The authors thank Marin Manuel and Olivia Goldman-Szwajkajzer for their help in taking the photographs. The authors also thank Arjun Masukar and Tobias Bock for proofreading the manuscript. Financial supports were provided by the Agence Nationale pour la Recherche (HYPER-MND, ANR-2010-BLAN-1429-01), the NIH-NINDS (R01NS077863), the Thierry Latran Foundation (OHEX Project), the French association for myopathy (grant number 16026) and Target ALS are gratefully acknowledged. Felix Leroy was the recipient of a &#34;Contrat Doctoral&#34; from the Ecole Normale Sup&#233;rieure, Cachan.

&#160;The experiments were performed in accordance with European directives (86/609/CEE and 2010-63-UE) and French legislation, and&#160;were approved by the Paris Descartes University ethics committee.
1. Spinal Cord Slice Preparation
<ol>
	<li>Prepare the following solutions daily or one day in advance. If kept overnight, bubble with 95% O2 and 5% CO2 and keep refrigerated in tightly closed bottles.
	<ol>
		<li>Prepare Low Na + artificial cerebrospinal fluid (ACSF): 3 mM KCl, 1 mM NaH2PO4, 230 mM sucrose, 26 mM NaHCO3, 0.8 mM CaCl2, 8 mM MgCl2, 25 mM glucose, 0.4 mM ascorbic acid, 1 mM Na-kynurenate, 2 mM Na-pyruvate. Bubble with 95% O2 and 5% CO2 (pH 7.4). Since Na-kynurenate is often hard to dissolve, make sure to purchase the one listed in the materials table.</li>
		<li>Prepare K-gluconate solution: 130 mM K-gluconate, 15 mM KCl, 0.05 mM EGTA, 20 mM HEPES, 25 mM glucose, 1 mM Na-kynurenate, 2 mM Na-pyruvate, adjusted to pH 7.4 with KOH.</li>
		<li>Prepare ACSF: 130 mM NaCl, 2.5 mM KCl, 2 mM CaCl2, 1 mM MgCl2, 1 mM NaH2PO4, 26 mM NaHCO3, 25 mM glucose, 0.4 mM ascorbic acid, 2 mM Na-pyruvate. Bubble with 95% O2 and 5% CO2 (pH 7.4).</li>
		<li>Prior to the beginning of the dissection, dissolve 2% agar in 80 ml of the K-gluconate solution, and keep warm at 60 &#176;C.</li>
	</ol>
	</li>
	<li>Intracardiac perfusion
	<ol>
		<li>Perform this preparation on female and male mice, ranging in age&#160;from P2 to P11.</li>
		<li>Anesthetize the mouse with an intraperitoneal injection of 0.1 ml of 25 mM pentobarbital sodium (50 mg/kg).</li>
		<li>Using needles or tape, immobilize the mouse on its back on a large petri dish filled with silicon. Use a dissecting microscope for the rest of the dissection.</li>
		<li>Holding the tip of the sternum, lift the chest and cut the diaphragm using fine scissors. Then open the chest on both sides by cutting through the ribs to expose the heart.</li>
		<li>Cut the right atrium before puncturing the left ventricle with a 27G needle.</li>
		<li>Perfuse with ice-cold low Na+ ACSF. After 30 sec, low Na+ ACSF should be seen flowing out of the atrium. Low amounts of sodium prevent the cells from spiking and reduce cellular death during dissection.</li>
		<li>Keep holding the needle in the heart under the dissecting microscope until the liver turns yellow when the blood is drained.</li>
	</ol>
	</li>
	<li>Spinal cord dissection
	<ol>
		<li>Decapitate the animal and put it on its stomach.</li>
		<li>Quickly remove the skin of the back (Figure 1A1). Make two cuts through the shoulders and going down the chest cage (Figure 1A2). Then cut the cord as low as possible in the caudal section in order to isolate the vertebral column with the beginning of the ribs from the lower part of the animal. Flip the animal again and remove the viscera still attached to the ribs.</li>
		<li>Transfer the vertebral column to another, smaller silicon-filled petri dish and use 4 insect pins to hold it dorsal side up (Figure 1A).</li>
		<li>Continuously perfuse the animal with carbogen-bubbled ACSF (at about 4 &#176;C) while performing a laminectomy of the dorsal side, and exposing the spinal cord from the rostral end (Figure 1B). To do so, insert the tip of fine scissors between the bone and the spinal cord and cut the bone little by little from the rostral end, making sure to stay away from the white matter. Alternate on each side while using tweezers to keep away the band of bone already cut&#160;(Figure 1B1).</li>
		<li>Using the smallest available spring scissors and tweezers, lift the dura and cut on both sides while holding the loose part of the dura in order to avoid damaging the spinal cord with the scissors. Cut along the rostro-caudal axis. 
		NOTE: The dura is a semi transparent continuous membrane; at this age the pia mater is too fragile and will come apart during the dissection and slicing (Figure 1B2).</li>
		<li>Once the dura is removed use a blunt glass or plastic tip to gently push the cord on the left side of the groove formed by the half-cut spinal column, and cut the ventral and dorsal roots on the right side, starting from the rostral side, furthest&#160;from where they enter the cord (a few mm at least, Figure 1B2).</li>
		<li>Repeat the operation on the left side, always going from rostral to caudal. If left-handed, start from the left side and then move to the right side.</li>
	</ol>
	</li>
	<li>Embedding in agar
	<ol>
		<li>Slip the cord out of the vertebral column. Use a smaller insect pin to pin down the cord with the dorsal surface up and delicately remove any piece of membrane still attached to it (Figure 1C1).</li>
		<li>Once cleaned, trim both ends (Figure 1C2). Insert a bended insect pin in the anterior part of the cord to manipulate the cord and note its orientation (arrow in Figure 1C2). Then transfer the spinal cord to an ice-cold K-gluconate solution.&#160; 
		NOTE: This solution mimics the intracellular composition of the CSF and will&#160;prevent the cells from dying of osmotic shock once the&#160;motoneurons will be cut26.</li>
