C or below. They prefer to inhabit in the channel with strong current, being abundant around the point areas, and they are found most abundantly at the depth of 5–60 m with gravel, sand or muddy bottom substrates. They tend to aggregate on rocky reefs, sunken trees or branch corals.

Difficulties remain in Oceania for a long term training of the hatchery technicians, half and round-pearl operators and marketing personnel. In Japan, the training at pearl farms including hatchery is a sort of traditional Japanese style (“shokunin katagi”) and normally done by verbal teaching and by occasional technical demonstrations, somewhat based on individual trainer's preference since the trainee is expected to be his successor. While a prospective trainee is working as a farm hand, he is given a choice to become a pearl operator and/or hatchery technician after signing agreement specifically prepared for him to protect company's (and trainer's) profit and secret.


2.1 Conditioning

If the spawners are kept at a pearl farm, they should be separated from other oysters. It is best to hang in pocket net or cage in deeper water to avoid unnecessary stress that stimulates untimely spawning. Select best quality spawners among the farm held oysters, looking healthy (e.g. firm mantle without dropping), without fouling organisms such as boring worms, having good nacre and colour of pearl layer, and with ripening gonads. It is good idea to select and separate about 100 spawners from other oysters for hatchery purpose. If they are kept in the conditioning tanks on land with a flow-through system, it is necessary to provide good water flow of at least 400–600% per day of water exchange rate for 1,000 l tank with 20 spawners and abundant food (mixed micro-algae species) of about 400–600 1 for the 1,000 l conditioning tank by dripping method.

2.2 Transportation

Polystyrene foam containers, either plastic reinforced (e.g. Esky box) or naked polystyrene foam box, have proven to be the best for a long distance transportation (up to 8 hours) of spawners either by air freight or by boat, because of its light weight and low fluctuation of inside temperature. Before transportation from the farm to hatchery, the spawners are pre-determined their sexes, level of maturation of gonad (spawning ripe, near ripe, poor and empty) by naked-eye inspection on site the farm. The gonad conditions are decided by the following criteria:

Spawning ripe:Gonad swollen towards the foot and bysal muscle, orange-yellow (female) and creamy white (ml) in colour
Near ripe:Gonad visible, sexes can be identified by a naked eye
Poor:Gonad not clearly visible, unable to identify sexes by a naked eye
Empty:Gonad empty, only muscle and gut colour visible

Only spawning ripe can be used for natural spawning and/or spawning induction program. However, spawners with near ripe and poor gonads can also be used for artificial maturation of ova and artificial activation of sperm followed by in vitro fertilization program.

When packing the spawners, they are wrapped with wet towel (with seawater) to maintain saturated humidity, which prevents the spawners from dehydration. Ice maker (blue gel/liquid) or ice block is included in the container to maintain inside temperature at around 22–25°C, which reduces metabolism of the spawner. 30–40 spawners can be packed in an 80l Esky for transportation from Vava'u to Nuku'alofa. On arrival, they are cleaned of any fouling organisms with a knife and scrubbing brush, then transferred to holding tank(s) or spawning tank(s). Spawners can be held in a raceway tank or round tank feeding raw seawater mixed with micro-algae (e.g. Tetraselmis spp. & Chaetoceros spp.) until spawning program. During spawning season, natural spawning often occurs immediately after the spawners are placed in the holding tank because of stresses from transportation, cleaning, drying and temperature shock. Therefore, preparation of equipment and tank setting for spawning program should be done for collecting fertilized eggs before the arrival of the spawners.


3.1 Preparation

Sea water needs to be filtered to 1 micron (preferably to 0.5micron); inline UV-sterilizer; air filters for clean air supply; thermometer; raceway tank (1,000 l) or round tanks (500 l) spawning; oyster rack to hold spawners upright position, shell opener and wooden wedges; scrubbing brushes; chisels/knives; gloves; 20–30 l plastic buckets/tanks for separating individual spawners; 20–25μ and 80–100μ mesh sieves for collecting and cleaning fertilized eggs; 10–20 plastic bucket for counting eggs; a small plunger; micropipette (0.2–5ml), rafter counting chamber, hand counter, Pasteur pipette; 500 l tanks for incubating eggs; glass tubing, air tubing and silicone tubing; binocular microscope.

