2.3 TECHNIQUES – GROWING ALGAE
The following sections provide the protocols used at the BBSR hatchery for culture of algae from stock to 100 litres vessels used for harvest. In order to ensure a contaminant free culture, sterile microbiological techniques are used during sub-culturing or start up of solutions. These techniques rely on the flaming of all equipment prior to use, and the continuous working by the flame during addition of nutrients, transfer of cultures, etc. At the Bermuda hatchery, a propane bottle, fitted with a nozzle is used as a flame source. Prior to opening of any culture flasks or nutrient bottles, the torch is turned on, placed on the workbench, and left on throughout the inoculation process. In this way, equipment is continuously flamed and “sterilized” when in use.
2.3.1 Master cultures
All algal culture system require a set of “stock” or “master” cultures; these are usually of about 125–250 ml in volume, and provide the reservoir of algal cells from which to start the larger-scale cultures used for feeding. Several centres, which specialize in the culture of algae can provide inoculum for stock cultures. However, it is also possible to isolate algal species from a specific body of water, and attempt to rear them under controlled conditions for feed. The isolation of singles cells of a species from natural live phytoplankton samples can be done using a capillary pipette and/or via a series of dilutions; this allows the separation of a selected species into a culture chamber with nutrient media. Thereafter, algae are cultured as detailed below.
At the BBSR hatchery, master algal cultures are purchased or received from various laboratories. The Culture Centre for Marine Phytoplankton in Bigelow (CCMP) has an extensive list of algal cultures, which may be purchased. Information may be obtained online at the following address: www.ccmp.bigelow.org. Cultures are sent in 15 ml plastic test tubes and shipped by courier, to minimize transport time and ensure a large number of healthy cells upon arrival. The procedure for starting algal cultures and growing them to high density in 125 ml flask is described below. The following section describes the start-up and sub-culturing of 500 ml, 4-litre and 100-litre cultures.
Prior to reception of purchased cultures, 125 ml Erlenmeyer flasks are cleaned and prepared with adequate salinity seawater (see Protocol–5). The seawater used is classified as low nutrient seawater (LNSW), collected from the Sargasso Sea by oceanographers at BBSR. It is collected in 5 percent HCL cleaned 50 litres carboys from a depth of 5 m using Niskin bottles. At the laboratory, it is left to age for about 2–3 weeks to help strip out any inorganic nutrients, and then subsequently filtered through a sterile 0.2 µm mini capsule filter (Pall Corp., Item #12122). Following this procedure, no detectable nitrogen (<0.04 µm) or phosphorus (<0.03 µm) is found. Erlenmeyer flasks (125 ml) are autoclaved with seawater, for sterilizing of both flasks and seawater. Full strength salinity (36 ppt) is used for all cultures, except for the diatoms, Chaetoceros species and T. pseudonana, which fare best in reduced salinity seawater (25 ppt). Once the autoclaving process is completed, cooled flasks are ready for inoculation. LNSW is used because it is available; however, finely filtered (0.2 to 1 µm) and autoclaved seawater should be adequate.
PROTOCOL–5
PREPARATION OF CULTURE FLASKS (125 ml – 500 ml)
1. Wash glass flasks, Pasteur pipettes and rubber stoppers with glass rods in 10 percent HCl (hydrochloric acid) bath.
2. Rinse 3 times with fresh water and do a final rinse with Q-water (de-ionized) and let dry. 3. Fill flask with appropriate volume of filtered seawater using a graduated cylinder. 4. For reduced salinity, dilute seawater with Q-water. Use the following equation to calculate the volume of Q-water required to reduce the salinity:
Total volume (ml)
X (ml seawater) = Y x ––––––––––––––––––––
Full strength salinity
where:
X = volume of seawater added, and
Y = new salinity
Volume of Q-water required = Total volume (ml) - X
so that if full strength salinity = 36 ppt, and total volume in flask is 50 ml, and a new salinity of 25ppt was required
X = 35 ml of seawater, and
Volume of Q-water = 15 ml
5. Label flasks with reduced salinity.
6. Close flask loosely with cotton plug and aluminium foil. Cotton plugs are made with cheesecloth material tied around absorbent cotton.
