This work was supported by grants from the National Natural Science Foundation of China (No. 81560198, 31660289).

All experimental protocols described were approved by the Animal Ethics Committee of Nanchang University (Nanchang, PR China, Ethical No.2017-010). All efforts were made to minimize the stress and pain of the experimental animals. The electrophysiological recordings performed here were carried out at room temperature (RT, 22&#8211;25 &#176;C).
1. Animals
<ol>
	<li>Use Sprague-Dawley rats (3&#8211;5 weeks old) of either sex. House the animals under a 12 h light-dark cycle and give them ad libitum access to adequate food and water.</li>
</ol>
2. Preparation of Solutions and Materials
<ol>
	<li>Solutions
	<ol>
		<li>Prepare artificial cerebrospinal fluid (ACSF) (in mM): 117 NaCl, 3.6 KCl, 1.2 NaH2PO4&#183;2H2O, 2.5 CaCl2&#183;2H2O, 1.2 MgCl2&#183;6H2O, 25 NaHCO3, 11 D-glucose, 0.4 ascorbic acid, and 2 sodium pyruvate. See Table 1.</li>
		<li>Prepare sucrose-ACSF (in mM): 240 Sucrose, 2.5 KCl, 1.25 NaH2PO4&#183;2H2O, 0.5 CaCl2&#183;2H2O, 3.5 MgCl2&#183;6H2O, 25 NaHCO3, 0.4 Ascorbic acid, and 2 sodium pyruvate. See Table 2.</li>
		<li>Prepare K+-based intracellular solution (in mM): 130 K-gluconate, 5 KCl, 10 Na2-phosphocreatine, 0.5 EGTA, 10 HEPES, 4 Mg-ATP, and 0.3 Li-GTP. See Table 3.</li>
		<li>Prepare Cs+-based intracellular solution (in mM): 92 CsMeSO4, 43 CsCl, 10 Na2-phosphocreatine, 5 tetraethylammonium (TEA)-Cl, 0.5 EGTA, 10 HEPES, 4 Mg-ATP, and 0.3 Li-GTP. See Table 4. 
		Note: All solutions must be prepared using distilled water. ACSF and sucrose-ACSF should be carbogenated (95% O2 and 5% CO2 mixture) prior to use to maintain an optimal pH of approximately 7.4, and the osmolality of these two solutions should be adjusted to 300&#8211;310 mOsm. Because ascorbic acid could affect calcium channels, this agent must be omitted if one would like to record calcium currents. The osmolality and pH of intracellular solutions should be measured and adjusted to 290&#8211;300 mOsm and 7.2&#8211;7.3, respectively. It is recommended to filter the intracellular solutions with 0.2 &#181;m filters and store the solutions as 1 mL aliquots at -20 &#176;C. Cs+ and TEA are applied in Cs+-based intracellular solution to block potassium channel, which is conducive to using the amplifier to hold the membrane potential steady at 0 mV when recording inhibitory postsynaptic currents (IPSCs).</li>
		<li>Prepare the 0.05% neurobiotin 488 solution. Dissolve 2 mg of neurobiotin 488 in 4 mL of K+-based intracellular solution and adjust the osmolality to 290&#8211;300 mOsm by using distilled water or sucrose if needed. 
		Note: 1 mM of sucrose increases osmolarity by 1 mOsm.</li>
		<li>Prepare 3% agar as a block for spinal cord. Dissolve 7.5 g of agar in 250 mL of purified water in a glass beaker, and then use a microwave to heat it until boiling and clear. Swirl the solution and pour the mixture into a 17.5 cm x 10.5 cm x 1.8 cm plastic box for solidification afterwards. Keep the agar at 4 &#176;C prior to use.</li>
		<li>Prepare 4% paraformaldehyde (PFA) for immunohistochemical processing. Mix 40 g of PFA powder to ~800 mL of heated (approximately 60 &#176;C) 1x PBS solution (130 mM NaCl, 7 mM Na2HPO4, 3 mM NaH2PO4). Slowly adjust the pH by adding 1 N NaOH drops until PFA powder is completely dissolved. Once the solution has become clear, adjust the total volume to 1 L with 1x PBS. Readjust the pH to 7.2&#8211;7.4 using 1 N HCl when needed, then filter the 4% PFA solution and store it at -20 &#176;C until use. 
		Note: PFA is toxic, so it is needed to wear masks, gloves as well as safety glasses. Conduct the process of preparing 4% PFA inside a ventilated hood. PFA powder can be completely dissolved at a pH of approximately 9&#8211;10.</li>
	</ol>
	</li>
	<li>Instruments
	<ol>
		<li>For a typical electrophysiological system, use an upright microscope equipped with infrared differential interference contrast (IR-DIC) and a high-resolution water-immersion objective, a CCD/CMOS camera, a patch-clamp amplifier, a micropipette holder and a micromanipulator allowing fine adjustment of the pipette position. An XY stage is also needed to move the microscope.</li>
		<li>Mount all equipment on a vibration isolation table surrounded by a Faraday cage. Connect a video monitor to the video camera to observe the neurons and visualize the micropipettes.</li>
	</ol>
	</li>
	<li>Micropipettes
	<ol>
		<li>Make recording electrodes from borosilicate glass capillaries using a micropipette puller. The typical pipette resistance ranges from 3-6 M&#937; when filled with intracellular solution.</li>
	</ol>
	</li>
	<li>Agar block
	<ol>
		<li>Prepare a 1.2 cm x 1.5 cm x 2.0 cm agar block. Trim the block into the shapes shown in Figure 1 as required.</li>
	</ol>
	</li>
</ol>
3. Acute Spinal Cord Slice Preparation
Note: Transverse or parasagittal spinal cord slices are prepared as previously described18,19,20.
<ol>
	<li>Prior to transcardial perfusion and spinal cord extraction, prepare ~500 mL sucrose-ACSF equilibrated with 95% O2 and 5% CO2 and cool the solution to ice-cold (0&#8211;4 &#176;C).</li>
	<li>Cool down all the dissection tools (e.g., dissecting scissors, iris scissors, toothed forceps, fine forceps, curved forceps) on the ice. The diagram of the preparation of acute spinal cord slice is shown in Figure 1.</li>
	<li>Transcardial perfusion
	<ol>
		<li>After a single injection (intraperitoneal, i.p.) of urethane (1.5 g/kg), wait for 2&#8211;3 min, and assess the anesthesia depth of rats by testing toe or tail pinch responses. Once a surgical plane of anesthesia is maintained, place the rat on crushed ice in the supine position. 
		Note:&#160;For producing profound anesthesia sufficient for non-perception of pain during thoracotomy, follow your local IACUC guidelines.</li>
		<li>Make an incision (3&#8211;4 cm) through the skin from the xiphoid process to the clavicle, and then make a transverse incision beneath the level of the xiphoid process.</li>
		<li>Grasp and raise the xiphoid process with a pair of curved forceps to expose the diaphragm completely. Make a transverse incision through the diaphragm, and then cut through the ribcage between the breastbone and ribs bilaterally to open the chest cavity. Use the curved forceps to grasp the xiphoid process so the ribcage is fixed, and the heart is sufficiently exposed.</li>
		<li>Hold the heart with another curved forceps gently, and then insert a 22 G needle through the left ventricle to the base of the aortic arch.</li>
		<li>Cut the right atrium with fine scissors immediately, and then start the perfusion of ice-cold, oxygenated sucrose-ACSF through a gravity system. 
		Note: Rapid and sufficient transcardial perfusion with ice-cold sucrose-ACSF can facilitate the rapid cooling of the spinal cord, while low sodium and calcium concentration may help alleviate excitatory toxicity and protect neuronal function. In addition, clearing red blood cells would be beneficial for reducing the background staining of biocytin when performing a morphological study. The perfusion is considered sufficient and satisfied as long as the fluid exiting the right atrium is clear and the color of the rat&#8217;s liver and paws is pale. The tip of the 22 G needle should be blunt to avoid rupturing the heart and aortic arch.</li>
	</ol>
	</li>
	<li>Spinal cord dissection and slicing
	<ol>
