My goal is to purify a 37kDa His-tagged protein that is expressed by BL21 E. coli cells. Initially I had followed the directions included in the HisLink Spin Protein Purification kit by Promega. The protein is being expressed (confirmed by SDS-PAGE) but is refusing to bind to the Nickel resin. In every of my past attempts, SDS-PAGE shows that a small amount of the His-tagged protein binds to the resin, but the majority of protein that does bind ends up coming off during the 1st washing. A very small amount is recovered after eluting with imidazole.
This led me to suspect that maybe the protein is bind too STRONGLY to the resin, so that it appears faintly in the 1st wash and the elution, however it is mostly still on the resin. I tried imidizole concentrations up to 600mM (elution buffer) and still no difference.
Past attempts for purification:
*All attempts included binding/washing buffers with NO imidizole at pH 8.0
*The kit is brand new (new resin, buffers, etc.)
1) Varying NaCl concentration in binding/wash buffers (0mM, 200mM, 600mM)
2) Denaturing conditions (8M Urea)
3) 100mM Tris binding/washing buffer (usually I use 100mM HEPES)
I don't even use the filter column included in the kits to do the washing/elution anymore. I have found protocols where pelleting the resin and transferring the supernatant is mentioned, and I'd rather save the filter columns for when I actually have a sample worth purifying through filter.
I am experimenting with using a high concentration of culture per purification. I have been using about 15mL of liquid culture per run, thinking about bumping it up to 100mL (suggested for low level expression proteins). Maybe an even higher imidizole concentration in the elution buffer, 1M?
I tried to be as detailed and concise in sharing my thoughts but probably missed most of the important bits. This has been a very frustrating journey...I will upload SDS-PAGE gel pics soon my SD card port is busted.. Any suggestions are greatly appreciated. Thank you.