PART III PRACTICAL EXERCISES

SCS/GFO/81/PE-1

MICROSCOPIC EXAMINATION OF PLANKTONIC ORGANISMS¹

1. INTRODUCTION

Planktonic organisms are best examined and identified with the aid of a microscope. Knowledge of the proper use and care of the microscope is needed for this exercise. Fresh mounts of plankton will be prepared by the participants.

2. OBJECTIVE

To demonstrate the proper use of the compound microscope and preparation of wet mounts.

3. MATERIALS

Compound microscope
Glass slide and cover slip
Medicine dropper
Toothpick
Plankton sample

4. PROCEDURE

1. With a medicine dropper, place a drop of the plankton sample on the glass slide. Place the cover slip on the drop gently with a toothpick.

2. Examine the slide under the low power objective and locate the organisms. Use the fine adjustment and high power objective of the microscope.

3. Identify the common organisms seen.

5. REFERENCE

Rimando, L.C. and R.C. Olaguer. 1980 Laboratory manual for biology. University of the Philippines at Los Banos, College, Laguna. 53p.

¹ Prepared by Rafael D. Guerrero III

SCS/GFO/81/PE-2

STERILIZATION OF CULTURE VESSELS AND OTHER MATERIALS¹

1. INTRODUCTION

Sterilization is defined by Hamilton (1973) as a process that ensures the total inactivation of all microbial life. This can be achieved by volatile and non-volatile disinfectants, moist air method by means of an autoclave or pressure cooker, dry heat method by means of hot-air oven and through filtration system to clear water supply and air supply for aeration. These methods are important in all phychological research not only to minimize bacterial contamination, but also to attain unialgal cultures. In the SEAFDEC Natural Food Section, the above methods have been adopted and is recommended to prospective algal culturists. However, it is important to point out that the above methods are not the only answer to attain sterility; what is needed is simple chemical cleanliness that forms the basis for most sterilization techniques.

2. OBJECTIVE

To acquire knowledge of the basic aseptic techniques used in algal culturing.

3. MATERIALS

Test tubes
Dextrose bottles
Glass tubings
Plastic tubings
Pipettes
Liquid and solid nutrient media
Flasks
Petri dishes
Gallon jars
Carboys
Vinyl tanks
Caserole

¹ Prepared by Ms. M. Rosales, SEAFDEC Aquaculture Department, Natural Food Project, Tigbauan, Iloilo, Philippines

4. METHODOLOGY/PROCEDURE

4.1 Sterilization of materials used in algal isolation

Sterilization of materials used in algal isolation is achieved by means of an autoclave at 121°C in pure saturated steam at 1 kg/cm² pressure.

1. Wash test tubes and Petri dishes thoroughly with soap and water.

2. Drain and dry Petri dishes in the tray and test tubes in the test tube rack.

3. Label and wrap Petri dishes tightly with material that permit easy access of steam. For the test tube, plug with cotton/ gauze.

4. Autoclave the Petri dishes and test tubes for 45 minutes.

5. Dip dry in the oven at 100°C for 20 minutes before using for isolation.

4.2 Sterilization of liquid and solid media

Sterilization of liquid and solid media is also achieved by means of an autoclave.

1. Wash 500 ml flasks/dextrose bottles thoroughly with soap and water.

2. Dip dry.

3. Plug flask/dextrose bottles with cotton/gauze.

4. Sterilize for 5 hours in hot-air oven with 100°C temperature.

5. Cool flask/dextrose bottles.

6. Pour the necessary amount of media needed in the sterilized flask/dextrose bottles.

7. Return the cotton/gauze plug in the flask/dextrose bottles.

8. Cover cotton/gauze plug with paper and tighten with rubber band.

9. Autoclave flask/dextrose bottles containing the media.

4.3 Sterilization of materials used in maintenance culture

For the maintenance cultures, culture vessels range from test tubes to 200-liter vinyl tanks. Test tube/flask cultures serve as stock cultures and starters for 1-liter dextrose bottles culture that will serve as starter for 3.5 gallon jars. The gallon jar cultures will then serve as starter for the vinyl tanks.

Adopt procedures (a) to (e) of Section 4.2 for test tubes, flasks, dextrose bottles and gallon jars.

For the carboys and vinyl tanks, sterilization is achieved by means of chemical methods. This chemical treatment is done if the mass-produced algal species is changed. The following steps are as follows:

1. Wash the carboy/vinyl tanks with soap and water.

2. Treat with 10 percent HCl for 2 days.

3. Rinse thoroughly with filtered freshwater.

However, for daily renewal of cultures, with the same algal species being cultured, adopt only steps (a) to (c) of the above procedure.

4.4 Sterilization of pipettes, glass tubings and plastic tubings

Smaller items such as pipettes, glass tubings and plastic tubings are sterilized in a separate system. The following steps are suggested:

1. Wash pipettes, glass tubings and plastic tubings with tap water.

2. Treat them overnight with HCl. Fifty percent HCl concentration is used for pipettes while 10 percent HCl concentration is used for glass tubings and plastic tubings.