		<li>Make a longitudinal incision (5 cm) on the back skin from caudal to rostral, then cut through the ribcage between the spine and ribs on either side in the state of perfusion.</li>
		<li>Make a cut at the caudal end of the spine, use scissors to cut away surrounding tissues, and isolate the lumbosacral segment of the spine rapidly.</li>
		<li>Transfer the lumbosacral segment to a glass dish containing ice-cold sucrose-based ACSF. With the ventral side upward, use fine scissors to cut through the vertebral pedicle bilaterally and expose the spinal cord carefully. Isolate a 2-cm long spinal cord with lumbosacral enlargement (L1&#8211;S3) and transfer the spinal section to another glass dish filled with cold sucrose-ACSF.</li>
		<li>Remove the meninges and the pia-arachnoid membrane under a dissecting microscope. Cut all the ventral and dorsal roots away as quickly as possible.</li>
		<li>Place the spinal cord on a previously trimmed agar block. To prepare transverse slices, attach the ventral side of the spinal cord to the agar and let the dorsal side toward the blade. To prepare parasagittal slices, attach the ventral side with superglue to the agar in a vertical direction as shown in Figure 1. Then, mount the agar block to a platform of a vibratome with superglue. Prepare 300&#8211;500 &#181;m transverse or parasagittal slices with an advance speed of 0.025 mm/s and a vibration frequency of 80 Hz.</li>
		<li>Use a plastic-trimmed pipette to transfer the slices onto nylon mesh in a storage chamber containing continuously oxygenated ACSF at 32 &#176;C for at least 30 min prior to recording. 
		Note: Take care to avoid injury to the spinal cord, especially the dorsal horn, when removing the meninges and spinal roots. The spinal cord should be sliced dorsal-ventrally. For the best results, the slices should be prepared rapidly (within 15&#8211;20 min). The thickness of a spinal slice should be no more than 600 &#181;m to satisfy cell visibility. In addition, the technique described above could be used to obtain horizontal spinal slices.</li>
	</ol>
	</li>
</ol>
4. Whole-cell Patch-clamp Recordings
<ol>
	<li>To conduct the whole-cell patch-clamp recordings from SG neurons, use K+-based intracellular solution for most recording cases, while applying Cs+-based solution only for the recording of inhibitory postsynaptic currents.</li>
	<li>Gently move a spinal cord slice to the recording chamber, and then maintain it with a U-shaped platinum wire attached with nylon threads firmly for optimal slice stability. Steadily perfuse the slice with bubbled ACSF at RT through a gravity system and set the perfusion rate at 2&#8211;4 mL/min to achieve sufficient oxygenation.</li>
	<li>Identify the region of SG (a translucent band) using a low-resolution microscope lens, choose a healthy neuron by using the high-resolution objective as the target cell, and adjust it to the center of the video monitor screen.</li>
	<li>Fill a micropipette with an appropriate volume of K+-based or Cs+-based intracellular solution as needed, insert the micropipette into the electrode holder, and ensure that the intracellular solution is contacting the silver wire inside the holder.</li>
	<li>Bring the micropipette into focus and immerse it into the ACSF using a micromanipulator, and then apply a mild positive pressure (~1 psi when measured with a manometer) to force the micropipette away from any dirt and debris.</li>
	<li>Move the micropipette towards the targeted neuron gradually. Release the positive pressure once the pipette approaches the neuron and a very small dimple forms on neuronal membrane to form a gigaseal.</li>
	<li>Alter the holding potential to -70 mV, which is close to the physiological resting membrane potential (RMP) of a cell. Then, apply a transient and gentle suction to the micropipette to rupture the membrane and create a good whole-cell configuration. 
	Note: After transferring the slice into the recording chamber, ensure steady perfusion for at least 5 min to clear the debris on the slice surface. It is worth noting that the ability to distinguish between healthy and unhealthy/dead neurons is of paramount importance for good sealing and stable recording. An unhealthy/dead neuron has a swollen or shrunken appearance, together with a visible large nucleus, while a healthy neuron is characterized by a 3-dimensional (3D) shape with a bright and smooth membrane, and its nucleus is invisible. To achieve a whole-cell configuration, it is essential to compensate for fast or slow capacitance step-by-step when necessary. At RT, the liquid junction potential is calculated to be 15.1&#8211;15.2 mV and 4.3&#8211;4.4 mV in K+-based and Cs+-based intracellular solution, respectively. In our studies, the recorded data were not corrected for liquid junction potential.</li>
	<li>Recordings of intrinsic membrane properties
	<ol>
		<li>Record passive intrinsic membrane properties: Record RMP immediately (within 20 s) after break in. Determine the neuronal input resistance by measuring the voltage response to a depolarizing current (10 pA, 500 ms) at RMP in current-clamp mode.</li>
		<li>Record firing properties: Test the firing pattern of each neuron in current-clamp with a series of 1 s depolarizing current pulses (25&#8211;150 pA with 25 pA increment) at RMP. Measure the threshold, amplitude and half-width of a single action potential offline.</li>
		<li>Record subthreshold current: To assess the somatic subthreshold currents, hold the membrane potential at -50 mV in voltage-clamp mode. Then, apply a series of hyperpolarizing voltage pulses of 1 s duration from -60 mV to -120 mV, with a 10-mV decrement.</li>
		<li>Record excitatory postsynaptic currents (EPSCs): Record EPSCs with the K+-based intracellular solution in voltage-clamp mode at a holding potential of -70 mV.</li>
		<li>Record IPSCs: Apply Cs+-based intracellular solution for recording IPSCs. Once the whole-cell configuration is established, hold the membrane potential at -70 mV for ~5 min, and then change the holding potential to 0 mV gradually. Wait a few minutes for stabilization, and then start to record the IPSC events. 
		Note: Only neurons with an RMP less than -50 mV and showing overshoot should be selected for further study. The series resistances in our study are typically 10-30 M&#937;, and a recording should be excluded once the series resistance changes by more than 20%. EPSCs could be confirmed with a bath application of 50 &#181;M APV and 20 &#181;M CNQX, while IPSCs could be confirmed with 10 &#181;M bicuculline and 1 &#181;M strychnine.</li>
	</ol>
	</li>
</ol>
5. Morphological study
<ol>
	<li>For morphological experiments, use a K+-based intracellular solution containing 0.05% neurobiotin 488.</li>
	<li>After maintaining a stable electrophysiological recording for at least 20 min, slowly remove the micropipette in the upward direction to allow the cell membrane to reseal and transfer the spinal cord slice to a container filled with 4% PFA. Fix the slices in 4% PFA at RT for 1 h and then at 4&#176;C overnight.</li>
	<li>Rinse the slices in PBS, and then immerse them in 50% ethanol for 30 min. After another three washes in PBS, mount the slices in their original thickness onto slides with a mounting medium.</li>
	<li>Use a confocal microscope (see Table of Materials) for image acquisition and neuronal 3D reconstruction. Scan neurons through a 20X lens with a z-stack of 1.5 &#181;m.</li>
</ol>