		<li>Once the spinal cord is within the intra-cellular solution, take the beaker with the agar out of the dry bath and cool it down on a mixture of ice and water.</li>
		<li>Keep stirring while measuring the temperature. When the temperature reaches 38 &#176;C immerse the spinal cord, holding it by the insect pin and place it rostral side down. Make sure the spinal cord is as straight as possible, away from the walls, caudal part slightly upward (Figure 1D1).</li>
		<li>Leave the beaker in the mixture of ice and water to allow the agar to solidify as quickly as possible. Make sure it stays in place and that the cord is as straight as possible (Figure 1D1).</li>
	</ol>
	</li>
	<li>Slicing
	<ol>
		<li>After solidification, cut the agar block containing the spinal cord in such a way that the base of the block is at a 35&#176; angle with the lumbar part of the cord (arrow in Figure 1D2). The dorsal surface should be facing away from the base (Figure 1D2). 
		NOTE: This is a critical step in the procedure, for maintaining the continuity of the motoneuron pools to the ventral roots from which they exit.</li>
		<li>Glue the block into the chamber of the vibratome using cyanoacrylate glue. Immerse it in K-gluconate solution and add slushed frozen K-gluconate solution to maintain the bath chilled (below 2 &#176;C).</li>
		<li>Cut 350 - 400 &#181;m thick slices of the lumbar region (identifiable by its curvature and larger diameter). In its proper orientation (see 1.4.1.), use the blade to cut from the dorsal to the ventral surface and make slices continuously more rostral. Typically, there are 4-5 suitable slices with ventral roots extending 2 mm or more. Use the following parameters: 10&#176; angle, 70 Hz vibration frequency&#160;and 10 mm/min speed of slicing.</li>
	</ol>
	</li>
	<li>Incubation
	<ol>
		<li>Transfer the slices to ACSF at 34 &#176;C. After approximately 30 min, cool the slices down to RT&#160;and begin the recording session.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 1" src="/files/ftp_upload/54525/54525fig1.jpg" /> 
Figure 1.&#160;Dissection 
A1: Removal of the skin of the back to expose the dorsal column. A2: Cutting of the shoulders and ribs to free the dorsal column. B1: Vertebral column pinned onto the silicon-filled petri dish (dorsal side up, caudal side left). B2: Same with the spinal cord exposed and dissected. C1: Spinal cord isolated (rostral side left). C2: Spinal cord ready to be embedded (ventral side up, rostral side left). Note the smaller insect pin on the rostral side. D1: Spinal cord in the agar beaker (rostral side dow, ventral side facing the bottom). D2: Cut agar block with the embedded spinal cord. Note the 35&#176; angle the spinal cord forms with the base of the block and the lumbar enlargement (Arrow). Scale bars 1 cm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54525/54525fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
2. Placing the Slice into the Chamber
NOTE: Ventral roots are of variable size.
<ol>
	<li>Prepare a box of various pipettes with tip diameters ranging from 40 to 170 &#181;m in advance. To prepare suction pipettes, prepare many pipettes with a long taper. Using a diamond knife, make a cut at different positions. Then under a dissecting microscope, break it by hitting the tip with tweezers.</li>
	<li>Remove the chamber from the recording microscope and place it under a dissecting microscope.</li>
	<li>Select a slice that contains a ventral root of sufficient length (2 mm or more) to be mounted on a suction stimulation electrode. Select the proper orientation of the slice with the ventral roots upward (Figure 2A) and delicately cut the agar around the ventral roots while leaving the rest of the agar around the slice (Figure 2B). 
	NOTE: Because the agar is firmer than the slice, this will allow the threads of the slice&#39;s anchor to rest on the agar rather than on the slice and thus the anchor will not damage the tissue. Make sure the threads of the slice&#39;s anchor are above the motoneuron pool (shown in red in Figure 2C).</li>
	<li>Mount the chamber back onto the microscope and continuously perfuse the recording chamber with ACSF at a rate of 1 - 2 ml/min, at RT. Using a glass pipette filled with ACSF and connected to a syringe, suck one of the ventral root (arrow in Figure 2C). In order to achieve good stimulation of the ventral root, the pipette tip needs to be tight around the ventral root. One pole should be in the stimulating electrode and the other one in the bath (or connected to the patch-clamping electrode reference).</li>
	<li>Achieve patch-clamp recording of the desired cell-type and record the effect of the ventral root stimulation as described previsouly10.
	<ol>
		<li>Here, use an amplifier for data acquisition. Filter whole-cell recordings at 3 kHz. Digitize at 10 kHz. Compensate bridge resistance in current-clamp mode.</li>
	</ol>
	</li>
</ol>
<img alt="Figure 2" src="/files/ftp_upload/54525/54525fig2.jpg" /> 
Figure 2. Ventral Root Preparation 
A: Lumbar spinal cord slice embedded in agar with the ventral root facing up. B: Lumbar spinal cord slice with the ventral roots freed from the agar. C: Lumbar spinal cord slice with a stimulating electrode tightly placed around the ventral roots (arrow). Note the location of the Renshaw cells expressing red fluorescence in the chrna2-Cre mouse28 crossed with the mouse reporter R26Tom 17. Scale bars 1 mm. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54525/54525fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