3.2 Procedure

Figure 1 shows diagram of spawning induction procedure. When using one 500 l round tank, select 20 spawners (10 females & males each) with spawning ripe gonads from the holding tank(s) and commence shell cleaning with using chisel and brush (removing fouling organisms and dirt as much as possible). The spawners should be kept out of the water before the cleaning process. After cleaning, rinse the spawners first with freshwater (rainwater) and then with filtered seawater and place them in the spawning tank(s). This shell cleaning process may take an hour. If they start opening (gaping), occasionally spray clean seawater to avoid drying out their tissues. Observe the spawners in the spawning tank(s) carefully and patiently. Male spawning normally precedes females and, if many males respond and release sperm in a short period (5–10 min.), remove spawning males from the tank before it becomes too cloudy to recognise female spawning. Take water sample to check whether the sperms are viable (active). Spawning females should be isolated from the other oysters and be kept individually in the 20–30 l buckets or tanks with filtered seawater. These females usually continue to release eggs after being removed and kept in the buckets/tanks where the water temperature is similar to the induction tanks. The fertilization occurs almost instantly in the spawning tanks, up to 10–15 minutes after the female spawning. About 15 minutes after fertilization, the first polar body will be seen under the microscope. When the female ceased releasing eggs, remove the oyster from the bucket/tank and return to the broodstock holding tank. It is good idea to record the spawned individuals (e.g. sex, size, tag #, date of spawning) for the next spawning program or broodstock conditioning. Take the water from the ml buckets/tanks, which contains full of viable sperms, and add to the buckets/tanks with the eggs, only if the fertilization seems to be poor. After confirming the fertilization, sieve the eggs with 80–100μ for removing large debris and collect the eggs with 20–25μ mesh screen. Rinse the eggs with filtered seawater to wash out sperm. Transfer the eggs to 10–20 l bucket for estimating the number of released eggs. Sampling eggs, counting eggs and computing eggs are as follows: take 2ml sample while gently plunging bucket with eggs, prepare 4–8 samples for counting, use Pasteur pipette and Rafter counting chamber; The estimated number of eggs in 10l bucket will be (X) × 10,000 × ½, where (X) = (X1+X2+ X3+…Xi) × 1/i

When the induction is carried outside the high spawning season, temperature shock treatment following the cleaning procedure should be required. After cleaning the shells, water temperature of the holding tanks are to be lowered to 21–22°c and kept for about 2–3 hours. Occasionally spray filtered seawater to avoid drying out their tissues if they are kept out of water in dry tanks in a cool room. Return the spawners in an induction tank with water temperature at about 31–32°C. Male spawning may commences within 15–30 minutes and female spawning follows the male spawning. If no ml responds, return all spawners to the cool room or cold water tanks. Repeat the above process until they spawn. If there is no response, return the spawners in the holding tanks and prepare for fresh batch of spawners for another induction trial. Feed spawners with excess amount of micro-algae while they are held in conditioning (conditioning) tanks with flow-through system.

Figure 1.

Figure 1. Induction Procedure

3.3 Summary of Induction Procedure

  1. Shell Cleaning & Sexing: select spawning ripe or best gonads. 10 females & 10 males
  2. Rinse with freshwater & filtered seawater
  3. Keep them in a cool room or cool tank (at 21~22°C) for 2~3 hrs
  4. Prepare temperature shock treatment if necessary; filtered seawater in 500~1,000 l tanks; raise water temperature at around 31~32°C
  5. Place male and female spawners; 10~20 spawners/500 l; no air & static water
  6. Wait for about 15~30 minutes for male spawning and females. If not spawned, return the spawners and feed them at ambient water temperature in holding tanks.

