Retinas from 5-, 8-, and 9-day-old chick embryos (embryonic day [E]5, E8, and E9) were used. For dense, stationary, and aggregate cultures, retinas were digested with trypsin (0.05% in calcium-magnesium free [CMF] solution) and after a brief centrifugation the retinas were dissociated in DMEM-5% FBS by mechanical aspiration with a large-bore pipette. For stationary cultures, the dissociated cells were plated in 24-well culture dishes and maintained in an incubator at 37°C in an atmosphere of 95% air and 5% CO2. When dense, stationary cultures were prepared for immunostaining, the cells were plated on poly-l-lysine–coated coverslips, which were maintained in 24-well plates. The number of cells seeded was 13.5 × 106, 6.3 × 106, and 6.6 × 106 per well in 0.5 mL of DMEM-5% FBS for E5, E8, and E9 retinas, respectively.
For explant cultures, retinas were cut into segments of approximately 1 to 2 mm2 and maintained in DMEM-5% FBS with rotation (80 rpm in a waterbath at 37°C). Usually, three retinas were used from E5, whereas only one each was used from E8 and E9, in 7 mL DMEM-5% FBS. The cultures were maintained in an atmosphere of 95% air and 5% CO2. For aggregate cell cultures, six E9 retinas were dissociated as described and diluted in 20 mL DMEM-5% FBS. The cell suspension was maintained in air-CO2, at 37°C with rotation (80 rpm). The culture medium was changed every 2, 3, or 4 days in stationary, explant, and aggregate cultures, respectively.
The dense cultures were maintained in an incubator until confluence (i.e., 2, 5, and 6 days for E5, E9, and E8, respectively) resulting in E5C2, E9C5, and E8C6 cultures, respectively. The explants followed the same scheme for dense, stationary cultures. The aggregate cultures were maintained for 8 days until further use.