1417cale100Microalgal culturing
Practical notes from the culturing short course held in Caen, 10-12 Feb 1999.
Ian Probert & Christine Klaas
General principles
A culture can be defined as an artificial environment in which the algae grow. In theory, culture conditions should resemble the alga’s natural environment as far as possible; in reality many significant differences exist, most of which are deliberately imposed.
A culture has three distinct components: a) a culture medium contained in a suitable vessel; and b) the algal cells growing in the medium; and c) air, to allow exchange of carbon dioxide between medium and atmosphere.
Extensive measures must be taken to keep pure unialgal cultures chemically and biologically clean. Chemical contamination may have unquantifiable, often deleterious, and therefore undesirable effects on algal growth. Biological contamination of pure algal cultures by other eukaryotes and prokaryotic organisms in most cases invalidates experimental work, and may lead to the extinction of the desired algal species in culture through out-competition or grazing. In practise, it is very difficult to obtain bacteria-free (axenic) cultures, and although measures should be taken to minimize bacterial numbers, a degree of bacterial contamination is often acceptable.
For an entirely autotrophic alga, all that is needed for growth is light + CO2 + H2O + nutrients + trace elements. By means of photosynthesis the alga will be able to synthesize all the biochemical compounds necessary for growth. Only a minority of alga seem, however, to be entirely autotrophic; many are unable to synthesize certain biochemical compounds (certain vitamins, for example) and will require these to be present in the medium. This condition is known as auxotrophy.
Based on their growth characteristics, two kinds of cultures can be defined. 
•	In limited volume (batch) cultures, resources are finite. When the resources present in the culture medium are abundant, growth occurs according to a sigmoid curve, but once the resources have been utilised by the cells, the cultures die unless supplied with new medium. In practise this is done by subculturing, i.e. transferring a small volume of existing culture to a large volume of fresh culture medium at regular intervals. 
•	In continuous cultures, resources are potentially infinite: cultures are maintained at a chosen point on the growth curve by the regulated addition of fresh culture medium. In practise, a volume of fresh culture medium is added automatically at a rate proportional to the growth rate of the alga, while an equal volume of culture is removed.
Physical parameters
Temperature
The temperature at which cultures are maintained should ideally be as close as possible to the temperature at which the organisms were collected; polar organisms (<10°C); temperate (10-25°C); tropical (>20°C). An intermediate value of 18-20°C is most often employed. Temperature controlled incubators usually use constant temperature (transfers to different temperatures should be conducted in steps of 2°C/week), although some models permit temperature cycling. In temperate regions ambient room temperature is generally acceptable for culturing purposes.
Light
Natural light is usually sufficient to maintain cultures in the laboratory. Cultures should never be exposed to direct sunlight (which may cause photopigment damage), and should therefore be placed next to a north-facing window (in the northern hemisphere). 
Artificial lighting by fluorescent bulbs is often employed for culture maintainance and experimental purposes. Light intensity should range between 0.2-50% of full daylight (= 1660 µE/s/m2), with 5-10% (c. 80-160µE/s/m2) most often employed. Light quality (spectrum) depends on type of bulb used (see manufacturers technical data), the most common types being ‘cool white’ or ‘daylight’ bulbs. Light intensity and quality can be manipulated with filters. Many microalgal species do not grow well under constant illumination, and hence a light/dark (LD) cycle is used (maximum 16:8 LD, usually 14:10 or 12:12).
Mixing
Mixing of microalgal cultures may be necessary under certain circumstances: when cells must be kept in suspension in order to grow (particularly important for heterotrophic dinoflagellates); in concentrated cultures to prevent nutrient limitation effects due to stacking of cells and to increase gas diffusion. It should be noted that in the ocean cells seldom experience turbulence, and hence mixing should be gentle. The following methods may be used: bubbling with air (may damage cells); plankton wheel or roller table (about 1 rpm); gentle manual swirling. Most cultures do well without mixing, particularly when not too concentrated, but when possible gentle manual swirling (once each day) is recomended.
Types of culture vessel
Culture vessels should have the following properties: non toxic (chemically inert); reasonably transparent to light; easily cleaned and sterilized; provide a large surface to volume ratio (depending on organism).
Certain materials which could potentially be used for culture vessels may leach chemicals which have a deleterious effect on algal growth into the medium. The use of chemically inert materials is particularly important when culturing oceanic plankton and during isolation. A list of materials which should be safe, inhibitory, or toxic to algal cultues is given by Stein (1973). Recommended materials for culture vessels and media preparation include:
•	Teflon (very expensive, used for media preparation);
•	Polycarbonate (expensive and becomes cloudy and cracks with repeated autoclaving);
•	Polystyrene (cheaper alternative to Teflon and polycarbonate, not autoclavable);
•	Borosilicate glass (has been shown to inhibit growth of some species).
