I am trying to purify a 10 kDa fungal protein with two predicted disulfide bonds by heterologous expression in E. coli with the aim of performing a pull-down assay to identify host targets. I had tried expressing the protein as a GST-tagged protein but had great difficulty solubilizing the protein. I had tried dropping expression temperature, inducer concentration, inducing at later stages of growth, adding reducing agents (10-20 mM B-Me, 0.25-5 mM DTT), adding 5% glycerol. Sarkosyl solubilization worked wonderfully but purification wouldn't work after solubilization. I tried adding Triton X-100 and CHAPS, changing salt concentration, changing the buffer composition, pH. Nothing worked.
I had prepared a his-tagged version of the protein for expression. Somehow, the his-tagged version is soluble. Although 50% of the protein degrades (likely his tag is degraded off) and ends up in the insoluble fraction, we still have a good yield of soluble, full-length protein. However, the yield from Ni-NTA purification is extremely small. I tried switching from 50 mM tris to 20 mM tris buffer, switching to 20 mM phosphate buffer, increasing the concentration of NaCl from 200 mM to 300, 400, 500 mM, adding 1-20 mM imidazole in the lysis buffer, decreasing the imidazole concentration in the wash buffer. I am still left with the vast majority of the protein in the flow-through. I am currently trying to add 10% glycerol and 10 mM B-mercaptoethanol as a last attempt before I'm out of ideas.
I had tried lysis and purifying under denaturing conditions (8M urea). Purification under denaturing conditions yielded 100% of the full length protein in the elution and zero protein in the flow through or wash.
If my protein is aggregating or improperly folded under native conditions but remains soluble and unable to bind the resin, could denaturing and refolding (slowly/stepwise) via dialysis return the protein to a state that would be able to bind the resin under non-denaturing conditions?
Thank you for any advice!
If you start with detergent-washed inclusion bodies from expression conditions that produce a lot of them, the protein will be nearly pure without any chromatography. You should then be able to get it homogeneous by Ni-NTA chromatography in 8M urea. Alternatively, if the protein is expressed in soluble form but extracted with 8M urea, you can do the Ni-NTA chromatography in 8M urea to purify it. Either way, you will then have lots of pure protein to explore refolding conditions and methods. If you manage to get the protein refolded and soluble, with proper disulfide bonds, a final gel filtration chromatography step can be used to remove residual aggregated protein.
Thank you for your response Adam. We were able to lyse the pellet in denaturing buffer (8M urea) and purify the protein successfully under denaturing conditions - all of the tagged protein remained bound to the resin and there was nothing in the flow through or wash fractions. If we were able to lyse and purify under denaturing conditions, elute the denatured protein, and then refold via dialysis - would we then be able to have the protein bind Ni-NTA under native conditions? The protein is soluble under native conditions and it does not bind the resin. Once refolded after denaturing, would it then be able to bind? I am assuming that it is perhaps misfolded or the his tag is masked, could refolding by dialysis change that?
Simple answer, yes it could. Do the experiment and you will be convinced or not.
I have had proteins that formed very large but soluble aggregates and found the same poor binding to IMAC as you describe. Using a resin with smaller beads increased the yield which indicates that the aggregates are not able to access the pores in the resin and only the ligands on the outside of the beads are used. In my case adding 2 M Urea made the aggregates smaller without denaturing the protein and significantly increased the amount that could bind to the IMAC column.
I am not sure , but checking this video may help
https://www.youtube.com/watch?v=dxdmmR7eRi4
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