Protein-protein gel shifts

Achim Recktenwald, PhD ARecktenwald at
Tue Aug 17 16:08:09 EST 1999

Larry <lca1 at> wrote in message
news:7paai9$r0u$1 at
> I've been trying (unsuccessfully) to perform a protein-protein
> gel shift to estimate the binding activity of my preps.  In
> short, I've been incubating equimolar amounts at room temp
> for 30 minutes, and then loading onto a native 4-12% Tris-Glycine
> gel.  However, when I silver-stain, I notice immediately that
> the larger protein complex formed forms a tight band, but also
> "smears" above which makes it difficult to quantitate how much
> bound.  Is there anyway to minimize this band-spreading when
> running native gels?  Thanks.

Where does the smearing start?  At the mol. weight of the larger protein or
of the sum of both protein masses?

If it's at the sum of both proteins, then your protein-protein complex is
not tight enough . This can easily happen, if one of the proteins is very
much smaller compared to the other, or if one component has a much higher
charge density than the other.
The complex dissociates during electrophoresis when the electric field
generates a much higher force on one protein than the other.

You have two options:
* you could play around with the pH of your gel, try to find a pH where the
effects are less noticeable.
* the other option is to perform a chemical crosslinking by incubating  your
protein-protein complex in a dilute glutaraldehyde solution ( 0.5 - max. 3%)
before running the gel. If your glutaraldehyde concentration is too low,
you'll see the same result as without. If it's too high, you'll obtain also
dimers of each protein.


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