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L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Experimental+study+of+the+spinothalamic+and+related+tracts.">Experimental study of the spinothalamic and related tracts.</a> Acta Psychiatr Neurol Scand. 26 (3-4), 371-396 (1951).</li><li>Sengul, G., Fu, Y., Yu, Y., Paxinos, G. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spinal+cord+projections+to+the+cerebellum+in+the+mouse.">Spinal cord projections to the cerebellum in the mouse.</a> Brain Struct Funct. 220 (5), 2997-3009 (2015).</li><li>Russier, M., Carlier, E., Ankri, N., Fronzaroli, L., Debanne, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A-,+T-,+and+H-type+currents+shape+intrinsic+firing+of+developing+rat+abducens+motoneurons.">A-, T-, and H-type currents shape intrinsic firing of developing rat abducens motoneurons.</a> J Physiol. 549 (Pt 1), 21-36 (2003).</li><li>Dourado, M., Sargent, P. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Properties+of+nicotinic+receptors+underlying+Renshaw+cell+excitation+by+alpha-motor+neurons+in+neonatal+rat+spinal+cord).">Properties of nicotinic receptors underlying Renshaw cell excitation by alpha-motor neurons in neonatal rat spinal cord).</a> J Neurophysiol. 87 (6), 3117-3125 (2002).</li><li>Mitra, P., Brownstone, R. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+in+vitro+spinal+cord+slice+preparation+for+recording+from+lumbar+motoneurons+of+the+adult+mouse.">An in vitro spinal cord slice preparation for recording from lumbar motoneurons of the adult mouse.</a> J Neurophysiol. 107 (2), 728-741 (2012).</li><li>Rothman, S. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+neurotoxicity+of+excitatory+amino+acids+is+produced+by+passive+chloride+influx.">The neurotoxicity of excitatory amino acids is produced by passive chloride influx.</a> J Neurosci. 5 (6), 1483-1489 (1985).</li><li>Olney, J. W., Price, M. T., Samson, L., Labruyere, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+role+of+specific+ions+in+glutamate+neurotoxicity.">The role of specific ions in glutamate neurotoxicity.</a> Neurosci Lett. 65 (1), 65-71 (1986).</li></ol>

The authors have nothing to disclose.