3.4 Artificial Maturation and in vitro Fertilisation

When the hatchery operation is conducted outside spawning season, it is difficult to obtain spawners with spawning-ripe or good gonads for induction program, then, artificial maturation and activation of gametes (ova and sperm) method, or stripping method, can be performed. Ova and sperm of the sacrificed oysters are collected separately in a beaker/flask and the former can be matured artificially in Ammonia solution and the latter activated instantaneously activated in the same Ammonia solution. Thus, the fertilization occurs in vitro. It is advised to select the ripest males and females for successful result. The following section shows how to prepare 0.5% seawater solution of 0.1 Normal NH4OH for this purpose.

3.4.1 Making Ammonia solution

Obtain full strength/concentrated Ammonia solution (Ammonium Hydroxide 25% w/v as NH3), which contains 25% by weight of ammonia and is 12.5 Normal, and should be diluted accordingly. If “880 ammonia” (NH4OH 28% w/v as NH3) is used, a different dilution should be done because “880 ammonia” is 14 Normal.

To make 0.1N - NH4OH (125ml original solution), take 1 ml of 12.5N - NH4OH concentrated solution and add 124 ml distilled H2O.

1/125=0.1/12.5 ----- 0.1N - NH4OH solution (125ml).

To make final concentration of 0.5% 0.1-N NH4OH (1000ml working solution), take 5 ml of 0.1N-NH4OH seawater solution and add 995 ml of filtered seawater.

0.5% = 0.5/100 ------ 0.005 × 0.1 N = 5/1,000-N (or 0.5×10-3-N) NH4OH seawater solution (1,000ml)

3.4.2 Artificial maturation of ova

  1. Strip female gonad with several incisions, but try not to gut the intestine and other organs.

  2. Collect eggs with 20–25μ mesh screen (use larger mesh such as 80μ to remove tissues and other debris.

  3. Introduce eggs into 0.5×10-3-N NH4OH solution.

  4. (take eggs samples and observe every 5~10 minutes)

  5. Wait for about 50~60 minutes

  6. Introduce sperm for fertilization

Stir well while waiting for maturation of the ova. Check the ova under microscope every 5–10 minutes. When the germinal vesicle becomes no longer visible or is difficult to recognize and the ova are round and, they are ready for fertilization.

3.4.3 Artificial activation of sperm and in vitro fertilization

  1. Strip male gonad with several incisions, but try not to gut the intestine and other organs.

  2. Collect sperm in a beaker/flask with 20–25μ mesh screen to remove tissues.

  3. Introduce 1–2ml of sperm into the container (0.5×10-3-N NH4OH solution with matured ova).

  4. Wait for 1~2 minutes.

  5. Wash the eggs thoroughly with filtered seawater (use 20–25μ screen for washing sperm off) and re-suspend the eggs in a 10–20l plastic bucket/container for counting eggs. Take extra samples to determine fertilization rate.

  6. Stock the eggs in 500–1,000 l incubator for hatching.

  7. The 1st polar body can be seen in about 15–20 minutes after fertilization, followed by the 2nd polar body (~30min.), 1st cleavage with 1st polar lobe formation, 2-cell stage, 2nd cleavage with 2nd polar lobe formation, 4-cell stage, division of D-cell (5-cell stage), 8-cell stage, Morula stage, Blastula stage (3~4hrs.), Gastrula stage (5~6hrs), Trochophore (8hrs.) and D-veliger stage (18~20hrs.).


4.1 Monospecific Micro-algae Culture

Pearl oyster hatchery requires monospecific micro-algae culture to provide food for the larvae and spat. Failure of monospecific micro-algae is mainly caused by human errors such as a lack of hygiene, careless handling and inappropriate equipment. A sense of hygiene is the most important for successful operation. The culture process is shown in flow-chart of micro-algae cultures (Fig. 2). Master stock culture should be obtained from a well-known organization which keeps micro-algae species for aquaculture purpose under axenic or non-axenic conditions (e.g.CSIRO Marine Laboratories, Hobart, Australia).

