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 neurons19. In addition, this agent could decrease the amplitude of Ih and further inhibit the excitability of SG neurons21. In support of these published data and the representative results that we show here, the currently described method is suitable for use in a wide range of electrophysiological studies.
As we noted previously, transcardial perfusion is a crucial element for obtaining healthy specimens. First, we use ice-cold solution for perfusion so the spinal cord can be rapidly cooled and the neuronal metabolism can be slowed22. Second, sucrose-substituted ACSF, a &#39;protective cutting&#39; solution with low Na+ concentration, can ameliorate passive Na+ influx and thus decrease neuronal edema through water entry23. Third, it is beneficial to obtain and analyze neuronal morphology because perfusion could minimize the background caused by biocytin22. For successful preparations, it is also important to use some antioxidants to reduce oxidative damage, which allows neuronal preservation24. Hence, in our protocol, we supplement ascorbic acid and sodium pyruvate, which are powerful antioxidants and can ameliorate edema in spinal cord slices effectively, in both ACSF and sucrose-ACSF. Also, in our experience, we can obtain healthy spinal cord slices successfully from neonatal as well as 3&#8211;10 weeks old SD rats. Thus, for this protocol to be successful, we recommend using SD rats that are less than 10 weeks old.
While performing &#39;ventral&#39; laminectomy and removing the meninges and spinal nerves, one should be patient and careful to avoid cutting, stretching or splitting the spinal cord. In some studies, spinal cord slices with attached dorsal roots have been used to evaluate the synaptic transmission SG neurons received peripherally25,26. The procedure of removing pia-arachnoid membrane is of technical difficulty in this case, and it requires a lot of patience.
This slice preparing technique also has some limitations. One clear drawback is that although acute slices preserve abundant synaptic connections, it could not reflect the real state and address what exactly happens in vivo. Thus, some studies have implemented in vivo recordings that are normally performed &#39;blind&#39;27,28,29. However, this in vivo approach is technically challenging, and it is difficult to tell whether a recording is performed from the soma or dendrite without sufficient experience. Another limitation of our current method is that sucrose-ACSF may not be sufficient for neuronal preservation when preparing slices from aging rodents. An updated approach using N-methyl-D-glucamine as a Na+ substitute has been proposed, and this optimized methodology could markedly improve morphological and functional preservation of neurons in acute slices30,31,32,33. Finally, SG neurons show different morphological and electrophysiological properties3. It seems difficult to interpret data obtained from whole-cell recordings while overlooking the heterogeneity. This limitation may be sidestepped by further verifying the morphological details of recorded neurons5 or using transgenic mice, which could help researchers identify specific neurons20,34. Furthermore, optogenetics, a novel tool allowing control of a sub-population of cells35, could be combined with whole-cell patch-clamp recording to study the role of specific ion channels or proteins and to investigate specific neuronal circuitry.
Overall, this preparation technique is an ideal way to investigate the electrophysiological, morphological, pharmacological, and biological characteristics of SG neurons, complemented by patch-clamp recording, immunofluorescent staining, specific agonists or antagonists, and the single-cell RT-PCR technique. Moreover, this approach can be applied together with paired patch-clamp recordings or optogenetics, and it is thus a valuable tool for illuminating the neuronal microcircuits.