Oblique slicing of the spinal cord is important since it allows for unilateral stimulation of motoneuron pools and Renshaw cells at a single vertebral segment in a reliable, comprehensive&#160;and specific way. Furthermore, it allows for a quick, elegant&#160;and non-ambiguous identification of recorded motoneurons. Next, we will highlight&#160;the advantages of this&#160;technique compared to other slice preparation methods, and then we will stress out&#160;the most common pitfalls to avoid while performing this procedure.
Most studies using transverse slices base their identification on intrinsic electrical properties 3,14,15,29. However, these vary greatly between motoneuron pools as well as between motoneuron subtypes within a given pool (alpha-, beta- and gamma-motoneurons30). Therefore, studies using size parameters are likely to exclude smaller gamma-motoneurons. On the other hand, Cooper and Sherrington described in 1940 a group of large nerve cells in the ventro-lateral grey matter of the lumbar spinal cord of monkeys and cats which had crossed ascending axons31,32. In addition to being close to the motoneuron&#160;somas, they noted that they appeared histologically indistinguishable from motoneurons. A recent article confirmed the existence of such cells in the mouse33. Due to the large size of these cells and their location, they could have been mistakenly included in former studies. Our identification criterion excludes putative spinal border cells, since their axon does not project in the ventral root, while allowing reliable identification of all motoneurons regardless of their&#160;size. Furthermore, analyzing the intrinsic electrical properties, (e.g., input resistance), is more time consuming than the simple observation of an antidromic action potential.
In addition to using intrinsic electrical properties, some studies using&#160;transverse slices, confirm their identification by performing post&#160;hoc analysis of the molecular marker for motoneurons (such as Islet-1/216) or by visualizing the morphology using biocytin labelings14,17. Such identifications are however tedious and do not provide a direct identification. They are therefore rarely systematically performed.
Some studies tried to make use of mouse lines expressing a genetically-encoded motoneuron&#160;fluorescent marker&#160;(Hb9-GFP mouse line18, ChAT-EGFP transgenic mice,&#160;which express eGFP in cholinergic cells including motoneurons19, or G85R SOD1-YFP transgenic mice, which strongly express the YFP fusion protein in motoneurons17). However, because the genetic marker expression is usually conditioned in time and since the slices are usually taken from embryos or juveniles, the expression of the marker may not be adequate at the age the study is performed. Other studies have used retrograde agent injection to visualize the motoneuron&#160;pool34. We also performed Cholera toxin beta injections into the soleus or the EDL (personal observation). Such labeling techniques are of great value to label a particular motoneuron&#160;pool but require tedious surgery a few days prior to the experiment. Furthermore, the young age at which the slice experiments are being done, makes it significantly harder to specifically target the muscles. Finally, there is a real concern of perturbing the motoneuron by injecting an exogenous agent.
Under optimal conditions we were able to obtain antidromic action potentials in virtually all of the recorded motoneurons. The 35&#176; angle used is critical to obtain ventral roots of sufficient length. In conclusion, the oblique slice preparation offers a quick and reliable way to identify every motoneuron, which is superior to other identification techniques.
As stated in the introduction, only two studies reported successful stimulation of the ventral roots in transverse slices. The first study recorded cervical neurons in the rat and stimulated the white matter where the ventral root stub emerges4. They successfully induced antidromic action potential in 85% of the recorded neurons. We believe their high success rate relied on the fact that at the cervical level, the axons and somas of motoneurons are on the same plane. This is not the case in more caudal slices. Even at the cervical level, their transverse slices only retained stubs of the ventral roots, which are less reliable to stimulate. The second study stimulated the chick embryo dorso-lumbar segment5. However,&#160;they used voltage sensitive dye recording, which lacked the spatial resolution to indicate the success of inducing an antidromic action potentials at the single cell level.
Our slice preparation also offers a superior way to stimulate Renshaw cells as stated in a recent paper28. Using the oblique slices, they were able to record monosynaptic Renshaw cells responses in 46% of the cases, which was a major improvement from the 10% success rate encountered using transverse slices35. Although we could not find a similar comparison for the stimulation of the motoneuron&#160;pools, this observation in the Renshaw cell leads us to believe that motoneurons also retain more connectivity.
Oblique slicing of the spinal cord is a highly demanding technique that requires much practice before reliably obtaining a preparation from which one can record. Using very fine and sharp tweezers and micro-scissors is essential. The time spent dissecting can be quite long (up to 1 hr) as long as the entire procedure is conducted in chilled carbogen-bubbled dissection medium. While some authors did not acknowledge any benefit of intracardiac perfusion in the preparation of slices36, we decided to maintain this step of the procedure. It is hard to objectively measure the benefit of intracardiac perfusion for the quality of the recordings, especially because it varies with age.
The most critical factor is the mouse age. We successfully recorded from mice between P2 and P11 using the above protocol. Past this age, the spinal cord myelin becomes too dense and the biggest cells (the motoneurons) die before slicing, most likely due to oxygen deprivation. After that window, we found it increasingly difficult to obtain healthy slices. Recently, new techniques have been reported to obtain healthy slices in older animals 17,36. They used extra additives to their dissecting and recording medium such as supplementary sources of energy (ethyl-pyruvate), lower Na+ and Cl- concentrations to prevent oncosis of the cells37,38 and polyethylene glycol to limit the effects of the extensive membrane transections that occur during slicing. With such enhancements, they were able to record from up to 6 month old animals.
In the future it would be interesting to attempt oblique slice preparations in adult animals to combine the advantages of the two techniques. Combining the two protocols should prove relatively easy and will provide a reliable tool to confirm motoneuron identity and study the population targeted by axon collaterals of motoneurons in adult mice. Additionally, one could try obtaining oblique slices in the embryo mouse. Noteworthy, our preparation also retains the dorsal roots, which can be stimulated to study the activation monosynaptic Ia innervation to the motoneurons, as well as the activation of dorsal spinal cord neuronal populations. In conclusion, ventral root stimulations offer a reliable tool to confirm motoneuron identity and to study the population targeted by motoneuron axon collaterals.

