Here are some of the conditions that I have gathered so far:
1. Fix cells with ice-cooled methanol in -20 freezer for 20 mins.
2. Fix cells with -20C methanol on bench (or in freezer) for 5 mins.
3. Some protocol also suggested adding 5mM EGTA to methanol.
Anyone know which one works best?
I always fix with -20°C methanol in the freezer for 4-5 min depends on the cell line! For our purposes it works pretty good!
Fixing with -20 methanol for 15 minutes in the freezer seems to work fine for me.Then wash 3 times with PBS. Also I include a 0.1% triton /PBS wash for 3 minutes, despite the fact that methanol alone is supposed to be sufficient to permeabilize the cell membrane.
I agree with the previous advices (MeOh at -20° for 10 minutes with HeLa cells). But some antibodies doesn't work well after metanol fixation (some epitopes are not preserved enough). That's why sometimes I perform 2% PFA at 37°C by adding directly few amount of concentrated PFA (16%) in the culture medium. Microtubules are preserved enough and the majority of antibodies work well.
Thomas
I usually use Methanol at -20C for 5 min, and it gives a very good results. As mentioned by others, be careful it you plan to stain with other antibodies, epitopes might be altered by methanol fixation.
Alternatively, I also had fairly good results with formaldehyde fixation and Triton permeabilization (3.7% formaldehyde 1% Triton X-100 in PBS for 5min at 37C). This could be a solution if the methanol fixation does not preserve other proteins you might be interested in.
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