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+-based intracellular solution</td>
		</tr>
		<tr>
			<td>NaH2PO4&#183;2H2O</td>
			<td>Sigma</td>
			<td>71500</td>
			<td>Used for the preparation of ACSF, sucrose-ACSF and PBS</td>
		</tr>
		<tr>
			<td>CaCl2&#183;2H2O</td>
			<td>Sigma</td>
			<td>C5080</td>
			<td>Used for the preparation of ACSF and sucrose-ACSF</td>
		</tr>
		<tr>
			<td>MgCl2&#183;6H2O</td>
			<td>Sigma</td>
			<td>M2670</td>
			<td>Used for the preparation of ACSF and sucrose-ACSF</td>
		</tr>
		<tr>
			<td>NaHCO3</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+-based intracellular solution</td>
		</tr>
		<tr>
			<td>Na2-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>CsMeSO4</td>
			<td>Sigma</td>
			<td>C1426</td>
			<td>Used for the preparation of Cs+-based intracellular solution</td>
		</tr>
		<tr>
			<td>CsCl</td>
			<td>Sigma</td>
			<td>C3011</td>
			<td>Used for the preparation of Cs+-based intracellular solution</td>
		</tr>
		<tr>
			<td>TEA-Cl</td>
			<td>Sigma</td>
			<td>T2265</td>
			<td>Used for the preparation of Cs+-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>Na2HPO4</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 patterns observed in SG neurons included tonic-firing, delayed-firing, gap-firing, initial-burst, phasic-bursting, single-spike and reluctant-firing, which have been described and categorized by previous studies.
Implementing this preparation, we also recorded subthreshold currents and spontaneously appearing currents in voltage clamp. Representative traces of subthreshold currents, including hyperpolarization-activated current (Ih), T-type calcium current (IT) and A-type potassium current (IA), are given in Figure 3A. These currents were obtained by holding cells at -50 mV and gradually stepping in 10-mV decrements from -60 to -120 mV. Ih was activated by hyperpolarizing voltage steps. However, IT and IA were activated by hyperpolarizing prepulses to release from inactivation followed with a depolarized voltage. Figure 3B, 3C show representative spontaneous EPSCs (sEPSCs) and IPSCs (sIPSCs) recorded from SG neurons, respectively. The amplitude and frequency of these synaptic events could be analyzed using the Mini-analysis software offline.
To characterize neuronal morphological features, parasagittal slices were applied because most of the SG neurons have significantly rostrocaudal spread of dendritic trees, and neurobiotin 488 was added to intracellular solutions. The size of neuronal soma and the extent and dimensions of their dendritic processes were evaluated after confocal microscopy imaging. As reported previously, SG neurons show morphological distinctions and could be categorized into central cells, radial cells, vertical cells, islet cells and unclassified cells. Representative micrographs of these cells are shown in Figure 4.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58479/58479fig1.jpg" /> 
Figure 1: Diagram for acute spinal cord slice preparation. After being deeply anesthetized with urethan (i.p.), rats are transcardially perfused with ice-cold carbogenated sucrose-ACSF. The spinal column is then quickly dissected, and a ventral laminectomy is performed. The meninges, pia-arachnoid membrane and attached spinal nerve roots are removed. Then, the spinal cord specimen is mounted on an agarose block. Transverse or parasagittal slices are cut with a vibratome as needed. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58479/58479fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/58479/58479fig2.jpg" /> 
Figure 2: Firing patterns of SG neurons. Firing patterns are determined by injecting a series of 1 s depolarizing current pulses into an SG neuron at RMP. The firing patterns may be classified as tonic-firing, delayed-firing, gap-firing, initial-burst, phasic-bursting, single-spike, and reluctant-firing. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58479/58479fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/58479/58479fig3.jpg" /> 
Figure 3: Voltage-clamp recordings in SG neurons.&#160;(A) Representative traces showing the response to hyperpolarizing current injection classified as Ih, IA and IT. The lower panel shows the evoking protocol for sub-threshold currents in voltage-clamp. (B) Representative traces of sEPSCs recorded from SG neurons at -70 mV in the absence and presence of 50 &#956;M APV and 20 &#956;M CNQX. Lower consecutive traces, which are shown in an expanded time scale before (left) and under (right) the action of APV and CNQX, correspond to a period indicated by a bar shown below the chart recording. (C)&#160;Representative traces of sIPSCs recorded from SG neurons in the absence and presence of 10 &#956;M bicuculline and 1 &#956;M strychnine at 0 mV. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58479/58479fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/58479/58479fig4.jpg" /> 
