This study was funded by the&#160;National Natural Science Foundation of China&#160;for Young Scholars (52207254 and 82301657) and the&#160;China Postdoctoral Science Fund (2022M711833).

The Institutional Animal Care and Use Committee approved all animal experiments&#160;and the studies were conducted in accordance with relevant animal welfare regulations.
1. Animals preparation
<ol>
	<li>Animals
	<ol>
		<li>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 &#176;C &#177; 2 &#176;C, humidity: 50%-60%, with a 12-h light/ dark cycle. Animals had free access to food and water.</li>
		<li>Label the motor neurons retrogradely: Inject Fluoro-Gold (FG) into the bilateral tibialis anterior and gastrocnemius muscle (2% in sterile saline, 50 &#181;L per muscle) to retrogradely label the motor neurons 2 days before the sacrifice.</li>
	</ol>
	</li>
	<li>Solutions
	<ol>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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 &#176;C water bath to maintain the liquid state.</li>
	</ol>
	</li>
	<li>Instrument preparation
	<ol>
		<li>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 &#176;C overnight in advance.</li>
		<li>Place the incubation chamber with nylon mesh in the oven at 45 &#176;C overnight in advance.</li>
		<li>Use low-melting point agarose to prefabricate a 35&#176; 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.</li>
	</ol>
	</li>
	<li>Intracardial perfusion
	<ol>
		<li>Anesthetize the rats with 2.5% tribromoethanol (160 &#181;L/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.</li>
		<li>When the proper anesthetization is confirmed, place the rats supine and immobilize them in the Petri dish filled with silica gel.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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.</li>
		<li>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&#39; liver and paws turn pale, and no blood flows out from the right atrium, good perfusion can be achieved.</li>
	</ol>
	</li>
	<li>Spinal cord dissection (Figure 1)
	<ol>
		<li>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.</li>
		<li>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).</li>
		<li>Use four insect pins to fix the spine by penetrating the paraspinal muscles (Figure 1B).</li>
		<li>Under the dissection microscope, cut the vertebral pedicles of both sides from the rostral end with the micro-scissor, which can be termed &#34;laminectomy&#34; (Figure 1C). Pay attention not to damage the spinal cord. Meanwhile, use the micro-toothed tweezers to lift the cut vertebral body.</li>
		<li>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!</li>
		<li>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).</li>
		<li>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 (&#62;1 mm), the vibratome cannot cut off the nerve root, which may lead to a serious tear in the spinal parenchyma.</li>
		<li>Use a micro-scissor to separate the lumbar enlargement to a length of 6-7 mm (Figure 1G).</li>
	</ol>
	</li>
	<li>Embedding in the low-melting agarose
	<ol>
		<li>Place the lumbar enlargement on the 35 &#176;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).</li>
		<li>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.</li>
		<li>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.</li>
		<li>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).</li>
	</ol>
	</li>
	<li>Slicing
	<ol>
		<li>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.</li>
		<li>Set the vibratome parameters: thickness: 350 &#181;m; speed: 0.14-0.16&#160;mm/s, amplitude: 1.0&#160;mm, and vibration frequency: 85 Hz.</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Incubation
	<ol>
		<li>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 &#176;C for 30 min, and then continue to incubate it at room temperature (RT) for another 30 min prior to recording. 
		&#8203;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.</li>
	</ol>
	</li>
</ol>
2. Patch-clamp recording with SCS (Figure 2)
<ol>
	<li>Preparations
	<ol>
		<li>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.</li>
		<li>Place a slice into the recording chamber. Use the U-shaped platinum wire with nylon threads to firmly stabilize the slice in place.</li>
		<li>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.</li>
		<li>Connect the pulse generator with custom-made electrodes (Figure 2A).</li>
	</ol>
	</li>
	<li>SCS configuration
	<ol>
		<li>Place the anode of SCS near the dorsal midline via the micromanipulation system (Figure 2B).</li>
		<li>Place the cathode of SCS near the dorsal root entry zone (DREZ) via the micromanipulation system (Figure 2B).</li>
	</ol>
	</li>
	<li>Cell targeting and imaging
	<ol>
		<li>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.</li>
		<li>Switch to a 60x objective lens to find a healthy neuron with a smooth and bright surface and invisible nuclei (Figure 2C,F).</li>
		<li>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).</li>
		<li>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.</li>
	</ol>
	</li>
	<li>Patch-clamp recording
	<ol>
		<li>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 M&#937;.</li>
		<li>Apply a small amount of positive pressure to the pipette to blow away the dust and cell debris.
		<ol>
			<li>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.</li>
			<li>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 (&#62;1 G&#937;). Then, the gigaseal is formed.</li>
		</ol>
		</li>
		<li>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.</li>
		<li>Switch to current-clamp mode by clicking the IC button on the software interface, and record the resting membrane potential (RMP).</li>
		<li>Apply the SCS for 1-2 s with the amplitude of 1-10 mA, while the pulse width and frequency are fixed at 210 &#181;s and 40 Hz, respectively. Determine the motor threshold by slowly increasing the stimulation amplitude until the first AP is observed.</li>
		<li>Distinguish delayed and immediate firing motor neurons using a 5 s depolarizing current injection around rheobase in the current-clamp mode10,11,12. 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).</li>
		<li>When SCS is turned off, continue to record the membrane potential for capturing the spontaneous APs firing.</li>
		<li>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 &#181;s, 2 Hz.</li>
	</ol>
	</li>
</ol>

