Practical course on counting, isolation and culturing of microalgae

Part II. Algal Isolation and Culturing Techniques

Edina Lengyel, Dr. Nóra Kováts, Prof. Dr. Judit Padisák

University of Pannonia

Department of Limnology

TÁMOP-4.1.1.C-12/1/KONV-2012-0015

Higher educational cooperation for the water sector

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Contents

1. Sampling for isolation ............................................................................................................ 4

1.1. Sampling of phytobenthos ............................................................................................... 4

1.1.1. Stones ....................................................................................................................... 4

1.1.2. Emerged macrophyta ................................................................................................ 5

1.1.3. Epipelon, episammon ............................................................................................... 5

1.2. Sampling of phytoplankton ............................................................................................. 6

2. Isolation and cultivation ......................................................................................................... 7

2.1. Equipment for the isolation and cultivation .................................................................... 7

2.2. Traditional microalgae isolation technique ................................................................... 11

2.2.1. Enrichment Cultures ............................................................................................... 11

2.2.2. Single-Cell Isolation Technique ............................................................................. 11

2.2.3. Agar ........................................................................................................................ 13

2.2.3. Dilution Technique ................................................................................................. 14

2.2.3. Gravity separation method ..................................................................................... 15

2.3. Cultivation of isolated algae .......................................................................................... 15

2.3.1. Batch cultures ......................................................................................................... 15

2.3.2. Continuous Cultures (chemostat) ........................................................................... 16

2.4. Culture media ................................................................................................................ 17

2.4.1. Chemical composition ............................................................................................ 18

2.4.2. Preparation of media .............................................................................................. 21

2.4.3. Recipes of media .................................................................................................... 22

2.5. Other limiting physical chemical factors ...................................................................... 22

2.5.1. Light intensity ........................................................................................................ 22

2.5.2. Temperature ........................................................................................................... 23

2.5.3. pH ........................................................................................................................... 24

3. Application of cultures ......................................................................................................... 25

3.1. Application in agriculture .............................................................................................. 25

3.2. Application for consumption ......................................................................................... 25

3.3. Industrial utilization ...................................................................................................... 25

3.4. Ecophysiological investigation ..................................................................................... 26

4. Sterilization .......................................................................................................................... 27

4.1. Sterilization methods ..................................................................................................... 27

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4.1.1. Heat treatment ........................................................................................................ 27

a) "Flame" method ........................................................................................................ 27

b) "Autoclaving" method .............................................................................................. 28

c) "Dry heat" method .................................................................................................... 29

d) "Pasteurization" method ........................................................................................... 29

e) "Tyndallization" method .......................................................................................... 30

4.1.2. Chemical treatment ................................................................................................ 30

a) Ethanol ..................................................................................................................... 30

b) Bleach ...................................................................................................................... 31

4.1.3. Physical treatment .................................................................................................. 31

a) "Filtration" method ................................................................................................... 31

4.1.4. Electromagnetic wave treatment ............................................................................ 31

a) "UV (ultraviolent radiation)" method ...................................................................... 31

4.2. Sterile techniques .......................................................................................................... 32

4.2.1. Laminar flow hood ................................................................................................. 32

4.2.2. Transferring liquid cell cultures ............................................................................. 33

5. Acknowledges ...................................................................................................................... 36

7. Appendix .............................................................................................................................. 41

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1. Sampling for isolation

The sampling of algae can be carried out from different water bodies: ponds, lakes, channels,

streams, rivers, seas, oceans, wetlands, etc. The sampling techniques depend on those algae

that are required to isolate (phytoplankton or phytobenthos). Materials used for sampling

equipment can include in both cases glass, plastics, tapes, special clothes.

1.1. Sampling of phytobenthos

The sampling must be carried out from the upper region of the water body (10-30 cm). Be

sure that the sampling point is covered by water for 4 weeks (sufficient time is needed for the

benthos to develop). There are several substrate types where the samples can be collected.

1.1.1. Stones

The stones (Fig. 1) should be placed on a tray. The benthos can be removed from the surface

with knife, scalpel or any kind of toothbrush using little amount of water (distilled water, de-

ionized water or from the water body) (Fig. 2). After it, the removed benthos should be

collected into a little vessel. The best vessel is made from glass ensuring the sufficient light

for the microorganisms.

Fig1. The phytobenthos developed on the surface of the stones

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Fig. 2. The process of the sampling phytobenthos

1.1.2. Emerged macrophytes

The substrate can be any kind of macrophytes, most commonly it is reed (Phragmites

australis) (Fig. 3). The sample should be collected from the 10-20 cm of the stem under the

water surface. Cut this part of the plant, place it on a tray and remove the benthos with knife,

scalpel or any kind of toothbrush.

Fig. 3. The phytobenthos developed on macrophytes

1.1.3. Epipelon, episammon

Make sure that only the benthos will be removed instead of the substrate. The sampling can be

carried out with pipettes (Fig. 4-5), sediment sampling equipment (Fig. 6-7) or spatula and

Petri dishes. In the cases of the latter, the Petri dishes should be placed on the sediment that

should be removed with a spatula after it (Ács and Kiss, 2004).

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Fig. 4-5. The sampling of phytobenthos with pipettes

Fig. 6-7. The sampling of phytobenthos with sediment sampling equipment

1.2. Sampling of phytoplankton

Samples should be collected from surface water (1-50 cm), from deep water by a bottle

(different form and size are available) or a water column using a special devices. The water

sample must be concentrated using a plankton net (~ 10 µm pore size) (Fig. 8) or by allow it

to precipitate to get more cell for the isolation process.

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Fig. 8. Plankton net

2. Isolation and cultivation

2.1. Equipment for the isolation and cultivation

Certain equipment is necessary for preparing stock solutions and culture mediums:

- glass wares (etc. Erlenmeyer flasks, reagent bottles, cylinders, Petri dishes, stirring rods,

tubes, beakers, pipettes, etc.) (Fig. 9).

8

Fig. 9. Glass wares for isolation and cultivation

- Plastic wares (e.g. pipettes, micro multiwell plates, Eppendorf tubes)

- Analytical balance

- Magnetic stirrer

- Gauze and cotton wool for the plugs (Fig.10)

Fig. 10. Cotton wool plugs in different sizes

- An autoclave is usually essential for sterilization, as being the most popular sterilization

methods. All of the mentioned materials should always be sterile.

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- Filtration equipment could also be needed for sterilization, if non-heat-resistant medium

(e.g. f/2 medium) is required. A vacuum source or filter syringe, filter holder, membrane

filters are the minimum materials for doing it.

