Jason,

Try dialyzing your protein in elution buffer minus the imidazole but with 100mM EDTA.

That worked for me.

Also, try Co instead of Ni or eluting your protein with low pH instead of imidazole.

Rationale behind this is that Ni can stick to the tag upon elution and cause solubility problems when the imidazole is removed.

Jason,

Try dialyzing your protein in elution buffer minus the imidazole but with 100mM EDTA.

That worked for me.

Also, try Co instead of Ni or eluting your protein with low pH instead of imidazole.

Rationale behind this is that Ni can stick to the tag upon elution and cause solubility problems when the imidazole is removed.

Hi Stefan,

Thanks for the suggestions. This problem is driving me mad. My protein has a pI of 8.4 and the reactions I'm trying require a pH of 8.5-8.7. Luckily I found another technique that labels at pH 7-9 so I've tried to put my protein in a buffer around pH 7.5. Unfortunately I can't dialyze out the imidazole without precipitation and having the concentration drop considerably.

Do you think EDTA will interfere with downstream labeling of amines with an activated ester? It would definitely need to be removed prior to labeling with a chelator/radiometal.

Also do you have a buffer recipe for the low-pH elution buffer used on Ni-NTA?

Thanks,

Jason

After dialysis in EDTA you can change the buffer to an EDTA free buffer. The metal will be gone.

A desalting column can speed this up.

For low pH elution, try 50 mM MES pH 6.0 and the other components you want to have in your buffer.

If you have to elute with more than 500mM imidazole you might need to go to even lower pH. One of my proteins comes down at pH 4, which is ridiculous.

Thanks, I'm going to try dialysis with EDTA. Most of my protein comes off in the first three 1 mL elution fractions with 400mM imidazole. If the dialysis w EDTA doesn't work to keep it from precipitating, I will look to lowering the pH. Thanks again for the help.

Phosphate buffer for dialysis as well as relatively high ionic strength (0.1-0.15 M Na/KCl) may turn out helpful. That worked with my proteins.

HI Stefan,

Just wanted to thank you for the advice. Adding EDTA to the dialysis worked to prevent the precipitation. It added an extra dialysis step, but it was worth it!

Thanks,

Jason

Hi Jason,

I am also having the same problem as yours. I am working on Dectin-1, a membrane protein for the structural analysis and dialysis after Ni column purification causes all the protein precipitated. I tried concentrating the protein without dialyzing imidazole and that time it worked somewhat. I am getting around 0.5 mg/ml (around 20micromolar) but I need more concentration. So I am also seeking the answer how to solubilize and make the protein stable in high concentration.

This phenomenon turned out to be the culprit with a similar problem i was having removing imidazole from concentrated protein after elution off a Ni-NTA column. Protein would immediately crash when imidazole was diluted below 350 mM.  I tested a range of pH, salt and glycerol conditions assuming it was a buffering issue but with no luck. Adding EDTA completely rectified the problem and protein was insensitive to removing imidazole (or variation of buffering conditions) once metal was chelated. Thanks Stefan.  

I came across the same problem. I used the Stefan suggestion and it worked perfectly. Thanks for sharing your expertise here. Best wishes

Thank you Stefan. I have no words to explain how happy I was when it worked out. To shorten the protocol, what I did was, mix EDTA directly to elution fraction at concentration of 10-20mM EDTA and dialyse to sufficient amount of intended buffer. 

Hi

I have a similar problem with imidazol in my elution buffer.actually i didn't use imidazol as a stabilizer r !i just used it to purified my His-tagged protein.the concentration of imidazol in elution buffer is high (500mM)and in addition  buffer contain salt stabilizer (NaCl300mM)and glycerol10%. Since Elimination the interference effect of imidazole with fluorescence spectrum ,dialysis of purified protein carried out .but result showed that my protein destabilized without imidazol.now i don't how i can solve this problem???!!!

Zahra,

High Imidazole in elution is common when nickle leaching off the beads. That causes the insolubility because imidazole and his tag are both chelating the nickle and removing the imidazole alone will make your sample insoluble. 

Try dialyzing your sample against 100mM EDTA and 500 mM NaCl buffered at the pH you want using the buffer you think is right. Glycerol shouldn't be a problem here although I wouldn't use it unless you really need it. Change the dialysis buffer at least three times and see if your protein is behaving now.

You can try several batches of buffer in parallel by splitting your sample and trying different buffers, pH, glycerol concentrations, but keep the EDTA at 100 mM. If your protein has an indigenous metal bound to it you probably should reduce the EDTA and try to figure the EDTA concentration that removes the nickle but leaves your indigenous metal in place. 

 Zahra,

another thing to look into, 500mM Imidazole is not very high. If i would use 500 mM on my protein then 95% would still stick on the column. I know 500 mM is what most protocols suggest as elution buffer, but if your protein comes off late and slow you might want to use 1-2 M to see if you can get it all off.

Should one remove 1 M imidazole from the sample after elution from a nickel column? In many cases it is good to have (an) additional purification step(s) following the elution of the sample from a Ni column. Size exclusion chromatography (SEC) is a convenient and effective way to remove the imidazole. If one finds that imidazole is an important variable in maintaining sample homogeneity, one can also simply reduce the concentration of imidazole from 1 M to a more reasonable level (0.02 – 0.2 M) for inclusion of crystallization reagents. In some cases a protein might not crystallize in the presence of imidazole. To remove the imidazole one can try precipitating the protein in ammonium sulfate and resuspending the sample in low salt buffer and purify by ion exchange or SEC. One can also remove imidazole by dialysis. In one instance (personal communication from Artem G. Evdokimov) decent crystals could only be obtained using 0.6 M imidazole/acetate buffer and two imidazole molecules were seen occupying hydrophobic spots on the protein surface. Sometimes the reason Ni column purified proteins precipitate without imidazole is that the Ni ions leak from the Ni column. The presence of imidazole may have been chelating this excess and trace Ni and once removed the remaining Ni Ni may cause the protein with a His tag to aggregate in solution. One may be able to remove this excess Ni using a Ni chelating resin (Hampton Research catalog number HR2-312). Or one can leave the imidazole buffer with the sample in a more reasonable concentration (0.02 – 0.1 M) or try citrate buffer (chelator) or EDTA. If you are working with a metalloprotein and need metals in the sample, using the chelating resin rather than leaving a chelating reagent in the sample is obviously a better choice.

if I want to keep my protein at -20 how much glycerol  dilution to the protein I should add and if I want to cleave the protein of interest after that from this recombinant protein how can I do it? Is anyone can help me in this.

https://www.researchgate.net/publication/309041540_Important_Factors_Influencing_Protein_Crystallization

Article Important Factors Influencing Protein Crystallization

i also hat often problems by using NHS or hydrazide for his-tagged proteins. tiny amounts form the elution still block. best worked to use maleimide to target SH groups, but after modifying lysines with trautsch reagents. this works excellent, u just have to add 5 mM EDTA to block the metals from oxidizing the SH groups...