Hello all,
I am interested in immobilizing an IgG polyclonal antibody to a solid phase
for affinity. I've tested the method with a commercially available
hydrazide-functional, crosslinked agarose to be the support. Hydrazide
should bind to aldehydes on periodate-treated glycoproteins, I'm told.
I tried to do that by periodate-treating a non-specific Rabbit IgG (RbIgG)
and exposing it to the gel for coupling, then using a mouse anti
rabbit~peroxidase conjugate as the target molecule, since that is easy to
detect in fractions at low concentrations. I eluted with acetic acid pH
~2.7, and neutralized the acidic fractions with Tris.
My problem emerges when the CarboLink gives me unacceptably high
backgrounds, i.e., exposing it to RbIgG that has not been periodate-treated
still apparently confers significant affinity for the target to the gel.
Gel that has not been exposed to the RbIgG apparently does not bind the
target.
The peroxidase conjugate is detectable in low concentrations, and I haven't
been able to account for all the activity that I measure in the solution
that I load onto the columns. I know its protein concentration is far lower
than it would be in practice, that is, even the molecule I'd be trying to
purify should be more concentrated at that point than is the conjugate I've
been using for practice.
Blocking the gel with BSA before exposing it to the RbIgG didn't seem to
work either (:-P), any thoughts?
