The present study was performed with the approval of and following the guidelines of the Animal Care and Use Committee of the Institute for Developmental Research, Aichi Developmental Disability Center (#2019-013), and Keio University (A2021-030). Timed pregnant ICR (wild-type) mice were obtained commercially (see Table of Materials). To observe the relationship between migrating cells and blood vessels, Flt1-DsRed mice, in which the endothelial cells express DsRed²², were used. Male Flt1-DsRed mice were crossed with female ICR mice (both mice were at the age of 8-24 weeks). All the animals were housed and manipulated under specific pathogen-free (SPF) conditions until sampling. The reagents and equipment used in this study are listed in the Table of Materials.

1. In utero electroporation

1. Prepare HEPES-buffered saline (HBS) solution by adding an equal volume of autoclaved water to commercially available 2x HBS.
2. There are two options of anesthetics; isoflurane (inhalation) and MMB (injection). In the case of using MMB, prepare it as follows. Add 0.75 mL of medetomidine (1 mg/mL), 2 mL of midazolam (5 mg/mL), and 2.5 mL of butorphanol (5 mg/mL) in 19.75 mL of autoclaved water²³.
   NOTE: This anesthetic solution becomes less active within 2 months. Store at 4 °C with light-shield.
3. Prepare atipamezole solution. Dilute 0.75 mL of atipamezole (5 mg/mL) with 24.25 mL of autoclaved water.
4. Prepare 0.2% (w/v) Fast Green solution in autoclaved water. Sterilize with a filter (pore size 0.22 µm).
5. Prepare plasmid using a conventional plasmid purification kit based on alkaline lysis and column purification²³ according to the manufacturer's instructions (see Table of Materials). Dissolve the purified plasmid from 100 mL bacteria culture in 30-50 µL of HBS to prepare the plasmid stock (3-5 µg/µL) (Supplementary File 1).
   1. Mix and dilute the plasmid stocks with HBS, and add 1/10 volume of 0.2% Fast Green to color the injectant. The components and final concentrations of the plasmids are determined according to their purpose (Table 1).
6. Make the micropipettes by pulling the glass capillaries using a pipette puller.
   NOTE: The tip of the pipette is cut obliquely using fine forceps. The outer diameter of the tip is approximately 70 µm. Mark the micropipettes at 2 µL intervals by sucking 2 µL water.
7. Inject MMB anesthetic at a dose of 10 μL/g body weight intraperitoneally into a pregnant mouse at an appropriate stage (following institutionally approved protocols).
   NOTE: The embryonic stages to be manipulated should be determined according to the purpose. To label future II/III pyramidal neurons, E14.5 is suitable. To label glial progenitors, electroporation must be performed on E15-16. Alternatively, the mice can be anesthetized by inhalation of isoflurane (1%-2%).
8. After the mouse is deeply anesthetized, place it on a warm plate at 38 °C.
   NOTE: Ensure that the mouse does not respond to pinching the tail.
9. After removing fur around the surgical site using depilatory cream, disinfect the surgical area several times in a circular motion with both an iodine-based scrub and 70% alcohol. Make a 2 cm long skin incision along the midline from the level of the 2^nd most posterior pair of nipples anteriorly, followed by a 2 cm midline incision in the abdominal wall along the linea alba (Figure 1A).
   NOTE: In the process of incision in the abdominal wall, lift it slightly with fine forceps so as not to cut the internal organs.
10. Place a sterile plastic sheet on the skin and a piece of sterile tissue paper on it (Figure 1A). Create a hole in the center to expose the surgical area.
    NOTE: The sterile plastic sheet and the sterile tissue paper are placed after making the incision because a wide view field is required to position the incision referring to the position of the nipples.
11. Wet the paper with sterile PBS, carefully observe the uterine horns from the incision, and draw out the uterine horn through the holes of the plastic sheet and paper.
12. Using a micropipette attached to an aspirator tube assembly, inject 1 µL of the plasmid mix into the left or right lateral ventricle through the uterine wall by expiratory pressure (Figure 1A).
    NOTE: Alternatively, plasmid can be injected by using a microinjector.
13. Pinch the head of the embryo with a tweezer-type electrode and apply electronic pulses using an electroporator (35 V for E14-16, 50 V for E17, 50 ms square pulses with 450 ms intervals, 5 times) (Figure 1A).
    NOTE: Do not pinch the head of the embryos strongly, which will damage the embryos. The actual current should be 40-70 mA.
14. Place the uterine horn back into the abdominal cavity.
15. Suture the abdominal wall using a 5-0 silk thread.
16. Apply atipamezole solution at a dose of 10 µL/g body weight intraperitoneally.
17. Suture the skin using a 5-0 silk thread.
    NOTE: The skin can be closed by using an autoclip.
18. Keep the manipulated mice on a warm plate for at least 30 min. The mice will start moving in the cage.
    NOTE: The procedure of in utero electroporation (from steps 1.7 to 1.16) should be done within 30 min. The typical operation time is 20 min.

