S.B.P is a recipient of a Loma Linda University Summer Undergraduate Research Fellowship.

The following protocol has been accepted and approved by the Institutional Animal Care and Use Committee (IACUC) of Loma Linda University. NIH guidelines for the ethical treatment of animals are followed in all animal experiments performed in the laboratory. All ethical standards were upheld by individuals performing this protocol.
1. Solutions
<ol>
	<li>Prepare artificial cerebral spinal fluid (aCSF).
	<ol>
		<li>Prepare fresh aCSF the evening before an experiment in 1 L batches using the following recipe: 7.250 g NaCl (124 mM), 0.224 g KCl (3 mM), 2.100 g NaHCO3 (25 mM), 0.069 g NaH2PO4&#160;&#8226; H2O (0.5 mM), 0.247 g MgSO4&#160;&#8226; 7H2O (1.0 mM), and 5.410 g D-glucose (30 mM), 0.221 g CaCl2&#160;&#8226; 2H2O (1.5 mM). Always add the CaCl2&#160;&#8226; 2H2O last.</li>
		<li>Dissolve the components into 1 L of deionized water. Stir for 20 - 30 min.</li>
		<li>Measure the pH of the aCSF and adjust to 7.40 &#177; 0.02 using small volumes (typically &#60;0.5 mL) of dilute NaOH, KOH, or HCl. 
		NOTE: Prepared aCSF can be stored in the refrigerator (4 &#176;C, pH 7.40 &#177; 0.02) up to three days before bacterial growth affects viability of the tissue during an experiment. Use 0.2 &#181;m filters to reduce contamination by fungus or bacteria present in the laboratory since experiments can last up to 36 h. For efficiency, the aCSF recipe described may be scaled up to 4 L batches following the same protocol and this volume will last for up to three days of experiments.</li>
	</ol>
	</li>
</ol>
2. Preparation of Dissection and Vibratome Rig
<ol>
	<li>Set up blade.
	<ol>
		<li>Use a fresh double-edged razor blades for cutting tissue on a vibratome. Wash the blade with 100% ethanol and rinse with deionized water before cutting or snapping the blade in half and inserting the blade into the mounting clamp on the vibratome.</li>
		<li>Change the blade every time a new tissue preparation is mounted on the vibratome for slicing or if it cuts into the paraffin slab while slicing. Any paraffin residue will make it nearly impossible to cut cleanly through neonatal neural tissue.</li>
	</ol>
	</li>
	<li>Set up paraffin wax slab.
	<ol>
		<li>Use any embedding-style paraffin wax. Place 10 g of embedding media in a heat-tolerant glass beaker and use low heat to melt the paraffin wax beads to liquid.</li>
		<li>Add approximately 0.5 g of graphite powder and mix solution thoroughly and uniformly into the liquid paraffin.</li>
		<li>Use a small plastic slab as the substrate for the paraffin. Cut a small (0.5 - 1 cm thick and approximately 2 x 2 cm2 wide) piece of plastic (polycarbonate or aluminum can also be used). Use a machinist&#39;s file to scratch grooves into the plastic asthis will ensure that the paraffin adheres to the plastic. 
		NOTE: Alternatively, use a heated 18 G hypodermic needle to place angled holes into the sheet to ensure that the paraffin-graphite mixture adheres to the plastic.</li>
		<li>Use the back of a cotton tip applicator to drip the paraffin-graphite mixture onto the plastic by repeatedly dipping the applicator into the molten paraffin-graphite mixture, dripping the slurry onto the plastic block, and building the paraffin-graphite to approximately 1.5 cm in thickness on top of the plastic.</li>
		<li>Once a sufficiently thick layer of the paraffin-graphite mix is deposited upon the block, set the block aside to cool and then shape to accommodate the brainstem-spinal cord.</li>
	</ol>
	</li>
</ol>
3. Dissection and Isolation of the Neuraxis
<ol>
	<li>Perform initial dissection and anesthetization. 
	NOTE: Animals used in this study may range in size from embryonic day 18 (E18) to postnatal day 10 - 20 in mice, or rats ranging from E18 to postnatal day 5/6. Rats or mice of any strain, treatment, or gender may be used, depending on the experimental design. Perform anesthesia and preliminary dissection under a fume hood since isoflurane is used (0.25 mL in a 25 mL chamber).
	<ol>
		<li>Weigh the pup and then place it in an anesthesia chamber containing 0.25 mL of isoflurane placed on a 2 x 2 inch2 piece of gauze. After the animal reaches a surgical plane of anesthesia (verified by toe pinch with no withdrawal reflex), pin the animal on a petri dish filled with paraffin or silicone elastomer. 
		NOTE: Neonatal rodents may also be anesthetized via cryoanesthesia13,14, isoflurane vaporization15, or injection16 balanced with oxygen.</li>
		<li>Place the animal ventral side down and make a midline incision from just behind the eyes to the mid-lumbar region of the spinal column with a number 10 or 11 scalpel blade or surgical scissors. Reflect the skin and decerebrate the animal at the level of the parietal/occipital suture (see Figure 1) and remove the skin transecting the animal below the diaphragm. Then remove the arms by cutting at the shoulder joint with scissors or a scalpel.</li>
		<li>Once the skin is removed, transfer the animal to a perfusion chamber with silicone elastomerin the bottom for further dissection and preparation of the brainstem-spinal cord.</li>
		<li>Bubble an aCSF reservoir (500 mL side-port bottle) continuously with chilled (4 &#176;C, pH 7.40 &#177; 0.02) aCSF and oxygenate with a mixture of 95% O2 and 5% CO2 (also called &#34;carbogen&#34;). During the dissection, use chilled aCSF to periodically flush the chamber, providing fresh oxygenated aCSF throughout isolation of the brainstem-spinal cord. 
		NOTE: Red blood cells can be seen in the remaining arteries and veins and, when sufficiently oxygenated, these are bright red.</li>
		<li>Use gravity-flow perfusion via tubing and a stopcock, to intermittently or continuously perfuse the tissue with oxygenated aCSF (bolus volume or via slow drip using a stopcock, approximately 0.5 - 1.0 mL/min). This oxygenates the tissue and maintains a clear surgical field.</li>
		<li>Remove excess fluids as needed by vacuum suction filtration system.</li>
		<li>Perform the dissection using a dissecting microscope. A continuous zoom model is preferred to allow fine adjustment of magnification and allow optimal visualization of the region of interest during each step of the dissection. 
		NOTE: Microsurgery tools recommended include a range of toothed tissue forceps, blunt forceps, #5 forceps, angled tissue forceps, a range of micro-dissecting scissors (spring scissors), and regular insect and micro-dissecting pins.</li>
		<li>Expose the brain via mid-line section of the skull along the sagittal suture then pin down the reflected flaps with micro-dissecting pins and pin the spinal column at the caudal end to the silicone-covered bottom of the dissection chamber using a 27 G needle. 
		NOTE: The rest of the dissection requires a dissection microscope to provide the clearest view of the tissue and landmarks used to ensure the maximum likelihood of obtaining rhythmic fictive output on the cranial rootlets of the brainstem and spinal rootlets. Perform these steps of the dissection using medium spring scissors or iris scissors and fine forceps.</li>
	</ol>
	</li>
	<li>Promptly transfer the isolated trunk of the animal to an aerated dissection chamber. Place the tissue dorsal side up with the rostral end (brain) facing the front of the chamber. Pin the tissue at the shoulders and most caudal end of the spinal cord.
	<ol>
		<li>Make a mid-sagittal incision through the skull following the parietal suture to avoid damaging the cortex and brainstem underlying the skull. 
		NOTE: Neonatal bone tissue is not fully calcified and is very brittle. The bone/tissue will be flexible to a degree, but tougher than the surrounding connective and muscle tissue.</li>
		<li>Use medium spring or iris scissors to snip the occipital sutures of the skull, beginning at the sagittal suture and working laterally. This will create &#34;flaps&#34; of skull that may be reflected and pinned to anchor the rostral part of the skull and provide some stability to the tissue (see Figure 2A).</li>
		<li>After reflecting the skull flaps, cut off the rest of the cerebral cortex, leaving the caudal portion of the cerebellum (vermis) relatively intact (Figure 2B).</li>
	</ol>
	</li>
	<li>Perform dorsal laminectomy.
	<ol>
		<li>Remove the musculature surrounding the skull and vertebral column using micro spring scissors and forceps. Remove tissue along the dorsal side of the rib cage, leaving the rib cage intact, as this will be pinned later to anchor the tissue during the ventral laminectomy, minimizing the chance of damage to the spinal cord.</li>
