PART III
PRACTICAL WORKS

Practica l,8,11
FIELD AND LABORATORY TECHNIQUES IN THE COLLECTION, PRESERVATION, CURATION, AND IDENTIFICATION OF MARINE BENTHIC ALGAE

by

Edna T. Ganzon-Fortes

1. INTRODUCTION

To be able to effectively acquaint one's self with the seaweeds, one must experience the collection of the specimens in the field and their preparation in the laboratory for the herbarium. Only through the practice of, handling and distinguishing the plants as they appear in nature or as, pressed or preserved specimens, can one develop the ease of identifying them. Reading the taxonomic descriptions of, the seaweed species will further enhance one's familiarity with the specimens and most importantly, this will serve as a check to the right identification of the species.

2. COLLECTION OF SEAWEED MATERIALS

The following are the usual supplies and materials necessary for seaweed collection:

-   Plastic bags
-   knife (or any scraping materials)
-   labelling materials (pentel pens, pencils, papers for label),
-   rubber bands
-   face masks and snorkels
-   booties or rubber shoes

The best time for collecting the seaweeds is during the hours of the falling tide. It is best to go to the collection site one to two hours before the time of the low tide as predicted by the tide table. This is to allow the collector considerable amount of time to observe the algae in their natural habitat, to record such observations, and to collect the specimens.

Provided with plastic bags, labelling materials and knife, the collector works by removing the complete plants {including the holdfast) from the substrate and placing them in plastic bags. Labels which include the place and date of collection, kind of substrate or habitat, should then be placed in the respective plastic bags. Delicate -or small materials should be placed in separate plastic bags or vials. Many specimens can be removed from their substrates by the hand but those closely adhering to rocks such as crustose or mat-forming species may be removed with the help of a knife or any scraping blade to secure the complete holdfast. However, those species which may adhere so closely to the rocks can be removed with the rock using a geologist's pick or any similar instrument.

3. PRESERVATION OF SEAWEED MATERIALS

Things needed:

-   5 – 10 percent solution of commercial formalin in sea water
-   pails with cover or large plastic bags
-   labelling materials

All animal components, rocks and other foreign materials should be removed from the collected seaweeds. Five percent solution of commercial formalin in sea water should be prepared. A stronger solution of ten per-cent should be prepared if the specimens to be fixed could not be processed immediately. Before addition of the formalin, water should be drained from the plastic bag. The formalin should then be poured to the seaweed materials inside the plastic bag in amounts just enough to fill the bottom of the bag. Additional formalin may be added if the material is bulky or fleshy. The fumes of the formalin would be enough to fix and preserve the algal materials. All these materials should be properly labelled, with information on the place and date of collection, name of the collector and pertinent observations on the character of the habitat. Such information should be written on a 7 × 12 cm sheets of quality paper or any substitute using indelible ink or pencil.

For transport of the plastic bags containing the preserved algal materials, they should be placed in tall pails with tight covers or in 2–3 layers of good quality large plastic bags (approximately 0.6 × 1.0 m in size) properly tied with rubber band to prevent any leakage of formalin.

4. LABORATORY PREPARATION OF SEAWEED MATERIALS

4.1 Preparation of exsiccatae materials

Things needed:

-   basins
-   galvanized sheet
-   forceps
-   dissecting needle
-   mounting papers of different sizes
-   paper cutter
-   data notebook
-   pencil
-   driers: cheese cloth, blotters, old newspapers
-   ventilators: corrugated aluminum sheet, corrugated cardboards
-   wooden pressers
-   strings

Upon return to the laboratory the preparation of preserved specimens may begin only after; leaving; the materials in the preservative for a few days to allow complete fixation. It would be convenient to work on the individual collecting bags. The specimens contained in each bag should be dumped into a flat-bottomed basin containing fresh water to wash off excess formalin and other foreign materials (i.e. sand, pieces of shells, etc.). The seaweeds would then be sorted out according to species. The sorted species should be placed on wide-mouthed jars or on trays. A collection number should be assigned to one species collected in one area at one time. This number should be recorded in a log notebook together with the information on the name of the species, date and place of collection and other pertinent ecological data. The same number should be written on the mounting sheet carrying the particular algal species.

