Xenopus primary neuron culture preparation

All experiments were approved by the Boston College Institutional Animal Care and Use Committee and performed according to national regulatory standards. Xenopus primary neuron cultures were obtained from embryonic neural tubes. Eggs collected from female X. laevis frogs were fertilised in vitro, dejellied and cultured following standard methods [44]. Embryos were grown to stage 22–24 [45], and neural tubes were dissected as described [46]. Three neural tubes were transferred for 10 minutes to an Eppendorf tube containing 150 μl CMF-MMR (0.1 M NaCl, 2.0 mM KCl, 1.0 mM EDTA, 5.0 mM HEPES, pH 7.4), centrifuged at 1000 g for 5 min, and 150 μl of Steinberg’s solution [58 mM NaCl, 0.67 mM KCl, 0.44 mM Ca(NO₃)₂, 1.3 mM MgSO₄, 4.6 mM Tris, pH 7.8] was added to the supernatant to follow with the tissue dissociation using a fire-polished glass Pasteur pipet. Cells were seeded on 60 mm plates pre-treated with 100 μg/ml poly-L-lysine and 10 μg/ml laminin; after 2 hrs the medium was replaced by plating culture medium (50% Ringer’s, 49% L-15 medium, 1% fetal bovine serum, 25 ng/μl NT3 and BDNF, plus 50 μg/ml penicillin/streptomycin and gentamycin, pH 7.4 and filter-sterilized) and kept for 24 hr before imaging.

The embryos were injected four times in dorsal blastomeres at two-to-four cell stage with 6 ng of the validated XMAP215 morpholino (MO; [47]), 10 ng of the validated tau MO [48], and/or 5 ng of a newly designed splice site MO for EB3 (3´CTCCCAATTGTCACCTACTTTGTCG5´; for verification see S5 Fig), in order to obtain a 50% knockdown of each.