Derivation of Dopaminergic Neurons (from Human Embryonic Stem Cells)

Gelatin Coating Culture

Vessels 1. Cover the whole surface of each culture vessel with Attachment Factor solution (1 mL for each well of a 6-well plate, 2 mL into each 60-mm dish, or 4 mL into each 100-mm dish) and incubate for 1 hour at room temperature. Wash once with distilled water before plating the MEF.

Note: AF is sterile 1X solution containing 0.1% gelatin, available from Invitrogen.

Thawing MEFs

1. Wearing eye protection and ultra low-temperature cryo-gloves, remove the vials of irradiated MEF from the liquid nitrogen storage tank using metal forceps. Note: Transfer the vials into a container with a small amount of liquid nitrogen if the vials are exposed to ambient temperature for more than 15 seconds between removal and step 3.
2. Briefly roll the vials containing MEF between your hands for about 10–15 seconds to remove frost and swirl them gently in a 37°C water bath. Do not submerge the vials completely.
3. When only a small amount of ice remains in the vials, remove them from the water bath. Spray the outside of the vials with 70% ethanol before placing them in the cell culture hood.
4. Pipet the thawed cells gently into a 15-mL conical tube using a 1-mL pipette.
5. Rinse the cryovial with 1 mL of pre-warmed MEF medium. Transfer the medium to the same 15-mL tube containing the cells.
6. Add 4 mL of pre-warmed MEF medium dropwise to the cells. Gently mix by pipetting up and down. Note: Adding the medium slowly helps cells to avoid osmotic shock.
7. Centrifuge the cells at 200 × g for 5 minutes.
8. Aspirate the supernatant and resuspend the cell pellet in 5 mL of pre-warmed MEF medium.
9. Remove 10 μL of cell suspension and determine the viable cell count using your method of choice.

Note: We recommend using the Countess® Automated Cell Counter for easy and accurate cell counting and viability measurements.

Plating MEFs

1. Centrifuge the MEFs at 200 × g for 5 minutes and aspirate the supernatant.
2. Resuspend the cell pellet in MEF medium to a concentration of 2.5 × 10⁶ cells/mL.
3. Aspirate the Attachment Factor solution from the coated culture vessels and wash the plates once with D-PBS.
4. Add the appropriate amount of MEF medium into each culture vessel (2.5 mL into each well of 6-well plate, 5 mL into each 60-mm dish, or 10 mL into each 100-mm dish).
5. Into each of these culture vessels, add the appropriate amount of MEF suspension (0.1 mL into each well of 6-well plate, 0.2 mL into each 60-mm dish, or 0.6 mL into each 100-mm dish). The recommended plating density for GIBCO® Mouse Embryonic Fibroblasts (Irradiated) is 2.5 × 10⁴ cells/cm².
6. Move the culture vessels in several quick back-and-forth and side-to-side motions to disperse cells across the surface of the wells and dishes. After plating the cells, place the vessels in a 37°C incubator with a humidified atmosphere of 5% CO₂. Use the MEF plates and dishes within 3–4 days of plating.