Historically, motoneuron recordings using sharp-electrode were conducted in vivo on large animals such as&#160;cats or rats1 or on an isolated whole spinal cord in&#160;mice2. The emergence of the patch-clamp recording technique during the 1980s, called for direct access to the motoneuron somas as sealing needed to be achieved under visual guidance. Thus, spinal cord slice preparation has been readily achieved since the early 1990s3. However, early slice preparation often did not allow for the stimulation of the ventral roots. To the best of our knowledge, only two studies have reported successful stimulation of the ventral roots in transverse slices, and none was obtained from mice4,5.
In this article we present a technique to achieve viable spinal cord slices of neonatal mice (P2 - P11) in which the motoneuron&#160;pool&#160;retains its&#160;ventral root departing&#160;axons. Ventral root stimulation triggers antidromic action potential back into the somas of the motoneuron&#160;pool&#160;exiting from the same ventral root. It also excites the motoneuron collateral targets,&#160;other motoneurons6-10 and the Renshaw cells11-13. Since only motoneurons&#160;send their axons down the ventral roots, we use the recording of antidromic action potentials as a simple and definitive way to physiologicaly identify motoneurons10.
In addition to using potentially non-inclusive or misleading electrophysiological and morphological&#160;criterions to confirm the motoneuron&#160;identity, recent studies on spinal cord motoneurons also relied on tedious and&#160;time-consuming post&#160;hoc stainings16. Such identification is usually performed only on a sample of the recorded cells. Other identification strategies rely&#160;on mouse lines in which the motoneurons express endogenous fluorescence17-19. However, using genetically encoded markers may be difficult at a young age when marker expression is still variable or if the study already requires using a transgenic mouse line. Alternatively, antidromic action potential recordings can be performed routinely on all mice from the onset of cell recording. Experimenters working on intact spinal cord preparations in the cat, rat and mouse, have reliably used such identification techniques since the 1950&#39;s1,2,20,21. In optimal conditions, we were able to elicit antidromic action potentials from virtually all of the recorded motoneurons.
Furthermore, ventral root stimulation can be used to reliably excite other motoneurons22,23&#160;or their targets. the&#160;Renshaw cells10,24,25. We present here applications of the ventral root stimulation in the form of antidromic action potential recordings from motoneuron somas, as well as excitation of Renshaw cells.

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			<td >Na-kynurenate</td>
			<td >ABCAM</td>
			<td >ab120256</td>
			<td >dissolves better then other brands</td>
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			<td >Sigma</td>
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			<td >NaH2PO4</td>
			<td >Sigma</td>
			<td >P5655</td>
			<td></td>
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			<td >sucrose&#160;</td>
			<td >Sigma</td>
			<td >S9378</td>
			<td></td>
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			<td >NaHCO3&#160;</td>
			<td >Sigma</td>
			<td >S6014</td>
			<td></td>
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			<td >CaCl2&#160;</td>
			<td >G Biosciences</td>
			<td >R040</td>
			<td></td>
		</tr>
		<tr >
			<td >MgCl2&#160;</td>
			<td >Quality Biological</td>
			<td >351-033-721</td>
			<td></td>
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			<td >glucose&#160;</td>
			<td >Sigma</td>
			<td >G5767</td>
			<td></td>
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			<td >ascorbic acid&#160;</td>
			<td >Sigma</td>
			<td >A5960</td>
			<td></td>
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			<td >Na-pyruvate&#160;</td>
			<td >Sigma</td>
			<td >P2250</td>
			<td></td>
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			<td >K-gluconate&#160;</td>
			<td >Sigma</td>
			<td >P1847</td>
			<td></td>
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			<td >EGTA&#160;</td>
			<td >Sigma</td>
			<td >E3889</td>
			<td></td>
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			<td >HEPES&#160;</td>
			<td >Sigma</td>
			<td >H4034</td>
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			<td >NaCl</td>
			<td >Sigma</td>
			<td >S9888</td>
			<td></td>
		</tr>
		<tr >
			<td >Agar</td>
			<td>Sigma</td>
			<td>A9799</td>
			<td></td>
		</tr>
		<tr >
			<td >QX-314</td>
			<td >Alomone</td>
			<td>Q150</td>
			<td></td>
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			<td >Mg-ATP</td>
			<td>Sigma</td>
			<td >A9187</td>
			<td></td>
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			<td >CsOH</td>
			<td>Sigma</td>
			<td >232041</td>
			<td></td>
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		<tr >
			<td >Na-GTP</td>
			<td>Sigma</td>
			<td >51120</td>
			<td></td>
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			<td >gluconic acid</td>
			<td>Sigma</td>
			<td >G1951</td>
			<td></td>
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		<tr >
			<td >Cesium hydroxide solution</td>
			<td>Sigma</td>
			<td >232041</td>
			<td></td>
		</tr>
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			<td >KOH</td>
			<td>Sigma</td>
			<td >P5958</td>
			<td></td>
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			<td >Vannas Spring Scissors - 2.5mm&#160;</td>
			<td >FST</td>
			<td>15000-08</td>
			<td>only use for cutting the dura, might get damaged if cutting bones</td>
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			<td >Stimulator</td>
			<td>A-M Systems</td>
			<td colspan="2">Isolated Pulse Stimulator Model 2100</td>
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			<td >Vibratome</td>
			<td >Campden</td>
			<td >Vibrating Microtome 7000 - Model 7000smz-2</td>
			<td></td>
		</tr>
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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.

