I'm trying to do immunocytochemistry and localize a protein that is known to reside in cell membranes. I plan on comparing this protein to a mutant version and (hopefully) demonstrate that the mutation causes localization problems.
The problem is that after 4% paraformaldehyde fixation (10min at room temp) and permeabilization with 0.5% Triton X-100, there appears to be no sign of membrane localization in the wild type cells. There's absolutely no signal from the nucleus of the cell, but the wild type protein seems to be scattered evenly throughout the cytoplasm, and there's no clear localization on the cell membrane. This is interestingly the exact same pattern I see in the mutant cells. Given that the mutation results in a truncated protein well before the transmembrane domain, I'm very surprised at this result.
I've seen articles claiming that Triton X-100 obliterates the entire membrane, and leaves any integral membrane proteins scattered around so they can't be visualized. If this is the case, perhaps any protein bound in to the membrane in the wild type cells has been washed away and thus would just appear like the mutant cells. Has anyone had this experience before? If so, what do you use to permeabilize cells with integral membrane proteins that you want to localize? I'm planning on trying again with 0.1% Tween-20 to see if that helps, unless of course there are other suggestions/recommendations.
Thanks!
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