I have done 2 purifications using a Ni-NTA column so far and each time we've noticed (based on my SDS-PAGE results) that lots of protein is collected in my flow-through (not binded to the resin).
We've tried incubating the resin with the protein at room temperature, rotating for 2 hrs. We've also tried using a gilson minipump to pump our solubilized protein solution into a column at a flow-rate of 2 mL/min.
Both procedures resulted in low yield of protein and lots protein located in the flowthrough.
Best binding occurs at pH 8.0. The more concentrated your protein the better it will bind. If you buffer contains amino acids or other inhibitors (see table 4 on page 74 of the QIAexpressionist.pdf from https://www.qiagen.com/us/resources/resourcedetail?id=79ca2f7d-42fe-4d62-8676-4cfa948c9435&lang=en) then you have to exchange the buffer before you can purify your protein. We exchange buffers using a pressure-based stirred cell concentrator. They come in many sizes from EMD millipore. You have to use a filter with an appropriate molecular weight cutoff to retain your protein in the concentrator.
In order to answer the question appropriately we need more information. How do you know that the protein of interest in present in the flow-through? What is your buffering system? pH? How much culture are you lysing prior to loading your resin? Do you clear your lysate with centrifugation? Do you filter sterilize your sample before loading the column? Have you tried both N and C terminal HIS6 tags with your protein construct (sometimes the termini interact with the protein decreasing the interaction with the Ni resin)?
There is a maximum binding capacity of protein to IMAC resins. So... Make sure you aren't overloading the column to begin with (~10-15mg/ml of HIS6 tagged protein is a good approximation). I usually have ~30mM Imidazole throughout the loading and column wash steps to prevent non-specific interactions with the resin. The pH should be between 7-8.5. Also, try recharging your IMAC resin with Ni2+ to see if that affects your purification.
I would try loading as a more dilute solution without any imidazlole in the buffer and use the link provided by Christopher to see if you have any inhibitors like EDTA in your buffer that may interfere with binding. I would try about 1 column volume of a low concentration of protein to see if that all binds first. If it comes right through the flow through, this would mean that the conformation of the protein is preventing binding, ie the His tag is masked, and that you did not overload your column. I had this problem with one of my proteins. The only way to overcome this is to unfold your protein using urea or guanidinium HCl, and then using the Ni NTA resin. Then you will have to purify it and refold it. I wouldn't recommend this unless the protein already is in inclusion bodies. Rather, I would move the His tag, or use standard chromatography like ion exchange, etc.
Thank you all for your answers.
First, I am purifying a protein that is mainly localized in inclusion bodies. Therefore, I am using a solubilization buffer composed of Tris pH 8.0 , NaCl and 8M Urea to solubilize the insoluble pellet (inclusion bodies) O/N. Once that is completed, I spin down the solution and collect the supernatant as my protein. Then using a minipump, I load my protein into my column at a flow rate of 2.0 mL/min. I then re-load the flowthrough once more and proceed to wash the column with solubilization buffer containing 20 mM imidazole to wash away any unbound proteins. After, I refold the protein using a matrix-assisted redox-based refolding protocol, which is basically done by washing the column with decreasing parts of solubilization buffer and increasing parts of refolding buffer. This is followed by eluting the protein with 200 mM imidazole. I confirmed that the protein is largely still found in the flowthrough, by collecting a sample of my flowthrough and it showed around the approx. molecular weight marker on my ladder.
how much beads are you using for purification, it depends on the quantity of your batch how much beads you should use.
As long as you don't have greater than 100mM Tris in your buffer you should be binding, but phosphate buffer would be better. Have you check if your protein is still on the Ni-NTA column? It may have precipitated when you removed the 8M Urea and so won't elute. It could have refolded but formed multimers (dimer, trimer, hexamer, etc...) that are difficult to elute with only 200mM imidazole. Try boiling some of the beads in SDS-PAGE buffer and running on a gel to see if it is still on the beads. Try eluting harder, in 300 mM or 400mM imidazole and soak for 5 minutes before draining the column. If it is precipitating on the column, try eluting the protein in 8M Urea and then refolding away from the column, by dilution into refolding buffer. I have had good success injecting fractions off Ni-NTA (that were still in 8M urea) into arginine refolding buffer. Does your protein have disulfide bonds... if so you need to have a refolding buffer that allows for disulfide reshuffling.
Yes, I believe it does have disulfide bonds. I was speaking with our collaborators on this project and they said to take a sample after the elution step to confirm that the protein is in fact coming off the column. I'm not sure how I feel about boiling the beads but I was thinking higher concentrations of imidazole could work. My collaborator once suggested to even go up to 500 mM but I wasn't convinced at the time. Also, I use 20 mM Tris in my buffer, pH'd at 8.0.
20mM Tris won't hurt binding.... but check your solution is really pH8.0 before adding to column (adjust if you must with NaOH or HCL)... 20mM is not a lot of buffering capacity to clamp the pH.
Barney: Some His-tags may be inaccessible or too weak to bind Ni-NTA. I noticed that you are using 20 mM imidazole in the loading/wash buffer, which is fine for other recombinant proteins but may be high enough to "elute" or prevent your protein from binding! So, remove that imidazole may help your protein to bind.
you can increase the NaCl concentration, 500 mM to even 800 mM. this helps in binding
Rotating in the cold is better. It improves the binding efficiency. And I would use less protein, as stated above to make sure you aren't overloading the resin.
maybe your his-tag gets, upon urea removal and thus maybe refolding, "buried" in the protein structure, thus preventing it from interacting well with the Nickel in the resin?
Try keeping your protein denatured in urea, purify, and then only remove the denaturing agent?
maybe try to equilibrate the column with less imidazole, then make a slow gradient to see at what imidazole concentration your protein elutes. maybe IMAC is not the best technique for your protein?
good luck!
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