Moesin Immunofluorescence in Brain Tissue

For Moesin immunofluorescence, frozen pieces of brain tissue from temporal cortex were sectioned at -20 ˚C and transferred to microscope slides. Samples were then warmed to room temperature and immediately incubated in 4% PFA at room temperature for 10 minutes. Slides were then rinsed in diH₂O and immersed in sodium citrate buffer (10 mM sodium citrate, 0.05% Tween 20, pH 6.0) and incubated above a 240 W LED light source (HTG Supply, Cat. No. LED-6B240) at 4°C for four hours to reduce lipofuscin autofluorescence. Next, slides were incubated in blocking solution (2% non-fat milk in PBS plus 0.3% TritonX (PBS_Tr)) at 4°C for 30 minutes. Following non-specific blocking, slides were incubated overnight in blocking solution containing primary antibodies. The following day, slides were rinsed three times in PBS_Tr and incubated in blocking solution containing secondary antibodies at room temperature for one hour. Next slides were rinse three times using PBS_Tr, mounted with DAPI containing media, and coverslipped. Brains were visualized by confocal microscopy (Zeiss LSM 780 NLO with Examiner, Zeiss LSM 810 with Airyscan), and ImageJ.¹¹¹ (link) Immunofluorescence was quantified by measuring average Moesin signal intensity within the nucleus of 50 neurons per biological replicate. For each sample, images were converted to 8-bit binary Z-projections using the Max Intensity projection setting and thresholded with the default parameters in ImageJ. Total fluorescence for each biological replicate was calculated by taking the product of the mean gray value and percent area for each of the 50 regions of interest selected and averaged. Antibodies, reagents, concentrations, and sources are listed in Table S11.