RE: Making reluctant antibodies work
In response to several requests for additional information, let me outline
both what I had done previously and what I have tried since.
The antibodies I'm looking at are made against a human interleukin, and I
have several of them. Some are made in mouse (IgG1), some in rat (mostly
IgG2a, but one is an IgG1). I have had great success with the mouse
antibodies using a rather routine IHC procedure: cut frozen sections of
known positive control tissue (mouse brain ectotopically expressing the
human interleukin after an intercranial injection of recombinant
adenovirus). The brains have been snap frozen using in OCT and after
sectioning are fixed for 10 min in acetone. Sections are stored at -80 C
and retrieved as needed. When used for IHC, the sections are brought to 4 C
before refixing for 5 min in acetone. The sections are then rinsed in TBS,
peroxidase and biotin blocked, and then incubated in primary (biotinylated)
antibody for 30 min at room temp. Sections are then incubated in Vector ABC
for 30 min. I have had success with both DAB and Vector NovaRED (I prefer
NovaRED because of possible DAB confusion with lipofuscin in the neurons,
and I believe DAB to be only slightly more sensitive).
I have also gotten good staining using a non-biotin conjugated primary along
with the DAKO ARKit to do the mouse on mouse staining (though no luck with
the Vector MOM Kit, which I believe is a pretty crummy product).
So, since the mouse antibodies were working on mouse tissue, it seemed a
snap to get the rat antibodies to work--not true! For these Abs, I followed
indentical protocol up to the primary incubation (using a non-conjugated
primary). A 30min incubation in a biotinylated anti-rat secondary antibody
was added (and yes, I know it works because I've used it on other
antibodies), followed by the ABC and DAB.
Since this didn't work, I've tried biotinylating the primary antibody to
follow exactly the protocol used for the mouse and haven't gotten any
staining that way either.
I then tried every digestion technique I could think of (pepsin, protenase
K, trypsin, 0.01% SDS) on the theory that it might undo some conformational
change due to fixation. It did chew up the tissue nicely, but failed to
help in staining.
Next I tried incubating the primary before fixation--while this produced
really lovely autolysis, it did not help with the staining.
So now I'm out of ideas. I know that the rat antibodies bind a different
epitope from the mouse Abs, but I'm not sure why that should make such a
difference...like I said in the first e-mail, I know the mAb binds in vitro,
in vivo, in ELISA and now it looks like it might even work in a Western
(naked, no SDS). I feel like I can tell my bosses that this one just isn't
going to work no matter how badly they want it to, but they don't seem to
want to accept that (especially since it works in everything else). So,
does anyone have any other suggestions or ideas about how to explain WHY it
doesn't work so I can move on to something more productive?
Sorry for the long winded message
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