In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation

All procedures connected to this protocol have been accepted by the appropriate authorities (e.g., Local Ethics Committee) and follow the national and international rules on animal welfare and management.

NOTE: Each participant involved in the procedure has to be properly trained in basic surgical procedures and has to have a valid license for performing animal experiments.

1. Anesthesia and premedication

1. Anesthetize a rat with intraperitoneal injections of sodium pentobarbital (an initial dose of 60 mg·kg^-1 for 6-month old male Wistar rats weighing 400‒550g).
   NOTE: This protocol is not limited to the indicated strain, sex or age of rats. Also, alternative anesthesia such as ketamine-xylazine mix, alpha-chloralose or fentanyl+midazolam+medetomidine can be used if more suitable for different research goals or when required by the ethics committee.
2. After approximately 5 min, check the depth of anesthesia by pinching the rat’s hind limb toe with blunt forceps. Proceed with the next steps of the protocol only when no reflex action is observed.
3. Inject 0.05 mL of atropine subcutaneously in order to reduce mucus production after intubation.
4. Inject subcutaneously 5 mL of phosphate buffer containing 4% glucose solution, NaHCO₃ (1 %) and gelatin (14%). This buffer will be absorbed by the cutaneous vessels throughout an experiment and will help maintain fluid balance.
5. Throughout the surgery, periodically check the animal for reflex actions and supplement anesthesia if required (10 mg·kg^-1·h^-1of sodium pentobarbital).

2. Surgery

1. Prepare the animal for surgical treatment by shaving fur over the dorsal part of the left hindlimb, from the ankle to the hip, the backside, from tail to the high thoracic segments, the left side of the chest, and the ventral side of the neck area above the sternum
2. Placement of the intravenous line
   1. Place the rat on its back on a closed-loop heating pad (and secure it with limb fixations).
   2. Using a 21 blade, make a longitudinal cut through the skin from a sternum to a chin.
   3. Hold the skin with forceps and separate it from the underlying tissue.
   4. Using blunt dissection techniques expose the right jugular vein. Carefully dissect the vein from surrounding tissues.
   5. Locate the part of the vein without branching points, slip two 4-0 ligatures beneath it.
   6. Make one loose knot on the proximal end of the previously identified non-branching segment of the vein and one loose knot on the distal end of this segment of the vein. Clamp the vein proximal to the heart, and then ligate the distal part of the vein.
   7. Using iris scissors, make an incision between the clamp and distant ligature. Hold a flap of the vein, and introduce a pre-filled catheter to the point where it is blocked by the clamp.
   8. While holding the vein and the catheter together with forceps, remove the clamp and push the catheter several millimeters into the vein. Secure both ends of the catheter to the vein, and add an additional fixation point to the skin.
3. Introduction of the tracheal tube
   1. Using blunt forceps separate the two mandibular glands covering the sternohyoid muscles. Separate sternohyoid muscles at the midline to expose the trachea.
   2. Slip three 4-0 ligatures beneath the trachea, then make two knots below the tracheal tube insertion point and one knot above.
   3. Locate the cricoid cartilage of the larynx and make an incision below the third tracheal cartilage.
   4. Insert a tracheal tube down the trachea and secure the tube in place with pre-prepared ligatures, then add an additional ligature to the skin.
   5. Place a small piece of cotton wool above the separated muscles, and suture the skin over the operated area.
4. Dissection of hind limb nerves
   1. Using 21 blade, make a longitudinal cut on the posterior side of the left hind limb, from the Achilles tendon to the hip.
   2. Grab the skin with forceps, and using blunt dissection techniques separate the skin from underlying muscles on both sides of the incision.
   3. Locate the popliteal fossa at the back of the knee joint, which is covered by the biceps femoris muscle, and using scissors make a cut between the anterior and posterior part of this muscle.
   4. Moving upwards cut two heads of the biceps femoris all the way to the hip to expose the sciatic nerve. Cauterize as needed to prevent bleeding.
   5. Identify the sural, tibial and common peroneal branches of the sciatic nerve.
   6. Using scissors, separate the lateral from the medial head of the gastrocnemius muscle to expose the tibial nerve and its branches.
   7. Using 55 forceps grab the distal end of the sural nerve, cut it distally and dissect as far as possible.
   8. Repeat the procedure with the common peroneal nerve.
   9. Using a blunt glass rod separate the tibial nerve from surrounding tissues, taking care not to damage the blood vessels, and cut it distally.
   10. Identify the medial gastrocnemius (MG) and the lateral gastrocnemius and soleus (LGS) nerves.
   11. Using 55 forceps, carefully dissect the MG and LGS nerves, disconnecting them from surrounding tissues, but maintaining their connection to the respective muscles.
   12. Place a saline-soaked piece of cotton wool under the exposed nerves.
   13. Close the skin over the operated area.
5. Laminectomy
   1. Using 21 blade make a longitudinal incision from the sacrum up to the thoracic vertebrae.
   2. Separate the skin from underlying muscles.
   3. Cut the longissimus muscle on both sides of the thoracic and lumbar spinous processes.
   4. Using blunt-edged scalpel retract the muscles from the spinal column to expose the transverse processes of each vertebra.
   5. Using blunt tip scissors cut the tendons of muscles connected to the transverse processes along the exposed spinal column. Apply hemostatic agents if necessary.
   6. Identify the Th13 vertebra as the lowest thoracic segment with rib insertion and using fine rongeurs remove spinous processes and laminae from Th13 to L2 vertebrae to expose lumbar segments of the spinal cord. Remember not to damage the L3 spinous process which will be used as a fixation point for spine stabilization.
   7. Remove the Th12 spinous process and smooth the vertebra dorsal surface as much as possible.
   8. Using blunt dissection techniques separate the muscles from Th11 vertebra to create holder insertion points.
   9. Place thin saline-soaked cotton wool over the exposed spinal cord segments.
   10. Move the rat to the custom made metal frame with two parallel bars and two adjustable arms with clamps to support and stabilize the spine.