7. Place flasks in autoclave and start cycle following manufacturer’s directions.
8. Wrap all pipettes, stoppers and 4 aeration rods and stoppers separately in aluminium foil. Label clearly on the foil the contents and put an arrow pointing to the fragile thinner end of the pipettes as an indicator. Make sure contents are completely wrapped in foil as they will not be sterile if there are any gaps.
The transfer of purchased stock cultures from 15 ml test tubes to 125 ml Erlenmeyer flasks is described in Protocol–6. Culture media used is F/2, and preparation for small culture volumes is outlined above (Protocol–5). Microbiological sterile techniques are used to transfer master algal stocks from 15 ml test tubes to 125 ml flasks. Depending on the density of each stock culture, one test tube is used to inoculate two flasks, yielding duplicates of each stock. Addition of media to the cultures is done using sterile microbiological techniques. For diatom species, an addition of autoclaved 3 percent sodium metasilicate is added for growth of the siliceous frustule. Flasks are closed with a cotton plug and wrapped with aluminium foil. They are maintained in the incubation chamber at T= 25 °C and on a 12-hour light cycle. Flasks are swirled once a day to prevent settlement and sticking of algae on the bottom; as well as to distribute the culture media throughout the solution. These are hereon referred to as master cultures. These master cultures must be sub-cultured frequently; some hatcheries do this weekly. At the BBSR hatchery sub-cultures are done monthly. Sub-culturing involves inoculating some cells from an old stock culture intro fresh culture medium. In this way cells can continue to grow and divide ensuring a healthy culture. If sub-culturing is not carried out, the algal cells in the stock culture will eventually die. It is important to take precautions to prevent contaminants from the air entering the stock cultures when sub-culturing. In this way, master cultures can be maintained indefinitely.
To start a new master culture, 10–20 ml of algae inoculum (depending on density) is taken from a master culture for inoculation of new flasks with new seawater and media. In the first instance, when cultures are inoculated with purchased stock, the lag phase is long, and it is found that an average of 1 month is required to achieve a density capable of inoculating a larger volume of seawater, as density of purchased stock is usually low. Protocol–6 describes the procedure utilized in the first inoculation of 125 ml master cultures with purchased stock, and the subsequent monthly sub-culturing for maintenance of master cultures.
PROTOCOL–6
INOCULATION OF 125 ml MASTER CULTURES
Preparation of flasks
1. Two 125 ml flasks for each start-up culture are cleaned as outlined in Protocol–5. 2. Flasks are filled with 50 ml of seawater with adequate salinity requirement.
3. Flasks and seawater are sterilized using autoclave procedures.
4. Culture media is prepared and sterilized as outlined in Appendix 9.
First inoculation of 125 ml flasks using purchased stock cultures
1. Upon receipt of stock cultures, open package and let cultures stand upright in incubation chamber, awaiting inoculation. Depending on state of received cultures, they can be left as received for 24 hours.
2. Prepare a work area, with a Bunsen burner close at hand, and F/2 solution mixed with vitamins and sodium metasilicate. Working under a hood is best.
3. Stock cultures usually come in 15 ml test tube with screw caps. Unscrew cap, keeping opening of test tube close to flame; discard cap. With other hand, hold 125 ml flask, remove cotton plug, keeping it in palm of hand.
4. Transfer stock culture to 125 ml flask without mouths of either container touching and remaining close to flame. Place cotton plug immediately back onto flask. Discard tube.
5. Label flask with algal species and date. This will allow you to maintain a tight schedule of re-inoculation.
6. Working close to flame, remove cap of culture media mixed with vitamins. Place cap on clean surface area and maintain media container close to flame. Using a sterile 1 ml pipette, remove 0.5 ml of nutrients from container. Flame mouth, flame cap and close, keeping pipette tip close to flame. Take 125 ml flask with inoculum, remove cotton plug and keep in palm of hand and add 0.05 ml of nutrients to algal inoculum (1 ml.l-1). Replace cotton plug immediately back onto flask. Discard pipette. Swirl flask to mix nutrients and algae.
