I’m encountering consistent failures in achieving successful DAPI nuclear staining and would appreciate your expertise in troubleshooting this issue.

I am doing a whole mount tissue immunofluorescence. Despite following standard protocols for cell fixation (4% paraformaldehyde) overnight and DAPI staining, I’m observing either weak signals or complete absence of nuclear fluorescence.

• Initial Protocol: Used SouthernBiotech DAPI Fluoromount-G (Cat. #0100-20), which I normally used in paraffin/frozen sections. But no nuclear fluorescence observed in 3D intact tissue imaging.
• Modified Approach:Switched to Sigma-Aldrich DAPI (Cat. #D9542).Prepared working solutions in ddH₂O: 0.5, 1, and 10 μg/mL. Tested immersion durations: 2 hrs vs. 48 hrs. Tissue size: 2×3×3 mm mammary tissue blocks. Imaging: Zeiss confocal (405 nm excitation). No nuclear fluorescence was observed.
• Control Experiment: Successful nuclear staining achieved with Sigma DAPI on frozen sections.

Could anyone experienced with 3D immunofluorescence (IF) staining share protocol recommendations? Are there critical steps I might be missing or optimization strategies I should implement in my workflow?

Thank you for sharing your expertise!