- The chemical ingredients are essential for preparing stock solution, culture mediums (agar

and liquids). There are several companies with many catalos for having these materials.

Before the order, make sure that the ingredients of the chemical constituents are studied,

because many of them could contain levels of trace metals or others, which may disturb

during the experiments or inhibit some species.

- An equipment to get de-ionized water (Fig. 11) for the mediums.

Fig. 11. Smart Purification Pack for Direct-Q 3

- A refrigerator (maybe with a freezer part) is necessary for maintaining stock solutions and

culture media.

- UV lamps for sterilizing the air and workspace.

- The most popular option for manipulating cultures and working with sterile equipments the

laminar flow hood which is also necessary in the laboratory.

- Microscopy for the isolation (Fig. 12). Dissecting and inverted microscopes are the most

widely used. Make sure that the microscopy has good optical lenses and sufficient

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magnification. The best magnification is from 40 up to 80. It can be supplemented with a

micromanipulator (Fig. 12) for make the isolation easier.

Fig. 12. Inverted light microscopy with micromanipulator

- Roller shakers may be required for keeping the cultures on the move (Fig. 13), because some

species could need it. Removable rolls are suitable for shaking of larger tubes.

Fig. 13. Shaker

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2.2. Traditional microalgae isolation technique

Many methods are available for isolating and purification algae. The methods usually depend

on the type of microalgae, type of used medium, source of sample, the goal of using of the

cultures.

2.2.1. Enrichment Cultures

Enrichment cultures are a preliminary step toward the single-cell isolations. A little amount of

culture medium or special nutrient solutions (mainly macronutrients) are added to the samples

that was collected previously, because natural samples are often deficient in one or more

nutrients. In some cases other limiting factors, such as trace metals can be added to the

samples. There are some special species that requires special supply, which could be also

added to the samples (Andersen and Kawachi, 2005):

- seeds for euglenoids

- fruits and vegetables for Oxyrrhis sp.

- soil extract and siliceous for diatoms

- dry grain of rice for mixotrophs

- any substrates for benthic forms, such as Pinnularia sp. (Fig. 14)

Fig. 14. Glass beads were used as substrates for diatoms

On these ways the growth of the required species is stimulated. The culture that consists of

mixed species can be checked time to time, until the cell density or biomass of the required

species increase. After it, one cell isolation technique can be easily applied.

2.2.2. Single-Cell Isolation Technique

The most common isolation method is the single-cell isolation by micropipette that is usually

carried out with Pasteur pipettes or glass capillary (Fig. 15) (Andersen and Kawachi, 2005).

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Fig. 15. Glass capillary for one-cell isolation

These materials can be heated and formed easily in a flame (Fig. 16), but make sure that the

thin extension do not break which could result unusable material. The end of the pipettes

should be smooth and round. Several micropipettes can be made to allow us using always new

ones. But this is not necessary, because the previously prepared pipettes can be reused after

sterilization.

Fig. 16. Preparation of the Pasteur pipettes for one-cell isolation (a: held in a flame; b-c: after

the glass become soft, remove it from the flame and pull it to get a thin tube; d: break the end;

e: unusable end; f: the perfect micropipette) (Soure: Andersen and Kawachi, 2005)

The aim of the one-cell isolation methods is to pick up only one cell from the samples,

deposit the cell without any kind of damage into a sterile droplet (originated mainly from the

applied medium), pick up the cell again and transfer to another droplet (Fig. 17). This

procedure should be repeated until the droplet is free from any contaminations (dust or

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microorganisms). After reaching this status, the cell can be safely placed into the culture

medium.

Fig. 17. Transfer the cell into new droplets (Source: Andersen and Kawachi, 2005)

This process can be easily performed under light microscope and with

micromanipulator. There are lots of options for placing the samples and the sterile droplets

under the microscope: glass or plastic Petri dishes, micro plates or microscope slides which

are transparent. There are two ways for picking up the cells: flexible, latex tube (to get a link

between the mouthpiece and the end of the capillary) or bulb.

2.2.3. Agar

Development of cells on agar plates is also a popular and common method. This method is

preferred mostly by coccoid algae and soil algae. For example, Pelagomonas and Peridinium

species usually do not grow on agar, while Synura, Chlamydomonas species grow well on it.

Most diatom species also like agar as medium contrary to dinoflagellates.

There are several studies about the concentration of agar. The agar should be between

0.8 and 2 % that is the best concentration for growth. Gupta and Agrawal (2007) studied

many physiological features of Nitzschia species on different agars, which also lead to this

conclusion. Hopkins (1969) observed that on dry agar, diatom motility was reduced very

much.

There are various available agars, but ordinary agar consist of agarose and agaropectin

that are contaminated with numerous impurities (Krieg and Gerhardr, 1981), while some of

them have water-soluble lytic agents (Allen and Gorham, 1981). To make plates, agar should

be dissolved in distilled water or mediums. Heat the solution and then cool it to solidify. You

can effuse the hot agar into Petri dishes, but make sure that the agar does not fill up to the ¾

of the dishes. After cooling down, streaking of the cells can be performed. But before it, little

time should be left to let the agar dry, because on its surface can form a condensed layer that

may inhibit the development of isolated colonies. Do not forgive to do all processes in sterile

work area! There are several methods for streaking, which were illustrated in Fig.18.

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Fig. 18. Streaking techniques (Source: László, 1996)

At the first streaking, a little amount of the collected samples should be streaked onto

the agar which leads to the development of mixed cultures. The streaking should be repeated

few times until the culture will consist of only one species. To reach it, every time a clearly

isolated colony (well-defined, unified colony) must be transferred and streaked onto the new

agar plates (Fig.19).

Fig. 19. The steps of streaking

Depending on the goal of the culturing, after getting pure culture, the culture can be grown

further on agar, or it can be transferred into liquid mediums. In this case, make sure that

always only one separated colony is transferred.

2.2.3. Dilution Technique

The dilution technique has been used for many years (Fig. 20), and it is effective for such

organisms that are rather abundant in the sample. The goal of this method is to place only one

cell into a new vessel. For this purpose, test tubes, flasks, multiwell plates can be used

(depending on the operator’s preference) (Throndsen, 1978). The knowledge of the biomass

or cell density of the required species in the samples could make the dilution technique easier.

If the cell density of the species is unknown, the best procedure is to make repeated (5-6)

serial dilutions of 1:10,

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Fig. 20. Dilution technique (Source: Andersen and Kawachi, 2005)

2.2.3. Gravity separation method

This method can be effective, when the goal is the separation of larger and smaller organisms.

