Differentiation and Characterization of Neural Progenitors and Neurons from Mouse Embryonic Stem Cells

Since their first derivation from the inner cell mass of the developing mouse blastocysts^1,2, mouse embryonic stem cells (mESC) have been used as powerful tools to study stem cell self-renewal and differentiation³. Furthermore, studying mESC differentiation leads to tremendous understanding of molecular mechanisms that may improve efficiency and safety in stem cell-based therapy in treating diseases such as neurodegenerative disorders⁴. Compared to animal models, this in vitro system provides many advantages including simplicity in practice and assessment, low cost in maintaining cell lines in contrast to animals, and relative ease in genetic manipulations. However, the efficiency and quality of differentiated cell types are often affected by different lines of mESCs as well as the differentiation methods^5,6. Also, the traditional assays to evaluate differentiation efficiency rely on qualitative examination of selected marker genes which lack robustness and they therefore fail to grasp global changes in gene expression.

Here we aim to use a battery of assays for systematic assessment of the neuronal differentiation. Using both traditional in vitro analyses on selected markers and RNA-seq, we establish a platform for measurement of the differentiation efficiency as well as the transcriptomic changes during this process. Based on a previously established protocol⁷, we generated embryoid bodies (EBs) through the hanging drop technique, followed by induction using supraphysiologic amount of retinoic acid (RA) to generate neural progenitor cells (NPCs), which were subsequently differentiated to neurons with neural induction medium. To examine the efficiency of the differentiation, in addition to traditional RT-qPCR and immunofluorescence (IF) assays, we performed RNA-seq and flow cytometry. These analyses provide comprehensive measurement of the progression of the stage-specific differentiation.

1. mESC culture

1. Coat a 10 cm tissue-culture-treated plate with 0.1% gelatin and allow the gelatin to set for at least 15–30 min before aspirating it out.
2. Seed γ-irradiated mouse embryonic fibroblasts (MEFs) one day before culturing the mESCs in the pre-warmed mESC medium (Dulbecco’s modified Eagle medium (DMEM) with 15% fetal bovine serum (FBS), non-essential amino acids, β-mercaptoethanol, L-glutamine, penicillin/streptomycin, sodium pyruvate, LIF, PD0325901 (PD), and Chir99021 (CH)).
3. Allow the γ-irradiated MEFs to settle and attach to the plate surface before culturing E14 cells.
4. Thaw E14 ESCs in 37 °C water bath and quickly transfer the cells in a 15 cm conical tube with warm mESC medium. Pellet the cells at 200 x g for 3 min and remove the supernatant.
5. Resuspend the cells in 10 mL of mESC medium and plate the cells on the culture plate containing the γ-irradiated MEFs seeded earlier. Incubate the cell culture in a 37 °C incubator under 5% CO₂.
6. For culture passaging, aspirate e the medium and wash the plate with sterile 1x PBS. Add enough 0.05% trypsin to cover the plate surface and incubate at 37 °C for 3 min.
7. Neutralize the trypsin with mESC medium and pipette to generate single-cell suspension. Centrifuge the cells at 200 x g for 3 min and remove the supernatant.
8. Count the cells with a hemocytometer or cell counter and seed about 5.0 x 10⁵ cells in a 10 cm culture plate.
9. Resuspend the cells in 10 mL of mESC medium, plate the cells on the gelatin-coated tissue culture plate and incubate cultures as described earlier.
   NOTE: It is recommended that mESCs be passaged every 2 days to prevent the cells from differentiating in their colonies. Phenol red in the medium functions solely as a pH indicator and depending on the cellular density, it can turn yellowish (more acidic), sooner than 2 days. Hence, it may be necessary to change the medium every day. The γ-irradiated MEFs will eventually die off after a couple of passages.

