Developing HiPSC Derived Serum Free Embryoid Bodies for the Interrogation of 3-D Stem Cell Cultures Using Physiologically Relevant Assays

1. Generation of Neural Progenitor Cells

1. Maintain hiPSCs derived from fibroblasts and PBMCs in 6-well plates on a γ-irradiated mouse embryonic feeder (MEF) cell layer in human iPSC medium supplemented with small-molecules (see the Materials Table).
   NOTE: Daily maintenance is a modified version of previously reported procedures^1,16.
   1. Plate 6 x 10⁵ MEFs onto each well of a 6-well tissue culture grade plate in 300 μL/well recommended medium (following manufacturer's protocol).
   2. After 48 h, replace MEFs medium with 300 μL/well hiPSC medium for 1 h before adding hiPSCs.
   3. Quickly thaw hiPSCs in a 37 °C water bath and slowly add 1 mL hiPSC medium dropwise. Transfer the hiPSCs and medium to a 15 mL conical tube. Add medium to bring the total volume to 5 mL. Centrifuge for 4 min at 129 x g, aspirate the supernatant, gently re-suspend the pellet in 1 mL hiPSC medium with ROCK inhibitor at 1:1,000 concentration.
   4. Plate the cell suspension (typically seeded at 300,000 cells/well) onto the MEF cell layer and incubate at 37 °C, 5% CO₂, 95% humidity.
   5. Monitor iPSC culture closely using a light microscope. After 48 h, remove cultures that have uneven edges, have grown to become cystic-like or appear yellowish-brown under the light microscope to minimize spontaneous differentiation.
      NOTE: After 7 – 10 days, hiPSC colonies should form. Colonies should be round, with clearly defined borders and uniform cell densities, and devoid of loosely packed non-uniform cells.
   6. Manually select round hiPSC colonies to clear the culture of spontaneously differentiated cells. Expand the colonies by placing them on fresh MEF layers. Expand 1:3 every 7 days when cultures near confluency and freeze^1,16.
2. After expansion and freezing cells (for backup), grow hiPSC colonies on plates until they reach 50 – 75% confluence.
3. Seven-days post plating, use mild enzymatic treatment (5-10 min with 300 μL of enzyme solution) and gentle trituration (2-4 times with a 1,000 µL pipet) to harvest and prepare hiPSCs for neural precursor cell differentiation.
4. Centrifuge the cell suspension at 129 x g for 4 min, aspirate the supernatant, and resuspend the pellet in 5 mL human iPSC medium. Transfer the cell suspension to a 0.1% gelatin coated 5 cm cell culture dish for 1 h at 37 °C to eliminate MEFs and to maximize the hiPSC yield. Transfer the medium (with non-adherent cells) to another 5 cm gelatin coated dish.
5. Transfer the non-adherent cells to a 15 mL centrifuge tube using a transfer pipet. Gently rinse the plates with 3 mL medium and add it to the 15 mL tube.
6. Centrifuge the cell suspension at 129 x g for 4 min, aspirate the supernatant, and gently resuspend the pellet in medium. Determine the cell count using an automated cell counter. Plate 9,000 cells/well in a 96-well low-adhesion V-bottom plate.
7. Centrifuge the plate at 163 x g for 3 min. Incubate the plate at 37 °C, 95% humidity and 5% CO₂. Using a pipette, change 50% medium every other day by removing half of the medium and replacing it with fresh chemically-defined differentiating medium 1 (DM1; Figure 1; Table of Materials) for 14 days.

2. Serum-Free Embryoid Body (SFEB) Induction (Figure 2)

1. Place 40 µm cell culture inserts into 6-well plates and add 1 mL DM1 medium at least 1 h prior to the addition of aggregates.
2. On day 14, transfer the aggregates with 20 µL medium using a 200 µL wide mouth tip onto 40 µm cell culture inserts and change the medium to DM2.
   NOTE: Cell culture inserts are specialized inserts that sit slightly off the bottom of the well and consist of a polytetrafluoroethylene membrane suspended across a plastic frame. This membrane is biocompatible and can efficiently sustain nutrient and oxygen transport to the SFEBs, which are placed on top. They are commonly used with organotypic hippocampal slice cultures taken from mouse or rat^7,17,18.
   1. Transfer 4-6 aggregates to one cell culture insert and allow adequate space between the aggregates. Remove as much excess solution as possible using a fine pipette tip (e.g. 200 μL tip).
      NOTE: It is acceptable to briefly disturb the SFEB as they will move on the cell culture insert as the solution in removed from around the SFEBs.
   2. Maintain the cultures in 1 mL of DM2 at 37 °C, 95% humidity and 5% CO₂. Using a pipette, change 75% medium every other day by removing three quarters of the medium and replacing with three quarters fresh medium. For example, remove 750 μL of medium and add 750 μL of fresh medium.
3. After 14 days of culture, switch to DM3 medium with 75% medium change every other day and for another 16 days.
4. To maintain the SFEBs on cell culture inserts beyond 30 days, change the medium every other day for 60, 90 and 120 days.
   NOTE: The SFEBs will grow to about 1,000 μm in diameter and are typically 100-150 μm thick¹.