Figure 2.

Figure 2. Micro-algae culture process

4.2 Preparation & precaution

There is a basic rule of preparation of equipment and precaution for commencing micro-algae culture in aseptic condition. Strict practice of hygiene must be considered the highest priority for successful work performance.

  1. Soak in freshwater and wash with detergent, brushing off dirt/wastes. If necessary, use hydrochloric Acid (5–10% solution) for cleaning the bottom of flasks by soaking. Collect the used hydrochloric acid in a glass bottle for re-use.
  2. Rinse with freshwater at least five repeats, completely wash off residue.
  3. Dry flasks upside down and avoid air-born dirt inside the flask
  4. Spray ethanol (70–80% solution), rinse with distilled water or rainwater filtered through 0.5~1μ filter, and wait for dry upside down.
  5. Put the lid on (aluminium foil) or place them in a dust-free cabinet for longer storage.
  6. Rinse with filtered seawater (UV-sterilized 0.2~0.5μ filtered seawater) before use
  7. Do not touch clean flasks & other equipment with dirty fingers
  8. Make sure that you wash your hand, particularly dirty fingernails and oily fingers, with soap and rinse off any chemical residues, and then spray ethanol before you commence work.
  9. Spray ethanol on the surface of culture containers /flasks/bench when entering the room.
  10. Keep the floor and bench clean and dry, if necessary, clean the floor with chlorinated freshwater
  11. Soak your feet in the chlorine bath before stepping into the room.
  12. Periodically check and clean air filter/air outlet of air-conditioner.
  13. Always keep the room door/windows closed, avoid unnecessary entry into the room.

4.3 Stock Culture (axenic non-aerated culture

100ml, 250ml or 500ml flasks are normally used for stock cultures and the culture medium is often made with artificial seawater. The flasks and nutrient media are to be autoclaved separately (for 10–25 min. at 105–121°C). For simplifying the procedure, however, they can also be sterilized in an autoclave altogether as one unit. If an autoclave or pressure cooker is not available, a gas/electric stove can be an alternative for sterilization of the media; the idea is the same as steam sterilization of milk bottles for babies (see Fig.2), where a flask with nutrient medium can be kept in the hot water (70–80°C) for about 15 – 30 minutes. The nutrient medium in the flask should not be boiled as this will alter the nutritional value of the medium. The purpose of this procedure is to minimize the risk of contamination by potential pathogens and other algal species in the flask before commencing the monospecific cultures.

Subculture of stock culture is the most important procedure to keep algal species alive for a long period and, whenever it is required, it must be ready to commence starter (high-density) culture. Bacterial contamination often occurs during this stock culture procedure, causing heavy bacterial contamination in subsequent starter and mass cultures and ruining the whole hatchery operations. Therefore, the inoculation is usually conducted in a laminar-flow chamber equipped with an air filter (0.2~1μ) and UV lighting to minimize contamination by air-born pollutant. If this equipment is not available, inoculation and subculture can still be conducted in a simple enclosed chamber with an open ceiling or a partitioned working bench using a Bunsen (gas) burner or an alcohol burner. This alternative technique has been successfully used to produce clean algal food during a commercial silver-lipped pearl oyster hatchery operation in a remote tropical island of the Torres Strait, Australia (Ito, 1992 & 1998a). Still, meticulous care is always needed in performing subcultures, surface-to-surface contact should be avoided when attempting transfer of micro-algae aseptically from one flask to the other. Touching sterile surface of culture medium and equipment will lead to the bacterial contamination. The stock cultures may be kept for 1–2 weeks, depending on the culture temperature and the algal species under low lights about 50μEm-2S-1 or 500–1,000 Lux (about 1m from 1×40 Watt white fluorescent tube), and they are further inoculated (1–10ml) into new stock culture media. For preparation of the nutrient medium of the stock culture, (see tables 1–4). All stock cultures are under non-aerated static conditions. However, this requires a gentle shaking once or twice a day to keep cells in suspension. It is essential to prepare at least one duplicate stock culture to avoid shortage of algal stocks caused by an accidental loss.