7. Using same technique, add sodium metasilicate (2 ml.l-1) if species is a diatom. 8. Place inoculated flask with nutrients in incubation chamber. Swirl daily.
Maintenance of master cultures – monthly sub-cultures
1. Once a month, prepare the same number of 125 ml Erlenmeyer flasks as already inoculated.
2. Fill flasks with 75 ml of seawater adjusted to adequate salinity.
3. Autoclave and let cool.
4. Prepare a workbench as above.
5. Do all transfers using sterile microbiological techniques.
6. Using a sterile 10 ml graduated pipette and bulb, pipette 10 ml of culture, maintaining pipette close to the flame.
7. Pick up one flask (check salinity) with other hand, remove foil and cotton plug, placing it on clean area, and add inoculum close to flame.
8. Quickly close flask with cotton plug after flaming the mouth of the flask and the cotton plug. Be careful not to put cotton plug too close to flame, it will burn.
9. Repeat 2 or 3 times with same culture and same pipette, so as to add an inoculum of 20–30 ml into the new flask.
10. Label with species and date.
11. Add nutrients accordingly.
12. Swirl and store in incubation chamber.
13. Swirl on a daily basis until next inoculation.
Once master cultures are established, larger volumes of algae can be cultured using this reservoir of various algal species. There are many different ways of culturing algae. These can be divided as batch culture, semi-continuous culture and continuous cultures. Batch culture is the most traditional method used for large-scale culture in bivalve hatcheries. Large volumes of algae are grown and harvested fully once desired cell density is achieved. Each new batch is inoculated from working culture flasks. It is a simple method, and a variety of containers can be used, ranging from 20-litre carboys to 3 m diameter tanks. Semi-continuous cultures refer to a system where part of the culture is harvested and used as food, and the amount taken is replaced with fresh culture medium (clean seawater and nutrients). After allowing 2–3 days for the remaining cells to grow and divide, the process is repeated. Semi-continuous cultures may be operated for up to 7–8 weeks. These types of cultures can crash because of a build-up of contaminants, bacteria or mismanagement. Minimizing contamination from any source is critical in semi-continuous cultures. Various containers can be used for semi-continuous cultures. The most common type of container is a sterile polyethylene bag. The bags are sealed and the inside is sterile. They can be inflated with sterile air to form the shape required before filled with seawater. The bags need to be supported by a frame. At the BBSR hatchery, 100 l vessels, described in the facilities Section 2.1 are used for semi-continuous cultures.
Continuous cultures may be maintained by using turbidostat culture, and by chemostat culture. In the former, the number of cells in the culture is monitored, and as cells divide and grow, an automatic system keeps the culture density at a pre-set level by diluting the culture with fresh medium. In the latter, a flow of fresh medium is introduced into the culture at a steady pre-determined rate. The surplus culture overflows into a collecting container, from which it can be taken and used as food. These systems are not commonly used in commercial hatcheries because they are expensive, very sensitive, and difficult to install and maintain.
Depending on the requirement for algae, they can be cultured using closely controlled methods on the laboratory bench top, for a few litres of algae to less controlled methods in outdoor tanks relying on natural light conditions, and producing thousands of litres. At the BBSR hatchery, several steps are taken to ensure a daily harvest of algal food of optimal quality to larvae and post-larvae scallops. Algae are first cultured in batches of 500 ml flasks and 4 l flasks; these 4 l cultures are used in turn to inoculate 100 l vessels, reared in a semi-continuous method and used for daily harvest.