There are two potential ways: centrifugation and settling. This method is frequently

performed to concentrate cells. Gentle centrifugation for shorter time let the diatoms and

dinoflagellates settling into the pellet, while the smaller microorganisms can be decanted from

the upper region. Keep always in mind that the centrifugation may cause damage in the cells,

so the time, the speed should be chosen carefully. If it necessary, the process of the

centrifugation can be repeated to get more concentrated sample. The settling is gentler

methods than centrifugation, because the damage of the cells can be avoided, but it need more

times.

2.3. Cultivation of isolated algae

2.3.1. Batch cultures

The batch cultures are a closed system (Fig. 21) in which the microorganism cells can growin

a fixed volume of nutrient culture medium under fixed environmental conditions (e.g.

temperature, pressure, irradiance, and pH) up to a certain density. The culture is started by the

inoculation of the required microalgae into the new medium. The generations of

microorganisms depend on the initial cell density. If the inoculum is less dense, more

generation can be grown before given nutrients are vanished. The batch culture has so many

advantages: low cost, ease of manipulation, volume of media, various potential manipulations.

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Fig. 21. Batch cultures

The growth rate of the microalgae population has three phases: lag phase, exponential

phase and stationer phase. The growth rates can be measured by the following equation

(Wood, 2005):

μ=(LnNt1 - LnNt0)/t1-t0

where N: population size

t: time

To estimate population growth rates, cell number must be counted. Alternatively, another

parameters can be measured, if it can be shown to be linearly correlated with cell number: in

vivo fluorescence, biomass (as dry weight, particulate organic material), optical density, the

concentrations of chlorophyll, protein, carbohydrate and lipid. If the goal is to estimate

population growth rates for diatoms, the use of proxy measures can be complicate. In most

cases, the average size of diatoms decreases because of the asexual mating (Paasche, 1973;

Armbrust and Chisholm, 1992).

2.3.2. Continuous Cultures (chemostat)

In continuous cultures, a fresh supply of medium is added continuously at the same rate. This

technique allows cultures to remain in exponential growth indefinitely. This kind of culture

can be performed in chemostat system (Fig. 22) (Gerhardt and Drew, 1994).

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Fig. 22. Chemostat system

The steady-state concentration of the culture can be determined by the concentration of a

single limiting nutrient (chemostate) or by the selection of a dilution rate (turbidostat).

At steady state the specific growth rate (m) of the population is determined by the dilution rate

(Wood, 2005):

µ = F / V = D

where F is the medium flow rate (L day-1

)

V is the volume of the culture vessel (L)

D is the dilution rate.

This equation assumes that the specific death rate can be ignored.

2.4. Culture media

The distribution of the algae is determined mainly by the chemical and physical features of

the environment. The optimal circumstances of the species are different, so there are various

culture mediums for isolation and cultivation of marine and freshwater species. The basic

ingredients and its concentrations in the media were determined long time ago. Recently,

numerous media exist due to the lots of modifications. In some cases, the modifications are

inevitable, because if just one necessary compound is added or taken away, it can change the

entire medium. For example in Lengyel et al’s study (2015), the applied soil extract was

previously purified with Ba(NO3)2 to eliminate its sulphate content via precipitation. This

addition also changed the applied medium. Basically, the formation and modifications of a

medium can be based on the analyses of the water in the original habitat, the detailed study on

the nutrient requirement of the microorganisms or on the consideration of ecological

parameters (Watanabe, 2005).

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The most common and popular media for cultivation are liquid (Fig. 23) and solidified

mediums due to their advantages: ease of manipulation, volume of media, various potential

manipulations. But some species preferred the agar, as medium, so liquid media are often

mixed with agar providing a solid medium on which these microalgae can grow.

Fig. 23. Autoclaved liquid media

The media consist of natural and/or artificial chemicals that are essential for microalgae to

grow. It includes nutrients, trace elements, vitamins and buffer compounds. Many algae need

special components, as soil extract in the growth media. All ingredients are responsible for

different intracellular biological process and needs.

2.4.1. Chemical composition

Based on the two criteria defined by Esptein (1972), 17 elements are essential that are

generally segregated into two groups: macro- and micronutrients (or trace elements). The

difference between the two categories is just the concentrations. Macronutrients are requested

in higher concentrations (10-2

- 10-4

M), while trace elements are sufficient in low levels (<10-

5 M). Macronutrients involve hydrogen, carbon, oxygen, nitrogen, potassium, calcium,

magnesium, phosphorous, sulfur, magnesium (Hopkins and Hüner, 2004). Silicate can also be

essential, if diatoms or silica-scaled chrysophytas are required to culture. Further beneficial

elements can be sodium, cobalt, selenium, etc. The chlorine, boron, iron, manganese, zinc,

copper, nickel and molybdenum belong to the trace elements (Hopkins and Hüner, 2004).

Nitrogen: Despite the fact that the atmosphere consists of approximately 80 % nitrogen, only

some bacteria and cyanobacteria can utilize it directly in gaseous form. Most algae absorb

nitrogen in inorganic nitrate (NO3-) or ammonium (NH4

+) form. But some studies concluded

that in some cases the algae can utilize also the organic nitrogen compounds, like amino acids.

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This element is essential for the microorganisms, because of its important role in

constitution of many molecules: proteins, enzymes, nucleic acids, hormones, chlorophyll,

vitamins, protoplasm protein. In the absence of nitrogen, the glucid:protid rate, the content of

lipids and carotins will increase in contrary to the chlorophyll content. Additionally, the

decrease of the photosynthetic activity, the formation of the reproductive cells and the cell

size will be expected.

Phosphorous:

Phosphorous is taken up mostly as inorganic (ortho) phosphate, but in some times it can be

adsorbed in organic phosphate (e.g. nucleic acid), too. This element is available in the soil

solution mainly in H3PO4 form, but the pH has a major role in its availability. Nitrogen,

silicate and phosphorus are generally required in a ratio of 16:16:1 (Brzezinski 1985).

According to the Redfield ratio, the chemical composition of the average phytoplankter is

106C:16N:1P (Redfield et al., 1963).

Phosphorous is part of the nucleic acid backbone and has an important role in

intermediary metabolism. It takes part in the constitution of ATP, the nitrate metabolism, the

synthesis and transformation of carbohydrates, the formation of chlorophyll and in cell

division.

