Culture Medium

In the Chlamy Center, nearly everything is kept on 1.2% agar slants containing Sueoka’s high salt medium with added acetate and yeast extract (we call this YA medium). The reasons for this are: a) all the strains will grow on it, including auxotrophs like arg and nic mutants; b) it’s easy and relatively cheap to make; c) it’s an enriched medium, so if you get contamination with fungi or bacteria, it will be apparent immediately.

This last point is very important. Bacteria can grow along with Chlamy on acetate or minimal medium without being detected. Some bacterial contaminants are very hard to get rid of, because they are sticky or rapidly motile, so it is far better to avoid getting them in the first place. Fungi are usually apparent on YA or acetate medium, but can lurk on minimal medium without showing up to a casual inspection. The quickest way to determine that a culture is free of living contaminants is to streak it extensively on an agar plate in the presence of acetate and yeast extract. The difference between a Chlamydomonas colony and a fungal or bacterial colony is obvious using a dissecting microscope within 4 days, and to the naked eye in a week or two.


Light, Containment and Temperature

Slants are stored in numerical order on a wire shelf in 4 x 10 epoxy coated steel wire tube racks at 22°C on a 14:10 light:dark cycle. Tubes are labeled with Avery® address labels with the strain name and date passed. Strains that can be maintained in the light are kept at ~40 μmol m-2 s-1 PPF under cool white florescent lights. Many photosynthetic mutants (dims and darks) are stored in either dim light (~2.5 μmol m-2 s-1 PPF) or complete darkness in black plastic tubs with lids.


Making Slants

The slants are made from 20 x 125mm screw-cap disposable culture tubes (we wash and reuse the caps) in Nalgene® Unwire tube racks (for 20 mm tubes). They are made with 9 ml of agar and slanted when they come out of the autoclave after final sterilization. This provides a larger surface area from which you can remove loopfuls of cells for transfer. Stab cultures are admittedly a little quicker to transfer, but they don’t give you nearly as good a volume of available cells to sample from. Also, it’s easier to inspect slant cultures to see whether they need to be transferred soon or are contaminated.

On slant making day, we prepare 6 liters of media and use a peristaltic pump, Tygon® tubing and a 16 cm stainless steel tube with a curved tip to dispense the agar. To make a smaller number of slants, prepare 1 liter of medium in an Erlenmeyer flask and autoclave just long enough to melt the agar. With our autoclave, it takes 15 minutes (plus slow exhaust). Although you can melt the agar on a hot plate it could easily boil over if not watched continuously. Dispense 9 ml aliquots into the tubes. This goes faster if you use a 25 ml disposable pipette with 9 ml increments marked with a Sharpie®. With our Fisherbrand® 25 ml disposable pipettes, we can dispense 36 ml with each pipette fill. Then cap, autoclave for 45 minutes and immediately tilt the racks to create the slants. We use a long, 1 cm thick piece of plastic that we lay the top of the tube racks on. It takes some practice to get exactly the right angle; what you want is for the agar to reach about 1-2 cm below the top of the tube. Let the tubes age for two weeks before using them, otherwise the agar surface will be wet and the Chlamy don’t seem to like this. Also, if the tubes were not autoclaved properly, this will give time for contaminants to grow up and become visible.


Preventing Loss of Cultures

We maintain two copies of each strain on two different floors at the University of Minnesota on the St. Paul campus that is sufficiently far apart that a fire, mite infestation or other disaster striking one would be unlikely to hit the other. High-temperature cut-off switches are installed in the walk-in growth chambers that will turn off the lights to prevent overheating if the air conditioning fails. The primary and secondary cultures are passed on a staggered schedule so they are not the same age.

Cultures are inspected on a regular schedule. Dims and darks tend to be finicky, so they are inspected more frequently. Some dark strains are passed once a month because many of these cultures are white and it is difficult to determine if they are alive or dead.


Transferring Cultures

In the Chlamy Center, depending on the strain, cultures typically last 2 to 12 months. Cultures need to be transferred to new media if they are:

  • changing from bright/dark green to olive drab
  • getting very pale
  • starting to bleach out at the top of the agar
  • drying out (agar shrinking substantially)

We go through the tubes one rack at a time and pull out anything that looks like it needs transferring and put it in a separate rack. This rack is filled with 20 tubes to be transferred, leaving two empty rows for the new slants. Labels are made and attached to the new tubes before they are transferred and they are kept paired in this rack until the new cultures have grown up. We usually allow one week for this before putting the new cultures away. The old cultures are then autoclaved and discarded.

New and old tubes are checked three different times to ensure they match:

  • when the label is put on the new tube,
  • when the cells are transferred from one tube to another, and
  • when the new tube is put back in the rack in place of the old one.


Cleaning up contaminated cultures

If, despite your best efforts, a strain gets contaminated, here’s a suggested triage system:

  1. Check a backup culture if you have one; if you have a good one, use it as a replacement and discard the contaminated culture.
  2. If you do not have a clean replacement copy, proceed to cleanup efforts.



