Hello everyone,
I have been using the Echelon lipid-coated beads for studying the interacting domain(s) of a purified recombinant protein (GST-tagged) and its truncated variants with some of the phosphoinositides present on the beads. I have encountered several issues and I would appreciate if I can get some advice on troubleshooting.
1) Before each pull down I block the beads with 3% BSA for one hour at 4C. I do this because I get a lot of background binding of my protein of interest to the control agarose non-coated beads. The blocking, I presume, does not seem to be working efficiently. Is there a better alternative to BSA for blocking?
2) One of my proteins is especially tricky to purify and it is present in very low concentrations. It is in a buffer supplemented with 10% glycerol and 500mM NaCl. Because of its lower concentration, I used bigger volume (50ul protein in 200ul binding buffer; 4x dilution). I did not observe binding of my protein to the beads, and I suspect that glycerol might be interfering. Could that be the case? Would increasing the dilution factor (let's say 50ul protein in 5ml of binding buffer per 25-50ul beads) be a good idea, or is there a ration between beads:binding buffer that I need to stick to?
3) Is it a good idea to change the tubes between washes?
4) Why is it so important to use fresh Laemmli buffer? Does it make a difference if it is 5x or 2x Laemmli?
Thank you.
2/10/2020 3:37:42 AM Reply