I am trying to purify my his-tagged protein of interest, disulfide isomerase. It is about 40kDa and is cloned in pET28a vector, at XholI and NdelI, and expressed in BL21.
I'm having issues with my protein purification.
My current protocol:
- 1L culture incubated at 37°C for 6 hours until OD600 reaches 1.5-2.0.
- IPTG is added to a final concentration of 1mM and is then further incubated for another 3-4 hours until OD600 reaches 2.2-2.7.
- I then lyse the cells using 1X binding buffer with 1X Protease inhibitor, no urea, 0.1mg/ml lysozyme, and DNase1.
- solubilize/sonicate with 1X binding buffer WITH 8M urea and 1X protease inhibitor.
- purify my protein by doing affinity chromatography using a nickel column with 1mL of Ni-Nta resin.
- I washed twice with 1XBB with 8M urea, 15-20mM imidazole, and 1X protease inhibitor.
- elute with 1XBB with 8M urea, 250mM imidazole, and 1X protease inhibitor.
During the solubilization step, I sonicate twice. After my first sonication and spinning down in the centrifuge for about 30min, my supernatant comes out super viscous and when I run it through my ni-nta column, it ends up clogging it and causing it to run super slow (1 drip = 5min). What can I do to fix this issue?
Should I try using 6M urea instead of 8M? Would that help with the viscosity?
Also, my elutions come out really dirty still with a lot of contaminant protein. How can I clean up my elutions?
My protocol for my wash calls for protease inhibitor, but I feel like that's a bit wasteful. Can I wash without using protease inhibitor first and then do another wash with the protease inhibitor? Or should I be using protease inhibitors in every wash?