I performed protein extraction from cell pellets using RIPA1X buffer and ROCHE's protease inhibitors cocktail cOmplete. When take it to Bradford protein quantification (Bio-rad) protocol, I get all my standards and samples absorbance readings at 595nm over 1.000 , even in the blank with only RIPA1X buffer and no proteins. Today I ran a test with RIPA buffer 1x and 0,1x with and without inhibitors with no mayor differences at the same concentrations, discarding the inhibitors as the problem source.
Results:
1x w/o inhibitors: 1.104
0,1x w/o inhibitors: 0.451
1x w/ inhibitors: 1.008
0,1x w/ inhibitors: 0.438
dH2O: 0.397
Run a standard curve with BSA and 0,1xr RIPA buffer getting a curve with r2=0,89 but weird as if it was kind of "saturated."
What can be the problem? I already checked all the variables and have no answer yet.
EDIT.
RIPA Lysis Buffer :
1% NP-40
0.1% SDS
50 mM Tris-HCl pH 7.4
150 mM NaCl
0.5% Sodium Deoxycholate
1 mM EDTA
You might like to read the discussion here: https://www.researchgate.net/post/Bio-rad_protein_assay_reagent_changes_color_in_RIPA_buffer
I have the same problema as you when using Bradford, the solution for me is to use BCA Protein Assay as soon as I do not have urea in my buffer. Moreover the concentration in Bradford in my conditions is always understimated compared to BCA
Bradford assay does not like detergent. 0.1% SDS should be ok, but NP-40 and sodium deoxycholate likely not, especially with all three present. See
http://www.sigmaaldrich.com/content/dam/sigma-aldrich/docs/Sigma/Bulletin/b6916bul.pdf
The BCA assay is far less affected by detergents (reducing agents will interfere with either of these assays). See
http://www.piercenet.com/instructions/2161296.pdf
for a bca compatible list.
Always aim to dilute your sample in a buffer compatible with the assay if at all possible. Use a micro-format of the assay if your concentrations are low. Remember, as long as you add the same amount of reagent to your standards and samples alike, it doesn't matter much what the relative amounts are, although reagent typically exceeds sample by 10 or 20 to 1. Less reagent usually gives better signal from low protein concentrations, but you run into signal/noise problems when its too low. Plate reader formats work well.
I made 1:10 and 1:5 dilutions for samples and for the buffer to made the standars. Get a acceptable calibration curve, but my samples get abasorbance bellow the limits. I guess they had not enough protein for the dilutions.
Thx everyone. I will try to change to BCA assay o look for another protein extraction protocol.
I would suggest you should also try a simple buffer exchange step on an aliquot of sample (if you have enough of it).
This would probably give you the best sample quality for either bradford or BCA.
As mentioned by Peter Holt, Coomasie Brilliant Blue solutions develop color with detergents (NP-40, deoxycholate are both detergents). This information was included in the Mariam Bradford's original paper (Bradford, M.M. (1976) A dye binding assay for protein. Anal. Biochem. 72:248-254). You should remove the detergent or use a different method such as Lowry or BCA methods.
Complement: Look at his reference
Compton SJ & Jones CG (1985) Mechanism of dye response and interference in the Bradford protein assay. Analytical Biochemistry 151(2):369 - 374. http://dx.doi.org/10.1016/0003-2697(85)90190-3
Not sure if mentioned but if this is for LC/MS proteomics analysis, just precipitate out proteins using acetone or chloroform methanol procedure and resuspend in whichever compatible buffer then measure concentration...
1% NP-40 of RIPA could also be a problem , bcz when u look at the compatibility chart of Bradford assay, more than 0.5 NP is non compatible.
Bradford reagent is not compatible with buffers containing detergents. I suggest DC protein assay by Bio-rad to use with RIPA or NP40.
the problem maybe the 0,5% Sodium Deoxycholate
look at, pag. 2 : http://www.bio-rad.com/LifeScience/pdf/Bulletin_9004.pdf
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