Potassium: Potassium is the most abundant cellular ion. This element is frequently deficient in

sandy soils due to its high solubility. Potassium can be provided in several ways: K2CO3,

K2HPO4, KNO3.

This element activates numerous enzymes, mainly in the process of photosynthesis

and respiration. Its absence has effect on the starch and protein synthesis, but it can be

compensated with sodium or rubidium tol certain extent. Potassium increases the permeability

of the plasma membranes. Additionally, this element has an important role in regulating the

osmotic potential of cells. Due to its high mobility, it serves to balance the charge of both

diffusible and non-diffusible anions.

Sulfur: Algae take it up, as the divalent sulfate anion.

Sulfur is an important constituent of proteins (cysteine and methionine), coenzymes

and vitamins (thiamine and biotin) which help the uptake of the divalent ions, while the

coenzymes have important role of the respiration and fatty acid metabolism. Furthermore, the

sulfur has a main role in the electron transfer reaction of photosynthesis (ferredoxin) and the

nitrogen fixation (Hopkins and Hüner, 2004). The sulphate is a competitive inhibitor of

molibdenate uptake (Cole et al, 1986) which has an important role of the NO3-

uptake by

being a component of enzymes involved in its reduction. The absence of sulfur is not a

common problem, because there are various bacteria that are able to oxidize sulfides or

decompose organic sulfur compounds.

Calcium: Depending on the taxon, this element can be macro- or micronutrient. Several algae

have higher amount of calcium in the cell wall which can be precipitate (CaCO3) on it

forming armor around the cell. Calcium is absorbed as the divalent cation.

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Calcium has an important role in the cell division, photosynthesis and the

physiological process of nitrogen fixing. Additionally, it is required for the physical integrity

and normal functioning of membranes. Calcium is implicated as a second messenger in

hormonal and environmental responses. It serves as regulating enzyme activities.

Magnesium: Magnesium is also taken up as the divalent cation. Magnesium can be provided

in several ways: mostly as MgCl2, MgSO4.

It is essential for algae due to the major role in the constitution of chlorophyll and

several enzymes. It is also needed to stabilize ribosome structure. Its deficiency inhibits the

cell division and contributes to its accumulation in the cell.

Chlorine: Generally chloride can be found in every alga, but the marine species contain higher

levels. It can be provided in several ways: mostly as MgCl2, NaCl, FeCl3.

Cl- has two key roles, in the photosynthetic oxygen evolution and on other side in the

charge balance across cellular membranes. Cl- is a counterion to several diffusible cations and

it can maintain electrical neutrality across the cellular membrane, due to its mobility. Cl- is

one of the principal osmotically active solutes in the vacuole, however, at high amount of this

ion, it becomes toxic element (Hopkins and Hüner, 2004). Survival of organisms and their

photosynthetical activity under salinity stress implies effective osmoregulation (Bauld, 1981). But salt

stress increases the accumulation of these toxic Cl- in chloroplasts causing decrease in

photosynthetic electron transport activities (Boyer, 1976). In surplus, chloride contributes to

the synthesis of abnormal chloroplasts, pirenoids and starchs.

Iron: Of all the trace elements, iron is the most essential ion for the algae. Iron can be

absorbed as ferric (Fe3+

) or ferreus (Fe2+

). The more common is ferreus due to it greater

solubility.

Iron has many important roles in the plants. Due to being a part of the catalytic group

for various enzymes (e.g. peroxidase, cytochrome, ferrdoxin, nitrogenase), iron are involved

in photosynthesis, nitrogen fixation, respiration. It contributes to the synthesis of chlorophyll.

In the absence of the iron, a simultaneous loss of chlorophyll and degeneration of chloroplast

structure is expected.

Boron: Boron is usually present in the waters as boric acid or H3BO3.

The role of the boron is not well-known yet contrary to the other trace elements. But

the effects of its deficiency was observed long time ago. Its deficiency results in inhibition in

cell division and elongation in primary and secondary roots in higher plants.

Copper: It is also taken up as the divalent cupric ion. Copper readily forms chelates (e.g.

humic acids). It is easily reduced to the cuprous form (Cu+), which is unstable.

Copper can be found in many oxidase. It is a cofactor for a variety of oxidative

enzymes, as plastocyanin, cytochrome oxidase. At high amount of this ion, it becomes toxic

element similar to the chloride ion.

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Zinc: It is also taken up as the divalent cation. This element is an activator of various

enzymes: ADH (alcohol dehydrogenase) catalyzes the reduction of acetaldehyde to ethanol; in

chloroplast and cytoplasm the CA (carbonic anhydrase) catalyzes the hydration of carbon

dioxide to bicarbonate in plants.

Manganese: Manganese is taken up as the divalent cation. Manganese serves as cofactor for

numerous enzymes, such as decarboxylase, dehydrogenase. Due to these enzymes this trace

element has a role in the respiratory carbon cycle. Manganoprotein takes part of the oxygen-

evolving complex in the chloroplast, where it accumulates charges during the oxidation of

water. Its deficiency slows down the growth of the algae, decreases the photosynthetic activity

and inhibits the liberation of oxygen (Hill reaction).

Molybdenum: In aquatic habitats, manganese can be found as the molibdate ion (MoO42-

).

It is required for the function of oxidase. Because of various enzymes (e.g. dinitrogenase,

nitrate reductase) require it, manganese has a role in the nitrogen metabolism.

Nickel: The role of this trace element is not clear, but it may be related to mobilization of

nitrogen and taken part in urease and hydrogenase enzymes.

2.4.2. Preparation of media

Media usually consist of three components: macronutrients, trace elements, and vitamins. All

ingredients should be prepared as stock solutions (Fig. 24), because direct combinations of

several compounds without any dilution in water may result undesirable precipitation. Stock

solutions must be sterilized, if they contain any substrates. If furry fungi or cloudy bacteria are

appeared, it must be discarded and re-prepared. The stock solution can be stored in

refrigerator.

Fig. 24. Stock solution in autoclavable vessels

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When preparation of stock solution and media are in progress, some point must be

kept in mind:

- Because considerable quantities of ammonia may be lost from the medium through

volatilization during autoclaving, ammonium should be added aseptically to the media after

autoclaving.

- Vitamins are heat-sensible; they should be added to the media after autoclaving.

- Phosphate stock solutions should never be stored in the polyethylene bottles.

- Silicate stock solutions should be stored in non-vitreous material (e.g. polyethylene, or

polycarbonate).

- If the Na2CO3 is required in larger amount, it should be added to the media after autoclaving,

because it readily precipitates.

