Microalgal culturing
Practical notes from the culturing short course held in Caen, 10-12 Feb 1999.
An RTF version oif this document can be down loaded by clicking here (click and hold on a MacIntosh) and select "save this link as".Ian Probert & Christine Klaas - with thanks to Franco Novarino
General principles
A culture can be defined as an artificial environment in which the algae grow. In theory, culture conditions should resemble the algas 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.
Physical parameters
TemperatureLight
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 湲/s/m2), with 5-10% (c. 80-160湲/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:
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):
Cleaning/sterilization of culture materials
Cleaning procedure
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 sterilisation methods and the choice depends on the purpose and material used:
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.
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).
X and
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:
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 25然 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 separately, for example). Most microalgae are capable of producing cell surface phosphatases which allow them to utilise 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 todays 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 _ 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
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:
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 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.
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 0.2痠 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 |
884然 |
1.0ml |
NaH2PO4.H2O |
5.0g/litre H2O |
36然 |
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 |
- |
11.7然 |
4.36g |
Na2EDTA.2H2O |
- |
12然 |
1.0ml |
CuSO4.5H2O |
9.8g/litre H2O |
0.04然 |
1.0ml |
Na2MoO4.2H2O |
6.3g/litre H2O |
0.03然 |
1.0ml |
ZnSO4.7H2O |
22.0g/litre H2O |
0.08然 |
1.0ml |
CoCl2.6H2O |
11.9g/litre H2O |
0.05然 |
1.0ml |
MnCl2.4H2O |
178.2g/litre H2O |
0.9然 |
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 |
- |
0.3然 |
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 |
884然 |
1.0ml |
NH4Cl * |
0.535g/litre H2O |
10然 |
1.0ml |
Na2glycerophosphate ** |
2.16g/litre H2O |
10然 |
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 |
- |
11.7然 |
37.22g |
Na2EDTA.2H2O |
- |
100然 |
0.5ml |
CuSO4.5H2O |
9.8g/litre H2O |
0.02然 |
1.0ml |
Na2MoO4.2H2O |
6.3g/litre H2O |
0.03然 |
1.0ml |
ZnSO4.7H2O |
22.0g/litre H2O |
0.08然 |
1.0ml |
CoCl2.6H2O |
11.9g/litre H2O |
0.05然 |
1.0ml |
MnCl2.4H2O |
178.2g/litre H2O |
0.9然 |
1.0ml |
H2SeO3 |
1.29mg/litre H2O |
0.01然 |
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 |
- |
0.3然 |
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 |
100然 |
1.0ml |
NH4Cl * |
0.535g/litre H2O |
10然 |
1.0ml |
Na2glycerophosphate * |
2.16g/litre H2O |
10然 |
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 |
- |
1然 |
3.72g |
Na2EDTA.2H2O |
- |
10然 |
1.0ml |
CuSO4.5H2O |
0.25g/litre H2O |
0.001然 |
1.0ml |
ZnSO4.7H2O |
28.75g/litre H2O |
0.1然 |
1.0ml |
CoCl2.6H2O |
2.38g/litre H2O |
0.01然 |
1.0ml |
MnCl2.4H2O |
19.8g/litre H2O |
0.1然 |
1.0ml |
H2SeO3 * |
1.29g/litre H2O |
0.01然 |
* 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 |
0.01然 |
1.0ml |
Biotin |
2.5mg/litre H2O |
0.001然 |
33.3mg |
Thiamine HCl |
- |
0.1然 |
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:
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______h olo-____ ________________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.
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 (0.2痠 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
or; a reach-in (fridge-like) incubator, with controllable temperature and illumination,
or; a cupboard type reach-in, or walk-in incubator (very expensive).
and/or; a laminar flow cabinet.
70% ethanol, a bunsen burner, sterile glass/plastic pipettes (2ml, 10ml, 25ml), pipette bulb.
either; an autoclave / pressure cooker (for moist heat sterilization)
or; an oven (for dry heat 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.
Novarino G. (1998) general aspects of microalgal cultivation. In, Report and abstracts of CODENET First Annual Meeting.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.