strange behavior in WESTERN BLOTTING: need explanation

Bipin K Dalmia bipin at
Thu Sep 2 12:59:09 EST 1993

first some context:

i routinely do western blotting by transfering proteins from a sds-page
gel to nitrocellulose. i have anti-sera against a protein called
glucoamylase. it is very high titre and i use a 1:3000 dilution. 

now, i have reason to believe (based on sequence homology) that my
anti-sera (with the polyclonals for glucoamylase) might cross-react with
another protein. so i tested for this. using a regular dilution (1:3000)
i saw a very faint but distinct and unequivocal band after staining. the
band was at the right place. so i figured i'd use more anti-sera to get 
a better cross-reaction. i increased the amount of anti-sera 20 fold (new
dilution 1:150) and this time i did not see even that faint band. i did
use positive controls so i know that my transfer/staining etc. were okay.

how do i explain this? is there a specific ratio of antigen to antibody
that would give a good reaction? the ratio would definitely matter for
immunoprecipitation but for immunoblotting i would guess that the more
antibodies i use, the stronger the reaction. could there be something in
the anti-sera (from rabbit) that might inhibit the binding? if so, why
did the positive control (glucoamylase) work so well. 

is there any quick way to test (without elisa equipment) how much
antibody to use?

please help :(
bipin k. dalmia               the other night i was lying on my bed, looking
bipin at             up at the beautiful stars, and i said to myself, 
n2.bkd at     'where the F*CK is my ROOF !!'

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