3. Preparation for the recording and stimulation

1. Vertebral column fixation and nerve arrangement
   1. Place the rat in the custom-made frame on a heating pad, connected to the closed-loop heating system to maintain the animal body temperature at 37 ± 1°C.
   2. Insert ECG electrodes under the skin and connect to an amplifier for heart rate monitoring.
   3. Using the skin flaps, form a deep pool over the exposed spinal cord.
   4. Using metal clamps, fix the vertebral column by putting clamps below Th12 transverse processes and at L3 spinous process.
   5. Make sure that the vertebral column is secured and arranged horizontally, and then apply dorso-ventral pressure on both sides of the column to retract the muscles.
   6. Fill the pool with warm (37 °C) mineral oil and maintain it at this temperature.
   7. Thread a 4-0 ligature through the Achilles tendon, lift and stretch the operated left hind limb so that the ankle is leveled with the hip.
   8. Using the skin flaps make a deep pool over the exposed tibial, MG and LGS nerves.
   9. Fill the pool with warm (37 °C) mineral oil.
   10. Place MG and LGS nerves on bipolar silver-wire stimulating electrodes and connect them to a square pulse stimulator. Use separate stimulation channels for each nerve.
2. Surface electrode placement
   1. Place a silver ball electrode on the left caudal side of the exposed spinal cord, with a reference electrode inserted in the back muscles, and connect both electrodes to the differential DC amplifier. The surface ball electrode will be used to record afferent volleys from nerves.
   2. Using a constant-current stimulator, stimulate the MG and LGS nerves with square pulses of 0.1 ms duration, repeated at a frequency of 3 Hz, and observe afferent volleys.
   3. Determine the threshold (T) for nerve activation, stimulate each nerve at approximately 3·T intensity, and record amplitude of afferent volley for each nerve.
   4. Move the surface electrode rostral and repeat the procedure to identify spinal segments at which amplitudes of the volleys are the highest for each nerve. After determining the maximum volley location, move the surface electrode to a safe distance from the spinal cord.
3. Muscle paralysis and forming a pneumothorax in order to reduce respiratory movements
   1. Paralyze the rat intravenously with a neuromuscular blocker and connect the tracheal tube to an external ventilator in line with a rodent-compatible capnometer (Pancuronium bromide, at an initial dose of 0.4 mg·kg^-1, supplemented every 30 min in doses of 0.2 mg·kg^-1)
   2. Monitor the end-tidal CO₂ concentration and maintain it at about 3‒4% by adjusting ventilation parameters (frequency, air pressure, and flow volumes).
   3. Make a longitudinal incision in the skin between the 5th and 6th rib on a side of the recording.
   4. Using blunt tip scissors cut the overlying muscles to visualize intercostal space between the ribs.
   5. Using small sharp scissors, make a small incision in the intercostal muscles and in the pleura, then insert a tip of a blunt edge forceps into the opening, taking care not to press on the lungs.
   6. Allow forceps to expand or insert a small tube to keep the pneumothorax open throughout the experiment.
   7. After the neuromuscular block, monitor anesthesia depth by checking ECG frequency, and supplement the anesthetic agent if the heart rate exceeds 400 bpm. Muscle paralysis and forming a pneumothorax in order to reduce respiratory movements, which will improve recording stability
4. Opening the dura and pia mater
   1. Using #55 forceps, gently lift the dura mater, and cut it caudally from the L5 segment, rostrally up to the L4 segment.
   2. Using a pair of ultra-thin 5SF forceps make a small patch in the pia covering the dorsal column, between the blood vessels, exactly at the level of the maximum afferent volley from the MG or the LGS nerve.
   3. Use small pieces of saline-soaked and dried gel foam to block bleeding if necessary.
5. tsDCS electrode placement
   1. Make a small incision in the skin on the ventral side of a rat abdomen at the rostro-caudal level corresponding to the location of L4-L5 spinal segments.
   2. Grab the exposed skin flap with a metal clip which will serve as a reference electrode.
   3. Place a saline-soaked sponge on the dorsal side of the Th12 vertebra. Make sure that the sponge size is equal to that of an active tsDCS electrode (circle-shaped stainless steel plate of 5 mm in diameter).
   4. Using a fine manipulator, press the sponge with an active tsDCS electrode to the bone and make sure that the entire surface of the electrode is pressed equally.
   5. Connect both reference and active tsDCS electrodes to a constant-current stimulator unit, capable of delivering a continuous flow of direct current.
6. Preparation of micropipettes
   1. Using a microelectrode puller, prepare a microelectrode.
      NOTE: Both filament and non-filament electrodes can be used, however, remember that the shank of the electrode must be long enough to reach the ventral horn while being thin enough not to compress the spinal cord while descending.
      1. Adjust the puller setting so that the shank entering the spinal cord is approximately 3 mm long, while the tip of the electrode is no more than 1‒2 µm in diameter and microelectrode resistance is between 10 and 20 MΩ.
   2. Fill the microelectrodes with 2M potassium-citrate electrolyte.
   3. Mount the prepared microelectrode on the micromanipulator allowing 1‒2 µm stepping movement and stereotaxic calibration.
   4. Connect the microelectrode to the intracellular amplifier with the reference electrode placed in the back muscles.
7.  After the neuromuscular block, monitor anesthesia depth by checking ECG frequency, and supplement the anesthetic agent so that the rat heart rate does not exceed 400 bpm.

