My cells won't grow

Start with the obvious

• Are the cells alive? Are they green and moving (if in liquid)? Did you get this culture through the mail in the middle of summer or winter? Dead cells usually become yellow, grey or colorless. Lack of movement may not mean dead cells, but probably means they are not happy.

• Will other cultures grow in the same medium or on the same plates? If not then you have probably made an error on making up the medium. Check the quantity of each of the stock solutions used. Could you have used a nitrogen or sulfur-free medium by mistake? If the medium contains a precipitate this suggests an error in making it up.

If you can eliminate these, then try the following:

• Is this a mutant strain with specific nutritional requirements? Is it a photosynthetic mutant needing acetate, an arginine-requiring strain? Some stocks cannot use nitrate as a N-source, does your medium offer another source, such as ammonium?

• Is the culture contaminated? Often a milky or yellowish tinge is a sign of trouble. Contaminated cultures often fail to swim and grow poorly. Not all bacterial contaminants are obvious, check under a compund microscope. If in doubt, follow the procedures given in the contamination FAQ

• Is the room temperature too high or low? Chlamy generally does best at 20-25ûC and even a short period of high temperature can have a significant effect

• Are they getting enough light? This is especially critical if you are growing the cells in the absence of acetate

My cells are contaminated

From Olivier Vallon

I have found an easy way of getting rid of fungal infections in cultures of Chlamydomonas reinhardtii. It takes advantage of the fact that C.r. can grow in the presence of methionine sulfoximine (MSX, a well known inhibitor of glutamine synthase), provided arginine is added to the medium, while most of our unwanted visitors cannot.

If a fungal infection appears on a plate and it seems difficult to pick a clean Chlamy inoculum, streak on a TAP plate supplemented with MSX (100 microM) and arginine (100 mg/l). If possible, place in bright light for 3-4 days, streak back on your usual medium (or a fresh TARG/MSX plate if you want to be extremely careful). This works fine even with heavily infected cultures. It seems to work also with bacterial infections.

I have no definite explanation why arginine allows resistance to MSX. In its absence, 50 microM MSX are enough to kill C.r. I suspect C.r. has a high ability to use arginine as exclusive ammonium source, bypassing NH3 and the need for glutamine synthase.

From Antonio Franco

To get rid of bacterial contaminants, I use antibiograms discs. I stripe a line of Chlamy cells in a TAP solid media and have several of these discs close of such a line. This way, I can use several antibiotics at the same time and in the same Petri dish. Tetracycline, Cefotaxime, Trimethoprim, erythromycin, ceftixozyme are fine. Chlamy cells isolated in this way, remain in pure cultures for a long time.

From Jason Frank

In liquid culture you can clean Chlamy by carefully centrifuging them down, washing the bottle walls carefully, repeating a few times, and letting them swim back to the top. I put aluminum foil around the bottle (so they don't swim down), and nab them with a pipette.

From Elizabeth Harris

We sometimes use ampicillin [50 micrograms/ml], especially in plates to be spread after biolistic transformation experiments where the risk of contamination is pretty high. However I find in general that brute force is a better cleanup technique. Streak cultures at low density on highly nutritive medium (e.g. containing yeast extract), where any contamination will be readily apparent. Using a fine glass needle under a dissecting scope, manipulate individual Chlamy cells away from the mass of contaminated culture. Inspect the plates daily under the microscope, and continue to move the algal cells away from the bacteria (or fungi). When you find a clean, uncontaminated colony, transfer it with a sterile toothpick to a fresh plate.

The Chlamydomonas Sourcebook has some additional suggestions for contamination control on pages 48-50.

Encouraging Chlamydomonas Strains to Mate

Mike Adams

I have found the following sequence to be very successful, even with paralyzed strains.

Grow the cells for one week on TAP plates under bright light

Scrape off into 10 mM HEPES, pH 7.4

Pass through a Dounce homogenizer, flattening all the clumps against the side

Bring to about 10^6 cells/ml

Place under bright light for 2-4 hours with constant bubbling or agitation

From Bob Hodson

I have pieced together procedures from various sources (all personal communications) --- Lib Harris (also source book), Pete Lefebvre, Bill Snell, and Susan Dutcher --- for mating Chlamy that are working routinely for us and accomodate cw- and pf strains as well as strains with various requirements such as arginine or nicotinamide. There is probably much room for modification, e.g. medium, light intensity, temperature, etc.