Figure 4: Representative morphology of rat SG neurons. According to soma sizes and dendrite properties shown in confocal microscopy images, SG neurons may be classified as the central cell (A), radial cell (B), vertical cell (C), islet cell (D) and unclassified cell (E). V: ventral; D: dorsal; R: rostral; C: caudal. Scale bar = 50 &#181;m. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/58479/58479fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component</td>
			<td>Molecular Weight</td>
			<td>Concentration (mM)</td>
			<td>g/L</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>58.5</td>
			<td>117</td>
			<td>6.84</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>74.5</td>
			<td>3.6</td>
			<td>0.27</td>
		</tr>
		<tr>
			<td>NaH2PO4&#183;2H2O</td>
			<td>156</td>
			<td>1.2</td>
			<td>0.19</td>
		</tr>
		<tr>
			<td>CaCl2&#183;2H2O</td>
			<td>147</td>
			<td>2.5</td>
			<td>0.37</td>
		</tr>
		<tr>
			<td>MgCl2&#183;6H2O</td>
			<td>203</td>
			<td>1.2</td>
			<td>0.24</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>84</td>
			<td>25</td>
			<td>2.1</td>
		</tr>
		<tr>
			<td>D-Glucose</td>
			<td>180</td>
			<td>11</td>
			<td>1.98</td>
		</tr>
		<tr>
			<td>Ascorbic acid</td>
			<td>198.11</td>
			<td>0.4</td>
			<td>0.08</td>
		</tr>
		<tr>
			<td>Sodium pyruvate</td>
			<td>110</td>
			<td>2</td>
			<td>0.22</td>
		</tr>
	</tbody>
</table>
Table 1: Recipe for ACSF.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component</td>
			<td>Molecular Weight</td>
			<td>Concentration (mM)</td>
			<td>g/L</td>
		</tr>
		<tr>
			<td>NaCl</td>
			<td>58.5</td>
			<td>117</td>
			<td>6.84</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>74.5</td>
			<td>3.6</td>
			<td>0.27</td>
		</tr>
		<tr>
			<td>NaH2PO4&#183;2H2O</td>
			<td>156</td>
			<td>1.2</td>
			<td>0.19</td>
		</tr>
		<tr>
			<td>CaCl2&#183;2H2O</td>
			<td>147</td>
			<td>2.5</td>
			<td>0.37</td>
		</tr>
		<tr>
			<td>MgCl2&#183;6H2O</td>
			<td>203</td>
			<td>1.2</td>
			<td>0.24</td>
		</tr>
		<tr>
			<td>NaHCO3</td>
			<td>84</td>
			<td>25</td>
			<td>2.1</td>
		</tr>
		<tr>
			<td>D-Glucose</td>
			<td>180</td>
			<td>11</td>
			<td>1.98</td>
		</tr>
		<tr>
			<td>Ascorbic acid</td>
			<td>198.11</td>
			<td>0.4</td>
			<td>0.08</td>
		</tr>
		<tr>
			<td>Sodium pyruvate</td>
			<td>110</td>
			<td>2</td>
			<td>0.22</td>
		</tr>
	</tbody>
</table>
Table 2: Recipe for sucrose-ACSF.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component</td>
			<td>Molecular Weight</td>
			<td>Concentration (mM)</td>
			<td>mg/100 mL</td>
		</tr>
		<tr>
			<td>K-gluconate</td>
			<td>234.2</td>
			<td>130</td>
			<td>3044.6</td>
		</tr>
		<tr>
			<td>KCl</td>
			<td>74.5</td>
			<td>5</td>
			<td>37.28</td>
		</tr>
		<tr>
			<td>Na2-Phosphocreatine</td>
			<td>453.38</td>
			<td>10</td>
			<td>453.38</td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>380.35</td>
			<td>0.5</td>
			<td>19.02</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>238.31</td>
			<td>10</td>
			<td>238.3</td>
		</tr>
		<tr>
			<td>Mg-ATP</td>
			<td>507.18</td>
			<td>4</td>
			<td>202.9</td>
		</tr>
		<tr>
			<td>Li-GTP</td>
			<td>523.18</td>
			<td>0.3</td>
			<td>15.7</td>
		</tr>
	</tbody>
</table>
Table 3: Recipe for K+-based intracellular solution.
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Component</td>
			<td>Molecular Weight</td>
			<td>Concentration (mM)</td>
			<td>mg/100 mL</td>
		</tr>
		<tr>
			<td>CsMeSO4</td>
			<td>228</td>
			<td>92</td>
			<td>2097.6</td>
		</tr>
		<tr>
			<td>CsCl</td>
			<td>168.36</td>
			<td>43</td>
			<td>723.95</td>
		</tr>
		<tr>
			<td>Na2-Phosphocreatine</td>
			<td>453.38</td>
			<td>10</td>
			<td>453.38</td>
		</tr>
		<tr>
			<td>TEA-Cl</td>
			<td>165.71</td>
			<td>5</td>
			<td>82.86</td>
		</tr>
		<tr>
			<td>EGTA</td>
			<td>380.35</td>
			<td>0.5</td>
			<td>19.02</td>
		</tr>
		<tr>
			<td>HEPES</td>
			<td>238.31</td>
			<td>10</td>
			<td>238.3</td>
		</tr>
		<tr>
			<td>Mg-ATP</td>
			<td>507.18</td>
			<td>4</td>
			<td>202.9</td>
		</tr>
		<tr>
			<td>Li-GTP</td>
			<td>523.18</td>
			<td>0.3</td>
			<td>15.7</td>
		</tr>
	</tbody>
</table>
Table 4: Recipe for Cs+-based intracellular solution.