The sheet of paper to be used for mounting should be cut according to the size of the species and should carry the collection number already assigned to the species to be mounted. The mounting paper should then be placed on top of a flat aluminum sheet and both would be immersed in a basin of clean water. The specimen should be arranged on the mounting paper while underwater to simulate their natural habit. Then the aluminum sheet should be lifted carefully from one side to allow the water to drain off gradually and to leave the specimens spread out and undisturbed. Final arrangement of the specimen may be made when out of the water with the use of forceps and dissecting needles.

The mounting sheet with the specimen should then be placed directly on top of the newspaper which is spread out on the blotter resting on a ventilator. A cheese cloth is placed on top of the specimen. Then, another newspaper and blotter is placed on top of the cheese cloth. A ventilator is further added on top of the pile. The same process is repeated for the rest of the mounted specimens until a sizeable pile is made. This pile should then be stacked between two wooden pressers. Enough pressure should be applied to the pile while trying it tightly.

The whole stack is left to dry in an oven-dryer under temperature of 65–707deg;c for about 24 hours depending on the thickness of the pile, If an oven-dryer is not available, frequent changing of the driers (such as the wet blotters, newspapers and cheese cloth) must be done until drying is complete

The crustose species of algae, on the other hand, should be dried directly in the air and preserved in this dry state in small boxes of suitable size. Articulated, calcareous algae that are so fragile/or so three-dimensional as to suffer badly from pressing, should also be air-dried and kept in boxes. They should, however, preferably be soaked for several days or weeks in a formalin solution containing 10 to 40 percent glycerin before being dried and kept in small boxes, Glycerin retains the flexibility of the genicula and prevents fragmentation.

The dried specimens (mounted or in boxes) should be provided with labels containing the following information: collection number, name of species, place and date of collection, collection and other pertinent ecological data if available. Gum arabic glue or any substitute may be used for those materials which do not stick to the mounting sheet. These exsiccate materials (dried herbarium materials) should be kept in the herbarium.

4.2 Preparation of slides of marine benthic algae

(a) Whole mounts: Microscopic forms of benthic algae, especially the epiphytes, are always mounted wholly. To prepare temporary mounts, the specimen is first washed carefully on a petri dish by changing the fresh water several times until the sand or mud particles are removed. The specimen is then transferred to a clean glass slide. The specimen is first stained with one percent aqueous aniline blue by adding one to two drops of the stain, acidifying it with one drop of one percent HC1 after about a minute, and then washing it with a drop or two of distilled water. Excess acidified stain and water may be blotted off with tissue paper. A drop of glycerin is added to the specimen before putting the cover slip. A nail polish is used to seal the edges of the cover slip.

(b) Cross-section mounts: The sections of the branches may be used for anatomical studies. The freezing microtome, if available, is one of the most useful equipment for preparation of sections. However, free hand sectioning may be easily substituted for the microtome although the sections may not be as good. Skill in making sections can be developed through practice.

For materials which are foliose, good sections may be produced by cutting a piece of the blade, then folding this several times and placing it between two glass slides in such a way that a part of the specimen is exposed beyond one end of the upper slide. The upper slide could then serve as a guide for cutting. New razor blades should be used. The blade may be run through the specimens using the end of the upper slide as a guide. The plane of cutting should be tilted towards the far end of the material in such a way that several cuts could be made before the upper slide is slid back a little to expose more of the materials for further cutting. Many such sections should be made and only the thin ones can be separated from the rest using a dissecting microscope.

For those materials with cylindrical or flattened branches the same technique can be used. Several of the branches should be placed between two slides as discussed above.

The sections are stained with aqueous aniline blue using the same procedure described for the “whole mounts”.

(c) Squash mounts: Materials which are soft and basically filamentous in construction easily lend themselves for this special technique. These usually include those materials which are slightly calcified. These are first decalcified using ten percent HC1 solution right on the slide or watch glass. The excess acid is removed using tissue paper as absorbent material. Drops of distilled water are further added to remove the excess acid. The excess liquid is blotted off using tissue paper.

