eoe sub articles

EoE Sub Articles

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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
<ol>
	<li>Anesthetize a rat with intraperitoneal injections of sodium pentobarbital (an initial dose of 60 mg&#183;kg-1 for 6-month-old male Wistar rats weighing 400&#8210;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.</li>
	<li>After approximately 5 min, check the depth of anesthesia by pinching the rat&#39;s hind limb toe with blunt forceps. Proceed with the next steps of the protocol only when no reflex action is observed.</li>
	<li>Inject 0.05 mL of atropine subcutaneously in order to reduce mucus production after intubation.</li>
	<li>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.</li>
	<li>Throughout the surgery, periodically check the animal for reflex actions and supplement anesthesia if required (10 mg&#183;kg-1&#183; h-1 of sodium pentobarbital).</li>
</ol>
2. Surgery
<ol>
	<li>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 the tail to the high thoracic (Th) segments, the left side of the chest, and the ventral side of the neck area above the sternum</li>
	<li>Placement of the intravenous line
	<ol>
		<li>Place the rat on its back on a closed-loop heating pad (and secure it with limb fixations).</li>
		<li>Using a 21-blade, make a longitudinal cut through the skin from the sternum to the chin.</li>
		<li>Hold the skin with forceps and separate it from the underlying tissue.</li>
		<li>Using blunt dissection techniques, expose the right jugular vein. Carefully dissect the vein from surrounding tissues.</li>
		<li>Locate the part of the vein without branching points and slip two 4-0 ligatures beneath it.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
	</ol>
	</li>
</ol>
3. Introduction of the tracheal tube
<ol>
	<li>Using blunt forceps, separate the two mandibular glands covering the sternohyoid muscles. Separate sternohyoid muscles at the midline to expose the trachea.</li>
	<li>Slip three 4-0 ligatures beneath the trachea, then make two knots below the tracheal tube insertion point and one knot above.</li>
	<li>Locate the cricoid cartilage of the larynx and make an incision below the third tracheal cartilage.</li>
	<li>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.</li>
	<li>Place a small piece of cotton wool above the separated muscles and suture the skin over the operated area.</li>
</ol>
4. Dissection of hind limb nerves
<ol>
	<li>Using a 21 blade, make a longitudinal cut on the posterior side of the left hind limb, from the Achilles tendon to the hip.</li>
	<li>Grab the skin with forceps, and using blunt dissection techniques, separate the skin from the underlying muscles on both sides of the incision.</li>
	<li>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.</li>
	<li>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.</li>
	<li>Identify the sural, tibial, and common peroneal branches of the sciatic nerve.</li>
	<li>Using scissors, separate the lateral from the medial head of the gastrocnemius muscle to expose the tibial nerve and its branches.</li>
	<li>Using 55 forceps, grab the distal end of the sural nerve, cut it distally, and dissect it as far as possible.</li>
	<li>Repeat the procedure with the common peroneal nerve.</li>
	<li>Using a blunt glass rod separate the tibial nerve from surrounding tissues, taking care not to damage the blood vessels, and cut it distally.</li>
	<li>Identify the medial gastrocnemius (MG) and the lateral gastrocnemius and soleus (LGS) nerves.</li>
	<li>Using 55 forceps, carefully dissect the MG and LGS nerves, disconnecting them from surrounding tissues but maintaining their connection to the respective muscles.</li>
	<li>Place a saline-soaked piece of cotton wool under the exposed nerves.</li>
	<li>Close the skin over the operated area.</li>
</ol>
5. Laminectomy
<ol>
	<li>Using a 21 blade, make a longitudinal incision from the sacrum up to the thoracic vertebrae.</li>
	<li>Separate the skin from underlying muscles.</li>
	<li>Cut the longissimus muscle on both sides of the thoracic and lumbar spinous processes.</li>
	<li>Using a blunt-edged scalpel, retract the muscles from the spinal column to expose the transverse processes of each vertebra.</li>
	<li>Using blunt tip scissors, cut the tendons of muscles connected to the transverse processes along the exposed spinal column. Apply hemostatic agents if necessary.</li>
	<li>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.</li>
	<li>Remove the Th12 spinous process and smooth the vertebra dorsal surface as much as possible.</li>
	<li>Using blunt dissection techniques, separate the muscles from the Th11 vertebra to create holder insertion points.</li>
	<li>Place thin, saline-soaked cotton wool over the exposed spinal cord segments.</li>
	<li>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.</li>
</ol>
6. Preparation for the recording and stimulation
<ol>
	<li>Vertebral column fixation and nerve arrangement</li>
	<li>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 &#177; 1&#176;C.</li>
	<li>Insert electrocardiogram (ECG) electrodes under the skin and connect to an amplifier for heart rate monitoring.</li>
	<li>Using the skin flaps, form a deep pool over the exposed spinal cord.</li>
	<li>Using metal clamps, fix the vertebral column by putting clamps below the Th12 transverse processes and at L3 spinous process.</li>
	<li>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.</li>
	<li>Fill the pool with warm (37 &#176;C) mineral oil and maintain it at this temperature.</li>
	<li>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.</li>
	<li>Using the skin flaps, make a deep pool over the exposed tibial, MG, and LGS nerves.</li>
	<li>Fill the pool with warm (37 &#176;C) mineral oil.</li>
	<li>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.</li>
</ol>
