The Institutional Animal Care and Use Committee approved all animal experiments and the studies were conducted in accordance with relevant animal welfare regulations.

1. Animals preparation

1. Animals
   1. Housing information: House male Sprague-Dawley rats (Postnatal 10-14 days, P10-P14) in a specific pathogen-free environment.
      NOTE: Room conditions were maintained at 20 °C ± 2 °C, humidity: 50%-60%, with a 12-h light/ dark cycle. Animals had free access to food and water.
   2. Label the motor neurons retrogradely: Inject Fluoro-Gold (FG) into the bilateral tibialis anterior and gastrocnemius muscle (2% in sterile saline, 50 µL per muscle) to retrogradely label the motor neurons 2 days before the sacrifice.
2. Solutions
   1. Prepare cutting solution: Mix 120 mM Choline Chloride, 2.6 mM KCl, 26 mM NaHCO₃, 1.25 mM NaH₂PO₄, 0.5 mM CaCl₂, 7 mM MgCl₂, 1.3 mM Ascorbic Acid, 15 mM Glucose. Pre-bubble the solution with 95% O₂ and 5% CO₂ (adjust to pH 7.4 with KOH) for 30 min before the dissection and slicing. Cool the solution with crushed ice.
   2. Prepare artificial cerebrospinal fluid (ACSF): Mix 126 mM NaCl, 3 mM KCl, 1.2 mM NaH₂PO4; 1.3 mM MgCl₂, 2.4 mM CaCl₂, 26 mM NaHCO₃, and 10 mM glucose. Pre-bubble the solution with 95% O₂ and 5% CO₂ for 30 min before the incubation.
   3. Prepare intracellular solution: Mix 126 mM K-Gluconate, 2 mM KCl, 2 mM MgCl₂, 0.2 mM EGTA, 10 mM HEPES, 4 mM Na₂ATP, 0.4 mM Na₂GTP, 10 mM K-Phosphocreatine, and 0.5% Neurobiotin (pH 7.25 and 305 mOsm/Kg). Cool the solution with crushed ice.
   4. Prepare low-melting agarose gel: Dissolve 4 g of agarose in 100 mL of cutting solution, and use a magnetic stirring rotor to fully dissolve it. At 30 min before embedding, heat the low-melting-point agarose in the microwave oven with high power for 1 min, and then transfer it into a 39 °C water bath to maintain the liquid state.
3. Instrument preparation
   1. Place crushed ice on the perfusion tray (Supplementary Figure 1A) at 10 min prior to perfusion. Place the anatomical tray (Supplementary Figure 1B) and cutting tray (Supplementary Figure 1C) with a water band at -80 °C overnight in advance.
   2. Place the incubation chamber with nylon mesh in the oven at 45 °C overnight in advance.
   3. Use low-melting point agarose to prefabricate a 35° slope and a 2 mm thick platform (Supplementary Figure 1D). After gel solidification, place them in the center of a 35-mm Petri dish to support the spinal cord in the next coming procedures.
4. Intracardial perfusion
   1. Anesthetize the rats with 2.5% tribromoethanol (160 µL/10 g) via intraperitoneal injection. Ensure the rats are fully anesthetized by verifying the lack of response to external stimuli, such as a gentle pinch of the toe.
   2. When the proper anesthetization is confirmed, place the rats supine and immobilize them in the Petri dish filled with silica gel.
   3. Cut a 5-mm longitudinal skin incision caudally to the xiphoid process, then fully expand the subcutaneous space. Cut a 2-cm longitudinal skin incision along the ventral midline to fully expose the outer chest wall, starting with the above-mentioned incision and ending with the top of the chest.
   4. Use toothed tweezers to lift the xiphoid (Supplementary Figure 2A), and then use fine scissors to cut the diaphragm. Cut the sternum along both sides of the xiphoid process to open the chest and expose the heart (Supplementary Figure 2B).
      NOTE: Be careful to preserve the internal thoracic vessels on both sides; otherwise, it may cause massive bleeding.
   5. Use toothless tweezers to lift the left ventricle. Insert a 22 G needle into the left ventricular apex along the longitudinal axis of the left ventricle (Supplementary Figure 2C). Meanwhile, observe the rhythmic blood pulsation in the perfusion tube, or the needle may puncture into the right ventricle, which may lead to a poor perfusion effect.