For strains heavily contaminated with bacteria, streak the culture on the most minimal medium on which the strain in question will grow; e.g., if it will grow on minimal, use HS or Tris-minimal; if it requires acetate, use HSA or TAP but omit the yeast extract. The idea is to enrich the Chlamy relative to the bacterial population before proceeding to the next step

If the contamination is slight, check under a dissecting scope to see if there are any uncontaminated areas, and try to recover a clean colony from there.

If there is slight but uniform contamination (Chlamy is still bright green, seems to outnumber the bacteria), then streak several plates of YA medium with it. Make one line across the plate with a wire loop or toothpick. Then, under a dissecting scope, use another sterile toothpick or glass needle to streak some Chlamy-rich areas perpendicular to this line, in hopes of getting a single colony that’s bacteria-free.

Revisit this plate daily, looking for green colonies. Continue to use a glass needle to try to chase the Chlamy away from the bacteria. If it’s apparent that the bacteria are overtaking the plate, you may need to use another tactic.

We don’t like using antibiotics to clean up cultures. It’s not an environmentally friendly practice, and often it doesn’t work very well either. However, if the enrichment technique mentioned above isn’t helping, sometimes a round on antibiotic medium will help and will at least buy you some time before the Chlamy are killed. Ampicillin is the best choice (doesn’t phase Chlamy at all). Neomycin is another one that doesn’t kill Chlamy but gets a lot of bacteria. Tetracycline is another possibility, but is inactivated by light so that’s somewhat incompatible with use on a photosynthetic organisms. If you happen to be working with a Chlamy mutant that’s already resistant to paromomycin, streptomycin, erythromycin etc., of course you can use that.

Another enrichment trick for Chlamy strains that can swim is to use phototaxis to concentrate them at the top of a liquid culture, then plate out cells from there.

Bottom line: patience, a dissecting scope and a glass needle usually works for bacterial cleanup. The other enrichment steps just facilitate your getting to the point where this is effective and rarely result in complete removal of the contaminant.



Fungal contaminants are actually easier to remove than bacteria, providing you catch them in time. However, many fungi seem to produce compounds that are toxic to Chlamy, so you need to work quickly.

You don’t want to shed fungal spores over your usual transfer area, so I recommend doing this first step at a lab bench in some other room where it won’t contaminate anything else.

If the plate is heavily and totally contaminated, use a sterile toothpick to wipe away some of the fungus from above a green patch of Chlamy. If you’re dealing with a plate that has some fungal colonies but not a solid mass, you can skip this step.

Use another toothpick to streak a Chlamy-rich suspension onto a fresh YA plate.

Leave this plate overnight. What you want is for the fungal spores that you inevitably transferred along with the Chlamy to germinate, but not to allow time for the fungus to sporulate again, so don’t let it go for several days.

Using a glass needle under the dissecting scope, gently chase some Chlamy cells down the outgrowing hyphae and collect them into a pile.

With a fresh toothpick, pick up this pile and streak it on a fresh YA plate.

Usually this is all you need to do, but be sure to watch that last plate closely for a few days to make sure the fungus doesn’t return.



If you find a tube or plate contaminated with several different types of bacteria, especially if the contamination seems to follow trails across the agar, you probably have mites.


Once these things get established, they are nearly impossible to eradicate. Don’t even think about using Parafilm® to prevent cross-contamination. Mites don’t care. They can also crawl between tubes if the tops are not screwed on tightly. We have found that screwing caps on tightly is helpful in slowly mite invasions.

If you have a backup of the same strain, dispose of the contaminated one by autoclaving. If you don’t have a backup, examine the plate or tube under a dissecting scope, pick off several of the cleanest looking Chlamy colonies you can find and put them on fresh plates. Make sure you don’t transfer any mite eggs along with them. (These are somewhat elongate, smooth microscopic blobs). After you’ve done the transfer, look at the new plates under the scope and make sure there’s nothing you don’t want on them. Then autoclave the original culture. Keep the new plates under quarantine and look at them daily; otherwise, treat them as for bacterial contamination above.

Remove everything from the shelf that the mite-infested plate was on and clean it with hot soapy water, Lysol®, and anything else you may have on hand.

Examine every tube or plate from this shelf under the dissecting scope to make sure you don’t have any more mites. Usually if there’s one, there will be more.

Continue to inspect these tubes and plates daily. If you have a real infestation, you will quickly become familiar with the life cycle. The eggs gradually develop some texture and then hatch into six-legged larvae. The larvae then undergo a metamorphosis into small eight-legged creatures (technically called “protonymphs”). These get bigger and go through two more stages to become adults. The specialist distinguishes these latter stages by the number of genital papillae they have; you don’t want to know this, you just want to get rid of them. The whole process takes a couple of weeks.

Also try to identify the source of the mites, such as from soil used by a nearby lab, or Drosophila flying in uninvited from a colleague’s lab. Mites like to eat fungi and they like humid conditions.