2.3.2 500 ml batch cultures
Once 125 ml cultures achieve higher densities of algae (6 000 cells.ml-1), inoculation to 500 ml flasks can be performed. These 500 ml cultures are in turn used as inoculum for 4 l cultures. Procedures for preparation, first inoculation, and maintenance of 500 ml flasks are outlined in Protocol–7. Flasks are filled with 250 ml of 1 µm filtered seawater (collected from the hatchery). Similar salinities as to those used for 125 ml flasks are used. Seawater and flasks are autoclaved. All procedures including transfer of inoculum, addition of media, or aeration pipette are conducted using sterile microbiological techniques. The inoculum of 35 ml is transferred using sterile microbiological techniques to the 500 ml flask. The exact amount is dependent on the master culture density. The denser the culture, the smaller the volume of inoculum required. The remaining volume in the master culture is used to re-inoculate a new 125 ml master culture flask (see Protocol–6). Culture media (F/2) and vitamins are added depending on volume of seawater inoculated; sodium metasilicate is added to diatom cultures. In 500 ml flasks, aeration is provided for mixing of culture media and algae, maintenance of algae in suspension and addition of CO2 for pH stability. For this reason, sterilized Pasteur pipettes are added prior to closure of flasks with cotton plug and aluminium cover. Flasks of 500 ml are connected to the airline in the algae container. In the first instance of inoculating a 500 ml from a master culture, approximately 2 weeks are required for a high-density culture, as lag phase is longer. At this time, 200 ml of the culture is transferred to a new 500 ml flask filled with 200 ml of sterile seawater. These are allowed to grow until cell density approaches 10 000 cells.µl-1 (1-2 weeks). Thereafter, once cultures are well established and dividing rapidly, 500 ml flasks are re-inoculated twice a week; this high frequency insures a healthy culture in continuous exponential phase. Maintenance of 500 ml flasks is achieved by transferring approximately 50–100 ml of algal culture from one 500 ml flask. The remaining 250–300 ml is used to inoculate a 4 l volume.
2.3.3 4 litres batch cultures
Algal cultures are reared to 4 l volumes for the purpose of obtaining a large inoculum required for the start-up of the semi-continuous 100 l cultures. Four litre Erlenmeyer flasks are too large to be sterilized in the existing autoclave at the Bermuda hatchery. For this reason, 1 µm filtered UV disinfected seawater is used. In the described facility, the seawater supply for the algae room needs to be set up, prior to inoculating 4 l flasks. Appendix 11 indicates the step by step procedure for obtaining 1 µm filtered UV disinfected seawater. Note: If UV disinfected seawater is not available, it is possible to sterilize seawater chemically, by using sodium hypochlorite (or commercially available
Chapter 2 – Algal cultures: facilities and techniques 53
chlorox), and neutralize it with sodium thiosulfate (1N solution). Appendix 12 provides the protocol for chemical sterilization. This was initially used at BBSR, and yielded satisfactory results.
Flasks are filled to 3.5 litres with UV disinfected seawater and closed with a rubber stopper (no. 10). One half of the 500 ml culture is used to inoculate this volume. Similarly culture media, vitamins and sodium metasilicate (for diatoms) is added to the new solution. For aeration of 4 l flasks, rubber stoppers are fitted with (diameter) glass rods. One rod equates the length of the flask, nearing the bottom used for bubbling, and the other is shorter, acting as a vent. Aluminium foil is used to cover the stoppers. Four litre flasks are connected to the airline in the algal container. Sub-inoculation of 500 ml flasks and 4 l flasks are conducted on the same day at the BBSR hatchery, twice a week (Monday and Thursday). In this way, healthy cultures in the exponential phase of growth are always available for inoculating larger 100 l cultures. Four litre cultures are allowed to grow for a period of 3–5 days before use as inoculum for 100 l cultures.
PROTOCOL–7
INOCULATION OF 500 ml FLASKS
Preparation of flasks
1. Two flasks for each algal species cultured are cleaned in a 10 percent HCl bath. They are rinsed three times with fresh water and have a final rinse with Q-water.
2. For a first inoculum from 125 ml master cultures, fill flasks to 200 ml. Use 1 ?m filtered seawater.
3. Close flask with cotton plug and foil, made as described in Protocol–6.
4. Sterilize flasks and seawater in autoclave.
5. For maintenance and sub-inoculation of 500 ml algal cultures, fill flasks to 350 ml using 1 ?m filtered seawater, adjusted to required salinity.
First inoculation of 500 ml flasks with 125 ml master cultures
1. When master cultures increase in density, inoculate 500 ml flask (filled with 200 ml of seawater) with 35 ml of master culture. Note: Remaining volume of master culture is used to re-inoculate a new 125 ml flask. See Protocol–6.