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E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Use+of+ascorbate+in+the+preparation+and+maintenance+of+brain+slices.">Use of ascorbate in the preparation and maintenance of brain slices.</a> Methods. 18 (2), 144-149 (1999).</li><li>Takasu, K., Ogawa, K., Minami, K., Shinohara, S., Kato, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Injury-specific+functional+alteration+of+N-type+voltage-gated+calcium+channels+in+synaptic+transmission+of+primary+afferent+C-fibers+in+the+rat+spinal+superficial+dorsal+horn.">Injury-specific functional alteration of N-type voltage-gated calcium channels in synaptic transmission of primary afferent C-fibers in the rat spinal superficial dorsal horn.</a> European Journal of Pharmacology. 772, 11-21 (2016).</li><li>Tian, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Excitatory+synaptic+transmission+in+the+spinal+substantia+gelatinosa+is+under+an+inhibitory+tone+of+endogenous+adenosine.">Excitatory synaptic transmission in the spinal substantia gelatinosa is under an inhibitory tone of endogenous adenosine.</a> Neuroscience Letters. 477 (1), 28-32 (2010).</li><li>Funai, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Systemic+dexmedetomidine+augments+inhibitory+synaptic+transmission+in+the+superficial+dorsal+horn+through+activation+of+descending+noradrenergic+control:+an+in+vivo+patch-clamp+analysis+of+analgesic+mechanisms.">Systemic dexmedetomidine augments inhibitory synaptic transmission in the superficial dorsal horn through activation of descending noradrenergic control: an in vivo patch-clamp analysis of analgesic mechanisms.</a> Pain. 155 (3), 617-628 (2014).</li><li>Yamasaki, H., Funai, Y., Funao, T., Mori, T., Nishikawa, K. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Effects+of+tramadol+on+substantia+gelatinosa+neurons+in+the+rat+spinal+cord:+an+in+vivo+patch-clamp+analysis.">Effects of tramadol on substantia gelatinosa neurons in the rat spinal cord: an in vivo patch-clamp analysis.</a> PLoS One. 10 (5), e0125147 (2015).</li><li>Furue, H., Narikawa, K., Kumamoto, E., Yoshimura, M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Responsiveness+of+rat+substantia+gelatinosa+neurones+to+mechanical+but+not+thermal+stimuli+revealed+by+in+vivo+patch-clamp+recording.">Responsiveness of rat substantia gelatinosa neurones to mechanical but not thermal stimuli revealed by in vivo patch-clamp recording.</a> The Journal of Physiology. 521 (Pt 2), 529-535 (1999).</li><li>Ting, J. T., Daigle, T. L., Chen, Q., Feng, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Acute+brain+slice+methods+for+adult+and+aging+animals:+application+of+targeted+patch+clamp+analysis+and+optogenetics.">Acute brain slice methods for adult and aging animals: application of targeted patch clamp analysis and optogenetics.</a> Methods in Molecular Biology. 1183, 221-242 (2014).</li><li>Ting, J. T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Preparation+of+Acute+Brain+Slices+Using+an+Optimized+N-Methyl-D-glucamine+Protective+Recovery+Method.">Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method.</a> Journal of Visualized Experiments. (132), e53825 (2018).</li><li>Li, J., Baccei, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Neonatal+Tissue+Damage+Promotes+Spike+Timing-Dependent+Synaptic+Long-Term+Potentiation+in+Adult+Spinal+Projection+Neurons.">Neonatal Tissue Damage Promotes Spike Timing-Dependent Synaptic Long-Term Potentiation in Adult Spinal Projection Neurons.</a> The Journal of Neuroscience. 36 (19), 5405-5416 (2016).</li><li>Ford, N. C., Ren, D., Baccei, M. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=NALCN+channels+enhance+the+intrinsic+excitability+of+spinal+projection+neurons.">NALCN channels enhance the intrinsic excitability of spinal projection neurons.</a> Pain. , (2018).</li><li>Cui, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Modulation+of+synaptic+transmission+from+primary+afferents+to+spinal+substantia+gelatinosa+neurons+by+group+III+mGluRs+in+GAD65-EGFP+transgenic+mice.">Modulation of synaptic transmission from primary afferents to spinal substantia gelatinosa neurons by group III mGluRs in GAD65-EGFP transgenic mice.</a> Journal of Neurophysiology. 105 (3), 1102-1111 (2011).</li><li>Yang, K., Ma, R., Wang, Q., Jiang, P., Li, Y. Q. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Optoactivation+of+parvalbumin+neurons+in+the+spinal+dorsal+horn+evokes+GABA+release+that+is+regulated+by+presynaptic+GABAB+receptors.">Optoactivation of parvalbumin neurons in the spinal dorsal horn evokes GABA release that is regulated by presynaptic GABAB receptors.</a> Neuroscience Letters. , 55-59 (2015).</li></ol>

