Fresh frozen spinal cord cracked and shattered in the center

Hi all,
Working with some human spinal cord. I took this fresh and embedded in OCT, and then froze in isopentane cooled by liquid nitrogen. We then stored in an airtight container at -80C before sectioning.

These are the sections we got

What’s the diagnosis?

  • Freezing was too slow
    • The white matter looks OK. This is a fairly large piece of tissue. Did the interior not freeze quickly enough and ice crystals formed and damaged the tissue?
  • Too cold at cryostat
    • We sectioned at -20C and then even brought it down to -13C. So I don’t think it was too cold, but I know tearing can happen with too cold blocks.
    • 10x discusses this for the Visium protocol:
  • Something else I’m not thinking about?

I’m inclined to think maybe not fast enough freezing. In which case, directly freezing the tissue in isopentane (not in OCT), might be the way to go.

@sshiers @liz

I feel pretty confident that your issue is the cryostat temperature, not your freezing technique. Tissue + OCT in mold over isopentane and liquid nitrogen is standard. We use powdered dry ice which isn’t standard and I did have issues with the grey matter shattering like that when I first started sectioning/histology on hSpinal cord… and it was completely resolved by optimizing the cryostat/sectioning conditions. For me, the chuck around 14 degrees, and chamber around -18 to -20 usually works great for me. I usually swap out the blade for a warmer one when I see the grey matter shattering like that, and you can even put your thumb against the sample to warm it slightly then try sectioning again. Faster speed works best - I think its just the abrupt difference in tissue density between white and grey matter.

Also - how long did you let the block sit in the cryostat before you started sectioning?

I actually am cutting spinal cord right now (weird timing :stuck_out_tongue: ). I just took a video (but it wont let me attach it). Cryostat chamber is -19, chuck holder is at -18 and my sections looked good. But I’ve been cutting this sample for a while now so its plenty acclimated to the chamber.

Very helpful comments @sshiers! Thank you. We’ll more carefully monitor temperature and try some of these tactics.

We’ll report back.

Do you prefer embedding fresh tissue directly in OCT vs. snap-freezing directly in isopentane then embedding in OCT after?

Perfect timing, while it’s fresh in your mind. We let it sit for at least 30 mins, and by the time those sections were made, we were at 1.5 hours.

You can attach a video using a Dropbox or Google Drive link if you want to do that.

I’ve never used the isopentane/liquid nitrogen technique so I cannot speak to that technique.
But I do prefer direct freezing of the sample, and then embedding in OCT later. The issue is that if the freezing is too slow, you get the ice crystal formation and I worry that the OCT slows down the freezing time.

We use powdered dry ice (we smash it up with a hammer) and then cover the tissues directly in the dry ice while in the OR. Then we transfer them to the pre-chilled tubes. They are kept in the -80. When we need samples for histology, we remove them from the -80 and place them on dry ice, and then embed them in OCT over dry ice using a metal mold (helps the oct freeze faster). We make thin, ring like layers around the specimen, let that freeze, and then add another layer and so on - that way we don’t risk thawing the sample. The metal mold helps a lot with that. It works great for us.

I am curious how you store your blocks of tissue - what air tight containers are you using? We place our OCT-embedded blocks of tissue in tiny ziplocks with index cards denoting the information for the sample… however, I have been looking for alternatives since I worry about air moisture in the bag as they move from the -20 cryostat, and -80.

Thanks for the details!
Glad to hear dry ice works for you.
I’ve done the isopentane mostly because it is recommended for fastest freezing and maintenance of morphology. Also, it’s the method recommended by Visium (10x). If it’s good enough for that, I figure it good enough for anything. 10x has great videos on the freezing and embedding process

https://pages.10xgenomics.com/sup-how-to-visium-tissue-prep.html?wchannelid=92q73hscm6&wmediaid=20jm5uauab

Do you have catalog numbers on those metal molds?

For storing blocks, this is what I’m using

The idea came from here:

For slide storage, I’ve even gone a little over the top:
I put slides in single-shot slide boxes with silica desiccant packets and vacuum sealed

In this way, I have no concerns about long-term stability and quality of the tissue. This is for fresh tissue. For PFA fixed with sucrose, I don’t do the vacuum, but I do the desiccant.

-Awesome I will try those containers and I really like the desiccant idea as well.

-I actually dont know where we ordered the metal molds from - I was cleaning out a drawer and found them years ago. Lol. But this looks similar to the set we have:

Note - they are reusable!

-I avoid storing pre-cut fresh frozen sections on slides at all costs. Probably for the exact reason you have to go to the extreme of the desiccant+vacuum bag… they are so sensitive to humidity changes and with the -80 door being opened and closed so often, the tissue is just not going to last. Instead, I only cut what I need for the immediate future (within the week), and use the slides immediately. I fix the sections on slides in cold 10% formalin for 15 minutes, dehydrate in 50-100% EtOH and then store them in 100% EtOH in the -20 and then use them over the week. I do this for RNAscope. For IHC, they get fixed, dehydrated, pap penned, and then put in block (10% NGS, 0.3% TX100 in 0.1M PB) until I need them (within a couple days)

I’ve also fixed, dehydrated and let them air dry in a drawer for days and that works too (for both RNAscope and IHC) - ACD tech support recommended that one to me.

https://utdallas.box.com/s/mpd59d6veyxaaw7kkr0kgkdpaw34fg3z

We had the cracking issue sectioning fixed tissue when using isopentane cooled to excessively low temperatures. My partner uses the isopentane technique for freezing rodent brains - she says it is critical to keep the isopentane around -55C. We put isopentane in a metal beaker, surrounded by dry ice, and use a low-temp thermometer to ensure the temperature is ~correct.

That’s an interesting insight. I thought the idea was to freeze as rapidly as possible, so as to avoid ice crystals, which is why people suggest using LN2 to cool the isopentane. I will try the method you recommend here.

And, as @sshiers has said, the fresh freezing offers a lot of flexibility downstream.

Yes, I often freeze my fixed rodent tissue on a weigh boat in liquid N2 - but this -55C strategy should be better for fresh or larger tissue. We’ve had cracking issues with fresh tissue or brains frozen at temperatures that were too cold. Good luck!