		<li>Carefully snip away the lateral processes of the vertebral laminae using medium spring scissors or fine iris scissors. Cut away any tissue overlying the pons and medulla. The vermis, cerebellum, pons, and beginning of the spinal cord will now be clearly visible (Figure 2C).</li>
	</ol>
	</li>
	<li>Perform ventral laminectomy.
	<ol>
		<li>Turn the tissue dorsal side down and pin at the rib cage and most caudal end of the spine. Pin the rostral side of the tissue down using the skull flaps (Figure 3A).</li>
		<li>Remove the ventral half of the rib cage, including the sternum and all abdominal organs using same scissors and forceps. Dissect away soft tissue attached to the ribcage exposing the ribs and spinal cord (Figure 3B).</li>
		<li>Remove the tongue, esophagus, trachea, larynx, and all other soft tissue and musculature overlying the base of the skull and the spinal column (as seen in&#160;Figure 3C).</li>
		<li>Dissect tissue overlying the hard pallet. Identify the hard palate by locating a rectangular plate of bone at the base of the skull with a V-shape indentation on it (Figure 3C). Cut along the midline of the palate, carefully lift it upwards, and perform a transverse cut to remove it (Figure 4A).</li>
		<li>Begin a ventral laminectomy by removing laminae exposing the ventral surface of the brainstem and the spinal cord from the first cervical vertebra to approximately thoracic vertebra 7 (T7) as seen in Figure 4B.</li>
		<li>At this stage, the ventral rootlets will be visible. Using small micro-dissection or spring scissors, take care to make cuts as close to the laminae and as far from roots&#39; origin at the spinal cord as possible; avoid stretching the roots.</li>
		<li>Snip 5 - 10 mm along both sides of the spinal column at the laminae. Do not cut too close to the spinal cord or into the vertebrae. This step allows removal of the cervical vertebrae to expose the spinal cord. Avoid cutting the rootlets.</li>
		<li>Snip rootlets approximately 20 - 25 mm (bilaterally) along the spinal column (approximately T7). Throughout this procedure avoid cutting into the spinal cord and manipulate the edges of the vertebrae as seen in Figure 4C.</li>
		<li>Remove C1, C2, and C3 by carefully lifting the rostral edge of each of the vertebrae and snipping closely underneath the bone. The closer to the bone that the cut is made, the longer the rootlet. Use hooked or bent forceps to aid achieving a firm grip on each cervical vertebra while making cuts (Figure 4C).</li>
		<li>Once the desired length of spinal cord is isolated and dissected from the vertebral column, make a transverse cut to remove the spinal cord (Figure 5A and Figure 5B).</li>
	</ol>
	</li>
	<li>Remove dura mater. 
	NOTE: The isolated brainstem-spinal cord can be used for en bloc in vitro recording as has been previously described3,7. With the brainstem-spinal cord removed from the vertebral column, the dura must be removed to provide optimal access for suction electrodes to perform recordings from the cut cranial and spinal rootles. Additionally, removal of the dura makes it possible to cleanly slice the brainstem and obtain rhythmically active thin slices for in vitro recording.
	<ol>
		<li>Carefully pin the isolated brainstem-spinal cord tissue dorsal side up to allow access to the dura surrounding the cranial rootles (Figure 5B). Pins should be applied through the lateral margin of the brainstem, rostral to the XII rootlets.</li>
		<li>Remove the dura from the dorsal surface of the brainstem by lifting the dura with fine forceps (#5) at the dorsal margin of the dura and cut from lateral-to-medial across the length of the brainstem with fine spring scissors. Be careful to avoid cutting into the brainstem itself. Gently lift blood vessels from the brainstem surface and cut to avoid impediment of the vibratome blade when sectioning the brainstem.</li>
		<li>Carefully dissect dura from the medial and lateral aspects of the brainstem, as the cranial nerve rootlets pass through the dura and are easily ripped away when the dura is lifted. Minimize the likelihood of the rootlets being pulled off by gently cutting around them with small spring scissors.</li>
		<li>Flip the brainstem-spinal cord so that the ventral surface faces up and the cranial rootlets are clearly visible. Dissect the dura from the dorsal side of the brainstem by gently lifting the dura directly over the area postrema, cutting from rostral to caudal. The area postrema will appear slightly pink due to the large number of microcapillaries perfusing this region.</li>
		<li>Use a fine insect pin to tease away any remaining dura and remove remaining blood vessels in close proximity to cranial and cervical rootlets.</li>
	</ol>
	</li>
</ol>
4. Slice Protocol
<ol>
	<li>After removing the dura, place the brainstem in the center of the paraffin platform on the plastic block (hereafter referred to as the &#34;cutting block&#34;). Pin the caudal end of the brainstem through the distal spinal cord using fine insect pins that have been trimmed to no more than 1 cm in length as shown in Figure 5C.</li>
	<li>Align the paraffin-covered cutting block with the pinned brainstem in the vibratome block holder so that the blade cuts perpendicular to the rostral face of the brainstem.</li>
	<li>Make an initial slice to remove the uneven, extraneous tissue on the rostral-most end. This initial cut can be 200 - 300 &#181;m typically but carefully avoid removal of IX, X, and XII cranial rootlets. This step will typically reveal the caudal extent of the facial nucleus (VII) and the glossopharyngeal rootlets and makes it possible to align the brainstem so that its face is par-planar to the vibratome blade. Make small adjustments as necessary to ensure par-planarity and make small cuts to remove tissue that is uneven. 
	NOTE: Glossopharyngeal (IX) rootlets will be visible at the lateral edge of the brainstem as one cuts each successive slice, and these provide a landmark for the rostro-caudal distance to the brainstem neural circuitry necessary for rhythm-generation. On the ventral side of the brainstem, the hypoglossal rootlets (XII) should also be visible slightly caudal to the cut face showing the IX rootlets. A final landmark will be the obex (the point in the human brain at which the fourth ventricle narrows to become the central canal of the spinal cord) on the dorsal face of the brainstem. With these three points of reference visible, the correct cutting plane is established (see Figure 6A) and a rhythmically-active slice will be reliably obtained.</li>
	<li>Continue cutting rostral-to-caudal until the IX rootlets are near the surface of the cut face of the brainstem.</li>
	<li>Refer to Figure 6 for landmarks and correct orientation. Adjust the paraffin-slab in the vibratome clamp so that the next angle of transection will create a very slight &#34;wedge&#34; shape to the cut slice and cut slices of approximately 100 - 200 &#181;m until the landmarks described can be clearly seen (Figure 6A). 
	NOTE: Cutting this slight wedge increases the number of XII motor neurons captured in the slice and maximizes the likelihood of obtaining rhythmic activity during recording. Using the landmarks described (rostral rootlets from the glossopharyngeal nerve bundle, rootlets from the hypoglossal nerve bundle, the obex) will ensure that the slice reproducibly contains at least a large portion of the pBC (Figure 6B).</li>
	<li>Cut a 300 - 500 &#181;m slice of brainstem from these rostral neuroanatomical markers to capture the pBC and associated transmission circuitry. 
	NOTE: An &#34;ideal&#34; slice will contain the most rostral rootlet of the hypoglossal rootlet bundle to reliably obtain inspiratory activity without resorting to surface recordings using a suction electrode over the rhythm-generating/transmitting loci within the brainstem.</li>
</ol>
5. Recording Procedures
<ol>
	<li>Place the slice in a recording chamber, perfuse continuously with aCSF (0.5 - 1.0 mL/min) and use suction or extracellular electrodes to record population activity from the XII rootlets or from the pBC, XII PMNs, or XII motoneurons. For detailed recording procedures, see previous publications on brainstem-spinal cord electrophysiology and patch clamping10,11.</li>
</ol>