The authors declare no conflicts of interest.

This protocol details the steps for preparing spinal cord slices, which we have used successfully when performing whole-cell patch-clamp experiments on SG neurons18,19,20,21. By implementing this method, we recently reported that minocycline, a second generation of tetracycline, could markedly enhance inhibitory synaptic transmission through a presynaptic mechanism in SG neurons<sup class="xref...

The substantia gelatinosa (SG, lamina II of the spinal dorsal horn) is an indisputably important relay center for transmitting and regulating sensory information. It is composed of excitatory and inhibitory interneurons, which receive inputs from the primary afferent fibers, local interneurons, and the endogenous descending inhibitory system1. In recent decades, the development of acute spinal cord slice preparation and the advent of whole-cell patch-clamp recording have enabled various studies on the intrinsic electrophysiological and morphological properties of SG neurons2,3,4 as well as studies of the local circuitry in SG5,6. In addition, by using the in vitro spinal cord slice preparation, researchers can interpret the changes in neuronal excitabilities7,8, the function of ion channels9,10, and synaptic activities11,12 under various pathological conditions. These studies have deepened our understanding of the role that SG neurons play in the development and maintenance of chronic pain and neuropathic itch.
Essentially, the key prerequisite to achieve a clear visualization of neuronal soma and ideal whole-cell patching using acute spinal cord slices is to ensure the excellent quality of slices so healthy and patchable neurons can be obtained. However, preparing spinal cord slices involves several steps, such as performing a ventral laminectomy and removing the pia-arachnoid membrane, which may be obstacles in obtaining healthy slices. Although it is not easy to prepare spinal cord slices, performing recordings in vitro on spinal cord slices has several advantages. Compared to cell culture preparations, spinal cord slices can partially preserve inherent synaptic connections that are in a physiologically relevant condition. In addition, whole-cell patch-clamp recording using spinal cord slices could be combined with other techniques, such as double patch clamp13,14, morphological studies15,16 and single-cell RT-PCR17. Therefore, this technique provides more information on characterizing the anatomical and genetic diversities within a specific region and allows for investigation of the composition of local circuitry.
Here, we provide a basic and detailed description of our method for preparing acute spinal cord slices and acquiring whole-cell patch-clamp recordings from SG neurons.

