I am having difficulty getting specific binding with my rat spinal cord immunofluorescence. My protocol is as follows:
1. Remove spinal cord and quick freeze in TissueTek OCT.
2. Cut tissue on cryostat, laying slices directly on gelatin coated slides.
3. Acetone (-20 C) fixation for 10-20 minutes.
4. PBS wash twice, 10 minutes each.
5. Blocking for an hour in blocking buffer (5% FBS, 0.2% Triton X-100, PBS) at RT.
6. Drain blocking buffer and incubate primary antibody mixed in blocking buffer for 2 hours at RT (have also tried overnight at 4 C).
7. Rinse slides in PBS 3 times, 10 minutes each.
8. Incubate secondary antibody and DAPI (0.5 micrograms/mL) in blocking buffer for 1 hour at RT.
9. Rinse slides in PBS 3 times, 10 minutes each.
10. Dehydrate with 50%, 80%, 90%, and 95% alcohol.
11. Mount with Vectashield mounting medium.
I have tried this method with many different primary antibodies, all validated for IF. The secondaries are from Vector Laboratories (Fluorescein and Texas-Red conjugated mouse/rabbit anti-IgG). My DAPI staining is not uniform across my slices; there are areas that you can see the nuclei very well, but others on the same slice are very dark). Any help would be appreciated!