1. Wearing eye protection and ultra low-temperature cryo-gloves, remove a vial of hESCs from the liquid nitrogen storage tank using metal forceps.
2. Immerse the vial in a 37°C water bath without submerging the cap. Swirl the vial gently.
3. When only a small amount of ice remains in the vial, remove it from the water bath. Spray the outside of the vial with 70% ethanol before placing it in the cell culture hood.
4. Transfer the cells gently into a sterile 15-mL conical tube using a 1-mL pipette. Rinse the vial with 1 mL of pre-warmed hESC medium to collect the remaining cells in the vial and add them dropwise to the cells in the 15-mL conical tube. Note: Adding the medium slowly helps cells to avoid osmotic shock.
5. Add 4 mL of pre-warmed hESC medium dropwise to the cells in the 15-mL conical tube. While adding the medium, gently move the tube back and forth to mix hESCs.
6. Centrifuge the cells for 5 minutes at 200 × g.
7. Aspirate the supernatant and resuspend the cell pellet in 5 mL of pre-warmed hESC medium.
8. Label the culture vessel containing inactivated MEFs with the passage number of the hESCs from the vial, the date and your initials.
9. Aspirate the MEF medium from the culture vessel containing the MEFs and gently add the resuspended hESCs into the vessel.
10. Move the culture vessel in several quick back-and-forth and side-to-side motions to disperse the cells across the surface of the vessel. Place the vessel gently into a 37°C incubator with a humidified atmosphere of 5% CO₂.
11. Replace the spent medium and examine the cells under a microscope daily. If feeding cells in more than one vessel, use a different pipette for each vessel to reduce the risk of contamination. Note:  hESC colonies may not be visible in the first several days.
12. Observe the hESCs every day and passage the cells whenever the colonies are too big or too crowded. The ratio of splitting depends on the total number of hESC in the culture vessel (approximately 1:2 to 1:4 for hESCs at the first time of recovery).

General Guidelines

• In general, split cells when the first of the following events occurs:
  • MEF feeder layer is two weeks old.
  • hESC colonies are becoming too dense or too large.
  • Increased differentiation occurs.
• The split ratio varies, but it is generally between 1:4 and 1:6.
• Occasionally hESCs grow at a different rate, requiring the split ratio to be adjusted. A general rule is to observe the last split ratio and adjust the ratio according to the appearance of the hESC colonies.
• If the cells look healthy and colonies have enough space, split them using the same
• ratio as the previous passage. If the cells are overly dense and crowded, increase the
• split ratio; if the cells are sparse, decrease the ratio.
• Generally, hESCs need to be split every 4–10 days based upon their appearance.

Passaging hESCs

1. Two days prior to passaging your hESC culture, prepare fresh MEF culture vessels.
2. Remove the culture vessel containing hESCs from the incubator. Mark differentiated colonies under a microscope using a microscopy marker and remove them by aspirating with a Pasteur pipette in the culture hood.
3. Add an appropriate amount of pre-warmed hESC medium into each culture vessel (2 mL for each 60-mm dish or 4 mL for each 100-mm dish).
4. Roll the StemPro® EZPassage™ Disposable Stem Cell Passaging Tool across the entire vessel in one direction (left to right). Rotate the culture vessel 90 degrees and roll the tool across the entire dish again.
5. Using a cell scraper, gently detach the cells off the surface of the culture vessel. Gently transfer the cell clumps into a 15- or 50-mL conical tube using a 5-mL pipette.
6. Rinse the culture vessel with an appropriate amount of pre-warmed hESC medium (1 mL for each 60-mm dish or 2 mL for each 100-mm dish) to collect remaining cells.
7. If some cell clumps are too big, pipet the cell solution up and down several times using a 5-mL pipette to break the cell clumps into smaller pieces.
8. Aspirate the MEF medium from each MEF culture vessel and replace it with an appropriate amount of pre-warmed hESC medium (5 mL for each 60-mm dish or 10 mL for each 100-mm dish).
9. Gently shake the conical tube containing the hESCs to distribute the cell clumps evenly and add an appropriate amount of hESC suspension into each MEF culture vessel. Note: The volume of hESC suspension added into each dish depends on the ratio of splitting (see General Guidelines, above).
10. Move the culture vessels in several quick back-and-forth and side-to-side motions to disperse the hESCs across the surface of the vessels. Place the culture vessels gently in a 37°C incubator with a humidified atmosphere of 5% CO₂.
11. Replace the spent medium daily. hESCs need to be split every 4–10 days based upon their appearance.