2. EB, NPC, and neuron differentiation

1. Perform culture passaging protocol mentioned earlier and count the cells (steps 1.7–1.10).
2. Hanging drop method (day 0)
   1. For a 10 cm cell culture plate, count roughly 2.5 x 10⁴ cells where 5.0 x 10² cells will be suspended in 20 µL differentiation medium (DMEM with 15% FBS, non-essential amino acids, β-mercaptoethanol, L-glutamine, penicillin/streptomycin, and sodium pyruvate). Roughly fifty 20 µL droplets containing the cells can be plated on one 10 cm plate.
   2. Aliquot the appropriate number of cells, and then centrifuge the cells at 200 x g for 3 min and remove the supernatant.
   3. Resuspend the cells in the appropriate volume of differentiation medium for a cell density of 5.0 x 10² cells per 20 µL (e.g., 2.5 x 10⁴ cells in 1 mL of differentiation medium).
   4. Using a micropipette or a repeater pipette, place 20 µL droplets of the cell suspension onto the lid of the tissue culture plate. Make sure that the droplets are not too close to one another to prevent them from merging.
      NOTE: The droplets can be plated on either a tissue-culture-treated attachment plate or suspension plate as they will be placed on the lid and not the plate itself. For a more feasible and sterile approach, plate the droplets on an attachment tissue-culture plate and transfer them to a suspension plate as described below.
   5. Fill up the plate with 5–10 mL of 1x PBS and carefully put the lid back on the plate. Incubate the culture in the 37 °C incubator.
      NOTE: PBS is added to the culture plate to prevent the droplets from drying up.
3. On day 2, use a micropipette to collect the droplets from the lid and place them in a 10 cm cell culture suspension plate filled with 10 mL of differentiation medium. Incubate the culture on an orbital shaker shaking at low speed in the incubator.
4. On day 4, to harvest the EBs, collect the cells, centrifuge at 200 x g for 3 min, and remove the supernatant.
   NOTE: The EBs can also be washed with 1x PBS as per the requirements of subsequent experiments.
5. To continue the procedure and induce the EB differentiation into neural progenitor cells (NPCs), prepare the differentiation medium with 5 µM retinoic acid (RA).
6. Remove the old medium by pelleting the EBs at 100 x g for 3 min or allow the EBs to settle before aspirating out the old medium. Add 10 mL of the differentiation medium containing 5 µM RA to the culture plate.
7. On day 6, replace at least half of the medium with fresh medium containing 5 µM RA by tilting the plate and pipetting out the medium as described above.
   NOTE: It is recommended that at least half of the medium be replaced with fresh RA-containing medium on days 5 and 7. Take note of the phenol red indicator; if it turns yellowish, it is best to replace all of the medium.
8. On day 8, harvest the NPCs by collecting the cells, centrifuging at 200 x g for 3 min, and removing the supernatant.
   NOTE: The NPCs can also be washed with 1x PBS according to the needs of subsequent experiments. If needed, NPCs can be frozen down and thawed again for later culture and analysis. If the NPCs are to be cultured, accutase can also be used as an alternative to trypsin.
9. To continue the procedure and to differentiate NPCs into neurons, collect NPCs in a 15 mL conical tube by centrifugation, dissociate them with trypsin and incubate them at 37 °C for 3 min. Pipette the NPCs to ensure that all NPC aggregates are dissociated and neutralize the trypsin with the medium.
10. Filter the cells with 40 μm nylon cell strainer and count the cells before plating them at a density of 1.5 x 10⁵/cm² in N2 medium (DMEM/F12 medium + 3 mg/mL glucose + 3 mg/mL lipid-rich bovine serum albumin (LBSA) + 1:100 N2 supplement + 10 ng/mL bFGF + 50 U/mL pen/strep + 1 mM L-glutamine) on a tissue-culture-treated plate for subsequent PCR and western blot experiments; or on a tissue culture chamber for immunofluorescence experiments.
11. On day 9, replace the old medium with a fresh N2 medium.
12. On day 10, switch the N2 medium with N2/B27 medium (50% DMEM/F12 and 50% neural basal, 3 mg/mL LBA, 1:200 N2 supplement, 1:100 B27 supplement, 50 U/mL pen/strep, and 1 mM L-glutamine).
13. On days 11–12, harvest the neurons as follows. Wash the cells with 1x PBS, add trypsin, and incubate the culture in the 37 °C incubator for 3 min before neutralizing the trypsin with medium and centrifuge at 200 x g for 3 min.