Understanding Motor Neuron Responses to Spinal Cord Stimulation

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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...

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, Gr&#233;goire Courtine&#39;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 experiment3,4,5, 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.

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		<tr>
			<td>Adenosine 5&#8217;-triphosphate magnesium salt</td>
			<td>Sigma</td>
			<td>A9187</td>
			<td></td>
		</tr>
		<tr>
			<td>Ascorbic Acid</td>
			<td>Sigma</td>
			<td>A4034</td>
			<td></td>
		</tr>
		<tr>
			<td>CaCl2&#183;2H2O</td>
			<td>Sigma</td>
			<td>C5080</td>
			<td></td>
		</tr>
		<tr>
			<td>Choline Chloride</td>
			<td>Sigma</td>
			<td>C7527</td>
			<td></td>
		</tr>
		<tr>
			<td>Cover slide tweezers</td>
			<td>VETUS</td>
			<td>36A-SA</td>
			<td>Clip a slice</td>
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		<tr>
			<td>D-Glucose</td>
			<td>Sigma</td>
			<td>G8270</td>
			<td></td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>Sigma</td>
			<td>E4378</td>
			<td></td>
		</tr>
		<tr>
			<td>Fine scissors</td>
			<td>RWD Life Science</td>
			<td>S12006-10</td>
			<td>Cut the diaphragm</td>
		</tr>
		<tr>
			<td>Fluorescence Light Source</td>
			<td>Olympus&#160;</td>
			<td>U-HGLGPS</td>
			<td></td>
		</tr>
		<tr>
			<td>Fluoro-Gold</td>
			<td>Fluorochrome</td>
			<td>Fluorochrome</td>
			<td>Label the motor neuron</td>
		</tr>
		<tr>
			<td>Guanosine 5&#8242;-triphosphate sodium salt hydrate</td>
			<td>Sigma</td>
			<td>G8877</td>
			<td></td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>Sigma</td>
			<td>H3375</td>
			<td></td>
		</tr>
		<tr>
			<td>infrared CCD camera</td>
			<td>Dage-MTI</td>
			<td>IR-1000E</td>
			<td></td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>Sigma</td>
			<td>P5405</td>
			<td></td>
		</tr>
		<tr>
			<td>K-gluconate</td>
			<td>Sigma</td>
			<td>P1847</td>
			<td></td>
		</tr>
		<tr>
			<td>Low melting point agarose</td>
			<td>Sigma</td>
			<td>A9414</td>
			<td></td>
		</tr>
		<tr>
			<td>MgSO4&#183;7H2O</td>
			<td>Sigma</td>
			<td>M2773</td>
			<td></td>
		</tr>
		<tr>
			<td>Micromanipulator&#160;</td>
			<td>Sutter Instrument&#160;</td>
			<td>MP-200</td>
			<td></td>
		</tr>
		<tr>
			<td>Micropipette puller</td>
			<td>Sutter instrument</td>
			<td>P1000</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-scissors&#160;</td>
			<td>Jinzhong</td>
			<td>wa1020</td>
			<td>Laminectomy</td>
		</tr>
		<tr>
			<td>Microscope for anatomy</td>
			<td>Olympus&#160;</td>
			<td>SZX10</td>
			<td></td>
		</tr>
		<tr>
			<td>Microscope for ecletrophysiology</td>
			<td>Olympus&#160;</td>
			<td>BX51WI</td>
			<td></td>
		</tr>
		<tr>
			<td>Micro-toothed tweezers</td>
			<td>RWD Life Science</td>
			<td>F11008-09</td>
			<td>Lift the cut vertebral body</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>Sigma</td>
			<td>S5886</td>
			<td></td>
		</tr>
		<tr>
			<td>NaH2PO4</td>
			<td>Sigma</td>
			<td>S8282</td>
			<td></td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>Sigma</td>
			<td>V900182</td>
			<td></td>
		</tr>
		<tr>
			<td>Na-Phosphocreatine</td>
			<td>Sigma</td>
			<td>P7936</td>
			<td></td>
		</tr>
		<tr>
			<td>Objective lens for ecletrophysiology</td>
			<td>Olympus&#160;</td>
			<td>LUMPLFLN60XW</td>
			<td>working distance 2 mm&#160;</td>
		</tr>
		<tr>
			<td>Osmometer&#160;</td>
			<td>Advanced&#160;</td>
			<td>FISKE 210</td>
			<td></td>
		</tr>
		<tr>
			<td>Patch-clamp amplifier&#160;</td>
			<td>Axon&#160;</td>
			<td>Multiclamp 700B</td>
			<td></td>
		</tr>
		<tr>
			<td>Patch-clamp digitizer</td>
			<td>Axon&#160;</td>
			<td>Digidata 1550B</td>
			<td></td>
		</tr>
		<tr>
			<td>pH meter&#160;</td>
			<td>Mettler Toledo&#160;</td>
			<td>FE28</td>
			<td></td>
		</tr>
		<tr>
			<td>Slice Anchor</td>
			<td>Multichannel system</td>
			<td>SHD-27H</td>
			<td></td>
		</tr>
		<tr>
			<td>Spinal cord stimulatior</td>
			<td>PINS</td>
			<td>T901</td>
			<td></td>
		</tr>
		<tr>
			<td>Toothed tweezer</td>
			<td>RWD Life Science</td>
			<td>F13030-10</td>
			<td>Lift the xiphoid</td>
		</tr>
		<tr>
			<td>Vibratome</td>
			<td>Leica</td>
			<td>VT1200S</td>
			<td></td>
		</tr>
		<tr>
			<td>Wide band ultraviolet excitation filter</td>
			<td>Olympus&#160;</td>
			<td>U-MF2</td>
			<td></td>
		</tr>
	</tbody>
</table>

