For fluorescent immunohistochemistry I fixed spinal cords in PFA, incubated in 30% sucrose, froze in OCT, and sectioned at 30um. All my sections are now on untreated charged glass slides that were immediately frozen at -20. I noticed they are easily falling off in wash buffer. Is there any way to help them stick after they have been mounted (emphasis on after!). I know you can use gelatin beforehand. Does heat or air drying help this late in the game? My cells of interest contain fluorescent signal prior to antibody labeling (alexa dyes and tdTomato), so for this reason I'm reluctant to leave the slides out at room temperature for too long.
Drying tissue sections on glass were useful for me. But I used unfixed brain tissue.
Also, try differently treated glass slides, maybe?
Hi Steve,
I know you emphasized on 'after the mounting' in your question but I was curious if you tried staining your slices first and then mounting them after the secondary? I am doing a lot of immunofluorescent experiments on spinal cord slices from mice and I get very consistent results in terms of background and better ab staining when I do the procedure on free-floating slices.
Thanks for your suggestion Prudhvi. Funnily enough, I was just considering this method. I see it has been catching on in the IHC world. Do you do something similar to this:
https://app.jove.com/v/61622/histological-based-stainings-using-free-floating-tissue-sections
The only downside (aside from having to mount tiny fragile slices) seems to be that is hard to keep track of the anatomy. Mounting straight to the slides has the advantage of being easier to track spinal segments and preserving the left and right orientation of the horns. Seems like you would never know if your free-floating sections flip. While I make transverse slices rostral to caudal I can put the slides right under the microscope to see which spinal segment I'm in. Do you have any method for segregating 30-100 free floating sections?
Hi Steve,
I don't know if the procedure in the link will work for SC slices given that they are too delicate. I agree with you that it is easier to find the anatomy of the SC if you mount the slices straight on to the slides. These are the things I do for IF staining in SC slices.
1. If you want to probe your tissue for specific proteins, the best thing to do is to collect the slices in series and put them in a multi-well plate. I also do transverse sections rostral to caudal but usually focus on segments L4-L6.
2. In order to distinguish the right from the left I make a cut along the ventral horn with a blade (since I do not need the ventral horn for my studies) after the tissue is incubated in 30% sucrose so it is firm enough following the dehydration.
3. I stain the sections in the multi-well plate without using the perforated chambers as shown in the video link.
4. Mounting the slices after the secondary ab and the PBS washes is not very difficult either.
30 slices can be easily done with the procedure I mentioned above but for 100 slices, it must be tedious.
I hope it helps. Let me know if you have any questions.
All the best.
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