7. Surface electrode placement
<ol>
	<li>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.</li>
	<li>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.</li>
	<li>Determine the threshold (T) for nerve activation, stimulate each nerve at approximately 3&#183;T intensity, and record the amplitude of the afferent volley for each nerve.</li>
	<li>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.</li>
	<li>Muscle paralysis and forming a pneumothorax in order to reduce respiratory movements</li>
	<li>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&#183;kg-1, supplemented every 30 min in doses of 0.2 mg&#183;kg-1)</li>
	<li>Monitor the end-tidal CO2 concentration and maintain it at about 3&#8210;4% by adjusting ventilation parameters (frequency, air pressure, and flow volumes).</li>
	<li>Make a longitudinal incision in the skin between the 5th and 6th rib on the side of the recording.</li>
	<li>Using blunt tip scissors, cut the overlying muscles to visualize the intercostal space between the ribs.</li>
	<li>Using small, sharp scissors, make a small incision in the intercostal muscles and in the pleura, then insert the tip of a blunt-edge forceps into the opening, taking care not to press on the lungs.</li>
	<li>Allow forceps to expand or insert a small tube to keep the pneumothorax open throughout the experiment.</li>
	<li>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 to reduce respiratory movements which will improve recording stability.</li>
</ol>
8. Opening the dura and pia mater
<ol>
	<li>Using #55 forceps, gently lift the dura mater and cut it caudally from the L5 segment, rostrally up to the L4 segment.</li>
	<li>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.</li>
	<li>Use small pieces of saline-soaked and dried gel foam to block bleeding if necessary.</li>
</ol>
9. Trans-spinal direct current stimulation (tsDCS) electrode placement
<ol>
	<li>Make a small incision in the skin on the ventral side of a rat&#39;s abdomen at the rostrocaudal level corresponding to the location of lumbar (L)4-L5 spinal segments.</li>
	<li>Grab the exposed skin flap with a metal clip, which will serve as a reference electrode.</li>
	<li>Place a saline-soaked sponge on the dorsal side of the thoracic (Th) 12 vertebra. Ensure that the sponge size is equal to that of an active tsDCS electrode (circle-shaped stainless steel plate of 5 mm in diameter).</li>
	<li>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.</li>
	<li>Connect reference and active tsDCS electrodes to a constant-current stimulator unit capable of delivering a continuous flow of direct current.</li>
</ol>
10. Preparation of micropipettes
<ol>
	<li>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.</li>
	<li>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&#8210;2 &#181;m in diameter, and microelectrode resistance is between 10 and 20 M&#937;.</li>
	<li>Fill the microelectrodes with 2M potassium-citrate electrolyte.</li>
	<li>Mount the prepared microelectrode on the micromanipulator, allowing 1&#8210;2 &#181;m stepping movement and stereotaxic calibration.</li>
	<li>Connect the microelectrode to the intracellular amplifier with the reference electrode placed in the back muscles.</li>
	<li>After the neuromuscular block, monitor anesthesia depth by checking electrocardiogram (ECG) frequency and supplement the anesthetic agent so that the rat heart rate does not exceed 400 bpm.</li>
</ol>
11. Motoneuron tracking and penetration
<ol>
	<li>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.</li>
	<li>Stimulate the medial gastrocnemius (MG) and the lateral gastrocnemius and soleus (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.</li>
	<li>Drive the micropipette into a selected patch in the pia with a mediolateral angle of 15&#8210;20&#176; (with a tip directed laterally).</li>
	<li>After descending below the surface, calibrate the microelectrode and compensate for 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.</li>
	<li>Proceed penetration with the microelectrode at 1&#8210;2 &#181;m steps and periodically use the buzz function of the intracellular amplifier to clear the electrode tip from any residue.</li>
	<li>Observe motoneuron penetration, which will be characterized by a sudden hyperpolarization of the recorded voltage trace and the appearance of an antidromic spike potential.</li>
</ol>
12. Recording motoneuron membrane and firing properties
<ol>
	<li>In a bridge mode of the intracellular amplifier, identify the motoneuron based on the &#34;all-or-nothing&#34; appearance of the antidromic action potential by stimulating respective nerve branches. Record 20 subsequent traces for later averaging.</li>
	<li>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.</li>
	<li>In a discontinuous current-clamp mode (current switch rate mode 4&#8211;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.</li>
	<li>Stimulate a motoneuron with 40 short pulses (100 ms) of hyperpolarizing current (1 nA) in order to calculate cell input resistance.</li>
	<li>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.</li>
	<li>Inject 500 ms square-wave pulses of depolarizing current, at increasing amplitudes in steps of 0.1&#8211;2 nA to evoke rhythmic discharges of motoneurons.</li>
</ol>
13. Trans-spinal direct current stimulation (tsDCS)
<ol>
	<li>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).</li>
	<li>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.</li>
	<li>During continuous current application, repeat steps 5.3&#8210;5.6 in 5 min intervals.</li>
	<li>Turn off the DC and continue to repeat steps 5.3&#8210;5.6 in 5-minute intervals until recordings become unstable or inclusion criteria are compromised.</li>
	<li>Terminate the experiment and euthanize the animal using intravenous administration of a lethal dose of pentobarbital sodium (180 mg&#183;kg-1).</li>
</ol>

