How can I improve protein purity without reducing yield? I encountered a problem that is quite common when trying to purify proteins: I lose significantly more protein when trying to increase purity. I am currently using Ni-NTA Magnetic Agarose beads from Qiagen, but I have tried several other types of beads, each returning similar elution profiles. In addition to changing beads, I have also tried varying conditions for the purification step as follows:
Wash the beads with 20 mM imidazole buffer, then elute with a gradient from 20 mM to 250 mM at 1mL/min over 30 min. With this method, I saw three contaminants that eluted at the same time as my protein: one with higher molecular weight, two with lower molecular weights. I changed the gradient to run for 60 min but obtained the same three contaminants. The contaminants begin eluting just prior to my protein, and complete their elution during the middle of my protein peak.
Wash the beads twice with 20 mM buffer, then elute with the same gradient described previously over the course of 30 min. With this protocol, I saw the same three contaminants eluting together with my protein, causing me to lose some of the protein.
Wash the beads at 20 mM, followed by another wash step at 40 mM, then run the usual elution gradient over the course of 30 min. In this case, I saw the same three contaminants again and lost even more of my protein.
Wash the beads with 50 mM buffer before eluting with the same gradient described. 80% of my protein was eluted, but it was accompanied by 100% of the contaminants. To separate the protein, I collected the 80% protein plus its accompanying contaminants in a 45 mL volume. I used dialysis to remove the imidazole and made a batch contact. Then I eluted three times with 1 mL 250 mM imidazole buffer each and found that 95% of my protein remained on the beads. At the end of this protocol, I got very little of my protein, but at least it was pure. However, solving the purity problem came at a high price in terms of quantity. My final yield was 0.3 mg from a 500 mL cell culture. I used 50 mM sodium phosphate and 500 mM NaCl with varying levels of imidazole for the buffers.
Can anyone offer suggestions for eliminating the contaminants while maintaining my protein? Additionally, I wonder why my protein elutes at such low imidazole concentrations (around 40 mM)? And in test protocol (iv) described above, why won't my protein elute from the beads?
It looks like the contaminants are interacting with the His-tag on your protein, causing it to elute at 40 mM imidazole. You could try to eliminate this interaction by using 6M urea in all your buffers.
I would like to do some structural studies with my protein, so I don't think it is possible for me to use urea.
You can remove the urea by dialysis after you purify the protein. Urea is only a mild denaturant, so your protein should be able to refold into the original structure.
Why don't you add another purification step? Size exclusion might work well depending on the exact sizes of the contaminants. In my experience, proteins at that expression level don't come clean with only one step.
I recommend using cobalt-loaded resin or Sigma His-Select, both of which I've found to have higher specificity.
In my experience, one Ni column seldom returns a pure protein. You can try ion exchange or gel filtration to further purify your protein.
I faced the same problem last year and was able to solve it using a His-tag purification kit from Thermo Scientific. You might try that kit as well.
I purify N-terminally His-tagged membrane proteins using Talon-cobalt columns under denaturing conditions (6 M Gu-Hcl to Urea) and my protein elutes at 0.5 mM-1 mM imidazole. There is a small yield, but it is quite pure. After that, I renature the protein by dialyzing it slowly in descending concentrations of urea, eventually ending with PBS. Since you are interested in the structural analysis but not the function of the protein, you can purify the protein under denaturing conditions suggested and later renature the protein.
Why does my protein lose its activity following affinity purification? I can partially purify my protein with Ni-His tag affinity chromatography, but activity assays show no activity. As a positive control, I used another pure protein of the same class that acts on the same substrate, so I don't believe the problem is with the assay reagents or technique. Why don't I see any activity? Could this be caused by co-purified products?
Did you use urea in the elution buffer? If so, did you refold the eluted protein? Are you certain that the His-tag doesn't interfere with the conformation of the protein? Your protein may be denatured or incorrectly folded following purification. Many proteins coordinate around zinc or another metal, so you may need to add some metal ions for full activity. Another possibility is that the high level of imidazole used to elute the protein from the affinity column may inhibit the activity of your protein. You can test this by adding some of the eluted protein to your positive control.
My protein is His(6X)-tagged and purified without urea. The buffer is Tris-HCl (pH7.2) and the activity of the positive control is not affected by high imidazole concentrations. There appears to be a product corresponding to the activity of the positive control protein in my test protein assays, despite running them in separate assay tubes. I tested all the buffers, cofactors (Mg and Mn), and reagents for contamination with the control protein, but there does not appear to be any contamination. Is it possible that two proteins in the same class react differently to differing levels of imidazole?
You should try reducing the imidazole concentration in your eluted protein to determine if that is causing the loss of activity. Did you assay the activity of your protein prior to applying it to the Ni-column? The His-tag may be influencing the conformation of the protein. If this is the case, you will need to move the His-tag to the other terminus. You could also try removing the imidazole from your purified protein by dialysis prior to testing for activity.