3. Wash them thoroughly with filtered freshwater.

4. Dip dry.

5. Place pipettes in a metal canister and wrap glass tubings and plastic tubings with an ordinary paper.

6. Sterilize pipettes in the oven for 5 hours at 100°C temperature.

7. Autoclave glass tubings and plastic tubings for 45 minutes.

8. Dry before using.

4.5 Sterilization of water supply

1. Pump water and pass through sand filter, then through 3.0 microns filter and through 2.5 microns filter.

2. Stock in a plastic tank/reservoir (optional).

3. Cover the tank with black plastic sheet to prevent algal growth.

4. Boil water in the desired volume of culture.

5. Store and cool in 4-liter flask for the culture of algal stock cultures. For the gallon jar cultures, the sterilized water is stocked in a covered 80-liter plastic container.

For the carboys and vinyl tanks, directly use the filtered water being stored in the tank.

5. REFERENCE

Hamilton, R.D. 1973 Sterilization. In J.R. Stein (editor) Handbook of Phycological Methods. Cambridge Univ. Press. Cambridge: 181–193

SCS/GFO/81/PE-3

PREPARATION OF VARIOUS CULTURE MEDIA AND STOCK SOLUTIONS¹

1. INTRODUCTION

Several media formulations have been used for algal cultivation depending upon the nutrient requirement of the cultured algal species. Usually, the chemical composition of a defined medium have been derived and modified from basic formulations of the pioneering phycologists. The changes have been adopted from the results of nutrient requirement experiments. These are basic phycological tests not only for academic purposes but also for algal mass production which is necessary for aquaculture practices.

In the SEAFDEC Natural Food Section, three nutrient media have been employed for algal mass production, namely: Guillard and Ryther's (1962) modified F medium, Walne's (1974) medium and Liao and Huang's (1970) modified medium. Refer to Appendix A for the chemical composition of each medium. These media were tested and found to be suitable for the algae being massproduced.

To facilitate media preparation, stock solutions are prepared 1000x the actual concentration to avoid tedious weighing since the chemical formulation consists of elements weighing in micrograms. In the tabulation, some of the media are prepared only 500x the actual concentration, this preparation is done to avoid bacterial contamination resulting from long storage.

For demonstration purposes, Walne's (1974) medium will be used.

2. MATERIALS

Dextrose bottles (1 000 ml and 500 ml)
Petri dishes
Test tubes
Pipettes
Flasks
Distilled water
Weighed chemical nutrients of Walne's Medium
Spatula
Wax paper
Agar

¹ Prepared by Ms. M. Rosales, SEAFDEC Aquaculture Department, Natural Food Project, Tigbauan, Iloilo, Philippines

3. METHODOLOGY/PROCEDURE

3.1 Liquid medium

1. Prepare sterilized items such as dextrose bottles (1 000 ml and 500 ml), stirring rod, medicine dropper and pipettes.

2. Autoclave 1 200 ml of distilled water for 45 minutes. In autoclaving, place 900 ml distilled water in a 1-liter dextrose bottle, 200 ml in a 500-ml dextrose bottle and the remaining 100 ml in a 250-ml flask.

3. Weigh the respective major ions, trace metals, and vitamins as provided in the tabulation of Appendix A.

4. Wrap and label weighed chemicals in wax paper in preparation for the mixing process.

5. Prepare the trace metals primary stock first, since these chemicals readily absorb moisture from air.

6. Dissolve the elements one by one in 100 ml of sterilized distilled water.

7. Mix well by means of a stirring rod.

8. Add a few drops of IN HCl until solution becomes clear.

9. For the vitamins primary stock, dissolve vitamins B₁ and B₁₂ in 200 ml of sterilized distilled water.

10. Mix well by means of a stirring rod.

11. For the major ions, dissolve and mix thoroughly the following: NaNO₃, Na₂EDTA, H₃BO₃, FeCl₃.6H₂O, MnCl₂.4H₂O and NaH₂PO₄.2H₂O in 900 ml of sterilized distilled water added in sequence and one at a time. Note that in dissolving the above ions, the phosphate source is added last in order to avoid precipitation.

12. When clear solution is achieved, add 100 ml of vitamins primary stock and mix well.

13. Add 1 ml trace metals stock and mix well.

14. Use 1 ml per liter culture.

3.2 Solid medium

The preparation is good for 1 liter solid medium.

1. Prepare sterilized Petri dishes, test tubes, two flasks and stirring rod.

2. Weigh 15–20 grams agar.

3. Place 75 percent of the total amount of water in one flask and the remaining 25 percent water in the other flask.

4. Place the weighed agar in a flask containing 75 percent water.

5. Add 1 ml of the enrichment nutrient, e.g., Walne's Medium in a flask containing 25 percent water.

6. Precook the agar in a boiling water bath. Stir the mixture while cooking.

7. Autoclave the two flasks for 45 minutes.

8. Cool both flasks to 56°C.

9. Mix thoroughly the contents of the two flasks.

10. Pour slowly into Petri dishes and test tubes, 18–20 ml/plate/ slant.

11. Store plates/slants at room temperature in an upside down position.

4. REFERENCES

Liao, I.C. and T.L. Huang. 1970 Experiments on the propagation and culture of prawns in Taiwan. Proceedings of the 14th Session of the Indo-Pacific Fisheries Council: 1–26

Guillard, R.R.L. and J.H. Ryther. 1962 Studies of marine planktonic diatoms. I. Cyclotella nana Hustedt and Detonula confervacea (Cleve) Gran. Can. J. Microbiol. 8: 229–239

Pringsheim, E.G. 1951 Methods for the cultivation of algae. In G.M. Smith (editor) Manual of Phycology. The Ronald Press Co., New York: 357–397

Walne, P.R. 1974 Culture of bivalve molluscs. The Whitefriars Press Ltd., London and Tondridge: 1–173.

