Dear all,
I am having considerable difficulty with separation of proteins on ief
gels. Specifically, I am looking for a 100kDa integral membrane protein
and a change in its phosphorylation state. Detection is by western
analysis. I have tried all combinations of the following additions to
increase the solubility of the protein in the gel and the loading
buffer:
2% Triton X-100
2% NP-40
2% CHAPS
9.5M urea
5% B-mercaptoethanol
Best results in terms of intensity of bands on a stained gel are with a
gel consisting of 4% acrylamide (37.5:1) and 5% ampholyte with the above
additions less BME. Sample buffer is the same without acrylamide but
including ampholytes and BME.
I focus in mini-gels at 500V for 4 hours. I can see plenty of bands on a
stained ief gel and on a stained PVDF membrane after semi-dry transfer
in 0.7% acetic acid. If I run the same samples on SDS PAGE I have no
problem getting a signal when blotting for this membrane protein when
the same amount of protein is loaded. If I blot my ief gels for soluble
cytoplasmic proteins I get good signals, and I observe bands shifting on
phosphorylation of these proteins. I am assuming that my membrane
protein (and maybe others, I haven't looked) is not entering the gel. I
have run ief gels with pH ranges from 4-6 and 3-10. The theoretical pI
(not necessarily worth much, I know) of my protein is 5.1, I am fairly
confident that the pH is such that it would enter the gel if it could.
Does anyone out there have any thoughts or comments on how I can improve
entry of membrane proteins into ief gels?
Thanks!
Will
