From Kosuke Shimogawara
By my experience, yield of the lytic activity is completely dependent with the efficiency of mating. To get good mating, I am using good mating strains, i.e., CC-620 and CC-621, both of them are available from the Chlamydomonas Genetic Center. You might also be very careful for the concentration of the cells during mating. If the concentration during mating is too high, the efficiency of mating seems to become worse (probably anaerobic environment is harmful).
The following is my protocol.
– Cultivate both mating type of the cells up to about 3×10^6 /mL in 250 mL of TAP in 1 L flask.
– Collect the cells by centrifugation at 3000 rpm 5 min.
– Resuspend the cells in 1L of TAP-N (NH4Cl was replaced with the same concentration of KCl) in 2 L flask.
– Induce gamete by gently shaking for 24 hr under light.
– Harvest the cells by centrifugation at 3000 rpm 5 min.
– Resuspend each cells in 200 mL of TAP-N.
– Mix both mating type of cells in 2 L flask (larger vessel seems better).
– Keep it under light without shaking for 1 to 2 hr to allow mating (vigorous shaking seems harmful).
– Remove the cells by centrifugation at 8000 rpm.
– Remove cell debris by centrifugation at max speed.
– Filtrate the medium by 0.45 um membrane filter unit.
– Keep in -80C until use.
From Karen Kindle
I think I have the opposite problem with autolysin – too much activity which can be a problem. We need to dilute our autolysin for many applications. I use high efficiency mating cells (CC620 and CC621) and grow them phototrophically, as described in the Sourcebook. They are then starved for nitrogen overnight and then the two mating types are mixed together for 1-2 hours. I pretty much follow the instructions, except I simply do a high speed (20K) spin, then filter through 0.45 micron filter, then freeze. Very simple except for manipulating the large amounts of culture supernatant.
From Bill Snell
Making g-lysin preps is not always easy. We have published a method in Experimental Cell Research (Buchanan and Snell, “Biochemical studies on lysin, a cell wall degrading enzyme released during fertilization in Chlamydomonas; ECR 179:181-193) that works almost all of the time. We understand why most details of the method are important and we can only guess about others. For example you need to have highly competent gametes (greater than 70-90% of them can fuse), we’re not sure why using cells at 108 is so important, but it is. Also we’re not sure about why the 0.5 hr mixing time is important, but it may have to do with the fact that g-lysin may bind to cell walls. If the cells are mixed long enough in the presence of lysin, the walls become degraded and are not sedimented when you clarify the lysin. Anyway, most times we can change some of these parameters and get good lysin. But, if our lysin preps start to be less efficient, we go back to being fastidious and it works.
From Elizabeth Harris
Heriberto Cerutti, a post-doc in the Boynton-Gillham lab, tells me that eliminating the final high-speed centrifugation prior to freezing the autolysin prep seems to result in higher activity. He thinks perhaps the enzyme is partly bound to cell debris. Since freezing at -80C would presumably kill any surviving Chlamydomonas cells in any case (as everyone who’s tried to preserve Chlamy by freezing can attest!), this step would seem to be non-essential. Has anyone else experimented with this?
From Gernot Gloeckner
We only remove the cells from the mating medium by centrifuging two times at 5000 rpm. Then we filtrate the supernatant through a 0.45 micrometer diameter filter. This preparation is stored at -20 ¯C. The activity of this Autolysin preparation seems to depend only on the mating eficiency. So I think, you need not to make a high speed centrifugation.
From Lisa Ellis
None of us in the Goodenough lab do the final high speed spin on our autolysin preps. We just spin out the cells and then freeze it. Any remaining Chlamy cells definitely DO die when frozen.