The same procedure described for the “whole mounts” is conducted for staining the squash mounts.

All slides should be labelled which always include the collection number, name of the species and structures emphasized in the slide.

5. IDENTIFICATION OF ALGAL SPECIES

As mentioned earlier, only through the practice of handling and distinguishing the plants as they appear in nature or as pressed or pre-served specimens, can one develop the ease of identifying them.

For beginners, it is best to familiarize oneself with pressed herbarium materials first, before going to the field to collect fresh materials. In the laboratory, you will be given representative specimens of the green, brown and red algae. Carefully study the morphological differences between the different genera and the taxonomic characters that distinguish each species of the same genus.

A dichotomous key is provided (see Appendix A), for you to be able to know how to use it in the identification of the different algal species. This artificial key is only applicable for the species presented, in the laboratory exercise. To confirm for the right identification of the species, you must read the taxonomic description of the species in the book, “Illustrated Seaweed Flora of Calatagan, Batangas” by Trono and Ganzon-Fortes (1980). If the description in the book fits that particular algal species, you are on the right track.

6. REFERENCES

Dawson, E. Y. 1966 Marine botany, an introduction. Holt Rinehart and Winston, Inc. New York, Chicago, San Francisco, Toronto, London. 37lp.

Trono, G. C. Jr., and E. T. Ganzon-Fortes. 1980 An illustrated seaweed flora of Calatagan, Batangas, Philippines. Filipinas Foundation Inc. and University of the Philippines Marine Sciences Center, Makati and Quezon City, Metro Manila. I15p.

APPENDIX A

DICHOTOMOUS KEY¹

¹ This artificial key is applicable only to the algal materials used in the laboratory exercise for the Training Course on Gracilaria Algae.

Practica 2–5, 9, 10, 12
INVENTORY AND PRODUCTION OF NATURAL SEAWEED STOCKS AT CALATAGAN, BATANGAS

by

Miguel D. Fortes

1. INTRODUCTION

In seaweed research, a considerable proportion of the ecological work is focused on the description of vegetation. The purpose of such a description is to enable people other than the worker to build a mental picture of the study area, allowing for comparison and classification of the different components of the vegetation. But before any detailed work can be started, it is important to know what algal species are present (floristic composition), the degree of abundance of each species, how they are distributed in time (seasonality) and space (zonation), and how they are functionally related to the other biotic components (community dynamics). However, a true “picture” of these structural and functional aspects of the seaweed vegetation can be made only with the use of efficient and effective methods for collecting, interpreting and presenting the data in a quantitative manner. Other information such as the areal topography and values obtained for environmental factors characteristic of the area may be essential for a realistic account of the place to be investigated.

For practical purposes, the participants will be divided into groups, each group to perform independently simple sampling techniques and growth rate measurements of important seaweed species in the study area,

2. A TAXONOMIC SURVEY OF THE SEAWEED FLORA OF BALONGBATO, CALATAGAN (Practicum 2)

Field Materials (per group) :

Writing slates, with pencil and ruler (4)
Plastic bags (small, 20 pcs.; big, 4 pcs.)
Pentel pens (4)
5 percent formalin solution (1 bottle) Rubber band (20 pcs.)

2.1 Methodology

Four different stations (Stations 1–4) will be surveyed to assess the component seaweed flora. At each station, each member of the group will collect at most three (3) representative samples of each species seen. Place the algae (free of adhering debris) inside plastic bags. Four out the water and seal the bags tightly. Note and record on the writing slates the dominant physical features of the habitat particularly in terms of the nature of the substratum, water depth and apparent visibility and wind and current directions. With pentel pens, label the plastic bags following the format given below:

Group No.:                                                                  
Station No. and Sites:                                                
Collector:                                                                    
Dates:                                                                         

Pickle the collected materials with small amount of 5 percent formalin - seawater solution. Place the collections inside the bigger plastic bag, label the bag as above, and store them in a cool, safe place. It is the responsibility of the group leader to assign who the one in-charge of the collections and recorded data will be. This will facilitate ease and convenience in the later treatment and collation of all data.