the ex vivo preparation of spinal cord slice for the whole-cell patch-clamp recording in motor neurons during spinal cord stimulation

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.

Thanks to the rigorous low-temperature maintenance during the fine operation (Supplementary Figure 1, Supplementary Figure 2,&#160;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 ...

Watch this Scientific Journal Video about Advancing Spinal Cord Stimulation - Exploring the Cellular Responses of Motor Neurons Through Patch-Clamp Electrophysiology at JoVE.com

Author Spotlight: Advancing Spinal Cord Stimulation - Exploring the Cellular Responses of Motor Neurons Through Patch-Clamp Electrophysiology 

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.

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

Spinal cord stimulation (SCS) can effectively restore locomotor function after spinal cord injury (SCI). Because the motor neurons are the final unit to ...

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JoVE Journal

Neuroscience

2.3K Views.  Tsinghua University. 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.

Video: Author Spotlight: Advancing Spinal Cord Stimulation - Exploring the Cellular Responses of Motor Neurons Through Patch-Clamp Electrophysiology 

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies of these neurons are located in the dorsal spinal cord and their axons follow stereotyped trajectories during embryonic development. Commissural axons initially project ventrally towards the floorplate. After crossing the midline, these axons turn anteriorly and project towards the brain. Each of these steps is regulated by the action of several guidance cues. Cultures highly enriched in commissural neurons are ideally suited for many experiments addressing the mechanisms of axon pathfinding, including turning assays, immunochemistry and biochemistry. Here, we describe a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. First, the spinal cord is isolated and dorsal strips are dissected out. The dorsal tissue is then dissociated into a cell suspension by trypsinization and mechanical disruption. Neurons are plated onto poly-L-lysine-coated glass coverslips or tissue-culture dishes. After 30 hours in vitro, most neurons have extended an axon. The purity of the culture (Yam et al. 2009), typically over 90%, can be assessed by immunolabeling with the commissural neuron markers DCC, LH2 and TAG1 (Helms and Johnson, 1998). This neuronal preparation is a useful tool for in vitro studies of the cellular and molecular mechanisms of commissural axon growth and guidance during spinal cord development.