Making Embryoid Bodies (EBs)

1. Culture the hESCs on MEF feeder cells until they are 90–100% confluent.
2. Roll the StemPro® EZPassage™ Disposable Stem Cell Passaging Tool across the entire vessel in one direction (left to right). Rotate the culture vessel 90 degrees and roll the tool across the entire dish again.
3. Using a cell scraper, gently detach the cells off the surface of the culture vessel. Gently transfer the cell clumps into a 50-mL conical tube using a 5-mL pipette. Note: Do not break the cells clumps into smaller pieces.
4. Add 1 mL of pre-warmed hESC EB medium into each well of 6-well plate, 2 mL into each 60-mm dish, or 3 mL into each 100-mm dish to collect remaining cells and add them to the 50-mL conical tube containing the hESC.
5. Centrifuge the cells for 5 minutes at 200 × g.
6. Aspirate the supernatant from the hESC pellet. Gently re-suspend the pellet with an appropriate amount of EB medium (15 mL for all the cells from one 60-mm dish or 40 mL for all cells from one 100-mm dish).
7. Transfer the cell clumps to an uncoated T-75 flask for a couple of hours. This allows the fibroblasts to differentially attach to the flask.
8. After a few hours, set the T-75 flask down at a tilted angle to allow the EBs to settle in one corner of the flask. Aspirate the EB medium and replace it with 40 mL of fresh EB medium. Transfer the cell clumps to a fresh T-75 flask and incubate them in a 37°C incubator with a humidified atmosphere of 5% CO₂.
9. Feed the EBs with EB medium every day for 4 days. When feeding, set the flask down at a tilted angle so that the EBs settle in one corner of the flask. Aspirate almost all spent EB medium, replace it with pre-warmed EB medium, and return flask to the incubator. Note:   Due to DNA release form dead cells, cell clumps may stick together. In this case, gently pipet the EBs up and down 2–3 times using a 5-mL pipette. This will help you clean the dead cells off the EB surface. If the EBs attach to flask, use a 5-mL pipette to blow the attached EBs off the bottom of the flask.

Differentiating EBs (Rosette Formation) and Midbrain Specification

1. After culturing the EBs in EB medium for 4 days, transfer the EBs from one T-75 flask into a 50-mL centrifuge tube and centrifuge for 3 minutes at 200 × g.
2. Aspirate the EB medium and resuspend the EBs in 10 mL of pre-warmed neural induction medium.
3. Centrifuge the EBs for 3 minutes at 200 × g.
4. Aspirate the supernatant and resuspend the EBs in 40 mL of pre-warmed neural induction medium. Transfer the EBs into a fresh T-75 flask and incubate the EBs in neural induction medium for 2 days in a 37°C incubator with a humidified atmosphere of 5% CO₂. After the EBs float in the neural induction medium for 2 days, they are ready to be differentiated. Note: If the EB attach to the flask, use a 5-mL pipette to blow the attached EBs off the bottom of the flask.
5. Dilute laminin in D-PBS to 20 μg/mL and coat ten 100-mm culture dishes using 2.5–3 mL of laminin for each dish. Incubate the laminin-coated culture dishes in a 37°C incubator for several hours. Note: Laminin may form a gel when thawed too rapidly. To avoid this, thaw slowly in the cold (2°C–8°C). Once thawed, aliquot into polypropylene tubes and store at –5°C to –20°C. Do not freeze and thaw laminin repeatedly.
6. After incubation, aspirate the laminin and add 10 mL of pre-warmed neural induction medium into each 100 mm dish.
7. Transfer the EBs from the T-75 flask into a 50-mL tube and centrifuge for 3 minutes at 200 × g.
8. Aspirate the supernatant and resuspend the EBs in 10 mL of pre-warmed neural induction medium.
9. Gently shake the 50-mL tube containing EBs to distribute the EBs evenly and add 1 mL of EB suspension into each laminin-coated culture dish.
10. Move the culture dishes in several quick back-and-forth and side-to-side motions to disperse the EBs across the surface of the dishes. Place the dishes gently in a 37°C incubator with a humidified atmosphere of 5% CO₂.
11. Feed the EBs every other day with fresh pre-warmed neural induction medium until early rosettes form (approximately 2–3 days).
12. To direct the neural precursors to the midbrain fate, feed the differentiating EBs every other day with neural induction medium containing 100 ng/mL FGF-8b and 200 ng/mL sonic hedgehog (SHH) for 5–6 days. Note: Plate the EBs at a density of 200–250 per one 100-mm dish. Generally, all EBs from hESCs cultured in one 100-mm dish can be plated into eight to ten 100-mm dishes. The variation is from the confluence of hESCs and efficacy of EB formation

