Dear Mukul,

It is very common to require an additional purification step after using Ni-NTA. It will do the bulk of the work, but some clean-up (e.g. SEC) is usually required (in my experience).

If you expect higher efficiency perhaps you could try using a greater quantity of Ni-NTA beads. You're using 500uL for 1L of TB culture. I imagine with TB you get a sizeable amount of protein. If you're saturating the beads then some protein will remain unbound.

The non-specific binding may be remedied by using a slightly higher concentration of imidazole in your binding buffer. At the moment you're using 10mM imidazole? With HisTrap columns from GE they recommend somewhere between 20 and 40mM imidazole in the binding buffer. Try that and see if it helps.

Good luck,

James

Hello Mukul,

I see two possibilities :

a) Your protein is truncated and the tag is cliped off by endogenous proteases of E. coli.

b) The structure of the protein is such that the tag is masked

In case b), you may try a different construct, for example putting the tag at the other end of the polypeptide, or including a linker between the tag and the polypeptide.

Concerning the non-specific bands, you can try washing with a somewhat nigher concentration of imidazole, e.g. 15 or 20 mM. You may also try Talon® Co2+ resin, which is somewhat more sepcific than Ni-NTA

Dear Mukul,

It is very common to require an additional purification step after using Ni-NTA. It will do the bulk of the work, but some clean-up (e.g. SEC) is usually required (in my experience).

If you expect higher efficiency perhaps you could try using a greater quantity of Ni-NTA beads. You're using 500uL for 1L of TB culture. I imagine with TB you get a sizeable amount of protein. If you're saturating the beads then some protein will remain unbound.

The non-specific binding may be remedied by using a slightly higher concentration of imidazole in your binding buffer. At the moment you're using 10mM imidazole? With HisTrap columns from GE they recommend somewhere between 20 and 40mM imidazole in the binding buffer. Try that and see if it helps.

Good luck,

James

Dear Mukul,

It is very common, that impurities remain after His-tag purification as these proteins naturally contain several His amino acids on their surfaces. Beside using higher (competing) imidazole concentrations, one might cope with this problem by using an engineered strain (see paper attached to this message). You should always use the minimum amount of resin material to bind your target protein; if I would like to have a really clean target protein I accept to loose some of it, and to overload the resin. Excess resin material will only provide binding capacity to contaminants. I would also assume like Pierre, that the portion of non-binding target protein might never bind as the tag is hidden or truncated or parts of the protein are mis-folded.

Good luck!

Daniel

Article Optimized E. coli expression strain LOBSTR eliminates common...

Hi there,

If your target protein doesn't bind efficiently to the resin (for whatever reason going from low level of expression or low affinity to hidden tag), you automatically tend to purify impurities from the lysate. If the problem is low affinity due to hidden tag then low chaotropic/detergent concentration may be tested to make tag more accessible.

Hi,

The main principle of Ni-NTA purification is the Ni2+ ions have affinity for histidine. Now every protein has certain number of histidine residues(there might be no histidne also), so it is possible that the other proteins are non specifically bound to the Ni resin and hence are eluted with your desired protein. Hence

1. I would suggest you to include 10mM imidazole in your extraction/lysis buffer, then 50mM imidazole in your wash buffer. and you can take 250-300mM imidazole in your elution buffer.  

2. Before loading the sample(protein), wash the resin properly with distilled water. Equilibrate it with wash buffer. if your column is 2ml, wash it 2x5 times. Then load your protein. 

3. Three hours is enough time for protein binding, If you are reusing your Ni beads try with new beads( if purifictaion is done manually). 

4. There are also chances that cells are not efficiantly lysed, or your protein is going into inclusion bodies( during expression) indicating faint desired band. You can go for efficient lysis of cells using either sonication or chaotropes in that case

5. Occurence of non specific bands in final elute is not a problem( happened wth me also), as you can always go for Size exclusion chromatography to get your pure protein, as well rremoval of salts etc.

Hope this helps!

To add to the very good advice above. A trick that is very easy to perform is to add fresh Ni-NTA resin to the unbound fraction and repurify. I have found that sometimes you get great binding in the second purification than the first. This trick has worked for me twice.

Let me known if it works!

The suggestions and observations that you have been receiving are all from well proven practices with IMAC.  Furthermore:  are you running your SDS-PAGE reduced versus non-reduced?  Does your protein have cysteines and you don't want to use DTT because it interferes with the NI?   TCEP is compatible with NI for reduction because phosphines do not use free thiols.  Have you considered a lower pH to protonate the HIs tau nitrogen rather than using Imidazole-only for elution/washing; assuming your protein solution will not precipitate but may remain soluble with some urea? Secondary and teritary amines such as Tris or cytoplasmic metabolites will partially/slowly reduce Ni even at 50 mM.  Have you titrated the sample with Ni resin  to see if you are overloading, under the conditions you use for binding, and getting the benefits of displacement chromatography?  If there is clipping then is the lower mw band the fragment containing the His tag and the upper band a covalent aggregate under non-reducing gel conditions?  To be sure your protein doesn't oxidize in situ in the gel, alkylate the cysteines after reduction then run the gel.  If you are familiar with blot-sequencing then the upper band can be analyzed to see if it is product related or use of Western blots if you have a polyclonal antibody against your protein and against poly His.  If the upper band is product related under reducing gel conditions then there may be a molecular biology issue.  

Almost everything is covered in above responses. I just have one doubt, which may not be such a big issue.

You did not mention what you are using for protease inhibition. If it is PMSF or protease cocktail inhibitor. The stability of PMSF is very limited at pH 8 ( half life of 35 mins at pH 8). Considering that 3 hours binding time seems too much to me. I faced similar issues with my protein where I used to get band of smaller size protein, supposedly degradation product of original one.

About large size impurity. I might be repeating it, but seems to me you need to do washing with higher imidazole concentration (>20mM). How did you come up with 300mM imidazole cocentration for elution? If you protein elutes at such high imidazole concentration you can increase imidazole in washing to much higher levels (although be careful, as prolonged washing may start slow elution of the intended protein).

Finally poor binding of targeted protein might be due to insufficient exposure of His6 tag (as mentioned above). You will need to use some chaotropes or make the tag to C-terminal in this case to check whether binding to resin improves.

Grant Shimamoto is right: use TCEP for preventing Cys oxydation during your quite long incubation.

In addition, you can also try to

- change your metal bound to your beads: Cobalt instead Nickel migth help to remove unspecific bound proteins.

- Add 10-20mM Imidazole in your extraction buffer and to equilibrate the column, and wash in a buffer with 50mM imidazole concentration.

- Make a new construct with a C-term His-tag.