Cultures of Chlamydomonas reinhardi

Unialgal cultures are available as nonmotile cells on agar slant tubes from Carolina or WardÕs. To remain viable, the cultures must be maintained at room temperature (never refrigerate algal cultures), and should be propagated every few months at most. Transfer is by loop just as with a bacterial culture. Solid Chlamydomonas medium is complex and can be purchased from a supplier, although the catalog may not list the item.

For long term culture of motile cells, Chlamydomonas medium is recommended. However, we have observed a failure of the cells to grow at all in CarolinaÕ medium, while they did fine in spring water alone. Spring water is reported to be unsuitable for long term culture, but is recommended for growth of large numbers of motile cells for experiments. Enriched media, including the Carolina medium, promote cell division but leave cells unsynchronized and they reflagellate poorly in deflagellation experiments.

SagerÕs minimal medium has been recommended to obtain large quantities of cells for deflagellation/reflagellation experiments. It does require bubbling with 5% CO2 and shaking. Synchronizing the cells with light and dark cycles helps, but is not necessary.

To maintain cell cultures long term, keep in suspension in a brightly lit area in enriched medium, and change out the suspension now and then by aseptically transferring a small amount of suspension to a sterile flask with fresh medium. Cultures can also be transferred to solid medium for long term maintenance.

To grow motile cells for flagellar growth experiments use spring water or SagerÕs, allowing several days for development of a densely populated culture. A wide, shallow, sealed container is best to promote gas exchange. Keep in a brightly lit location, sunlight preferred.

Cultures of mixed freshwater organisms ("pondwater")

In appropriate weather, collect fresh samples from a freshwater source such as drainage ditch, river, creek, stagnant pool, pond, or lake. Include muck from the bottom, plant material including algae, and plant debris. A very healthy (i.e., green and stinky) pond will provide a wealth of organisms.

In cold weather or after heavy rains freshly collected samples may be disappointing. In old cultures, diversity may be stimulated by exposure to strong light and addition of a source of nutrients such as fish food flakes. The result will be a proliferation of bottom feeders, however, and the variety of photosynthetics will be severely limited. A pool (wading pool, tub) of fresh water spiked with samples from natural sources and maintained in strong light at room temperature may serve as a source.

Cultures of Naegleria gruberi (strain NEG)

Freeze-dried cysts can be obtained from the American Type Culture Collection. Order #30223 (Naegleria gruberi, strain NEG), and #3637 (Xanthomonas maltophilia, the food source). It takes a long time to get them to excyst from a freeze-dried preparation. Allow several days for activity to appear, and at least a week and a half if large numbers are to be grown.

Details on the biology and laboratory culture of Naegleria can be found in Fulton, Chandler, Amebo-flagellates as Research Partners: The Laboratory Biology of Naegleria and Tetramitus. Meth. Cell Physiol. 4: 341, 1970. The amoebae are propagated on agar plates. Two agar preparations are recommended, namely "NM" or "PM."

For NM plates, prepare (in grams/liter) Difco Bacto-peptone 2.0; Dextrose, 2.0; anhydrous K2HPO4, 1.5; anhydrous KH2PO4, 1.0; Difco Bacto-agar 20.0. Prepare PM plates in the same way, but double the concentration of peptone. Pour about 40 ml medium per 100 mm dish. PM is reported to grow a richer bacterial lawn, providing easier visualization of plaques, and faster growth of amoebae. Cultures are reported to be more heterogeneous than on NM, however. In our experience, there isnÕt much difference.

For large quantities of amoebae (mass plates), prepare a dense suspension (A650 of 0.6) of Xanthomonas maltophilia and spread a few hundred microliters over the plate. Prepare a suspension of amoebae or cysts and spread over the plate. Alternatively, mix the suspensions or simply harvest amoebae and bacteria from another plate and spread them together. We have not worked out appropriate concentrations of amoebae or volumes to use to obtain mass plates by a desired time.

Large quantities of vegetative amoebae are ready for harvest when the mass plate shows the beginning of plaque formation. Sparsely populated areas will be separated by ridges that upon inspection prove to be large quantities of feeding amoebae. When plaques begin to form, the amoebae in the bacteria-free regions will form cysts. Plates of viable cysts can be obtained by wrapping the plate with Parafilmª and leaving at room temperature until all cells have encysted. Plates should then be used within a few days or refrigerated for future use.

Propagation of a culture for maintenance of the cell line is best accomplished by preparation of an edge plate. Bacteria alone are spread on a plate and allowed to grow overnight at room temperature. The next day a loopful of amoebae should be obtained aseptically from an active culture and lightly streaked near one edge of the plate. Within a day a plaque will form at which time the plate should be sealed with Parafilmª and refrigerated. By stopping growth early, the number of generations is kept to a minimum and the likelihood of changes to the original strain due to selection pressures is kept to a minimum.

Edge plates can be pemitted to develop at room temperature and used as sources of either cysts or active amoebae, however quantities are lower and there is a risk of the plate drying out. Cysts remain viable for months, but the cultures die out when the plates are allowed to dry.

