I am working on human oligodendrocytes precursor cells, studying their differentiation into oligodendrocytes. I have a problem recently in staining O4, I do live staining 30 min incubation with primary antibody at 37 degrees, then fix using 4% PFA for 15 min, wash 3 times with PBST then block with 5 % goat serum for an hour and secondary antibody incubation for 1 hr, then wash again with PBST 3 times. I get positive staining which are bright and visible fluorescent oligodendrocytes but not all cells in my fields are stained not even a weak stain they are totally missing though I can see them as moderate to highly branched oligodendrocytes in the phase contrast images. If you have any feedback, I just what to know what is the technical issue behind this problem.
Hello, I am not sure which antibody you are using but I had the same problem with the mouse anti-O4 IgM from R&D(MAB1326). The tricks are to avoid any permeabilization before the secondary antibody to avoid internalization of the complex but more importantly, if you use this antibody is to use an anti-IgM secondary instead of a classic anti-IgG, when I introduced these changes it was like night and day. Here is my procedure:
1. Live staining at 37°C for 30min-1h
2. Wash in PBS (not PBST!!)
3. 10min fixation in 2%PFA
4. Wash in PBS
5. 1h anti-IgM secondary in 5% serum (important: don't use any detergent like Triton)
At this point if you want to counterstain with another marker you can proceed with permeabilization and blocking as normal.
I hope this helps.
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