Chaos with Paramecium

Obtain one or more Chaos amoebae in a small volume of culture medium as described in section I of the Appendix. Place the drop in the center of a clean slide and confirm in the dissecting microscope that the transfer was successful. An ameoba can be readily seen with the unaided eye as a bright white spot in the drop. Add one drop of concentrated Paramecium.

The wet mount must be prepared carefully otherwise the amoeba will be crushed. Make an especially high Vaselineª "ledge," and add additional culture medium to the drop if necessary so that the coverslip will contact medium without being pushed down far enough to crush the Chaos. Gently press down the coverslip starting at one edge, trying to keep the amoeba to the center of the chamber. Once contact has been made, keep an eye on the amoeba itself. Stop pressing down once the coverslip has made contact with the cells. If a thinner preparation is desired, it might be possible to press down further once the cell has begun to spread on the surface.

This preparation should be observed immediately if predator/prey activity is to be seen. Paramecium will congregate around the Chaos, and eventually be engulfed. It is not unusual to see one cell with a half dozen or more food vacuoles with Paramecium moving in them.

Chlamydomonas deflagellation and experimental design

Label 15 ml Falcon or Corning capped disposable centrifuge tubes appropriately, and to two of them add 30 mg each dry colchicine. Prepare a stock solution of 1 mg/ml cycloheximide or emetine in spring water (latter should be protected from light). The deflagellated cultures will consist of deflagellated Chlamydomonas with 3 mg/ml colchicine, 10 µg/ml cycloheximide or emetine, or no treatment (positive control). For nondeflagellated cultures, suspensions of motile cells will receive the same treatments.

Deflagellation by pH shock

Start with 100 ml of a rather dense culture of motile Chlamydomonas. While stirring vigorously and monitoring pH, use 0.5 N acetic acid to lower pH to 4.5 within 30 sec. Let the culture stir for 30 sec. to 1 min. Quickly bring the pH back to 6.8 with 0.5 N KOH. Distribute in 6 x 15 ml tubes and centrifuge at room temperature, full speed in a model HN (swinging-bucket rotor). Sample cells during the centrifugation to determine if the deflagellation was successful. Drawback - cells often don't re-grow flagella.

Decant the supernatants by pouring. Flagella are poured off with the supernatant. Resuspend and combine two pellets in a total of 10 ml spring water, add to one of the colchicine tubes, and mix. Resuspend and combine two more pellets as above, add 100 µl 1 mg/ml cycloheximide or emetine, mix. Resuspend and combine the remaining two pellets as above for the positive control.

Mechanical deflagellation

An alternative deflagellation procedure that is more reliable has been use of a Polytron type tissue homogenizer to mechanically amputate flagella. On the Tekmar device, homogenization of 50 ml cells in a Potter-Elvejhem vessel at setting 70 for 30 sec was effective in removing flagella from 95% of cells. Partial flagella do remain, however.

Add inhibitors to suspensions of motile Chlamydomonas from the same original culture as above. Keep in 50 ml flasks to provide a good surface area, in good light (avoid heating the cultures with artificial light).

Naegleria transformation experiments

Prepare suspensions of amoebae or cysts from plates prepared as described in the culture instructions for Naegleria. Use spring water or 0.1M Tris-Cl, pH 6.5. Cysts tend to adhere to the agar surface. They can be hosed off of the surface with a jet of water from a pasteur pipet or automatic pipetor, or removed by adding water to the plate and using a rubber policeman to free them. Cysts should be relatively bacteria-free, so one wash should be sufficient to prepare them for an experiment.

Amoebae are easier to remove than cysts. Mass plates in which plaques are just barely appearing provide the best yields, however a wire loop can be used to remove amoebae from an edge plate. Either way, the better preparations start off with a lower density of bacteria. Amoebae should be washed one or two times by centrifugation at half speed for a couple of minutes in a clinical centrifuge, and a cell count performed.

Conditions that promote differentiation include suspension in liquid and absence of bacteria (starvation). The trigger has not been precisely identified, however. In fact, the type of substrate and the cell density appear to affect the process as well. The differentiation process itself is fascinating to watch. Experiments can be designed to study the trigger also.

To follow the differentiation process we prepare a wet mount of suspension and examine the cells in dark field at 40 to 100x and at 400x in phase contrast. We periodically tally up numbers of cysts, active amoebae, and, later, flagellates. Individual amoebae can be seen to contract to the point that they superficially resemble a cyst, then they begin to wiggle and/or spin in place as flagella develop. The fully differentiated form is elongated (football-shaped), has two flagella equal in length to the long axis of the cell, and is very motile. Motile forms come up from the surface of the slide, so at 400x they are easily missed. Mounts should be periodically examined at both low and high magnifications.

Experiments can be performed by varying cell density, concentration of bacteria, and treating the suspensions various ways. We have performed the Chlamydomonas flagellar regeneration experiment on Naegleria with interesting results. Colchine treatment actually appears to stimulate differentiation, and flagella fully develop in the transformants.

Paramecium with stained yeast

Stained yeast show up well in the microscope in dark field yet are also visible in bright field. When ingested by Paramecium they permit observation of the acidification of food vacuoles, as the dye color changes from red to purple as the pH drops. For our purposes 10 ml of suspension is more than sufficient.

The original recipe called for 30% compressed yeast and 0.3% Congo red dye in distilled water, with gentle boiling for 10 min. We have had great success using either 3 grams dry yeast or several loopfuls of plate-grown yeast and 0.3 grams dye in 20 or 30 ml distilled water in a small (50 ml) flask. A stopper is dropped into the flask but not pressed into place, and the flask is placed in a beaker with water poured up to the level of yeast suspension. The preparation is heated for at least 10 minutes in a microwave oven set to "simmer." The volume can be reduced by removing the stopper and heating without boiling - otherwise cavitation makes a mess of the inside of the microwave oven.

Obtain concentrated Paramecium as described in part I (biological materials). Pipet a drop or two onto a clean glass slide, preferably under a dissecting microscope at low power (5 to 7 x). Confirm the presence of a large number of cells, and add a drop of cooled stained yeast suspension. Yeast should be added side by side and partially mixed so that the drops overlap, with a portion of the Paramecium drop devoid of yeast.

Prepare a normal wet mount, pressing gently on the edge of the coverslip in closest proximity to the liquid, then working around the perimeter. With practice, a thin layer of suspension can be sandwiched between slide and coverslip without squeezing out the suspension and without leaving a large air bubble.

After a few minutes the cells should congregate along an edge or in a corner of the chamber. They can be readily observed at all dry magnifications when quieted down. It is acceptable and may be benefical to prepare these slides well in advance of a lab session. Progression of food vacuoles to the purple color may take several hours.