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

Nanyang Technological University

Research

JoVE Journal

Neuroscience

13.9K Views.  First Affiliated Hospital of Nanchang University. 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.

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

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 ...

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of intracellular events that lead to closure of cyclic-nucleotide-gated channels in the cell membrane. The resulting change in membrane potential leads in turn to reductions in the amount of neurotransmitter release from both rod and cone synaptic terminals. To measure how the light-evoked change in photoreceptor membrane potential leads to downstream activity in the retina, scientists have made electrophysiological recordings from retinal slice preparations for decades1,2. In the past these slices have been cut manually with a razor blade on retinal tissue that is attached to filter paper; in recent years another method of slicing has been developed whereby retinal tissue is embedded in low gelling temperature agar and sliced in cool solution with a vibrating microtome3,4. This preparation produces retinal slices with less surface damage and very robust light-evoked responses. Here we document how this procedure can be done under infrared light to avoid bleaching the visual pigment.

Here we describe a procedure for generating dark-adapted slices of the mouse retina for electrophysiological recordings.

Our visual experience is initiated when the visual pigment in our retinal photoreceptors absorbs photons of light energy and initiates a cascade of ...

Whole Cell Patch Clamp for Investigating the Mechanisms of Infrared Neural Stimulation

It has been demonstrated in recent years that pulsed, infrared laser light can be used to elicit electrical responses in neural tissue, independent of any further modification of the target tissue. Infrared neural stimulation has been reported in a variety of peripheral and sensory neural tissue in vivo, with particular interest shown in stimulation of neurons in the auditory nerve. However, while INS has been shown to work in these settings, the mechanism (or mechanisms) by which infrared light causes neural excitation is currently not well understood. The protocol presented here describes a whole cell patch clamp method designed to facilitate the investigation of infrared neural stimulation in cultured primary auditory neurons. By thoroughly characterizing the response of these cells to infrared laser illumination in vitro under controlled conditions, it may be possible to gain an improved understanding of the fundamental physical and biochemical processes underlying infrared neural stimulation.

Infrared nerve stimulation has been proposed as an alternative to electrical stimulation in a range of nerve types, including those associated with the auditory system. This protocol describes a patch clamp method for studying the mechanism of infrared nerve stimulation in a culture of primary auditory neurons.

It has been  demonstrated in recent years that pulsed, infrared laser light can be used to  elicit electrical responses in neural tissue, independent of ...

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 ...

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 ...

