Hello Everyone,

So I usually purify hexahistadine KRas (1-166) through BL21-A1 expression system. This is the truncated g-domain of the protein. After affinity purification, I run it through FPLC SEC column and end up with a yield around 2.5 mg / Liter culture. 

Currently, we are trying to purify a new construct- a "cysteine light" version, with introduced cysteines on the surface in order to conduct screens. 

The yield for these constructs have been significantly lower (~0.5 mg / L).

I generally notice protein precipitation  / aggregation during crude protein concentration (after affinity elution and buffer exchange to remove imidazole). 

I've tried avoiding thiol reactivity by increasing TCEP in all my buffers up to 5mM. I've increased my NaCl concentration to 500mM, and I've kept everything at 4C. I even added 5% glycerol to my elution buffer to aid protein stability. 

The isoelectric point of the protein is around 5, and for the Cys light constructs I've been using 20mM Tris pH 8.0 for the buffer. For the original construct, I used 40mM HEPES pH 8.0. 

Should I revert back to the HEPES buffer? Any advice on how to increase yield? 

General purification scheme is as follows: Freeze down bacterial pellets, resuspend in lysis buffer with protease inhibitors and lysozyme (1mg/mL). Sonicate, then spin down. Bind to nickel resin at 4C for around an hour. Wash resin via batch purifiation scheme. Add 250mM imidazole elution buffer, and incubate 30 min at 4C rotating. Add beads to column, and collect elution at 4C. Buffer exchange to remove imidazole. Concentrate, then polish using FPLC. 

Any advice would be appreciated.

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