After fixing brain, spinal cord, nerve, or DRG, how do you store the whole, unsectioned tissues?
Right now, I keep fixed (4% PFA) tissues in 1x PBS with 0.1% sodium azide at 4ºC. I keep these for weeks to months.
I also do this for fixed, free-floating sections (40-50 um).
I haven’t formally evaluated whether the duration of storage in these conditions affects performance in immunohistochemistry.
I know some people will use a cryoprotectant (e.g. glycerol-dmso) for storage at -20C or -80C.
This review goes over many options for human brain biobanking
What do you all do? What is the basis for your choice? Do you note a difference between storage at 4C in PBS vs freezer-temp storage?
My fixed (perfused or drop fixed) samples go straight into cryoprotectant (30% sucrose) until they sink, and then are frozen in OCT over dry ice and stored at -80. Although I am sure storage at -20 would be fine.
When I was at NeuroMab, we always stored our fixed sections (for free floating IHC) in TBS + 0.1% Sodium Azide at 4C. I mostly did immunoperoxidase IHC back then and the azide replaces the hydrogen peroxide pre-treatment step because it also blocks endogenous peroxidases. I don’t know how long the sections keep like that because I went through them so fast, but I expect they keep for a long time.
Thanks @sshiers for continuing to share your IHC wisdom.
So before sectioning, your approach is:
Fix (immersion or perfuse)
30% sucrose cryoprotectant
Block in OCT and then store as a block in OCT at -80C.
That’s sensible and efficient, and then the tissue is ready to go for sectioning and is preserved as well as can be at -80C.
For after sectioning, it sounds like you stored just in buffer with azide. That is also what we do, but I’m wondering about the long-term preservation of that method. There is some suggestion that the antigenicity declines over time. See this paper:
It got me thinking about storing sections in something besides buffer and keeping at a lower temperature.
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 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.
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.
You might find this paper interesting about fresh vs fixed
They evaluated acetone vs. PFA fixed fresh sections vs. FFPE.
This is consistent with my experience. I think PFA fixation is best all around. Acetone has not been good in my hands.
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.
Oh, the block is super easy to pop off the chuck. If you just rest the chuck (with block attached) on your palm (in the cryostat) - with the bottom of the chuck touching your palm that is. Give it like 10-20 seconds and then pop the block off. It just needs the slightest bit of warmth on the bottom of the metal to easily remove. I cover the used block with prechilled tin foil, place it back in the labeled bag over dry ice and return to the -80. I am going to try those little storage containers you linked! I will order today!
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.
I only use 10% formalin. We use methanol for Visium and that seems fine for general H&E stain, but I have never tried it for IHC. Morphology looks the same to me between fresh frozen and perfused.
I had to run a really crazy optimization protocol recently to make sure a set of our antibodies was compatible with a company’s fixation/antigen retrieval protocol. It was absolutely insane and really brought into question my knowledge on how all this stuff works because I thought there is no way me hair drying (yes I brought in my hair drier) an unfixed section of human DRG for 5 minutes, and then baking it at 37 degrees for 1 hour (still unfixed) + fixing + antigen retrieval #1 + antigen retrieval #2, + fixing again, then IHC would ever work. But honestly it gave me one of the most incredibly beautiful IHCs with virtually no background that I’ve done with those abs. I’m currently trying to tease that protocol apart to figure out what the key component was because I really want to implement it for finicky antibodies. Completely blew my mind tbh