Whole-cell Patch-clamp Recordings for Electrophysiological Determination of Ion Selectivity in Channelrhodopsins

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate electrical processes in target cells. In this context, ion selectivity of a channelrhodopsin is of particular importance. This article describes the investigation of chloride selectivity for a recently identified anion-conducting channelrhodopsin of Proteomonas sulcata via electrophysiological patch-clamp recordings on HEK293 cells. The experimental procedure for measuring light-gated photocurrents demands a fast switchable &#8211; ideally monochromatic &#8211; light source coupled into the microscope of an otherwise conventional patch-clamp setup. Preparative procedures prior to the experiment are outlined involving preparation of buffered solutions, considerations on liquid junction potentials, seeding and transfection of cells, and pulling of patch pipettes. The actual recording of current-voltage relations to determine the reversal potentials for different chloride concentrations takes place 24 h to 48 h after transfection. Finally, electrophysiological data are analyzed with respect to theoretical considerations of chloride conduction.

This article describes how the ion selectivity of channelrhodopsin is determined with electrophysiological whole-cell patch-clamp recordings using HEK293 cells. Here, the experimental procedure for investigating chloride selectivity of an anion-selective channelrhodopsin is demonstrated. However, the procedure is transferable to other channelrhodopsins of distinct selectivity.

Over the past decade, channelrhodopsins became indispensable in neuroscientific research where they are used as tools to non-invasively manipulate ...

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 ...

Application of Automated Image-guided Patch Clamp for the Study of Neurons in Brain Slices

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a challenging and low-throughput technique due to its complexity and high reliance on user operation and control. This manuscript demonstrates an image-guided automatic patch clamp system for in vitro whole-cell patch clamp experiments in acute brain slices. Our system implements a computer vision-based algorithm to detect fluorescently labeled cells and to target them for fully automatic patching using a micromanipulator and internal pipette pressure control. The entire process is highly automated, with minimal requirements for human intervention. Real-time experimental information, including electrical resistance and internal pipette pressure, are documented electronically for future analysis and for optimization to different cell types. Although our system is described in the context of acute brain slice recordings, it can also be applied to the automated image-guided patch clamp of dissociated neurons, organotypic slice cultures, and other non-neuronal cell types.

This protocol describes how to conduct automatic image-guided patch-clamp experiments using a system recently developed for standard in vitro electrophysiology equipment.

Whole-cell patch clamp is the gold-standard method to measure the electrical properties of single cells. However, the in vitro patch clamp remains a ...

Auditory Brainstem Response and Outer Hair Cell Whole-cell Patch Clamp Recording in Postnatal Rats

The outer hair cell is one of the two types of sensory hair cells in the mammalian cochlea. They alter their cell length with the receptor potential to amplify the weak vibration of low-level sound signal. The morphology and electrophysiological property of outer hair cells (OHCs) develop in early postnatal ages. The maturation of outer hair cell may contribute to the development of the auditory system. However, the process of OHCs development is not well studied. This is partly because of the difficulty to measure their function by an electrophysiological approach. With the purpose of developing a simple method to address the above issue, here we describe a step-by-step protocol to study the function of OHCs in acutely dissociated cochlea from postnatal rats. With this method, we can evaluate the cochlear response to pure tone stimuli and examine the expression level and function of the motor protein prestin in OHCs. This method can also be used to investigate the inner hair cells (IHCs).

This protocol describes methods to record the auditory brainstem response from postnatal rat pups. To examine the functional development of outer hair cells, the experimental procedure of whole-cell patch clamp recording in isolated outer hair cells is described step-by-step.

The outer hair cell is one of the two types of sensory hair cells in the mammalian cochlea. They alter their cell length with the receptor potential to ...

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

Summary

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Chapters in this video

Dissection and Culture of Commissural Neurons from Embryonic Spinal Cord

Patch Clamp Recordings from Mouse Retinal Neurons in a Dark-adapted Slice Preparation

Whole Cell Patch Clamp for Investigating the Mechanisms of Infrared Neural Stimulation

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

Whole-cell Patch-clamp Recordings in Brain Slices

The Preparation of Oblique Spinal Cord Slices for Ventral Root Stimulation

Whole-cell Patch-clamp Recordings for Electrophysiological Determination of Ion Selectivity in Channelrhodopsins

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

Application of Automated Image-guided Patch Clamp for the Study of Neurons in Brain Slices

Auditory Brainstem Response and Outer Hair Cell Whole-cell Patch Clamp Recording in Postnatal Rats