I am making a total protein extract from a bacterium. In the final buffer I diluted the sample with 7M Urea, 2M Thiourea, 4% NP40, Tris, PMSF, EDTA and DTT. I have tried to quantify the sample with BCA and Bradford, but it interferes. Anyone have any system for quantifying protein extracts with these conditions?
I am using a method that a researcher for my lab told me. You spot the protein and BCA dilution in a PVDF membrane using a dot-blot system from BioRad and after you stained with ponceau red.
you can also try with lysis buffer (9 Murea, 2% CHAPS, 13 mM DTT, 1% IPG buffer) and then you go for Quantification of proteins by the Bradford method
Hi all,
thanks for all your answer. I attach the protocol that the BGI company for the proteomics service send me for obtain the extract of protein. I don´t have more experience with total extract, I think that one possibility it´s resuspend the protein in the buffer without the DTT, take a sample for the quantification and after these adding the DTT, what you think?.
Thanks.
César.
Thiols and detergents often impede protein quantification. Try precipiation (see attachment)
HI there, this is a routine issue for protein quantification as there is NO set way to properly quantify your protein using the current assays - something will ALWAYS interfere. BCA can tolerate detergents, but only so much, however they are limited by the addition of reducing, chelating and phospholipds! Its mind boggling. An article that shows how each method interferes - Bailey & Noble (2009) Methods in Enzymology, clearly shows there is no complete method. New technologies are out there to help overcome this, in particular Infra-Red based systems, that measure the Amide bond as opposed to relying on the protein sequence. Perhaps this is something worth investigating - www.emdmillipore.com/directdetect
Butterfly procedure-Protein quantitation using bound Coomassie blue stain
Samples (5-20 μl) and BSA standards (1–10 or 1-20 μg) are spotted on filter paper strips (0.5x1.5 cm, Whatman 3MM).
Filter papers (numbered in pencil) are stained with Coomassie brilliant blue G-250 (Bio-Rad, USA; 2.5 g/L in 40% methanol, 10% acetic acid)
and unbound dye is washed with destain solution (20% methanol, 7% acetic acid).
The papers are transferred to 24-well plates, one sample per well.
Bound Coomassie stain is extracted from the filter papers with 1 ml of 3% SDS.
200 µl from each sample are transferred to a 96-well plate.
The absorbance of the solution is measured at 590 nm.
Protein amounts are determined using the bovine serum albumin calibration curve.
Additional notes:
STAINING AND WASHING STEPS MUST BE PERFORMED IN A CHEMICAL HOOD!
The SDS extraction is faster with gentle shaking at 37 degrees than at room temperature. But if it's the end of the working day, and you're not running your gels until the next day anyway, you can just leave the samples overnight on the bench.
It is very important to run duplicates.
Also, the volume of sample depends on how much protein you think you have. When I used to do yeast lysates, I had to dilute them to be in the linear range.
Personally, I think it is a tedious and inaccurate method. But it's cheap and sometimes nothing else works like in your case..
I am using a method that a researcher for my lab told me. You spot the protein and BCA dilution in a PVDF membrane using a dot-blot system from BioRad and after you stained with ponceau red.
Our experience is that Bradford + 2% Urea is really not accurate.
We also perform a blotting of protein on membrane like César.
i found a Bradford Method to Estimate in presence of SDS
if you add 3µl of Sample Dilution Factor will be 100, at 5µl its 60 and at 1 its 300
i did not get good results by pipetting only 1µl thou
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