Issues with Bradford Assay

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dhkwak
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Issues with Bradford Assay

Post by dhkwak » Mon Apr 19, 2010 7:05 pm

Hey all,

I have had some troubles with the Bradford method and wanted to know if anyone who has sufficient experience with this procedure could provide some insight.

First, I had produced my own Coomassie dye (using CBB G250) solution and had noticed that in solution (before being bound to protein), the solution color is a dark green instead of a brown-red color. I am wondering why this is occuring. I had added 0.01 % (w/v) CBB G250, 4.7% (w/v) EtOH, and 8.5% (w/v) o-H3PO4, as described in the original Bradford publication. Does the ortho-phosphoric acid yield results different from other forms of phosphoric acid?

Second, I had measured my protein concentration on one day and then again on another day and found that the concentration on one day was lower than that on the second day. I am unsure why this had happened. My protein sample concentration is about 100 to 200 ug per mL, which was stored in +4C in polypropylene eppendorf 1.7 mL centrifuge tubes. I'm confused about this and will be implementing the concentration assay again, but any insight would be very much appreciated. Thanks!

DK

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JackBean
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Post by JackBean » Mon Apr 19, 2010 7:26 pm

what other phosphoric acids are you reffering to?

what about precipitation? Did you mix the tube before?
http://www.biolib.cz/en/main/

Cis or trans? That's what matters.

dhkwak
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Post by dhkwak » Mon Apr 19, 2010 7:56 pm

Hey JackBean,

Thanks for the reply. I was too quick to ask about orthophosphoric acid vs. other forms of phosphoric acid, as they are essentially the same. I'm sorry about that.

I don't suspect very much precipitation and I did not see signs of such. I accidentally wrote 100-200 ug per mL, when it is actually 100-200 ug per 250 uL. I am using BSA in PBS, which is highly soluble in water. Unfortunately, I did not mix before measuring, except for pipetting several times up and down the tip before aliquoting. During each of the two separate measurements, I had a control BSA solution, which had a fixed concentration and was placed in similar conditions (+4C, same pipet-mixing) and that concentration was measured to be constant. I am wondering why the experimental concentration was so different.

Thanks again for the input.

dhkwak
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Post by dhkwak » Tue Apr 20, 2010 4:56 pm

Hey guys,

I had run some small experiments to see what I was doing wrong and though I am still in the process of more definitively figuring out what had happened, here are my thoughts:

The CBB-G250 I had used is in the green form. CBB-G250 can take on three forms: acidic (red-brown), neutral (green), and basic (blue). Because I had not adjusted the pH of my dye solution for the acidic form, the neutral CBB-G250 was not binding with the same affinity as it would for the acidic form. Moreover, when adding the dye solution to my protein sample solution, I observed some precipitation (thanks JackBean! :)) and I am not completely sure why I had seen precipitation and whether it was from the dye or protein (I had filtered the dye beforehand and did not observe any precipitation of the protein when aliquoting). BSA is reported to be very soluble in water (about 40 mg/mL) and I believe the pI is around 5-7. My protein was in PBS of pH 5 and 7 (as well as 9) and I had not seen any precipitation but I also only had orders of 100-200 ug per 250 uL (400-800 ug/mL).

Anyway, for anyone working with the Bradford assay and deciding to make their own dye solution, make sure the solution is acidic (turns red) and has a pH of about 1-1.5. Good luck all!

Edit:

Here is a helpful page to visit:

http://www.chem.iitkgp.ernet.in/faculty ... rdExp2.pdf

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