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Durgs and solutions</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Atropinum sulfuricum</td>
			<td>Polfa Warszawa</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Glucose</td>
			<td>Merck</td>
			<td>346351</td>
			<td>-</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Merck</td>
			<td>106329</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Pancuronium Jelfa</td>
			<td>PharmaSwiss/Valeant</td>
			<td>-</td>
			<td>Neuromuscular blocker</td>
		</tr>
		<tr>
			<td>Pentobarbital sodium</td>
			<td>Biowet Pu<img alt="Equation 1" src="/files/ftp_upload/22707/22707eq01.jpg" style="margin: auto;" />awy Sp. z o.o</td>
			<td>-</td>
			<td>Main anesthetic agent</td>
		</tr>
		<tr>
			<td>Pottasium citrate</td>
			<td>Chempur</td>
			<td>6100-05-06</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Tetraspan</td>
			<td>Braun</td>
			<td>-</td>
			<td>HES solution</td>
		</tr>
		<tr>
			<td>Surgical equipment</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>21 Blade</td>
			<td>FST</td>
			<td>10021-00</td>
			<td>Scalpel blade</td>
		</tr>
		<tr>
			<td>Cauterizer</td>
			<td>FST</td>
			<td>18010-00</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Chest Tubes</td>
			<td>Mila</td>
			<td>CT1215</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Dumont #4 Forceps</td>
			<td>FST</td>
			<td>11241-30</td>
			<td>Muscle forceps</td>
		</tr>
		<tr>
			<td>Dumont #5 Forceps</td>
			<td>FST</td>
			<td>11254-20</td>
			<td>Dura forceps</td>
		</tr>
		<tr>
			<td>Dumont #5F Forceps</td>
			<td>FST</td>
			<td>11255-20</td>
			<td>Nerve forceps</td>
		</tr>
		<tr>
			<td>Dumont #5SF Forceps</td>
			<td>FST</td>
			<td>11252-00</td>
			<td>Pia forceps</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>FST</td>
			<td>11008-13</td>
			<td>Blunt forceps</td>
		</tr>
		<tr>
			<td>Forceps</td>
			<td>FST</td>
			<td>11053-10</td>
			<td>Skin forceps</td>
		</tr>
		<tr>
			<td>Hemostat</td>
			<td>FST</td>
			<td>13013-14</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Rongeur</td>
			<td>FST</td>
			<td>16021-14</td>
			<td>For laminectomy</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>FST</td>
			<td>15000-08</td>
			<td>Vein scissors</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>FST</td>
			<td>15002-08</td>
			<td>Dura scissors</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>FST</td>
			<td>14184-09</td>
			<td>For trachea cut</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>FST</td>
			<td>104075-11</td>
			<td>Muscle scissors</td>
		</tr>
		<tr>
			<td>Scissors</td>
			<td>FST</td>
			<td>14002-13</td>
			<td>Skin scissors</td>
		</tr>
		<tr>
			<td>Tracheal tube</td>
			<td>-</td>
			<td>-</td>
			<td>Custom made</td>
		</tr>
		<tr>
			<td>Vein catheter</td>
			<td>Vygon</td>
