Slime Mold Culture and Use Prepared by W. J. Sundberg

Plant Biology, SIUC

This information is provided for participants in the Microbiology for High School Teachers workshop for use in their classrooms and may not be published or presented elsewhere. For questions please contact the author, Dr. W. J. Sundberg.

The slime mold (Myxomycete) Physarum polycephalum is a very useful teaching tool. Its plasmodial stage (see accompanying life cycle diagram) is easy to grow and manipulate with a minimal amount of equipment. With a minimum of effort, it can be preserved and stored in the dormant sclerotial stage (see accompanying life cycle diagram) for use in subsequent semesters.

With slime mold, one can dramatically illustrate cytoplasmic streaming, a morphogenetic shift from the growth to the reproductive phase, and differentiation (of sporangia). If desired, the plasmodium can also be employed to test the rates of migration and the effects of various chemicals and other environmental parameters on streaming and migration. Further, spore germination and motile (flagellated) cell structure and function can be studied with the microscope.

I. Culture of the Plasmodium (Modified Camp Culture)

The technique described below can be used to produce large amounts of plasmodium in a short time (approximately one week). The resulting culture serves as a source for (a) the subsequent manufacture of new "starter" material (sclerotia) for future use and (b) inoculum material for various subculture procedures described for class use herein.

Materials needed

- Covered container (e.g., large turtle dish, refrigerator crisper, plastic ice cream container, etc.)

- Small turtle bowl or finger bowl, about 2 inches tall, heavy enough not to float

- Paper toweling (preferably white; filter paper can be used but paper towering offers greater tensile strength -and other advantages)

- Old fashioned oatmeal flakes (not instant, precooked, or pretreated)

- Tap water

- Plasmodial sclerotia (the dry, dormant form of the vegetative phase). Initially obtainable from biological supply houses.

Adapted in part, with the authors' permission, from: Miller, D. M., and J. D. Anderson. 1971. Migration and biopotentials in slime mold plasmodia. Experiments in Physiology and Biochemistry 4: 183-202.

Procedure

1. Invert the small bowl and cover its bottom with a paper towel. Hold under running water to wet and wash the towel.

2. Place the small, paper-covered, inverted bowl into the larger container. Add a small amount of water--to a depth of about 3 mm--to the large container, thus creating a moat. The paper toweling on the small bowl must contact the water. Excess water promotes undesirable bacterial growth.

3. Place a piece of the sclerotium on or near the center of the paper-covered, inverted bowl. Subsequent cultures, if desired, can be started from small explants of the growing plasmodium.

4. Cover the large container to maintain the humidity and place in the dark at room temperature (20-26°C). Activation of the sclerotium takes about 3-5 hours.

5. Once the plasmodium (or at least most of it) has migrated off of the piece of inoculum paper (usually 4-12 hours at most), remove and discard the inoculum paper to reduce the potential for contamination. Add 1 or 2 oat flakes at or near the advancing front. Cover and incubate as before (see step 4).

6. After 24-48 (72) hours~, remove the cover and gently wash the culture. The purpose of this wash is to remove excess bacterial build up and excess slime material and other waste products produced by the slime mold.

To wash: (Do in subdued light, not in window light) Use a gentle stream of tap water from a faucet , with a nonsplash attachment. It may be helpful or more useful to use a short length of rubber tubing attached to the faucet. Wash well by removing and holding the inverted paper towel-covered bowl under the water stream and allowing the water to run over the culture and toweling. Larger cultures require longer and more thorough washing times. Be patient with this step. Drain for 5-10 minutes.

7. Add more oats. Place the oats on the advancing front if possible. The amount of oats used is dependent on the size of the culture--the larger the culture, the more oats that should be used. If too few oats are added, rapid migration occurs. However, overfeeding enhances the chances for contamination.

8. Rinse the large container several times, add fresh moat water, and replace the small bowl. Cover and incubate as above (see step 4).

9. Wash and feed the culture daily. A well developed culture should obtain in 3-4 days and can be maintained for 1-1/2 to 2 weeks or more. If growth is not vigorous, wash twice a day (feed once). Washing vigorously is the "trick" to maintaining large cultures. If the plasmodium migrates down into the moat, wash away all mold in or on the moat water.

II. Production of New Sclerotia for Storage

Using a clean, well developed, paper towel (Camp) culture, the dormant form of the vegetative stage, called a sclerotium (see accompanying life cycle diagram), can be produced. Because they take up a minimum of space, can be easily revived (often up to a year or more later if kept refrigerated), and develop directly on revival into new plasmodia, the sclerotia represent a simple mechanism of live specimen storage and retrieval. A simple method for sclerotium production is summarized below.

1. Prepare a paper towel (Camp) culture as described in Part I above.

2. Carefully remove the oat flakes. They can then be used as inoculum to start another paper towel culture.

3. Wash and drain well the plasmodium left on the paper towel.

4. Remove the paper towel from the finger bowl and cut away most of the excess paper from the margins. Temporarily place the remaining paper with plasmodium up on a clean newspaper or a clean paper towel in order to remove some additional moisture.