APPENDIX A

I. Liquid Media

II. Solid Media

III. Stock solutions

SCS/GFO/81/PE-4

USE OF THE HEMACYTOMETER FOR COUNTING PHYTOPLANKTON¹

1. INTRODUCTION

The chief use of the hemacytometer is for making red blood cell, white blood cell, and blood platelet counts. It can also be an inexpensive tool for counting algae with sizes of 2–30 u in diameter with densities up to 50× 10⁷ cells/ml.

The improved Neubauer hemacytometer consists of a thick rectangular slide with an H-shaped trough forming two counting areas (Fig. 1). With the cover slip in place, each area forms a chamber with a depth of 0.1 mm.

The total ruled area of one chamber is 9 mm². The four corner squares (1 mm × 1 mm) are subdivided into 61 smaller squares and the centre square (1 mm × 1 mm) is subdivided into 25 smaller squares, each has an area of 0.04 mm² (.2 mm × .2 mm) (Fig. 2).

2. OBJECTIVES

2.1 To demonstrate the use of hemacytometer for counting phytoplankton.

2.2 To determine the population densities of algal cultures using the hemacytometer.

3. MATERIALS

Improved Neubauer hemacytometer, microscope, algal test organisms (Chaetoceros sp. and Tetraselmis sp.), Lugol's iodine solution, cover slips, tissue paper, test tubes, test tube rack, pasteur pipettes, bulbs.

¹ Prepared by Ms. E. Aujero, SEAFDEC Aquaculture Department, Natural Food Project, Tigbauan, Iloilo, Philippines

Fig. 1 Macroscopic view of the hemacytometer

Fig. 2 Ruling and dimensions of one chomber of the improved Neubauer hemacytometer

4. PROCEDURE/METHODOLOGY

4.1 Sample preparation

1. Label two test tubes: Chaetoceros and Tetraselmis.
2. Put 5–7 ml of the cultured algae in the labeled test tubes.
3. Add one drop of Lugol's iodine to both tubes.
4. Mix and let tubes stand in rack.

4.2 Filling the chamber

1. Clean the hemacytometer and the cover slip with soap and water or with rubbing alcohol. They must be free from dust, lint or grease.

2. Place the cover slip centrally over the ruled areas.

3. Using a clean pasture pipette, put a drop of well-mixed algal sample in the “V” groove of the metal surface. Fill both chambers.

4. Check for the evenness of cell distribution under low power magnification (Fig. 4).

5. If the following should occur, repeat steps (a) to (d); presence of air bubbles, overflowing, underfilling, and uneven distribution of cells (Fig. 3).

6. Allow cells to settle for 3–5 minutes before counting.

4.3 Counting

1. For cells greater than 6 u and not too dense cultures, the total count is made in any one block (A, B, C, D, E) as shown in Fig. 2*

   • Start at the top left square and count only those cells which lie within or touching the boundary line as shown in Fig. 5.

   • Make a duplicate count in the corresponding block in the second chamber.

   • Record the counts in individual 1 mm² blocks.

* For minute and dense populations, count the cells in the five small squares in the centre block (E Block).

Fig. 3 Properly and improperly charged counting chamber

Fig. 4 Distribution of cells

Score board for each small square Total count is 48

Fig. 5 Method of counting cells

4.4 Calculation

(a) If all the cells in the individual blocks are counted, the density (d) would be:

(b) If the cells were counted only in the five small squares in the centre block (E Block), the following formula may be used:

where 10 = the 10 squares of the 2 chambers and

4×10^-6 = the volume of sample over the small square area which is equivalent to .004 mm³ (.2 × .2 × .1) expressed in cm³ (ml).

4.5 Data to be gathered

1. Cell count of Chaetoceros sp. using counting procedures (a) and (b).

2. Cell count of Tetraselmis sp. using counting procedures (a) and (b).

5. REFERENCES

AO Bright line hemacytometer counting chamber, American Optical, Buffalo, N.Y., 1425, 19pp.

Guillard, R.L. 1973 Division rates. In Phycological Methods, Janet R. Stein (ed.), Cambridge University Press, Cambridge, 289–311pp.

