Culturing

Protocols for Isolation and Culturing of Freshwater Algae

Algal cultures are essential when conducting competition studies, bioassays, assessment of zooplankton food preferences, and determination of algal life histories. They are also necessary for molecular systematic work. Algal cultures may be "unialgal," which means they contain only one kind of alga, usually a clonal population (but which may contain bacteria, fungi, or protozoa), or cultures may be "axenic," meaning that they contain only one alga and no bacteria, fungi or protozoa. There are four major techniques for obtaining unialgal isolates: streaking, spraying, serial dilution, and single-cell isolations. Streaking and spraying are useful for single-celled, colonial, or filamentous algae that will grow on an agar surface; cultures of some flagellates, such as Chlamydomonas and Cryptomonas may also be obtained by these procedures. Many flagellates, however, as well as other types of algae must be isolated by single-organism isolations or serial-dilution techniques.

Culture Collections

UTEX The Culture Collection of Algae [1]

Canadian Phycological Culture Centre (CPCC) [2]

SAG Culture Collection, University of Göttingen [3]

Culture Collection of Algae and Protozoa (CCAP) [4]

A list of Algal Culture Collections [5]

Algae Depot [6]

Ward's Natural Science Algal Cultures [7]

Training Website

Online Seminar on cell culture - controlling contamination [8]

Useful Websites

Phycological collection catelogues [9]

I. Species Isolation

The first goal when establishing any new culture of algae is to isolate the species and generate a ‘clean’ monoculture. It involves the elimination of contaminant species, especially those that can outcompete the target species. There are three general methods that can be used for isolation: single-cell isolation by micropipette, agar streaking, and atomized spray onto agar plates. After one of these techniques is successful at isolating a focal taxon, then cells of the separates species are grown in three different culture media in hopes that one of these is sufficient for algal growth.

A. Pre-screening of metazoans: For water or sediment samples taken from a natural lake or stream, it may be necessary to first screen out larger organisms such as zooplankton or insects using a 500 µm mesh sieve.

B. Initial observation and ID

1. To identify the target species, put the sample containing into a plastic 60×15mm Petri dish if you were looking for large filamentous algae (you can see algal mat); put the sample into a multi-well plate or on a microscope slide if you were looking for small algae (i.e., diatom, blue-green, green coccoids). 2. Use a dissecting microscope for Petri dish and a compound microscope for multi-well plates and slides. Locate the target species under low-level magnification on a microscope. 3. Use the software imaging system to take a picture of the target species and put a ruler on the image. This image may be useful later on for determining the success of culture methods, and for comparing morphology of the taxon between field populations vs. cultured populations.

C. Single-cell isolation by micropipette

1. Prepare 3 or more sterile droplets in multi-well plate (see Fig.1), or on a microscope slide for five culture mediums, respectively.

2. Prepare glass Pasteur pipette (very fine-tipped pipette):

a) Hold the Pasteur pipette in one hand, and a forceps holds in the other hand to support the tips. b) Heat the Pasteur pipette in a flame of a Bunsen burner (gas flame) and rotate it to provide even softening as the pipette warms to the melting point. c) When the heated area is sufficiently soft, remove the pipette from flame and simultaneously pull it to produce a thin tube. d) Cool the glass for a couple of seconds. Next, reposition the forceps to the thin area, approximately where the weight of the tip bends downward. Remove the end with a slight tugging and bending motion. The broken end of pipette should be smooth and round, not jagged or broken. e) You can vary the diameter of the finely pulled tip by changing the speed of pulling; the diameter of a slowly-pulled tip will be greater than that of a rapidly-pulled tip. You would want a narrow diameter tip if you are trying to isolate very small algae, but a larger diameter tip is required for large cells. Try to match the diameter of the pipette tip to the size of the algal cells to be isolated.