<table><tbody><tr><td>NaCl</td><td>Sigma</td><td>S7653</td><td>Used for the preparation of ACSF and PBS</td></tr><tr><td>KCl</td><td>Sigma</td><td>60130</td><td>Used for the preparation of ACSF, sucrose-ACSF, and K&lt;sup&gt;+&lt;/sup&gt;-based intracellular solution</td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma</td><td>71500</td><td>Used for the preparation of ACSF, sucrose-ACSF and PBS</td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma</td><td>C5080</td><td>Used for the preparation of ACSF and sucrose-ACSF</td></tr><tr><td>MgCl&lt;sub&gt;2&lt;/sub&gt;&amp;middot;6H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma</td><td>M2670</td><td>Used for the preparation of ACSF and sucrose-ACSF</td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Sigma</td><td>S5761</td><td>Used for the preparation of ACSF and sucrose-ACSF</td></tr><tr><td>D-Glucose</td><td>Sigma</td><td>G7021</td><td>Used for the preparation of ACSF</td></tr><tr><td>Ascorbic acid</td><td>Sigma</td><td>P5280</td><td>Used for the preparation of ACSF and sucrose-ACSF</td></tr><tr><td>Sodium pyruvate</td><td>Sigma</td><td>A7631</td><td>Used for the preparation of ACSF and sucrose-ACSF</td></tr><tr><td>Sucrose</td><td>Sigma</td><td>S7903</td><td>Used for the preparation of sucrose-ACSF</td></tr><tr><td>K-gluconate</td><td>Wako</td><td>169-11835</td><td>Used for the preparation of K&lt;sup&gt;+&lt;/sup&gt;-based intracellular solution</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;-Phosphocreatine</td><td>Sigma</td><td>P1937</td><td>Used for the preparation of intracellular solution</td></tr><tr><td>EGTA</td><td>Sigma</td><td>E3889</td><td>Used for the preparation of intracellular solution</td></tr><tr><td>HEPES</td><td>Sigma</td><td>H4034</td><td>Used for the preparation of intracellular solution</td></tr><tr><td>Mg-ATP</td><td>Sigma</td><td>A9187</td><td>Used for the preparation of intracellular solution</td></tr><tr><td>Li-GTP</td><td>Sigma</td><td>G5884</td><td>Used for the preparation of intracellular solution</td></tr><tr><td>CsMeSO&lt;sub&gt;4&lt;/sub&gt;</td><td>Sigma</td><td>C1426</td><td>Used for the preparation of Cs&lt;sup&gt;+&lt;/sup&gt;-based intracellular solution</td></tr><tr><td>CsCl</td><td>Sigma</td><td>C3011</td><td>Used for the preparation of Cs&lt;sup&gt;+&lt;/sup&gt;-based intracellular solution</td></tr><tr><td>TEA-Cl</td><td>Sigma</td><td>T2265</td><td>Used for the preparation of Cs&lt;sup&gt;+&lt;/sup&gt;-based intracellular solution</td></tr><tr><td>Neurobiotin 488</td><td>Vector</td><td>SP-1145</td><td>0.05% neurobiotin 488 could be used for morphological studies</td></tr><tr><td>Agar</td><td>Sigma</td><td>A7002</td><td>3% agar block was used in our protocol</td></tr><tr><td>Paraformaldehyde</td><td>Sigma</td><td>P6148</td><td>4% paraformaldehyde was used for immunohistochemical processing</td></tr><tr><td>Na&lt;sub&gt;2&lt;/sub&gt;HPO&lt;sub&gt;4&lt;/sub&gt;</td><td>Hengxing Chemical Reagents</td><td></td><td>Used for the preparation of PBS</td></tr><tr><td>Mount Coverslipping Medium</td><td>Polyscience</td><td>18606</td><td></td></tr><tr><td>Urethan</td><td>National Institute for Food and Drug Control</td><td>30191228</td><td>1.5 g/kg, i.p.</td></tr><tr><td>Borosilicate glass capillaries</td><td>World Precision Instruments</td><td>TW150F-4</td><td>1.5 mm OD, 1.12 mm ID</td></tr><tr><td>Micropipette puller</td><td>Sutter Instrument</td><td>P-97</td><td>Used for the preparation of micropipettes</td></tr><tr><td>Vibratome</td><td>Leica</td><td>VT1000S</td><td></td></tr><tr><td>Vibration isolation table</td><td>Technical Manufacturing Corporation</td><td>63544</td><td></td></tr><tr><td>Infrared CCD camera</td><td>Dage-MIT</td><td>IR-1000</td><td></td></tr><tr><td>Patch-clamp amplifier</td><td>HEKA</td><td>EPC-10</td><td></td></tr><tr><td>Micromanipulator</td><td>Sutter Instrument</td><td>MP-285</td><td></td></tr><tr><td>X-Y stage</td><td>Burleigh</td><td>GIBRALTAR X-Y</td><td></td></tr><tr><td>Upright microscope</td><td>Olympus</td><td>BX51WI</td><td></td></tr><tr><td>Osmometer</td><td>Advanced</td><td>FISKE 210</td><td></td></tr><tr><td>PH meter</td><td>Mettler Toledo</td><td>FE20</td><td></td></tr><tr><td>Confocol microscope</td><td>Zeiss</td><td>LSM 700</td><td></td></tr></tbody></table>