2.4.3. Recipes of media

As it was already been mentioned, there are numerous cultures and uncountable media. For

example according to World Catalog of Algae (Komagata et al., 1989), there are about 11,000

strains, classified into 3,000 species, that are maintained in 40 culture collections representing

16 countries. The Culture Collection of Algae at Goettingen University (international

acronym SAG) has a huge diversity of about 1600 species of microscopic algae (about 2400

strains). 51 media are currently available there.

In this chapter, the most common and popular media are included. The whole recipes

can be read in Appendix. The following media are used and recommended for the certain

taxon:

- Bacillariophycean /=DIAT/ for diatoms.

- BG11 for Cyanobacteria

- Brackish Water Medium (= 1/2 SWES) for brakish species

- Dunaliella Medium (= Dun.) for Dunaliella species

- Euglena Medium (= Eg "Euglena gracilis") for Euglena species

- Spirulina Medium (=Spirul.) for Spirulina sp

- Volvox Medium (= Vol.) for Volvocales

- Ochromonas Medium (= Ochr.) for Ochromonas species

- Desmidiacean Medium (= MiEB12 "Micrasterias + Erddekokt + Vitamin B12") for

Desmidiales

- Bourelly media for Cyanobacteria

- Modified Bourelly media for Cyanobacteria living in saline lakes

2.5. Other limiting physical chemical factors

2.5.1. Light intensity

The biomass of the phytoplankton is mainly regulated by light intensity. The irradiance has a

major role in the photosynthesis. The relationship can be described with the P-I

(photosynthesis-Irradiance) curves that is not a linear relation (Webb et al., 1974; Platt et al.,

1981; Hill, 1996). Generally, there are two types: non-inhibited or inhibited photosynthesis

(Fig. 25). The photosynthetic activity initially increases with the irradiance, and then it

23

approaches asymptotic maximum (β=0) or it can peak (β>0) reaching a maximal value (Ps or

Pm). The light intensity, where the maximum can be observed, usually is called as

photoadaptation parameter (Ik). The species that can tolerate shading can be described with

lower Ik, in contrary to the light-demanding species having higher values. The initial slope of

the curve (α) is also very informative and useful parameter to characterize this kind of feature

of the algae.

Fig. 25. The characteristic P-I curves (Source: Platt et. al., 1981)

Every species can have optimum irradiance for growth that should be kept in mind

when isolation and cultivation are in progress. For example, Nitzschia frustulum is a good

light competitor similar to many cyanobacteria (10-231 µmol m-2

s-1

) and most green algae

(85-510 µmol m-2

s-1

) (Padisák, 2004; Reynolds, 1988). Similarly, Anabaena minderi and A.

torques as the members of the phytobenthos exhibited the highest growth rates at low

irradiances (75 µmol m-2

s-1

) indicating their adaptation to low light conditions (de Tezanos

Pinto and Litchman, 2010). In most cases, natural light is sufficient to maintain cultures in the

laboratory. But make sure that the cultures should never be exposed to direct sunlight,

because it may cause irreversible photooxidative damage. Based on numerous studies

involving diatoms, photosynthesis is usually inhibited at moderate intensities around 600-

1200 µmol m-2

s-1

, or at higher level (>1200 µmol m-2

s-1

) (Taylor, 1964; Whitney and Darley,

1983). As artificial light source, there are several possibilities. The daylight or cool-white

fluorescent tubes are the most popular ones.

An other important factor is the selection of the light:dark cycle. There are lots of

variations, as 14:10, 12:12: 16:8, etc. It can be adjusted to the natural conditions where the

isolated species lived.

2.5.2. Temperature

Temperature has a major role in many biological processes. Low temperatures may cause

irreversible changes in membrane fluidity (Falkowski and Raven, 1997). On the other side,

the photosynthetic efficiency may be reduced because of denaturation of enzymes at high

24

temperatures (Hopkins and Hüner 2004). The photosystem II and the oxygen evolving

complex inactivate and the fluidity of the thylakoid membranes changes (Falkowski and

Raven 1997; Hopkins and Hüner 2004). Salinity, light and temperature are important

environmental variables influencing species abundance via affecting the photosynthetic

process (Oppenheim, 1991; Underwood et al., 1998; Underwood, 1994). During the isolation

and cultivation, the cultures should be maintained close to the temperature observed in their

natural habitat: for example polar organisms (<10°C); temperate (10-25°C); tropical (>20°C).

Similarly to the light intensity, species have different temperature optima. During the

cultivation, keep it in mind. For example Nitzschia species prefer higher temperature (~30 °C)

(Barker, 1935), while Navicula lanceolata typically occurs in cold waters (Hofmann et al.,

2013). But after the cultivation and experiments, if the goal is just to maintain the culture for

potential utilization in the future, lower temperature should be used. This condition allows us

to slow the biological and physiological processes of the culture down and ensure the survival

of the culture.

For adjust the temperature, several equipments can be used. Permanent temperatures

can be kept constant by a circulating water bath, air conditioner or other incubators.

2.5.3. pH

The control of pH in culture media is important, because algae grow only within narrow pH

range, it prevents the formation of precipitates, and the pH influences the available C source

for photosynthesis. The dissociation of carbon dioxid is pH-dependent (Fig. 26). It is very

important for the algae, because they can utilize only free CO2 or HCO3-. Generally the pH of

natural waters varies between 7 and 8 serving sufficient available C.

Fig. 26. The pH-dependence of CO2, HCO3

- and CO3

2-

The pH level is easy to maintain in the culture media by buffers. It can be evolved with HCl,

bicarbonate, Tris (Tris-hydroxymethyl-aminomethane), or glycylglycine. The media can be

overwhelmed during the process of autoclaving (high temperatures drive CO2 out of the

solution causing a shift in the bicarbonate buffer system and an increase in pH), or in very

dense cultures of microorganisms (enough CO2 is taken up leading similar effect). After the

autoclaving and cooling, the CO2 will re-enter into the solution from the atmosphere. In the

case of very dense culture, the pH/CO2 level can be controlled by air bubbling or having a

25

large surface area of media exposed to the atmosphere. When bubbling should be applied, the

air must be passed through 0.2µm pores-size filter to keep sterile conditions.

3. Application of cultures

3.1. Application in agriculture

Algae have been used in fertilization for a long time ago due to their high nutrient content,

like potassium. Nowadays, cyanobacteria also are utilized for agriculture work, because one

of their groups is capable to N2 fixation that can be essential for the crops. In Japan, the

production of rice increased with 20-22 %, while in India it was approximately 30% after

cyanobacteria-treatment. For fertilization, several methods can be applied: plowing of fresh,

living algae into the soil; after various preparation of the algae (fermentation, cremation);

solution of algae (e.g. in dust form).