      NOTE: Toothed tweezer should not be used; otherwise it may cause extra blood leakage from the tweezer holding site.
   6. Use fine scissors to cut the right atrium (Supplementary Figure 2C), then manually inject 100 mL of ice-cold perfusing fluid at a rate of about 2 mL/s within 1 min.
      NOTE: When rats' liver and paws turn pale, and no blood flows out from the right atrium, good perfusion can be achieved.
5. Spinal cord dissection (Figure 1)
   1. Place the rat in the prone position, and cut the spine at the anterior superior iliac spine (about L4 vertebral level) and the curvature shifting point of the thoracic column (about T6 vertebral level), respectively (Figure 1A). Then, immediately place the isolated spine in the oxygenated ice-cold perfusing solution to wash off the residual blood and fat tissue; this procedure is beneficial for keeping the operative field clean in the subsequent procedures.
      NOTE: The paraspinal muscles should be reserved, which is conducive for the subsequent fixation of the spine by using an insect pin.
   2. Immediately transfer the isolated spine to the anatomical tray (Figure 1B) with the dorsal side up and the rostral end close to the operator. Fill the anatomical tray with 50 mL of continuously oxygenated ice-cold cutting solution (Figure 1B).
   3. Use four insect pins to fix the spine by penetrating the paraspinal muscles (Figure 1B).
   4. Under the dissection microscope, cut the vertebral pedicles of both sides from the rostral end with the micro-scissor, which can be termed "laminectomy" (Figure 1C). Pay attention not to damage the spinal cord. Meanwhile, use the micro-toothed tweezers to lift the cut vertebral body.
   5. After the laminectomy, do not separate the spinal cord from the spinal canal immediately. Instead, use a micro-scissor to cut the dura mater along the dorsal midline, which is conducive for nutrient uptake between cells and oxygenated ACSF (Figure 1D).
      NOTE: Never tear the dura mater. Only cutting the dura mater by micro-scissor is permitted; otherwise, the nerve root and the spinal parenchyma will be seriously damaged!
   6. Lift the rostral part of the spinal cord, then carefully cut the nerve root with about 1 mm reserved (Figure 1E). After separating the spinal cord from the vertebral canal, use 2 insect pins to fix the spinal cord with the ventral side up (Figure 1F).
   7. Use a micro-scissor to cut the dura mater along the ventral midline (Figure 1F). Cut off the redundant nerve roots with about 1 mm reserved.
      NOTE: The nerve is much more tenacious than the spinal cord. If the reserved nerve root is too long (>1 mm), the vibratome cannot cut off the nerve root, which may lead to a serious tear in the spinal parenchyma.
   8. Use a micro-scissor to separate the lumbar enlargement to a length of 6-7 mm (Figure 1G).
6. Embedding in the low-melting agarose
   1. Place the lumbar enlargement on the 35 °slope (Figure 1H) with the dorsal side up and caudal end down. Use an absorbent filter paper to remove abundant water on the tissue surface (Figure 1H).
   2. Slowly pour the molten agarose gel into the Petri dish containing the lumbar enlargement (Figure 1I).
      NOTE: Do not pour too fast, or bubbles will accumulate in the gel.
   3. Place the above Petri dish in the ice-water mixture to cool the gel as soon as possible, which is conducive to maintaining the activity of cells.
   4. Trim the gel into a 15 mm x 10 mm x 10 mm cube and mount it on the specimen disc with superglue (Figure 1J).
7. Slicing
   1. Place the specimen disc into the pre-frozen cutting tray, then pour the ice-cold cutting solution (Figure 1K). Continuously bubble with 95% CO₂ and 5% O₂ into the cutting tray.
   2. Set the vibratome parameters: thickness: 350 µm; speed: 0.14-0.16 mm/s, amplitude: 1.0 mm, and vibration frequency: 85 Hz.
   3. Harvest 2-3 suitable slices per animal. Record 1-2 healthy FG+ motor neurons per slice, with a range of 5-6 cells per animal.