Martinez, M.P., Chakroff, J.B. Pantastico. 1975 Direct Phytoplankton counting technique using the hemacytometer. In Phil. Agriculturist 59. 43–50

SCS/GFO/81/PE-5

ALGAL ISOLATION BY AGAR PLATING AND INOCULATION TECHNIQUES¹

1. INTRODUCTION

The isolation of an algal unit into a medium suitable for growth is required to establish a unialgal culture. There are several methods of isolation, i.e., capillary pipette, agar plating, cover glass attachment, repeated subculture, etc. The choice depends on the algal type and size.

The method described below is applicable to many species. The source of algae for isolation is a water sample containing mixed plankton population.

2. OBJECTIVES

2.1 To demonstrate algal isolation and inoculation techniques.

2.2 To produce unialgal cultures from a mixed plankton sample using the agar plate method.

3. MATERIALS

Microscope, Pasteur pipettes and bulbs, Erlenmeyer flasks, slides, cover slips, test tubes, Petri dishes, cotton plugs, alcohol lamp, wire loop, test tube rack, Walne's medium, F medium, agar, water sample containing mixed plankton population.

¹ Prepared by Ms. E. Aujero, SEAFDEC Aquaculture Department, Natural Food Project, Tigbauan, Iloilo, Philippines

4. PROCEDURES

All media, glasswares and other materials should be sterile.

1. Media preparation

   • Prepare agar plates and agar slants containing Walne's and F growth media separately, solidified with 1.5–2 percent agar.

   • Prepare liquid media in flasks using Walne's and F formulations.

2. Place 1–2 drops of water sample on one side of the agar plate.

3. Using aseptic technique, use a flamed wire loop to make parallel streaks of the suspension on the agar.

Fig. 1. Pattern of streaks on agar surface; starting point is shown on tip of arrow.

4. Cover, invert the plate and incubate for 6–8 days on lighted shelves.

5. Check the colonies for growth of desired species. With a wire loop take a sample and place it in a drop of sterile water on a slide. Cover with a cover slip and examine under the microscope if the desired species has been isolated.

6. Repeat the streaking procedure on agar plates from a single colony and allow colonies to develop.

   Repeat procedures (d) and (e) and check if colonies are unialgal.

7. Transfer algal units from the desired colony/colonies to liquid medium in flasks and agar slants.

Fig. 2. Pattern of streak on agar slant

8. Leave on lighted shelves for 6–8 days; gently swirl liquid cultures daily.

   Cultures on agar slants and flasks can be kept as stock cultures and/or scaled up to larger volumes using enriched media.

9. Data to be gathered:

   • algal species that predominate in Walne's medium
   • algal species that predominate in F medium
   • all isolated algal species

5. REFERENCE

Hoshaw, W. and R. Rosowski. 1973 Methods for microscopic algae. In Handbook of Phycological methods, Janet R. Stein, Ed., Cambridge University Press, Cambridge, pp. 53–68

SCS/GFO/81/PE-6

HARVESTING, PRESERVATION AND UTILIZATION TECHNIQUES
FOR PHYTOPLANKTON¹

1. INTRODUCTION

A continuous supply of natural food is a must for successful aquaculture operation. Maintenance of live algal cultures are affected by weather conditions, hence, techniques of preservation have to be developed. Other advantages of maintaining algae in preserved states are: reduction of time and space, reduction in the toxic effect of feeding large volumes of algal culture medium to the larvae, ease of distribution and storage of cultures in large numbers and preservation of the genetic constitution of the strains over a long period of time.

Prior to preservation, algal cultures must be harvested. Several methods are available; however, selection of the most suitable depends on the ultimate use of the algal cells, cost, and efficiency of the process.

The methods presented below for harvesting and preservation are some of the less expensive and reasonably dependable.

2. OBJECTIVE

To demonstrate algal harvesting and preservation techniques.

3. MATERIALS

Centrifuge, 1-liter beakers, stirring rod, Pasteur pipettes, algal cultures (Chaetoceros sp. and/or Tetraselmis sp.), IN NaOH, freezer, plastic bags, plastic tubings, rubber bulbs, magnetic stirrer (optional), magnetic stirring bars (optional), pH meter, IN HCL.

¹ Prepared by Ms. E. Aujero, SEAFDEC Aquaculture Department, Natural Food Project, Tigbauan, Iloilo, Philippines

PROCEDURES

4.1 Harvesting

Two methods are presented:

1. Centrifugation - to be observed.

   1. Resuspend cell concentrate in small volume of diluted seawater (ca. 80%) and proceed to procedures 4.2 and 4.3)

2. Chemical flocculation

   1. Take the initial pH of the culture.

   2. Transfer the algal culture to a one-liter beaker or larger container.

   3. Add IN NaOH to the algae in 1–2 ml portions accompanied by vigorous stirring for 5 minutes followed by gentle stirring for 15 minutes or until floc forms.

   4. Allow to settle for one hour.

   5. Inspect the supernatant for clarity.

   6. Drain off the supernatant without disturbing the cells by using plastic tubing.

   7. Neutralize the algal concentrate to its initial pH with IN HCl.

The harvested algal concentrate may be utilized for feed or for starters or may be preserved for later use.