3.2. Application for consumption

People, who live at the border of seas (mainly in Asia, Europe) consume marine algae

(Péterfi, 1977). It can be used for soups, salads, bakery, etc. In Japan, Porphyra tenera

(Rhodophyta), Monostroma sp. and Enteromorpha sp. (Chlorophyta) are basic gastronomic

ingredients for soups, meals and fishes. Gloeopeltis sp. (Rhodophyta) Alaria sp., Laminaria

sp., Undaria sp. (Phaeophyta) are used for mainly cooking. In South America, Ulva and

Durvillea sp., while in the Scandinavian area the Porphyra sp., Chondrus sp. and Gelidium

sp. are also popular. In Africa, the people traded with Spirulina pratensis originated from

Lake Chad. For the human consumption, the cultivation of the mentioned algae is carried out

in several countries.

Marine species (mainly macroalgae) have already been used in foraging for long time

ago. At low tide, high biomass of algae can be collected. The harvested algae usually are

mixed with the forage of pets. On this way, the forage was enriched with carotins, vitamins,

amino acids, proteins, minerals, etc.

3.3. Industrial utilization

Algae can be utilized as biofuel that is an alternative to fossil fuel. Several companies and

government agencies are funding efforts to reduce capital and operating costs and make algae

fuel production commercially viable. Like fossil fuel, algae fuel releases CO2 when burnt, but

unlike fossil fuel, algae fuel and other biofuels only release CO2 recently removed from the

atmosphere via photosynthesis as the algae or plant grew. The energy crisis and the world

food crisis have ignited interest in algaculture for making biodiesel and biofuels.

The photobioreactor and fermentor technology are used over the last two decades by

Martek Biosciences Corporation (MacIntyre and Cullen, 2005). Generally, the

photobioreactors are more versatile, expensive, complicated. Martek designed this system to

produce stable isotopically labeled compounds (e.g. carbon, hydrogen, nitrogen). It can be

used to produce pigments, fatty acids, and bioactive molecules. Additionally, in order to

product biomass, photobioreactors have been designed for life support in outer space, removal

of various compounds from water, production of gas vesicles in cyanobacteria, CO2 removal,

hydrogen production, and macroalgal production (Behrens, 2005).

26

Another important application of the algae can be found in the medicine mainly due to

their antibiotic effects. For example, Chaetomorpha okamurai is strongly antibacterial.

Additionally, the algae have higher vitamin content: in Phyllophora nervosa (Rhodophyta)

the B12, in chlorophytas the B1, B

6 and E vitamin, while in Phaeophyta the C vitamin is

significant. But they have more several important features, like nutrients, other assimilation

products, etc.

3.4. Ecophysiological investigation

Microalgae play a major role in the ecology of the planet, they account for about 48 % of the

total primary production (Field et al., 1998) and supporting food webs in all water type.

Salinity, temperature, pH and conductivity have both direct and indirect effects on the

composition and biomass of phytoplankton and phytobenthos by effecting other factors (e.g.

nutrients), controlling cellular processes, causing osmotic and ionic stress, regulating the

photosynthesis and the growth of the species (Hasegawa et al., 2000; Munns, 2002; Sudhir

and Murthy, 2004; Sullivan and Currin, 2000). To understand the underlying processes,

ecophysiological investigations are needed. Due to these studies, we can get helpful

information for example about the optimums, the tolerance, the acclimatization and the

adaptation capabilities of the species (or assemblages). These kinds of investigations can be

carried out in situ and ex situ measurements. In laboratory, the batch cultures and the

chemostat system are the most popular tools for measure growth or photosynthetic activity of

the species. But lots of incubation systems are also developed. One of them is the

photosynthetron (Fig. 27).

Fig. 27. Photosynthetron: a potential incubation system for ecophysiological investigation

27

This incubation system consists of 9 cells with 9 different irradiances possibility. The PAR

(photosynthetically active radiation) usually is provided by daylight fluorescent tubes at both

sides of the photosynthetron. There are many mirrors at the inner walls for multiplying and

uniforming the irradiance. Permanent temperatures can be kept constant by a circulating water

bath.

On molecular level, there are many possibilities to recognize the biological processes

that are underlaying the investigated parameters. For example, we can get effective results and

information about the membrane fluidity, nutrient uptake kinetics, photosynthesis, pigment

synthesis, etc. Furthermore, the cultures can be used to get information about the phylogenetic

relationships due to the lots of genetic (DNA) examinations.

4. Sterilization

Sterilization is a process for remove or kill all microorganisms (virus, bacteria, and other

algae) found on the different equipments and in the applied reagents. This process is very

important and necessary when monoculture is desired. For minimize the biotic contaminations

sterile equipments and sterile techniques are usually combined. Although both of them have

easy procedures and require relative short time, precautions must be taken during and after

them. The sterilized equipment can be easily re-contaminated for example from the air, hands

or the surface of the work area in which there could be lots of spores and microorganisms. In

order to avoid the re-contamination the possibility of these kinds of dangers should be always

kept in mind.

4.1. Sterilization methods

Several sterilization methods and techniques exist, but in this chapter the most general and

common ones will be mentioned.

The methods can be classified in four main groups:

4.1.1. Heat treatment

a) "Flame" method

- general information: direct heat with fire. Mostly, Bunsen burner was used.

- utility: suitable for sterilization of surfaces: inoculating loops and needles, glass

pipettes (Pasteur pipettes), glass Erlenmeyer flask (Fig. 28).

- disadvantage: cannot be used for non-heat-resistant materials: numerous plastic equipments.

28

Fig. 28. Inoculating loops and glass capillaries used for isolation of microorganisms

b) "Autoclaving" method

- general information: 2 atm. steam pressure, 121 °C for 10-60 min. without boiling liquids.

According to the general directive, 10-20 min. should be enough for small volume (e.g. test

tubes of 18 mm diameter), while 1 hour for large volume (>10 liters of liquid). Above 100 °C

all vegetative forms of microorganisms and spores will be killed. The autoclave is the most

popular and effective methods, so there are several available autoclave types in the trade (with

different capacity) (Fig. 29), in this case the best suitable solution (for the given purpose) can

be chosen.

Fig. 29. Front view and top view of autoclave

29

- utility: suitable for liquids, agars, glass, metal and some plastic equipments.

