Hello, I am trying to purify a recombinant his-tagged protein (MW: 29.5 kd, pI:5.7, mostly composed of beta strands).
It has been expressed from BL21(DE3) competent cell. I am using 50 mM Tris, pH 8, 2 mM EDTA, 0.5% Triton X-100 with the addition of Complete Mini Protease Inhibitor Tablets as lysis buffer. As my protein forms inclusion bodies, I use 50 mM Tris, pH 8, 6 M GuHCl, 10 mM DTT as resuspension buffer. For refolding, I am using 50 mM Tris, pH 8, 4-0 M GuHCl, 0.4-0.1 M Arginine, 3 mM Reduced Glutathione, 0.9 mM Oxidized Glutathione. I equilibrate my column with 50 mM Tris, pH 8, 500 mM NaCl and then wash it with 50 mM Tris, pH 8, 1 M NaCl, 0.1% Triton X-100 (again wash with equilibration buffer to remove Triton). From there, I elute the protein from the resin using elution buffer (50 mM Tris, pH 8, 250 mM Immidazole). All the fractions are then pooled and dialyzed against 20 mM HEPES, pH 7.8
My protein shows very nice dark band on SDS-PAGE gel (from the elution fractions). I have tried to concentrate the protein using Amicon ultra-15 centrifugation filter unit and vivaspin 6 (MWCO 3kDa). I quantify my protein following Bradford assay using BSA standard. The problem starts after this point. Right after I finish concentrating the protein, it starts aggregating. I believe, there is something wrong with the solution. I have tried to store it at different storage condition. However, none of these worked. The concentration of my protein significantly decrease at all conditions. ( I can still see my protein on gel but bands are light).
I was wondering if anyone has experienced this kind of situation and what can be done to solve this. I would really appreciate any kind of help. Thanks in advance!
Some proteins have limited solubility. In such a case, avoid concentrating the protein beyond the limit.
It may help to include a detergent or to adjust the salt concentration or pH.
Thank you so much for your suggestions. I am looking for a suitable detergent but I am really confused about which one would be the best ( I have been reading about using a combination of strong and mild detergent also) and at which step I should start adding it, also what concentration would be best to start . Is there anything you can suggest on this please?
I think you can increase the salt concentration (0, 300, 600, etc mM) / lower the pH and monitor the aggregation states using DLS. After determining the critical concentration or pH for the aggregation, you can modify the protein solution before amicon ultra-centrifugation
If the protein is precipitating because of exposed hydrophobic patches, then you may be able to prevent it by adding a non-denaturing detergent such as n-octylglucoside , dodecylmaltoside, or CHAPS. These are just a few of the many detergents that are compatible with maintaining protein structure.
From your description it appears that you are attempting on-column refolding followed by elution using high imidazole buffer and then dialysis into storage buffer. Its not clear what salt strength you have in your 20mM HEPES pH 7.8 storage buffer. Also, how many cysteine residues does your construct have? You might alter the ratio of reduced and oxidized glutathione you are using during refolding. Often, switching to cystamine/cystieamine redox couple can also help. There are several strategies you might attempt here. One method is to use fast dilution method of refolding, rather than dialysis. In your final buffer you may consider adding glycerol or PEG. Also, avoid over-concentrating your protein. You might read this paper to get an alternative idea for in vitro refolding. Check the protein structure and see if mutating any residue could improve the stability/ solubility of the protein.
Article Structural Insights into N-Terminal IgV Domain of BTNL2, a T...
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