3. Characterization of mESCs and differentiated cells

1. Alkaline phosphatase (AP) assay
   1. Use a kit to assess alkaline phosphatase activity (see the Table of Materials).
   2. Remove the medium from the culture plate and wash the ESCs with 1x PBS.
   3. Add 1 mL of the fix solution (consists of formaldehyde and methanol) provided with the kit to the plate and incubate it at room temperature for 2–5 min.
      NOTE: Over-incubation in the fix solution can compromise the AP activity.
   4. Remove the fix solution, wash the ESCs with 1x PBS and leave some amount of PBS in the plate.
      NOTE: Keep the ESCs moist in PBS to not compromise the AP activity.
   5. Prepare the AP solution by mixing the A, B, and C substrate solutions at 1:1:1 ratio. Mix A and B solutions first and incubate the mixture at room temperature for 2 min before adding the C solution.
   6. Remove the 1x PBS and add the AP solution prepared earlier.
   7. Incubate the ESCs for about 15 min in the dark by wrapping the culture plate with aluminum foil or performing step 3.5 in a dark room.
   8. Monitor the reaction and remove the reaction solution when the solution turns bright to avoid non-specific staining.
   9. Wash the ESCs twice with 1x PBS.
   10. Prevent the sample from drying by covering the ESCs with 1x PBS or mounting medium.
       NOTE: A red or purple stain will appear for AP expression. The plate can be stored in a 4 °C refrigerator.
2. RT-qPCR
   1. Collect cells at various stages by following steps 1.6–1.7 and 2.4 for ESCs and EBs and NPCs, respectively.
   2. Isolate RNA using RNA, DNA, and protein extraction solution (see Table of Materials).
   3. Generate cDNA with a reverse transcriptase kit (see the Table of Materials) and follow the manufacturer’s manual.
3. Fixation and embedding
   1. Harvest EBs and NPCs as described above (step 2.6) and fix them with 4% paraformaldehyde (PFA) solution in 1x PBS for 30 min at room temperature.
   2. Remove the PFA and wash the sample with 1x PBS for 5 min.
   3. Place the sample in a serial dilution of 1x PBS, 10%-, 20%-, and 30%-sucrose solutions at 25‒28 °C where the sample is transferred to the next solution after 30 min of incubation.
      NOTE: The sample can be stored in the 30% sucrose solution at 4 °C before continuing with the embedding step.
   4. Wet the pipette tip with sucrose solution before placing the sample (without stacking them) at the center of the cryo-mold and pipette out the excess liquid.
      NOTE: Filter paper can also be used to remove the excess solution. Wetting the pipette tip with sucrose solution is important to prevent the EBs and NPCs from sticking to the walls of the tip.
   5. Carefully add optimum cutting temperature (OCT) solution to the mold without resuspending the samples and remove excess bubbles with a pipette.
   6. Place the mold with the sample on a laboratory mixer at low speed to mildly agitate the sample for 15 min. This helps to settle the EBs and NPCs to the bottom if they are resuspended in the OCT solution.
   7. Quickly freeze the sample by placing the mold in liquid nitrogen or on dry ice.
      NOTE: Samples can be stored in a -70 °C freezer before continuing to the next step.
4. Cryosectioning
   1. Set the cryostat to cool down to -20 to -18 °C before transferring the sample to the instrument.
   2. Detach the frozen OCT block from the mold and secure it on the holder with a little OCT solution placed on the surface of the holder.
   3. Align the OCT block so that the EBs and NPCs are closest to the blade to ensure that the sample is not lost during sectioning.
   4. Carefully section 10 μm of the block and pay close attention to the slices that contain the sample.
   5. Quickly place the OCT slice containing the sample onto the tissue-embedding glass slide and allow the OCT slices to air-dry for 1 h at room temperature.
      NOTE: The samples can be stored at -70 °C for later use.
5. Immunofluorescence (IF)
   1. Block OCT sections or culture chambers containing neurons in 10% normal donkey serum/0.1% Triton X-100 in 1x PBS for 1 h at room temperature.
   2. Incubate the samples in primary antibody diluted in 5% normal donkey serum/0.05% Triton X-100 in 1x PBS overnight at 4 °C.
   3. Wash samples in 1x PBS/0.1% Triton X-100 thrice for 5 min each wash.
   4. Incubate the samples with secondary antibody diluted in 5% normal donkey serum/0.05% Triton X-100 in 1x PBS for 1 h at room temperature.
   5. Wash samples in 1x PBS/0.1% Triton X-100 thrice for 5 min each wash then incubate the samples in 1 μg/mL DAPI.
   6. Mount the samples with a coverslip and some mounting medium and allow it to dry.
   7. Observe the samples under a fluorescence microscope.
6. RNA-seq analysis
   1. Collect the cells at various stages and perform RNA extraction (see step 3.2).
   2. Prepare the cDNA libraries, perform deep sequencing and data analysis according to the protocol described in Wang et al.⁸.
   3. Perform the Gene Ontology (GO) analysis using the R package, clusterProfiler.
7. Flow cytometry
   1. Collect ESCs by following the steps 1.6–1.7 and resuspend the cells in medium. Collect the EBs and NPCs by following step 2.4 and resuspend the cells in medium.
   2. Filter the cell suspension using the 40 μm nylon cell strainer into a new 15 mL conical tube.
   3. Measure the GFP signal of the samples using the flow cytometer (performed by the institution’s core facility).