This video demonstrates a method to dissect and culture commissural neurons from E13 rat dorsal spinal cord. Dissociated commissural neurons are useful to study the cellular and molecular mechanisms of axon growth and guidance.

Commissural neurons have been widely used to investigate the mechanisms underlying axon guidance during embryonic spinal cord development. The cell bodies ...

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate a method to fabricate suction electrodes that are used to examine CPG activity, or fictive locomotion, in dissected rodent spinal cords. The rodent spinal cords are placed in artificial cerebrospinal fluid and the ventral roots are drawn into the suction electrode. The electrode is constructed by modifying a commercially available suction electrode. A heavier silver wire is used instead of the standard wire given by the commercially available electrode. The glass tip on the commercial electrode is replaced with a plastic tip for increased durability. We prepare hand drawn electrodes and electrodes made from specific sizes of tubing, allowing consistency and reproducibility. Data is collected using an amplifier and neurogram acquisition software. Recordings are performed on an air table within a Faraday cage to prevent mechanical and electrical interference, respectively.

A demonstration of the fabrication and use of an extracellular suction electrode used to measure electrophysiological recordings of neonatal rodent spinal cords in vitro

Development of neural circuitries and locomotion can be studied using neonatal rodent spinal cord central pattern generator (CPG) behavior. We demonstrate ...

Chicken Embryo Spinal Cord Slice Culture Protocol

Slice cultures can facilitate the manipulation of embryo development both pharmacologically and through gene manipulations. In this reduced system, potential lethal side effects due to systemic drug applications can be overcome. However, culture conditions must ensure that normal development proceeds within the reduced environment of the slice. We have focused on the development of the spinal cord, particularly that of spinal motor neurons. We systematically varied culture conditions of chicken embryo slices from the point at which most spinal motor neurons had been born. We assayed the number and type of motor neurons that survived during the culture period and the position of those motor neurons compared to that in vivo. We found that serum type and neurotrophic factors were required during the culture period and were able to keep motor neurons alive for at least 24 hr and allow those motor neurons to migrate to appropriate positions in the spinal cord. We present these culture conditions and the methodology of preparing the embryo slice cultures using eviscerated chicken embryos embedded in agarose and sliced using a vibratome.

Slice cultures facilitate the manipulation of embryo development by gene and pharmacological perturbations. However, culture conditions must ensure that normal development can proceed within the reduced environment of the slice. We illustrate a protocol that facilitates normal spinal cord development to proceed for at least 24 hr.

Slice cultures can facilitate the manipulation of  embryo development both pharmacologically and through gene manipulations. In  this reduced system, ...

Spinal Cord Transection in the Larval Zebrafish

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability to reinitiate spinal neurogenesis. However, some anamniotes including the zebrafish Danio rerio exhibit both sensory and functional recovery even after complete transection of the spinal cord. The adult zebrafish is an established model organism for studying regeneration following spinal cord injury, with sensory and motor recovery by 6&#160;weeks post-injury. To take advantage of in vivo analysis of the regenerative process available in the transparent larval zebrafish as well as genetic tools not accessible in the adult, we use the larval zebrafish to study regeneration after spinal cord transection. Here we demonstrate a method for reproducibly and verifiably transecting the larval spinal cord. After transection, our data shows sensory recovery beginning at 2&#160;days post-injury (dpi), with the C-bend movement detectable by 3 dpi and resumption of free swimming by 5 dpi. Thus we propose the larval zebrafish as a companion tool to the adult zebrafish for the study of recovery after spinal cord injury.

After spinal transection, adult zebrafish have functional recovery by six weeks post-injury. To take advantage of larval transparency and faster recovery, we present a method for transecting the larval spinal cord. After transection, we observe sensory recovery beginning at 2&#160;days post-injury, and C-bend movement by 3 days post-injury.

Mammals fail in sensory and motor recovery following spinal cord injury due to lack of axonal regrowth below the level of injury as well as an inability ...