Materials which should generally be avoided during microalgal culturing include all types of rubber, and PVC. The following types of culture vessel are most commonly used for microalgal culturing (but may not prove suitable for all species):
•	Erlenmeyer flasks (glass or polycarbonate) with cotton, glass, polypropylene, or metal covers.
•	tubes (glass or polycarbonate) with cotton, glass, polypropylene or metal caps.
•	polystyrene tissue culture flasks, purchased as single-use sterile units (eg. Iwaki, Nunc, Corning).
Cleanling/sterilization of culture materials
Cleaning procedure
•	scrub (abrasive brushes not appropriate for most plastics) and soak with warm detergent (not domestic detergents, which leave a residual film on cultureware – use laboratory detergent such as phosphate-free Decon);
•	rinse extensively with tap water;
•	soak in 10% HCl for 1 day-1 week (not routinely necessary, but particularly important for new glass and polycarbonate material);
•	rinse extensively with distilled and finally bidistilled water;
•	leave inverted to dry in a clean, dust-free place.
Sterilization
Sterilization can be defined as a process which ensures total inactivation of microbial life (NB. not the same as desinfection, which is defined as an arbitrary reduction of bacterial numbers). The primary purpose of sterilization is to prevent contamination by unwanted organisms, but it may also serve to eliminate unwanted chemicals. There are several sterilization methods and the choice depends on the purpose and material used:
•	Gas  (ethylene oxide, often used for disposable plastic material);
•	Moist heat
Autoclave or pressure cooker (1-2 Bar, 121°C, in pure saturated steam).
For sterility the steam must penetrate the material (wrap in material that allows access of steam - kraft paper or aluminium). Autoclave steam may introduce chemical contaminants; glass and polycarbonate vessels should be autoclaved containing a small amount of bidistilled water which is poured out (thus diluting contaminants) under sterile conditions immediately prior to use. Never close vessels (risk of implosion); use cotton wool bungs, or leave screw caps slightly open.
•	Dry heat 
Oven (at least 2 hours at 160°C). Cover neck of vessel with aluminium foil to maintain sterility on removal from oven. 
Flame (heating to incandescence for metal) 
Problems: few materials stand high temperatures (glass, teflon, silicone, metal, cotton). 
•	Radiation
X and g - radiation (industrial applications for disposable plastics that cannot stand more then 60°C, not always effective)
UV radiation 240-280 nm (not often used for culture materials).
 Transfer protocol 
The following procedures should always be used when preparing media or transferring cultures:
•	work in clean place or preferably in a laminar flow cabinet (cabinet must be turned on at least 30 minutes before transfer; if equipped with UV lamps, leave on overnight prior to use).
•	clean working surface with 70% alcohol (ethanol/isopropanol) prior to and after use.
•	clean hands with disinfecting soap and rinse with 70% alcohol prior to all operations. 
•	when not using a laminar flow cabinet (and to be safe even when using a cabinet), sterilise (flame) the neck of vessel of origin before and after transfer (not possible with some plastic vessels, which must, therefore, be opened in a laminar flow cabinet).
•	pipettes must be clean and sterile; use autoclavable tips for repeating pipettes (eg. Gilson), pre-wrapped sterile single use plastic/glass pipettes, or if using non-sterile glass pipettes (with cotton plugs), sterilise in the flame before use.
The frequency of culture transfer depends on species and culture conditions (it is advisable to follow growth for each unfamiliar species to get a feeling). Transfers should be conducted as population growth slows, preferably before population growth ceases (stationary phase). Culture transfers (usually 1-5 % of culture volume into fresh sterile medium) can be conducted directly (pouring, not used when vessel neck is flame sterilized), or more often with a pipette.
Culture media
Media for the culture of marine phytoplankton consist of a seawater base (natural or artificial) which may be supplemented by various substances essential for microalgal growth, including nutrients, trace metals and chelators, vitamins, soil extract and buffer compounds.
Seawater base
The quality of water used in media preparation is very important. 
Natural seawater can be collected nearshore, but its salinity and quality is often quite variable and unpredictable, particularly in temperate and polar regions (due to anthropogenic pollution, toxic metabolites released by algal blooms in coastal waters). The quality of coastal water may be improved by ageing for a few months (allowing bacterial degradation of inhibitory substances), by autoclaving (heat may denature inhibitory substances), or by filtering through acid-washed charcoal (which absorbs toxic organic compounds). Most coastal waters contain significant quantities of inorganic and organic particulate matter, and therefore must be filtered before use (eg Whatman no. 1 filter paper). 
The low biomass and continual depletion of many trace elements from the surface waters of the open ocean by biogeochemical processes makes this water much cleaner, and therefore preferable for culturing purposes. 
Collect seawater from the front of the boat (or pump from subsurface) to avoid contamination. Seawater can be stored in polyethylene carboys, and should be stored in cool dark conditions.
Artificial seawater, made by mixing various salts with deionized water, has the advantage of being entirely defined from the chemical point of view, but they are very laborious to prepare, and often do not support satisfactory algal growth. Trace contaminants in the salts used are at rather high concentrations in artificial seawater because so much salt must be added to achieve the salinity of full strength seawater. Some of the more successful artificial seawater media that have been developed include the ESAW medium of Harrison et al. (1980), and the AK medium of Keller et al. (1987).