preparation of acute spinal cord slices for whole-cell patch-clamp recording in substantia gelatinosa neurons

Recent whole-cell patch-clamp studies from substantia gelatinosa (SG) neurons have provided a large body of information about the spinal mechanisms underlying sensory transmission, nociceptive regulation, and chronic pain or itch development. Implementations of electrophysiological recordings together with morphological studies based on the utility of acute spinal cord slices have further improved our understanding of neuronal properties and the composition of local circuitry in SG. Here, we present a detailed and practical guide for the preparation of spinal cord slices and show representative whole-cell recording and morphological results. This protocol permits ideal neuronal preservation and can mimic in vivo conditions to a certain extent. In summary, the ability to obtain an in vitro preparation of spinal cord slices enables stable current- and voltage-clamp recordings and could thus facilitate detailed investigations into the intrinsic membrane properties, local circuitry and neuronal structure using diverse experimental approaches.

Acute spinal cord slices were prepared according to the diagram shown in Figure 1. After slicing and recovery, a spinal cord slice was transferred to the recording chamber. Healthy neurons were identified based on soma appearance using IR-DIC microscopy. Next, the action potentials of SG neurons were elicited by a series of depolarizing current pulses (1 s duration) when neurons were held at RMP. As shown in Figure 2, the firing ...

Watch this Scientific Journal Video about Preparation of Acute Spinal Cord Slices for Whole-cell Patch-clamp Recording in Substantia Gelatinosa Neurons at JoVE.com

Preparation of Acute Spinal Cord Slices for Whole-cell Patch-clamp Recording in Substantia Gelatinosa Neurons

Here, we describe the essential steps for whole-cell patch-clamp recordings made from substantia gelatinosa (SG) neurons in the in vitro spinal cord slice. This method allows the intrinsic membrane properties, synaptic transmission and morphological characterization of SG neurons to be studied.

Recent whole-cell patch-clamp studies from substantia gelatinosa (SG) neurons have provided a large body of information about the spinal mechanisms ...

preparation-acute-spinal-cord-slices-for-whole-cell-patch-clamp

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Neuroscience

13.9K Views.  First Affiliated Hospital of Nanchang University. Deze methode kan helpen verduidelijken van de spinale mechanismen die ten grondslag liggen aan sensorische transmissie, nociceptieve regelgeving en chronische pijn of pijn ontwikkeling Het belangrijkste voordeel van deze techniek is dat het kan toestaan dat ideale neuronale bewaring en het kan nabootsen in vivo voorwaarden tot op zekere hoogte. Begin met het opnemen van het opnemen van elektroden van borosilicaat glazen microcapillaries met behulp van een haarvatentrekker. Vul vervolgens een ...

Video: Preparation of Acute Spinal Cord Slices for Whole-cell Patch-clamp Recording in Substantia Gelatinosa Neurons

Acute Dissociation of Lamprey Reticulospinal Axons to Enable Recording from the Release Face Membrane of Individual Functional Presynaptic Terminals

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated calcium channels (VGCCs) located in the release face membrane, is the trigger for vesicle fusion and neurotransmitter release. Crucial to the rapidity of synaptic transmission is the spatial and temporal synchrony between the arrival of the action potential, VGCCs and the neurotransmitter release machinery. The ability to directly record Ca2+&#160;currents from the release face membrane of individual presynaptic terminals is imperative for a precise understanding of the relationship between presynaptic Ca2+&#160;and neurotransmitter release. Access to the presynaptic release face membrane for electrophysiological recording is not available in most preparations and presynaptic Ca2+&#160;entry has been characterized using imaging techniques and macroscopic current measurements &#8211; techniques that do not have sufficient temporal resolution to visualize Ca2+ entry. The characterization of VGCCs directly at single presynaptic terminals has not been possible in central synapses and has thus far been successfully achieved only in the calyx-type synapse of the chick ciliary ganglion and in rat calyces. We have successfully addressed this problem in the giant reticulospinal synapse of the lamprey spinal cord by developing an acutely dissociated preparation of the spinal cord that yields isolated reticulospinal axons with functional presynaptic terminals devoid of postsynaptic structures. We can fluorescently label and identify individual presynaptic terminals and target them for recording. Using this preparation, we have characterized VGCCs directly at the release face of individual presynaptic terminals using immunohistochemistry and electrophysiology approaches. Ca2+ currents have been recorded directly at the release face membrane of individual presynaptic terminals, the first such recording to be carried out at central synapses.

Recording Ca2+ currents at the presynaptic release face membrane is key to a precise understanding of Ca2+ entry and neurotransmitter release. We present an acute dissociation of the lamprey spinal cord that yields functional isolated reticulospinal axons, permitting recording directly from the release face membrane of individual presynaptic terminals.