<ol>
	<li>Hilaire, G., Duron, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Maturation+of+the+Mammalian+Respiratory+System.">Maturation of the Mammalian Respiratory System.</a> Physiological Reviews. 79, 325-360 (1999).</li><li>Pinkerton, K. E., Joad, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+Mammalian+Respiratory+System+and+Critical+Windows+of+Exposure+for+Children's+Health.">The Mammalian Respiratory System and Critical Windows of Exposure for Children's Health.</a> Environmental Health Perspectives. 108, 6 (2000).</li><li>Smith, J. C., Ellenberger, H. H., Ballanyi, K., Richter, D. W., Feldman, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Pre-B&#246;tzinger+complex:+a+brainstem+region+that+may+generate+respiratory+rhythm+in+mammals.">Pre-B&#246;tzinger complex: a brainstem region that may generate respiratory rhythm in mammals.</a> Science. 254, 726-729 (1991).</li><li>Smith, J. C., 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=Respiratory+rhythm+generation+in+neonatal+and+adult+mammals:+the+hybrid+pacemaker-network+model.">Respiratory rhythm generation in neonatal and adult mammals: the hybrid pacemaker-network model.</a> Respiration Physiology. 122, 131-147 (2000).</li><li>Ramirez, J. M., Schwarzacher, S. W., Pierrefiche, O., Olivera, B. M., Richter, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Selective+lesioning+of+the+cat+pre-B&#246;tzinger+complex+in+vivo+eliminates+breathing+but+not+gasping.">Selective lesioning of the cat pre-B&#246;tzinger complex in vivo eliminates breathing but not gasping.</a> The Journal of Physiology. 507, 895-907 (2004).</li><li>Butera, R. J., Rinzel, J., Smith, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Models+of+Respiratory+Rhythm+Generation+in+the+pre-B&#246;tzinger+Complex:+II.+Populations+of+Coupled+Pacemaker+Neurons.">Models of Respiratory Rhythm Generation in the pre-B&#246;tzinger Complex: II. Populations of Coupled Pacemaker Neurons.</a> Journal of Neurophysiology. 82, 398-415 (1999).</li><li>Suzue, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Respiratory+rhythm+generation+in+the+in+vitro+brain+stem-spinal+cord+preparation+of+the+neonatal+rat.">Respiratory rhythm generation in the in vitro brain stem-spinal cord preparation of the neonatal rat.</a> Journal of Physiology (London). 354, 173-183 (1984).</li><li>Feldman, J. L., Del Negro, C. A., Gray, P. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Understanding+the+rhythm+of+breathing:+so+near,+yet+so+far.">Understanding the rhythm of breathing: so near, yet so far.</a> Annual Review of Physiology. 75, 423-452 (2013).</li><li>Rekling, J. C., Shao, X. M., Feldman, J. L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrical+Coupling+and+Excitatory+Synaptic+Transmission+between+Rhythmogenic+Respiratory+Neurons+in+the+PreB&#246;tzinger+Complex.">Electrical Coupling and Excitatory Synaptic Transmission between Rhythmogenic Respiratory Neurons in the PreB&#246;tzinger Complex.</a> Journal of Neuroscience. 20, 113 (2000).</li><li>Rousseau, J. -. P., Caravagna, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Electrophysiology+on+Isolated+Brainstem-spinal+Cord+Preparations+from+Newborn+Rodents+Allows+Neural+Respiratory+Network+Output+Recording.">Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording.</a> Journal of Visualized Experiments. , e53071 (2015).</li><li>Segev, A., Garcia-Oscos, F., Kourrich, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Whole-cell+Patch-clamp+Recordings+in+Brain+Slices.">Whole-cell Patch-clamp Recordings in Brain Slices.</a> Journal of Visualized Experiments. (112), e54024 (2016).</li><li>Koizumi, H., 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=Functional+Imaging,+Spatial+Reconstruction,+and+Biophysical+Analysis+of+a+Respiratory+Motor+Circuit+Isolated+In+Vitro.">Functional Imaging, Spatial Reconstruction, and Biophysical Analysis of a Respiratory Motor Circuit Isolated In Vitro.</a> Journal of Neuroscience. 28, 2353-2365 (2008).</li><li>Danneman, P., Mandrell, T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Evaluation+of+five+agents/methods+for+anesthesia+of+neonatal+rats.">Evaluation of five agents/methods for anesthesia of neonatal rats.</a> Laboratory Animal Science. 47, 386-395 (1997).</li><li>Umezawa, N., 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=Orexin-B+antagonized+respiratory+depression+induced+by+sevoflurane,+propofol,+and+remifentanil+in+isolated+brainstem-spinal+cords+of+neonatal+rats.">Orexin-B antagonized respiratory depression induced by sevoflurane, propofol, and remifentanil in isolated brainstem-spinal cords of neonatal rats.</a> Respiratory Physiology &amp; Neurobiology. 205, 61-65 (2015).</li><li>Bierman, A. M., Tankersley, C. G., Wilson, C. G., Chavez-Valdez, R., Gauda, E. B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Perinatal+hyperoxic+exposure+reconfigures+the+central+respiratory+network+contributing+to+intolerance+to+anoxia+in+newborn+rat+pups.">Perinatal hyperoxic exposure reconfigures the central respiratory network contributing to intolerance to anoxia in newborn rat pups.</a> Journal of Applied Physiology. 116, 47-53 (2014).</li><li>Formenti, A., Zocchi, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Error+signals+as+powerful+stimuli+for+the+operant+conditioning-like+process+of+the+fictive+respiratory+output+in+a+brainstem-spinal+cord+preparation+from+rats.">Error signals as powerful stimuli for the operant conditioning-like process of the fictive respiratory output in a brainstem-spinal cord preparation from rats.</a> Behavioural Brain Research. 272, 8-15 (2014).</li><li>Negro, C. A. D., 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=Sodium+and+Calcium+Current-Mediated+Pacemaker+Neurons+and+Respiratory+Rhythm+Generation.">Sodium and Calcium Current-Mediated Pacemaker Neurons and Respiratory Rhythm Generation.</a> Journal of Neuroscience. 25, 446-453 (2005).</li><li>Johnson, S. M., Koshiya, N., Smith, J. C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Isolation+of+the+Kernel+for+Respiratory+Rhythm+Generation+in+a+Novel+Preparation:+The+Pre-B&#246;tzinger+Complex+"Island".">Isolation of the Kernel for Respiratory Rhythm Generation in a Novel Preparation: The Pre-B&#246;tzinger Complex "Island".</a> Journal of Neurophysiology. 85, 1772-1776 (2001).</li><li>Funk, G. D., 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=Functional+respiratory+rhythm+generating+networks+in+neonatal+mice+lacking+NMDAR1+gene.">Functional respiratory rhythm generating networks in neonatal mice lacking NMDAR1 gene.</a> Journal of Neurophysiology. 78, 1414-1420 (1997).</li><li>Campos, M., Bravo, E., Eugen&#237;n, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Respiratory+dysfunctions+induced+by+prenatal+nicotine+exposure.">Respiratory dysfunctions induced by prenatal nicotine exposure.</a> Clinical and Experimental Pharmacology and Physiology. 36, 1205-1217 (2009).</li><li>Ballanyi, K., Volker, A., Richter, D. W. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Anoxia+induced+functional+inactivation+of+neonatal+respiratory+neurones+in+vitro.">Anoxia induced functional inactivation of neonatal respiratory neurones in vitro.</a> NeuroReport. 6, 165-168 (1994).</li><li>Ruangkittisakul, A., 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=High+Sensitivity+to+Neuromodulator-Activated+Signaling+Pathways+at+Physiological+[K+]+of+Confocally+Imaged+Respiratory+Center+Neurons+in+On-Line-Calibrated+Newborn+Rat+Brainstem+Slices.">High Sensitivity to Neuromodulator-Activated Signaling Pathways at Physiological [K+] of Confocally Imaged Respiratory Center Neurons in On-Line-Calibrated Newborn Rat Brainstem Slices.</a> Journal of Neuroscience. 26, 11870-11880 (2006).</li><li>Rybak, I. A., 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=Modeling+the+ponto-medullary+respiratory+network.">Modeling the ponto-medullary respiratory network.</a> Respiratory Physiology &amp; Neurobiology. 143, 307-319 (2004).</li><li>Ruangkittisakul, A., Kottick, A., Picardo, M. C. D., Ballanyi, K., Del Negro, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Identification+of+the+pre-B&#246;tzinger+complex+inspiratory+center+in+calibrated+'sandwich'+slices+from+newborn+mice+with+fluorescent+Dbx1+interneurons.">Identification of the pre-B&#246;tzinger complex inspiratory center in calibrated 'sandwich' slices from newborn mice with fluorescent Dbx1 interneurons.</a> Physiological Reports. 2, (2014).</li><li>Paxinos, G., Watson, C. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=.">.</a> The Rat Brain in Stereotaxic Coordinates. , (1997).</li><li>Franklin, K. B. J., 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=.">.</a> The Mouse Brain in Stereotaxic Coordinates. , (2008).</li></ol>