2. Preparation of slices

NOTE: The freshness of the slices is the key to the success of this experiment. The slices must be prepared (from steps 2.5-2.12) within 2 h.

1. Prepare the Hanks' balanced salt solution with Ca²⁺ and Mg²⁺; HBSS(+). Add 5 mL of CaCl₂ (14 g/L, autoclaved) and 5 mL of MgSO₄.7H₂O (20 g/L, autoclaved) to 500 mL of HBSS(-). Store at 4 °C. Bubble the solution with 95% O₂ + 5% CO₂ gas for 5 min on ice before use.
2. Prepare the culture medium. At least 2.5 mL/dish is needed. Add 2 mL of fetal bovine serum, 25 µL of L-glutamine (200 mM), 200 µL B27 (50x), 50 µL of penicillin + streptomycin (10 K units/mL and 10 mg/mL) to 8 mL of Neurobasal medium.
3. Prepare 3% low melting temperature agarose gel. Pore 50 mL of HBSS(+), then add 1.5 mg of low melting temperature agarose, and autoclave it. Melt the agarose in a microwave oven and keep it at 50 °C in a water bath incubator prior to use.
4. Add 1.9 mL of culture medium to a glass base dish. Put a cell culture insert on it carefully to not trap air bulbs under the filter, then add 500 µL of culture medium onto the filter (Figure 1B). Place it on ice.
5. After anesthetizing the manipulated mice by inhalation of 5% isoflurane, sacrifice them by cervical dislocation (following institutionally approved protocols) at appropriate stages to be observed.
   NOTE: Two days (48 h) and 3 days after electroporation at E14.5 are suitable for observing the multipolar-bipolar transition and locomotion mode, respectively.
6. Take out the embryos and place them in a Petri dish with cold PBS.
7. Select the embryos by the expression of GFP or other fluorescent proteins under a microscope.
8. After decapitation of the embryos, dissect out the brains⁶ under a dissecting microscope and keep them in cold HBSS(+).
9. Pour the melted 3% low melting temperature agarose gel into a cryomold (see Table of Materials), and then put the brains in it and roll them several times. Place it at room temperature until the agarose completely solidifies, and then place it on ice.
10. Slice the brains coronally at the thickness of 350 µm using a vibrating microtome (see Table of Materials).
11. Arrange the slices on the filter (maximum 6 slices per filter).
12. Withdraw the culture medium from the inside and outside of the filter cup (600 µL in total). In this step, the slices are pressed and fixed onto a filter. After 5 min, add 100 µL of culture medium to the inside of the cell culture insert.

3. Time-lapse imaging

1. Keep the culture chamber mounted on the stage of an inverted laser scan microscope at 37 °C using the heater of the chamber or a hood enclosing the microscope at least 30 min before imaging to prevent Z drift. Supply humidified 5% CO₂ + 40% O₂ gas to the culture chamber.
2. Set the culture dish with slices prepared above in the culture chamber.
3. Using a long working distance 20x lens (NA = 0.45), acquire around 10 Z-stack images with 5 µm steps every 15-30 min for 24 h.
   NOTE: The use of a motorized XY stage is highly recommended. This enables us to obtain images from multiple objective fields. Typically, we obtain 10-20 images from different objective fields in one time-lapse experiment.

4. Imaging analysis

1. Convert XYZT images to XYT by the maximum projection of the Z axis using ImageJ or other software.
   NOTE: If the slices slid during the observation, adjust the position of the cells within the images using StackRed, an ImageJ plugin, with its function of Rigid Body transformation (see Table of Materials).
2. Trace the trajectory of GFP- or other fluorescent protein-positive cells either manually or automatically. Manual and automatic tracing can be performed using MTrackJ²⁴ and TrackMate^25,26 (ImageJ plugins), respectively. Automatic tracing using TrackMate is efficient for analyzing many cells in an unbiased manner. However, manual tracing using MTrackJ remains useful for the accurate tracking of individual cells.