2. Use microbiological sterile techniques for transfer.
3. Prepare a work area, with a Bunsen burner close at hand, F/2 solution, vitamin solution and sodium metasilicate solution. Keep 5 and 10 ml sterile pipettes and 3-way pipette bulb nearby.
4. Using a 10 ml Pasteur pipette and bulb, transfer 30–35 ml of master culture to 500 ml flask, remaining close to flame at all times. Do not touch mouth of either flask with pipette. Plug flask quickly after transfer.
5. Label flask with algal species and date. This will allow you to maintain a tight schedule of re-inoculation.
6. Working close to flame, remove cap of culture media mixed with vitamins. Place cap on clean surface area, and maintain media container close to flame. Using a sterile 1 ml pipette, remove 0.5 ml of F/2 from container (nutrients added need to equate 1 ml of nutrient per litre of culture). Flame mouth, flame cap and close, keeping pipette tip close to flame. Take 500 ml flask with inoculum, remove cotton plug and keep in palm of hand and add nutrients to algal inoculum. Replace cotton plug immediately back onto flask. Discard pipette. Swirl flask to mix nutrients and algae
54 Installation and operation of a modular bivalve hatchery
7. Using same technique, add sodium metasilicate if species is a diatom at 2 ml of sodium metasilicate per litre of culture. In 500 ml flasks, 1 ml of sodium metasilicate is added. 8. Remove one Pasteur pipette, from autoclaved packet and keep close to flame. Take 500 ml flask in one hand, remove cotton plug and maintain in palm of hand, place Pasteur pipette in flask and plug.
9. Take flask to lightbank shelf and connect Pasteur pipette to airline. Regulate air bubble so as to have good mixing.
Maintenance of 500 ml cultures
1. 500 ml cultures initially inoculated with master cultures will take some time to reach required densities (2–4 weeks depending on strength of inoculum).
2. Once required densities are obtained, 500 ml cultures are inoculated twice a week, always maintaining cells in exponential phase of growth.
3. Prepare 500 ml flasks as above, but filling with 350 ml of seawater.
4. Transfer inoculum from 500 ml flask directly from flasks using microbiological techniques, working by the flame and avoiding for the mouths of the flask to touch, thus preventing contamination.
5. Label flasks with algal species and date.
6. Add nutrients using techniques described above, but with higher volumes For F/2 add 1 ml.l-1, and for sodium metasilicate add 2 ml.l-1.
7. Add Pasteur pipette for aeration using techniques described above.
8. Connect Pasteur pipette to aeration. Tetraselmis sp. has a tendency to stick to the bottom, if not well aerated, so more vigorous aeration is usually required for this species.
PROTOCOL–8
INOCULATION OF 4 LITRES FLASKS
Preparation of flasks
1. Clean two flasks for each algal species cultured using commercial grade bleach (5 percent chlorox). If needed, soak in seawater and bleach solution. Rinse well in fresh water.
2. Fill flasks at time of inoculation with 3 litres of 1 ?m UV disinfected seawater. If salinity needs to be adjusted, adjust using fresh water (see Protocol–5).
3. Wrap rubber stoppers fitted with glass rods, acting as aerating tubes, in foil and autoclave.
Inoculation of 4 l flasks with 500 ml cultures
1. When algal density in 500 ml flasks reach density >10 000 cells.ml-1, use 2/3 of culture to inoculate new 4 l flask. Note: Remaining volume of 500 ml culture is used to reinoculate a new 500 ml flask. See Protocol–7.
2. Use microbiological sterile techniques for transfer.
3. Prepare a work area, with a Bunsen burner close at hand, F/2 solution, vitamin solution and sodium metasilicate solution. Keep 5 and 10 ml sterile pipettes, and 3-way pipette bulb nearby.
4. Take 500 ml flask in one hand, remove cotton plug, keeping mouth of flask close to flame. With other hand, flame mouth of 4 l flask.
5. Transfer inoculum from 500 ml flask directly into 4 l flask; avoid the mouths of the flask to touch, thus preventing contamination.
6. After transfer, replace cotton plug quickly on 500 ml, even if discarded later. This ensures you to maintain a clean area, and prevent cross-contamination.
