I have stored free floating sections at -80 in 30% sucrose - one of our collaborators here suggested it. The few antibodies I used on them still looked great. However, the tissue did get tangled (probably because I put way too many in a tube together) so I had section loss. Is there a reason you are cutting so many sections at once? I usually just cut what I need for the experiment and put the block back in the -80.
That’s good to know. I try to slice all at once because I was under the impression that, for fresh tissue at least, going back and forth with temperature changes (-20c to 80C) would degrade the morphology and maybe affect things like RNA. Also it’s just easier to get it all done and move on to other tissues. For fixed tissue, we could keep the block in -80C after sectioning, yes. But if we’re sectioning, I figure, might as well make all the things and be done. Hence my desire to find a storage solution for sections. I’ll try the sucrose.
How do you store partially used blocks? Do you take it off the chuck or just put it in the freezer while on the chuck? If the latter, that’s one of my reasons for avoiding keeping remaining tissue in the freezer - it uses up a chuck.
Yes, those steps are correct. I honestly prefer using fresh frozen tissue for everything now though and I rarely use perfused/pre-fixed tissues. Antibodies can be so finicky when it comes to fixation state that I would rather just optimize the fixation times for sections of one fresh frozen specimen, over perfusing a bunch of different animals with different volumes/flow rate/etc of fix to figure out which one works best.
I see your point. I’ve thought about the same. Do you not find the morphology of fixed tissue better than fresh (and then fixed on slide)? The PFA-fixed tissue just looks nicer in my hands. Maybe we need to get better with the freezing.
What other fixatives do you use? I’ve toyed with acetone recently. I’m not impressed. Probably factor that matters most is duration of PFA fixation. I never do antigen retrieval.