Optic Nerve Explant Culture Protocol

Optic nerve explant cultures were prepared from mice aged postnatal day (P)7–11, as described previously (Hawkins and Butt 2013 (link)). In brief, optic nerves were carefully dissected and maintained in pre-warmed (37 °C) and pre-gassed (95 % O₂/5 % CO₂) dissecting media, constituted of Dulbecco’s Modified Eagle Medium (DMEM) (Sigma) medium, supplemented with 4 mM l-glutamate, 10 % foetal bovine serum (FBS; Invitrogen) and 0.1 % gentamicin (Invitrogen, Life Technologies Ltd., Paisley). From this point on optic nerves were kept under sterile conditions and cut into 1–2 mm fragments in filter sterilized pre-warmed dissecting media, using a scalpel blade. For further dissociation, optic nerve fragments were triturated and nerve fragments transferred onto laminin (Invitrogen) and poly-l-ysine (Sigma) coated coverslips and incubated in dissecting medium at 37 °C in 95 % O₂/5 % CO₂ overnight to allow the adhesion of the explants. After 24 h, the dissecting media was substituted with a low serum (0.5 %) modified Bottenstein and Sato (B&S) culture medium, supplemented with 10 ng/ml recombinant human PDGF-AA (R&D Systems) and 0.1 % gentamicin. After 3–4 days in vitro (DIV) the medium was replaced to B&S media supplemented with 0.5 mM dibutyryl cAMP for up to 10 DIV. Explanted cells were then cultured in B&S media with 0.1 % gentamicin for up to 15 DIV, changing media every 3–5 days.