4.2 Preservation

1. Sundrying

   1. Spread the algal concentrate thinly on trays and expose it to sunlight.

   2. Store dried algae in covered bottles and keep at room temperature.

2. Freezing

   1. Pour the algal concentrate into small plastic bags, seal and freeze.

4.3 Utilization

1. For feed

   1. Use the frozen algae for feed.

   2. Use the dried algae for feed.

2. For starters

   1. Thaw the frozen sample in a waterbath at room temperature.

   2. Suspend the cells in the culture medium, provide light and aeration.

5. REFERENCES

Aujero, E. and O. Millamena. 1979 Viability of frozen algae used as food for larval penaeids. Quarterly Research Report, 4th Quarter, Vol. III, No. 4.

Hom-Hansen. 1973 Preservation by freezing and freeze-drying. In Handbook of Phycological Methods, Janet R. Stein (ed.), Cambridge University Press, Cambridge. 195–205pp.

Millamena, P. and E. Aujero. 1978 Preserved algae as food for Penaeus monodon larvae. Quarterly Research Report, 4th Quarter, Vol. II., No. 4

Ukeles. 1976 Cultivation of plants. In Marine Ecology Vol.III, O. Kinue (ed.) 462–464pp.

SCS/GFO/81/PE-7

CULTURE OF ALGAE AND PLOTTING OF GROWTH CURVES¹

1. INTRODUCTION

Phytoplankton are a necessary part of the diet of many aquatic organisms. Many algal species have been used as food organisms in the culture of penaeid protozoea (Dawson, 1966). Skeletonema costatum has been widely used both in extensive and intensive hatchery systems (Simon, 1978). Other species used include Tetraselmis chuii and Chaetoceros calcitrans. Mass production techniques for the algae should be exercised to have a continuous supply of feed. This may depend on the quality of feed and its culture medium. A dependable method for culturing sufficient quantities of phytoplankton foods for different stages should be noted.

The method described is suggested for cultures from 1-liter to gallon containers and 20-liter carboys. Three culture media are used, namely: Walne (1974), Guillard and Ryther (1962) and Liao and Huang (1970).

2. OBJECTIVES

2.1 To culture phytoplankton in one-liter culture containers, gallons and 20-liter carboys.

2.2 To determine growth curves of the phytoplankton at 4-hour intervals.

3. MATERIALS

1-liter dextrose bottles
Gallon containers
20-liter carboys
Glass tubing
Plastic tubing
Cotton
Haemacytometer
Cover slips
Pipettes
Hand tally counter

¹ Prepared by Ms. F. Escritor and Ms. S. Javellana, SEAFDEC Aquaculture Department, Natural Food Project, Tigbauan, Iloilo, Philippines

4. PROCEDURE

1. Wash dextrose bottles, gallons and carboys with soap and water. Soak in muriatic acid, dry and sterilize at 100°C for 5 hours.

2. Prepare stock solutions for culture media. Walne's medium is used for culture of green algae while Guillard and Ryther's medium is for diatoms. However, for gallons and carboys, Liao and Huang's media is used.

3. Put one liter of seawater in each sterilized dextrose bottle.

4. Put glass tubing attached to an aeration system by plastic tubing for aeration purposes.

5. Fertilize with Guillard and Ryther's medium as instructed in Laboratory Exercise No. 3.

6. Examine inoculum as to condition of the cells and population density and presence of contaminants. Inoculum should only be from a pure culture with high population density, young cells and motile, specifically for the green phytoflagellate.

7. Compute volume of inoculum and initial density of 50 000 cells/cc with the following formula:

Where N - volume of inoculum
X - desired initial density
Y - actual count of source
V - volume of culture medium

8. Count algae at 4-hour intervals using the haemacytometer.

9. Plot algal counts to show the growth curves.

10. Harvest at exponential growth phase for inoculum for gallons.

11. Culture in gallons with the same method and harvest at exponential growth phase for inoculum for carboys.

12. Data to be collected:

    1. Algal counts at 4-hour intervals.
    2. Growth curves showing the following phases in the graph (Fogg, 1965).

       Lag phase - slow algal growth.

       Exponential growth phase - cell multiplication is rapid and cell numbers increase.

       Phase of declining relative growth - cell numbers remain more or less stationary.

       Death phase - algal culture collapses and nutrients are already utilized.

5. REFERENCES

Fogg, G.E. 1965 Algal cultures and phytoplankton ecology. Univ. Wisconsin Press

Dawson, Y. 1966 Marine botany, Halt, Rinehart and Winston, Inc. pp. 27–46

Guillard, R.R.L. and J.R. Ryther. 1962 Studies of marine planktonic diatoms. I. Cyclotella nana Hustedt, and Detonula confervacea (Cleve) Gran. Can. Jour. Microbiol. 8: 229–239

Liao, I.C. and T.L. Huang. 1970 Experiments on propagation and culture of prawns in Taiwan. Proceedings of the 14th Session of the Indo-Pacific Fisheries Council. pp 1–26

Simon, C.M. 1978 The culture of the diatom, Chaetoceros gracilis and its use as food for penaeid protozoeal larvae. Aqua. 14: 105–113

Walne, P.R. 1974 Culture of bivalve molluscs. The Whitefriars Press Ltd., London and Tondridge. pp. 1–173

SCS/GFO/81/PE-8

CULTURE OF BRACHIONUS PLICATILIS¹

1. INTRODUCTION

Development of techniques for the mass culture of zooplankton is an important prerequisite for successful larval rearing of fish larvae to meet aquacultural needs. Many fish biologists in the field of seed production, have investigated the technique for mass culture of the marine rotifer, Brachionus plicatilis Muller and the procedure of culture has been rapidly improved (Furukawa and Hidaka, 1972; Hirata, Kanazawa, Yamamidori and Yasuda, 1973; Hirayama and Kusano, 1972; Hanayama and Ogawa, 1972; Nagawa, Oohara, Kitamura and Nakagawa, 1972).