5. Clean and dry a large container.

6. Place the now-trimmed paper towel on supports (small beakers, etc. are useful) in the large container so that the paper can slowly dry out via evaporation. The margins of the paper should be free--not touching the supports or the sides of the large container.

7. Secure the cover of the large container leaving a fairly small opening (tilt the cover) so that slow evaporation will occur. Slow drying is the "secret" to sclerotial formation.

8. Incubate in the dark. As the paper dries out, the plasmodial mass will condense and finally sclerotize.

9. After 1-3 days (when dry and sclerotized), remove the paper. Cut away most of the excess. Place the dry sclerotium in a clean petri dish or similar container for storage. If the sclerotium is large, it can be carefully cut into smaller pieces. Store in a refrigerator (do not freeze) until needed again.

III. Preparation of Plasmodial Cultures for Observation of Cytoplasmic Streaming and Amoeboid Movement

The slime mold is one of the best organisms for demonstrating or observing cytoplasmic streaming. Its flow is rapid, relatively easy to see, and bidirectional (= shuttle flow). To study cytoplasmic streaming and to make large numbers of

short-lived cultures for individual student study, the following procedure is recommended.

1. Prepare non-nutrient 2-3% agar (= water agar) petri dishes. If they are to be kept (stored) or used only for 1-3 days, although cleanliness is required, sterilization should not be necessary.

2. Add some plasmodium (prepared as in Part I above) to the solid agar surface. Although inoculum C below has worked best (produced the largest and most vigorous and lasting migrating plasmodia) for us, any of the following inocula can be used:

A. Small amounts of slime mold that have migrated on to the moat water surface

B. A piece of the paper with slime mold on it cut from

the side of the bowl

C. An oat flake with the slime mold growing on it

3. Cover and incubate in an upright position in the dark for 12-24 hours before use. In order to enhance migration, the cultures should be kept covered and in the dark when not in use. Overexposure to light may reduce plasmodial activity.

Alternatively, by using a very thin (less than one mm thick) layer of agar in the petri dish, streaming may be observed by inverting the dish (with cover in place) and viewing through the plate bottom and agar with the microscope using its lower power objectives.

IV. Observation of Plasmodial Cultures

To study cytoplasmic streaming and amoeboid movement (the latter at the advancing front), petri dish cultures prepared as in Part III above are most effective. Remove the petri dish cover and observe with a compound microscope at lower magnifications. With proper care and if the working distance is not too short, even the high dry lens can often be used for short periods. However, it fogs due to the humidity with extended and continuous use. A dissecting microscope with transmitted light will also work, but is not as effective due to its generally lower magnification power.

The slime mold exhibits shuttle (bidirectional) flow or cytoplasmic streaming. The average length of time that flow occurs toward the advancing front and away from the advancing front, as well as that between reversals, can be determined with a stop watch.

If a micrometer is available, a rough estimate of distance traveled in one direction by the cytoplasm can be calculated. Procedure: Measure the time it takes an identifiable particle to travel from point A to point B--a known distance--on the micrometer scale. Divide that figure into the total average

time of unidirectional flow (in that direction). Multiply the result times the distance traveled from point A to point B on the micrometer scale. Result: approximate total distance traveled in one direction.

Observations on cytoplasmic streaming can be used to initiate discussions of general topics such as:

A. hypotheses invoked to explain cytoplasmic streaming

B. hypotheses invoked to explain amoeboid motion

C. viscosity and its effects on flow

D. surface resistance and its effects on flow in tubes (applies equally well to slime mold "veins", veins and arteries in the circulatory system, garden hoses, pipes, etc.)

V. Studies of Plasmodial Migration

The following procedures can be used to study various aspects;

of plasmodial migration (distances moved, rates of movement, etc.); effects of environmental factors on movement, etc.) at the macroscopic level.

A. The Petri Plate Method

1. Prepare and incubate petri plates of plasmodium as outlined above (see Part III)

2. After 12-24 hours and using a wax pencil, outline the position of the advancing front of the plasmodium.

3. Incubate in the dark again. The period of time can be adjusted to fit a class schedule, but should be at least a few hours and if possible, should not be longer than 24 hours.

4. Outline the new position of the advancing front and compare the difference. Qualitative differences will be obvious. Rough quantitative data (distance of movement, rate of movement, etc.) can be obtained via measurement or calculation. However, because of the lack of directional control of movement, and the variability thus introduced, the quality of such data is questionable. (It would provide a good vehicle for initiation of discussions on experimental design and the importance of control of variables).

B. The Race Track Method

This technique offers a more controlled system for studying migration. It relies upon the fact that the advancing front of the plasmodium will neither migrate

across waxed surfaces nor back over the same surface a second time.