The authors have nothing to disclose.

Adapting the protocol presented here into an en bloc or slice workflow is advantageous for laboratories and studies that would like to utilize either en bloc brainstem-spinal cord and/or thin slice preparations for electrophysiology recordings. The dissection and slice method presented, combined with methods previously reported by others17,18,19, will allow reproducible preparation of robust and viable tissue that is widely adaptable to a range of experiments using the rodent hindbrain or spinal cord. The dissection presented is highly detailed and includes dorsal and ventral laminectomy as well as extensive detail regarding orientation of the brainstem-spinal cord when preparing to cut slices. The detailed dissection and slice protocol presented allows the researcher to include all circuitry necessary for the generation and transmission of inspiratory rhythm. The main neural populations/structures that can be captured using this method include: the hypoglossal premotor neurons, the hypoglossal motor nucleus, and the preB&#246;tzinger complex. Note that regions of the brainstem that contain central CO2 sensors or expiratory rhythm-generating circuits (RTN/pFRG) are not included in the preparation detailed here20,21. Once the en bloc preparation or slice has been obtained, rhythmic activity may be obtained using suction electrodes, surface electrodes, or single-cell recording methods such as extracellular unit or patch-clamping methods3,10,11.
The goal of this protocol is to provide a clear, easy-to-follow method for generating either a brainstem-spinal cord preparation or a thin, rhythmically-active slice. This protocol should be valuable to both experienced and new researchers interested in incorporating rhythmic brainstem/slice preparations in their armamentarium, as there are several critical steps detailed within the procedures that facilitate capture of robust, reproducible, and long-lasting rhythmically active slices.
Once the researcher has become comfortable with identification of the anatomical landmarks and the skills necessary for dissection, the speed of the dissection will increase, and the procedures can be optimized for each individual investigator. The protocol detailed here was taught to the senior author (CGW) during his post-doctoral work with Jeffrey Smith, the originator of the rhythmic brainstem slice preparation3. To ensure rhythmic activity and viability in both the slices and en bloc preparations, all procedures to generate a slice should be finished within approximately 30 min&#160;from beginning of the procedure to a slice in the recording chamber. With continuous perfusion, there is very little impact on tissue viability but minimizing the time for dissection allows for longer recording and data acquisition. For respiratory studies, a slice that is as thin as approximately 280 &#181;m thick3 will have rhythmic, robust inspiratory activity, but slices may range up to 550 &#181;m17,18,19,22. Careful practice is required to develop the necessary skills to perform this procedure and minimize the variability in thickness and rostral-caudal position of each slice. Including or excluding certain cell populations will influence the quality and robustness of rhythmic activity obtained in later recording as both excitatory and inhibitory nuclei in the brainstem can impact the &#34;quality&#34; of the rhythm recorded from the slice3.
Finally, it is also critical that aCSF is prepared exactly according to protocol, and is at a standardized pH20, temperature, and oxygenation level. Anoxic preparations will not be rhythmically active and will affect data collection21.
There are several key steps in the procedure that facilitate an investigator capture of viable and rhythmically active preparations. During the ventral laminectomy, keeping the dorsal rib cage intact allows extra leverage and provides more tissue for securing the preparation while performing the relatively delicate isolation of the brainstem-spinal cord from the vertebral column. The dissection steps detailed here allow the investigator to easily produce en bloc preparations or thin slices so, by necessity, extra steps are detailed that can be omitted to obtain just a thin slice. Other useful tips include, while performing the ventral laminectomy, it is important to gain a firm grip on the vertebral body to lift it upwards while severing the nerve rootlets. Once the brainstem-spinal cord has been dissected and extracted, the dura mater and remaining vasculature must be dissected off of the surface of the neural tissue. Blood vessels and dura are tough and can prevent the vibratome blade from cutting a clean slice. Dissection of the dura is not vital for the en bloc preparation, but is recommended to optimize nerve-suction electrode coupling. Finally, careful and methodical practice in the use of the vibratome is necessary for cutting a clean, reproducible slice. This protocol provides a very comprehensive list of detailed steps for the dissection and cutting procedures. The emphasis here is to provide a step-by-step detailed list for an investigator to use as a reliable foundation and training tool from which they can modify their laboratory&#39;s procedures as needed.
As a whole, this methodology represents a thirty-year evolution of methods of in vitro electrophysiology. The scope of this paper focuses on the standardization of en bloc and slice brainstem-spinal cord recordings that are typically used for studying inspiratory activity in P0-P5 rats and mice22. However, this dissection process may be utilized across a range of animal sizes, ages, and species for a wide variety of studies, including E18 rats/mice, neonatal rats and mice (postnatal days 0 to 6), and adult mice. Future studies may use the methods presented to streamline protocols across species and ages, allowing studies to interrogate mechanisms in rhythm generation or other physiological process across both species and ages. Ultimately, changing the orientation, thickness, or location of the slice or en bloc preparation will influence rhythmic activity23. Extensive literature has been published in the past concerning the neural populations captured by slice procedures and there are stereotaxic rat and mouse atlases available to provide more detail concerning the neural circuitry discussed here24,25,26. The procedures presented here provide extensive detail with clear illustrations that allow a new investigator to learn to cut rhythmic slices from landmarks readily visible under a laboratory dissecting scope.