4. Motoneuron tracking and penetration

1. Place the afferent volley recording electrode back on the dorsal surface of the spinal cord, caudally to the location of the recording site, at the level of the L6 segment.
2. Stimulate the MG and LGS nerves with electrical 0.1 ms pulses at a frequency of 3 Hz, and 3T intensity, to activate all the axons of alpha-motoneurons within a selected nerve.
3. Drive the micropipette into a selected patch in the pia with a medio-lateral angle of 15‒20° (with a tip directed laterally).
4. After descending below the surface, calibrate the microelectrode and compensate its capacitance and voltage offset, and continue penetration of the spinal cord when all parameters are stable. An antidromic field potential of the motoneuron pool will be visible at the microelectrode voltage trace while approaching a dedicated motor nucleus during stimulation of the respective nerve.
5. Proceed penetration with the microelectrode at 1‒2 µm steps, and periodically use the buzz function of the intracellular amplifier to clear the electrode tip from any residue.
6. Observe motoneuron penetration which will be characterized by a sudden hyperpolarization of the recorded voltage trace and appearance of an antidromic spike potential.

5. Recording motoneuron membrane and firing properties

1. In a bridge mode of the intracellular amplifier, identify the motoneuron on the basis of the “all-or-nothing” appearance of the antidromic action potential by stimulating respective nerve branches. Record 20 subsequent traces for later averaging.
2.  Implement a strict inclusion criterion to ensure high-quality data: resting membrane potential of at least -50 mV in amplitude; action potential amplitudes greater than 50 mV, with a positive overshoot; membrane potential stable for at least 5 min prior to recording.
3. In a discontinuous current-clamp mode (current switch rate mode 4–8 kHz) of the intracellular amplifier, evoke an orthodromic action potential in a motoneuron using 0.5 ms intracellular depolarizing current pulses. Repeat at least 20 times for offline averaging.
4. Stimulate a motoneuron with 40 short pulses (100 ms) of hyperpolarizing current (1 nA) in order to calculate cell input resistance.
5. Stimulate a motoneuron with 50 ms square-wave pulses at increasing amplitudes to determine the rheobase value as the minimum amplitude of depolarizing current required to elicit a single spike.
6. Inject 500 ms square-wave pulses of depolarizing current, at increasing amplitudes in steps of 0.1–2 nA to evoke rhythmic discharges of motoneurons.

6. Trans-spinal direct current stimulation (tsDCS)

1. While maintaining a stable penetration of the motoneuron, start the polarization procedure by trans-spinal application of direct current. Adjust the current intensity and application time to the experiment design (e.g., 0.1 mA for 15 min).
2. Immediately after switching on the DC, observe the motoneuron membrane potential. Anodal polarization (the active electrode as an anode) should result in depolarization of the membrane potential, while cathodal polarization (the active electrode as a cathode) should evoke an opposite effect. Observe whether a change in the resting membrane potential in response to DC stimulation is constant, which ensures that electrical field intensity is not affected.
3. During continuous current application, repeat steps 5.3‒5.6 in 5 min intervals.
4. Turn off the DC and continue to repeat steps 5.3‒5.6 in 5 min intervals until recordings become unstable or inclusion criteria are compromised.
5. Terminate the experiment and euthanizethe animal using intravenous administration of a lethal dose of pentobarbital sodium (180 mg·kg^-1).