3. Pick up a cell from the sample using the prepared Pasteur pipette.

a) Attach the micropipette to a pipette bulb. b) Place a Petri dish of algae on the stage of a dissecting microscope (or slides on compound microscope) and locate the single cell/colony/filament to be isolated. Then find the tip of the micropipette and move it to the vicinity of the alga, then suck it up into the pipette tip, then stop the suction. Try to avoid sucking up any other algae. c) Remove the pipette from the dish, then blow the liquid+algae into one of the drops of medium on a multi-well plate. d) Washing procedure: with the same technique, a clean micropipette (the previously used micropipette can be redrawn to form a new tip, and the heat required to melt the glass is sufficient to sterilize it, assuming that no contaminating liquid is further up in the pipette. The micropipette can continue to be used until all of the pulled portion has been consumed.) is then used to pick up the cell again, and transfer it to a second sterile droplet. Repeat this process until a single algal cell can be confidently placed into culture medium (Fig.1).

e) Filaments can be grabbed with a slightly curved pipette tip and dragged through soft agar (less than 1%) to remove contaminants. It is best to begin with young branches or filament tips which have not yet been extensively epiphytized.

D. Single-cell isolation with use of agar: Some algal species are able to grow on agar very well, and thus isolation via agar is the preferred isolation method for many coccoid algae and soil algae. Following is a list of algae that can be isolated on agar: Chlamydomonas; Pavlova; Synura; Tetracelmis.

1. Sterilize the loop in gas flame first. Load the bacterial loop with a small amount of sample, and then run the loop along the Petri dish following certain pattern (Fig. 2). 2. Inoculate algae in Petri dishes with three different kinds of medium because we don’t know which kind of medium is most suitable for algal growth and the algae has to be left on the agar for a while. 3. After streaking, seal the Petri dishes with parafilm, and the agar plate is incubated until colonies of cells appear. Repeat the process until a single-species colony appears. 4. Isolate the colonies from agar plate with a drawn-out micropipette or bacterial loop, and rinse in a liquid medium to free the cell. Sterilize the ring before pick another algal colony.

E. Atomized cell spray technique: In this technique, a stream of compressed air is used to disperse algal cells from a mixture onto the surface of a Petri plate containing growth medium solidified with agar.

1. Hold a Petri plate about 18 inches from the two Pasteur pipettes. One of these is attached to an airline via a hose, and mounted onto a ringstand. The other pipette is suspended tip-up into a container holding the algal mixture. The other pipette is suspended tip-up into a container holding the algal mixture. The airflow from the first pipette creates a vacuum that draws a stream of algae-containing liquid up from the container through the second pipette. A liquid suspension containing target species is atomized with forced sterile air so that cells are scatted onto the plate (Fig. 3). The spray should be done in a sterile hood. 2. Spray the algal mixture at three different kinds of Petri dishes. 3. After about a week you should see algal growth. You want to pick the colonies as early as possible, so look for colonies under stereo scope at 25-50 X. 4. Incubate the plates, and after colony formation, select cells for culturing next step.

F. Culture: after transfer through 3-10 drops, transfer the algae into well plates with growth medium.

1. If the algae was isolated by single-cell isolation technique, prepare a 48-well plate with 3 culture mediums. Do 8-10 replicates for each medium. Find at least 24 single cells and put the single cell into the culture medium. Label flasks in the following form: Scientific name + strain number + culture medium. Put it on shaker table. 2. If the algae was isolated by striking or spray technique, first choose the most suitable medium. Then prepare a 48-well plate with the most suitable medium. Isolate colony with Pasteur pipette and rinse the pipette in the medium. 3. Keep the original sample and the isolation flasks and Petri dishes in incubator at least for one week in case the target species fail to grow in well plates. Add some culture medium to the original raw sample. 4. Set up the growth chamber with illumination of 30-60 μmol·m-2·s-1. Place algae into growth chamber and allow growth to occur for 2 weeks. 5. For newly isolated cells, a light-dark cycle should be used until continuous light growth is established. A light-dark cycle between 12:12 and 16:8 is used. 6. Examine them with dissecting microscope for signs of growth or contamination.

