I have been modifying a protein through a radical reaction, which conjugates to tyrosine. As such, measuring protein concentration by absorbance at 280 nm is not working anymore. So I was told, "Just do a Bradford!" So I did a Bradford assay using BSA for my standard. The BSA absorbs at 595 nm just like it's supposed to. However, my protein has a peak at 660 nm and no peak at all at 595 nm. The conjugation absorbs at 550 nm and it must be what is causing problems, but that absorbance is not showing at all. What can interact with Coomassie G-250 to make it absorb at 660 nm?
Weird problem. Maybe you should use BSA that has also been through your radical reaction as the standard.
Adam B Shapiro,
That was suggested by PI and ran that reaction today. It turns out that modified BSA and the unmodified protein of interest do not show a peak at 660 nm. So now I'm at a loss. I'm curious if it's concentration-dependent, so I'll try that next.
So I had buffer exchanged my protein into a buffer for a protease digest before concentrating. That buffer contains 0.5% SDS, which I thought was low enough to not interfere with the assay. However, my modified BSA shows a peak at 660 nm when in the digest buffer and the solution turns green.
Avoid having detergent in the Bradford assay, if possible. If it can't be avoided, make sure to have a consistent amount in all samples and standards. Here is a compatibility chart for Bradford reagent.
https://zageno.com/l/ts-bradford-assay
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