With its high reproductive potential, the rotifer lends itself easily to mass culture. At present, mass culture of the rotifer is performed mostly according to the experiences by using the green phytoplankton as its food. Many experiences in Japan tell us that a marine species of Chlorella and baking yeast are the suitable food for the mass production of these organisms. At the Phycology Section of the Aquaculture Department, SEAFDEC, larval food production involves the mass culture of Brachionus plicatilis using Chlorella and bread yeast indoors and outdoors.

2. OBJECTIVES

2.1 To demonstrate the culture techniques used in the mass culture of Brachionus plicatilis

2.2 To determine population density of the rotifer, B. plicatilis at a certain culture period.

2.3 To demonstrate counting technique/procedure for monitoring B. plicatilis cultures.

¹ Prepared by Ms. S. Javellana and Ms. F. Escritor, SEAFDEC Aquaculture Department, Natural Food Project, Tigbauan, Iloilo, Philippines

3. PREPARATION OF CONICAL TANKS

3.1 Materials

Muriatic acid (HCl)
Conical tanks
Airstones
Cross tee
Plastic tubing

3.2 Methodology

1. Prepare 10 percent muriatic acid by adding 100 ml muriatic acid/liter of seawater.

2. Soak the conical tanks in 10 percent muriatic acid overnight and wash thoroughly with freshwater before use.

3. Fill the tank with seawater up to the desired volume.

4. Provide aeration through the plastic tubing attached to aeration system. A plastic tubing extends from each valve to the tank terminating in an airstone. Aeration can be regulated through the tee valves.

However, availability of food and culture of green phytoplankton like Chlorella virginica/Dunalliela sp./Tetraselmis sp. must be scheduled before stocking in order to have enough food supply throughout the culture/ experiment.

Also, preparation of food dosage must be taken into consideration.

For Chlorella, a population density of 30 × 10⁴ cells/ml is maintained but for Dunalliela sp./Tetraselmis sp. a feeding density of 5 × 10⁴ is maintained and for bread yeast, dissolve 0.25 g - 600 ml and add 100 ml/conical tank.

4. COUNTING PROCEDURE

The Sedgwick-Rafter (S-R) counter is the most commonly employed device for zooplankton counting like the rotifer, Brachionus plicatilis. The counter is 50 mm long by 20 mm wide by 1 mm deep. The total area is 1 000 mu² and the total volume is 1 × 10¹² cubic micron or 1 000 mu³ or 1 ml. The 10 × objective is generally used because the depth of the chamber normally precludes the use of 40 × or 100 × objectives.

The total number of zooplankton in the S-R counter is calculated by counting all the organisms found in the counter. Population counts are on per ml basis. A three random sampling is recommended to get an accurate count.

4.1 Materials

Sedgwick-Rafter counter
Hand tally counter
Tuberculine syringe (1 ml capacity)
Lugol's solution
Compound microscope
Beakers or plastic cups (100 ml capacity)

4.2 Methodology

1. Using a beaker/plastic cup (100 ml) get a sample of the culture after stirring vigorously.

2. Add one to two drops of Lugol's solution* to the sample to paralyze the organisms.

3. Transfer 1 ml of the well-mixed sample by the use of a tuberculine syringe to the S-R counter.

4. Let it stand for 2–3 minutes to permit settling of the zooplankton before counting.

5. Start counting from one of the corners of the counting chamber, passing through one or four divisions at a time, in order to avoid counting the same organisms twice.

6. To calculate the concentration of the organisms:

Where C = actual counts of organisms
F = number of sampling

* Composition of Lugol's solution:

5. MONITORING

Daily monitoring of population density of both the feed (algae) and the test organism (rotifer) is done to maintain feeding concentration of the algae (feed) and growth of the test organisms.

Sampling is done every 24 hours from start of culture.

5.1 Materials

5.2 Methodology

1. Get sample for both the algae and the rotifer by using a beaker or plastic cup after stirring vigorously. One individual is recommended to do the sampling throughout the duration of the experiment to minimize sampling errors.

2. Fix the samples with a few drops of Lugol's solution.

3. Count the food (algae) density using the hemacytometer. One individual is recommended to do the counting to minimize counting errors.

4. Counting of the population density of the test organism can be done later using the Sedqwick-Rafter counter. One individual also is recommended to do the counting to minimize errors.

