Eliminating Protein/Protein Interactions for Western Blot
Posted 07 December 2012 - 09:22 AM
The predicted size is ~95 kDa, but we consistently see a single, clear band at ~150 in a range of samples.
I've performed a knockdown experiment that seemed to confirm the 150 kDa band is our protein. Post translational modification or alternative transcripts do not appear to explain the discrepancy.
I've been stumped, but I found a paper yesterday that indicated the protein forms a 158 kDa complex with another protein, and that that complex is difficult to break up. In the past I've increased BME and increased heating time/temperature without effect.
I want to run a western blot today with the harshest possible conditions for denaturing, but I'm finding a lot of conflicting info with google.
My short list of conditions to try is:
LDS 4x sample buffer + 2.5% BME, heat 70 degrees for 10 minutes (our default prep)
LDS 4x sample buffer + 50 mM DTT, heat 70 degrees for 10 minutes
LDS 4x sample buffer + 8 M Urea + 50 mM DTT, heat 70 degrees for 10 minutes
LDS 4x sample buffer + 8 M Urea + 50 mM DTT, heat 95 degrees for 10 minutes
This is with the Xcell II blot module, NuPage MOPS running buffer, NuPage transfer buffer, and nitrocellulose filter paper.
Are these conditions stupid/are there others I should be trying? Is there any reason I can't run Urea and non-Urea samples on the same gel? Do I need to add Urea to our ladder (Biorad Precision Plus Dual Color)?
Posted 07 December 2012 - 08:33 PM
The usual concentration for DTT is up to 150 mM or 300 mM for B-ME, which will give the same number of reducing sulphur groups. Boiling at 95 with 150 mM DTT should be enough to denature most proteins in my experience. Ensure that both of these are fresh as they will quickly degrade in solution unless frozen.
Posted 10 December 2012 - 10:12 AM
I'll try the 150 DTT + boiling @ 95 too.