Hi there, I try to get a membrane protein RAGE in monomeric/dimeric form. After overexpression e.coli and Ni-agarose purification I have a high order oligomer (~5*10^6 Da, light scattering result).
Protein purification is performed in denaturing conditions (Lysis & wash buffers: 8 M Urea, 0.1 M Tris, 0.3 M NaCl, 5 mM DTT 40 mM Imidazole; elution 0.1 M Tris, 0.3 M NaCl, 5 mM DTT 500 mM Imidazole). On SDS-PAGE I have a monomeric form, but this protein is denatured.
Protein need for incorporation in nanodiscs, so I need it monomeric form. Can I use such detergent like sodium cholate? Is it compatible with urea? Can I elute the protein in 0.5% of sodium cholate?
Thanks!
If the membrane protein was not aggregated when it was dissolved in 8M urea (something you should check), washing out the urea without adding a detergent caused it to aggregate. It may not be possible to disaggregate with detergent once it has already aggregated. You may have to disaggregate it with urea, or start a fresh prep. Add the detergent while the protein is still in urea and bound to the column, then gradually reduce the urea concentration while maintaining the detergent concentration at several-fold above the critical micellar concentration. Once the urea is gone, then elute the protein.
Dear Adam B Shapiro, thank you for the quick response! During protein extraction should I keep the detergent in the solution?
Can I run SEC without detergent for checking result? Maybe should I do a native PAGE?
Whether or not you use detergent in the cell extraction depends on where in the cell the protein is to be found - in the membranes, in inclusion bodies, or as an insoluble aggregate. Since you did the chromatography in 8M urea, I'm assuming you started out with inclusion bodies o an insoluble aggregate. You don't need to use detergent to extract inclusion bodies, but a gentle detergent wash of the inclusion bodies may help clean off contaminating proteins. If the protein is not in inclusion bodies but just needs some help to get into solution, extracting with detergent may do the job.
Membrane proteins are usually not soluble in the absence of detergent or chaotrope. If you try to run the protein on SEC without detergent, it is probable that the protein will precipitate inside the column. A solution to this problem is to use a detergent in the chromatography buffer that has a small micelle size, such as CHAPS, at several-fold the CMC. The high detergent concentration assures that each micelle has only one protein molecule. The small micelle size avoids causing the apparent molecular weight of the protein to be elevated.
There are 3 important considerations if you use Tween-20. (1) If the concentration used is higher than the critical micellar concentration (CMC), the protein will be embedded in micelles and will elute as if it were a larger protein. (2) The low CMC of Tween-20 makes it hard to remove, in case it is not compatible with the nanodisc preparation. (3) If you do use it, be careful about the source. Oxidation of Tween can lead to formation of protein-destroying peroxides.
Earlier you mentioned using sodium cholate. That is related to CHAPS and may be a good choice because of its small micelle size, as long as it is compatible with nanodisc preparation.
Dear Adam, thank you for your answer!
I will use buffer for SEC with sodium cholate. Since the CMC of sodium cholate is 9-15 mM and it forms a small micelles (900-1300) I can prepare buffer with ~ 12 mM sodium cholate. If the protein will be embedded in micelles the molecular mass will not much higher.
I have a question about sample preparation for SEC. After Ni sepharose purification my sample buffer contain 8M urea. I will change buffer using centrifugal filter concentrators for buffer that I will use for SEC. Is the concentration of sodium cholate in this buffer the same as SEC buffer (12mM) or higher? The protein in this sample will be concentrated.
The goal is to prevent aggregation of the denatured protein when the urea is removed. Therefore, the detergent must be added to the protein while it is still in urea. The concentration to add should probably be several times the CMC to make sure that there is enough so that each micelle contains only one protein molecule. You might consider using 50-100 mM cholate. The cholate will not be concentrated by ultrafiltration because of the small micelle size. You may not need to use such a high concentration in the SEC column, but the concentration should remain above the CMC. Do not concentrate the protein any more than necessary, since higher concentration favors aggregation.
I tried to do it, but experiment wasn't successful. You can see at attached files SEC of RAGE in 18 mM sodium cholate and nanodiscs that were prepared using collected RAGE.
In case of RAGE SEC, target protein was observed in the second peak 12.25 (Mw ~ 600 kDa, RAGE 46 kDa). I couldn't understand is it result of sodium cholate or RAGE formes oligomers in sod ch. solution.
I investigated nanodiscs they didn't contain RAGE. Maybe it is result a high MW of RAGE in sodium cholate.
I didn't check if the RAGE was not aggregated when it was dissolved in 8M urea . It is my mistake. Is it possible to retain agregate for protein during ni-sepharose purification in 8M urea? How to completly destroy agregate?
I have performed such experiment for SDS-PAGE samples: I used sample buffer with DTT and SDS but I didn't heat sample. And my protein didn't come into gel. Maybe I have a protein complex that i couldn't destroy.
It is possible for a protein to remain aggregated even in 8M urea. You can try a stronger chaotrope, such as guanidine HCl or guanidinium isothiocyanate.
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