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Frozen Neuron Thaw and Culture Protocol
for Vials of One Million Cells

Materials required but not supplied (recommended examples)
 a. Substrate: glass coverslips ( # 63-3031, www.carolina.com ); 
     or 24-well plastic (Corning #3526), or BrainBits ( #PDL ).
 b. Medium (Neurobasal, 50x B27, 0.5 mM glutamine or Glutamax, 
     all Invitrogen).
 c. 15 ml sterile polystyrene or PET centrifuge tube (Corning #430055).
 d. 70% ethanol.
 e. Water bath, 37oC .
 f. Hemacytometer (VWR #15170-079), trypan blue (Sigma #T8154) 
    and inverted phase contrast microscope. 
 g. Sterile culture-grade water (e.g. 18 Megohm).
 h. 5% CO2, 37oC humidified incubator (Even better, 5% CO2, 9% O2, 
     eg. Forma 3130).
 i. Sterile hood with vacuum source.
 j. Poly-D-lysine hydrobromide (Sigma P6407) dissolve with sterile 
    18 MΩ H2O to 100µg/ml. Store in PET or polystyrene tube 
    (not polypropylene) at -20oC.  Thawed no more than 2 times.

Preparation of substrates (Neurons are adhesion dependent.  Surfaces must be
coated with adhesive >1-20 hr. before thawing cells. Brainbits offers prepared
substrates, #PDL).
 k. In sterile hood, coat sterile cell culture polystyrene or glass
     with sterile PDL(0.15ml/cm2, 50µg/ml). Let sit for 1 to 20 hr.  Do                           
     not let dry.
 l.  Aspirate with sterile stip.
 m. Rinse one time with sterile water and let substrate dry in sterile
      hood (can be stored at 4oC for a few days, best if used immediately). 
      Alternatively, buy prepared PDL coated substrates from
      BrainBitsLLC (#PDL) .

Preparation of medium (about 0.25 ml/cm2 substrate) (can be stored at 4oC for 1 week)
Growth medium: Neurobasal, 2% B-27, 0.5mM Glutamax or complete NbActiv4. 
Bring to room temperature.

Plating frozen - thawed neurons
1.  Thaw one tube at a time (leave others frozen) for a minimum amount of time in
     37° C  waterbath (target 3 min). Remove immediately when thawed. 
     Do not vortex or shake vigorously.  Wipe outside of cryovial with 70% ethanol.

2.  Transfer 0.21 ml to a 50 ml centrifuge tube without creating bubbles. 

3.  Add 1.91 ml room temperature medium dropwise over a period of 30 seconds.
     Mix gently as you add medium. Gently mix by inversion two times.
 
4. Immediately plate 0.2 ml/cm2 substrate (range 0.15-0.5 ml/cm2) or 2 ml per dish containing           
    3 precoated slips. Gently mix cell suspension every 2 minutes in between aliquots.                
  
5. Incubate plated cells 37°C, 5% CO2,  9% O2, (ambient oxygen will work also) .  Also place            
    more medium in incubator with loose cap to equilibrate pH for step 7.

6. Mix leftover cells from step 4 and aliquot 30 µl of cells into 30 µl of trypan blue. 
    Count cells with hemacytometer (10-10x fields).

7. After 0.7-3 hrs incubation, remove culture from incubator for medium exchange.
    Fill pipet with fresh medium equilibrated in 37°C, 5% CO2 , 0.2 or 0.4 ml fresh
    medium/cm2 (for 35 mm dish use 2.0 mL). Drain and aspirate medium over cells and
    immediately add fresh medium. Be careful not to let cells dry out.  Be careful to add
    medium gently from side so that adherent cells are not disturbed.  Quickly observe
    phase bright adherent cells with 10 or 20x objective mag to ensure attachment to substrate.

8. After 4 days in culture or longer, cells are ready for your use.

9. One-half medium should be exchanged once or twice a week with equilibrated medium.

10. Measure viability at 4 to 5 DIV.  Exchange medium with warm Hibernate E/B27/gln.  Count        
      live, phase-bright cells with processes.  Small, bright cells are dead or apoptosing.

Alternatively a live-dead fluorescence assay can be conducted as follows:
Viability assay: viability=green cells per unit area/(total cells plated per unit area) or
survival (green cells/(green + red cells)

 a. Rinse 2 times with PBS or HBSS, 0.4 ml/2 cm2  of  substrate

 b. From an acetone stock of 15 mg/ml fluorescein diacetate (Sigma #F7378), add 15 μl
     (1:100 dilution of the stock into 1.5 ml HBSS).  From an aqueous stock of 4.6 mg/ml
     propidium iodide (Sigma #P4170), add 15 μl of the stock into the same
     1.5 ml HBSS (1:100 dilution).  Add 40 ul of that dilution to each well with 0.4 ml HBSS 
     (further 1:10 dilution).

 c. After about 1 min., count using Nikon B1A, G1B filters or other blue excitation
     appropriate for fluorescein fluorescence.  Green cells are live.  Small red nuclear 3
     stain indicates a dead cell.

 d. If desired, fix and stain with 0.25% Coomassie blue R in ethanol/HAc/ water (45/10/45),
     1 min., rinse with 10% HAc, aspirate and dry.