Nutrients,  trace metals, and chelators
The term ‘nutrient’ is colloquially applied to a chemical required in relatively large quantities, but can be used for any element or compound necessary for algal growth. 
Table 1. The average concentrations of constituents of potential biological importance found in typical seawater (after Brand, 1990). 
Element	Average Molar concentration (range in brackets)	
Group A		
Na+
K+
Mg2+
Ca2+
Cl-
SO42-
HCO3-
BO33-	          4.7 x 10-1
        1.02 x 10-2
          5.3 x 10-2
        1.03 x 10-2
          5.5 x 10-1
          2.8 x 10-2
          2.3 x 10-3
          4.2 x 10-4	
Group B		
Br-
F-
I03-
Li+
Rb+
Sr2+
Ba2+
MoO42-
VO43-
CrO42-
AsO43-
SeO42-	         8.4 x 10-4
         6.8 x 10-5
         4.4 x 10-7
         2.5 x 10-5
         1.4 x 10-6
         8.7 x 10-5
            1 x 10-7
         1.1 x 10-7
         2.3 x 10-8
            4 x 10-9
         2.3 x 10-8
         1.7 x 10-9	
Group C		
NO3-
PO43-
Fe3+
Zn2+
Mn2+
Cu2+
Co2+
SiO44-
Ni2+	           3 x 10-5 (10-8 to 4.5 x 10-5)
        2.3 x 10-6 (10-7 to 3.5 x 10-6)
           1 x 10-9 (10-10 to 10-7)
           6 x 10-9 (5 x 10-11 to 10-7)
           5 x 10-10 ( 2 x 10-10 to 10-6)
           4 x 10-9 (5 x 10-10 to 6 x 10-9)
           2 x 10-11 (10-11 to 10-10)
           1 x 10-4 (10-7 to 1.8 x 10-4)
           8 x 10-9 (2 x 10-9 to 1.2 x 10-8)	
Group A: concentrations of these constituents exhibit essentially no variation in seawater, and high algal biomass cannot deplete them in culture media. These constituents do not, therefore, have to be added to culture media using natural seawater, but do need to be added to deionized water when making artificial seawater media.
Group B: also have quite constant concentrations in seawater, or vary by a factor less than 5. Because microalgal biomass cannot deplete their concentrations significantly, they also do not need to be added to natural seawater media. Standard artificial media (and some natural seawater media) add molybdenum (as molybdate), an essential nutrient for algae, selenium (as selenite), which has been demonstrated to be needed by some algae, as well as strontium, bromide and flouride, all of which occur at relatively high concentrations in seawater, but none of which have been shown to be essential for microalgal growth. 
Group C: all known to be needed by microalgae (silicon is needed only by diatoms and some chrysophytes, and nickel is only known to be needed to form urease when algae are using urea as a nitrogen source). These nutrients are generally present at low concentrations in natural seawater, and since microalgae take up substantial amounts, concentrations vary widely (generally by a factor of 10 to 1000). All of these nutrients (except silicon and nickel in some circumstances) generally need to be added to culture media in order to generate significant microalgal biomass.
Nitrate is the N source most often used in culture media, but ammonium can also be used, and indeed is the preferential form for many algae since it does not have to be reduced prior to amino acid synthesis, the point of primary intracellular nitrogen assimilation into the organic linkage. Ammonium concentrations greater than 25µM are, however, often reported to be toxic to phytoplankton, so concentrations should be kept somewhat low.
Inorganic (ortho)phosphate, the P form preferentially used by microalgae, is most often added to culture media, but organic (glycero)phosphate is sometimes used, particularly when precipitation of phosphate is anticipated (when nutrients are autoclaved in the culture media rather than seperately, for example). Most microalgae are capable of producing cell surface phosphatases which allow them to utilize this and other forms of organic phosphate as a phosphorus source.
The trace metals which are essential for microalgal growth are incorporated into essential organic molecules, particularly a variety of coenzyme factors which enter into photosynthetic reactions. Of these metals, the concentrations (or more accurately the biologically available concentrations) of Fe, Mn, Zn, Cu and Co (and sometimes Mo and Se) in natural waters may be limiting to algal growth. Little is known about the complex relationships between chemical speciation of metals and biological availability. It is thought that molecules which complex with metals (chelators) influence the availability of these elements. Chelators act as trace metal buffers, maintaining constant concentrations of free ionic metal. It is the free ionic metal, not the chelated metal, which influences microalgae, either as a nutrient or as a toxin. Without proper chelation some metals (such as Cu) are often present at toxic concentrations, and others (such as Fe) tend to precipitate and become unavailable to phytoplankton. In natural seawater, dissolved organic molecules (generally present at concentrations of 1-10mg l-1) act as chelators. The most widely used chelator in culture media additions is EDTA (ethylenediaminetetraacetic acid), which must be present at high concentrations since most complexes with Ca and Mg, present in large amounts in seawater. EDTA may have an additional benefit of reducing precipitation during autoclaving. High concentrations have, however, occasionally been reported to be toxic to microalgae. As an alternative the organic chelator citrate is sometimes utilised, having the advantage of being less influenced by Ca and Mg. The ratio of chelator:metal in culture medium ranges from 1:1 in f/2 to 10:1 in K medium. High ratios may result in metal deficiencies for coastal phytoplankton (ie. too much metal is complexed), and many media therefore use intermediate ratios.
