I am working on a membrane protein. After I mix the lysate from the solubilization of membrane from pichia yeast and nickel beads, I washed thoroughly with 20 mM and 40 mM imidazole, and elute with 350 mM imidazole. After I ran the SDS-PAGE, i saw a major band and many other upper bands. The purity is only about 50%. I want to further purify my protein.
First I tried to use the Q column. But I am not able to separate the major peak and other peaks.
Second, i tried to add a TEV site before His8. I cut with TEV, and further load into Nickel column. However, the cut protein is still not pure.
Can anyone suggest anything to improve the purity of my protein to 80%?
Can i load my his-tag protein into the second nickel column to further purity my protein?
thanks.
Do you use pPICZ expression system to induce your protein? I used to express secretory membrane protein with pPICZalpha vector. Single band was obtained without problem.
Hi there,
You might consider more steps in your protocol: going from 40mM to 350mM imidazole is a big step. Either you could include more steps at 100 an 200mM (for instance) or set up a gradient of imidazole to increase the resolution.
What detergent do you use? If it's something with small micelles, you could try size exclusion (or sample with Triton on column for Xlarge proteins).
Other option would be hydrophobic interaction (Octyl/Phenyl etc.). But on that one I usually lost lots of activity. But it depends of course on the protein and how will you do it.
I agree that some kind of affinity column would be best if the imidazlole gradient doesn't work. You can also try cobalt resin for the His tag purification. It has less non specific sticking.
I second on trying Cobalt resin, it is said to have higher specificity than the nickel for His-tags. Otherwise, if your "upper bands" have quite a large size difference, a size exclusion chromatography might work.
Alternatively, I think your other bands might be other membrane associated proteins that complexes with your protein of interest. If they are unwanted, you can probably add in or increase the detergent concentration to try and dissociate them from your POI or purify your protein under reducing conditions (only if you can fold them back though)
You may add 10 mM Imidazol to lysis buffer, increase salt concentration for washing. If still not pure enough, you may try a gel-filtration by a superdex 200 column. If you are doing protein purification under denaturing conditions, you may wash and elute by lowering pH of the buffer instead of increasing imidazol concentration.
Hi, edward,
I think you are right. my other bands might be other membrane associated proteins that complexes with my protein of interest. So you think that i can add in the detergent concentration to try and dissociate them from my protein? I used 1.5% DDM to solubilize the membrane and used 0.02% DDM in the purification process. How do i do this? So also I add BME as the reducing conditions? THANKS.
Hai
Hi Edward,
First, I would try to determine the nature of the contaminants. Are they product related? Do you have a specific antibody (or you could try the anti His antibodies) to your protein? Otherwise you could try mass spec. I suppose I'm wondering if the upper bands are modified forms of your protein.
good luck!
Vince
many times membrane proteins bind to lectin columns because they are glycosylated. I would try Concanavalin A or Wheat Germ Agglutinin.
Hi Hai,
Apart from adding in detergents in your purification, you can also consider reducing the salt concentration (commonly at 300-500mM for His tag purification). The high salt concentration used during purification is to reduce non-specific binding to the nickel, but it might increase hybrophobic interactions between proteins, which in your case of membrane proteins, it might cause it to bind tighter. Im not too familiar with DDM, if it works for you, go ahead, if not, perhaps try 1% triton-x or 0.1% SDS.
Cheers
Edward
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