As a representation of our method, we performed an EB, NPC, and neuron differentiation experiment on E14 cells. E14 cells were cultured on γ-irradiated MEFs (Figure 1A) until the γ-irradiated MEF population diluted out. We confirmed the pluripotency of the E14 cells by performing Alkaline Phosphatase (AP) staining (Figure 1B) and later RT-qPCR (see below) for Nanog and Oct4 markers. The γ-irradiated MEF-free E14 cells were then induced for differentiation using the protocol outlined in Figure 2A. Briefly, differentiation media droplets of 20 µL containing 500 cells were seeded on the lid of the culture plate (see protocol section 2 for details). The EBs formed were then collected and placed in suspension in fresh differentiation media. From day 4 to day 8 of differentiation, 5 μM RA was added to the culture plates to induce NPCs. Differentiated EBs showed round shape and their size continued to increase during differentiation (Figure 2B). At day 8, the NPCs were harvested and trypsinized, and then the resulting single cell suspension was plated in a tissue culture chamber in DMEM/F12 medium with N2 supplement and later in B27 supplement. By day 10, NPCs differentiating into neurons appear to have elongated-cell shape (Figure 2B).

To further evaluate our differentiation experiment, we performed immunofluorescence (IF) experiments on E14 NPCs at day 8 and E14 neurons at day 12. We observed positive staining for nestin in NPCs and neurofilament (NF) signal for neurons (Figure 3A). Alternatively, RT-qPCR and RNA-seq confirmed the induction of NPC marker genes and loss of pluripotency genes in NPCs (Figure 3B,D,F). As a quantitative method to test the success of ESC differentiation, we differentiated a mouse ESC line expressing a Sox1 promoter-driven GFP reporter⁹, followed by flow cytometry analysis on ESCs and NPCs. We found that 58.7% of total cells at the NPC stage are GFP-positive while the GFP signal is 0.0% at the ESC stage (Figure 3C). To profile the transcriptomic changes during differentiation, RNA-seq experiments for E14 ESCs, EB day 3, and NPC day 8 were performed and revealed gene clusters associated with the respective stages (Figure 3D). The genes in the RNA-seq heat-map were sorted based on their expression levels to identify differentially expressed genes in the different stages during differentiation. Gene Ontology (GO) analysis for the four gene clusters showed that these clusters correspond to distinct cellular functions or pathways indicating that the three cell stages of mESC neuronal differentiation each have a group of genes that are highly expressed in their respective stage but not others (Figure 3E). For example, genes in Cluster 3 are highly expressed in E14 NPCs compared to other stages and correspond to pathways related to neuronal development. Clusters 1, 2, and 4 do not contain highly expressed genes related to any germ layer lineage specifications but they are related to cellular growth and proliferation. Thus, the RNA-seq and accompanying GO analysis showed that the E14 cells have differentiated into the neuronal lineage by day 8 of differentiation.