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal degeneration. Lack of growth cone formation, regeneration, and loss of additional myelinated axonal projections within the spinal cord greatly limits neurological recovery following injury. To assess how central myelinated axons of the spinal cord respond to injury, we developed an ex vivo living spinal cord model utilizing transgenic mice that express yellow fluorescent protein in axons and a focal and highly reproducible laser-induced spinal cord injury to document the fate of axons and myelin (lipophilic fluorescent dye Nile Red) over time using two-photon excitation time-lapse microscopy. Dynamic processes such as acute axonal injury, axonal retraction, and myelin degeneration are best studied in real-time. However, the non-focal nature of contusion-based injuries and movement artifacts encountered during in vivo spinal cord imaging make differentiating primary and secondary axonal injury responses using high resolution microscopy challenging. The ex vivo spinal cord model described here mimics several aspects of clinically relevant contusion/compression-induced axonal pathologies including axonal swelling, spheroid formation, axonal transection, and peri-axonal swelling providing a useful model to study these dynamic processes in real-time. Major advantages of this model are excellent spatiotemporal resolution that allows differentiation between the primary insult that directly injures axons and secondary injury mechanisms; controlled infusion of reagents directly to the perfusate bathing the cord; precise alterations of the environmental milieu (e.g., calcium, sodium ions, known contributors to axonal injury, but near impossible to manipulate in vivo); and murine models also offer an advantage as they provide an opportunity to visualize and manipulate genetically identified cell populations and subcellular structures. Here, we describe how to isolate and image the living spinal cord from mice to capture dynamics of acute axonal injury.

An Ex Vivo Laser-induced Spinal Cord Injury Model to Assess Mechanisms of Axonal Degeneration in Real-time

We present a protocol utilizing two-photon excitation time-lapse microscopy to simultaneously visualize the dynamics of axon and myelin injuries in real time. This proposed protocol permits studies of both intrinsic and extrinsic factors which can influence central myelinated axon fate after injury and contribute to permanent clinical disability.

Injured CNS axons fail to regenerate and often retract away from the injury site. Axons spared from the initial injury may later undergo secondary axonal ...

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by repeated, time-lapse in vivo imaging in live rodents. These studies were made possible by straightforward surgical procedures, which enabled optical access for a prolonged period of time without repeat surgical procedures. In contrast, analogous studies of the spinal cord have been previously limited to only a few imaging sessions, each of which required an invasive surgery. As previously described, we have developed a spinal chamber that enables continuous optical access for upwards of 8 weeks, preserves mechanical stability of the spinal column, is easily stabilized externally during imaging, and requires only a single surgery. Here, the design of the spinal chamber with its associated surgical implements is reviewed and the surgical procedure is demonstrated in detail. Briefly, this video will demonstrate the preparation of the surgical area and mouse for surgery, exposure of the spinal vertebra and appropriate tissue debridement, the delivery of the implant and vertebral clamping, the completion of the chamber, the removal of the delivery system, sealing of the skin, and finally, post-operative care. The procedure for chronic in vivo imaging using nonlinear microscopy will also be demonstrated. Finally, outcomes, limitations, typical variability, and a guide for troubleshooting are discussed.

A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

Studies in the mammalian neocortex have enabled unprecedented resolution of cortical structure, activity, and response to neurodegenerative insults by ...

The Preparation of Oblique Spinal Cord Slices for Ventral Root Stimulation

Electrophysiological recordings from spinal cord slices have proven to be a valuable technique to investigate a wide range of questions, from cellular to network properties. We show how to prepare viable oblique slices of the spinal cord of young mice (P2 - P11). In this preparation, the motoneurons retain&#160;their axons coming out from the ventral roots of the&#160;spinal cord. Stimulation of these axons elicits back-propagating action potentials invading the motoneuron&#160;somas and exciting the motoneuron&#160;collaterals within the spinal cord. Recording of antidromic action potentials is an immediate, definitive and elegant way to characterize motoneuron identity, which surpasses&#160;other identification methods. Furthermore, stimulating the motoneuron collaterals is a simple and reliable way to excite the collateral targets of the motoneurons within the spinal cord, such as other motoneurons or Renshaw cells. In this protocol, we present antidromic recordings from the motoneuron somas as well as Renshaw cell excitation, resulting from ventral root stimulation.

