Developing Neuron Balls Using a Hanging Drop Culture Technique

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. Preparation of neuron balls as hanging drop culture (Days in vitro (DIV) 0-3)

NOTE: The procedures described here for the preparation of neuron ball culture are based on the method previously reported by the Sasaki group with some modifications. We also adopted several procedures from the Banker method for dissociated culture.

1. Confirming the following before starting the dissection
   1. Prepare all the required solutions and sterilize them by autoclave/filtration in advance.
   2. Keep ready all the instruments and materials to be used in each step of this cortical neuron culture.
   3. Spray and wipe the laminar air flow cabinet, dissection table, stage plate of stereomicroscope, scissors, and forceps with 70% ethanol.
2. Euthanize the mouse upon application of CO₂ and dissect the abdomen to obtain E16 embryos.
3. Remove the brains from embryos carefully with the help of fine tips of forceps and transfer them into 60 mm cell culture dishes containing 4 mL of 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid, HEPES Buffered Salt Solution (HBSS).    
   NOTE: The dissection medium HBSS contains 10 mM HEPES (pH 7.4), 140 mM sodium chloride (NaCl), 5.4 mM potassium chloride (KCl), 1.09 mM di-sodium hydrogen phosphate (Na₂HPO₄), 1.1 mM potassium dihydrogen phosphate (KH₂PO₄), 5.6 mM D-glucose, and 5.64 µM phenol red.
4. Remove the scalp, cut the olfactory bulb, separate cortices from each cerebral hemisphere using the fine tips of forceps under a stereomicroscope, and transfer to another 60 mm dishes containing fresh HBSS. Use at least 3-5 embryos for each separate neuron ball culture.
5. Cut the cortices into small pieces with microdissecting spring scissors in a laminar flow cell culture hood.
6. Transfer minced cortices to a 15 mL tube and trypsinize the minced cortices in 4 mL of 0.125% trypsin in HBSS for 4.5 min in a water bath at 37 °C.
   NOTE: This trypsinization time is critical for efficient neuron culture as the increasing time (> 4.5 min) leads to many more dead neurons.
7. Transfer the cell aggregates to a new 15 mL tube containing 10 mL of HBSS by a sterile transfer pipette and incubate at 37 °C for 5 min. Repeat this step one more time.
8. Transfer the cell aggregates to a new 15 mL tube containing 2 mL of Neurobasal media containing GlutaMax, B27 supplement (NGB medium), 0.01% DNase I and 10% horse serum.
9. Triturate the trypsinized cortices by repeatedly pipetting them up and down (3-5 times) using a fire-polished fine glass Pasteur pipette.    
   NOTE: The diameter of a fire-polished fine glass Pasteur pipette is very important as described in the Banker method paper. If the pipette is too narrow to pass cortices, prepare pipettes possessing 2-3 different sizes, and try from a larger pipette.
10. For preparing neuron balls, take the above cell suspension and adjust the cell density to 1 x 10⁶ cells/mL using an NGB medium.
11. Culture the cortical neurons as hanging drops containing 10,000 cells/drop (1 drop is 10 µL) inside the upper lids of 10 cm culture dishes.
12. Add 7 mL of phosphate-buffered saline (PBS) to the bottom part of culture dishes, then keep in an incubator for 3 days at 37 °C with 5% CO₂ under humidified conditions to allow for neuron ball formation.