The respiratory neural network of the brainstem provides a fertile domain for understanding the general characteristics of rhythmic neural networks. In particular, the interest is in the development of neonatal rodent breathing and understanding how the breathing rhythm develops. This may be done using a multi-level approach, including in vivo whole animal plethysmography, in vitro en bloc nerve recordings, and in vitro slice recordings that contain the breathing rhythm generator. Reductionist in vitro en bloc and slice recordings are an advantageous method to use when interrogating the mechanisms behind respiratory rhythmogenesis and neural circuitry in the brainstem-spinal cord region of developing rodents. The developing respiratory system includes approximately 40 cell types, characterized by firing pattern, including those of the central respiratory1,2. The central respiratory network includes a group of rhythmically active neurons located in the rostral ventrolateral medulla1,3. Mammalian respiratory rhythmogenesis is generated from an autorhythmic interneuron network dubbed the preB&#246;tzinger complex (pBC), which has been localized experimentally via both slice and en bloc preparations of neonatal mammalian brainstem-spinal cords3,4,5,6,7,8. This region serves a similar function to the sinoatrial node (SA) in the heart and generates an inspiratory timing system to drive respiration. From the pBC, the inspiratory rhythm is carried to other regions of the brainstem (including the hypoglossal motor nucleus) and spinal motor pools (such as the phrenic motor neurons that drive the diaphragm)9.
Rhythmic activity may be obtained using brainstem spinal cord en bloc preparations or slices from a variety of cell populations, including C3-C5 nerve rootlets, XII nerve rootlets, hypoglossal motor nucleus (XII MN), hypoglossal premotor neurons (XII pMN), and the pBC3,10,11,12. While these methods of data collection have been successful across a handful of laboratories, many of the protocols are not presented in a way that is fully reproducible for new researchers entering the field. Obtaining viable and rhythmically active en bloc and slice preparations requires an acute attention to detail through all steps of the dissection and slice cutting protocol. Previous protocols extensively describe the various recording procedures and electrophysiology, yet lack detail in the most critical part of obtaining a viable tissue preparation: performing the brainstem-spinal cord dissection and slice procedure.
Efficiently obtaining a rhythmically-active and viable en bloc or slice preparation brainstem-spinal cord electrophysiology recordings requires that all steps be performed correctly, carefully, and swiftly (typically, the whole procedure related here can be performed in approximately 30 min). Critical points of the brainstem-spinal cord electrophysiology protocol that have not been previously well described include the dissection of nerve rootlets and the slicing procedure on the vibratome. This protocol is the first to stepwise visually communicate the brainstem-spinal cord dissection for both new researchers and experts in the field. This protocol also thoroughly explains surgical techniques, landmarks, and other procedures to assist future researchers in standardizing slices and en bloc preparations to contain the exact circuitry desired in each experiment. The procedures presented here can be used in both rat and mouse neonatal pups.

<table><tbody><tr><td>NaCl</td><td>Fisher Scientific</td><td>S271-500</td><td></td></tr><tr><td>KCl</td><td>Sigma Aldrich</td><td>P5405-1KG</td><td></td></tr><tr><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td><td>Fisher Scientific</td><td>BP328-1</td><td></td></tr><tr><td>NaH&lt;sub&gt;2&lt;/sub&gt;PO&lt;sub&gt;4&lt;/sub&gt; &amp;bull;H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma Aldrch</td><td>S9638-25G</td><td></td></tr><tr><td>CaCl&lt;sub&gt;2&lt;/sub&gt;&amp;bull;2H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma Aldrich</td><td>C7902-500G</td><td></td></tr><tr><td>MgSO&lt;sub&gt;4&lt;/sub&gt;&amp;bull;7H&lt;sub&gt;2&lt;/sub&gt;O</td><td>Sigma Aldrich</td><td>M7774-500G</td><td></td></tr><tr><td>D-Glucose</td><td>Sigma Aldrich</td><td>G8270-1KG</td><td></td></tr><tr><td>Cold-Light source Halogen lamp 150 W</td><td>AmScope</td><td>H2L50-AY</td><td></td></tr><tr><td>Dissection Microscope</td><td>Leica</td><td>M-60</td><td></td></tr><tr><td>Vibratome 1000 Plus</td><td>Vibratome</td><td>W3 69-0353</td><td></td></tr><tr><td>Magnetic Base</td><td>Kanetic</td><td>MB-B-DG6C</td><td></td></tr><tr><td>Isoflurane, USP</td><td>Patterson Veterinary</td><td>NDC 14043-704-06</td><td></td></tr><tr><td>Sword Classic Double Edge Blades</td><td>Wilkinson</td><td>97573</td><td></td></tr><tr><td>Histoclear</td><td>Sigma-Aldrich</td><td>H2779</td><td></td></tr><tr><td>Dumont #5 Fine Forceps</td><td>Fine Science Tools</td><td>11254-20</td><td></td></tr><tr><td>Dumont #5/45 Forcep</td><td>Fine Science Tools</td><td>11251-35</td><td></td></tr><tr><td>Scalpel Blades #10</td><td>Fine Science Tools</td><td>10010-00</td><td></td></tr><tr><td>Scalpel Handel #3</td><td>Fine Science Tools</td><td>10003-12</td><td></td></tr><tr><td>Spring Scissors Straight&amp;nbsp;</td><td>Fine Science Tools</td><td>15024-10</td><td></td></tr><tr><td>Narrow Pattern Forcep Serrated/straight</td><td>Fine Science Tools</td><td>11002-12</td><td></td></tr><tr><td>Castroviejo Micro Dissecting Spring Scissors; Straight</td><td>Roboz</td><td>RS-5650</td><td></td></tr><tr><td>Vannas Scissors 3&quot; Curved</td><td>Roboz</td><td>RS-5621</td><td></td></tr><tr><td>Insect pins, 0</td><td>Fine Science Tools/8840604</td><td>26000-35&amp;nbsp;</td><td></td></tr><tr><td>Insect pins, 0, SS</td><td>Fine Science Tools</td><td>26001-35</td><td></td></tr><tr><td>Insect pins, 00</td><td>Fine Science Tools</td><td>26000-30</td><td></td></tr><tr><td>Insect pins, 00, SS</td><td>Fine Science Tools</td><td>26001-30</td><td></td></tr><tr><td>Insect pins, 000</td><td>Fine Science Tools</td><td>26000-25</td><td></td></tr><tr><td>Insect pins, 000, SS</td><td>Fine Science Tools</td><td>26001-25</td><td></td></tr><tr><td>Minutien pins, 0.10 mm</td><td>Fine Science Tools</td><td>26002-10</td><td></td></tr><tr><td>Minutien pins, 0.15 mm</td><td>Fine Science Tools</td><td>26002-15</td><td></td></tr><tr><td>Minutien pins, 0.2 mm</td><td>Fine Science Tools</td><td>26002-20</td><td></td></tr><tr><td>Fisher Tissue prep Parafin&amp;nbsp;</td><td>fisher</td><td>T56-5</td><td></td></tr><tr><td>Graphite&amp;nbsp;</td><td>fisher&amp;nbsp;</td><td>G67-500</td><td></td></tr><tr><td>Delrin Plastic&amp;nbsp;</td><td>Grainger</td><td>3HMT2</td><td></td></tr><tr><td>18 Gauge Hypodermic Needle</td><td>BD</td><td>305195</td><td></td></tr></tbody></table>

preparation of rhythmically-active in vitro neonatal rodent brainstem-spinal cord and thin slice

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a signal driving the rhythmic contraction of inspiratory muscles. Rhythmic neural activity generated in the pBC and carried to other neuronal pools to drive the musculature of breathing may be studied using various approaches, including en bloc nerve recordings and transverse slice recordings. However, previously published methods have not extensively described the brainstem-spinal cord dissection process in a transparent and reproducible manner for future studies. Here, we present a comprehensive overview of a method used to reproducibly cut rhythmically-active brainstem slices containing the necessary and sufficient neuronal circuitry for generating and transmitting inspiratory drive. This work builds upon previous brainstem-spinal cord electrophysiology protocols to enhance the likelihood of reliably obtaining viable and rhythmically-active slices for recording neuronal output from the pBC, hypoglossal premotor neurons (XII pMN), and hypoglossal motor neurons (XII MN). The work presented expands upon previous published methods by providing detailed, step-by-step illustrations of the dissection, from whole rat pup, to in vitro slice containing the XII rootlets.

