Good Day.
I have two rather lengthy issues I hope some of you will be able to help me
resolve.
There's been ongoing discussion (5 years) in my department about IHC
antibody controls.
What do you use?
I've been using what seems to be the most logical to me - normal IgG's from
the same host, at the same concentration as the primary antibody.
One researcher here uses normal IgG's, but titer's it to a concentration
that doesn't give background - usually 1ug/ml - regardless of the
concentration of the primary.  I don't see the point of this.   Another
researcher uses just antibody diluents - here we use 0.1% BSA.
I think using IgG's controls for protein binding (non-specificity), and
using only diluents controls for any background staining not due to the
antibodies (endogenous biotin or peroxidase).

My second issue is one regarding an IHC protocol that's giving me specific
false positive staining.  I'm looking at hexon staining in human tumor
xenographs in nude mice.  I have a goat polyclonal and use goat IgG's as a
control - both at 10ug/ml.  I usually get distinct hexon staining in tumor,
but always get some positive cells at the periphery, looks like in adipose
tissue, in both primary and control antibody sections.  I omitted the
antibodies and it looks like the binding is coming from the secondary swine
or rabbit anti-goat.  So i tried a mouse adsorbed secondary, and I tried
adding 2 to 10% mouse serum to the blocking step and to the secondary
antibody dilution.  No consistently helpful results - my false positive (or
background) did not go away.  What should I try next?  The problem isn't a
bad one - it's just bothersome.

Thank you all for your thoughts.
Jen

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