We have been resolving proteins and blotting to nitrocellulose then cutting
out the band of interest in order to affinity purify polyclonal antibodies.
We generally Western blot 30 gels, each with a 120 mm long well loaded
with 0.1 ug of protein per mm. Our binding buffer is either PBS with 0.3%
Tween 20, 5.0% nonfat dry milk, and protease inhibitors or 0.05M Tris/HCl pH
8.0, 0.5M NaCl, 0.3% NP-40, 5.0% nonfat dry milk, and protease inhibitors. We
have bound our serum at dilutions ranging from 1:5 to 1:50. The only elution
buffer we have tried is 3M MgCl2, 0.03M Hepes/NaOH, 25% ethylene glycol, final
pH about 7.0. Between binding and elution, we wash with PBS and 0.3% Tw or
0.3% NP-40. Following 10 cycles of binding and elution, we desalt our eluted
antibodies using PD10 columns and assay for antibody activity. Sometimes we
have good activity and other times we have virtually no activity. I suspect
that the difficulty is with efficient elution of the bound antibody.
I would be most interested in other people's experiences with this technique,
especially regarding the buffer used for elution.