Culturing Primary Cortical Neurons

Coat culture dishes with 0.05mg/mL Poly-D-lysine overnight at 37^oC.

Wash culture dishes with sterile water x2 then leave to dry.

Sedate pregnant mouse with 120mg/kg Euthanyl and confirm sedation prior to proceeding with the next step.

*The neurons must be plated within approximately 4 hours of sedation.

Soak pregnant mouse with 70% ethanol then dissect open the abdominal cavity.

Isolate uterus with embryos and place in a tube of 1X PBS.

In a laminal flow hood, dissect the brains of E14.5-15.5 embryos in Hank's Balanced Salt Solution (HBSS) and isolate each brain in 700uL of HBSS.

Gently pipette up/down x10 to mix cortices then add 20uL trypsin for every embryo and incubate on a rotator at 37^oC for 20 minutes.

Add 300uL Solution A per embryo and carefully pipette up/down x5 then centrifuge at 2500g for 5 minutes at 4^oC.

Solution A: 2.75mL Neurobasal media (unsupplemented) + 150uL trypsin inhibitor + 100uL DNAse1

Remove supernatant and carefully resuspend pellet in 300uL Solution B per embryo by pipetting up/down x10.

Solution B: 2.55mL Neurobasal media (unsupplemented) + 200uL trypsin inhibitor + 250uL DNAse1

Centrifuge at 2500g for 5 minutes at 4^oC then remove supernatant.

Resuspend pellet in 1mL Neurobasal complete per embryo.

Optional trypan blue cell exclusion assay: Mix 100uL trypan blue + 100uL 1X PBS + 20uL resuspended cells. Count with haemocytometer.

Plate neurons on pre-coated dishes at desired confluency.

Change neurobasal medium at 4 days in vitro.

Fix neurons at 7 days in vitro by removing media and replacing with 4% paraformaldehyde for 10 minutes at room temperature.

Wash neurons with 1X PBS x2 to remove traces of paraformaldehyde then store in 1X PBS at 4^oC.