Plates often become contaminated with fungus. I have not tried fungicidal additives. To clean up the culture, I've done the following.

From dried cultures or old plates, the emergence of Naegleria is very difficult to predict. The best results seem to be obtained by depositing a suspension of cysts on a plate with bacterial lawn established. Keep the plate very wet, adding sterile water every other day. Plaques may take several weeks to show up. Edge plates usually become active within a day or two of removal from the refrigerator.

Naegleria often grow unpredictably, and it can be difficult to guarantee a pure population of cysts and/or amoebae in sufficient numbers, without experience. It is suggested that plates be prepared daily, in sufficient numbers for the class, starting at least a week in advance of the first planned laboratory session. Prior to the first lab session, resuspend a plate and check if it has a satisfactory ratio of cysts to amoebae. Use older plates if more cysts are needed, use younger plates if more amoebae are needed.

Cysts are very obvious to the initiated, at all magnifications at or above 40x. They are especially obvious in dark field mode. Cysts are round, 10 µm in diameter, and appear dark in bright field viewing, very refractile in dark field. Amoebae are recognizable to the trained eye in all modes, but require high contrast, dark field, or phase contrast to be recognized. They are of the same average diameter as cysts but are more irregular. Confirm the viability of a culture by looking for viable amoebae in a suspension at 400x, phase contrast.

Cultures of Paramecium caudatum and/or multimicronucleatum

Cultures purchased from a supplier are usually ready for use immediately. Cultures can be maintained indefinitely, however extreme care must be taken to prevent contamination with a population of rotifers. Rotifers are ubiquitous, and can be introduced to cultures on pipets, by directly handling wheat seeds, and possibly by using culture jars that were not sufficiently cleaned. Unfortunately, cultures from suppliers often come contaminated.

Expand cultures by swirling a culture to mix the contents and pouring some of the water into a fresh jar. Add spring water and one or two boiled wheat seeds. Periodically change some of the water, remove old husks and debris, and add new wheat seeds.

Paramecium tend to congregate around the food source, so that populations are the most dense among the debris around the wheat seeds or the edge of the culture jar. In the most active cultures the population becomes dense throughout the medium. To harvest the cells one can carefully place the culture under a dissecting microscope so as not to disturb the contents, and collect Paramecium from the most densely populated areas. Frequently they are sufficiently concentrated "as is." To concentrate them further, centrifuge a volume of culture water that has been sampled this way, using a clinical centrifuge. Two or three minutes at 200 x g or so brings them down without harming them. The cells can then be re-suspended in a smaller volume for use in wet mount preparations.

After harvesting, simply replace the spring water that was removed and replace the wheat seeds if necessary. Cultures should be rejuvenated now and then by pipetting cells in a fairly large volume of water into a fresh jar and adding fresh spring water and wheat seeds. Carolina does cell medium that is especially prepared for culture of Paramecium, but this may not provide any advantage over spring water and is more expensive.

Cultures of Pelomyxa (Chaos) carolinensis

A recommended source of cultures, water, and food is the Carolina Biological Supply Co. Obtain well in advance, since the protists may be fragmented or killed during transportation, especially during the summer months. Cultures can be kept indefinitely, and numbers of ameobae increased by expanding the number of cultures and feeding with Paramecium.

To simply maintain viable cultures, replace about half the water with spring water once or twice a month, add a fresh boiled wheat seed, and remove old husks or other debris. Amoebae should survive on the food chain from the wheat germ alone, however they will do better when given a pipetful of Paramecium now and then.

Amoebae are best identified by placing the open jar or fingerbowl on an opaque surface under a dissecting microscope at low power, with strong illumination from the side, angled down. Amoebae appear white and opaque, often with obvious pseudopodia. They will be on the order of 0.5 to 1 mm or more in diameter.

To separate amoebae from accumulated algae and/or debris, swirl to create a gentle vortex and examine the center of the jar in a dissecting microscope at low power, with illumination from the side. Amoebae are denser than other materials. They tend to drop to the bottom quickly and accumulate in the center of the culture jar, from which they can be collected using a pipet. This method can be used for obtaining individual amoebae for slide preparations, in fact, with practice one can obtain an individual amoeba with very little accompanying water in the tip of a pipet.

To expand cultures, swirl a culture to mix contents and immediately pipet or pour some of the water into a fresh container, either a fingerbowl or clean widemouth culture jar (old plastic jars from suppliers work just fine). Alternatively, let amoebae accumulate in the center and remove them individually.

Drop amoebae in a fresh jar with spring water and one or two boiled wheat seeds. Add Paramecium for food until a food chain develops. To obtain large numbers of amoebae for a class, check the cultures frequently and maintain fairly high numbers of Paramecium (they should be readily visible throughout the culture). Extra cultures that will not be maintained indefinitely do not require wheat seeds, provided the population of Paramecium is maintained by frequently additions.