The method presented here allows a researcher interested in obtaining rhythmically active slices of brainstem to reproducibly and reliably cut a viable, robust slice that will allow recording of fictive motor output for many hours. All of the minimally necessary neural circuit elements for generating and transmitting inspiratory rhythm can be captured in a thin slice using this method. These elements include: the preB&#246;tzinger Complex, premotor neurons projecting to the hypoglossal motor neurons (pXII MNs) and the hypoglossal motor neurons (XII MNs), and hypoglossal nerve rootlets. The pBC, XIIn, and C4 nerve rootlets are commonly used for inspiratory rhythm recordings, as illustrated in Figure 7.
Successful use of this procedure will produce a viable and rhythmically active en bloc preparation in 10 - 15 min, or a rhythmically-active slice in &#60;30 min. After isolation of the en bloc brainstem-spinal cord or a thin slice, 15 min of equilibration in the recording chamber is sufficient for the production of fictive motor output. In the case of the slice, increasing the extracellular [K+] to approximately 8 - 9 mM will produce robust neural drive that can last for 24 to 36 h in this laboratory&#39;s experience. The preparation should be continuously superfused with carbogenated (95% O2/5% CO2) and heated aCSF (~ 27 &#176;C). Successful dissections are performed quickly and avoid pinching, stretching, or damaging nerve rootlets used for recordings. For optimal results, all steps in the procedure are performed swiftly and the tissue must be bathed or continuously perfused with carbogenated aCSF when performing the dissection and tissue isolation (as described above). For extensive details concerning the neuroanatomy and precise atlases of the brainstem, please see the work of Ruangkittisakul et al.13,14, Ballanyi15 and colleagues. The protocol presented here is one method that has proven reliable in the senior author&#39;s laboratory and this report provides a graphical, step-by-step method for generating these slices relying only upon surface landmarks visible on the brainstem or within the brainstem as detailed here.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/58870/58870fig1.jpg" /> 
Figure 1: Initial dissection for the brainstem-spinal cord extraction. (A) Anesthetized pup pinned for dissection. Dashed lines indicate incision guidelines for sagittal incision into skin and transverse cuts caudal to eyes and below the diaphragm. (B) Guide for removing skin and forelimbs from animal. (C) Dorsal aspect of skinned rodent. Dashed lines indicate incision guidelines for skull-flap &#34;clamshell&#34; exposure. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58870/58870fig1large.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/58870/58870fig2.jpg" /> 
Figure 2: Stages of the dorsal laminectomy. (A) Dashed lines indicate incision guidelines for removing the cerebrum. (B) Result after removing tissue. Dashed lines indicate incision guidelines for dorsal laminectomy. (C) Exposed brainstem-spinal cord following dorsal laminectomy. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58870/58870fig2large.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/58870/58870fig3.jpg" /> 
Figure 3: Stages of initial ventral dissection. (A) Ventral side of a skinned rodent with severed forebrain and abdomen. Dashed lines indicate incision guidelines for removing the xyphoid process and opening the rib cage to subsequently remove abdominal and thoracic cavity organs. (B) Abdominal and thoracic cavity organs removed. (C) Oropharyngeal structures removed. Dashed lines indicate incision guidelines for hard palate removal. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58870/58870fig3large.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/58870/58870fig4.jpg" /> 
Figure 4:&#160;Stages of the ventral laminectomy. (A) Hard palate and remaining skull parts removed. Incision guidelines indicate how to remove remaining tissue surrounding the brainstem-spinal cord. (B) First cervical vertebra (C1) exposed. (C) Demonstration on lifting C1 and making a transverse cut to the vertebral body to remove it. Nerves are carefully cut flush to the dorsal side of the vertebral body before removing it. This is the most critical step in the dissection. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58870/58870fig4large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/58870/58870fig5.jpg" /> 
Figure 5:&#160;Preparation of brainstem-spinal cord for slicing. (A) C1-C3 removed from spinal cord. Incision guidelines indicate where to sever the brainstem-spinal cord from the caudal spinal cord. (B) Ventral side of brainstem-spinal cord with labeled nerve rootlets. Incision guidelines indicate recommended transection or remaining rostral structures at the ponto-medullary before recording from the brainstem-spinal cord region. (C) Paraffin slab setup for slicing on the vibratome. The slicing region includes tissue between cranial nerves IX and XII. Do not place micro-pins into the slicing region. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58870/58870fig5large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 6" class="xfigimg" src="/files/ftp_upload/58870/58870fig6.jpg" /> 
Figure 6: Landmarks used for obtaining slices with an intact hypoglossal motor nucleus and PreB&#246;tzinger Complex. (A) Transverse view of brainstem-spinal cord after initial cut has been made rostral to the glossopharyngeal (IX) rootlets. Incision guideline indicates transection level just caudal to the hypoglossal (XII) rootlets. (B) Orientation of paraffin block to blade before cutting a slice containing the pBC and the hypoglossal motor nucleus. Creating a trapezoidal &#34;wedge&#34; shape cut maximizes the likelihood of capturing enough inspiratory neurons in the slice to obtain rhythmic activity during recording. The wedge will include the pBC, the hypoglossal premotor neurons, and the hypoglossal motor neurons, which collectively provide the necessary circuitry to transmit rhythmic drive, which is an index of respiratory rhythmogenesis. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58870/58870fig6large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 7" class="xfigimg" src="/files/ftp_upload/58870/58870fig7.jpg" /> 
Figure 7: Representative recordings from en bloc or slice preparations. (A) Integrated trace from XII rootlet. (B) Integrated trace from the pBC. Recording from the pBC using an en bloc preparation will require a blind patch clamp. (C) Integrated trace from the C4 nerve rootlet. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/58870/58870fig7large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice at JoVE.com

Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice

This protocol both visually communicates the brainstem-spinal cord preparation and clarifies the preparation of brainstem transverse slices in a comprehensive step-by-step manner. It was designed to increase reproducibility and enhance the likelihood of obtaining viable, long lasting, rhythmically-active slices for recording neural output from the respiratory regions of the brainstem.

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preB&#246;tzinger complex (pBC), which produces a ...

preparation-rhythmically-active-vitro-neonatal-rodent-brainstem

Universidad San Sebastian

Research

JoVE Journal

Neuroscience

11.4K Views.  Loma Linda University. This protocol both visually communicates the brainstem-spinal cord preparation and clarifies the preparation of brainstem transverse slices in a comprehensive step-by-step manner. It was designed to increase reproducibility and enhance the likelihood of obtaining viable, long lasting, rhythmically-active slices for recording neural output from the respiratory regions of the brainstem.

