I am doing a Blue Native gel followed by a Western to detect protein complexes.
My detection antibody only recognizes a denatured epitope so I have to denature proteins after the gel is done. I tried doing that "in gel" by soaking in 2% SDS and DTT before transferring and the Western was blank. I can clearly see proteins on the membrane by Ponceau S, just no signal. Any recommendations to denaturing proteins on a PVDF? I have read that detergents can wash proteins off of PVDF easily so I can't do anything too harsh... Help!
Is performing a 2D electrophoresis out of the question? You could cut out the gel strip the band is in, denature the strip by boiling in SDS, Mercapto, etc... overlay it on an SDS-Gel, run it, transfer and then blot.
Those kits that AVEA posted look really nice! If you are looking for something quick you can also do a detergent fractionation. Digitonin is commonly used and I have used it with good success.
My general protocol:
To extract a cytoplasmic fraction, first make a lysis buffer that has a very low concentration of detergent (40 ug/ml) and load with cells for a short amount of time (2 min). Remove and retain sup and load a higher concentration of detergent to extract membrane proteins. I have used 1% digitonin for 30 minutes for this step. All steps are performed at 4 degrees C, rocking.
For reference, I use primary CGN's attached to a bromosilicate plate, at a density of ~4 million cells.
Well, if you are doing a western blot, you will not be seeing every band available. You will be using an antibody that is directed against your protein of interest. Since this antibody (should) only bind to your protein, you can then use a secondary antibody (chemilum. etc..) to visualize it, w/out getting other protein bands, right?