- Embryonic (E14) rats or embryonic (E13) mice
Media and Reagents
- Water, distilled (Cat. no. 15230-162)
- Hanks’ Balanced Salt Solution (HBSS) without Ca2+ and Mg2+ (Cat. no. 14170-112)
- Dulbecco’s Phosphate-Buffered Saline (D-PBS) without Ca2+ and Mg2+ (Cat. no. 14190)
- D-Glucose (Sigma, Cat. no. G8270)
- Penicillin-Streptomycin (Cat. no. 15070-063)
- Ascorbic Acid (Sigma, Cat. no. A4034)
- StemPro® Accutase® Cell Dissociation Reagent (Cat. no. A11105-01)
- Trypan Blue (Cat. no. 15250-061)
- Natural Mouse Laminin (Cat. no. 23017-015)
- Poly-L-Ornithine (Sigma, Cat. no. P4957)
- L-Glutamine (Cat. no. 25030-081)
- Neurobasal® Medium (Cat. no. 21103-049)
- B-27® Serum-Free Supplement (Cat. no. 17504-044)
- Heat-inactivated Fetal Bovine Serum (FBS) (Cat. no. 10438)
- Microdissecting instruments (sterilized)
- Small dissecting scissors
- Medium dissecting scissors
- Dumont forceps, straight
- Dumont forceps, angled or curved
- Curved microdissecting scissors
- Spatula, Moria perforated spoon with holes (e.g., Moria MC17)
- Dissecting microscope (e.g., Leica MZ6 or Zeiss Stemi 2000)
- Curved scalpel blade (e.g., BD Bard-Parker no. 23 or 24)
Make a 10-mg/mL stock solution of poly-L-ornithine in distilled water. Filter-sterilize using a 0.22-μm filter and store for up to 12 months at −20°C.
Ascorbic Acid Stock Solution
Make a 200-mM stock solution of ascorbic acid in D-PBS. Filter-sterilize using a 0.22‑μm filter. Protect from light and store for up to 12 months at −20°C.
For 100 mL of dissection buffer, aseptically mix the following components. The buffer can be stored at 4°C for 1 week. Add ascorbic acid solution before use.
|Ascorbic Acid Solution||0.1 mL|
For 100 mL of differentiation medium, aseptically mix the following components. The medium can be stored at 4°C for 1 week and add ascorbic acid solution before use.
|Neurobasal® Medium||98 mL|
|B-27® Supplement||2 mL|
|Ascorbic Acid Solution||0.1 mL|
Matrix for Midbrain Neural Cell Culture
- Prepare a 1:500 dilution of poly-L-ornithine in distilled water for a final concentration of 20 μg/mL.
- Add 2 mL of 20 μg/mL poly-L-ornithine solution to a 35-mm dish (0.5 mL for a 4-well plate or slide, 0.25 mL for 8-well slide).
- Incubate the culture vessel at 37°C in a humidified atmosphere of 5% CO2 for at least 2 hours.
- Rinse the culture vessel once with distilled water.
- Prepare a 1:100 dilution of laminin in distilled water for a final concentration of 10 μg/mL.
- Add 2 mL of 10 μg/mL laminin solution to a 35-mm dish (0.5 mL for a 4-well plate or slide, 0.25 mL for a 8-well slide).
- Incubate the culture vessel at 37°C in a humidified atmosphere of 5% CO2 for at least 2 hours. Store at 4°C until use.
Note: You may coat the plates in advance and store them at 2-8°C, wrapped tightly with Parafilm®, for up to 4 weeks.
Except for the initial steps of collecting the uterine horns, work under sterile conditions in a laminar flow hood, or add antibiotics (penicillin/streptomycin at standard concentrations) to reagents. Perform the steps in a timely manner, and keep the tissue cooled on ice and immersed in ice-cold buffers throughout the procedure.
- Using aseptic technique, collect the uterine horns from a time-pregnant rat (staged to E14 of gestation) or mouse (staged to E13 of gestation).
- Submerge uterine horns in a 100-mm petri dish containing ice-cold, sterile HBSS, and carefully rinse 2–3 times with 15 mL ice-cold, sterile HBSS.
- Transfer the uterine horns to a clean 100-mm petri dish containing dissection buffer.
