Culture of mouse/rat primary neurons

Isolation and Culture of Hippocampal Neurons

Day 1.      Dry coverslip soak in concentrate HNO₃, O/N Day2.

1. Wash coverslip with autoclaved water extensively (2 ltrs of water should be consumed, changing every hour or so for 5 times at least). Small volume and more frequent wash would be more efficient than less frequent but large volume wash.
2. Bake coverslip in oven at 200℃ for 2hrs (this will serve to sterilize coverslip as well as get rid of any trace amount of HNO3). Or wash with EtOH once. Flame coverslips and place them in the dish of interest (i.e. 6-well plate or 12-well plate).

Day 3.

1. Spread freshly prepared poly L-lysine (1 mg/ml in TRIZMA buffer) on the coverslip. 1 ml/well for 6-well plate, 0.5 ml/well for 12-well plate. Make sure PLL is completely covering the surface of coverslip.
2. Leave it on RT or in the incubator (3hrs – O/N).

Day 4.      1-2 hrs before you start dissection

1. Wash coverslip with sterile water 3 times, and then add NM5 media (1ml per well of 6-well plate, 0.5 ml for 12-well). Leave it in the incubator for preconditioning.
2. Transfer dissection medium to dishes and tubes, keep on ice or 4℃.
3. Spray dissection tools with 70% EtOH.
4. Dissection out hippocampus from pups. (During all the steps, tissue should be immersed in CHILLED dissection media).
5. Add 200ul Papain into 2ml of DM for digest.
6. Incubate for 20 min at 37℃.
7. Fire polish 2-3 Pasteur pipettes to narrow down opening at three different levels.
8. Rinse hippocampus with 10 ml of warm NM5, twice. Keep 2 ml in the tube.
9. Triturate with the pipette with widest opening first BE VERY VERY GENTLE (this is the MOST crucial step- it is your work so far and the life off a mother and its pups, so have patience). Once you see hippocampi sheared into softer form then go to next level of pipette and triturate 4-5 times. This will break the hippocampi into small parts. Finally triturate 3-5 times with the finest pipette and it should give you a homogenous suspension.
10. Run through a cellstrainer.
11. Count number of cells.
12. Plate the cells accordingly, 250K-300K/ml/well (6-well plate) or 80K-100K/0.5ml/well (12-well) for live imaging. (After 2-3 hrs you should see most of the neurons adhered to the coverslips and few of them will have neuritis, if you see this you have done well so far.)

Day 5.      Replace the media with NM0.

Then, change the media once a week (change only half of the media, i.e. if you volume is 4 ml then take 2ml out and add 2.5ml to compensate evaporation).

All media for culturing neurons should add 2% B27 before use.

Reagents and solutions

Gluta-MAX: 5 ml/tube, store at -20 (culture room, -20)

Pen.strep: 5 ml/tube, store at -20 (culture room, -20)

B27: 1 ml/tube, store at -20 (culture room, -20) (Gibco 17504-044)

FBS: 10 or 25 ml/tube, store at -20 (culture room, -20)

Papain: 100U/ml in DM (200ul/tube, store at -20) (Worthington, LK003176).

TRIZMA: one pk in 1L ddH2O (filter, store at 4) (power: sigma T-1194)

Poly L-lysine (PLL): 1 mg/ml in TRIZMA buffer (prepare before use, filter) (P2636-1G, Monica’s -20 icebox)

Neurobasal media: 500 ml (Gibco21103, cold room)

Dissection Medium, DM (500 ml)

50 ml 10X HBSS w/o Ca and Mg (Gibco 14185-052) 5ml Pen.strep (Gibco 15140122)

5 ml Na Pyruvate (Gibco 11360070)

10 ml HEPES (Gibco 1530080)

5 ml glucose (sigma: G-8769 2.5M 45%) 425 ml ddH2O

(Filter, store at 4)

NM5 (500 ml)

465 ml Neurobasal media 25 ml FBS

5 ml Gluta-MAX (invitrogen: 35050-061)

5 ml Pen.strep

(Store at 4, add 2% B27 before use)

NM0 (500 ml)

490 ml Neurobasal media 5 ml Gluta-MAX

5 ml Pen.strep

(Store at 4, add 2% B27 before use)

Dissection Tools

Scissors

Tweezer (straight and angle) Portable pipette

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Note:

1. Usually we use tissue from E18 pups for neuron culture, but E19-P0 works too, with a little lower survival rate.

2. This protocol works for both mouse and rat.

3. For cortical neuron culture, plate higher density of neuron, e.g. 500-600K/well (6-well plate) or 160-200K/well
(12-well plate).

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