We are overexpressing different phosphomutants (serine mutated to alanine or glutamate) of a protein by lentiviral infection in C6 glioma cells. When we stain the cells by immunocytochemistry we see a clear overexpression for all different phosphomutants.
However, when we lyse the cells (1,5x Laemmli buffer, samples boiled and sonicated afterwards) and do a western blot with the same antibody, we see strong differences between the different mutants. Some bands are as faint as the control band, some as strong as the overexpressed wildtype protein, some in between. We already tried different antibodies recognizing different epitopes and also did a dot blot to exclude insoluble or degraded fractions, but the picture is still the same.
Does anyone have an idea what the reason could be for this discrepancy?
Thanks,
Britta
Thanks for your input.
@Sue Sha: the pH of the Laemmli buffer is o.k. and it is lysing the wildtype protein just fine. Or which pH did you mean?
@ Yang Liu: our protein can form multimers up to 24mers, but according to literature the phosphorylation actually shifts the equlilibrium towardssmaller 6-12x oligomers. So when the wildtype is denatured I can't explain why the phosphomimics are not...
Western Blot shows denatured proteins (due to denaturing SDS-Page). Depending on your immunocytochemistry protocol you may have a differently denatured or even naturally folded protein. Antibodies detect 3D-structures and denatured proteins will show different 3D structures. You should check first what your antibody is made for (e.g. WB, ELISA, FACS, IHC, ...), the supplier will tell you.
The antibodies we use all work for both IHC and WB. I also suspect some different 3D structures but I would rather have expected then that the signal in IHC would differ.
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