Isolating DA Progenitors

1. Label all differentiating colonies containing rosettes using a microscope marker.
2. Using a 200-μL pipette tip pointing to the center of each marked colony, blow off the cells in rosettes.
3. Use a 10-mL pipette to transfer the detached cell clumps into a 50-mL centrifuge tube. Note: You can combine the cell clumps from five 100-mm dishes into one 50-mL tube.
4. Centrifuge the cells for 3 minutes at 200 × g.
5. Aspirate the supernatant and resuspend the cell clumps in 40 mL of neural expansion medium containing 100 ng/mL FGF-8b and 200 ng/mL SHH.
6. Transfer the cell clumps to a T-75 flask and place the flask in a 37°C incubator with a humidified atmosphere of 5% CO₂. The rosettes will roll up to form neurospheres after about 1 day in the incubator.
7. Replace half of the neural expansion medium containing 100 ng/mL FGF-8b and 200 ng/mL SHH with fresh medium every other day.

Note: Contaminating non-neural cells tend to attach to the flask. When changing the medium, set the flask down at a tilted angle to allow the neurospheres to settle in one corner of the flask. Aspirate half of the neural expansion medium and use a 10-mL pipette to transfer the neurospheres with the rest of the spent neural expansion medium to a fresh T-75 flask. Add 20 mL of pre-warmed fresh neural expansion medium to the flask and incubate in a 37°C incubator with a humidified atmosphere of 5% CO₂. You can perform this procedure several times to purify the neural cells.


DA Neuron Differentiation

1. Coat the surface of the culture vessel (with or without cover slips) with poly-L-ornithine working solution at 20 μg/mL in distilled water (14 mL for T-75, 7 mL for T-25, 3.5 mL for 60-mm dish, 2 mL for 35-mm dish) and incubate the vessel overnight at room temperature.
2. Wash the poly-L-ornithine-coated vessel 4 times with distilled water, and then coat it with laminin working solution at 10 μg/mL in D-PBS without calcium or magnesium (14 mL for T-75, 7 mL for T-25, 3.5 mL for 60-mm dish, 2 mL for 35-mm dish). Incubate the culture vessel for 3 hours at 37°C.  Note: You may coat the culture vessels in advance, replace the laminin solution with D-PBS without calcium or magnesium, and store them wrapped tightly in Parafilm for up to 1 week. Make sure that the culture vessels do not dry out.
3. After the neurospheres float in neural expansion medium for 6–8 days, transfer them into a 15-mL tube and centrifuge for 5 minutes at 200 × g.
4. Aspirate the supernatant and incubate the neurospheres in pre-warmed StemPro® Accutase® Cell Dissociation Reagent for 10 minutes at 37°C.
5. Gently pipet the cell clumps up and down to break the larger clumps into a single cell suspension.
6. Centrifuge the cells for 5 minutes at 200 × g and aspirate the supernatant.
7. Resuspend the cells in 10 mL of pre-warmed neural differentiation medium.
8. Repeat steps 6 and 7.
9. Aspirate the laminin from the coated culture vessels and plate the dissociated DA progenitors.
10. Incubate the cells in a 37°C incubator with a humidified atmosphere of 5% CO₂ and replace the spent medium with fresh neural differentiation medium every other day.
11. You can evaluate DA neuron differentiation 3–4 weeks after plating.

Limited Use Label License No. 557:   Protocol for Producing Dopaminergic Neurons (from human embryonic stem cells)

The use of the method described in the protocol ‘Derivation of Dopaminergic Neurons (from human ES cells)’ to produce dopaminergic neuronal cells for in vitro applications including drug discovery and development, toxicology testing and consumer products testing may require a license from GE Healthcare under US 7,763,463.