I have been trying to stain G4 with BG4 antibody for quite a while now. I tried the method that was used previously in our lab, which involved first blasting cells in 75mM KCl for 30min and then sedimenting nuclei with Cytospin. This is where it constantly goes wrong for me, because not every "boat" in the machine will sediment the nuclei flat, in majority of cases they will kind of look crinkled on the cover slip. I guess the Cytospin itself might be the problem because it is quite old and every time I use it, getting flat nuclei is really random (does not depend on the "boat" that I use). So instead I tried fixing cells growing on cover slips with 4% PFA for 10min and then permeabilising with 0.5% Triton X for 10min. The antibody binding was not very good. Would it make more sense to fix cells with methanol instead?
Methanol fixation preserves protein epitopes differently than PFA. Since BG4 is an antibody against G4 structures, methanol might help maintain the integrity of these secondary DNA structures better than PFA, which can crosslink and potentially mask them. You could also try an acetone/methanol mix (1:1) if methanol alone is too harsh. Another alternative is using a milder fixation like ice-cold 70% ethanol, which may strike a balance between antigen preservation and structural integrity.
If methanol fixation improves nuclear morphology but still doesn’t give strong BG4 staining, you might need to optimize blocking conditions or antibody incubation times.
Attapong Sinkitjasub thank you very much for your recommendations. I managed to figure out why cytospin was not working and I also changed the extraction buffer. I still have to pre-extract nuclei because either anti-G4 or anti-FLAG bind something outside the nuclei if I fix whole cells. However after I extract and sediment nuclei, I proceed with 100% methanol fixation and now I get very clear foci after BG4 antibody staining, much better than after PFA fixation.
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