I have tested several induction conditions for expressing my protein, I am using the pet28b vector(C terminal-his tagged), expressing in E.coli BL21(DE3). First I started with 0.1mM/ 0.2mM /0.5mM/ 1mM IPTG at 37C for 5 hours and took a sample every one hour but unlucky the 0.1mM after 1 hr formed inclusion bodies in the cell debris after cell lysis. Second I tried 0.1mM/ 0.05mM/0.02mM IPTG at 16C for 21 hrs and took samples at 13/16/18/21 hrs but still 0.1mM & 0.05mM showed inclusion bodies at 13 hrs while 0.02 almost show no prominent induction. I also worked on petSumo vector (N-terminal his tagged) & still have the inclusion bodies.
The problem is that I even did an uninduced culture with the same conditions (overnight at 16C) as a control. It's expression pattern was lower than the induced but still most of the protein is in inclusion bodies. My protein was synthesized with optimum codon usage for E.coli to avoid low expression but now I can't get it in a soluble form and I usually start induction after reaching OD 0.5-0.6. Is there anything I can do to avoid this? I tried adding 30% glycerol to the cell lysis buffer: the supernatent showed a faint band of the protein compared to samples without but still most was in Inclusion bodies and not enough for protein purification. Just wondering if there is anything I can optimize to avoid this, or since even the uninduced has inclusion bodies, is there no hope? I want to avoid as much as possible denaturation/renaturation process.