Dear Bhaskar,

In which buffer did you equilibrate the column or device you have used for size exclusion chromatography? When you did not add any Imidazole to these buffers you should already have changed the buffer of your samples by this step.

You can stain a native PAGE with normal Coomassie staining procedure. For an easy visible band on the gel you require about 1µg of protein.

Good Luck,

Manuel

The imidazole will go after dialysis or purification of your proteins.

You can use tris glycine buffer at optimum pH.

The most important think in Native PAGE is use the protein in their native form, Do dialysis at their optimum pH, so the protein can stable. Remove all other chemicals from the protein before running the gel.

If you did dialysis, you would obvious eliminate imidazole. If some stay, it would normally behave as HIS+ in intact protein sample, considering sample buffer for native PAGE is around 6.8. Hence, imidazole would influence less the overall separationon Native PAGE. For optimum separation and preservation of active, run the native-page at low voltage with a cool-buffer solution to avoid overheating. Use normal R250 staining solution and avoid acetic acid greater than 7%.

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.

Dear Bhaskar,

I do agree with Louis that with dialysis membrane filter (centricon tube of 50 KDa cut off) lower size molecules gets eliminated out of the concerned solution if washed many times. as concentrated solution still retains same concentration of imidazole (but increases the concentration of molecules higher than 50 KDa) one should dilute the concentrate of first run with required buffer and re-concentrate it

@ Manuel Twachtmann: I initially used Ni-NTA purification so i had used Imidazole . Protein got eluted at 150mM Imidazole. Then i equilibriated my SEC column with buffer having 20mM imidazole. So in this case my protein samples after SEC would obviously have less Imidazole. I was thinking if i could dilute the concentrate with buffer having no imidazole and run it 3-4 times and then concentrate it instead of dialysis. I think dialysis can sometimes lead to loss of some protein which stick to the membrane. Would it be of any advantage if i avoid dialysis or are both methods suitable?

i recommend to remove imidazole after elution itself by dialysis, may be treat with DTT then only add coomasie stain.Together may not be good. DTT treating time also important, not too long , then you may not see anything sometime

Dear Bhaskar,

In my opinion you are fine with 20mM Imidazole in our samples in order to run them in a Native PAGE. There are protocols which actually use imidazole as buffer system in native PAGE (http://www.nature.com/nprot/journal/v1/n1/full/nprot.2006.62.html).

If I were you then I just would give it a try this time. For future His-tag purification I would do a SEC afterwards without any imidazole and then concentrate the protein for storage at -80°C. Thats how we do it in our lab but take care it is allways depending on your specific protein.

All the best

@manuel twachtmann: Thank you once again for your suggestion. have followed it and found it useful. Regarding protein storage, do you suggest i flash freeze aliquots of my protein and store at -80 or directly store at -80. I do not want to add glycerol as a cryoprotectant as i have to again dialyze to remove the glycerol or do buffer-exchange, which may lead to loss of some protein. I generally keep my protein at 4deg for 1-2 weeks. after that i am still in a fix about what to do?

Great that my suggestions helped you !

Since the proteins we work with are quite stable even at room temperature, we normally just freeze them in aliquots at -80°C.

However, if your protein is not as stable then it might be better to flash freeze in liquid nitrogen before you put them to -80°C.

I have learned that you have to be careful with your choice of buffer . For example phosphate buffers can make a big jump in their pH upon freezing, which then results in degradation of your protein. His-taq purification often uses phosphate buffers! Personally, I store my proteins in Tris Buffer at neutral pH with some salt (50mM NaCl).

Maybe someone else can comment on the buffer choice for long term storage.

Here is some more information:

http://www.embl.de/pepcore/pepcore_services/protein_purification/storage_purified_proteins/

Good Luck,

Manuel

But even Tris buffers specially tris-hcl has a high variability in pH with respect to temperature. Tris and Tris-HCL are they different?

Dear Bhasker,

Tris-Cl (tris base) and tris HCl are two different buffers.

If you have sufficient amount of your protein you can try to freeze it without glycerol and do freez thaw study up to 5 cycle by running the sample on gel. One time experiment will give you the answer.

Buffer to be used will depend on the pI of the protein.

Good luck.

Nice discussion, though old quesion, I wanted to respond to clarify my doubt. Since you've concentrated your protein using Amiron filters with kD cutoff, during the same step, if you solvent-exchange with cold buffer with salt n pH of your choice, will this not remove imidazole from your protein besides concentrating it? Just a hunch.