If I remember correctly buffers should be pH 8. Did you use low imidazole in the buffers?

Is it some prepackaged column that you are using, or just the agarose slurry loaded in small columns from the kit? Did you wash and equilibrate the agarose before usage?

If you have slurry try adding it to the lysate and shaking for an hour instead of incubating lysate on the column.

Are you sure the protein is not stuck to the column?

Did you use His-tag antibodies for the western? Do you know in fact that this protein is tagged?

Is it 6xHis? Such cases are not frequent but if there's no other explanation it is possible that tag is not exposed in the native conditions due to protein tertiary structure. You can try purifying under denaturing conditions in parallel to see if it is the case.

Hi Anvesh,

I would normally start with 10-20 mM imidazole in the equilibration as well as the washes buffers. The other thing, as Oleksandr suggested, have you use anti-his in your western blot? and also it could be that the His -tag is not exposed in native condition.

Good luck!

Oleksandr Galkin:

The recommended pH from Clontech was pH 7.4  Hence used the buffers 7.4-7.5. Its a prepacked gravity column from clontech. And yes i did the incubation with gentle shaking on an orbital shaker. I did optimize protein expression and check them with the his tag antibodies. I was able to see no to minimal expression in uninduced and good expression in induced. I will try changing my buffers to pH 8 and try again. I will get back to you how it worked. Thank you!

Rajaei Almohammed: 

Thank you for the suggestion. However, all my protein is in the flow thru as it did not bind to the Ni at all. I did see some binding (less than 5%) which got eluted in the wash buffer. However, i will try your suggestion and get back to u. Thank you!

Angelo toto:

I had 0.5M NaCl in my lysis buffer and 0.3M in my wash buffer. May be i might have to increase pH of my lysis buffer to 8.0 to see if it will change the conformation of my protein to make its His tag more accessible to the Ni resin.

Hi there,

There are 2 obvious explanations: the histag has been cleaved (but it wouldn't explain such low affinity for the 5% full length protein) and/or the histag is unaccessible for the immobilized Nickel in the protein structure. To test these hypotheses: run a WB with cell lysate and probe it with anti his antibody(it will tell you if your protein is actually tagged and if it's not then you may try adding protease inhibitors to your cells prior lysis) ) and add some non denaturant concentration of chaotropic agent (preferentially urea) or detergent (Triton X100) to the lysate and to every solution used for affinity chromatography in order to make the tag accessible. An other solution is to construct and express the protein with the tag at the opposite end(hoping for a better behavior toward affinity column).

Hi Dominique,

Thank you for the suggestion! I am actually doing the experiment right now. I will update you with the result tomorrow. Also, the plasmid is a gift from a researcher at NIH. So, i am not sure if i can do the reconstruction. 

Western blotting showed the presence of His tagged protein in Flow thru (90 %) and first Wash (10%). There is no protein in any of the elutes. I am not sure why that is happening. Trying with buffers (pH 8) and NaCl (0.5 M). Did not add any imidazole to my Equilibration buffer.

Note: The wash buffer has 30 mM and Elute has 300 mM imidazole. The experiments are going on now. Will update soon if this has any positive affect. Or else will try purifying using denaturing conditions.

Hello everyone,

The binding worked when i incubated my protein overnight with the Ni-resins on an orbital shaker at 4C. However, i have a lot of non-specific binding (about 30 % of proteins in the lysate bound to the column). I might have to optimize the wash conditions to achieve better elution of the protein. Please let me know if any of you guys have better suggestions which could help speed up my work.