04 September 2019 2 9K Report

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!

More David Christy's questions See All
Similar questions and discussions