Has anyone done free-floating spinal cord sectioning/staining using fixed, but not frozen tissue using a vibratome?
For example, I noticed Andrew Todd does much of his IHC this way, and the images look great. Any thoughts? Is it much better than frozen fixed?
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I’ve used Andrew Todd’s vibratome method for VGLUT2 and VGAT staining in the spinal cord, following https://www.ncbi.nlm.nih.gov/pubmed/20817353. The methods look pretty similar to the paper you linked except for the 0.5um semi-thin sections which I’ve never done. Anyway, I didn’t really think the images were that much better than with fixed frozen sections. A lot of people say that the morphology is better preserved with the fixed and unfrozen approach, so perhaps if you were doing a study focusing on fine details of morphology then it might make a bigger difference. However, for my purposes I find that the fixed-frozen method with free-floating immuno can produce some really nice looking images.
Thanks a lot @liz. That’s exactly what I was looking for. If it’s not that much better, it’s likely not worth the effort. Would you be willing to share your free floating IHC protocol here? We do a standard BSA one, but sometimes the background is higher than I’d like.
Also, do you have any tips for the washes? I’ve used some netwells in 24 well format, but they seem to damage the tissue. If I do it just in 24 wells alone, without nets, it takes longer and doesn’t fully remove buffers/antibodies. @zhzhj131421 removes slices one by one using a little hook. takes forever but he gets the results. if you have any insights, that’d be great.
Thanks!
Oh, my protocol is pretty standard as well. Using 40um frozen sections, I wash 3 x 20 min. with PBS, then permeablize for 1 hour with 0.3% Tx-100 in PBS (This step might not actually be necessary since the blocking and Ab buffers have 0.3% Tx-100 as well, but I guess I’m a little superstitious about deviating from a working protocol). I block using 5-10% normal animal serum of the species in which my secondary Ab was produced, diluted in PBS-Tx (block for 2-4 hours depending on what else is going on that day). I have had background issues as well when using BSA, although I know some people use it routinely with good results. For antibody buffers, I use 3% animal serum in PBS-Tx- there is usually some leftover blocking buffer that I can just dilute down. I do incubation overnight at 4C then wash 3x the next day, and then my secondary for 1-1.5hr at room temp. Wash again 3x then I mount with a paintbrush. Of course incubation times and conditions may vary with Ab. But this general protocol seems to work for most things.
For section handling, I use the netwells (12 well format) and just accept that I’m going to lose some sections to damage. I don’t have too many problems with tissue damage in P21 or older mouse spinal cord sections, but I tried some P7 tissue recently and they had much more damage from the netwells. So if your tissue is fragile then maybe use the hook or paintbrush method to transfer. With a paintbrush you can probably pick up a bunch at once so it saves some time. Alternatively, Corning seems to have a wide variety of different well inserts made of different materials and meshes so maybe it’s just a matter of finding the right product for fragile tissue. This is going to sound weird, but I know our lab uses pantyhose (like pieces of the material, not the whole garment) to hold and support live spinal cord slices while they’re incubating prior to ephys recording, and I’ve often wondered if I could build some small holders our of this material for IHC use.
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Thanks for sharing @liz. That’s very similar to my protocol. I may try the other kinds of buffers (non-BSA).
Re: new kinds of wells. yes, I’ve thought of something similar. In addition to keeping the tissues intact, I also want to minimize reagent use. The netwells use a lot more antibody than I would if directly staining in wells. It’s a trade off. I can envision like a thin wire frame with pantyhose material around it. That could work. I’ll try some time. Maybe this summer.
Hi Alex, I think the sections from fiexed, but not frozen tissues using vibratome are often used for the electron microscopy. It is convenient to do immunofluorescence using frozen thin sections.
Hi @liz and @achamess, stumbled upon this pretty old thread. Hope you are all still in business in academia. I am looking into using Netwells for managing mouse brain sections. Is it the consensus that they may damage the tissue? If yes, is it due to abrasion from the net material or from clinging to the well wall or something third?
Best regards
Dang
Hi Dang,
I’m almost exclusively in the spinal cord, but I have used the netwells for mouse brain sections as thin as 20um without noticeable damage. Additionally, my colleagues use them for ~40um brain sections regularly, and they haven’t had any issues with damage. I think their opinion is that the netwells produce less damage than repeatedly manipulating each section with a brush. Hope that helps!
Edit: I should add that this is in the context of adult tissues. Neonatal CNS tissue tends to be a lot more fragile, so damage from netwells could be increased in that case.
Hi @liz
Thank you so much for your very fast and helpful reply. What would you recommend for washing e.g. neonatal CNS tissue?
If it’s important to do free-floating sections, I’d probably just try the netwells and see how it works out for you. It could be totally fine, but I’ve never done anything with neonatal brains before so I can’t speak from experience. You could also consider mounting the sections directly on slides rather than free-floating.
I’ll try out that.
Thanks a lot