I am trying to detect phospho EGFR in Hep3B cells by western blot exposed to varying concentrations of an EGFR inhibitor. I get one band at ~170kDa as expected, but also a band at around 130kDa. I have read that EGFR is usually glycosylated and the glycosylation brings the mw from 135kDa to ~175kDa.
Is it possible I am detecting the deglycosylated version? Why would it be deglycosylated in normal conditions? I think both bands are specific as they are both affected by the dose of the inhibitor. I haven't yet tried looking at total EGFR (non phospho). I am using normal measures to prevent degradation e.g. protease inhibitor and keeping lysates on ice.
Thanks
Did you try to detect purified glycoproteins instead of the cell lysates? Did you use a monoclonal or polyclonal Ab as your primary Ab against your glycoproteins? Do not you think protease-mediated degradation can not occur inside the cell? Did you use purified non-glycosylated and glycosilated proteins as controls?
You may have detected EGFR that has been translated but not yet glycosylated, a biosynthetic intermediate, in other words. Or you may have detected proteolysis at a hypersensitive location. If you have access to monoclonal antibodies with N-terminal and C-terminal epitopes, you can use them to distinguish between these possibilities.
Glycosylated proteins give broad bands in SDS-PAGE because of charge heterogenity in the polysaccharide. The unglycosylated precursor gives crisp bands.
You can also try to stain the sugar moiety with the periodic acid/Schiff-reaction (PAS), however, instead of the classical Schiff-reagent I'd use modern fluorescent stains (e.g. Thermofisher Pro-Q Emerald) to increase sensitivity.
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