Taxonomic identification and curation of the collected specimens will be done in the laboratory as scheduled.

3. SIMPLE RANDOM SAMPLING: A STRUCTURAL ANALYSIS OF THE MACROBENTHIC VEGETATION SOUTH OF ALVAREZ FARM, CALATAGAN (Practicum 3)

Field Materials (per group);

0.25 m² metal quadrant (1)
Marker with float (1)
Writing slate with pen and ruler (4)
Plastic bags (small, 20 pcs.; big, 1 pc.)
Pentel pens (4)
5 percent formalin solution (1 bottle)
Data sheet 1 (10 pcs.)

3.1 Methodology (Practicum 3)

Within a previously delimited portion of the reef flat, select randomly, by means of the “closed-eyes-turn-around or CETA” method, the exact position of your specific plot. Walk towards the marker but be careful not to disturb too much the bottom sediments as this will make the visibility of the water poor (also making your ocular observation and collection difficult). Horizontally, lay down the quadrat frame with the marker exactly at its center. With your face masks on, observe carefully the delimited portion of the vegetation. YOU NEED NOT SWIM. Collect, just outside your plot, representative samples of each plant species (seaweed and seagrass) seen inside the quadrat. Place the. materials separately inside plastic bags and number the bags (if you can identify the species, label! the bags using the name of the species). Seal the bags tightly. They will serve as samples to facilitate tentative identification of the corresponding species present inside your plot. Get the following parameters:

1. Density - For each of the individually growing, non-tcreeping species, count the total number of individuals inside the 0.25 m² quadrat. DO NOT CONSIDER THE SMALLER QUADRAT SQUARES. Enter your counts in the table previously prepared on the writing slate (see suggested format, DATA SHEET 2). For species which are either too densely aggregated or which creep, a good measure of abundance is;

2. Frequency - Without moving the quadrat, count the number of smaller squares inside the frame in which a species occurs. Do the same for all the other species. (Which frequency type are you going to use - shoot or rooted?). Enter the results in the same table as for density.

3. Coyer - Estimate carefully the proportion of the substratum occupied by each species (not by each individual of a species) within the 0.25 m² plot. Be guided by the following scale (Domin scale¹ :

1 For practical purposes, this scale is to be copied on the slate.

Enter your ratings for each species in the appropriate column in the same table.

Do procedures (a) - (c) using two other randomly chosen quadrats (plots) within the study area. Enter your findings as before and be extra careful not to mix your data. Immediately after the fieldwork, transfer all recorded data on the data sheets provided.

Add sufficient amount of 5 percent formalin-seawater solution to the collected samples. Seal the bags tightly. Check the labels. Place all materials inside the big plastic bag and label the bag as before.

3.2 Computations

From the data obtained, compute for the following:

(a) Density (D):

	where Di	=	density of species i
	ni	=	total number of individuals of species i
	A	=	total area sampled (m2)

(b) Relative species density (RD) :

	where RDi	=	relative density of species
	Di	=	density of species i
	TD	=	sum of the densities of all the species

(c) Frequency (f) :

	where fi	=	frequency of species i
	ji	=	number of smaller quadrats in which species i occurred
	k	=	total number of smaller quadrats taken

(d) Relative Frequency (Rf) :

	where Rfi	=	relative frequency of species i
	fi	=	frequency of species i
	Tf	=	sum total of the frequencies of all the species

(e) Coverage (C) :

	where Ci	=	coverage of species i
	ai	=	the total area covered by species I
	A	=	total habitat area sampled

(f) Relative coverage (RC) :

	where RC1	=	relative coverage of species i
	Ci	=	coverage of species i
	TC	=	total of the coverage of all the species

(g) Importance Value (IV):

IV_i = RD_i + Rf_i + RC₁

3.3 Laboratory processing and collation of gathered data (Practica 10 and 12)

After the laboratory identification of the seaweeds present in the quadrate (plots)), enter the data in DATA SHEET (1). Based on the Group Data, answer individually ,(not as a group) the following guide questions (please write your answers only in the spaces provided below each question):

1. How many species represent each of the 4 major divisions of seaweeds?

   1. Cyanophyta (blue-green algae)                         
   2. Chlorophyta (green algae)                                 
   3. Phaeophyta (brown algae)                                
   4. Rhodophyta (red algae)                                     

2. Which of the division represented had the highest (and lowest) “importance value”? Which species? Give probable reasons for the observed differences.

