The Ex vivo Preparation of Spinal Cord Slice for the Whole-Cell Patch-Clamp Recording in Motor Neurons During Spinal Cord Stimulation

Summary

This protocol describes a method using a patch-clamp to study the electrical responses of motor neurons to spinal cord stimulation (SCS) with high spatiotemporal resolution, which can help researchers improve their skills in separating the spinal cord and maintaining cell viability simultaneously.

Abstract

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to execute sensorimotor behaviors, directly studying the electrical responses of motor neurons with SCS can help us understand the underlying logic of spinal motor modulation. To simultaneously record diverse stimulus characteristics and cellular responses, a patch-clamp is a good method to study the electrophysiological characteristics at a single-cell scale. However, there are still some complex difficulties in achieving this goal, including maintaining cell viability, quickly separating the spinal cord from the bony structure, and using the SCS to successfully induce action potentials. Here, we present a detailed protocol using patch-clamp to study the electrical responses of motor neurons to SCS with high spatiotemporal resolution, which can help researcher improve their skills in separating the spinal cord and maintaining the cell viability at the same time to smoothly study the electrical mechanism of SCS on motor neuron and avoid unnecessary trial and mistake.

Introduction

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Andreas Rowald et al. reported that SCS enables lower limb locomotor and trunk function within a single day¹. Exploring the biological mechanism of SCS for locomotor recovery is a critical and trending research field for developing a more precise SCS strategy. For example, Grégoire Courtine's team demonstrated that excitatory Vsx2 interneuron and Hoxa10 neurons in the spinal cord are the key neurons to response to SCS, and cell-specific neuromodulation is feasible to restore the rat walking ability after SCI². However, few studies focus on the electrical mechanism of SCS at a single-cell scale. Although it is well-known that the suprathreshold direct current stimulus can elicit the action potentials (APs) in the classic squid experiment^3,4,5, how the pulsed alternating electrical stimulation, such as SCS, affects the motor signal generation is still unclear.

Given the complexity of intraspinal neural circuits, appropriate selection for cell population is important for investigating the electrical mechanism of SCS. Although SCS restores motor function by activating the proprioceptive pathway⁶, the motor neurons are the final unit to execute the motor command, derived from integrating proprioception information afferent input⁷. Therefore, directly studying the electrical characteristics of motor neurons with SCS can help us understand the underlying logic of spinal motor modulation.

As we know, patch-clamp is the golden-standard method for cellularly electrophysiological recording with extremely high spatiotemporal resolution⁸. Therefore, this study describes a method using a patch clamp to study the electrical responses of motor neurons to SCS. Compared with the brain patch-clamp⁹, the spinal cord patch-clamp is more difficult due to the following reasons: (1) The spinal cord is protected by the vertebral canal with tiny volume, which requires very fine micromanipulation and rigorous ice-cold maintenance to obtain better cell viability. (2) Because the spinal cord is too slender to be secured on the cutting tray, it should be immersed in low-melting point agarose and trimmed after solidification.

Hence, this method provides technical details in dissecting the spinal cord and maintaining the cell viability at the same time so as to smoothly study the electrical mechanism of SCS on motor neurons and avoid unnecessary trials and mistakes.

Protocol

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.

Results

Thanks to the rigorous low-temperature maintenance during the fine operation (Supplementary Figure 1, Supplementary Figure 2, and Figure 1), the cell viability was good enough to perform subsequent electrophysiological recordings. To simulate the clinical scenario as much as possible, we used micromanipulation to place the SCS cathode and anode near the dorsal midline and DREZ, respectively (Figure 2), which could initiate neural signal in ...

Discussion

The movement information modulated by SCS is finally converged to the motor neurons. Therefore, taking the motor neurons as the research target may simplify the study design and reveal the neuromodulation mechanism of SCS more directly. To simultaneously record diverse stimulus characteristics and cellular responses, a patch-clamp is a good method to study the electrophysiological characteristics at a single-cell scale. However, there are still some difficulties, including how to maintain cell viability, how to quickly s...

Materials

Name	Company	Catalog Number	Comments
Adenosine 5’-triphosphate magnesium salt	Sigma	A9187
Ascorbic Acid	Sigma	A4034
CaCl2·2H2O	Sigma	C5080
Choline Chloride	Sigma	C7527
Cover slide tweezers	VETUS	36A-SA	Clip a slice
D-Glucose	Sigma	G8270
EGTA	Sigma	E4378
Fine scissors	RWD Life Science	S12006-10	Cut the diaphragm
Fluorescence Light Source	Olympus	U-HGLGPS
Fluoro-Gold	Fluorochrome	Fluorochrome	Label the motor neuron
Guanosine 5′-triphosphate sodium salt hydrate	Sigma	G8877
HEPES	Sigma	H3375
infrared CCD camera	Dage-MTI	IR-1000E
KCl	Sigma	P5405
K-gluconate	Sigma	P1847
Low melting point agarose	Sigma	A9414
MgSO4·7H2O	Sigma	M2773
Micromanipulator	Sutter Instrument	MP-200
Micropipette puller	Sutter instrument	P1000
Micro-scissors	Jinzhong	wa1020	Laminectomy
Microscope for anatomy	Olympus	SZX10
Microscope for ecletrophysiology	Olympus	BX51WI
Micro-toothed tweezers	RWD Life Science	F11008-09	Lift the cut vertebral body
NaCl	Sigma	S5886
NaH2PO4	Sigma	S8282
NaHCO3	Sigma	V900182
Na-Phosphocreatine	Sigma	P7936
Objective lens for ecletrophysiology	Olympus	LUMPLFLN60XW	working distance 2 mm
Osmometer	Advanced	FISKE 210
Patch-clamp amplifier	Axon	Multiclamp 700B
Patch-clamp digitizer	Axon	Digidata 1550B
pH meter	Mettler Toledo	FE28
Slice Anchor	Multichannel system	SHD-27H
Spinal cord stimulatior	PINS	T901
Toothed tweezer	RWD Life Science	F13030-10	Lift the xiphoid
Vibratome	Leica	VT1200S
Wide band ultraviolet excitation filter	Olympus	U-MF2