We show how to prepare oblique slices of the spinal cord in young mice. This preparation allows for the stimulation of the ventral roots.

Electrophysiological recordings from spinal cord slices have proven to be a valuable technique to investigate a wide range of questions, from cellular to ...

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing organisms, combined with the embryonic optical clarity, served as initial compelling attributes of this model. Over the past two decades, the success of this model has been further propelled by its amenability to large-scale mutagenesis screens and by the ease of transgenesis. More recently, gene-editing approaches have extended the power of the model.
For neurodevelopmental studies, the zebrafish embryo and larva provide a model to which multiple methods can be applied. Here, we focus on methods that allow the study of an essential property of neurons, electrical excitability. Our preparation for the electrophysiological study of zebrafish spinal neurons involves the use of veterinarian suture glue to secure the preparation to a recording chamber. Alternative methods for recording from zebrafish embryos and larvae involve the attachment of the preparation to the chamber using a fine tungsten pin1,2,3,4,5. A tungsten pin is most often used to mount the preparation in a lateral orientation, although it has been used to mount larvae dorsal-side up4. The suture glue has been used to mount embryos and larvae in both orientations. Using the glue, a minimal dissection can be performed, allowing access to spinal neurons without the use of an enzymatic treatment, thereby avoiding any resultant damage. However, for larvae, it is necessary to apply a brief enzyme treatment to remove the muscle tissue surrounding the spinal cord. The methods described here have been used to study the intrinsic electrical properties of motor neurons, interneurons, and sensory neurons at several developmental stages6,7,8,9.

Zebrafish In Situ Spinal Cord Preparation for Electrophysiological Recordings from Spinal Sensory and Motor Neurons

This manuscript describes methods for electrophysiological recordings from spinal neurons of zebrafish embryos and larvae. The preparation maintains neurons in situ and often involves minimum dissection. These methods allow for the electrophysiological study of a variety of spinal neurons, from the initial electrical excitability acquisition through the early larval stages.

Zebrafish, first introduced as a developmental model, have gained popularity in many other fields. The ease of rearing large numbers of rapidly developing ...

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in adult mammalian animals remains elusive. In this study, we describe an experimental procedure for measuring calcium dynamics through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy. This method enables recordings and quantifications of neural activity in bilateral mouse brain regions one at a time in the same experiment for a prolonged period in vivo. Key aspects of this method, which can be completed within an hour, include minimally invasive surgery procedures for creating dual optical windows, and the use of 2P imaging. Although we only demonstrate the technique in the S1 area, the method can be applied to other regions of the living brain facilitating the elucidation of structural and functional complexities of brain neural networks.

Imaging Neural Activity in the Primary Somatosensory Cortex Using Thy1-GCaMP6s Transgenic Mice

We describe an experimental procedure for measuring neuronal activity through dual optical windows above bilateral primary somatosensory corticies (S1) in Thy1-GCaMP6s transgenic mice using 2-photon (2P) microscopy in vivo.

The mammalian brain exhibits marked symmetry across the sagittal plane. However, detailed description of neural dynamics in symmetric brain regions in ...

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic tectum, is a popular model to study how neural circuits self-assemble. The ability to carry out whole cell patch clamp recordings from tectal neurons and to record RGC-evoked responses, either in vivo or using a whole brain preparation, has generated a large body of high-resolution data about the mechanisms underlying normal, and abnormal, circuit formation and function. Here we describe how to perform the in vivo preparation, the original whole brain preparation, and a more recently developed horizontal brain slice preparation for obtaining whole cell patch clamp recordings from tectal neurons. Each preparation has unique experimental advantages. The in vivo preparation enables the recording of the direct response of tectal neurons to visual stimuli projected onto the eye. The whole brain preparation allows for the RGC axons to be activated in a highly controlled manner, and the horizontal brain slice preparation allows recording from across all layers of the tectum.

Preparations and Protocols for Whole Cell Patch Clamp Recording of Xenopus laevis Tectal Neurons

In this paper, we discuss three brain preparations used for whole cell patch clamp recording to study the retinotectal circuit of Xenopus laevis tadpoles. Each preparation, with its own specific advantages, contributes to the experimental tractability of the Xenopus tadpole as a model to study neural circuit function.

The Xenopus tadpole retinotectal circuit, comprised of the retinal ganglion cells (RGCs) in the eye which form synapses directly onto neurons in the optic ...