7. Flame stopper (used while cleaning of flask) and mouth of 4 l flask. Close flask quickly.
8. Label flasks with algal species and date.
9. Add nutrients, by pipetting required volume from stock solution, continuously ensuring that pipettes, and flasks are close to the flame. Quickly cap bottles, and transfer nutrients to 4 litres, flaming the mouth of the flask, before addition of nutrients and after. In between addition, keep flask closed with stopper. If you need to put stopper down, make sure it is put on a clean area. For F/2 add 1 ml.l-1, and for sodium metasilicate add 2 ml.l-1.
10. When nutrient addition is complete, unwrap rubber stopper with aerating rods, and flame. Replace plain rubber stopper with aerating stopper.
11. Connect aerating stopper to aeration tube on shelf by lightbank. Tetraselmis sp. has a tendency to stick to the bottom, if not well aerated, so more vigorous aeration is usually required for this species.
Remember:
Always use a different pipette for each algal culture species to avoid cross-contamination.
Use a different pipette for each nutrient solution (F/2, vitamin, sodium metasilicate).
Note: For ease of inoculation, it is best to inoculate one species at a time. Do 500 ml sub inoculation on same day as 4 l. For example: Start with T-Iso, sub-inoculate two new 500 ml and two 4 l. Add nutrients, aeration, connect to air supply. Clean bench, discard pipettes, and work with second algal species.
2.3.4 100 litres cultures: semi-continuous method
The volume of algae required in hatchery operations, mainly for larval, post-larval, and broodstock purposes, is harvested daily from 100 l vessels. The set-up at the BBSR hatchery yields approximately 120 l of algae a day.
Preparation and inoculation procedure for 100 l vessels are given in Protocol–9. Similarly to 4 l flasks, 100 l vessels are filled with 1 µm filtered UV disinfected seawater. These large-scale cultures are reared on a semi-continuous cycle. This yields a culture in a continuous exponential phase of growth, and minimizes labour. As a culture is harvested daily, and decreases in volume, new water and nutrients are added; this boosts the culture growth, such that algal densities reach 12 000 cells.ml-1 within three days. A first inoculum is given initially, and thereafter, addition of new water and nutrients done on a regular basis allows the culture to be maintained for at least one month or more depending on cleanliness of the culture and of the techniques.
Details of the procedure are given in Protocol–9. For a first inoculum, vessels are filled to 25 litres or 50 litres depending on the strength of the inoculum. The lower the strength, the smaller the volume of new seawater inoculated. A 4 l flask is used for inoculum. Culture media, vitamins and sodium metasilicate are added dependent on volume. Algal cultures are allowed to grow for a period of 5 days. At this time, addition of seawater is done, and vessels are filled. Cultures are allowed to grow for a period of 3–5 days; at which time, daily harvest of cultures for the hatchery complex is possible. One hundred litre vessels are harvested down to 25 litres in approximately 4–5 days. When 25 litres of culture remain, 75 litres of new 1 µm UV disinfected seawater is added with culture media. It usually takes approximately 3 days to reach harvest density. In this way, semi-continuous cultures can be maintained on the average 4–6 weeks depending on cleanliness of techniques.
PROTOCOL–9
INOCULATION AND SEMI-CONTINUOUS CULTURE OF 100 LITRES VESSELS
Preparation of vessels
1. Allocate two vessels to each algal species.
2. Clean vessels outside using a fresh water hose and commercial grade bleach. If needed, muriatic acid can be used. Care must be taken when using muriatic acid. Special attention is given to the bottom of the cone, valve area, and rim of the cone and other edges when cleaning these vessels. If needed, the bottom of the cone is soaked in seawater and chlorox solution for a few days.
3. Rinse well with fresh water.
4. Fill vessels with UV disinfected seawater prior to inoculation.
Inoculation of 100 l vessels with 4 l cultures
1. Connect 100 l vessel to airline and turn slight airflow on. It is important to do this, prior to addition of water or algae, as if there is no airflow, water will pass through the Tygon tubing and soak the bacteria filter. If this occurs, the bacteria filter needs to be replaced.
