Epifluorescence microscope

iPSCs (passage 14–20) were cultured on Matrigel-coated coverslips in a 24-well plate. Cells were fixed with 4 % paraformaldehyde (Electron Microscopy Sciences) for 15 min, permeabilized with 0.1 % Triton-X-100 (Sigma) solution for 20 min and blocked with 0.3 % BSA (Sigma) solution for 1 h at room temperature. Cells were then incubated with primary antibodies diluted in 0.3 % BSA solution overnight at 4 °C (Table 2), and appropriate secondary antibodies for 1 h, and counterstained with DAPI for 10 min. Coverslips were mounted onto slides with SlowFade Gold Antifade (ThermoFisher Scientific) and imaged with an epifluorescence microscope (Keyence).

Cells were fixed
after 4 or 24 h of culture by washing twice (70 μL, 5 min) with
warm DPBS and fixing with warm 4% paraformaldehyde for 30 min (50
μL) followed by two washes with DPBS (70 μL, 5 min).
For immunostaining, fixed cells were permeabilized with 0.1% Triton
X100 in DPBS for 15 min at room temperature and washed twice with
DPBS. Samples were blocked with 1% bovine serum albumin (BSA) in DPBS
for 30 min at room temperature. The primary antibody was diluted with
1% BSA solution (see dilutions of each antibody in Supplementary Table 2), and 70 μL was added in each
well. After overnight incubation at 4 °C, samples were washed
three times with DPBS + 0.5% Tween 20 and then phalloidin staining
and the secondary antibody were added (diluted in 1% BSA solution;
see final dilutions of each secondary antibody in Supplementary Table 2). Finally, samples were washed three
times with DPBS + 0.5% Tween 20 and imaged using an epifluorescence
microscope (Keyence).