Fixation and Sectioning of Mouse Eyes

Anesthetized mice were transcardially perfused with a fixative solution containing 4% paraformaldehyde in 80 mM PIPES (pH 6.8), 5 mM EGTA, and 2 mM MgCl₂. Eyes were enucleated and post-fixed in the same solution for two hours at room temperature. After fixation, dissected eyecups were embedded in 2.5% low-melt agarose (Precisionary) and cut by a Vibratome (VT1200S; Leica) into 100 µm thick slices as described previously^{52 (link)}. Agarose sections were blocked in PBS containing 5% donkey serum and 0.5% Triton X-100 for 1 h at room temperature before staining with primary antibody in blocking buffer overnight at 4 °C. After primary antibody staining, sections were washed three times in PBS and incubated with secondary antibody in blocking buffer overnight at 4 °C. Finally, sections were washed three times in PBS and nuclei were stained with 10 µg/ml Hoechst (H3569; Thermo Fisher Scientific) for 30 min at room temperature. Finally, sections were washed three times in PBS, and mounted onto slides with Immu-Mount (Thermo) and coverslipped. Images were taken with a confocal microscope (Eclipse 90i and A1 confocal scanner; Nikon) with a 60× objective (1.4 NA Plan Apochromat VC; Nikon) using Nikon NIS-Elements software. Image analysis and processing was performed with ImageJ.