G. Growth rates and growth period

Measure the fluorescence value of each well as an estimate of cell density. Do the measurement at the same time everyday (e.g. 2pm everyday). Plot log fluorescence against time to get an idea which medium is most suitable for specific algal species and what is the maximum growth rate for each species given saturating nutrients. Or calculate the growth rate using the equation as shown below. Enter the growth rate into spreadsheet. Write down the days for a specific species to reach maximum fluorescence. r=(lnF_2-lnF_1)/(〖Day〗_2-〖Day〗_1 )

Notes: It is not uncommon for the target species to grow in the initial stages of isolation but die after one or more transfers to fresh culture medium. This often indicates that the culture medium is lacking a particular element or organic compound, which is not immediately manifest. Alternatively, the organism may be accumulating wastes that poison its environment, causing death.

II. Establish Stock Culture

The stock cultures are prepared to maintain a complete algal library.

As long as a successful unialgae culture is gained, algal transfer and cultivation should be performed in laminar flow hood to protect cultivation from contamination.

A. Set-up in the laminar flow hood

1. Turn off UV lights, turn on regular lights if needed. 2. Run hood blower for 3 minutes to clean the air in the hood. Swipe the hood stage with 70% EtOH. 3. Everything that goes INTO the hood must be cleaned with 70% EtOH. This includes your hands. Spray them down before starting. Wipe off anything you intend to use in the hood with 70% EtOH, bring stuff into the hood. 4. When finished, remove all of your media/solution bottles (tightly capped), tubes, etc from the hood. Wipe down the bench with 70% EtOH.

B. Check the library algae plate for contaminating algae under a microscope before using it to inoculate stock cultures

1. Before using the microscope, spray a Kimwipe with 70% EtOH and wipe down the stage. Do this also when you are done to prevent media spills from spreading between plates. 2. Prepare a wet mount slide. Place 1-2 drops of dH2O on a clean slide using Pasteur pipette. Burn the loop, wait five seconds to let it cool down and then scrape some algae from the old culture, rinse the algae in the water drop, and then cover it with coverglass. Try to avoid bubble in the slide when put the coverglass on it. Observe the algae under microscope at 400× magnification. For tiny algal species, try to observe it at 1000× magnification.

C. Agar plates culture

1. Sterilize hands by spraying with ethanol, then sterilize the inoculation loop in a Bunsen burner at such an angle that the whole wire glow red, and allow it to cool in air for 5 seconds. Prepare 5 agar slants with suitable mediums. 2. Run the loop along the agar plates with the desired algae in zigzag motion, and then sterilize the loop again before inoculating the next agar plate. Cap the plates. Do at least 5 replicates for each strain. Label it and put it into growth chamber. 3. Sterilize the surface and hands in between each inoculation to make sure that there isn’t contamination. Make sure that there is no liquid on the surface when you take out the next dish. Liquid is a great medium for contamination! 4. Check library cultures for fungi growth and contamination every 4-6 weeks. Redo library if necessary, we usually transfer algae every 4-6 months.

D. Backup culture

1. Some algae does not grow on agar surface. Therefore every time when we transfer algae to agar plates, we will also transfer algae to liquid culture as a backup of algal strain. 2. Run the ring along the plate with the desired algae (be sure to sterilize the ring first) and then put the ring in the liquid media (swish around a bit, but don’t touch the sides of the flask with the ring). Sterilize the ring again before inoculating the next flask. Keep a sterile cotton ball in the flask opening when not in use. Do 5 replicates.

• • Notes: When discarding of either the stock or the active cultures, autoclave and then discard. The petri dishes do melt, so place them in an autoclavable bag first.

III. Initial Characterization of Single Species

A. Growth rates and growth period

       These have already been done in Section I.G.

