His-Tag purification - contaminated protein

[Elayoe]

Hallo,

I have a problem with the purification of my His-tagged protein:
After the purification I can see additional bands in the gel, so I don't have 100% pure protein. I used to have no problems with that. I learned the purification not where I do it now, but I use the same buffers and do everything exactly the same way.

The additional bands I see are:
1. bands smaller than my protein
2. bands smaller than my protein that I can already see in the lysate
3. bands bigger than my protein I can see in the lysate
4. bands bigger than my protein which are at the very top of the gel (therefore must be very hugh)

1. could be due to degradation but it never happend before... All buffers are cooled in ice before usage so the room temperature shouldn't make a difference.
2. and 3. could be because some interaction or unspecific binding, but why didn't that happen before???
4. could be DNA/RNA????

I need to have pure protein for my assays so I need to remove all these additional bands!

A bit more information about how I do the purification:
I express my protein in E.coli DE3 BL-21 RPL and lyse them with the following buffer:
20 mM Tris; 500 mM NaCL; 10% Glycerol; 20 mM Imidazole; pH 8.0.
And I am also adding 1 mM PMSF, 2 mM mercaptoethanol, 1% Triton, Lysozyme and DNase.
I keep it stirring for 2 h at 4C and after centrifugation load it onto a Ni-NTA-column and wash with cold buffer before I elute it with cold buffer.

As I said, I do the protocol exactly like I learned it but now I get these additional bands and have no idea what I can do...

Hope you can help me. If you need to know anything else, just ask...
Thanks in advance.

[kevin_a]

Nickle columns can bind lots of endogenous protein from bacteria. You could be seeing those.

Are your fractions clear at the end of your wash steps?
If your lysate is too viscous you might not be able to throughly wash the column.

Have you done the Western to determine if any of the bands are degradation products?
Obviously the larger ones are not. If you do have degradation, you might need to use an inhibitor cocktail instead of just PMSF.

You can try a Cobalt column to reduce/elminate some of the endogenous bacterial proteins and improve purity.

You might also use Ion Exchange columns to purify the full length protein if your bands are caused by degradation. Pierce has some nice membrane based spin columns.

[ednot]

Hello

I suggest you to add Imidazole in the Binding Buffer (5 -40mM) or try to increase the ionic strenght in the buffer (>1M Nacl), or try to increase the Tris concentration (NaH2PO4 favor the binding of proteins on imac, but TRIS does no enhance, in contrast avoids the binding).

Other thing that you can try, is to dilute your eluted protein(that contains imidazole) to different amounts of imidazole, and try to re-purify in IMAC.
For example, is your protein is in 500mM Imidazole, dilute 10X son now is on 50mM Imidazole and apply it again on an IMAC and see what you eluted in the second run.

I recommend Ni-NTA than IMAC Sepahrose 6FF, because is more selective on 6his residues, the resin from Amersham Binds more protein but also more contaminants.

And dont forget, to try different metal ions, i use Zinc for more selective Binding, and cupper for a strong binding.

Best Regards

Ed.

[irkendna]

I get these extra bands too. I've read that pre-treating the column with a low-concentration (~5-10 mM) imidazole buffer helps the column to selectively bind your protein. I also suggest step-wise increasing the imidazole concentration while washing the column after the protein has bound (my procedure is to pour through the protein solution first, then wash with a 10 mM, 15 mM, and 20 mM imidazole buffer to wash away non-specifically bound protein, then a final elution with 300 mM imidazole to extract my protein.)

Yours,

D