1. For inoculum, prepare an actively growing Camp culture as described in Part I above.

2. Prepare one or more "race tracks" as follows:

  A B C

a. Fill the bottom of a flat tray

or pan (one with a cover, like

a rectangular cake pan) with

non-nutrient, 2-3% agar (water

agar) and allow the agar to

solidify. If kept covered and

used in 1-3 days, although

cleanliness is required,sterilization

should not be necessary.

b. Place single parafilm strips (waxed paper should also work) on the agar surface along the entire length of the pan at A, B and C (see diagram above). The remaining exposed linear strips of agar are the "race tracks". Their width and number per pan can be controlled by varying the width of the parafilm strips.

3. Inoculate the race tracks by placing a piece of paper toweling covered with slime mold (part of the Camp culture) on the agar at one end of the track (see diagram above at A). DO NOT FEED. Incubate in the dark at room temperature.

4. As soon as the plasmodium (or the major portion of it) has migrated off of the inoculum paper (toward C in the diagram), remove the paper. DO NOT FEED. Mark the position of the advancing front by marking the waxed paper or by sticking a clean dry piece of toothpick vertically into the edge of the track.

5. Incubate in the dark as before (see Part III-3).

6. Mark the new position(s) of the advancing front(s) and measure the distance traveled. Alternatively, one could allow the plasmodia to "race" the entire distance down the tracks stopping the exercise when the first one reached the end. (Remember, bookmaking is illegal'). In either case, migration rates can be easily calculated.

Some of the variables that affect migration rates and might be experimentally manipulated and tested include:

a. temperature

b. width of "race track" (size of advancing front)

C. A Modified Race Track Method for Testing the Effects of Chemicals on Migration

The effect of chemicals (ions, organic compounds [such as caffeine, etc.], etc.) on migration can be tested with a slight modification of the race track method as described below.

1. Prepare inoculum and race track(s) as described above in part V-B, 1-2. Use at least two (preferably four) tracks--one (or two) as experimental track(s) and one (or two) as control track(s).

2. Using a sterile scalpel , cut out A B C

and completely remove all agar

from a small part (across the

entire width) of each race track.

Sterilize the scalpel by dipping into

95% alcohol; then flame and let

the alcohol burn itself out. Let cool

slightly. Repeat this process three

or more times. Allow to cool at least

30 seconds or more before use.

3. P1ace the agar removed from

each track in a separate heat

resistant test tube.

4. To the test tube, add a few

ml of a solution containing:

a. the chemical to be tested--for experimentals

b. the same solvent without the chemical to be tested--for controls

5. Gently heat the test tubes (in a water bath or with a Bunsen burner) until the agar is melted and the chemicals are mixed with it.

6. Pour the agar back into the trough created by initial removal of the agar and allow it to solidify.

7. Inoculate and incubate the race tracks as in Part V-B, 3-5.

8. Observe and record the effects of the modified agar(s) and their chemical contents (in the troughs) on plasmodial migration.

VI. Differentiation of Sporangia

Under the proper environmental conditions--including slow drying and exposure to light, the plasmodium will shift from continued vegetative growth to reproduction. This morphogenetic shift results in the formation of many multilobed, spore containing sporangia (see the accompanying life cycle). Once mature, the intricate structure of the sporangia and the spores can be studied, if desired, with the microscope. The procedure described-below can be used to readily demonstrate the sporangial development process.

1. Prepare an actively growing Camp culture as described in part I.

2. Carefully remove the oat flakes (they can be used to start another Camp culture).

3. Place the paper towel and any plasmodium on it (either still over the inverted bowl or draped over a beaker or some other support) in a clean and dry large container.

4. Cover the large container leaving a fairly small opening so that very slow evaporation will occur.

5. Incubate in the light at room temperature. Sporangia will begin to form and mature within 1-2 days.

VII. Study of Spore Germination and Swarm Cell Structure and Movement

Although small in size, it is possible to study some facets of slime mold spore germination with the aid of a compound microscope equipped with a high dry or oil immersion objective or both. Likewise, the structure (flagellum, nucleus, nucleolus, contractile vacuole, etc.) and activity (swimming and amoeboid motions) of the flagellated product of germination--the swarm cell--can also be observed.

1. Use the sporangia produced as described in Part VI as the spore source.

2. Free the spores by carefully placing the sporangia in a vial or small bottle. Cap the bottle and shake vigorously to break the fragile sporangial walls and release the dark powdery spores. Remove the large pieces of sporangial remnants. The spores can then be stored and will retain their viability for some time (often a year or more) in a small clean vial.

3. To obtain spore germination:

a. Place some of the spore powder in a dropping bottle with clean distilled water (about 1/3 full). Depending on the additives in it, tap water might also work.

b. To wet the spores, shake the capped dropping bottle vigorously several times over a period of 5-10 minutes.

c. Incubate at room temperature.

d. Prepare and examine fresh water-mount slides using some of the debris at the bottom of the bottle. Test at intervals of 12, 24, 48, etc. hours to determine when spore germination occurs.

The best clue that germination is occurring or has recently occurred will be the presence of numerous, apparently uniflagellate, hyaline motile cells—the swarm cells. (These cells actually have two flagella but one is so short that it is not usually seen).