Synaptic transmission is an extremely rapid process. Action potential driven influx of Ca2+&#160;into the presynaptic terminal, through voltage-gated ...

Electrophysiological and Morphological Characterization of Neuronal Microcircuits in Acute Brain Slices Using Paired Patch-Clamp Recordings

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with simultaneous intracellular biocytin filling allows a correlated analysis of their structural and functional properties. With this method it is possible to identify and characterize both pre- and postsynaptic neurons by their morphology and electrophysiological response pattern. Paired recordings allow studying the connectivity patterns between these neurons as well as the properties of both chemical and electrical synaptic transmission. Here, we give a step-by-step description of the procedures required to obtain reliable paired recordings together with an optimal recovery of the neuron morphology. We will describe how pairs of neurons connected via chemical synapses or gap junctions are identified in brain slice preparations. We will outline how neurons are reconstructed to obtain their 3D morphology of the dendritic and axonal domain and how synaptic contacts are identified and localized. We will also discuss the caveats and limitations of the paired recording technique, in particular those associated with dendritic and axonal truncations during the preparation of brain slices because these strongly affect connectivity estimates. However, because of the versatility of the paired recording approach it will remain a valuable tool in characterizing different aspects of synaptic transmission at identified neuronal microcircuits in the brain.

Patch-clamp recordings and simultaneous intracellular biocytin filling of synaptically coupled neurons in acute brain slices allow a correlated analysis of their structural and functional properties. The aim of this protocol is to describe the essential technical steps of electrophysiological recording from neuronal microcircuits and their subsequent morphological analysis.

The combination of patch clamp recordings from two (or more) synaptically coupled neurons (paired recordings) in acute brain slice preparations with ...

Optical Control of a Neuronal Protein Using a Genetically Encoded Unnatural Amino Acid in Neurons

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to photo-regulate endogenous proteins expressed in their native environment. Here, we present an approach to optically control the function of a neuronal protein directly in neurons using a genetically encoded unnatural amino acid (Uaa). By using an orthogonal tRNA/aminoacyl-tRNA synthetase pair to suppress the amber codon, a photo-reactive Uaa 4,5-dimethoxy-2-nitrobenzyl-cysteine (Cmn) is site-specifically incorporated in the pore of a neuronal protein Kir2.1, an inwardly rectifying potassium channel. The bulky Cmn physically blocks the channel pore, rendering Kir2.1 non-conducting. Light illumination instantaneously converts Cmn into a smaller natural amino acid Cys, activating Kir2.1 channel function. We express these photo-inducible inwardly rectifying potassium (PIRK) channels in rat hippocampal primary neurons, and demonstrate that light-activation of PIRK ceases the neuronal firing due to the outflux of K+ current through the activated Kir2.1 channels. Using in utero electroporation, we also express PIRK in the embryonic mouse neocortex in vivo, showing the light-activation of PIRK in neocortical neurons. Genetically encoding Uaa imposes no restrictions on target protein type or cellular location, and a family of photoreactive Uaas is available for modulating different natural amino acid residues. This technique thus has the potential to be generally applied to many neuronal proteins to achieve optical regulation of different processes in brains. The current protocol presents an accessible procedure for intricate Uaa incorporation in neurons in vitro and in vivo to achieve photo control of neuronal protein activity on the molecular level.

Here, a procedure to selectively activate a neuronal protein with a short pulse of light by genetically encoding a photo-reactive unnatural amino acid into a target neuronal protein expressed in neurons in culture or in vivo is presented.

Photostimulation is a noninvasive way to control biological events with excellent spatial and temporal resolution. New methods are desired to ...

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have validated the utility of this method for enhancing neuronal preservation and overall brain slice viability. The implementation of this technique by early adopters has facilitated detailed investigations into brain function using diverse experimental applications and spanning a wide range of animal ages, brain regions, and cell types. Steps are outlined for carrying out the protective recovery brain slice technique using an optimized NMDG artificial cerebrospinal fluid (aCSF) media formulation and enhanced procedure to reliably obtain healthy brain slices for patch clamp electrophysiology. With this updated approach, a substantial improvement is observed in the speed and reliability of gigaohm seal formation during targeted patch clamp recording experiments while maintaining excellent neuronal preservation, thereby facilitating challenging experimental applications. Representative results are provided from multi-neuron patch clamp recording experiments to assay synaptic connectivity in neocortical brain slices prepared from young adult transgenic mice and mature adult human neurosurgical specimens. Furthermore, the optimized NMDG protective recovery method of brain slicing is compatible with both juvenile and adult animals, thus resolving a limitation of the original methodology. In summary, a single media formulation and brain slicing procedure can be implemented across various species and ages to achieve excellent viability and tissue preservation.

Preparation of Acute Brain Slices Using an Optimized N-Methyl-D-glucamine Protective Recovery Method

This protocol demonstrates the implementation of an optimized N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. A single media formulation is used to reliably obtain healthy brain slices from animals of any age and for diverse experimental applications.

This protocol is a practical guide to the N-methyl-D-glucamine (NMDG) protective recovery method of brain slice preparation. Numerous recent studies have ...