In today’s aerobic ocean, iron is present in the oxidized form as various ferric hydroxides and thus is rather insoluble in seawater. While concentrations of nitrogen, phosphorus, zinc and manganese in deep water are similar to plankton elemental composition, there is proportionally 20 times less iron in deep water than is apparently needed, leading to the suggestion that iron may be the ultimate geochemically limiting nutrient to phytoplankton in the ocean (Brand, 1986). Very little is known about iron in seawater or phytoplankton uptake mechanisms due to the complex chemistry of the element.  Iron availability for microalgal uptake seems to be largely dependent on levels of chelation. It is highly recommended that iron be added as the chemically prepared chelated iron salt of EDTA rather than as iron chloride or other iron salts; the formation of iron chelates is relatively slow, and iron hydoxides will form first in seawater, leading to precipitation of much of the iron in the culture medium.
Apparently as a result of the extreme scarcity of copper in anaerobic waters, copper did not begin to be utilised by organisms until the earth became aerobic and copper increased in abundance. Consequently copper does not seem to be an obligate requirement, algae either not needing it, or needing so little that free ionic copper concentrations in natural seawater are sufficient to maintain maximum growth rates (Brand, 1986). Copper may indeed be toxic, particularly to more primitive algae, and hence copper, if added to culture media at all, should be kept at low concentrations.
The provision of manganese, zinc and cobalt in culture medium should not be problematical since even fairly high concentrations are not thought to be toxic to algae.
Vitamins
Roughly ½ of all microalgal species tested have been shown to have a requirement for vitamin B12, which appears to be important in transferring methyl groups and methylating toxic elements such as arsenic, mercury, tin, thallium, platinum, gold, and tellurium (Brand, 1986), around 20% need thiamine, and less than 5% need biotin.
It is recommended that these vitamins are routinely added to seawater media. No other vitamins have ever been demonstrated to be required by any photosynthetic microalgae. 
Soil extract
•	Prepared by heating, boiling, or autoclaving a 5 to 30% slurry of soil in fresh water or seawater and subsequently filtering out the soil. 
Soil extract has historically been an important component of culture media. The solution provides macronutrients, micronutrients, vitamins, and trace metal chelators in undefined quantities, each batch being different, and hence having unpredictable effects on microalgae. With increasing understanding of the importance of various constituents of culture media, soil extract is less frequently used. Soil extract should only be used on a non-experimental basis.
Buffers
The control of pH in culture media is important since certain algae grow only within narrowly defined pH ranges, and in order to prevent the formation of precipitates. Except under unusual conditions, the pH of natural seawater is around 8. Because of the large buffering capacity of natural seawater (due to a bicarbonate buffering system, HCO3 being present at c. 2.2M) it is quite easy to maintain the pH of marine culture media. The buffer system is overwhelmed only during autoclaving, when high temperatures drive CO2 out of solution and hence cause a shift in the bicarbonate buffer system and an increase in pH, or in very dense cultures of microalgae, when enough CO2 is taken up to produce a similar effect. As culture medium cools after autoclaving, CO2 reenters solution from the atmosphere, but certain measures must be taken if normal pH is not fully restored:
•	The pH of seawater may be lowered prior to autoclaving (adjustment to pH 7-7.5 with 1M HCl) to compensate for subsequent increases. 
•	Certain media recipes include additions of extra buffer, either as bicarbonate, Tris (Tris-hydroxymethyl-aminomethane), or glycylglycine to supplement the natural buffering system. Tris may also act as a Cu buffer, but has occasionally been cited for its toxic properties to microalgae. Glycylglycine is rapidly metabolized by bacteria and hence can only be used with axenic cultures. These additions are generally not necessary if media are filter sterilized, unless very high cell densities are expected. 
•	The problem of CO2 depletion in dense cultures may be reduced by having a large surface area of media exposed to the atmosphere relative to the volume of the culture, or by bubbling with either air (CO2 concentration c.0.03%) or air with increased CO2 concentrations (0.5 to 5%). Unless there is a large amount of biomass taking up the CO2, the higher concentrations could actually cause a significant decline in pH. When bubbling is employed, the gas must first pass through an in line 0.2µm filter unit (eg. Millipore Millex GS) to maintain sterile conditions. For many microalgal species, aeration is not an option since the physical disturbance may inhibit growth or kill cells.