Confirmation of Motoneuron Identity Using Antidromic Action Potentials
Cell targeting
Motoneurons are found in the ventral horn (visible in red in Figure 2C). Start from the bundle of axons forming the ventral root and go up until the bundle disperses fully and one starts seeing large cells (long soma axis, above&#160;20 &#181;m). Achieve whole-cell recording of a round healthy looking cell using an electrode of initial resistance of 3 to 4 M&#937;. The internal solution used contained: 140 mM K-gluconate, 6 mM KCl, 10 mM HEPES, 1 mM EGTA, 0.1 mM CaCl2, 4 mM Mg-ATP, 0.3 mM Na2GTP. Adjust the pH to 7.3 with KOH, and the osmolarity to 285 - 295 mOsm by adding distilled water.
Confirmation of the motoneuron identity
Once whole-cell recording is achieved, apply a single biphasic stimulation of the ventral root (1 -&#160;50 V, 0.1 -&#160;0.3 msec) to elicit an antidromic action potential in the recorded cells of the ventral cord. Figures 3A1 and 3A2 show antidromic action potentials in voltage-clamp or current-clamp, respectively. When increasing the stimulation intensity from 1 to 5 V (Figure 3A1) and from 10 to 15 V (Figure 3A2) the antidromic action potential appears in an all-or-nothing fashion with a latency of 0.9 msec and 1.1 msec, respectively. Given that only motoneurons send their axons through the ventral roots, the antidromic action potential is a proof of the cell identity.
In a minority of motoneurons (about 10%), ventral root stimulation failed to elicit antidromic action potential but rather elicited orthodromic action potentials. Figures 3B1 and 3B2 show orthodromic action potentials in voltage-clamp or current-clamp, respectively. When increasing the stimulation from 20 to 30 V (Figure 3B1) and from 25 to 40 V (Figure 3B2), an action potential appears, following a previous excitatory post-synaptic current (EPSC) or excitatory post-synaptic potential (EPSP). The latencies of the action potentials are longer (5.1 msec and 5.3 msec, respectively). In these two cells, stimulating the ventral root elicited a feed-forward excitation (motoneurons send axon collaterals to themselves and to other motoneurons) that was strong enough to elicit an orthodromic action potential. In such cells, the failure to elicit an antidromic action potential (characterized by its all-or-nothing fashion and a latency shorter than 5 msec) does not allow us to conclude that these cells are motoneurons. Note that the required stimulation intensity to elicit&#160;feed-forward excitation is higher than the one needed to elicit an antidromic action potential. All stimulations shown here are 0.1 msec but sometimes, longer stimulation (up to 0.3 msec) may be required to elicit an antidromic action potential.
<img alt="Figure 3" src="/files/ftp_upload/54525/54525fig3.jpg" /> 
Figure 3. Motoneuron&#160;Responses to Ventral Root Stimulation 
A1: 1 V (red trace) and 5 V (black trace) stimulations. A2: 10 V (red trace) and 15 V (black trace) stimulations. B1: 20 V (red trace) and 30 V (black trace) stimulations. B2: 25 V (red trace) and 40 V (black trace) stimulations. Black dots indicate stimulus timing. Double arrowheads emphasize the difference in latencies between ortho- and antidromic spikes. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54525/54525fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Renshaw Cell Stimulation
Cell targeting
Renshaw cells are found in the ventral horn27,28. Visualize the axon bundles with differential interference contrast (DIC) imaging using a 20X and&#160;a 63X objectives. Start from the bundle of axons forming the ventral root and go up until the axons start to disperse but are still discernible (Figure 4A, arrows). Aim for the cells of intermediate size (about 10 - 15 &#181;m in diameter, Figure 4A, arrowheads). Achieve whole-cell recording of a round healthy-looking cell using an electrode of initial resistance of 5 to 7 M&#937;. Depending on the experiment, we used either Cs+-based or K+-based internal solutions10. The Cs-based solution prevented large potassium currents that were visible around -45 mV. In some experiments, we also added 5 mM QX-314 to block the sodium channels responsible for spiking in the recorded cell. The Cs+-based solution contains: 125 mM Cs-gluconate, 5 mM QX-314 Cl, 10 mM HEPES, 10 mM EGTA, 1 mM CaCl2, 4 mM Mg-ATP, and 0.4 mM Na-GTP, with the pH adjusted to 7.3 with CsOH. The K+-based solution should be used to analyze,&#160;both in voltage- and current-clamp, the Renshaw cell responses to ventral root stimulation in conditions approximating as much as possible the physiological ones. This solution contains:&#160;125 mM K-gluconate, 10 mM HEPES, 1 mM EGTA, 0.1 mM CaCl2, 4 mM Mg-ATP, 0.4 mM Na-GTP, with the pH adjusted to 7.3 with KOH. The stimulus intensity varied between 10 and 100 V and its duration varied between 50 and 300 &#181;sec.&#160;Bipolar pulses were used in all cases.
Response of the Renshaw cells to ventral root stimulation
Stimulation of the ventral root triggers the corelease of glutamate and acetylcholine from motoneuron&#160;collaterals onto the Renshaw cells10. Feed-forward inhibition mediated by GABA and glycine is also recruited10. Figure 4B displays responses to the single stimulation of the ventral root. The cell was recorded in a voltage-clamp mode with the cesium-based solution, and maintained at a&#160;voltage of -45 mV. QX-314 in the intracellular solution prevented the cell from firing and GABA and glycine responses were blocked by 3 &#181;M gabazine and 1 &#181;M strychnine, respectively. The inward synaptic current in Figure 4B is the sum of the glutamatergic and nicotinic currents.
<img alt="Figure 4" src="/files/ftp_upload/54525/54525fig4.jpg" /> 
Figure 4. Renshaw Cell Stimulation from the Ventral Root 
A: Images of the Renshaw cell pool acquired using an oblique condenser in the same slice taken at two different magnifications (20X and 40X objectives). Note the motoneuron axons merging into the ventral root (arrow). Putative Renshaw cells are indicated with arrowheads. Scale bars: 100 &#181;m in A1, 50 &#181;m in A2. B: Voltage-clamp recordings of a Renshaw cell following stimulation of the ventral root (black dot). The red trace is the average of the grey ones. Holding voltage was set at -45 mV. QX-314 in the intra-cellular solution prevented the cell from firing and GABA and glycine responses were blocked by 3 &#181;M gabazine and 1 &#181;M strychnine, respectively. Black dot indicates stimulus timing. <a href="https://www-jove-com.remotexs.ntu.edu.sg/files/ftp_upload/54525/54525fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about The Preparation of Oblique Spinal Cord Slices for Ventral Root Stimulation at JoVE.com