Video: Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice

An Optogenetic Approach for Assessing Formation of Neuronal Connections in a Co-culture System

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are reprogrammed from human fibroblasts using episomal vectors. Colonies of iPSCs can be observed 30 days after initiation of fibroblast reprogramming. Pluripotent colonies are manually picked and grown in neural induction medium to permit differentiation into neural progenitor cells (NPCs). iPSCs rapidly convert into neuroepithelial cells within 1 week and retain the capability to self-renew when maintained at a high culture density. Primary mouse NPCs are differentiated into astrocytes by exposure to a serum-containing medium for 7 days and form a monolayer upon which embryonic day 18 (E18) rat cortical neurons (transfected with channelrhodopsin-2 (ChR2)) are added. Human NPCs tagged with the fluorescent protein, tandem dimer Tomato (tdTomato), are then seeded onto the astrocyte/cortical neuron culture the following day and allowed to differentiate for 28 to 35 days. We demonstrate that this system forms synaptic connections between iPSC-derived neurons and cortical neurons, evident from an increase in the frequency of synaptic currents upon photostimulation of the cortical neurons. This co-culture system provides a novel platform for evaluating the ability of iPSC-derived neurons to create synaptic connections with other neuronal populations.

A protocol to generate a co-culture system consisting of neurons derived from induced pluripotent stem cells (iPSCs), primary cortical neurons and astrocytes is described. This co-culture system allows detection of the formation of synaptic contacts and circuits between new, iPSC-derived neurons and pre-existing cortical neurons expressing channelrhodopsin-2.

Here we describe a protocol to generate a co-culture consisting of 2 different neuronal populations. Induced pluripotent stem cells (iPSCs) are ...

Developmental Biology

An Organotypic Slice Assay for High-Resolution Time-Lapse Imaging of Neuronal Migration in the Postnatal Brain

Neurogenesis in the postnatal brain depends on maintenance of three biological events: proliferation of progenitor cells, migration of neuroblasts, as well as differentiation and integration of new neurons into existing neural circuits. For postnatal neurogenesis in the olfactory bulbs, these events are segregated within three anatomically distinct domains: proliferation largely occurs in the subependymal zone (SEZ) of the lateral ventricles, migrating neuroblasts traverse through the rostral migratory stream (RMS), and new neurons differentiate and integrate within the olfactory bulbs (OB). The three domains serve as ideal platforms to study the cellular, molecular, and physiological mechanisms that regulate each of the biological events distinctly. This paper describes an organotypic slice assay optimized for postnatal brain tissue, in which the extracellular conditions closely mimic the in vivo environment for migrating neuroblasts. We show that our assay provides for uniform, oriented, and speedy movement of neuroblasts within the RMS. This assay will be highly suitable for the study of cell autonomous and non-autonomous regulation of neuronal migration by utilizing cross-transplantation approaches from mice on different genetic backgrounds.

This protocol describes an organotypic slice assay optimized for the postnatal brain and high-resolution time-lapse imaging of neuroblast migration in the rostral migratory stream.

Neurogenesis in the postnatal brain depends on maintenance of three biological events: proliferation of progenitor cells, migration of neuroblasts, as ...

Whole Cell Recording from an Organotypic Slice Preparation of Neocortex

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex. Because of the lack of specific pharmacological agents for these channels, we have taken a genetic approach to manipulating channel expression. We use an organotypic culture preparation (16) in order to maintain cell morphology and the laminar pattern of cortex. We typically isolate acute neocortical slices at postnatal days 8-10 and maintain the slices in culture for 3-7 days. This allows us to study neurons at a similar age to those in our work with acute slices and minimizes the development of exuberant excitatory connections in the slice. We record from visually-identified pyramidal neurons in layers II/III or V using infrared illumination (IR-) and differential interference contrast microscopy (DIC) with whole cell patch clamp in current- or voltage-clamp. We use biolistic (Gene gun) transfection of wild type or mutant potassium channel DNA to manipulate expression of the channels to study their function. The transfected cells are easily identified by epifluorescence microscopy after co-transfection with cDNA for green fluorescent protein (GFP). We compare recordings of transfected cells to adjacent, untransfected neurons in the same layer from the same slice.

This is a protocol to prepare and maintain a neocortical slice preparation in organotypic culture for the purpose of making electrical recordings from pyramidal neurons.

We have been studying the expression and functional roles of voltage-gated potassium channels in pyramidal neurons from rat neocortex.  Because of the ...

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

The mouse is an excellent model organism to study mammalian brain development due to the abundance of molecular and genetic data. However, the developing mouse brain is not suitable for easy manipulation and imaging in vivo since the mouse embryo is inaccessible and opaque. Organotypic slice cultures of embryonic brains are therefore widely used to study murine brain development in vitro. Ex-vivo manipulation or the use of transgenic mice allows the modification of gene expression so that subpopulations of neuronal or glial cells can be labeled with fluorescent proteins. The behavior of labeled cells can then be observed using time-lapse imaging. Time-lapse imaging has been particularly successful for studying cell behaviors that underlie the development of the cerebral cortex at late embryonic stages 1-2. Embryonic organotypic slice culture systems in brain regions outside of the forebrain are less well established. Therefore, the wealth of time-lapse imaging data describing neuronal cell migration is restricted to the forebrain 3,4. It is still not known, whether the principles discovered for the dorsal brain hold true for ventral brain areas. In the ventral brain, neurons are organized in neuronal clusters rather than layers and they often have to undergo complicated migratory trajectories to reach their final position. The ventral midbrain is not only a good model system for ventral brain development, but also contains neuronal populations such as dopaminergic neurons that are relevant in disease processes. While the function and degeneration of dopaminergic neurons has been investigated in great detail in the adult and ageing brain, little is known about the behavior of these neurons during their differentiation and migration phase 5. We describe here the generation of slice cultures from the embryonic day (E) 12.5 mouse ventral midbrain. These slice cultures are potentially suitable for monitoring dopaminergic neuron development over several days in vitro. We highlight the critical steps in generating brain slices at these early stages of embryonic development and discuss the conditions necessary for maintaining normal development of dopaminergic neurons in vitro. We also present results from time lapse imaging experiments. In these experiments, ventral midbrain precursors (including dopaminergic precursors) and their descendants were labeled in a mosaic manner using a Cre/loxP based inducible fate mapping system 6.

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

A method to generate organotypic slices from the E12.5 murine embryonic midbrain is described. The organotypic slice cultures can be used to observe the behavior of dopaminergic neurons or other ventral midbrain neurons.

The  mouse is an excellent model organism to study mammalian brain development due  to the abundance of molecular and genetic data. However, the ...

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to neuroblasts able to move along the rostral migratory stream (RMS) towards the olfactory bulb (OB). This long-distance migration is required for the subsequent maturation of newborn neurons in the OB, but the molecular mechanisms regulating this process are still unclear. Investigating the signaling pathways controlling neuroblast motility may not only help understand a fundamental step in neurogenesis, but also have therapeutic regenerative potential, given the ability of these neuroblasts to target brain sites affected by injury, stroke, or degeneration.
In this manuscript we describe a detailed protocol for in vivo postnatal electroporation and subsequent time-lapse imaging of neuroblast migration in the mouse RMS. Postnatal electroporation can efficiently transfect SVZ progenitor cells, which in turn generate neuroblasts migrating along the RMS. Using confocal spinning disk time-lapse microscopy on acute brain slice cultures, neuroblast migration can be monitored in an environment closely resembling the in vivo condition. Moreover, neuroblast motility can be tracked and quantitatively analyzed. As an example, we describe how to use in vivo postnatal electroporation of a GFP-expressing plasmid to label and visualize neuroblasts migrating along the RMS. Electroporation of shRNA or CRE recombinase-expressing plasmids in conditional knockout mice employing the LoxP system can also be used to target genes of interest. Pharmacological manipulation of acute brain slice cultures can be performed to investigate the role of different signaling molecules in neuroblast migration. By coupling in vivo electroporation with time-lapse imaging, we hope to understand the molecular mechanisms controlling neuroblast motility and contribute to the development of novel approaches to promote brain repair.