- Under a dissection microscope placed in a laminar flow hood, dissect each embryo from the uterine sac and remove the amniotic membranes.
- Use a Morian-type perforated spoon to transfer the embryo to a clean sterile petri dish containing ice-cold dissection buffer.
- Confirm the gestational age by measuring and recording the crown rump length of the embryos (10–12 mm for E14 rat or E13 mouse embryos). Exclude any malformed or otherwise damaged embryos.
- Decapitate each fetus using microdissection scissors or a scalpel.
- Hold the tissue with forceps near the forebrain or hindbrain region to avoid damage to the midbrain region of interest. Carefully dissect and remove the overlying scalp tissue to isolate the brain.
- Place the isolated brain in a clean 60-mm petri dish containing dissection buffer on ice.
- Stabilize the brain with forceps near the forebrain or hindbrain regions, carefully remove and discard the fore- and hindbrain regions using a scalpel or microscissors. Make the rostral cut close to the forebrain vesicles and thalamic region, and the caudal cut at the isthmus region.
Dissecting the Ventral Midbrain
- Steady the obtained midbrain tube with forceps exclusively at the posterior midbrain region marked by the convex curvature at the dorsal midline.
- Use small microscissors or the very tip of a curved scalpel blade to gradually dissect open the midbrain tube along the dorsal midline.
- Carefully open the now characteristically butterfly-shaped tissue flap.
- Use forceps to thoroughly remove any remaining overlying meningeal tissue.
- Trim the outermost (most dorsal) areas of the midbrain tube by dissecting away two thirds of the tissue on each side (approximately lateral/posterior to the sulcus limitans as an anatomical landmark).
- Transfer the resulting tissue piece (~0.3 mm × 1.0 mm in dimension) into a conical tube containing cold dissection buffer kept on ice. Use ~0.2 to 0.5 mL of buffer volume for each piece of VM tissue.
- Wash the pieces of VM tissue in cold dissection buffer (e.g., 15 mL of buffer in a 15-mL conical tube) by letting the tissue pieces sink to the bottom of the conical tube. Aspirate the medium, and fill the tube with fresh buffer.
- Aspirate the buffer and add 1 mL StemPro® Accutase® for every 10 pieces of VM tissue. Incubate the tissues 3-15 minutes at 37°C. Observe the digestion process and determine the optimal duration by test dissociation and homogenization. Avoid over-digestion,
- Using fire-polished Pasteur pipets with decreasing diameter, gently dissociate the tissue pieces by pipetting the tissue up and down for a total of ~20 times. Alternatively, you may dissociate and homogenize the tissue by first using a pipettor with a 1000-μL tip, followed with a 200-μL tip. Avoid excessive formation of air bubbles during mechanical dissociation of VM tissue, as it reduces cell viability.
- If large pieces of tissue remain in the solution, selectively homogenize the pieces separately.
- Optional: Pipet the cell suspension through a cell strainer cap or through a 35- to 70-μm mesh. To minimize loss of cells from this filtering step, flush the filter membrane with a small volume of medium after the cell suspension is passed.
- Centrifuge the cell suspension at 4°C for 3–5 minutes at 200 × g. Aspirate the supernatant.
- Resuspend the cells with differentiation medium. Use 200 μL of differentiation medium for every 10 pieces of midbrain originally isolated.
- Using aliquot of the cell suspension, determine the cell concentration and viability by dye exclusion method (Trypan Blue). Use the quantity of live cells counted for calculating the cell concentration. Cell viability needs to be >80%, and should ideally range from 95-100%. DA neurons are among the most fragile cells in the solution. While the cultures will contain neuronal cell types after relatively harsh treatment, the number of DA neurons will be low.
- Keep the cell suspension on ice or at 4°C until use.
Culturing Midbrain Neural Cells
- Aspirate the laminin from the poly-L-ornithine and laminin-coated culture plate and plate midbrain neural cells in differentiation medium at a density of 2 × 105-5 × 105 cells/cm2.
- Culture the cells in an incubator, changing medium every other day.
- Culture the cells for 3-10 days, then check for DA neurons by immunocytochemical staining using antibodies against the neuronal maker β-III-tubulin, and the dopaminergic markers tyrosine hydroxylase (TH)