3. What is the agronomic or maricultural implication of your findings?

4. What could be the ecological and methodological implication(s) of the differences in the group results?

5. How is field exercise (2) related to what you are doing in you respective country/institution?

6. Suggest or comment on the methodology we used.

DATA SHEET 1

Group No,                           Quadrat No.                      
Habitat Description                                                       
Station No,                     Site                                        
Total Area Sampled           Total No. of Plots              

4. SYSTEMATIC SAMPLING: BIOMASS AND ZONATION OF THE MACROBENTHIC VEGETATION AT BUROT, CALATAGAN (Practicum 4)

Field Materials (per group):

0.25 m² metal quadrat (1)
100 m transect line, calibrated (1)
2 m sighting poles, calibrated (2)
plastic bags (small. 50 PCS., big, 2 pcs.)
pentel pens (4)
5 percent formalin-seawater solution (1 bottle)
plastic basins (2)
1 m wooden pegs(3)
writing slates (4)

4.1 Methodology

Two members of the group will initially lay the 100 m transect line straight and perpendicular to the shoreline, with the 0 m end pegged near the watetline. At 5 m intervals (start from 0 m), lay down the metal quadrat with its left side coinciding with the transect line. With your face masks on, observe the enclosed “sample” of the Vegetation. YOU NEED NOT SWIM; JUST BEND AND LOOK DOWN: Collect, by simply “Weeding” all the plant species (seaweeds and seagrasses) present inside the plot. This is quadrat 1 (Q1). Place them inside plastic bags and label the bags following the format given below:

Group No.            
Q              Interval        
Date                              

Pour the water out and seal the bags tightly,

Do the same procedure for 9 other quadrats. Be very careful not to mix and confuse the collections. On shore, pickle the collected materials with small amount of 5 percent formalin-seawater solution and reseal tightly. Inspect the labels and place all the collections inside the bigger bag and label as before.

The “sighting” method is employed to determine the contour of the bottom. In a group, the members will be assigned as sighter recorder, pole holder and coordinator. (Details of the method will be discussed and demonstrated in the laboratory). At 2 m intervals along the transect line. get the following data: distance from 0 point (m), pole reading (cm), water depth (cm) and time (hrs on min). Record the results in the table prepared on the writing slate. (If circumstances permit, the collected samples may be sorted out and pre-dried in the vicinity of the working site. Final processing of the materials will be done in the laboratory.

4.2 Biomass estimate (Practica 9 and 10)

Follow the flow-diagram below to estimate the dry biomass of the collected seaweed materials:

Enter all the results in DATA SHEET 3¹

¹ TDW is the Total Dry Weight, i.e., for species with fresh weights greater than 20 gm, TDW = (Dry Weight of Subsample) × (Total Fresh Weight/20).

4.3 Zonation and bottom contour (Practica 9 and 10)

From the data, note the spatial distribution of. the seaweed species from the first to the last quadrat along your transect line.

To determine the bottom contour (profile), graph the values obtained from the “sighting” method. Follow the figure format; given below:

4.4 Results and discussion (Practicum 12)

In the spaces provided, individually answer the following guide questions:

1. Which seaweed species has the highest (and lowest) biomass in the 4rea your group studied?

2. Can you detect a change in the species composition (i.e., presence of absence of species) and biomass of the seaweeds from the first to the last quadrat? What could be the probable reasons for your observation?