Media preparation
The salinity of the seawater base should first be checked (30-35‰ for marine phytoplankton), and any necessary adjustments (addition of fresh water/evaporation) made before addition of enrichments.
Always use reagant grade chemicals and bidistilled (or purer) water to make stock solutions of enrichments. Gentle heating and/or magnetic stirring of stock solutions can be used to ensure complete dissolution. When preparing a stock solution containing a mixture of compounds, dissolve each individually in a minimal volume of water before mixing, then combine and dilute to volume.
Seawater, stock solutions of enrichments and the final media must be sterile in order to prevent (or more realistically minimize) biological contamination of unialgal cultures. Several methods are available for sterilization:
•	autoclaving is the most widely used technique for sterilizing culture media, and is the ultimate guarantee of sterility (including the destruction of viruses). A commercial autoclave is best, but pressure cookers of various sizes are also suitable. Sterility requires 15 minutes at 1-2 Bar pressure and a temperature of 121°C in the entire volume of the liquid (ie. longer times for larger volumes of liquid; approximately 10 min for 100ml, 20 min for 2l, 35 min for 5 l). Bottles containing media should be no more than ¾ full, and should be left partially open or plugged with cotton wool or covered with aluminium foil. Ensure the heating elements are covered with distilled water, and the escape valve should not be closed until a steady stream of steam is observed. After autoclaving, the pressure release valve should not be opened until the temperature has cooled to below 80°C. 
Autoclaving is a process which has many effects on seawater and its constituents, potentially altering or destroying inhibitory organic compounds, as well as beneficial organic molecules. Because of the steam atmosphere in an autoclave, CO2 is driven out of the seawater and the pH is raised to about 10, a level which can cause precipitation of the iron and phosphate added in the medium. Some of this precipitate may disappear upon re-equilibriation of CO2 on cooling, but both the reduced iron and phosphate levels, and the direct physical effect of the precipitate may limit algal growth. The presence of EDTA and the use of organic phosphate may reduce precipitation effects. Addition of 5% or more of distilled water may also help to reduce precipitation (but may affect final salinity). The best solution, however, if media are autoclaved, is to sterilize iron and phosphate (or even all media additions) seperately and add them aseptically afterwards.
Autoclave steam may contaminate the media (i.e. with trace metals from the autoclave tubing). Autoclaving also produces leaching of chemicals from the medium receptacle into the medium (silica from glass bottles, toxic chemicals from plastics). Autoclaving in well-used Teflon or polycarbonate vessels reduces leaching of trace contaminants.
Autoclaving will cause evaporation of water, and hence an increase in salinity (usually of c. 1‰). Distilled water can be added prior to autoclaving to compensate for this increase.
•	Pasteurization (heating to 90-95°C for 30 minutes) of media in Teflon or polycarbonate bottles is a potential alternative, reducing the problems of trace metal contamination and alteration of organic molecules inherent with autoclaving. Pasteurization does not, however, completely sterilize the seawater; it kills all eukaryotes and most bacteria, but some bacterial spores probably survive. Heating to 90-95°C for at least 30 minutes and cooling, repeated on two successive days (‘tyndallisation’) may improve sterilization efficiency; it is assumed that vegetative cells are killed by heat and heat resistant spores will germinate in the following cool periods and be killed by subsequent heating.
•	Ultraviolet radiation can be used to sterilize seawater, but very high intensities are needed to kill everything in the seawater (1200 W lamp, 2-4h for culture media in quartz tubes). Such intense UV light necessarily alters and destroys the organic molecules in seawater and generates many long lived free radicals and other toxic reactive chemical species (Brand, 1986). Seawater exposed to intense UV light must, therefore, be stored for several days prior to use to allow the level of these highly reactive chemical species to decline.
•	Sterilization by commercial microwave apparatus is another option. Microwave sterilization has not, as yet, been widely employed in culture media preparation due to uncertainties about sterilization efficiency. Trials should be conducted before use of this method to ensure sterility of seawater.
•	Sterile filtration is probably the best method of sterilizing seawater without altering the chemistry of the seawater, as long as care is taken not to contaminate the seawater with dirty filter apparatus. Sterilization efficiency is, however, to some extent reduced compared with heat sterilization methods. Membrane filters of 0.2µm porosity are generally considered to yield water free of bacteria, but not viruses. 0.1µm filters can be used, but the time required for filtration of large volumes of culture media may be excessively long. The filtration unit must be sterile: for small volumes (<50ml) pre-sterilized single use filter units for syringe filtration (eg. Millipore Millex GS) can be used; for volumes up to 1 litre reusable autoclavable self-assembly filter units (glass or polycarbonate) with 47mm cellulose ester membrane filters (eg. Millipore HA) can be used with suction provided by a vacuum pump; for larger volumes an in-line system with peristaltic pump and cartridge filters may be the best option.
Filter units (particularly disposable plastic systems), and the membrane filters themselves can also leak toxic compounds into the filtrate. The first portion of filtrate (eg. 5% of volume to be filtered) should be discarded to alleviate this problem.