- disadvantage: cannot be used for non-heat-resistant materials (e.g. vitamins). Trace metal

contaminations are possible. The pH might change.

- caution: After autoclaving, because of the steam, the materials could be wet, so dry-heat

procedure is also required. The system is under pressure, so be sure that the caps of the

vessels are ease off and enough free places are left in the flasks for steam and potential

boiling. Furthermore, for labeling vessels autoclave tape is recommended or any kind of

steam-and heat resistant material. Finally, after the sterilization the door should not be opened

until the pressure is zero, and the temperature is under 100 °C. In most cases it is necessary to

get the liquid mediums out of the autoclave and leave it cool to room temperature quickly to

avoid the formation of precipitants.

c) "Dry heat" method

- general information: Sterilization at 150-250 °C for 3-5 hours with hot air. This method

needs higher temperature and longer time then autoclave. 150 °C for 3 hours is the most

acceptance condition. Some autoclave involves this method, too. But several types of drying

ovens are available (Fig. 30).

Fig. 30. Front view of drying oven

- utility: suitable for dry goods, glass and metal equipments.

- disadvantage: cannot be used for non-heat-resistant materials and liquids.

- caution: The materials should be covered (mostly aluminum foil). After the sterilization the

door should not be opened until the temperature is under 60 °C.

d) "Pasteurization" method

- general information: The methods ensures that the liquids are not exposed to higher

temperature (>100 °C). During the method, the liquid is raised to a given temperature (66-80

°C), held for at least 30 min, and then quickly cooled down (<10 °C). (Fig. 31)

30

Fig. 31. The schematic flowchart of the pasteurization

- utility: suitable for liquids with heat-labile components.

- disadvantage: It is not a complete sterilization process, because this method was originally

developed for food items.

e) "Tyndallization" method

- general information: Similar to the pasteurization, but the process is repeated three times

(Fig. 32), which is required to kill cysts.

Fig. 32. The schematic flowchart of the tyndallization

- utility: suitable for liquids with heat-labile components.

- disadvantage: time-consuming.

4.1.2. Chemical treatment

a) Ethanol

- general information: 50-70 % solution.

- utility: for general disinfection. The surface of the workplace or other equipment can be

cleaned with it.

- disadvantage: It is not a complete sterilization process, there are lots of resistant

microorganisms.

31

b) Bleach

- general information: 1-5 mL to 1 L water for several hours. Smaller amount of bleach is

very effective, and most of the microorganisms are killed. For shorter time, more concentrated

solution should be applied.

- utility: for large volume of water for aquaculture.

- disadvantage: It is not a complete sterilization process, there are lots of resistant cysts.

Furthermore, its neutralization is required with usually sodium thiosulfate.

4.1.3. Physical treatment

a) "Filtration" method

- general information: ≤0.2 µm pore size filter should be used. Various filters are available

(e.g. pore size, diameter, composition) (Fig. 33). The best filter is a membrane filter, because

it is autoclavable. It is important to note that there could be micro defects on its surface

leading some microorganism pass through.

Fig. 33. Various filters used during isolation and cultivation

- utility: suitable for liquids with heat-labile components (e.g. vitamins).

- disadvantage: Only small volume could be sterilized in this way. The viruses can pass

through.

4.1.4. Electromagnetic wave treatment

a) "UV (ultraviolent radiation)" method

- general information: 260 nm, at least 5-10 min (Fig. 34). UV radiation is suitable for use in

laboratory contrary to x-rays or gamma rays.

32

Fig. 34. Sterilization with UV

- utility: for sterilizing surfaces (vessels, pipettes, etc.) and working area or little amount of

liquid.

- disadvantage: There are some ultraviolent-sensitive plastics. Due to the fact that UV is

damaging human eyes, the exposure should be avoided.

4.2. Sterile techniques

4.2.1. Laminar flow hood

This is the most popular option for manipulating cultures and working with sterile

equipments. In the best case, the laminar flow (Fig. 35) hood should be placed in a small,

closed room, which is isolated from the laboratory.

Fig. 35. Vertical laminar flow hood

33

In the room and/or in the laminar flow hood should be equipped with UV lamps for pre-

sterilizing the air and the workplace before any kind of manipulations start. It could be

combined with chemical treatments (e.g. ethanol). To minimize the potential contaminations,

the air circulation and any movements should be reduced. Even under the laminar flow hood,

any motion should be slow and the operator should avoid crossing hands, crossing above the

materials, waving materials. Two types of the boxes exist: horizontal and vertical. There are

several supplementary which the hood could be equipped with: electrical plug in, severe gas

supplies, etc. Both of them ensure any kind of dust and potential microorganisms originated

from the air cannot enter the working area. In the hood usually there is a filter that should be

controlled and cleaned periodically. To increase the safety, the person, who work under the

hood can wear latex gloves which could be easily changed more times. The sterility can be

intensified, if the positions of the applied materials are well-chosen which depends on the

person (right- or left-handed) (Fig. 36).

Fig. 36. The right location of the equipments in the laminar flow hood (Soure: Kawachi és

Noël, 2005)

4.2.2. Transferring liquid cell cultures

Various procedures exist for transferring cultures into fresh ones. The following example is a

general process which can be used in algological or also in bacteriological laboratories

(László, 1996; Barker, 1935; Kawachi és Noël, 2005). The example suggests right-hand

person, the existence of laminar hood flow, the use of autoclaved materials and glass Pasteur

pipettes.

1. Clean the materials (e.g automatic pipettes) and the workplace with ethanol or any kind of

disinfectant.

2. Be sure that all vessels are labeled and place the materials in the right positions.

3. Turn on the laminar flow hood and the UV lamp inside the boxes. Turn on the UV lamp in

the room. Hold it at least for 10-15 minutes before starting the manipulations.

4. After the sufficient time, turn on the UV lamps.

34

5. With your left hand pick up the pipette holder, remove its cap with your right hand using

your palm and two outer fingers. Make sure that every process happens near the Bunsen

burner (the best distance is 20 cm) (Fig. 37 a-b).

6. After shaking the holder with your left hand for coming the pipettes out of it (few cm is

enough), catch and remove the pipettes with your right hand (thumb, index and middle

fingers) (Fig. 37 c).

Fig. 37. The process of sterile technique from 5-7 points (source: Kawachi és Noël, 2005)

7. Replace the cap. Be very careful about that the pipette does not touch any surface (Fig.

37d). Place the holder back in its starter positions with your left hand.