2. For a first inoculation, fill vessel to 25–50 litres, depending on the density of the inoculum. The denser the 4 l culture, the greater an initial volume of water can be used. 3. When density in the 4 l flasks reaches an algal count >10 000 cells.ml-1, use culture to inoculate 100 l vessel. Note: Take care not to use the very bottom of the culture, as it often contains some precipitate and detritus.
4. At this time, sterile microbiological techniques are no longer required. A 4 l culture is simply poured into a 100 l vessel. Also, diatoms appear to grow well in full salinity in these large volumes, and there does not seem any need to adjust the salinity.
5. Label vessel with date and algal species.
6. Add nutrients using 25 ml graduated cylinder or 10 ml pipettes. For 100 l vessels, F/2 solution is taken directly from purchased containers; equal parts of A and B are mixed, according to manufacturer’s instructions.
7. When nutrient addition is complete, place lid on top to avoid any detritus from falling and adjust aeration so as to create gentle mixing of nutrients throughout. Again, more vigorous mixing is needed for Tetraselmis species.
8. Allow for algal cultures to grow until dense. At this time, fill vessel with UV disinfected seawater. If volume of seawater added is 75 litres, add nutrient volume in accordance to 75 litres volume. For example: in Bermuda, 14 ml of nutrients in total is added to 100 l; during semi-continuous culture, if 75 ml of new seawater is added, only 10.5 ml of nutrients would be added.
9. Allow algal cultures to grow. When densities required are reached begin harvesting using valve at the bottom of the cone. Culture should be harvested within 6–7 days. Do not harvest below 25 l.
Semi-continuous culture method
1. Once culture volume has decreased to 25 litres, add 75 litres of new UV disinfected seawater and adequate nutrient volumes.
2. Label with date of added water.
3. Allow cultures to grow for 3 days before harvest.
4. Note: Cultures can be boosted at any time; if a culture of 50 litres volume needs to be reboosted, add 50 litres of seawater and according nutrients.
5. Although sterile techniques are not used for these large cultures, cleanliness is a must to avoid cross-contamination between vessels.
2.3.5 Monitoring of algal cultures
Algal cultures should be examined daily for clumps or aggregations of cells on the bottom. Colour of the culture is most important, and with experience, one can quickly determine if a culture is healthy. Microscopic examination of the algal culture should be done routinely using a compound microscope. Cultures should have cells of uniform size that are not clumped together, and are actively swimming if the species is motile (for e.g. Isochrysis sp., Tetraselmis sp.). If the cells are clumped, cell walls broken, more than one species present, or a species other than algae present, or if the culture is badly contaminated with bacteria, it should be discarded. The culture vessel should be well cleaned before next use. A bad odour emanating from a culture vessel usually indicates bacterial contamination. Algal cultures used in bivalve hatcheries are not axenic (bacteria-free). In order to have a healthy algal culture, bacterial levels must be kept under control, since they can depress growth of the algae and cause cultures to crash before reaching harvestable densities. Cultures with high level of bacteria should not be fed to larvae but should be discarded; they could be used for broodstock, if needed. If a severe bacterial contamination occurs in stock cultures, every effort should be made to clean the culture with antibiotics, or a new culture should be ordered from an algal culture centre. There are various methods for determining levels of bacteria contamination. Appendix 12 describes bactopeptone testing.
At the BBSR hatchery, a daily routine check of the algal cultures, giving results of visual inspection, is reported on an “Algal culture check” list (see Appendix 13). Monitoring of algal cultures under the microscope is done during periods of sub-culturing or harvesting. During the latter, algal cell density is estimated to calculate the volume required for feeding. There are two methods used most commonly in hatcheries to estimate algal cell density, haemocytometer and coulter counter. Coulter counters are expensive but useful machines. Sometimes used machines can be purchased from hospitals or factories. The time saved and the accuracy of the counts is superior to that when using the haemocytometer. At the Bermuda hatchery, funds were not available for a coulter counter, and a haemocytometer cell is used. This cell was initially developed to count blood cells, and consists of a thick glass slide with two chambers. A special coverslip is placed over these two chambers giving a total volume of 0.1 mm3 per chamber. The chambers are divided into a grid, to aid in counting cells within the area (Appendix 14). Before counting motile algal species, 1 or 2 drops of 10 percent formalin should be added to a 50 ml sample. The coverslip is mounted over the chambers, and the chambers are filled with the algal sample using a Pasteur pipette. Care is taken not to introduce any air bubbles, as the number of algae estimated is dependent on the exact volume of the chambers. Protocol–10 outlines the procedure used for estimating the number of algal cells using a haemocytometer cell.