Figure 1: E14 ESCs in culture. (A) The light microscope images show E14 cells (black arrow) growing in colonies atop the γ-irradiated MEFs. E14 colonies continue to proliferate as seen in the colony size difference between day 1 and day 3 cultures. (B) Confirmation of the pluripotency of E14 ESCs by the alkaline phosphatase (AP) stain. Purple arrows indicate the mESCs that were positive for AP stain. Please click here to view a larger version of this figure.

Figure 2: E14 differentiation into EBs, NPCs, and neurons. (A) The schematic summarizes the major steps for differentiating E14 cells into EBs, NPCs, and neurons. (B) E14 ESCs cultured in medium without LIF and 2i in a suspension plate form individual spheres of EBs visible at day 2 where they continue to grow and expand in size in subsequent days. RA is added at day 4 of differentiation to induce the differentiation into NPCs. After 4 days of induction, these NPCs are plated for differentiation into neurons, which are shown in the bottom panel. Please click here to view a larger version of this figure.

Figure 3: Characterization of differentiated cells. (A) Immunofluorescence images in the top panel show NPC-containing EBs at day 8 probed for nestin (green) and nuclei (DAPI, blue). The bottom panel shows immunofluorescence images for neurons at day 12 probed for neurofilament (green). The red box in the merged images are zoomed in 3x for better view. (B) RT-qPCR analysis showing the pluripotency markers (Nanog and Oct4) and NPC markers (Pax6, NeuroD1, and Nes) of E14 ESCs and NPCs. Error bars are mean ± SD. (C) E14 cells expressing a Sox1 promoter-driven GFP reporter were differentiated into NPCs. Both the ESCs and NPCs at day 8 were quantified for positive GFP fluorescent signal with flow cytometry. (D) The heatmap shows the z-scores for the determined FPKM of the genes expressed in the ESC, EB, and NPC stages. Four distinct gene clusters were identified signifying groups of genes that are differentially expressed in either the ESC, EB, or NPC stage. (E) GO analysis was performed using the R package, clusterProfiler, on the four clusters identified in the RNA-seq. (F) The graph shows the FPKM values for three other pluripotency markers, Sox2, Klf4, and Myc as well as neural markers, NeuN, Map2, and Tubb3 for E14 cells at the ESC, EB day 3, and NPC day 8 stages. Please click here to view a larger version of this figure.

The method for neural differentiation of mouse embryonic stem cells has been established for decades and researchers have continued to modify the previous protocols or create new ones for various purposes^7,10,11. We utilized a series of assays to comprehensively analyze the efficiency and progress of the differentiation stages of mESCs to neurons, which may be used in analysis of other lineage differentiation of mouse or human ESCs. Furthermore, our approaches have proved to be useful tools to evaluate the impact of specific genes or pathways on neuronal differentiation in vitro⁸.

With our methods, pluripotent un-committed ESCs treated with retinoic acid (RA) commit to the neural lineage with a high efficiency and are further induced to generate neurons⁷. To improve the successful differentiation of ES cells into neural cells and reduce the heterogeneity, it is important to keep the ES cells in an undifferentiated state¹². Non-proliferative MEFs, treated with γ-irradiation or mitomycin C, function to maintain the pluripotency of the ESCs and provide a scaffold for their growth^13,14. To obtain consistent results, we start each differentiation experiment with mESCs cultured on γ-irradiated MEFs. After a few passages, the γ-irradiated MEFs die out and the culture eventually becomes homogenous for mESC cells. Alternatively, the mESCs can be pre-plated on gelatin for about 45 min before γ-irradiated MEFs are seeded to better remove them on the next passage. Leukemia inhibitory factor (LIF) has long been used to maintain the pluripotency state of cultured mouse ES cells by activating the JAK/STAT pathway^15,16,17. More recently, PD0325901 (PD, a MEK inhibitor) and CHIR99021 (CH, a GSK inhibitor) were found to provide additional pluripotency maintenance of the ES cells^3,18. In our protocol, we culture mESCs with these inhibitors together with LIF to maintain high pluripotency of mESCs.

