Have run into a similar problem.
Cleared the background color by running 2M NaOH together with 0.2M EDTA.
Better replace BMT with TCEP (1 mM).
Also keep in mind that adsorption is pH dependent, that is the higher
the pH, the better is adsorption.
Many proteins adsorb irreversibly above pH 7.0.
If you reduce the pH, say to 5.2-5.5, not only you make adsorption less
stronger (hence, column capacity may drop down), but you will at the
same time prevent cysteine oxidation.
You can also increase [imidazole] in the equilibration buffer to reduce
adsorption.
HTH,
Nadir
Pr. Nadir T. Mrabet
Structural& Molecular Biochemistry
Nutrigenex - INSERM U-954
Nancy University, School of Medicine
9, Avenue de la Foret de Haye, BP 184
54505 Vandoeuvre-les-Nancy Cedex
France
Phone: +33 (0)3.83.68.32.73
Fax: +33 (0)3.83.68.32.79
E-mail: Nadir.Mrabet<at> medecine.uhp-nancy.fr
On 13/01/2012 01:42, Michael Thompson wrote:
Katherine,
You are not alone. I have inadvertently destroyed a GE HisTrap column with high
concentrations of proteins that contain many exposed cysteines. In my case the
Co2+ resin turned a very dark purplish-brown and the protein appeared to have
crashed out on the column. I didn't try to strip it, because I figured it was
done for anyway, so I can't tell you any more about the problem. Here's how I
explained it to myself (whether or not this is actually right I'm not 100%
sure, but it makes sense in my head). The columns I was using have a maximum
concentration of 5mM for DTT and 10mM for B-mercaptoethanol. So that seems like
the column can handle 10mM thiol groups. If you have a protein with many
cysteines and it is very highly concentrated (as was the case for me) then you
are adding considerably more thiol groups to the solution. This abundance of
thiols reduces the metal on the column, and disaster ensues. For me, repeating
the same prep with less DTT (3mM vs. 5mM) in the buffer fixed the issue. If you
are concerned about your protein oxidizing at lower concentrations of DTT or
BME, the other alternative is to switch to TCEP. The IMAC columns can tolerate
higher concentrations of TCEP, and it is a far superior reducing agent (more
stable, more reductive, etc.)...but also a lot more expensive (although you can
get away with using much less because it works so much better).
HTH,
Mike
----- Original Message -----
From: "Katherine Sippel"<[email protected]>
To: [email protected]
Sent: Thursday, January 12, 2012 4:01:10 PM GMT -08:00 US/Canada Pacific
Subject: [ccp4bb] Metal won't strip from IMAC
Hi all,
I've run into a bit of a protein purification conundrum and wondered if anyone
had encountered a similar situation. I've exercised all of my google-fu and
can't find anything. It's a fairly straightforward setup; His-tagged protein
and Talon Co2+ resin, load lysate, wash with 5 mM imidazole, elute with 150 mM
imidazole. There is protein in the elution fractions as would be expected. The
strangeness occurs when I try to regenerate the column. Using the standard
protocol of 25 mM MES, 100 mM NaCl pH 5 doesn't change the color of the resin
back to light pink the way it should with a regenerated column. I try stripping
with the suggested 0.2M EDTA, still pink, 0.5M EDTA, still pink, 8 M urea plus
4% CHAPS and then EDTA, still pink, 1 M NaOH then EDTA, still pink. I've
checked the resin using a Western (with a really specific monoclonal Ab) and it
seems that my protein has somehow irreversibly bound to the column and is
preventing the metal from releasing the sepharose. I've even tried competing
the protein off with excess Co2+ and Mg2+ (the endogenous divalent bound
cation).
Clearly the solution is swapping to a Ni column, but this is really bugging me
now. Has anyone run into this problem with IMAC before?
Background: The protein does bind divalent cations (Mg and Mn) with low
affinity (~1 mM) and has a ridiculous number of cysteines (10 in 416 residues
total). There is 1 mM BME and 1 mM MgCl2 in all of the buffers.
Thanks,
Katherine