Figure 3.

Figure 3. An alternative way of sterilization of flasks for stock culture.

4.4 Starter Culture (axenic aerated culture)

After 1–2 weeks of culture, the stock culture can be inoculated (100–300ml) into starter (high-density) culture, normally 21 – 31 in volume. The aim is to obtain a high-density culture (e.g. 8–15×106cells/ml of micro-algae) under sterile (axenic) aerated and higher irradiation; 100– 250μEm2S-1 or more than 4,000 Lux (in front of 2–40 Watt white fluorescent tubes). Subculture for making and extra starter culture and/or a mass culture is best preferred during the exponential phase, which may take 5–7 days after inoculation, because the young and viable algal cells are the ones most desirable for further cultures. The flasks with nutrient medium and all fittings should be sterilized (e.g. by autoclaving) before use.

4.5. Mass Culture (non-axenic aerated culture)

Once a starter culture reaches an exponential phase with high density (6–10 × 106 cells/ml), it can be either subcultured to begin another starter culture and/or used to inoculate (1–2 l) into a new 30–60 l mass culture. When the mass culture reaches greater than 2 × 106 cells/ml, it can be either harvested for feeding larvae, or subcultured for expanding into a large-scale mass culture (500 l or 1,000 l). Attention should be placed on minimizing contamination; however, it is inevitable that bacterial contamination will occur during mass culture. When foam appears on the surface and/or the algae clumps and the water becomes cloudy, these indicate heavy contamination and the culture should be discarded. All the culture tanks, lids and aeration tubings must be washed with chlorine, rinsed thoroughly with freshwater, and alcohol (75 – 85%) is sprayed during a drying process. For using mass culture tanks of 500 l or larger capacity, freshwater is used for the final rinsing after washing with chlorine.

4.6 Selecting Micro-algae Species

The nutritional aspects of micro-algae in the mariculture of bivalve molluscs have been well documented (e.g. Brown et al. 1989). The culture of micro-algae as a food for the larvae and spat of pearl oysters is one of the most important aspects for hatchery operation. Although many algal species have been used as food, they are not equally successful in supporting the growth of particular animals. The reasons for this are related to differences in the size, digestibility and particularly nutritional value of algae. The nutritional value depends primarily on the biochemical composition of the algae and the specific nutritional requirements of the feeding animal. The following four live micro-algae species are commonly used as standard food for the pearl oyster larval and spat rearing.

4.6.1 T.ISO (= Tahitian clone of Isocrysis sp.)

T.ISO is one of the best known micro-algae species for aquaculture in the subtropical and tropical conditions. This species shows fast growth (3–5 × 106cells/ml after 3–5 days of inoculation from starter culture) and survives for a long period while maintaining high density during the stationary phase (6–9 × 106 cells/ml for 7–10 days) under a relatively low light intensity, indicating that this species is tolerant of the bacterial contamination and has been regarded as a suitable food for a long term larval rearing. Nutritionally good but lacking one of the 3-HUFA (High Unsaturated Fatty Acids), particularly lacks 22:6 3 known as DHA. This suggests that it may be wise to avoid using this species as a single food source during larval rearing. Combinations of T.ISO and Chaetoceros spp. or Pavlova spp. have been used for pearl oyster hatchery operations in Australia.

4.6.2 Pavlova species

Pavlova salina has recently been introduced to subtropical and tropical hatchery operations. This species is quite similar to P. lutheri. P.lutheris is the cold water species with excellent nutritional composition. P.salina, originally isolated from the Sargasso Sea, is a thermal tolerant, tropical species of Pavlova, considered to be a good food source for tropical pearl oyster hatchery operations (Jeffrey and Garland, 1989). However, extra efforts are required to obtain a good culture because the initial growth (log phase) is very slow (7-0 days to attain 3–6 × 106cells/ml) with a short peak period after reaching stationary phase (3–4 days to attain 6–8 × 106 cells/ml), and because this species is much less tolerant of the bacterial contamination compared to T.ISO. Providing good light sources, minimizing bacterial contamination and appropriate subculture timing are essential. It is well known that the dead cells of P.lutheri release a toxin that contributes to larval mortality in a poor larval rearing management.

