300 micron thick brain slice mounting?

Hi Santiago,

I'm not sure what you're using as a mounting medium, but perhaps the most immediate solution is to use a more viscous fluid. Although the coverslip presses down on the tissue and compresses it, we've had no problems mounting thick slices in either vectashield or 90% glycerol in PBS (add dabco or npropylgallate or ascorbic acid as an antifade), coverslipping and then sealing with nail polish or dpx.

For some studies, the compression by the coverslip was unacceptable, and I very recently overcame this problem two ways:

you can either buy silicone gasket spacers to support (and in some cases even secure) the coverslip;

or, you can achieve the same effect by using broken coverslips (2 layers of #1 coverslip glass is almost exactly 300um).

I recently spent an off afternoon making a bunch of slides into "chambers" for this purpose. A diamond glass cutting pen was enormously useful for neatness, but you really could just do it by carefully breaking the coverslips with forceps.

Alberto Sepulveda Rodriguez

I also just remembered that we've successfully used dental wax or blu-tac to build a chamber when we had to mount specimens in BABB or methyl salicylate.

Those could also work for your purposes here.

Victoria Wu

I've used parafilm as a spacer. Since its waxy nothing should get past it and it works well as a gasket of sorts. You can also make thick layer of this by layering the parafilm. But the trick is to put your sample right in the middle of a solid piece of parafilm that has a hole cut out for your sample. Press the parafilm down around the sample. If 1 or 2 layers of parafilm is not thick enough, you can use rubber cement. It is not toxic, dries fast and has no background fluoresce. you can pair this with the parafilm by surrounding your sample with parafilm and sample hole in middle. then layer the rubber cement on top and once it dries, put your sample and mounting media in the middle. Add coverslip. Should stick. Good luck.

Santiago Elías Charif

Thanks to all of the researchers that have been answering to my inquiry. I'm taking notes of every possible alternative that you recommend!

Jenelle Wallace

I have the exact same problem - thanks to everyone on this thread for your suggestions!

Jesus Bautista-Garrido

Thank you so much for everyone's input. I will definitely try the wax method. I wonder if I could use orthodontic/braces wax? it is super malleable and I would assume easy to use for this purpose. I will pass along what I fund out.

Thanks again!

Santiago Elías Charif

We have published in 2020 (Reference at the end) a work that employs the following method. I am very thankful for your suggestions. I hope is understandable. You can reach me for more details.

Cut a square of Parafilm and fold it by the half (you'll get a rectangle). Cut it in the middle, keep one of the squares. The idea is that it has to be slightly larger than a squared coverslip (not more than 1-2 milimiters).

Keep it pressed with something heavy (like a book) against a table for a couple of minutes.

Try to draw, with a thin point marker, the shape of the slice on the parafilm square (do not place the slice). It does not have to be tight, you'll need to build a little pool after.

Use a blade from a stanley knife or utility to cut the marked shape, bit by bit, without dragging or tearing the parafilm.

Stick the parafilm square onto a gelatin covered glass slide. Use fast dry adhesive (in Argentina is named "La Gotita", in USA is named "Super Glue"). For this, place a droplet in each corner, and then spread it, trying not to react the border of the empty, cutted space (otherwise, when you place the parafilm it will leak and the tissue will not adhere).

Fill the "pool" (empty spaced shaped like the slice) with the mounting medium of choice (we use glycerol in PBS). For coronal sections of motor cortex slices from paraformaldehyde fixed brain of adult mice), empirically test the right volume. Try 80 ul. It does not have to leak or cover the parafilm. Just enough to form a drop that reaches the edges of the sectione parafilm shape. NOTE: it is important that the 400 um thick slice is flat; for this, you need to fixate it on a 24 well plate. We tried to fixate it in an 1.5ml eppendorf tube but the slice resulted in a bent shape and the microscope visualization gave us out of focus sections.

Place the slice with a fine hair brush, trying not to overflow the mounting medium.

Place a square coverslip carefully, again, trying not to displace the mounting medium and trying to keep the slice in the center of the "pool".

Press it and apply a generous amount of transparent nail polish in the border of the coverslip-parafilm. You can place a small weight on the center of the coverslip while drying, don't forget this.

Let dry in darkness, and it can be stored horizontally at 4°C. Do not change its orientation to avoid slice displacement or bubble formation.

Hope it works!

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