Most stock solutions of culture medium additions can be sterilized separately by autoclaving, although vitamin stock solutions are routinely filtered through 0.2µm single use filter units (eg. Millipore Millex GS), since heat sterilization will denature these organic compounds. Filter sterilization of all additions may reduce uncertainties about stability of the chemical compounds and contamination from autoclave steam, but absolute sterilization is not guaranteed. Stock solutions can be stored in ultraclean sterile glass, polycarbonate or Teflon tubes/bottles. In order to minimize effects of any microbial contaminations, all stock solutions should be stored in a refrigerator at 4°C, except vitamin stocks which are stored frozen at –20°C and thawed immediately prior to use.
Some culture medium recipes
The recipes of 3 media which have proved successful for the culture of coccolithophores are given. In practise dilutions of f/2 and K medium  (eg. 10%-->f/20, K/10) are sufficient to maintain good coccolithophorid growth. A variety of alternative marine culture media recipes are given by Stein (1975), and on the web pages of the major culture collections (eg. CCMP, Utex).
__________________________________________________________________________________________
f/2-Si (Guillard, 1975)
To 996ml of sterile seawater aseptically add:
Quantity	Compound	Stock solution (sterile)	Final conc. in medium	
1.0ml	NaNO3	75.0g/litre H2O 	884µM	
1.0ml	NaH2PO4.H2O	5.0g/litre H2O	36µM	
1.0ml	f/2 trace metal solution	(see recipe below)	(see below)	
1.0ml	f/2 vitamin solution	(see recipe below)	(see below)	

f/2 trace metal solution
To 950ml distilled H2O add:
Quantity	Compound	Stock solution	Final conc. in medium	
3.15g	FeCl3.6H2O	-	11.7µM	
4.36g	Na2EDTA.2H2O	-	12µM	
1.0ml	CuSO4.5H2O	9.8g/litre H2O	0.04µM	
1.0ml	Na2MoO4.2H2O	6.3g/litre H2O	0.03µM	
1.0ml	ZnSO4.7H2O	22.0g/litre H2O	0.08µM	
1.0ml	CoCl2.6H2O	11.9g/litre H2O	0.05µM	
1.0ml	MnCl2.4H2O	178.2g/litre H2O	0.9µM	
Make up to 1 litre with distilled H2O, sterilize (autoclave or filter) and store in fridge.

f/2 Vitamin solution
To 950ml distilled H2O add:
Quantity	Compound	Stock solution	Final conc. in medium	
1.0ml	Vit. B12 (cyanocobalamin)	0.5g/litre H2O	0.37nM	
1.0ml	Biotin	5.0mg/litre H2O	2.0nM	
100.0mg	Thiamine HCl	-	0.3µM	
Make up to 1 litre with distilled H2O, filter sterilize into plastic vials and store in freezer.
__________________________________________________________________________________________
K(-Si) (Keller et al., 1987)
To 994 ml of sterile seawater aseptically add:
Quantity	Compound	Stock solution (sterile)	Final conc. in medium	
1.0ml	NaNO3	75.0g/litre H2O 	884µM	
1.0ml	NH4Cl *	0.535g/litre H2O	10µM	
1.0ml	Na2glycerophosphate **	2.16g/litre H2O	10µM	
1.0ml	TRIS-base (pH7.2) ***	121.1g/litre H2O	1mM	
1.0ml	K trace metal solution	(see recipe below)	(see below)	
1.0ml	f/2 vitamin solution	(see recipe below)	(see below)	
* should not be autoclaved (volatile when heated)
** inorganic orthophosphate can be substituted (particularly if not autoclaving)
*** can be omitted (particularly if not autoclaving, or if cell density will not be very high)
K trace metal solution
To 950ml distilled H2O add:
Quantity	Compound	Stock solution	Final conc. in medium	
4.3g	(Na)FeEDTA	-	11.7µM	
37.22g	Na2EDTA.2H2O	-	100µM	
0.5ml	CuSO4.5H2O	9.8g/litre H2O	0.02µM	
1.0ml	Na2MoO4.2H2O	6.3g/litre H2O	0.03µM	
1.0ml	ZnSO4.7H2O	22.0g/litre H2O	0.08µM	
1.0ml	CoCl2.6H2O	11.9g/litre H2O	0.05µM	
1.0ml	MnCl2.4H2O	178.2g/litre H2O	0.9µM	
1.0ml	H2SeO3	1.29mg/litre H2O	0.01µM	
Make up to 1 litre with distilled H2O, sterilize (autoclave or filter) and store in fridge.
 f/2 Vitamin solution
To 950ml distilled H2O add:
Quantity	Compound	Stock solution	Final conc. in medium	
1.0ml	Vit. B12 (cyanocobalamin)	0.5g/litre H2O	0.37nM	
1.0ml	Biotin	5.0mg/litre H2O	2.0nM	
100.0mg	Thiamine HCl	-	0.3µM	
Make up to 1 litre with distilled H2O, filter sterilize into plastic vials and store in freezer.
