I'm trying to detect a very low abundance 18kDa protein from whole
tissue extracts by western blotting.
Here's my western protocol:
Semi-dry transfer for 20min. (Pre stained marker shows efficient
transfer)
Wash with PW (purified water) for 5min and then with
KPLs 0.2% detector block for 1 hour at RT (as instructed).
Apply the primary at 1:100 dilution in detector block for 1 hout RT.
Then wash with PBS 1% Tween-20 3X15min.
My secondary is an anti-goat AP from southern biotech used at 1:100000
in detector block incubated for 1 hour RT. Then repeat the washes and
wash with assay buffer (100mM Tris 100mM NaCl pH 9.5) 2X5min.
Incubate with ready-to-use CDP star for 5min and expose on film.
The problem is that my negative control (no primary) is showing very
high background that is like reverse image on film even at 1min
exposure. I cannot clarify whether this detector block is any good. I'm
not using BSA since I've read somewhere that BSA is interfering with AP
detection. The dilution of the secondary is low as it is and I'm not
sure whether going at 200000 or 500000 will be any good.
Any ideas or tips will be greatly appreaciated
Thanks
Sotis