The Preparation of Oblique Spinal Cord Slices for 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 ...

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Nanyang Technological University

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Neuroscience

9.4K Views.  Centre de Neurophysique, Physiologie et Pathologie, Université Paris Descartes. We show how to prepare oblique slices of the spinal cord in young mice. This preparation allows for the stimulation of the ventral roots.

Video: The Preparation of Oblique Spinal Cord Slices for Ventral Root Stimulation

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

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

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

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

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled samples with little to no tissue damage enables us to analyze cell morphology and to compare individual samples to each other, hence allowing the identification of mutant anomalies.
In the demonstrated dissection method the nervous system remains mostly inside the adult fly. Through a dorsal incision, the abdomen and thorax are opened and most of the internal organs are removed. Only the dorsal side of the ventral nerve cord (VNC) and the cervical connective 
(CvC) containing the big axons of the giant fibers (GFs)1 are exposed, while the brain containing the GF cell body and dendrites remains2 in the 
intact head. In this preparation most nerves of the VNC should remain attached to their muscles. 
Following the dissection, the intracellular filling of the giant fiber (GF) with a fluorescent dye is demonstrated. In the CvC the GF axons are located at the dorsal surface and thus can be easily visualized under a microscope with differential interference contrast (DIC) optics. This allows the injection of the GF axons with dye at this site to label the entire GF including the axons and their terminals in the VNC. This method results in reliable and strong staining of the GFs allowing the neurons to be imaged immediately after filling with an epifluorescent microscope. Alternatively, the fluorescent signal can be enhanced using standard immunohistochemistry procedures3 suitable for high resolution confocal microscopy.

Whole Mount Preparation of the Adult Drosophila Ventral Nerve Cord for Giant Fiber Dye Injection

An in vivo dissection of the adult Drosophila ventral nerve cord (VNC) is demonstrated. This particular dissection method causes little damage to the VNC allowing the subsequent labeling of the giant fiber neurons with fluorescent dye for high resolution imaging.

To analyze the axonal and dendritic morphology of neurons, it is essential to obtain accurate labeling of neuronal structures. Preparing well labeled ...

Preparation of Parasagittal Slices for the Investigation of Dorsal-ventral Organization of the Rodent Medial Entorhinal Cortex