5. Determine the population density of the food (algae) from the stock culture to be used as feed.

5.3 Computation

1. If algal count is within the range of the feeding density then no algae will be added.

   Example No. 1: Actual algal count = 298 × 10³
   Desired feeding density = 300 × 10³

2. If algal count is less than the range of the feeding density, then food will be added. Algal population density count from stock culture must be done first before counting:

   

   Where:	desired	=	desired feeding density
   	actual	=	actual algal count in the tank
   	algal count	=	algal count of stock culture
   	volume	=	volume of the tank

   Example No. 2:

   Desired density is	=	300 × 103
   Actual algal count is	=	190 × 103
   Algal count of stock culture is	=	10 000 × 103
   Volume	=	15 li

   Therefore:

   

   An equal volume of culture medium in the tank must be drained before adding the cultured feed to maintain the level of seawater in the tank.

3. If algal count is more than the range of the feeding density, a certain volume of the water will be drained using the formula:

   

   = ×

   Therefore: Volume to be drained = Volume of the tank - X

   Example No. 3:

   Desired density is 300 × 10³

   Volume of the tank is 15 li

   Actual count is 500 × 103

   Therefore: 15 li - 9 li = 6 li (Volume to be drained)

   After draining is done, an equal amount of seawater must be added to the tank to maintain the water level.

Draining procedure:

Draining of water is done by 2 individuals.

1. Use 30 μ mesh net to prevent the entrance of the test organism.

2. Siphon the desired volume of water to be drained using a 3/8 diameter plastic tubing with a beaker (1 li cap.) below to measure the volume of water.

6. HARVESTING OF BRACHIONUS CULTURE

6.1 Materials

Rubber hose
30 and 40 μ mesh nylon nets
Pail

6.2 Methodology

1. Drain the culture using the rubber hose with 40 and 30 μ nylon nets below respectively. The detritus will be almost completely removed and only the rotifers will be concentrated in the nets.

2. Wash again several times with seawater to remove other contaminants like diatoms.

3. After washing, the rotifer obtained can be used as feed to the larvae and others will be used to start the culture again.

7. REFERENCES

Alon, N.C. 1974 Simplified culture of the rotifer, Brachionus plicatilis Muller using yeast diets and EDTA. Aquaculture.

American Public Health Association. 1971 Standard methods for the examination of water and wastewater, 13th Edition, APHA…, New York, pp. 734–735

Hirata, H. 1974 An attempt to apply an experimental microosm for the mass culture of marine rotifer, Brachionus plicatilis Muller. Mem. Fac. Fish. Kagoshima Univ. Vol. 23. pp. 163–172

Hirayama, K. and S. Ogawa. 1972 Fundamental studies on physiology of rotifer for its mass culture - I. Filter Feeding of Rotifers. Bull. Jap. Soc. Sci. Fish., 3(11): 1207–1214

Hirayama, K., Takogi Kenzo and Kimura Hiroshi. 1979 Nutritional effect of eight species of marine phytoplankton on population growth of the rotifer, Brachionus plicatilis. Bull. Jap. Soc. Sci. Fish., 45(1): 11–16

SCS/GFO/81/PE-9

QUALITY ANALYSIS OF ARTEMIA CYSTS¹

1. INTRODUCTION

The quality of Artemia cysts varies widely for different commercially available brands. An important criterion used by buyers in selecting a specific cyst-product to be fed for their cultured species is the amount of nauplii per gram product. Quality evaluation of Artemia cysts may be done using the following indices: hatching percentage and hatching efficiency.

Hatching percentage is the ratio of the hatched cysts over the unhatched full cysts. Hatching efficiency, on the other hand, is the quantity of cysts that should be incubated to produce a million nauplii. Determination of these values would greatly help prevent wasteful hatching and maximize the use of Artemia.

2. OBJECTIVE

To evaluate Artemia cyst quality by determination of the hatching percentage and hatching efficiency.

3. MATERIALS

1 g Artemia cysts
Graduated cylinder, 100-ml capacity, 4 pieces
Micropipette, with 3–5 pieces plastic tips
Test tubes, 5-ml capacity, 30 pieces
Rotator axle
Petri dishes, 6–10 pieces
Medicine droppers, 2 pieces
Wash bottle
Parafilm or small plastic sheets
Rubber bands
Dissecting microscope
Chemicals: Lugol's Iodine
                  IN NaOH
                  NaOCl

¹ Prepared by Ms. R. Figueroa, SEAFDEC Aquaculture Department, Natural Food Project, Tigbauan, Iloilo, Philippines

4. PROCEDURE

4.1 Hatching percentage (H%)

Note: In the determination of hatching percentage, only full cysts are considered while empty cysts are excluded.

