I would recommend that you use a higher pH than 7: pH 8 is normal for running nickel affinity columns.

The reason for this is that the nickel affinity column relies on the binding of neutrally charged histidine to the nickel ion (via a lone pair on one of the nitrogens). The pKa of histidine (i.e. the point at which 50% of the histidine is protonated on both nitrogens) is 6.0. Therefore, if you run at pH 7, between 5-10% of histidines will be protonated, and will repel from the positively charged nickel. Result: more of your protein will flow through. At higher pH, this proportion will be reduced, and you get far more specific binding.

It is also worth considering the pH of your buffer at the temperature that you will run the experiment at. Most of us make buffers at room temperature, and purify protein at 4 C. Almost all buffers change pH with temperature, and usually pH increases as the temperature drops (this is especially the case with Tris, a common choice for IMAC buffers). You may well find that, at the temperature that you are running the experiment, your buffer is at a higher pH than you think. Many people who use Tris pH 8.0 (RT), like I do, are really purifying at pH 8.6!

As a result, it might not be so surprising that your protein is pH sensitive if you are working at low temperatures.

Hope this helps....

I think 7.5 pH is best for Ni-NTA interaction with His residues.

Have you checked this pH??

What about pH 7.0??

below pH 7.4 and above 8 the binding reduces dramatically. I purified more than 300 His- tagged proteins up to now.

Thanks Birendra.....

This time I will try with pH 7.0 as my CFE is in pH 7.0...From next time I will take care of your point...

@Birendra Singh: Dear Sir, its quite impressive that you work a lot on proteins. Please explain a little bit more about what happened with cploumn or protein at pH below 7.4 and above pH8 which result into less purification.

@Piyush: If if you just want to do assay with your enzyme then its ok, you can purify your proein at pH 7. it will result into lesser amount but enough.

Simple question is how much protein and at what quality (purity) you really need. Either the binding is reduced or stay same, if you need 1 mg and you get it it is OK.

I was purifying His-tag proteins at pH from 11 to 7 and it just worked. Dymanic binding capacity of most NI-NTA media is ranging from 10-40mg/ml. So, if your you reduced binding even ten times you might get what you want.

I would recommend that you use a higher pH than 7: pH 8 is normal for running nickel affinity columns.

The reason for this is that the nickel affinity column relies on the binding of neutrally charged histidine to the nickel ion (via a lone pair on one of the nitrogens). The pKa of histidine (i.e. the point at which 50% of the histidine is protonated on both nitrogens) is 6.0. Therefore, if you run at pH 7, between 5-10% of histidines will be protonated, and will repel from the positively charged nickel. Result: more of your protein will flow through. At higher pH, this proportion will be reduced, and you get far more specific binding.

It is also worth considering the pH of your buffer at the temperature that you will run the experiment at. Most of us make buffers at room temperature, and purify protein at 4 C. Almost all buffers change pH with temperature, and usually pH increases as the temperature drops (this is especially the case with Tris, a common choice for IMAC buffers). You may well find that, at the temperature that you are running the experiment, your buffer is at a higher pH than you think. Many people who use Tris pH 8.0 (RT), like I do, are really purifying at pH 8.6!

As a result, it might not be so surprising that your protein is pH sensitive if you are working at low temperatures.

Hope this helps....

If your recombinant protein is 6xHis-tagged you can get good results at pH 7

I think Nicolas explained the rational about selection of pH for His residues binding to your protein. The maximal binding is in between 7.5-8. His-tagged proteins will bind in low and high pH as well, however the yield might decrease.

Anything higher than pH 6.5 (pKa of histidine is ~6-6.3) works for the histidine tag binding to nickel. Whether your protein tolerates a wider range of pH is a question that is answered by optimization. We use pH 7.4 for all over our proteins with no problems.

I've used pH 7.4 with no difficulty what so ever to purify tens of milligrams of protein, like others have been pointing out. Since your protein will destabilize after pH 7.5, I recommend using pH 7.4-7.3, and you should be just fine. Remember to have 150 mM NaCl in there, and you can start out at 10 mM imidazole to reduce non specific binding.

Hope it works out brilliantly for you.

From my experience I agree with Michael Sehorn. I remember in an old Qiagen protocol (before 2000, may be), in a denaturing extraction protocol the Ni-NTA resin after protein binding was washed with a solution C , containing 8M urea-tris phosphate, pH 6.5. The protein elution was obtained by a pH gradient with a sol D (pH 5.2) , Sol E (pH 4.5) to finish with a sol F (0.2 M acetic acid, 0.2 NaCl). The last solution then disappeared from any protocol.

Hello All......Thank you so much for the answers and views....

I am completely new to this...

Can you please tell me about the concentration of buffers I can use with this.....????

My CFE is in 0.2M K.phosphate........

For the components of your IMAC buffer, you should use a buffer (phosphate or Tris are commonly used) at 10-50 mM (I use 20 mM, others use different amounts). You need to add some salt (usually NaCl, sometimes KCl). 50-1000 mM is commonly used: most people use either 150 mM (isotonic with the human serum) or 500 mM. The purpose of the salt is to minimise protein-protein interactions, specifically between your protein of interest and weak binders. This improves purity.

Many protocols recommend that you add about 20 mM imidazole to the binding buffer. Again, this often improves purity.

Obviously, if your protein requires some additive (e.g. cofactor for an enzyme), then you should probably keep some in the IMAC buffers.

Two things to remember: firstly, imidazole is a buffer in its own right, and the salt is _very_ basic. It is a good idea to properly buffer an imidazole stock with HCl, or your 20 mM phosphate/250 mM imidazole "pH 7.4" may be more like pH 9-10!

Secondly, the preferred buffer (the chemistry of the buffer can matter as much as the pH), salt concentration, and additives are different for every protein, and if the protein is challenging, you need to determine them individually for your protein. I had a good example where the addition of the right additive at 5 mM turned a protein from precipitating within 5 minutes of elution from IMAC, into a protein that was stable for enough hours to complete the purification.

i have purified protein at pH 7.0, as i have found that our protein/enzyme was active at that pH but not at 8.0.......that gives sufficient amount of protein for further characterization...........

Thank you all for the views...

I have tried 0.2 M buffer, but it didn't worked.

So I lowered down to 0.05M buffer, pH 7.0 and the elution is great.

Although there are some non-specific bands that I am trying to optimize.

Thanks again ...:)

Hi, for my protein (pI~9) I used lysis/binding buffer at pH=5,3 and it worked fine, then I eluted with buffer at pH 7,5 and got pure protein. When I used lysis buffer pH=7,5 I was always getting a lot of contaminating proteins interacting with my protein of interest. Maybe it's worth trying?

Hey,

You mean to say that I have to increase the concentration of my binding buffer.????

No no, I meant that you can try using buffers at different pH, but first check the isoelectric point, otherwise your protein may precipitate. For one protein I used binding buffer at pH 8.8, for other 5.3 and the binding to Ni-NTA was good. What's more, regulating pH may be the easiest step to get rid of contaminating proteins.

Good luck with your protein!