Lab notes · Physarum polycephalum
How to Grow Slime Moulds
The purpose of this document is to help others learn how to culture, maintain, and experiment with slime moulds (Physarum polycephalum).
Solid culture
Agar Culturing
Materials required:
- Petri dish
- Agarose (molecular biology grade, if possible)
- Parafilm
- Slime moulds – liquid culture or agar culture
- Autoclaved oats
- Sterile double-distilled water
Procedure:
- Make a 1.5% agarose gel by mixing 1.5 gms of agarose in 100 mL of sterile double-distilled water.
- Autoclave the mixture to make it sterile (pressure cookers can be used for this purpose).
- Pour it into Petri dishes and let it set. Do this in a laminar air flow hood or a still air box to try to reduce the contamination.
- Spread out some autoclaved oats on the agar surface.
- Cut out a small piece of the slime mould from a previous agar culture or a filter paper culture. If you are reviving the slime mould from a filter paper culture, place the cut piece on oats and add a few drops of water to hydrate the slime mould (refer below on how to make filter paper cultures)
- Close the Petri dish and seal it with Parafilm to prevent it from drying out.
- Keep the slime mould at 25°C in a dark space.
- You should get good growth in a few days. Try playing around with how much food you provide to see what works for your purpose. Add more oats if you see slime mould has covered all the provided oats. It’s a good practice to subculture the slime mould every week or, if you find your slime mould has overgrown, to maintain a healthy stock.
Long-term storage
How to make filter paper cultures?
Materials required:
- Petri dish
- Sterilised filter paper
- Parafilm
- Slime moulds – liquid culture or agar culture
- Autoclaved oats
- Sterile double-distilled water
Procedure:
- Cut a sterilised filter paper to make it fit inside the Petri dish.
- Use enough water to wet the filter paper, around 2 ml.
- Place some autoclaved oats on the filter paper.
- Inoculate the slime mould onto oats on the wet filter paper.
- Seal the Petri dish using Parafilm.
- Let the slime mould grow in the dark at 25°C for 2–3 days.
- Once it has spread around more than half of the filter paper surface, remove the parafilm and keep the slime mould in the incubator. This should lead to moisture loss and the filter paper and slime mould drying up.
- When the filter paper and slime mould are dry, seal the Petri dish with parafilm and keep it in a dry and dark place at 25°C for long-term storage.
- Some slime mould filter paper cultures were kept in -80 for long-term storage.
- The lifetime of filter paper cultures is supposed to be around a year. From the data that I have so far, the filter paper cultures
Scaling up
How to make liquid cultures?
Composition of 100 ml GPH (Glucose-Peptone-Hemin) media:
| Ingredient | Amount |
|---|---|
| Glucose (G) | 1 g |
| Peptone (P) | 1 g |
| Hemin Solution (H) (3 mg/mL) | 150 ul |
| Distilled Water (MilliQ) | 100 ml |
Preparation of Hemin Solution:
- Weigh the hemin powder directly in the 15 mL Falcon tube. Stabilise the tube using a stand before weighing the hemin. Weigh around 15 mg of hemin.
- Dissolve Hemin in 5 mL of DMSO. The hemin concentration will be 3 mg/mL.
- Sterifilter the hemin solution through a 0.22 µm RC (regenerated cellulose) membrane filter.
Notes:
Do not use PVDF membrane filters, as DMSO is incompatible with them; the solution will go through the membrane, dissolving the membrane in the process.
This will be a time-consuming process. Make sure you have multiple filters and 1 mL syringes. Pour the solution into a Petri dish to make it easier to get the solution into the syringe.
- Wrap the Falcon tube with aluminium foil as both DMSO and hemin are light sensitive, and keep it in the 4℃ section of the refrigerator. Since the melting point of DMSO is 18℃, the hemin solution will be frozen. Keep the hemin solution outside to thaw for about 30 minutes before use. It is recommended to let it melt slowly.
Preparation of GPH media:
- Weigh 1g of Glucose (G) & Peptone(P) each in a 250 ml bottle and dissolve them in 100 ml of Milli-Q water.
- Adjust the pH of the solution to 6 by adding the appropriate amount of HCl.
Note:
If you prepare a 200 ml solution of Glucose (2g) and Peptone (2g), add 400 ul of 2N HCl to lower the pH from around 6.5 to 6. This method has worked well so far. Split the media into two 250 ml bottles with 100 ml each.
- Autoclave the media.
- Add 150 ul of 3 mg/mL hemin (H) solution to the media just before slime mould inoculation.
Inoculation:
- Maintaining Liquid Culture: If you are inoculating from an existing liquid culture, then aliquot 1 ml of slime mould into a 1.5 ml Eppendorf tube and let it settle. Cut the end of a 200 ul pipette tip (to minimise membrane damage) and take out 200 ul of slime mould for inoculating the new media. This should grow into a significant amount within 3 days of inoculation. You can adjust the amount of slime mould used for inoculation based on your requirements. 5 days is the recommended period of subculturing. Try to subculture within 7 days or before the media turns murky.
- Restarting Liquid Culture:
- For restarting the liquid culture, you can start by using slime mould grown on 1.5% Non-Nutrient Agarose (NNA)(molecular biology grade). If you have to revive a dried filter paper culture, then cut a small piece of slime mould containing paper and place it on the NNA and add ~200 ul of autoclaved water. Give enough oats for it to grow. It is recommended to let the slime mould grow for at least 3 days before inoculation. This is because if used before, it was observed that the slime mould turns white and dies. You can test whether slime mould will survive if inoculated from a culture less than 3 days old. A good rule of thumb would be to use slime mould that has turned orange. This is because the colour would be a better indicator of the physiological state of slime mould than time. When inoculating, try to avoid including oat pieces as much as possible.
- For a cleaner inoculum, you can start by growing them on a Glucose-Peptone (GP) or GPH agar – the same relative compositions as the liquid media with 1.5% agarose. Slime mould usually grows in a mat-like appearance on these nutrient agars. Wait for the slime mould to grow for at least 3 days. Scrape it out using the flat end of the spatula for inoculation.
Note:
This is a minimal media with no buffering agents. You can modify this protocol by referring to the articles mentioned in the references or by trial and error.
References:
- Adrian Fessel (2019). Topological dynamics of small-degree networks. Dissertation.
- Brewer, E. N., Kuraishi, S., Garver, J. C., & Strong, F. M. (1964). Mass culture of a slime mould, Physarum polycephalum. Applied Microbiology, 12(2), 161-164.
- Daniel, J. W., & Rusch, H. P. (1961). The pure culture of Physarum polycephalum on a partially defined soluble medium. Microbiology, 25(1), 47-59.