Hello everyone. We are recently starting to perform reversible biotinylation experiments. I created a protocol customized to our needs based on different sources (see below). In this protocol, we want to see the internalization of a receptor and for this we biotinilate at 4ºC the membrane proteins with Sulfo-NHS-SS-Biotin and later a treatment is carried out at 37ºC in Hek-293 cells transfected with the membrane receptor. After the treatment cells are returned to 4ºC and the S-S bond of the biotin is reduced with glutathione. This compound does not cross the membrane so it will only split the biotin that is marking proteins on the extracellular side, in such a way that when the lysates are prepared the only remaining protein is the biotinyled one (if any).

For the proper functioning of the technique we used a Biotin control (biotinylation of cells and then lysate) and a Stripping control (biotinylation and cleave of the biotin with glutathione and then make the lysates). The biotin control should give a strong band since all our protein of interest are biotinylated. The stripping control should not give band since we have cleaved all the biotins and we should not rescue anything. My result is exactly the opposite. I find little signal in the biotinylated control, and a lot in the stripping control. It has happened to us 3 times and I don't understand what's going on. In the image, the negative control are untransfected Hek cells and the positive control are transfected and lysated HEK-293 cells without biotin. In addition, after doing the lysates we did not find tubulin in the samples so it seems that the rescue of biotin is being adequate.

We are using Ez-Link Sulfo-NHS-SS-Biotin (21331, Thermo) and L-Glutathione reduced (G6013, Sigma).

Any ideas about what may be going on? Thank you very much

If you need any additional detail about the protocol, do not hesitate to ask.