PROTOCOL–10
ESTIMATING CELL DENSITY USING A HAEMOCYTOMETER CELL
1. Collect 10–20 ml of algal culture in a scintillation vial.
2. Add 2–3 drops of 10 percent formalin to culture if flagellate species to stop it from swimming. Mix sample thoroughly.
3. Mount cover slip on haemocytometer cell.
4. With a Pasteur pipette, retrieve 1 ml of sample, and introduce a drop into the chamber at the edge of the cover slip. Do not force sample in, allow it to run by capillary action. Make sure not to have any air bubbles in cell.
5. Fill grooves of cell completely with algal sample.
58 Installation and operation of a modular bivalve hatchery
6. Allow 1 or 2 minutes for cells to settle out on bottom of counting chamber.
7. Using a counter, count number of algal cells in at least three of the 25 squares. Count all cells lying within the square or overlapping the lines on the right-hand or bottom sides. Each square measures 0.2x0.2 mm.
8. Calculate the average number of cells per square.
9. To obtain the cell density: Average number of cells/square x 250. This gives you number of algal cells per µl.
10. To obtain the number of cells per ml multiply above number by 1 000.
Explanation:
Each cell is 0.004 mm3
Average number of cells is per 0.004 mm3
To obtain average number of cells per mm3, multiply by 250. 1 mm3= 1 µl
There are 1 000 mm3 in 1 ml. Multiply average by 1 000 to obtain number of cells per ml.
2.3.6 Alternate feed for spat
The production cost of microalgae using conventional phototrophic means, as described above, is high, ranging from 20–50 percent of hatcheries’ operating costs (Coutteau and Sorgeloos, 1992). Nutritionally adequate alternatives have been sought that may be more cost-effective than on-site algal production. Some of those tested include spray-dried, heterotrophically grown microalgae (Langdon and Onal, 1999), and microencapsulated artificial diets (Laing, 1987). Although these show potential in future rearing of bivalve spat, they are not commercially available. Other off the shelf alternatives that show more promise include microalgal concentrates; these are produced by centrifugation and refrigerated at 2–4 °C for 1–8 weeks. They have been used successfully as part of mixed or complete diets for larval or juvenile bivalves (Heasman et al. 2000). Such microalgal concentrates can either be prepared by hatcheries with existing infrastructure to produce microalgae and concentrates on-site, or by large specialized algal production facilities for sale to hatcheries (Brown and Robert, 2002). It was shown that not all algal species lend themselves to the process of concentration, notably the flagellates, P. lutheri and Isochrysis sp. (T-Iso) are easily damaged and deteriorate rapidly (Heasman et al. 2000). Another alternative is the use of dry, non-live microaglae which can be purchased commercially; studies conducted by several authors (Langdon and Onal, 1999; Davis and Campbell, 1998) found that a mixed live algal diet of T-Iso and C. calcitrans supplemented with spray-dried microalgae enhanced juvenile mussel growth; and that a mixed spray-dried algal diet of Schizochytrium and Spirulina contained the biochemical constituents necessary to satisfy the nutritional requirements of mussels.
In light of the restricted capacity of the algal culture facility at BBSR, replacement of live algal diet with commercially available dry algal mixture was tested on scallop spat. Results of a short study on growth of E. ziczac show that food ration composed solely of dry algae meet the nutritional requirements of larger zigzag spat (>3 mm); shell growth was seen to increase by 1.2 mm per week in the first three weeks. This growth was comparable or better to that of spat fed live algae. On the other hand, smaller spat (<3 mm), seemed to fare better in the long term when fed live algae. These results led to the routine use of commercially purchased dry algae for older spat, prior to transfer to grow-out sites. Dry microalgae are purchased from Reed Mariculture (www.instant algae.com).