Purified proteins using CHAPS/Urea method for running 2D-DIGE (compatible buffer).
Need to quantitate using Bradford so final concentration of sample is 50ug/10ul.
HOWEVER, when I diluted my stock sample in water (as per the 2D-DIGE experts advice - they like to quantitate in water) to do the Bradford assay, it didn't appear to solublize well (got a clearish pellet in the bottom of the tube) and when I did the Bradford assay I got a protein concentration reading of say 10 ug/ul.
SO, I tried diluting my stock sample in PBS, and it appeared to dissolve much better, there was no precipitation or pellet in the tube, but I only got 1/2 the concentration (say 5ug/ul) of the EXACT same amount of sample.
I must have a final concentration of 50ug/10ul for the DIGE reaction to work properly. If I base my dilution on the readout from dissolving in the protein in water does this result in a 2-fold OVERestimate? Or if I base my dilution on the readout from dissolving the protien PBS does this result in a 2-fold UNDERestimate?
Do I go with the water or the PBS reading?
The PBS versus water does not affect the standard curve nor the Bradford dye.
It's just what stays soluble in my sample when i try to dilute it for the Bradford assay.
What would you do?
Has anyone encountered this?
Oh - and if there is TOO MUCH protein in the final sample for DIGE, it will preferentially label the OVERabunant protiens and I won't get good results. If there is TOO LITTLE protein in the final sample for DIGE, the labelling reaction will not proceed well. Therefore, I really need the right range of protein!
Edited by Labscience, 10 November 2009 - 11:35 AM.