Mounting many DRG for cryosectioning - share your tips

How does everyone mount their DRG for cryosectioning?

I was taught by @tberta to put a piece of filter paper down on a chuck, and then lay out a grid array of DRG on the filter paper and then put OCT over it. That’s worked pretty well over the years. Sometimes it’s hard to get all the DRG in one section because of their different geometries, but you can really pack a lot in one section (16-20 for me).

What does everyone else do? Are there other methods people like? Also, is there a case to make for doing one at a time?

@runDRG @tonellor @esypek @ShanTan @sshiers @thicunha

I don’t do anything fancy - but I like your idea of using the filter paper since getting all DRGs in the same plane for cryosectioning can be difficult. I usually make a small flat surface of OCT in the mold, freeze it, lay several DRGs on that and cover immediately in OCT.

When cryosectioning, I take a section periodically and look at it under a Brightfield microscope to make sure I have at least one decent looking section in the field of view. A lot of times they are not oriented perfectly, or you get a lot of nerve root sections so checking this way helps a lot! :slight_smile:

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I like this idea! I usually mounted fewer per block just to make it easier to keep track which DRGs are which, since I was interested in comparing between levels.

I think using filter paper is not a good idea. I did that a long time ago and ended up with tiny pieces of paper tangled in my DRG sections. Fragments of the dorsal root that are contiguous with the DRG are magnets for all kind of dirt and will make mounting the DRG sections on the slide difficult and nerve-wracking.

This is what I do. Put some OCT in the chuck and freeze it. Your chuck should have a mark on it so you always mount it in the same orientation. Cut the OCT with the cryostat to get a flat surface. Then lay your DRG on that surface as you want them. Cover with OCT and freeze. Mount the chuck with the same orientation. Now you can cut your DRG and you will get several sections on the same cut.

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@marvizon I like that! Indeed, I have the issue with filter paper getting sliced onto the slice.
When you’re laying the DRG on the frozen OCT, you’re doing this all in the cryostat or on dry ice? Also, do you chill the DRGs (in sucrose solution) so that they don’t melt the OCT? If it’s -80C that’s not a big deal. I’m afraid of the DRG freezing onto the OCT too fast and then I won’t be able to position them properly. I make grids of 4x4 or 5x4 DRG so I can basically do an entire experiment on one slide.

I use free-floating sections, so there is no need to organize the DRG on the chuck in any way.

Here is a method we often use and may be a help for you. We place an aluminum block on dry ice. The reason to use the metal block is because it has a higher specific heat than the dry ice, so it would absorb the heat from the sample very quickl. On top of the aluminum block we place a glass coverslip. Here you could draw a grid on the coverslip to organize your DRG. Then we place de DRG (from a 30% sucrose cryoprotcting solution) on the coverslip. Put a drop of OCT on the coverslip and it will freeze very quickly around the DRG thanks to the aluminum block. The last step is to gently push the drop of OCT with the embedded DRG off the coverslip (with forceps). That drop has a flat surface that can be “glued” onto the flat surface of OCT on top of the chuck using a small amount of OCT (when it freezes it will attach the DRG to the chuck).

I developed this method to be able to section electrophysiology spinal cord slices (400 um thick) into 25 um-thick cryostat sections, but it works for many other small samples that need to be positioned very precisely for cutting in the cryostat.

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Wow! That’s ingenious. Thanks for sharing. I’ve got plenty of things to try now. I’m glad I asked about this question. Thank you!

You are welcome!

Don’t forget to cut the OCT on the chuck with the cryostat as I described in the first response to create a flat surface parallel to the plane of cutting of the blade of the cryostat.

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Wanted to check back in. I tried a number of methods, but actually what worked best was putting the DRGs in the bottom of a cryo-mold. Previously i had avoided this because I was worried about the DRGs moving around when I poured OCT on top. But Judy Golden in Gereau lab told me the trick - you need to dry the DRG on a Kimwipe or filter paper and dab off all the excess sucrose so that it adheres to the base of the cryo-mold. I did this, aligned the DRG into a 3 x 3 array and then put the OCT on top. It worked perfectly. And because the DRG are directly on the bottom face, you can cut the entire DRG through the full thickness because the tissue is sufficiently elevated off the chuck by the molded OCT.

I can post video/photos next time I do this. I feel a little silly for not trying this sooner but for years I’ve just thought it wouldn’t work. Little tricks make the difference.

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I also put the DRGs at the bottom of the cryo-mold. I pour a little OCT in; place and orient the DRGs; then fill with OCT.

To ensure I know which DRG is which, I alternate between horizontal and vertical orientation of DRGs (90-degree rotation, looking down; these are mainly L4-L5 mouse DRGs, so elongated). I record which are which and write it on the cryo-mold. The orientation starts with horizontal and ends with a vertical DRG, so even if things get flipped my records ensure I can identify the DRGs.

I always include DRGs from different mice/groups in an individual block, so they are all treated the same throughout the process and to enable blinding during microscopy/analysis.

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Thanks @adgaudet. That’s also a great trick about the orientation so you know which is which. Will spare the mental spatial gymnastics I when making my slices to try to get them all the same orientation on the slide :slight_smile:

I agree 100% with putting all the DRG that you will compare on the same section.