1. Have on hand at all times lawns on TAP (tris-acetate-phosphate with 8 mM ammonium) supplemented with 0.4% yeast extract, 2 mM arginine, and 10 mM MOPS (1.5% agar unless otherwise stated) = "rich plates". pH is 6.8 and the result of using acetic acid (1 ml per liter) instead of acetate, and additional sodium hydroxide (about 1.4 ml of 4M per liter). MOPS helps to control pH. Tris has a pK of about 8, and with it as the only buffering agent the medium tends to rise to that value and be toxic. MOPS has a pK around 7 (but is twice as expensive). Arginine is added to make sure that arg- strains have enough, and it can be omitted. Yeast extract provides supplements for some of the "standard" mapping strains (e.g. CC2645) and also helps reveal contamination in cultures (but also encourages it). The lawns are grown at room temperature (20C) under dim light and are ready in 5 days; other temp and lighting conditions will probably work equally well, but growth time will vary accordingly. These stocks are replaced every two weeks, but may be useful longer than that, especially if MOPS is used.

2. Scrape off about 1/3 of the lawn and suspend in 1.0 ml N-TAP (no MOPS or N source) in 24-well culture dish. Break up clumps with repeated pipeting. Use 0.5 ml to make lawn of cells on N-TAP containing 1 mM ammonium = "lowN plates". Incubate 2 days @ 25C under moderate light (2000 f-c fluorescent). Even arg- and nic- strains will survive and mate well. Plate remaining 0.5 ml on rich plates to refresh stocks.

3. Scrape off about 1/3 of lowN lawn and suspend in 4 ml half-strength N-TAP in 25-ml flask. Goal is about 2 million cells per ml. Shake in water bath at 25C under bright light (2, 150W flood lamps about 20 cm away) for 4 hours (2 hours may be enough; mating activity declines overnight but not to zero).

4. Mix 0.5 ml of each mating type in 1.5-ml sterile Eppendorf tube. Incubate stationary at 25C under bright light for appropriate time (see below). Rest tubes in the necks of 25-ml conical flasks filled with water and bathed with water to control temp. The appropriate time depends on strains. Very active walled strains such as CC124- and CC125+ can be plated in an hour or two, and longer incubation gives clumps that are hard to break up. cw- strains probably can also be plated fairly soon if the other strain mates well. For pf strains let mating go for an hour, spin cells just enough to pellet (2000 rpm, 1 min), remove most of the medium, vortex cells gently up in the residual medium, and let mating proceed another 3 hours to overnight as needed. Plate 0.5 ml portions.

From Bill Snell

In response to the inquiries about using IBMX for activating gametes of C. reinhardtii, folks should be aware that not all strains will activate with dibutyryl cAMP and IBMX. For example, we are unable to activate strain 21gr using this combination, and instead the cells resorb their flagella and otherwise look funny. On the other hand gametes activate perfectly well if we use 10 mM dibutyryl cAMP (50 mM stock is made in N-free medium and stored frozen) and 0.15mM papaverine. We use 100% DMSO as the solvent for the 15 mM papaverine stock solution. And we warm the DMSO to 60 degrees to get the papaverine in solution. Also, we always make up fresh papaverine, and we find it is important to use DMSO that is fresh. Thus we use DMSO sold in small vials by Sigma. We find that cell concentration is not very important but if money is an object, and one wants to use less dibutyryl cAMP, cells activate just as well at 2 x 107 cells per ml as they do at 2 x 109 cells/ml.

From Elizabeth Harris

In response to a query about getting cell wall deficient strains to mate:

Save a little of your mating mixture, 0.5-1 ml or so, in a tube and leave it in bright light overnight, then check to see if there's a zygospore pellicle formed. This will look like a somewhat reticulate network around the walls of the tube, and on the surface of the liquid - try it with a wild type x wild type cross done in parallel, so you have something to compare it to. If you *don't* see a pellicle in your cw mating, then you have an advance indication that you may not have gotten good mating efficiency, and you can go ahead and repeat the cross. This saves waiting for maturation and visible zygotes on the plates.

In my experience, there are two types of problems with cw crosses. First, some of the cw strains have a high proportion of "bald" cells with no flagella. This is particularly true of the original Davies cw15 isolates, in the Chlamydomonas Genetics Center collection as CC-277 and CC-278. However, CC-400 and CC-406 are among the better ones in this respect, as they were selected originally for swimming and mating ability.

Second, we've had problems with crosses involving cw15 and arg7 in combination. In this case, mating may occur at reasonable efficiency, and zygospores are formed, but germination and product survival are very poor. I don't have a good workaround for this, and would be interested in hearing the experience of others.

From Carol Hwang,

I would add that if the cells on the plate are 6-7 days old, they should have depleted the nitrogen and be gametic so all you need is for them to be flagellated to mate them. I try to suspend cells by smearing them on the side of the tube then bringing medium up by the loopful to dissolve the smear (Paul Levine compared this to making a flour paste for gravy) and I don't vortex the cells because this can shear off flagella. I also look at my plates under a dissecting scope before wrapping them because frequently I can see zygotes by their relatively enourmous cell diameter.