8. Incubation
   1. Use cover slide tweezers to clip a slice (Figure 1L) and place it into the incubation chamber filled with continuously oxygenated ACSF. Place the incubation chamber in a water bath at 32 °C for 30 min, and then continue to incubate it at room temperature (RT) for another 30 min prior to recording.
      ​NOTE: The above procedures, from anesthesia to obtaining the first slice, should be completed within 20-30 min to retain the viability of cells as much as possible. The motor neurons in each slice can maintain their viability for approximately 6-7 h.

2. Patch-clamp recording with SCS (Figure 2)

1. Preparations
   1. Perfuse the recording chamber with continuously bubbled ACSF at a rate of about 1-2 mL/min. Adjust the flow rate individually via the control panel on the peristaltic pump.
   2. Place a slice into the recording chamber. Use the U-shaped platinum wire with nylon threads to firmly stabilize the slice in place.
   3. Use an infrared differential interference contrast microscope (IR-DIC) to observe the slice. Under the 4x objective lens, confirm that the length of the dorsal root is about 1 mm. Find the area where the dorsal root enters the parenchyma, then move the central field of vision to this area.
   4. Connect the pulse generator with custom-made electrodes (Figure 2A).
2. SCS configuration
   1. Place the anode of SCS near the dorsal midline via the micromanipulation system (Figure 2B).
   2. Place the cathode of SCS near the dorsal root entry zone (DREZ) via the micromanipulation system (Figure 2B).
3. Cell targeting and imaging
   1. Use IR-DIC with a 10x objective lens roughly find the dorsolateral region of the motor column, where most motor neurons are located. Then, move the central field of vision to this area.
   2. Switch to a 60x objective lens to find a healthy neuron with a smooth and bright surface and invisible nuclei (Figure 2C,F).
   3. Slightly turn down the IR intensity and turn on the fluorescence light source. Switch the light filter to the wide band ultraviolet excitation filter (Figure 2D,G), to select an appropriate FG-positive (FG+) motor neuron (Figure 2E,H).
   4. Use suction electrodes to apply 1x motor threshold stimulation to the dorsal root. If an evoked action potential is detected in the motor neurons (Supplementary Figure 3), it confirms that the activity of the dorsal root is intact. If not, this slice should be discarded.
4. Patch-clamp recording
   1. Fill the micropipette with the intracellular solution and insert it into the electrode holder. Use the micromanipulation system to lower the pipette into the ACSF bath. The pipette resistance ranges from 5-8 MΩ.
   2. Apply a small amount of positive pressure to the pipette to blow away the dust and cell debris.
      1. Lower the electrode to approach the cell. When the pipette touches the surface of FG+ neuron, a small indentation of the membrane becomes visible at the level of the tip. Release the positive pressure.
      2. Then, apply a small amount of negative pressure to the pipette using a syringe. This creates a small amount of suction that pulls the cell membrane into contact with the glass pipette. Always pay attention to the total resistance on the software interface until the resistance value increases to gigaohms (>1 GΩ). Then, the gigaseal is formed.
   3. Clamp the membrane potential at -70 mV, then press the fast capacitance compensation button on the software interface of the amplifier. Gently apply a transiently negative pressure to rupture the cell membrane, then press the slow capacitance compensation button on the software interface of the amplifier. At this point, a good whole-cell configuration is obtained.
   4. Switch to current-clamp mode by clicking the IC button on the software interface, and record the resting membrane potential (RMP).
   5. Apply the SCS for 1-2 s with the amplitude of 1-10 mA, while the pulse width and frequency are fixed at 210 µs and 40 Hz, respectively. Determine the motor threshold by slowly increasing the stimulation amplitude until the first AP is observed.
   6. Distinguish delayed and immediate firing motor neurons using a 5 s depolarizing current injection around rheobase in the current-clamp mode^10,11,12. Immediate firing motor neurons: Low rheobase can induce immediate and repetitive firing with stable firing frequency; Delayed firing motor neurons: High rheobase can induce a delayed onset for repetitive firing with an accelerating firing rate (Figure 3).
   7. When SCS is turned off, continue to record the membrane potential for capturing the spontaneous APs firing.
   8. Perform voltage-clamp recordings voltage-clamp recordings for excitatory postsynaptic current (EPSC) when SCS is on and off. The stimulation parameter is 1x motor threshold, 210 µs, 2 Hz.