			<td>1261.201</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Vessel cannulation forceps</td>
			<td>FST</td>
			<td>18403-11</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Vessel clamp</td>
			<td>FST</td>
			<td>18320-11</td>
			<td>For vein clamping</td>
		</tr>
		<tr>
			<td>Vessel Dilating Probe</td>
			<td>FST</td>
			<td>10160-13</td>
			<td>For vein dissection</td>
		</tr>
		<tr>
			<td>Sugrgical materials</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Gel foam</td>
			<td>Pfizer</td>
			<td>GTIN 00300090315085</td>
			<td>Hemostatic agent</td>
		</tr>
		<tr>
			<td>Silk suture 4.0</td>
			<td>FST</td>
			<td>18020-40</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Silk suture 6.0</td>
			<td>FST</td>
			<td>18020-60</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Equipment</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>Axoclamp 2B</td>
			<td>Molecular devices</td>
			<td>discontinued</td>
			<td>Intracellular amplifier/ new model Axoclamp 900A</td>
		</tr>
		<tr>
			<td>CapStar-100 End-tidal CO2 Monitor</td>
			<td>CWE</td>
			<td>11-10000</td>
			<td>Gas analyzer</td>
		</tr>
		<tr>
			<td>Grass S-88</td>
			<td>A-M Systems</td>
			<td>discontinued</td>
			<td>Constant current stimulator</td>
		</tr>
		<tr>
			<td>Homeothermic Blanket Systems with Flexible Probe</td>
			<td>Harvard Apparatus</td>
			<td>507222F</td>
			<td>Heating system</td>
		</tr>
		<tr>
			<td>ISO-DAM8A</td>
			<td>WPI</td>
			<td>74020</td>
			<td>Extracellular amplifier</td>
		</tr>
		<tr>
			<td>Microdrive</td>
			<td>-</td>
			<td>-</td>
			<td>Custom made/replacement IVM/Scientifica</td>
		</tr>
		<tr>
			<td>P-1000 Microelectrode puller</td>
			<td>Sutter Instruments</td>
			<td>P-1000</td>
			<td>Microelectrode puller</td>
		</tr>
		<tr>
			<td>SAR-830/AP Small Animal Ventilator</td>
			<td>CWE</td>
			<td>12-02100</td>
			<td>Respirator</td>
		</tr>
		<tr>
			<td>Support frame</td>
			<td>-</td>
			<td>-</td>
			<td>Custom made/replacement lab standard base 51601/Stoelting</td>
		</tr>
		<tr>
			<td>Spinal clamps</td>
			<td>-</td>
			<td>-</td>
			<td>Custom made/replacement Rat spinal adaptor 51695/Stoelting</td>
		</tr>
		<tr>
			<td>TP-1 DC stimulator</td>
			<td>WiNUE</td>
			<td>-</td>
			<td>tsDCS stimulator</td>
		</tr>
		<tr>
			<td>Miscellaneous</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>1B150-4 glass capillaries</td>
			<td>WPI</td>
			<td>1B150-4</td>
			<td>For microelectrodes production</td>
		</tr>
		<tr>
			<td>Cotton wool</td>
			<td>-</td>
			<td>-</td>
			<td>-</td>
		</tr>
		<tr>
			<td>flexible tubing</td>
			<td>-</td>
			<td>-</td>
			<td>For respirator and CO2 analyzer connection</td>
		</tr>
		<tr>
			<td>MicroFil</td>
			<td>WPI</td>
			<td>MF28G67-5</td>
			<td>For filling micropipettes</td>
		</tr>
		<tr>
			<td>Silver wire</td>
			<td>-</td>
			<td>-</td>
			<td>For nerve electrodes</td>
		</tr>
	</tbody>
</table>