Whole-cell Patch-clamp Recordings in Brain Slices

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the neuron. In this configuration, the micropipette is in tight contact with the cell membrane, which prevents current leakage and thereby provides more accurate ionic current measurements than the previously used intracellular sharp electrode recording method. Classically, whole-cell recording can be performed on neurons in various types of preparations, including cell culture models, dissociated neurons, neurons in brain slices, and in intact anesthetized or awake animals. In summary, this technique has immensely contributed to the understanding of passive and active biophysical properties of excitable cells. A major advantage of this technique is that it provides information on how specific manipulations (e.g., pharmacological, experimenter-induced plasticity) may alter specific neuronal functions or channels in real-time. Additionally, significant opening of the plasma membrane allows the internal pipette solution to freely diffuse into the cytoplasm, providing means for introducing drugs, e.g., agonists or antagonists of specific intracellular proteins, and manipulating these targets without altering their functions in neighboring cells. This article will focus on whole-cell recording performed on neurons in brain slices,&#160;a preparation that has the advantage of recording neurons in relatively well preserved brain circuits, i.e., in a physiologically relevant context. In particular,&#160;when combined with appropriate pharmacology, this technique is a powerful tool allowing identification of specific neuroadaptations that occurred following any type of experiences, such as learning, exposure to drugs of abuse, and stress. In summary, whole-cell patch-clamp recordings in brain slices provide means to measure in ex vivo preparation long-lasting changes in neuronal functions that have developed in intact awake animals.

This protocol describes basic procedural steps for performing whole-cell patch-clamp recordings. This technique allows the study of&#160;the electrical behavior of neurons, and when performed in brain slices, allows the assessment of various neuronal functions from neurons that are still integrated in relatively well preserved brain circuits.

Whole-cell patch-clamp recording is an electrophysiological technique that allows the study of the&#160;electrical properties of a substantial part of the ...

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 ...

Subcellular Patch-clamp Recordings from the Somatodendritic Domain of Nigral Dopamine Neurons

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding these fundamental functions will shed light on the information transfer in these neurons. Dendritic patch-clamp recordings provide the possibility to directly examine the electrical properties of dendrites and underlying voltage-gated ion channels. However, these fine structures are not easily accessible to patch pipettes because of their small diameter. This report describes a step-by-step procedure to collect stable and reliable recordings from the dendrites of dopaminergic neurons in acute slices. Electrophysiological measurements are combined with post hoc recovery of cell morphology. Successful experiments rely on improved preparation of slices, solutions and pipettes, adequate adjustment of the optics and stability of the pipette in contact with the recorded structure. Standard principles of somatic patch-clamp recording are applied to dendrites but with a gentler approach of the pipette. These versatile techniques can be implemented to address various questions concerning the excitable properties of dendrites.

Subcellular patch-clamp recordings offer the possibility to investigate the functional properties of dendrites. However these fine structures are not easily accessible due to their small diameter. The protocol described here aims to facilitate the collection of stable and reliable recordings from the dendrites of dopamine neurons in slices in vitro.

Dendrites of dopaminergic neurons receive and convey synaptic input, support action potential back-propagation and neurotransmitter release. Understanding ...

Preparation of Acute Slices from Dorsal Hippocampus for Whole-Cell Recording and Neuronal Reconstruction in the Dentate Gyrus of Adult Mice

Although the general architecture of the hippocampus is similar along its longitudinal axis, recent studies have revealed prominent differences in molecular, anatomical and functional criteria suggesting a division into different sub-circuits along its rostro-caudal extent. Owing to differential connectivity and function the most fundamental distinction is made between the dorsal and the ventral hippocampus, which are preferentially involved in spatial and emotional processing, respectively. Accordingly, in vivo work regarding spatial memory formation has focused on the dorsal hippocampus.
In contrast, electro-physiological in vitro recordings have been preferentially performed on intermediate-ventral hippocampus, largely motivated by factors like slice viability and circuit integrity. To allow for direct correlation of in vivo data on spatial processing with in vitro data we have adapted previous sectioning methods to obtain highly viable transverse brain slices from the dorsal-intermediate hippocampus for long-term recordings of principal cells and interneurons in the dentate gyrus. As spatial behavior is routinely analyzed in adult mice, we have combined this transversal slicing procedure with the use of protective solutions to enhance viability of brain tissue from mature animals. We use this approach for mice of about 3 months of age. The method offers a good alternative to the coronal preparation which is frequently used for in vitro studies on dorsal hippocampus. We compare these two preparations in terms of quality of recordings and preservation of morphological features of recorded neurons.

We present a protocol to prepare acute-slices from the dorsal-intermediate hippocampus of mice. We compare this transversal preparation with coronal slicing in terms of quality of recordings and preservation of morphological features of recorded neurons.

Although the general architecture of the hippocampus is similar along its longitudinal axis, recent studies have revealed prominent differences in ...

Understanding Motor Neuron Responses to Spinal Cord Stimulation

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.

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.

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

Patch-Clamp Recordings from the Dendrite of a Dopaminergic Neuron in a Brain Slice

Source: Engel, D. <a href="https://app.jove.com/v/54601">Subcellular Patch-clamp Recordings from the Somatodendritic Domain of Nigral Dopamine Neurons.</a> J. Vis. Exp. (2016).
This video demonstrates a technique for patch-clamp recording from the dendritic membrane surface. This helps to directly examine the electrical properties of dendrites and underlying voltage-gated ion channels in dopaminergic neurons.

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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Preparation of Acute Spinal Cord Slices for Whole-cell Patch-clamp Recording in Substantia Gelatinosa Neurons

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