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

Neuroblast migration is a fundamental event in postnatal neurogenesis. We describe a protocol for efficient labeling of neuroblasts by in vivo postnatal electroporation and subsequent visualization of their migration using time-lapse imaging of acute brain slices. We include a description for the quantitative analysis of neuroblast dynamics by video tracking.

The subventricular zone (SVZ) is one of the main neurogenic niches in the postnatal brain. Here, neural progenitors proliferate and give rise to ...

Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to stimuli remain to be fully understood. To overcome the difficulty of accessing the brainstem in the whole animal, isolation of the brainstem and part of the spinal cord is performed. This preparation is maintained in artificial cerebro-spinal fluid where gases, concentrations, and temperature are controlled and monitored. The output signal from the respiratory network is recorded by a suction electrode placed on the fourth ventral root. In this manner, stimuli can be directly applied onto the brainstem, and the effect can be recorded directly. The signal recorded is linked to the inspiratory signal sent to the diaphragm via the phrenic nerve, and can be described as bursts (around 8 bursts per minute). Analysis of these bursts (frequency, amplitude, length, and area under the curve) allows precise characterization of the stimulus effect on the respiratory network. The main limitation of this method is the viability of the preparation beyond the early post-natal stages. Thus, this method greatly focuses on the study of the whole network without the peripheral inputs in the newborn rat.

The central respiratory drive is located in the brainstem. Spontaneous respiratory motor output from an isolated brainstem-spinal cord is recorded by placing an electrode on the fourth ventral root. This experimental approach is valuable for pharmacological investigations or the assessment of respiratory challenges and genetic manipulations on rhythmic motor behavior.

While it is well known that the central respiratory drive is located in the brainstem, several aspects of its basic function, development, and response to ...

An Alternative Approach to Study Primary Events in Neurodegeneration Using Ex Vivo Rat Brain Slices

Despite&#160;numerous studies that attempt to develop reliable animal models&#160;which reflecting the primary processes underlying neurodegeneration, very few have been widely accepted. Here, we propose&#160;a new procedure&#160;adapted from the well-known ex vivo brain slice technique, which offers a closer in vivo-like scenario than in vitro preparations, for investigating the early events triggering cell degeneration, as observed in Alzheimer&#39;s disease (AD). This variation consists of simple and easily reproducible steps, which enable preservation of the anatomical cytoarchitecture of the selected brain region and its local functionality in a physiological milieu. Different anatomical areas can be obtained from the same brain, providing the opportunity to perform multiple experiments with the treatments in question in a site-, dose-, and time-dependent manner. Potential limitations&#160;which could affect the outcomes related to this methodology are related to the conservation of the tissue, i.e., the maintenance of its anatomical integrity during the slicing and incubation steps and the section thickness, which can influence the biochemical and immunohistochemical analysis. This approach can be employed for different purposes, such as exploring molecular mechanisms involved in physiological or pathological conditions, drug screening, or dose-response assays. Finally, this protocol could also reduce the number of animals employed in behavioral studies. The application reported here has been recently described and tested for the first time on ex vivo rat brain slices containing the basal forebrain (BF), which is one of the cerebral regions primarily affected in AD. Specifically, it has been demonstrated that the administration of a toxic peptide derived from the C-terminus of acetylcholinesterase (AChE) could prompt an AD-like profile, triggering, along the antero-posterior axis of the BF, a differential expression of proteins altered in AD, such as the alpha7 nicotinic receptor (&#945;7-nAChR), phosphorylated Tau (p-Tau), and amyloid beta (A&#946;).

An Alternative Approach to Study Primary Events in Neurodegeneration Using Ex Vivo Rat Brain Slices

We present a method which can&#160;provide further insights into the early events underlying neurodegeneration and based on the established ex-vivo brain technique, combining the advantages of in vivo and in vitro experiments. Moreover, it represents a unique opportunity for direct comparison of treated and untreated group in the same anatomical plane.

Despite&#160;numerous studies that attempt to develop reliable animal models&#160;which reflecting the primary processes underlying neurodegeneration, ...

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 ...

In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively thin to properly visualize and access neurons for patch-clamping or imaging, and in vitro examination of brain circuitry is limited to only what is physically present in the acute slice. To maintain the benefits of in vitro slice experimentation while preserving a larger portion of presynaptic nuclei, we developed a novel slice preparation. This &#8220;wedge slice&#8221; was designed for patch-clamp electrophysiology recordings to characterize the diverse monaural, sound-driven inputs to medial olivocochlear (MOC) neurons in the brainstem. These neurons receive their primary afferent excitatory and inhibitory inputs from neurons activated by stimuli in the contralateral ear and corresponding cochlear nucleus (CN). An asymmetrical brain slice was designed which is thickest in the rostro-caudal domain at the lateral edge of one hemisphere and then thins towards the lateral edge of the opposite hemisphere. This slice contains, on the thick side, the auditory nerve root conveying information about auditory stimuli to the brain, the intrinsic CN circuitry, and both the disynaptic excitatory and trisynaptic inhibitory afferent pathways that converge on contralateral MOC neurons. Recording is performed from MOC neurons on the thin side of the slice, where they are visualized using DIC optics for typical patch-clamp experiments. Direct stimulation of the auditory nerve is performed as it enters the auditory brainstem, allowing for intrinsic CN circuit activity and synaptic plasticity to occur at synapses upstream of MOC neurons. With this technique, one can mimic in vivo circuit activation as closely as possible within the slice. This wedge slice preparation is applicable to other brain circuits where circuit analyses would benefit from preservation of upstream connectivity and long-range inputs, in combination with the technical advantages of in vitro slice physiology.

Integration of diverse synaptic inputs to neurons is best measured in a preparation that preserves all pre-synaptic nuclei for natural timing and circuit plasticity, but brain slices typically sever many connections. We developed a modified brain slice to mimic in vivo circuit activity while maintaining in vitro experimentation capability.

In vitro slice electrophysiology techniques measure single-cell activity with precise electrical and temporal resolution. Brain slices must be relatively ...

Electrophysiological Recordings of Brainstem-Spinal Cord Preparations from a Newborn Rodent

Source: Rousseau, J., et al. <a href="https://app-jove-com.bdigitaluss.remotexs.co/v/53071">Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording.</a> J. Vis. Exp. (2015).
This video demonstrates electrophysiological recordings of isolated brainstem-spinal cord preparations from newborn rodents. The preparation is placed in a recording chamber, and the inspiratory bursts from the brainstem are recorded under oxygenated and oxygen-deprived conditions using the fourth ventral nerve root in the spinal cord.

Preparation of Rhythmically-active In Vitro Neonatal Rodent Brainstem-spinal Cord and Thin Slice

Summary

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Chapters in this video

An Optogenetic Approach for Assessing Formation of Neuronal Connections in a Co-culture System

An Organotypic Slice Assay for High-Resolution Time-Lapse Imaging of Neuronal Migration in the Postnatal Brain

Whole Cell Recording from an Organotypic Slice Preparation of Neocortex

Organotypic Slice Cultures of Embryonic Ventral Midbrain: A System to Study Dopaminergic Neuronal Development in vitro

In vivo Postnatal Electroporation and Time-lapse Imaging of Neuroblast Migration in Mouse Acute Brain Slices

Electrophysiology on Isolated Brainstem-spinal Cord Preparations from Newborn Rodents Allows Neural Respiratory Network Output Recording

An Alternative Approach to Study Primary Events in Neurodegeneration Using Ex Vivo Rat Brain Slices

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

In Vitro Wedge Slice Preparation for Mimicking In Vivo Neuronal Circuit Connectivity

Electrophysiological Recordings of Brainstem-Spinal Cord Preparations from a Newborn Rodent