Isolation, Culture and Long-Term Maintenance of Primary Mesencephalic Dopaminergic Neurons From Embryonic Rodent Brains

NOTE: The animals were maintained and handled in compliance with the institutional guidelines and all animal procedures were approved by the Imperial College's Animal Welfare and Ethical Review Body (AWERB) and the Home Office and Harvard University Institutional Animal Care and Use Committee (IACUC), in compliance with federal and state regulations.

1. Reagent and Equipment Setup

1. Thaw, aliquot and store 1 mg/ml Laminin solution at –80 ºC. Dissolve 2 µl in 1 ml DMEM/F12, resulting in a coating concentration of 1-2 µg/cm².
   NOTE: Thaw Laminin slowly on ice and dissolve in cold DMEM/F12 and immediately add it to the plates/coverslips.
2. Dilute 75 ml of 0.01% Poly-L-ornithine in 425 ml PBS and store at 4 ºC.
   NOTE: The shelf life of the working solution is up to one month.
3. Dissolve 25% (wt/vol) BSA in PBS, pH 7.4, and store at 4 ºC for up to a year.
4. Prepare 50% FBS (in HBSS) deactivation media.
5. Prepare complete medium by adding 50 U/ml penicillin and streptomycin, 1x N2 supplement, 5% (vol/vol) FBS, 0.36% D-(+)-Glucose (wt/vol), 0.25% BSA (wt/vol) to DMEM/F12 and store at 4 ºC.
   NOTE: The shelf life is up to 10 days.
6. Place coverslips in boiling 70% (vol/vol) ethanol for at least 30 min to remove any traces of grease and autoclave.
7. Place coverslips in 24-well plates and add 500 µl Poly-L-ornithine solution to each well. Incubate 1 hr under the tissue culture hood at RT and then, wash 3 times with 500 µl water. Add 500 µl laminin solution to each plate and incubate O/N in the humidified tissue culture incubator at 37 ºC.
   NOTE: While no washes are necessary after laminin treatment, washing the poly-L-ornithine less than three times results in death of the cells 24 h after culturing.
8. Polish the tips of glass pipettes over natural gas or torch flame.
   NOTE: After this step the diameter of the tip should be about half of its normal size.
9. Cut the caps of 1.5 ml microcentrifuge tubes, using an autoclaved scissor and autoclave.

2. Dissection of Embryonic Ventral Midbrain

1. Narcotize timed-pregnant (E12.5) dams by CO₂ inhalation and then euthanize by cervical dislocation.
2. Spray the abdomen with 70% ethanol and make an abdominal incision until the uterine sacs are all exposed. Using forceps cut the vaginal attachment and remove uterus. Place the uterus in ice-cold HBSS, in a 100 mm Petri dish.
3. Remove embryos from the uterus and amniotic sac, using forceps (Figure 1C). Place embryos in fresh ice-cold HBSS.
   NOTE: The rectangle in Figure 1D indicates the location of embryonic ventral midbrain.
4. Place the embryos under a dissection microscope (10x magnification) and dissect the mesencephalic arch by cutting the brain at the isthmus and mesencephalic-diencephalic boundary region, using the bioscissor and forceps (Figure 1E, please refer to Figure 1A for lines of incision).
5. After taking out the entire midbrain, make a cut in the mediodorsal midbrain (Figure 1F).
6. Remove meninges, using two forceps (Figure 1H).
   NOTE: CRITICAL STEP: This step is essential for optimal neuronal yield and survival. Since the culture conditions are optimal for brain cells, most of the cells from the surrounding tissue die after plating and the apoptotic signal from this tissue may damage integrity of the cultures.
7. Flatten the midbrain tissue on the Petri dish to observe the butterfly shape and cut approximately half of each wing, using a sterilized shaving blade (Figure 1H,I).
   NOTE: Figure 1J depicts the isolated ventral midbrain.
8. Transfer the piece of ventral midbrain into ice-cold HBSS in a 15 ml conical tube and proceed with the next embryo.
   NOTE: Steps 2.1-2.8 should not take more than 1 hr. Keeping the embryos beyond this time frame on ice decreases cell viability significantly.

3. Dissociation of Ventral Midbrain Cells

1. Transfer the pieces of ventral midbrain under a laminar flow hood. Remove the HBSS and add 1 ml pre-warmed (37 ºC) 0.05% trypsin-EDTA into the conical tube (volumes enough for up to 12 pieces of ventral midbrain). Incubate the tissue at 37 ºC for 5-10 min.
2. Remove trypsin-EDTA under the hood and add 1 ml de-activation medium (the serum will deactivate trypsin) to the tissue.
3. Remove the deactivation medium and wash the tissue in 1 ml complete medium twice.
   NOTE: Avoid removing all the medium from the tube and pipetting the tissue, to prevent discarding the dissociated cells.
4. Add 1ml complete medium and triturate the tissue with a fire-polished glass pipette. Avoid formation of bubbles and continue triturating until single cell suspension is achieved.
   NOTE: Usually, 8-10 passes through the restricted tip are required for complete dissociation.
5. Centrifuge the cells at 400 x g for 5 min at RT and remove the medium.
6. Re-suspend the pellet in 1 ml complete medium by slowly pipetting up and down for up to four times.

4. Plating Ventral Midbrain Cells

1. Mix 10 µl of the cell suspension with 90 µl Trypan blue solution and count the number of cells with a hemocytometer.
   NOTE: Viability of the cells can also be checked at this stage.
2. Place sterilized microcentrifuge tube caps in 100 mm Petri dishes and transfer the Poly-L-ornithine/Laminin coated coverslips, using forceps, one at a time on top of the caps.
   NOTE: Do not wash the coverslips and avoid drying the laminin, since this may cause uneven cultures and decrease in survivability of the cells.
3. Adjust the volume to 1,500 cells per 1 µl by adding complete medium and add 100 µl (total of 150,000 cells) of the cell suspension on top of each coverslip (the volume is optimal to cover the surface of the coverslip without spillage). Close the Petri dishes and incubate them for 1 hr in a humidified tissue culture incubator (37 °C, 5% CO₂).
   NOTE: CRITICAL STEP: This step is required for enhancing the attachment and viability of the cells and for increasing the number of coverslips per embryo. Alternatively, the cells can be transferred directly into the wells (containing coverslips) but the number of cells should be increased to 350,000 per well (instead of 150,000). In other words, using this method, 8-10 coverslips can be acquired, while direct transfer yields about 3-4 coverslips because of the cells, which attach to the plates (beside or underneath the coverslips). The average viability of the cultures, using this method was around 90%.

5. Culture Growth and Maintenance

1. After 1 hr of incubation, carefully transfer the coverslips including the medium into 24-well plates, containing 400 µl pre-warmed (to 37 ºC) complete medium (total volume: 500 µl). Incubate the cells O/N at 37 ºC.
2. Within 24 hr after incubation of the wells, gently add 500 µl complete medium into each well.
   NOTE: The first few hours is the most critical stage for survival of the dopaminergic neurons and the number of surviving cells does not change significantly beyond this point.
3. Do not add additional media to the cultures for the first two weeks or until the medium becomes yellow. If culturing for more than two weeks or medium color changes to yellow, exchange half of the media (500 µl) with fresh complete pre-warmed media (typically every two weeks).
   NOTE: Dopaminergic neurons within the cultures would survive for more than 6 weeks under these conditions.