3. Try to relate your answer in (2) with the contour of the substratum in your area.

4. What is the agronomic or maricultural implication of your Findings?

5. How is field Exercise 3 related to the specific fishery or seaweed problem in your country/institution?

6. Suggest or comment on the methodology used.

DATA SHEET 2

Group No,                           Quadrat No.                      
Habitat Description                                                       
Station No,                     Site                                        
Total Area Sampled           Total No. of Plots              

5. GROWTH RATE STUDIES ON GRACILARIA AND EUCHEUMA AT CALATAGAN, BATANGAS (Practicum 5)

Field Materials and Instruments (per group) :

5.1 Methodology

From the seed(ling) stock of Gracilaria, two members of each group will get test “plants” by careful breaking of healthy thalli. Get the most practical and common sample weight and length for all 35 “branch portion”. Be careful not to cut the growing tips of the samples. Record and immerse the pieces in seawater shaded from the direct heat of the sun. Prepare a corresponding number of tags for the plants.

From the seedstock of Eucheuma, the same group members will weigh 35, 100 gm fresh portions (samples) of healthy thalli. Record, Immediately after weighing, tag each test plant and tie securely with a “tie-tie”. If the culture lines are not yet ready, immerse them in seawater as above.

Within a predetermined site, the other two members from each group will lay out the two 10 m lines (abaca rope and monofilament), pegged at the ends, parallel and 0.5 m away from each other, The distance of the lines from the bottom should be estimated in such a way that they will not be exposed to air during low tide nor will they be covered by the shifting sandy-silty substratum. (The height of the Eucheuma nets and plants nearby would be a measure of the desired distance).

Once the lines are set, and the seedlings and their tags are ready, tie the test plants carefully along the lines. Tie those of Eucheuma at the “notches” pre-made for the purpose (30 cm interval). For the Gracilaria seedlings, partly “unwind” the strands of the rope at the marked points (also 30 cm interval) and insert carefully the seedlings. Gently release your grip from the rope so that the strands will just hold firmly, not obliterate, the test plants. Tag the plants and record, including all information you think are relevant to this portion of the exercise.

For the environmental variables, the two group members who set the culture lines will lay down two pairs of clod cards (preweighed), one at each end of the line. In addition, tie securely a maximum-minimum thermometer (pre-set), alongside the lower portion of a peg which supports your line. The cards and the thermometer are to be retrived 30 minutes before departure time. Record both the time of immersion (T_i) and retrieval (T_r). Record the results upon recovery of the materials.

Using a strip of pH paper, determine the pH of the surface water at the region near the ends of the lines. Similarly, determine the salinity, using the refractometer, at those points. Record your results.

5.2 Measurement of growth rates (Practicum 5, continuation)

Field Materials and Instruments (per group):

Writing slates, with pencil and ruler (4)
Plastic basin (1)
Aluminum plates (10)
Tissue paper (1 roll)
Platform balance (2 to be shared)
Data Sheet (3) (5)

5.2.1 Methodology

Two members from each group will retrieve the abaca rope holding the Gracilaria test plants. Gently place the rope in the basin 3/4 full of seawater. Using the same balance used during the initial phase of the study, weigh consecutively each of the samples, noting particularly the number of each written on the tags. Record the resulting weights. Measure the length along the long axes of the plants. Record.

Carefully retrieve the Eucheuma test plants from the monofilament line. Place gently the plants, with their tags but without the “tie-ties”, in a basin with seawater. Weigh each plant and record the results.

Checklist all materials and instruments after use.

5.3 Discussion

Based on the data obtained (See Data Sheet 3 ), answer individually (not as a group) each of the guide questions (Plase write your answers only in the spaces provided):

1. How do the growth rates of Gracilaria and Eucheuma compare? Cite probable reason(s) for your answer.

2. What do the standard deviation(s) and standard error (SE) you obtained for the growth rates of the two species suggest?

3. What is the agronomic or maricultural implication of your findings?

4. What could be the ecological implication(s) of the differences/similarities in the group results?

5. How is field exercise (4) related to specific fishery or sea weed problem in your country/institution?

6. For practical purposes, please comment or suggest on the methodology used.

DATA SHEET (3)

Group No.                 Growth Period: From           To         (                 Days)
Habitat Description                                                                                        
Station No.               Site                                         Date                                
Remarks                                                                                                         

Practica 6 and 7
DEMONSTRATION ON THE POND CULTURE OF SEAWEEDS; MONITORING OF GROWTH RATES AND ECOLOGICAL PARAMETERS

by

Gavind C. Trono. Jr.

