When collecting cell lysates for a Western blot, how do I induce di-sulfide bonds?

I would like to conduct a simple dimerization experiment for some protein I'm collecting from a cultured cells. My thought is, that if I'm running a non-reducing, denaturing PAGE gel, then removing beta-mercaptoethanol/DTT from the sample buffer should be enough to allow di-sulfide bonds to form.

I have seen several authors incubate the cells first with a drug called BSS that diffuses across membranes and creates protein cross-links. I may be wrong, but the use of this drug seems more appropriate when trying to follow up with an immunoprecipitation or a pull-down assay.

If anyone has any experience with this, could you please enlighten me?

For the introduction of new cystine bonds, you will probably need to maintain an appropriate redox potential when dialyzing your cell-lysate to allow your cystine bonds to reform. To break the current di-sulfide bonds, we have been adding in Disulfide Bond Isomerases (DsBC to be precise) which will allow your lysate to get to an appropriate equilibrium.

As for examining for disulfide bond formation that has already occured between proteins in your lysate, removing reducing equivalents like DTT and beta-mercaptoethanol should do the trick. You can even denature your proteins by incubating them in LDS or Laemmli buffer at 90C as those components will not reduce your cystine bonds. In our experience the bands typically run at the expected molecular weight on a SDS-PAGE.

Watch the video: Immunoprecipitation IP principles and troubleshooting (January 2022).