The tissue is fixed in 4% PFA/PBS (no methanol) and ground up for approx. 5 min. in 1% Triton-X 100 and 40 mM Sodium Citrate for subsequent flow cytometry counting of free nuclei - then washed and stained as per usual (2 washes after the primary stain. I’ve used different incubation times, temperatures, PBS types, secondaries (Alexa Fluor 647 and 488), freshly fixed (1-2days) and older brains, nothing works. I use up to 1:10 dilutions of MAB377 in a solution of roughly 10^6 cells/ml. If anything, the nuclei are darker than the tissue debris under the fluorescence microscope, it is as if nothing gets in. I have used two batches of MAB377 (Merck Millipore) – they seem to have both been frozen (the latter batch very briefly on its way to me) but surely that won’t totally destroy the antibody? Any ideas or alternatives?
Test the antibody on tissue fixed with other fixing solution. Even in frozen tissue samples (following the procedures for obtain cryostat sections). Test another antibody against other nuclear antigen to test how is your tissue preserved. As a final option, increase the time in triton X-100 an the wash the sample in tap water if your probe allow it. Good luck.
Thanks very much! I will try all this and let you know if (and if so, what) worked!
How long are you fixing for? Maybe adjust your fixation/personalisation steps
Hi All, thanks for your responses. As an update, it may have been the fixative indeed - the protocol seems to work with specimens fixed in formalin that was _first_ dissolved in distilled water, and had 10x saline added afterwards (I had previously cooked up the PFA in saline, which seems to diminish its fixation abilities according to a histologist I talked to). But I still need to confirm that this really is the main problem. The fixation times are really long - weeks up to 1 year - and fixation in the "wrong" formalin seems to very quickly degrade the ability of the sample to stain. Hopefully I'll figure it out soon and I'll post a final solution then :-).
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