I am trying to stain free-floating brain slices with a FITC secondary antibody. I think its well known that the brain has autofluorescence and I have attempted to limit this with preincubation with sodium borohydride.
I used a serial dilution of the primary antibody (1:50, 1:100, 1:200, 1:400, and 1:500). Several literary sources that cited the secondary antibody in tissue sections had good results with a concentration of 1:100, which seemed high to me.
I am staining for a potassium channel which is a membrane-bound channel. Because the sections were not paraffin-embedded, I did not include an antigen retrieval step. In two other attempts, I did include an antigen retrieval using an antigen retrieval buffer (from sigma, I think) and another attempt with SDS.
I also have attempted blocking with the serum that the secondary produced in (donkey). I used a high concentration (10%) but still had a significant background.
Would blocking in BSA work better? Should I block with donkey serum AND BSA?
What percentage of BSA should I use? I have read several reports using between 3 and 5%.