1. POND CULTURE OF CAULERPA

This exercise will be done at the Experimental Caulerpa pond at Balong Bato, Calatagan, Batangas. Each group shall obtain about five kg of C. lentillifera seedstocks. The modified broadcast method will be applied for this purpose. Select, from the seedstock a handful of planting materials. Arrange these in a bundle and mould the base of the bundle with a fist-size clayey soil or mud. Broadcast or distribute the moulded seedstock evenly in the pond.

In addition each grup shall plant three one-meter plots to monitor the biomass production in the pond using the following treatments:

Plot 1: plant with 0.10 kg of seedstock
Plot 2: plant with 0.25 kg of seedstock
Plot 3: plant with 0.50 kg of seedstock

Each plot must be marked by enclosing it with a string. The biomass in each plot will be harvested during the follow-up trip and the amount of bio-mass compared.

Record, the salinity, pH and water temperature in the pond. Write a brief report on the results of this field exercise.

2. POND CULTURE OF GRACILARIA

This exercise will be done at the Gracilaria experimental pond in Palo Bandera, Calatagan, Batangas. Seedstocks of G. verrucosa and Gracilaria sp from Bacoor Bay, Cavite will be used for this purpose.

Each group of five will obtain about 15 kg of Gracilaria seedstock. Divide this material into five and ten kilogram lots, Cut the Gracilaria into 10–15 cm pieces. Each group shall be assigned two ponds to seed using the following treatments:

Broadcast the seedstocks evenly in the ponds. Apply the fertilizer three days after planting.

Each group shall monitor the growth rate of Gracilaria using the “rope method” used in India. Untwist the rope and insert cuttings of Gracilaria at 10 cm intervals. Then trim the seedling to uniform length. Record the initial length of the seedlings. Plant a seeded rope in each experimental pond tying the two ends of the rope to wooden stakes as support. The seeded rope must be 15 cm to 20 cm from the bottom of the pond.

Record the ecological parameters in the ponds: salinity, pH and water temperature.

During the follow-up trip, harvest all the Gracilaria in each pond. Record the biomass produced in each pond. Measure the increase in length of the plants attached to the rope. Compute for the increase in biomass and increase in length of plants in each pond. Determine the daily growth rates of the plants. Write a short report on the results of the exercise using data gathered from the six experimental ponds.

Practicum 13
EXTRACTION OF AGAR

by

Nemesio Montano and Reynita L. Veroy ¹

¹ Research Associates, Marine Sciences Center, University of the Philippines, Diliman, Quezon City

1. MATERIALS AND APPARATUS

Bleached seaweed
Waterbath
Pots
Shallow dish pans
Cheese cloth
Oven
Stirrers
Beakers
Mill

2. PROCEDURE

A known amount of dry bleached seaweed is washed in freshwater to remove the foreign matter. The clean weed is cut into pieces and heated with 700 ml water in a boiling water bath for two hours with occasional stirring. The thick paste is filtered through a clean cheese cloth. The filtrate is poured into shallow dish pans and allowed to freeze overnight. The frozen gel is thawed the next day, and washed with cold water. The final product is dried in a 60°C oven or sundried and then ground to a powder of the desired mesh.

Practicum 14
ANALYTICAL METHODS: DETERMINATION OF MOISTURE CONTENT AND PHYSICAL PROPERTIES

by

Reynita L. Veroy and Nemesio Montano

1. DETERMINATION OF MOISTURE, FOREIGN MATTER CONTENT OF RAW SEAWEED

1.1 Moisture content

Moisture content is determined using a moisture balance, A known amount of sample is exposed to the IR lamp of the balance at 60–70°C for 30 minutes. Moisture content is read directly.

1.2 Foreign matter content of raw seaweed

Fifty to one hundred grams of seaweed sample is washed very thoroughly and dried overnight in an oven at 60°C. The difference in weight before and after washing and drying represents moisture and foreign matter.