__________________________________________________________________________________________
BWM (blue water medium) Brand (unpublished)

To 995.5 ml sterile seawater, aseptically add:
Quantity	Compound	Stock solution (sterile)	Final conc. in medium	
1.0ml	NaNO3	8.5g/litre H2O 	100µM	
1.0ml	NH4Cl *	0.535g/litre H2O	10µM	
1.0ml	Na2glycerophosphate *	2.16g/litre H2O	10µM	
1.0ml	BWM trace metal solution	(see recipe below)	(see below)	
0.5ml	BWM vitamin solution	(see recipe below)	(see below)	
* should not be autoclaved (volatile when heated)
** can be substituted for inorganic orthophosphate

BWM trace metal solution
To 950ml distilled H2O add:
Quantity	Compound	Stock solution	Final conc. in medium	
0.367g	(Na)FeEDTA	-	1µM	
3.72g	Na2EDTA.2H2O	-	10µM	
1.0ml	CuSO4.5H2O	0.25g/litre H2O	0.001µM	
1.0ml	ZnSO4.7H2O	28.75g/litre H2O	0.1µM	
1.0ml	CoCl2.6H2O	2.38g/litre H2O	0.01µM	
1.0ml	MnCl2.4H2O	19.8g/litre H2O	0.1µM	
1.0ml	H2SeO3 *	1.29g/litre H2O	0.01µM	
* optional
Make up to 1 litre with distilled H2O, sterilize (autoclave or filter) and store in fridge.

BWM Vitamin solution
To 950ml distilled H2O add:
Quantity	Compound	Stock solution	Final conc. in medium	
1.0ml	Vit. B12 (cyanocobalamin)	13.55g/litre H2O	0.01µM	
1.0ml	Biotin	2.5mg/litre H2O	0.001µM	
33.3mg	Thiamine HCl	-	0.1µM	
Make up to 1 litre with distilled H2O, filter sterilize into plastic vials and store in freezer.
__________________________________________________________________________________________
Some aquatic microalgae grow well on solid substrate. 
A 3% high grade agar can be used for the solid substrate. The agar and culture medium should not be autoclaved together, because toxic breakdown products can be generated. The best procedure is to autoclave 30% agar in deionized water in one container and nine times as much seawater base in another. After removing from the autoclave, sterile nutrients are added aseptically to the water, which is then mixed with the molten agar. After mixing, the warm fluid is poured into sterile petri dishes, where it solidifies when it cools. The plate is inoculated by placing a drop of water containing the algae on the surface of the agar, and streaking with a sterile implement. The plates are then maintained under standard culture conditions. This method may be particularly effective for cleaning cultures infected with bacteria, clean colonies of the algal species being isolated from the plate into fresh liquid medium.
Considerations in the selection of culture media
Two approaches to selection of media composition:
•	In theory it is best to work on the principle that if the alga does not need the addition of any particular chemical substance to the culture media (ie. if it has no observable positive effect on growth rate), don’t add it. 
•	In practise it is often easier to follow well known (and presumably, therefore, well tried) media recipes, and safer to add substances ‘just in case’ (providing they have no observable detrimental affect on algal growth). 
When choosing a culture medium, the natural habitat of the species in question should be considered in order to determine its environmental requirements: 
Table 2. Considerations in the selection of culture media (after Brand, 1986).
Habitats
Eutrophic, coastal, variable environment, well-mixed, nutrient rich		
Oligotrophic, oceanic, predictable environment, stratified, nutrient poor	
Organisms
r-selected, rapid growth rate, wide environmental tolerance, highly autotrophic, high biomass/low diversity communities		
K-selected, slow growth rate, narrow environmental tolerance, tendency towards auxotrophy/mixotrophy/photoheterotrophy, low biomass/high diversity communities
	
     ________________diatoms__________________
                                          ____hetero-______coccolithophores______holo-____
                                                                   ________________dinoflagellates________________
	
Culture media
f/2, GPM, IMR, ESAW                  	
  K, BWM      	
                      ??????	
To date, culture techniques have been quite successful in culturing the r-selected species from eutrophic habitats, but quite poor at culturing the K-selected species from oligotrophic habitats. The media recipes currently available should be adequate for the culture of diatoms, some coccolithophorids (particularly coastal heterococcolithophorids) and few coastal dinoflagellates, the exact choice for a particular species therefore dependant on trial and error (taking into account the above discussion). K and BWM media are perhaps the best available options for successful culturing of oceanic coccolithophorids and dinoflagellates, but further advances, including the reduction in concentration of many nutrients, and the possible use of organic nutrients, should also be considered.
•	It must be remembered that in culturing in general there are (within limits) no right and wrong methods; culture media have only developed by culturers trying out various additions (usually based on theoretical considerations), and hence innovation is actively encouraged.