1. Weigh 250 mg Artemia cysts.

2. Incubate cysts in 100 ml seawater for 24–48 hours at room temperature.

3. Pipette five 100 μl subsamples into Petri dishes.

4. Fix subsamples with one to two drops of Lugol's Iodine.

5. Count the nauplii and emerging nauplii (umbrella stage) under a dissecting microscope.

6. Place one drop NaOH and four drops NaOCl to the unhatched cysts.

Data to be gathered

N = number of hatched cysts, including the umbrella stage

C = decapsulated full cysts

4.2 Hatching efficiency

1. Weigh three samples of 250 mg cysts.

2. Place the cyst samples in separate graduated cylinders and hydrate in 80 ml seawater for one hour.

3. Adjust the volume of seawater to 100 ml.

4. Pipette ten 250 μl subsamples from each cylinder into the test tubes and add seawater up to the mark.

5. Cover tubes with parafilm or plastic pieces tide with rubber bands.

6. Arrange tube into the rotator axle and rotate for 24–28 hours at 30°C under continuous illumination.

7. Fix samples in tubes with Lugol's Iodine.

8. Count the number of nauplii.

Data to be gathered

N = number of hatched nauplii

Hatching efficiency (a and b)

(a) N × 4 × 4 × 100 = number of nauplii/g cysts

(b)

5. REFERENCES

Sorgeloos, P. et al. 1978 The use of Artemia cysts in Aquaculture: The concept of “hatching efficiency” and description of a new method for cyst processing: 715–721.
In Proc. 9th Ann. Meeting WMS. (ed.) Avault, J.W., Jr. Louisiana State University, Baton Rouge (LA-U.S.A.): 807pp.

Vanhaecke, P. and P. Sorgeloos. 1981 International study on Artemia. XIX. Hatching data on 10 commercial sources of brine shrimp cysts and re-evaluation of the “hatching efficiency” concept. Paper presented at the World Mariculture Society-Technical Sessions, Seattle (WA-USA), March 8–10, 1981.

SCS/GFO/81/PE-10

DECAPSULATION OF ARTEMIA CYSTS¹

1. INTRODUCTION

The presence of a thick outer shell (chorion) in the cysts often causes problems in the use of Artemia as food for fish and crustaceans. To solve this, the decapsulation technique was developed. Decapsulation is an oxidation process where the outer layer of the cyst is dissolved in hypochlorite, without affecting the viability of the embryo. Removal of the chorion disinfects the cysts and increases hatching rate, hatching efficiency and energy content. Furthermore, decapsulated eggs may be fed directly to some fish and crustacean larvae.

2. OBJECTIVE

To demonstrate the decapsulation technique for Artemia cysts using sodium hypochlorite (NaOCl)

3. MATERIALS

5 g Artemia cysts
Graduated glass cylinder, 1-L
                                        100-ml
                                        10-ml
Beaker, 1-L
Measuring pipette, 5-ml
Refractometer with refractive index
Tissue paper
Wash bottle
Laboratory thermometer
Distilled water
Ice
Chemicals: NaOCl
                  1 N NaOH
                  0.1N HCI

¹ Prepared by Ms. R. Figueroa, SEAFDEC Aquaculture Department, Natural Food Project, Tigbauan, Iloilo, Philippines

4. PROCEDURE

4.1 Preparation of decapsulation solution

Composition of decapsulation solution: NaOH, NaOCI and seawater.

Ratio: For every gram of cysts, 20 ml decapsulation solution is needed.

1. Determine the strength of hypochlorite

   1. Pipette 5 ml NaOCI
   2. Determine the refractive index using a refractometer
   3. Compute the activity of NaOCI (g/L) using the following formula:

      For new solution:

      Y = 3 000X - 4 003

      Where Y = Activity and X = Refractive Index

      For old solution:

      Y = 2707X - 3611

      e.g. (new solution)

      Y = 3000 (1.352) - 4003
      = 4056 - 4003
      = 53 g/L

2. Determine the amount of NaOCI needed to decapsulate the cysts from the formula:

   

   Ratio: For every 2 cysts, 1 g active product is needed.
   e.g., for 5 g cysts:
   

3. Determine the amount of IN NaOH needed:

   Ratio: For 40 ml decapsulation solution, 1.0 ml 1 N NaOH is needed.

4. Determine the amount of seawater to be added:

   From the preceding example (5 g cysts):

   Total volume of decapsulation solution = 100 ml

   Vol. of NaOCI = 47.16 ml

   

   Therefore, volume of seawater = 100 - 49.66 = 50.34 ml

4.2 Steps in decapsulation of Artemia cysts

1. Hydrate the cysts for one hour.

2. Add the decapsulation solution and allow to react for 7–15 minutes. Keep the temperature below 40°C. Add ice if necessary.

3. Wash the cysts very well to remove the excess hypochlorite. Dipping in 0.1N HCI is necessary if decapsulated cysts are stored in the refrigerator for future use.

   Incubate cysts in transparent container (temperature 25–28°C) if newly hatched nauplii are needed.

5. REFERENCES

Bruggeman, E. et al. 1979 Improvements in the decapsulation of Artemia cysts: 309–315. In Cultivation of fish fry and its live food. EMS Spec. Publ. No. 4 (eds.) Styczynska-Jurewicz, E. et al. Institute for Marine Scientific Research Bredene (Belgium), 534pp.

Sorgeloos, P. et al. 1977 Decapsulation of Artemia cysts: a simple technique for the improvement of the use of brine shrimp in aquaculture. Aquaculture 12(4): 311–316