2. GEL STRENGTH MEASUREMENT

Materials and apparatus: 3.6 g sample, top-loading balance, beakers, thermometer, water bath, heater, gel tester

Procedure:

Prepare a 2% solution of agar by dissolving a 3.6 g sample in 140 g distilled water contained in a tared 250 ml beaker and heat in a boiling water bath. Add more hot distilled water until the solution weighs 180 g. Reheat the solution, then let it cool. Allow the solution to set overnight. Transfer the gel formed to a crystallizing dish and measure its gel strength by means of a gel tester.

3. VISCOSlTY MEASUREMENT

Materials and apparatus: 3.75 g sample, top-loading balance, beakers, thermometers, viscometer, heater, water bath

Procedure:

Prepare a 1.5 percent solution of the agar by dissolving a 3.75 g sample in 200 g of hot distilled water. When everything has dissolved, add more boiling distilled water until the weight of the solution is 250 g. Transfer the solution to an electrolytic beaker and determine the viscosity of this solution at 70–75° with a viscometer. (Keep the 1.5 percent agar solution for the determination of gelling and melting temperature).

4. DETERMINATION OF GELLING TEMPERATURE AND MELTING TEMPERATURE

Materials and apparatus: 1.5 percent agar solution, glass beads (2 mm in diameter), lead shots, test tubes (20 × 125 mm), thermometer, water bath, heater, clamps

Gelling temperature :

Procedure:

Add 25 ml of 1.5 percent agar solution (70–75°C) into a 20 × 125 mm test tube. Allow the solution to cool at a rate of 2°C/min with a thermometer immersed in the solution. Introduce glass beads (2 mm in diameter) at the surface of the solution at intervals. Note the temperature when the beads fail to sink. (Keep the set-up for the determination of melting temperature).

Melting temperature:

Heat in a water bath containing the gel (with lead shot on the surface) in such a way as to give a 1° rise in temperature/minute. Note the temperature at which the lead shot sinks.

Praeticum 15
SPECTROPHOTOMETRIC METHODS AND SULFATE DETERMINATION

by

Reynita L. Veroy and Nemesio Montano

1. FLUORIMETRIC DETERMINATION OF SULFATE

Reagents %

1. Acridine orange : 1.54 × 10^-4 M stock solution (23.2 mg/500 ml distilled H₂O)

2. Agar solutions : 50 mg/50 ml distilled H₂O stock solution;
   10 ml of stock solution diluted to 100 ml

Apparatus and equipment:

Fluorescence spectrophotometer, cuvettes, pipettes

Procedure;

To an aliquot of 50 ml 1.54 × 10^-4 M acridine orange, add 0.5 ml of the agar working solution and make up the mixture to 50 ml. Prepare several of such solutions using increasing amounts of the seaweed extract and measure the fluorescence intensity of each solution at 540 nm (excitation wavelength = 405 nm).

In a graphing paper, plot the relative Fluorescence Intensity (%) versus the number of ml agar solution added. From the graph, compute the relative equivalent mass (REM) per anionic site for each sample¹.

¹ For computations, refer to: E.C. Laserna, G.J.B. Cajipe, R.L. Veroy and A.H. Luistro, “Spectrofluorimetric assay of carrageenan and agar from Philippine seaweeds”, Kalikasan, Phil. J. Biol. 7; 110–116 (1978).

2. IR SPECTRA OF AGAR

Materials and apparatus : Liquid mercury, 120 mg of agar sample, 30 ml low-form porcelain crucibles, pipettes, vacuum oven, IR Spectrophotometer

Procedure;

Dissolve 120 mg of the sample in 60 ml of boiling distilled water. In a 30 ml low-form porcelain crucible, slowly pour 5 ml of clean Hg. Then add 5–7 ml of the prepared solution. Evaporate off the water in a vacuum oven at 40°C. When the film is completely dry, pour off the Hg and mount the film on a piece of cardboard with a 30 mm × 12 mm hole. Record the spectrum of the film on an IR Spectrophotometer.

Practica 16 and 17
POST HARVEST HANDLING OF SEAWEEDS

by Gavino C. Trono, Jr.

Record the length of time required in drying. Determine the moisture content of each treatment. How much of the dry weight of the unwashed and unsorted lots is represented by foreign materials compared to the washed and sorted materials?

Which of the treatments produced the best results? Write a short report.