We're trying to image TIM4 using a primary conjugated antibody that is affected by fixation, so we try to keep fixation to a minimum. The samples are fixed for 2 hours before cryo-protecting in 30% sucrose and final freezing in OCT in a Ethanol-Dry Ice slurry. This initial process is the same for the "wholemount" and 10 um cryosections.
Wholemount samples were thawed and sectioned by vibratome at 500 um and stained for 5 days at 4 C with gentle shaking.
Cryosections were sectioned at 10 um and not fixed again before proceeding with the staining procedure. Also, the slides were air dried for less than 30 minutes at room temperature. The antibodies were incubated O/N at 4 C.
We see staining in the "wholemount" samples, but not in the cryosections. Of note is that we use another marker that often colocalizes with TIM4 as observed by flow cytometry which works perfectly, so I don't think it's due to not having the right sections so to speak.
Could the air drying of cryosections have an influence on antigen masking? Should we increase the incubation time of the cryosections perhaps?
if the epitope is PFA-sensitive, 5 min in 4% PFA on ice may be sufficient (some use even 2% or even 1% PFA). 2h may be too long. also, wholemounts in your case are 50x thicker so the signal you observe comes from 50 stacks (similar to compression of 50 optical sections into one bright image), while 10u is .. 10u, the signal maybe not strong enough
Hi Igor O. Nasonkin, thank you for your suggestion. I forgot to mention the 2 hour fixation is done at 4 C. When you mean fixation for 5 min on ice, do you mean to fix the whole tissue sample for 5 min or just the sections? Or would it be better to snap freeze the tissue sample, cryosection, and then fix the sections afterwards perhaps?
Your slices should be kept frozen until you are ready to use them. When pulled out of the freezer, put them in blocking solution and begin the staining procedure. Also, what kind of tissue are you working with? I can only speak for muscle and brain cryo/staining.
I found snap freezing of tissues embedded in OCT and direct sectioning without using any fixative is much better option to prevent autofluorescence.
Elliot J. Glotfelty Thank you for your suggestion and I think this does help although I've repeated it with unfixed tissues; we are working on subcutaneous fat pads, so they are a pain to section in the first place.
Shahid Nazir when you snap freeze a tissue sample -
1) do you cover the fresh tissue in OCT and then submerge them in dry ice cooled isopentane?
or
2) freeze the sample first in the dry ice cooled isopentane and then cover with OCT and freeze again?
The only experience snap freezing I have so far is for protein extraction so we didn't care about morphology when submerged in liquid nitrogen.
It all depends on the target antigen. if you are targeting immune cell markers. this method will work well. Fixation affects detection of immune cell markers and will create autofluorescence. i will just cover the fresh tissue in OCT and then snap freeze in liquid nitrogen and store at -80C. there should not be any direct contact between OCT/tissue and liquid nitrogen. This method works well for immunofluorescence study only where you are not much bothered about morphology. If you want to go for classical IHC then you need to modify it.
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