trans-spinal direct current stimulation of a rat&#39;s spinal motoneuron

Source: B&#261;czyk, M., et al. <a href="https://app-jove-com.remotexs.ntu.edu.sg/v/61439">In Vivo Intracellular Recording of Type-Identified Rat Spinal Motoneurons During Trans-Spinal Direct Current Stimulation.</a> J. Vis. Exp. (2020).
This video describes the in vivo intracellular recording of rat motoneurons under trans-spinal direct current stimulation. The protocol describes how to measure membrane properties and record the rhythmic firing of motoneurons before, during, and after anodal or cathodal polarization of the spinal cord.

Trans-Spinal Direct Current Stimulation of a Rat&#39;s Spinal Motoneuron

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.
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Begin by placing a saline-soaked sponge on the thoracic-12 vertebra of a rat prepared for the procedure.
Position an electrode on it for trans-spinal direct current stimulation or tsDCS, which involves applying a direct current across the spinal cord.
To measure the intracellular recording of a target motoneuron during tsDCS, insert a microelectrode into the spinal cord at the location of its origin.
Assess the motoneuron&#39;s baseline membrane potential and the frequency of electrical impulses to ensure the microelectrode is securely placed for accurate measurements during tsDCS.
Apply anodal tsDCS, where positively charged ions enter the neuron as the current flows from the anode through the tissue to the cathode.
Apply cathodal tsDCS, where negatively charged ions enter the neuron as the current flows in the reverse direction.
Record the changes in the motoneuron&#39;s electrical properties during anodal and cathodal tsDCS.

trans-spinal-direct-current-stimulation-of-a-rat-s-spinal-motoneuron

Nanyang Technological University

Research

JoVE Encyclopedia of Experiments

Neuroscience

Neurophysiology

Stimulation and Modulation Techniques

128 vues. Source : Bączyk, M., et al. Enregistrement intracellulaire in vivo de motoneurones spinaux de rat identifiés de type lors d’une stimulation transspinale à courant continu. J. Vis. Exp. (2020). Cette vidéo décrit l’enregistrement intracellulaire in vivo de motoneurones de rats sous stimulation transspinale à courant continu. Le protocole décrit comment mesurer les propriétés de la membrane et enregistrer l’activation rythmique des motoneurones avant, pendant et après la polarisat...

Vidéo: Stimulation transspinale à courant continu du motoneurone spinal d'un rat - Concept

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method described here enables the investigator to measure long-lasting (from minutes to hours) high-quality intracellular responses from single Drosophila R1-R6 photoreceptors and Large Monopolar Cells (LMCs) to light stimuli. Because the recording system has low noise, it can be used to study variability among individual cells in the fly eye, and how their outputs reflect the physical properties of the visual environment. We outline all key steps in performing this technique. The basic steps in constructing an appropriate electrophysiology set-up for recording, such as design and selection of the experimental equipment are described. We also explain how to prepare for recording by making appropriate (sharp) recording and (blunt) reference electrodes. Details are given on how to fix an intact fly in a bespoke fly-holder, prepare a small window in its eye and insert a recording electrode through this hole with minimal damage. We explain how to localize the center of a cell's receptive field, dark- or light-adapt the studied cell, and to record its voltage responses to dynamic light stimuli. Finally, we describe the criteria for stable normal recordings, show characteristic high-quality voltage responses of individual cells to different light stimuli, and briefly define how to quantify their signaling performance. Many aspects of the method are technically challenging and require practice and patience to master. But once learned and optimized for the investigator's experimental objectives, it grants outstanding in vivo neurophysiological data.