4.6.3 Chaetoceros species

Chaetoceros muelleri (formerly called as C.gracilis) and C.calcitrans have good nutritional value (Volkman et al., 1989) and have been widely used for pearl oyster hatcheries. Although they are not thermally tolerant, they grow without adverse affect at around 25–28°C. If they are suited as food sources with a combination of P.salina or T.ISO in tropical hatchery conditions. Where a good airconditioning facility is available (with room temperature ranging from 20–23°C), C.muelleri and C.calcitrans are recommended for use together with P.lutheri for larval and spat rearing.

4.7 Preparation of Micro-algae Culture Media

Guillard's f/2 medium (Guillard and Ryther, 1962) and Modified-f medium are the most commonly used for microalgae cultures in Oceania. However, these nutrient media involve a rather complicated preparation technique using expensive chemicals, particularly in the preparation of trace metal solutions. A ready-to-use trace metal mix powder (Clewat-32™), which can be stored at room temperature for a long period, is used for the micro-algae cultures (indoor) to reduce the amount of time and to simplify preparation and handling (see Tables 1–3).

For large-scale outdoor cultures (500 l and larger volume), a powdered nutrient mix (e.g. Aquasol™ or Phostrogen™ gardening fertiliser) can be used (Table 4). In addition, the nutrient medium working solutions should be made up in simplified ways to minimize handling mistakes by hatchery staff during the operations. For example, the Aquasol at 32g/l solution with phosphate-strengthened nutrient medium (Modified-MI medium) has been used for the outdoor mass culture of T.ISO, P.salina, Chaetoceros spp. and Tetraselmis spp. in commercial-scale hatchery operations (see Table 4 for application rates).

Table 1. Culture medium.

Micro-algae nutrient mediaConcentration (mg/l)
 Stock/Starter CultureMass Culture
Vitamin B10.1 (100μg)0.05-0.01
Vitamin B120.0002 (0.2μg)0.0001–0.00002
D-biotin0.001 (1μg)0.0005-0.0001

* Clewat-32™ is the powdered trace metal mix and the commercial product of Teikoku Chemical Co. Ltd., Japan.
The trace metal contents (per 1kg) of Clewat-32 is as follows:

3.8g of FeCl36H2Oas Fe
7.7g of MnCl2-4H2Oas Mn
1.6g of ZnCl2as Zn
0.07g of CuSO4-5H2Oas Cu
6.3g of (NH4)6Mo7as Mo
24.7g of H3BO3as B
0.23g of CoCl2-6H2Oas Co

** Sodium Metasilicate (Na2SiO39H2O) is required for diatom culture (e.g. Chaetoceros spp.)

Table 2. Preparation of the working solutions of culture media.

Solutions of culture mediaStock Culture (ml)Starter Culture (l)Mass Culture (l)
 200ml medium2.5 l medium30 l medium
Solution A (ml)0.22.515
Solution B (ml)1.012.515
Solution C (ml)0.22.515
Solution D (ml)0.22.515

1.Solution A (Main Nutrients):NaNO3 100g
Na2HPO4-12H2O 14g
NaHCO3 12.6g
Na2-EDTA 18.1g
Add distilled water to make up 1 litre solution.Store in the fridge.
2.Solution B (Trace Metals): Clewat-32100g 
Add distilled water to make up 1 litre solution.Store in the fridge.
3.Solution C (Vitamin Mix): Vitamin B1100mg 
(take 10ml from B1 Original Solution*)  
Vitamin B120.2mg 
 (take 0.2ml from B12 original solution*)
 (take 1ml from D-biotin original solution*)
Add distilled water to make up 1 litre solution.Store in the fridge.
4.Solution D (Silicate for Diatom)Na2SiO3-9H2O15mg 
Add distilled water to make up 1 litre solution.  
* Vitamin Original solution: Vitamin B11g
(Always store in the fridge)(Add distilled water to make up 100ml solution)
  Vitamin B120.1g 
(Add distilled water to make up 100mL solution)  
(Add distilled water to make up 100ml solution) 