3. Determining SFEB Composition

1. Detach the SFEBs from the insert by gently pipetting medium using a 200 µL wide mouth pipette tip. To transfer the SFEBs, use a wide mouth pipette tip to gently suction the bodies into the pipet tip. After loading SFEB into the pipet tip, gently transfer to 12-well plates and wash with 300 μL phosphate buffered saline (PBS) per well.
   NOTE: SFEBs will be suspended in solution in the 12-well plates. This is important to allow full penetration of the fixative, the blocking solution, and antibodies. If using multiple SFEBs for staining, up to 10 SFEBs can be added per well of a 12 well plate. This will conserve solutions and antibodies.
2. Fix SFEBs with 300 μL per well 4% paraformaldehyde at room temperature for 30-45 min. Wash fixed SFEBs twice with 300 μL PBS, 3-5 min each wash.
   Caution: Wear appropriate personal protective equipment when handling paraformaldehyde.
   NOTE: Probe for immunoreactivity to markers to determine maturity, cell type etc. For marking neurons in this study chicken-Tuj1 was used at 1:500, for additional markers please see Table 2 and references^1,16.
3. Permeabilize and block with 0.1% Triton-X100 in PBS and 10% normal donkey serum (blocking solution; 300 μL per well) for 1 h at room temperature.
4. Remove the blocking solution and treat SFEBs with 300 μL primary antibodies in blocking solution. Incubate at 4 °C overnight. Wash SFEBs thrice (3-5 min per wash) in 300 μL PBS containing 0.1% Triton-X.
5. Prepare secondary antibodies 1:1000 in blocking solution. Incubate SFEBs for 1 h at room temperature with secondary antibodies. Wash SFEBs with 300 μL PBS, 3 times for 3-5 min each.
   NOTE: At this stage SFEBs can be prepared for imaging.
6. To image, pipette SFEBs onto a glass slide using a wide-bore pipette. Remove excess solution around the SFEB using gentle suction.
   NOTE: For similar staining conditions, multiple SFEBs can be placed on the same glass slide.
7. Add a drop of mounting medium on the SFEBs, place a glass coverslip, and gently press to ensure that there are no air bubbles. Allow the mounting medium to harden and proceed to imaging and quantification (step 3.8).
   NOTE: After the mounting medium hardens, the SFEBs are ready for imaging.
8. Perform cell quantification by taking multiple regions of interest (ROI) across the SFEB and using a z-stack combined with a maximal projection image through each ROI on a standard confocal microscope as described in references^1,16.
   NOTE: Whole SFEBs can be quantified using image-based tiling (see references^1,16 for details). Since the SFEBs thin to approximately 100 μm, there is minimal scattering of light in the tissue and thus there is no need for 2-photon microscopy. Earlier uses of this protocol with fibroblast-derived iPSCs produced a SFEB fate map that revealed complex and diverse structure with subpopulations of interneurons with transcriptional identities that resembled CGE and anterior forebrain fates^1,16.

4. Recording Neuronal activity in SFEBs Using MEAs

1. Prepare 12-well MEA plates as per manufacturer's instructions.
2. Wash the MEA plates 3 times for 5 min with sterile H₂O under aseptic conditions to clean. Wash for 5 min with 75% ethanol and then with 100% ethanol to sterilize the plate.
3. Bake the plate inverted in an oven for 4-5 h at 50 °C to complete the sterilization process.
4. Add 500 µL 0.2% polyethyleneimine solution (PEI) to each well and incubate 1 h at room temperature. Aspirate PEI and wash the wells 4x with sterile distilled H₂O. Air dry in the hood overnight.
5. Prepare a 20 µL/mL solution of laminin in L15 medium and add 10 µL laminin to the center of each well.
   NOTE: Do not coat the surrounding reference and grounding electrodes.
6. Add sterile dH₂O to the surrounding reservoirs to prevent medium or laminin evaporation. Incubate the plate for 1 h at 37 °C.
7. Add SFEBs (see sections 1 and 2) to the laminin coated MEA plates by gently suctioning them into a wide mouth pipette tip and transfer them. Add 200 µL DM2 medium to each well and incubate overnight at 37 °C, 95% humidity and 5% CO₂.
8. Add an additional 200 µL medium after 24 h and allow it to recover for 30 min at 37 °C, 95% humidity, 5% CO₂ before reading the plate.
9. Place the MEA plate in the plate reader (preheated to 37 °C) to record neuronal activity. Record activity for 10 min with the associated MEA recording software. See reference¹⁹ for details.
10. Generate raw data-continuous streams and raster plots of neuronal activity using statistical analysis software¹⁹.
    NOTE: MEA recordings are taken 7 days post SFEB plating. For this study, recordings were taken for 10 min to detect network burst activity, continuous trace, and raster plots. SFEBs can be maintained long-term on MEA plates and can be recorded over longer periods of time using environmentally controlled conditions (Figure 6).