Electrophysiological Method for Recording Intracellular Voltage Responses of Drosophila Photoreceptors and Interneurons to Light Stimuli In Vivo

Sharp microelectrodes enable accurate electrophysiological characterization of photoreceptor and visual interneuron output in living Drosophila. Here we show how to use this method to record high-quality voltage responses of individual cells to controlled light stimulation. This method is ideal for studying neural information processing in insect compound eyes.

Méthode électrophysiologique de réponses d&#39;enregistrement intracellulaire tension de<em&gt; Drosophila</em&gt; Photorécepteurs et Interneurones à Light Stimuli<em&gt; In Vivo</em

Voltage responses of insect photoreceptors and visual interneurons can be accurately recorded with conventional sharp microelectrodes. The method ...

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JoVE Journal

Neurosciences

Transcranial Direct Current Stimulation (tDCS) in Mice

Transcranial direct current stimulation (tDCS) is a non-invasive neuromodulation technique proposed as an alternative or complementary treatment for several neuropsychiatric diseases. The biological effects of tDCS are not fully understood, which is in part explained due to the difficulty in obtaining human brain tissue. This protocol describes a tDCS mouse model that uses a chronically implanted electrode allowing the study of the long-lasting biological effects of tDCS. In this experimental model, tDCS changes the cortical gene expression and offers a prominent contribution to the understanding of the rationale for its therapeutic use.

Transcranial Direct Current Stimulation tDCS in Mice

Transcranial direct current stimulation (tDCS) is a therapeutic technique proposed to treat psychiatric diseases. An animal model is essential for understanding the specific biological alterations evoked by tDCS. This protocol describes a tDCS mouse model that uses a chronically implanted electrode.

Stimulation transcrânienne courant continu (CDV) chez la souris

Transcranial direct current stimulation (tDCS) is a non-invasive neuromodulation technique proposed as an alternative or complementary treatment for ...

Enregistrement électrophysiologique extracellulaire dans le cortex moteur de souris

Multichannel Extracellular Recording in Freely Moving Mice

The protocol aims to uncover the properties of neuronal firing and network local field potentials (LFPs) in behaving mice carrying out specific tasks by correlating the electrophysiological signals with spontaneous and/or specific behavior. This technique represents a valuable tool in studying the neuronal network activity underlying these behaviors. The article provides a detailed and complete procedure for electrode implantation and consequent extracellular recording in free-moving conscious mice. The study includes a detailed method for implanting the microelectrode arrays, capturing the LFP and neuronal spiking signals in the motor cortex (MC) using a multichannel system, and the subsequent offline data analysis. The advantage of multichannel recording in conscious animals is that a greater number of spiking neurons and neuronal subtypes can be obtained and compared, which allows the evaluation of the relationship between a specific behavior and the associated electrophysiological signals. Notably, the multichannel extracellular recording technique and the data analysis procedure described in the present study can be applied to other brain areas when conducting experiments in behaving mice.

Pleins feux sur l’auteur : Progrès de l’enregistrement extracellulaire multicanal pour l’étude de l’activité neuronale chez les souris se déplaçant librement 

The protocol describes the methodology of extracellular recording in the motor cortex (MC) to reveal extracellular electrophysiological properties in freely moving conscious mice, as well as the data analysis of local field potentials (LFPs) and spikes, which is useful for evaluating the network neural activity underlying behaviors of interest.

Enregistrement extracellulaire multicanal chez des souris se déplaçant librement

The protocol aims to uncover the properties of neuronal firing and network local field potentials (LFPs) in behaving mice carrying out specific tasks by ...

Trans-Spinal Direct Current Stimulation of a Rat's Spinal Motoneuron

Transcription

Trans-Spinal Direct Current Stimulation of a Rat's Spinal Motoneuron