Table 3. Culture medium for mass culture

Micro-algae nutrient culture mediaCultureVolume (I)
Vitamin Mix10ml20ml50ml100ml

* Na2SiO3-9H2O is only required for diatom culture.

Table 4. Modified culture medium for outdoor mass culture of micro-algae.

Micro-algae nutrient culture mediaCultureVolume (I)
Aquasol (32g/l sol.)60ml50ml100ml250ml500ml
Na2HPO4-12H2O(3.5g/l sol.)30ml50ml100ml250ml500ml
Na2SiO3-9H2O (15g/l sol.)*30ml50ml100ml250ml500ml

* Na2SiO3-9H2O is only required for diatom culture.
** For up to 60 litre culture volumes, Aquasol solution volume added is 1ml/l. For greater than 60 litre culture volumes, this is reduced to 0.5ml/l (=1/2 strength).
Nutrient composition (W/V) of the Aquasol is as below (see the container label for more details):

N asNH4-PO4 ----------- 1.8% P asNH4-PO4 ----------- 4% (total water soluble)
N as       KNO3 ----------- 2.6%K as     KNO3 ----------- 7.8%
N as       (NH2)CO ------- 18.6%K as     KCl ----------- 10.2%
TotalN ---------------------- 23%Total     K ----------- 18%

Trace metals (W/V)

Zn ----------- 0.05%Cu ----------- 0.06%
Mo ----------- 0.0013% Mn ----------- 0.15%
Fe ----------- 0.06%B ----------- 0.011%

4.8 Counting Micro-algae

Using a Pasteur pipette, collect a sample from the vial to be placed on the counting chamber (Haemocytometer). Centre the coverslip over the counting chamber (Fig. 4). Place the tip of the pipette in one of the two grooves and release the sample from the pipette into the groove. It is important that no air bubbles form otherwise the estimated count is incorrect. Keep releasing the sample until both grooves have been filled and there are no air bubbles. Also, be sure that there is not too much of the sample between the coverslip and the counting chamber. The coverslip should not be able to slide freely.

Figure 4.

Figure 4. Counting Chamber (Haemocytometer)

The counting chamber (Haemocytometer) consists of two grids on which the microalgae are to be counted. Use low magnification (x40) on the microscope to first find one of the grids. Then, use the high power (x100) to focus on the centre of the grid that has the highest number of boxes - 25 boxes (each with 0.2mm × 0.2mm) consisting of 16 small boxes each (0.05mm × 0.05mm). Using a counter, count how many micro-algae are within the centre grid; large area-(1) (1mm×1mm), consisting of 400 small boxes as shown in Fig. 5. The gap between the coverslip and the counting chamber is 0.1mm (0.01cm).

Figure 5.

Figure 5. The grid of the counting chamber

Thus, the volume of this space (large area -(1)) is; 1mm×1mm×0.1mm=0.1mm3= 0.0001cm3=10-3ml. If a total of α cells are counted in the space over this area-(1), the estimated number of cells per ml in the sample is: α×104. In order to obtain the algal density in the culture, get average counts from at least 4 replicates of grids (samples): (α1+α2+α3+ α4) cells per ml.

It is best to count in a pattern going up and down or across and back. If the microalgae are touching the line of the box, it is to be included in a count. If a size of a cell body is very small such as green-algae(e.g. Nannochloropsis spp.) and it is only touching the outside of the borderline, do not include in a count (see Fig. 6).

Figure 6.

Figure 6. Determining micro-algae cell counts

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