Spinal cord stimulation can help patients restore locomotion functions after injury. Motor neurons are the final unit to execute sensorimotor behaviors. Studying the electrical response of motor neurons can aid in understanding the underlying logic of spinal motor modulation.

Patch-clamp is the golden standard method for cellular electro physiological recordings with the extremely high spatial temporal resolution. Therefore, this study describes a method using a patch-clamp to simultaneously record diverse stimulus characteristics and cellular responses of multi neurons at a single-cell scale. Compared with the brain patch-clamp, the spinal cord patch-clamp is more difficult, because the spinal cord is well protected by the vertebral canal with tiny volume.

Therefore, this protocol provides technical details in quickly micro dissecting the spinal cord, and rigorous ice-cold maintenance to obtain better cell viability. Patch-clamp allows a unique understanding of the synaptic transmissions and the understanding of action potentials and coding pattern activities by the spinal cord stimulation. After exposing the heart of an anesthetized rat, cut the right atrium using fine scissors and inject 100 milliliters of chilled perfusing fluid at approximately two milliliters per second within one minute.

Position the rat in the prone position, cut the spine at the anterior superior iliac spine and the curvature shifting point of the thoracic column. Immediately place the isolated spine into the oxygenated ice cold perfusing solution to wash off residual blood and fat tissue. Then transfer the isolated spine to the anatomical tray with its dorsal side up and the rostral end near the operator.

Fill the tray with 50 milliliters of continuously oxygenated ice cold cutting solution. Under a dissection microscope, cut the vertebrae pedicles on both sides from the rostral end using micro scissors. Next, delicately sever the dura mater along its dorsal midline with micro scissors.

Lift the rostral part of the spinal cord carefully and cut the nerve root, leaving about one millimeter reserved. After the spinal cord has been separated from the vertebral canal, secure it with the ventral side facing up using two insect pins. Trim any excessive nerve roots, leaving about one millimeter intact.

Then separate the lumbar enlargement to a length of six to seven millimeters using a micro scissor. After dissecting the spinal cord and separating the lumbar enlargement, place the lumbar enlargement on a 35 degree slope with the dorsal side up and the caudal side down. Absorb any excess water on the tissue surface using an absorbent filter paper.

Carefully pour the molten agarose gel into the Petri dish containing the lumbar enlargement. Position the Petri dish in the ice water mixture to ensure rapid gel cooling. Shape the gel into a cube of 15 by 10 by 10 millimeters and mount it onto the specimen disc using super glue.

Adjust the vibratome settings to a thickness of 350 micrometers, a speed of 0.14 to 0.16 millimeters per second, an amplitude of one millimeter, and a vibration frequency of 85 hertz. Now, use the cover slide tweezers to clip a slice and place it into the incubation chamber filled with continuously oxygenated artificial cerebrospinal fluid. To begin, place the prepared spinal cord slice in the recording chamber.

Then place the anode of the micro-manipulation system near the dorsal midline and the cathode close to the dorsal root entry zone for proper spinal cord stimulator positioning. Now switch to a 60X objective lens to identify a healthy neuron, having a smooth and bright surface without visible nuclei. Slightly decrease the infrared intensity, turn on the fluorescent light source, and adjust the light filter to the wideband ultraviolet excitation option.

Now select an appropriate Fluoro-Gold positive motor neuron, and gradually lower the electrode until it is near the neuron. As the pipette contacts its surface, look for a small membrane indentation at the tip level and release the previously applied positive pressure. Then apply negative pressure to the pipette using a syringe and wait until the resistance value rises to gigaohms on the software interface.

Clamp the membrane potential at minus 70 millivolts. Press the fast capacitance compensation button on the amplifier's software interface. Then apply a brief negative pressure to rupture the cell membrane and press the slow capacitance compensation button on the amplifier's software interface.

Apply the spinal cord stimulation for one to two seconds, keeping the amplitude between one and 10 milliamperes. Determine the motor threshold by gradually increasing the stimulation amplitude until the first action potential is observed. Distinguish delayed and immediate firing motor neurons using a five-second depolarizing current injection around rheobase in the current clamp mode.

After turning off the spinal cord stimulation, continue recording the membrane potentials to capture the spontaneous action potentials firing. When the amplitude of spinal cord stimulation was raised, the membrane potential was increased, and action potentials were triggered every 10 to 20 pulses. After the spinal cord stimulation was turned off, the neuron fired a series of spontaneous action potentials for a short period of time.

Then the resting membrane potential returned to minus 65 millivolts.