8. With your left hand pick up the pipettes carefully (thumb and index finger) at the cotton-

plugged end of the pipette (Fig. 38a). Pick up and insert the bulb on it (Fig. 38b-c).

35

Fig. 38. The process of sterile technique in the 8th points (source: Kawachi és Noël, 2005)

9. At this point the heat treatment (Flame method) of the pipettes is an option, but not

necessary.

10. Hold the pipettes in your right hand, while pick up the culture vessel with your left hand.

11. With your right hand, using your palm and small fingers, remove the cap of the vessel

(Fig. 39a).

12. With your left hand, flame the opening of the vessel and slowly rotate its lip in the flame

(at least 45 degrees) (Fig 39.b).

13. Slowly insert the tip of the pipette into the liquid (Fig. 39c). Slowly draw a cell suspension

into the pipette. Be sure that the liquid level does not reach close to the cotton plug.

14. With you left hand flame again the vessel (Fig. 39d).

15. With your right hand (small and palm fingers), replace the cap of the vessel (Fig. 39e).

Fig. 39. The process of sterile technique from 11 to 15 points (source: Kawachi és Noël,

2005)

16. Move your left hand to the vessel that will be inoculated. The similar procedures should

be done like those described above.

17. Discharge slowly and carefully the cell suspension into the new vessel, and then remove

the pipette.

18. Flame the vessel and replace its cap.

36

20. Bring the pipette close to the discard container (filled with water). With your left hand

(your thumb and index finger), pick up the pipette with at its cotton-plugged end and remove

the bulb with your right hand.

21. With your left hand place the pipette into the container, being sure that the tip is immersed

in the water.

22. Turn off the Bunsen burner, remove all the materials, and cleaned the work area with

ethanol.

23. Switch off the hood, and turn on the UV lamps for at least 10-15 minutes.

5. Acknowledgements

The present study materials is supported by TÁMOP-4.1.1.C-12/1/KONV-2012-0015

37

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40

41

7. Appendix

Table 1. The recipe of Bacillariophycean medium

Ingredients Stock solution (g/100 ml) Nutrient solution (ml)

Ca(NO3)2 x 4H2O 0.2 20

K2HPO4 0.1 10

MgSO4 x 7H2O 0.1 25

Na2CO3 0.1 20

Na2SiO3 x 9H2O 0.1 50

Fe-citrate 0.1 10

Citric acid 0.1 10

Soil extract 30

De-ionized or distilled water 820

B12 vitamin (5 x 10-6

g/l)

Micronutrient solution: 5

ZnSO4 x 7H2O 0.1 1

MnSO4 x 5H2O 0.1 2

H3BO3 0.2 5

Co(NO3)2 x 6H2O 0.02 5

Na2MoO4 x 2H2O 0.02 5

CuSO4 x 5H2O 0.0005 1

FeSO4 x 7H2O 0.7 g

EDTA (titriplex III) 0.8 g

De-ionized or distilled water 981

42

Table 2. The recipe of Brakish medium

Ingredients Stock solution (g/100 ml) Nutrient solution (ml)

NaNO3 7.5 1

NaH2PO4 0.5 1

Na2SiO3 x 9H2O 3 1

Filtered seawater 1000

Vitamin solution 1

vitamin B12 1 mg 1

Biotin (hygroscopic) 0.1 mg 10

thiamin HCl 200 mg

Micronutrient solution: 1

ZnSO4 x 7H2O 2.2 1

MnCl2 x 4H2O 18 1

CoCl2 x 6H2O 1 1

Na2MoO4 x 2H2O 0.63 1

CuSO4 x 5H2O 0.98 1

FeCl3 x 6H2O 3.15 g

Na2EDTA 4.36 g

De-ionized or distilled water 1000 ml

43

Table 3. The recipe of BG 11 medium

Ingredients Stock solution (g/100 ml) Nutrient solution (ml)

NaNO3 15 10

K2HPO4 x 3H2O 0.4 10

MgSO4 x 7H2O 0.75 10

Na2CO3 0.2 10

CaCl2 x 2H2O 0.36 10

Ferric ammonium citrate 0.06 10

Citric acid 0.06 10

EDTA (dinatrium-salt) 0.01 10

De-ionized or distilled water 919

Micronutrient solution: 1

ZnSO4 x 7H2O 287 mg

MnSO4 x H2O 169 mg

H3BO3 61 mg

(NH4)6Mo7O24 x 4H2O 12.5 mg

CuSO4 x 5H2O 2.5 mg

De-ionized or distilled water 1000 ml

44

Table 4. The recipe of Bourelly medium (Krienitz and Wirth, 2006)

Ingredients Stock solution

(g/100 ml)

Nutrient solution

(ml)

KNO3 10 2

K2HPO4 1 4

MgSO4*7H2O 1 3

Ca(NO3)2 1 3

NaHCO3 1.68 10

Fe-EDTA

FeSO4*7H2O 0.7 0.5

EDTA (Triplex III) 0.9 0.5

Biotin 0.0033 10

B12 Vitamin 0.0005 10

Thiamin 0.0005 10

Micronutrient 0.5

MnCl2 x 4H2O 0.0099 g

CoSO4 x 7H2O 0.00028 g

CuSO4 x 5H2O 0.00005 g

ZnSO4 x 7H2O 0.00073 g

H3BO3 0.00031 g

(NH4)6Mo7O24 x 4H2O 0.00018 g

NiSO4 x 6H2O 0.00263 g

NH4VO3 0.00015 g

De-ionized or distilled water 100 ml

45

Table 5. The recipe of modified Bourelly medium

Ingredients Stock solution

(g/100 ml)

Nutrient solution

(ml)

KNO3 10 2

K2HPO4 1 4

MgSO4*7H2O 1 3

Ca(NO3)2 1 3

NaHCO3 1.68 10

Fe-EDTA

FeSO4*7H2O 0.7 0.5

EDTA (Triplex III) 0.9 0.5

Na2CO3 300 mg

NaCl 15000 mg

Biotin 0.0033 10

B12 Vitamin 0.0005 10

Thiamin 0.0005 10

Micronutrient 0.5

MnCl2 x 4H2O 0.0099 g

CoSO4 x 7H2O 0.00028 g

CuSO4 x 5H2O 0.00005 g

ZnSO4 x 7H2O 0.00073 g

H3BO3 0.00031 g

(NH4)6Mo7O24 x 4H2O 0.00018 g

NiSO4 x 6H2O 0.00263 g

NH4VO3 0.00015 g

De-ionized or distilled water 100 ml