Western Blot Secondary Antibody Diluent - basic question (Jan/17/2011 )
Hi, so I've got a basic question about western blotting. For the most part, I block my western blot in 4% nonfat milk, and then dilute both antibodies in that blocking buffer and have success. I recently purchased a new antibody, however, for which the data sheet says not to use milk as my blocking agent--to use BSA instead. So I tried to perform a blot--blocked with 5% BSA in TBST and then diluted both primary (Rabbit) and secondary (Goat anti-rabbit) antibodies in 1% BSA-TBST (no sodium azide in either dilution)--and I got nothing. I'm pretty confident that the actual blot worked (markers look good), but I am wondering if I was right to dilute my antibodies in 1% BSA...particularly the secondary? What do you dilute your antibodies in in such a situation? Is there anything I need to be particularly careful about?
Hola, I think that the important step is the blocking, so , having your membrane blocked, you can add your antibodies in any solution, PBs, PBS Tw20, PBS Tw20 BSA. If the dilution recommendation says donīt use milk, use BSA.This is , I have read in any time, in the case of any specific antibodies as anti phosphoproteins, for instance, because milk has some of them and you lost antibody activity in the solution.Buena suerte
Diluting primary antibody in 1%BSA is correct, but my lab dilutes the secondary straight in TBST, no BSA at all. However, I would imagine it would work anyways. You may want to try again with no BSA in the secondary dilution, though I would suspect that something else is the true culprit
So after a few more tries with new ECL, new secondary, I did a dot blot and have decided that the primary antibody simply isn't working and got a refund for it. Thanks for the input though!