|From:||"Anna-Karin Robertson" |
I have a very general question about dilution buffers for primary and
secondary antibodies. What is normally used? I have looked at several
protocols and see no clear or common principle.
I am staining, and double staining, on mouse tissues with primary
antibodies, usually rat or hamster, and sometimes these antibodies are
biotinylated. I use normal immunhistochemistry or immunofluorescence. If I
have a goat-anti-rat as a secondary antibody I block with goat serum first,
before the primary antibody iss applied. If I have a avidin-labelled
secondary step I block with BSA before applying the primary antibody.
I know that a lot of people use BSA or dried milk in their dilution buffers,
but if I am using a secondary step being a streptavidin- or
avaidin-conjugated fluorochrome, I realize I must avoid anything containing
biotin, i.e. serum (?), milk, BSA (?) etc in order not to lower or block the
binding. Should avoid this in the dilution of the primary antibody as well?
But if I use only PBS, won't I risk getting I high background? The antibody
diluent from immunohistochemistry from Pharmingen seem to contain only PBS
and sodium azide, though.
Is there a difference between what is applied in immunofluorescence compared
to normal immunohistochemistry?
I realize each specific case must be tested out, but I would be very
grateful for some general advice.
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