B. Biovolume

1. Make a wet slide of algal culture in laminar flow hood and observe it with microscope to make sure it is a unialgae cultivation without contamination of other algae. If contamination occurred, re-isolate target species. 2. Observe the algal culture under microscope, measure cell dimension (i.e., diameter, length, width, depth). Calculate biovolume with reference to Hillebrand et al.’ paper. Record the biovolume in spreadsheet.

C. Images

1. Take some pictures of the species, put rulers on the image, and save them with the taxonomic name of species. 2. Compare it with the algal image of field sample to ensure normal morphology.

D. Resources limitation

1. Set up nutrient gradients with regards of nitrogen, phosphorus and silica in well plates, separately. Inoculate algae from stock culture. 2. Measure the fluorescence every day during culture period. 3. Plot growth rates against nutrients. Find out the nutrient level where the growth rate is zero. Record the nutrient value as R* (i.e., N*, P*, Si*) in spreadsheet.

IV. Entry into Library Datasheet

A. Identify the target algae at the species level. Record species name and the origin of the species into the algae library spreadsheet. The origin includes habitat and locality, name of isolator, and isolation date. B. Record morphological characteristics of algae and culture medium into spreadsheet, which will be done in section III. C. Record conditions for routine serial transfer into spreadsheet, which will be done in section IV.

V. Maintenance and Batch Culture

The object of maintenance is to retain a healthy, physiologically, morphologically, and genetically representative population. Therefore we need to subculture the organism at the end of its exponential growth phase. Batch cultures are used for experiments and are made right before the experiments.

A. Check the cultures for contamination using a microscope and to purify them if contaminated.

B. Liquid culture

1. Prepare several 250ml flasks with medium half filled. 2. Run the ring along the plate with the desired algae (be sure to sterilize the ring first) and then put the ring in the liquid media (swish around a bit, but don’t touch the sides of the flask with the ring). Sterilize the ring again before inoculating the next flask. Keep a sterile cotton ball in the flask opening when not in use. 3. Once the 250 ml flasks are prepared, you can move them to the fridge with the cultures. Make sure to sterilize the area before bringing the flasks into the room. It usually takes 5-10 days before any algae are visible in the cultures.

C. Transfer

1. Prepare some agar slants or liquid medium depending on what previous substrate the target algae grow on. 2. Run the sterilized loop on the previous agar plates, streak in a zig-zag pattern on the new agar plates. Do 5 replicates. 3. Use pipette transfer 5-10 ml of algal suspension into the new liquid medium. Do five replicates. 4. Keep the previous stock culture until you make sure the new stock culture is successful.

D. Transfer interval

1. A safe transfer interval can be estimated by using one quarter the time that a strain can maximally survive. 2. For sensitive strains, the shortest transfer cycle is 1-2 weeks. 3. For some green and blue-green algae which are maintained at low light and temperature, the transfer cycle can be 6 months. 4. To reduce the risk of loosing strains, retain subcultures from at least two different transfer dates. 5. Label each tube with transfer date if transferred at different time; label the whole rack with transfer date if transferred at the same time.

E. Culturing conditions regarding morphological maintenance

1. After inoculation into fresh medium, place the culture under optimal condition before it is relocated into the stock under suboptimal conditions. 2. Optimal post-transfer condition generally is elevated temperature (20°C or more), and elevated light intensity of up to double the stock maintenance intensities (20-60 μmol·m-2·s-1).

F. The agar slants should be placed in test tube rack. Labeling has to be in the following form:

1. Scientific name + strain number + culture medium + transfer date. 2. Also label each tube with transfer date if transferred at different time; label the whole rack with transfer date if transferred at the same time.

G. Watch out for overheating problem and temperature fluctuation caused by light/dark cycle.

H. Humidity is not allowed to be higher than 60% to avoid fungal growth.

VI. Culture Vessels Cleaning Methods

A. Immerse the vessels overnight in a neutral detergent bath (especially algal cells are strongly attached to inner surface of the glassware)

B. Scrub with a brush and sponge

C. Rinse several times with running water and rinse with deionized water finally

D. Dry in an area protected from dust