Pure, axenic cultures
If biological contaminants appear in a culture, the best remedy is to isolate a single cell from the culture with a micropipette, and try to establish a new, clean clonal culture. Alternatively the culture can be streaked on an agar plate (see below) in the hope of attaining a colony free of contaminants. Neither of these methods work well, however, for eliminating bacteria that attach firmly to the surface of microalgae. Placing a test tube of microalgal culture in a low-intensity 90 kilocycles/sec ultrasonic water bath for varying lengths of time (a few seconds to tens of minutes) can sometimes physically separate bacteria without killing the algae, making it easier to obtain an axenic culture by micropipette isolation.
Often, however, to achieve an axenic culture, antibiotics must be used. Best results appear to occur when an actively growing culture of algae is exposed to a mixture of penicillin, streptomycin and gentamycin for around 24 hours. This drastically reduces the growth of bacteria while allowing the microalgae to continue to grow, increasing the chances of obtaining an axenic cell when using micropippette or agar streaking isolation. Different algal species tolerate different concentrations of antibiotics, so a range of concentrations should be used (generally 50-500 mg/l). Other antibiotics that can be used include chloramphenicol, tetracycline, and bacitracin. Antibiotic solutions should be made with distilled water and filter-sterilized (0.2µm filter units) into sterile  tubes, and should be stored frozen until use. Another approach is to add a range of antibiotic concentrations to a number of subcultures and then select the culture that has surviving algal cells but no surviving bacteria or other contaminants. Sterility of cultures should be checked by microscopic examination and by adding a small amount of sterile bacterial culture medium (eg. 0.1% peptone) to a microalgal culture and observing regularly for bacterial growth. Absence of bacterial growth does not, however, ensure that the microalgal culture is axenic, since the majority of bacteria do not respond to standard enrichments. In reality there is no way of demonstrating that a microalgal culture is completely axenic. In practise, therefore, axenic usually means ‘without demonstrable unwanted prokaryotes or eukaryotes’. Some microalgal cultures may die when made axenic, probably due to the termination of obligate symbiotic relationships with bacteria.
Equipment checklist
•	a north-facing window (in the northern hemisphere) in a room in which temperature is not too high (normally <25°C, depending on type of algae being grown), supplementary fluorescent lights,
or; a reach-in (fridge-like) incubator, with controllable temperature and illumination,
or; a cupboard type reach-in, or walk-in incubator (very expensive).
•	bench space (preferably in a clean lab), 
and/or; a laminar flow cabinet.
70% ethanol, a bunsen burner, sterile glass/plastic pipettes (2ml, 10ml, 25ml), pipette bulb.
•	detergent (laboratory, not domestic), 10% HCl, and distilled water for cleaning cultureware.
•	culture vessels; borosilicate glass Erlenmeyer (or other shape) flasks with cotton wool bungs / glass covers, or pre-wrapped sterile polystyrene culture flasks.
•	seawater (for natural based media), reagant grade chemicals and bidistilled water for media enrichments, an analytical balance (1mg sensitivity), spatula, hot-plate magnetic stirrer, sterile glass / polystyrene / polycarbonate tubes/bottles for media enrichments, media recipients – 1 litre glass/polycarbonate/Teflon bottles, fridge/freezer, pH meter, salinity meter.
•	equipment sterilization:
either; an autoclave / pressure cooker (for moist heat sterilization)
or; an oven (for dry heat sterilization)
•	media sterilization
either; an autoclave / pressure cooker
or; a microwave oven
or; filter apparatus (see discussion above)

References & further reading

Brand L. E. (1986). Photosynthetic picoplankton. Canadian Bulletin of Fisheries and Aquatic Sciences 214, 205-233.
Brand L. E. (1990). The isolation and culture of microalgae for biotechnological applications. In: Isolation of biotechnological organisms from nature, Labeda D. P. (ed.). McGraw-Hill, New York, pp. 81-115.
Guillard R. R. L. (1975).  Culture of phytoplankton for feeding marine invertebrates. In: Culture of marine invertebrate animals, Smith W. L. and Chanley M. H. (eds.). Plenum, New York, pp. 29-60.
Guillard R. R. L. and Keller M. D. (1984). Culturing dinoflagellates. In: Dinoflagellates, Spector D. L. (ed.). Academic, New York, pp. 391-442.
Harrison PJ, Waters RE & Taylor FJR (1980). A broad spectrum artificial seawater medium for coastal and open ocean phytoplankton. J. Phycol. 16, 28-35.
Keller M.D., Selvin R. C., Claus W and Guillard R. R. L. (1987). Media for the culture of oceanic ultraphytoplankton. J. Phycol. 23, 633-638. 
Provasoli L., McLaughlin J. J. A. and Droop M. R. (1957). The development of artificial media for marine algae. Archiv für Mikrobiologie. 25, 392-428.
Stein J. R. (1973). Handbook of phycological methods: Culture methods and Growth measurements. Cambridge University Press, London and New York. 448pp.