I am purifying a recombinant protein (MW 37 kD) using IMAC. I used the Bradford colorimetric assay (by adding a drop of the solution from the column to Coomassie blue G-250) on the wash and elution fractions from the column, and observed the change of color to blue, suggesting the presence of protein. However, when I run SDS-PAGE there are no bands in the wash and elution fractions, while there are bands in the flow-through (including that of my protein). I get the same result after repeating the experiment. What could be a reason for this and the apparent false positive of the Bradford assay ? My Wash and Elution buffers are composed of HEPES, Nacl and Immidazole.
http://www.bio-rad.com/webroot/web/pdf/lsr/literature/Bulletin_6852.pdf has a reagent compatibility chart for various protein assays. Note that you should always measure protein concentration against a reagent blank that has the same composition as your sample, except for protein.
Protein precipitation with CHCl3/MeOH (doi:10.1016/0003-2697(84)90782-6) is a reliable way to prevent buffer interference in assays. Even simpler is spotting samples onto a PVDF membrane and then staining (doi:10.1006/abio.1994.1595 or doi:10.1016/S0003-2697(03)00255-0).
Hi Saman,
Normally, imidazole, HEPES, and NaCl in your solution are compatible with the Bradford assay. However, they have an upper limit. For example, 200 mM for imidazole, 100 mM (pH 7.5) for HEPES, and 5 M for NaCl.
You can check to see if it goes beyond these limits? Did you prepare or buy the Bradford reagent? If you bought it, you can check MSDS for compatibility values for the chemicals you use.
Hi,
In the study you mentioned, do you perform this single step purification at FPLC instrumentation?
If you see the FPLC-UV responses (peaks) of these wash and elution fractions, this may be also clue for confirmation of your colorimetric assay.
Do you have the possibility to perform BCA, biuret or Lowry assay for protein quantification alternatively ? In case of non-responsing from one of these test, than we may consider false positiveness at Bradford assay...
As Fatih Beris and Engelbert Buxbaum indicated, the concentrations of your buffers are critical and this case you may perform appropriate dilution using your column conditioning (loading) buffer or perform an online/offline buffer exchange using desalting resins to see their effects on protein quantification.
Thanks,
Emir
Fatih Beris , my Imidazole concentration is indeed 250 mM in my elution buffer. That might be the reason for the false positive, I shall redo it with reduced concentration. Thanks a lot for pointing that out.
İsmail Emir Akyildiz , I did not see any peaks on the FPLC response either.
Engelbert Buxbaum thank you for the sources.
Hello, 1st thing I want to suggest to change the % of gel used for SDS-PAGE. Stain the gel overnight with coomassie and destain on next day. I have also went through same problem with small 32 kDa. 2nd may be the concentration of separated protein is very less hence u can not see the protein bands in SDS-PAGE.
Measure protein using Lowry or Bradford assay along with a series of standards (e.g., BSA) and blank. Alternatively you can also measure by easy to use highly sensitive amidoschwarz protein assay (see reference below):
[Heda GD, Kunwar U, Heda RP (2014). A modified protein assay from micrograpm to low nanogram levels in dilute samples. Anal Biochem 445C:67-72].
Run a gradient gel (ex. 5-20%). Stain with colloidal coomassie blue, destain with water. If you can measure your protein by assay, you ought to find protein(s) on gel.
Hi Saman Salike,
Let us discuss that step by step:
1. the apparent false positive of the Bradford assay; does not mean you have proteins because also the buffer components react with bradford reagents. to exclude that you need to check if the elution buffer give the same color with bradford as the samples.
2. on your SDS-PAGE there were no bands in the wash and elution fractions, while there were bands in the flow-through (including that of my protein). that could means only one thing there was no binding with IMAC and I suggest that you check your protocol or change the methods, in the past I had great experience with Ni-NTA agarose beads if you are purifying a his-tagged proteins.
3. in case if you are sure about the IMAC protocol and that binding with your protein, then the problem is the protein concentration and sensitivity of Coomassie blue, I would suggest to use stain-free gel as the sensitivity is much higher than coomassie and also does not require any staining/destaining time.
let me know if I can help you with anything.
best of luck,
From this info it might be difficult to figure out your problem. since you have several variables. The problem could be with the buffer you used (some buffers are incompatible with SDS). It could also be with your gel itself, do you use molecular weight markers? are they seenable at the end of the run? You might also be using a gel percentage that's large enough to allow your protein flow through the gel.
Good luck!
Hi,
According to your explanation, your protein of interest did not bind on to IMAC beads. You should optimize your conditions. There are more than one proteins in your sample solution. Therefore, it is normal to see blue coloration.
I suggest you increase the first incubation time and decrase the protein concentration of the sample. Shake your sample and IMAC beads gently during the incubation too.
Good luck!
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