Another critical factor to achieve successful differentiation is the quality of EBs. We perform the differentiation of E14 cells by the hanging drop method, which has been applied by other investigators^5,19,20. With this method, single ES cells are allowed to suspend in the differentiation medium droplet for 2 days where they spontaneously aggregate and form EBs. The resulting EBs are typically more well-defined in terms of their morphology (Figure 2B) compared to the method of suspending isolated mESCs in medium, which results in EB sizes in a much wider range in our experience (data not shown). To prevent EBs from attaching to the plates, it is important to do a low speed rotation starting from day 3 continuing through the differentiation process. The EBs are induced to differentiate into NPCs by treating them with RA. The resulting NPCs from RA treatment at EB day 4 are typically heterogenous for neural cells such as oligodendrocytes and astrocytes²¹. Using RT-qPCR or immunofluorescence, the NPC population can be probed for neuron and other neural lineage markers such as Gfap for astrocytes and Olig2 for oligodendrocytes. To further induce the differentiation of NPCs into neurons, the NPCs are cultured in optimal neuron medium where the most important components are the N2 and B27 supplements. N2 supplement mainly functions to help the NPCs to commit to the neuronal lineage while the B27 supplement functions to maintain the longevity of the neurons.

The samples can be collected across the differentiation period (e.g., ESC stage, EB day 2, EB day 4, NPC day 6, NPC day 8, neurons day 10, and neurons day 12) to track the differentiation process by performing RT-qPCR for pluripotency and ectoderm markers. Comparing the pluripotency markers such as Oct4, Nanog, Sox2, Klf4, and Myc between the different cell stages will verify the pluripotency of the mESCs (Figure 3B,F). To investigate the efficiency of the neural differentiation, markers for the mesoderm layer such as Hand1, Snai1, and Tbxt; and endoderm layer such as Eomes and Gata4 can also be probed for (data not shown). Further verification can be performed with immunofluorescence (IF) probing for NPC or neuronal markers (Figure 3A). However, these methods are not quantitative and biased toward the selected markers. To overcome these limitations, we incorporate flow cytometry and RNA-seq analyses (Figure 3C‒F). The cell line used in the flow cytometry experiment is a Sox1-GFP E14 cell line, which was used specifically in this experiment to assess the quality of the NPC differentiation procedure. Sox1 is one of the earliest specific neuronal marker during neuroectoderm development²² hence making it an excellent marker for NPC lineage. Sox1 can be probed for using RT-qPCR or Western blot to evaluate the NPC population. These analyses are particularly beneficial to investigate the differentiation defect caused by gene manipulation or chemical treatment.

It is important to note that there are a few limitations to our protocol presented here. First of all, we are only presenting the comprehensive analysis for one wild-type mESC cell line. Other ESC lines originating from mice or humans might require changes and further optimization in the protocol to ensure successful and efficient neuron differentiation. Secondly, we present an in vitro neuron differentiation method, which naturally bears its own set of limitations. As mentioned before, EBs are treated with a supraphysiological level of RA to drive them towards the NPC lineage. The resulting NPCs are then placed in neuron-optimum media to mimic the physiological conditions and encourage neuron lineage commitment, growth, and longevity. Here, N2 and B27 supplements are used to culture neurons but other supplements are also available such as NS21²³ for similar purposes, which may alter the success and efficiency of neuron differentiation. These conditions are synthetically reconstituted in the cell culture assays, which may not fully represent physiological conditions. The quality of the EBs, NPCs, and neurons highly depend on the starting mESCs. mESCs that have been passaged for too many times and kept in culture for more than 1 week typically start to lose pluripotency and may not successfully undergo differentiation. Thus, maintaining the mESCs in an optimal condition is key in ensuring that they can effectively differentiate into EBs, NPCs, and neurons. Other neuron culture methods such as 3D models have also been proposed to better mimic physiological conditions^24,25,26 sometimes at the expense of throughput and feasibility^27,28. We believe our protocols are useful to characterize these 3D culture models.