Computation in the brain relies on neurons responding appropriately to their synaptic inputs. Neurons differ in their complement and distribution of membrane ion channels that determine how they respond to synaptic inputs. However, the relationship between these cellular properties and neuronal function in behaving animals is not well understood. One approach to this problem is to investigate topographically organized neural circuits in which the position of individual neurons maps onto information they encode or computations they carry out1. Experiments using this approach suggest principles for tuning of synaptic responses underlying information encoding in sensory and cognitive circuits2,3.
The topographical organization of spatial representations along the dorsal-ventral axis of the medial entorhinal cortex (MEC) provides an opportunity to establish relationships between cellular mechanisms and computations important for spatial cognition. Neurons in layer II of the rodent MEC encode location using grid-like firing fields4-6. For neurons found at dorsal positions in the MEC the distance between the individual firing fields that form a grid is on the order of 30 cm, whereas for neurons at progressively more ventral positions this distance increases to greater than 1 m. Several studies have revealed cellular properties of neurons in layer II of the MEC that, like the spacing between grid firing fields, also differ according to their dorsal-ventral position, suggesting that these cellular properties are important for spatial computation2,7-10.
Here we describe procedures for preparation and electrophysiological recording from brain slices that maintain the dorsal-ventral extent of the MEC enabling investigation of the topographical organization of biophysical and anatomical properties of MEC neurons. The dorsal-ventral position of identified neurons relative to anatomical landmarks is difficult to establish accurately with protocols that use horizontal slices of MEC7,8,11,12, as it is difficult to establish reference points for the exact dorsal-ventral location of the slice. The procedures we describe enable accurate and consistent measurement of location of recorded cells along the dorsal-ventral axis of the MEC as well as visualization of molecular gradients2,10. The procedures have been developed for use with adult mice (&gt; 28 days) and have been successfully employed with mice up to 1.5 years old. With adjustments they could be used with younger mice or other rodent species. A standardized system of preparation and measurement will aid systematic investigation of the cellular and microcircuit properties of this area.

We describe procedures for preparation and electrophysiological recording from brain slices that maintain the dorsal-ventral axis of the medial entorhinal cortex (MEC). Because neural encoding of location follows a dorsal-ventral organization within the MEC, these procedures facilitate investigation of cellular mechanisms important for navigation and memory.

Computation in the brain relies on neurons responding appropriately to their synaptic inputs. Neurons differ in their complement and distribution of ...

Spinal Cord Transection in the Larval Zebrafish

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

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

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

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

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

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

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

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

Preparation of Primary Mixed Glial Cultures from Adult Mouse Spinal Cord Tissue

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information regarding glial activities at the cellular and molecular levels has been obtained through in vitro cell culture systems. The in vitro systems utilized, mainly include primary glia derived from neonatal brain cortical tissue and immortalized cell lines. However, these systems may not reflect the characteristics of spinal cord glial cells in vivo. In order to further investigate the roles of spinal cord glial cells in the development of peripheral nerve injury-induced neuropathic pain using a culture system that better reflects the in vivo condition, our laboratory has developed a method to establish primary spinal cord mixed glial cultures from adult mice. Briefly, spinal cords are collected from adult mice and processed through papain digestion followed by myelin removal with a density-gradient medium. Single cell suspensions are cultured in complete Dulbecco&#39;s modified Eagle media (cDMEM) supplemented with 2-mercaptoethanol (2-ME) at 35.9 oC. These culture conditions were optimized specifically for the growth of mixed glial cells. Under these conditions, cells are ready to be used for experimentation between 12 - 14 d&#160;(cells are usually in log phase during this time) after the establishment of the culture (D 0) and can be kept in culture conditions up to D 21. This culture system can be used to investigate the responses of spinal cord glial cells upon stimulation with various substances and agents. Besides neuropathic pain, this system can be used to study glial responses in other diseases that involve pathological changes of spinal cord glial cells.

The development of neuropathic pain involves pathological changes of spinal cord glial cells. A reliable glial culture system derived from adult spinal cord tissue and designed for studying these cells in vitro is lacking. Therefore, we show here how&#160;to establish primary mixed glial cultures from adult mouse spinal cord tissue.

It has been well-accepted that spinal cord glial responses contribute significantly to the development of neuropathic pain. Tremendous information ...

Hydraulic Extrusion of the Spinal Cord and Isolation of Dorsal Root Ganglia in Rodents

Traditionally, the spinal cord is isolated by laminectomy, i.e. by breaking open the spinal vertebrae one at a time. This is both time consuming and may result in damage to the spinal cord caused by the dissection process. Here, we show how the spinal cord can be extruded using hydraulic pressure. Handling time is significantly reduced to only a few minutes, likely decreasing protein damage. The low risk of damage to the spinal cord tissue improves subsequent immunohistochemical analysis. By performing hydraulic spinal cord extrusion instead of traditional laminectomy, the rodents can further be used for DRG isolation, thereby lowering the number of animals and allowing analysis across tissues from the same rodent. We demonstrate a consistent method to identify and isolate the DRGs according to their localization relative to the costae. It is, however, important to adjust this method to the particular animal used, as the number of spinal cord segments, both thoracic and lumbar, may vary according to animal type and strain. In addition, we illustrate further processing examples of the isolated tissues.

Here, we present a protocol for hydraulic extrusion of the spinal cord as well as identification and isolation of specific dorsal root ganglia (DRGs) in the same rodent. Compared to standard spinal cord isolation methods, this method is significantly faster and reduces the risk of tissue damage.

Traditionally, the spinal cord is isolated by laminectomy, i.e. by breaking open the spinal vertebrae one at a time. This is both time consuming and may ...

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

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

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

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

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

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

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

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

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

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

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.

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 ...