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

Summary

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.

Abstract

Mammalian inspiratory rhythm is generated from a neuronal network in a region of the medulla called the preBötzinger 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.

Introduction

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 respiratory^1,2. The central respiratory network includes a group of rhythmically active neurons located in the rostral ventrolateral medulla^1,3. Mammalian respiratory rhythmogenesis is generated from an autorhythmic interneuron network dubbed the preBötzinger complex (pBC), which has been localized experimentally via both slice and en bloc preparations of neonatal mammalian brainstem-spinal cords^3,4,5,6,7,8. 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 pBC^3,10,11,12. 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.

Protocol

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

1. Prepare artificial cerebral spinal fluid (aCSF).
   1. 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 • H₂O (0.5 mM), 0.247 g MgSO₄ • 7H₂O (1.0 mM), and 5.410 g D-glucose (30 mM), 0.221 g CaCl₂ • 2H₂O (1.5 mM). Always add the CaCl₂ • 2H₂O last.
   2. Dissolve the components into 1 L of deionized water. Stir for 20 - 30 min.
   3. Measure the pH of the aCSF and adjust to 7.40 ± 0.02 using small volumes (typically <0.5 mL) of dilute NaOH, KOH, or HCl.
      NOTE: Prepared aCSF can be stored in the refrigerator (4 °C, pH 7.40 ± 0.02) up to three days before bacterial growth affects viability of the tissue during an experiment. Use 0.2 µm 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.

2. Preparation of Dissection and Vibratome Rig

1. Set up blade.
   1. 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.
   2. 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.
2. Set up paraffin wax slab.
   1. 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.
   2. Add approximately 0.5 g of graphite powder and mix solution thoroughly and uniformly into the liquid paraffin.
   3. 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'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.
   4. 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.
   5. 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.

3. Dissection and Isolation of the Neuraxis

1. 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).
   1. 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 cryoanesthesia^13,14, isoflurane vaporization¹⁵, or injection¹⁶ balanced with oxygen.
   2. 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.
   3. 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.
   4. Bubble an aCSF reservoir (500 mL side-port bottle) continuously with chilled (4 °C, pH 7.40 ± 0.02) aCSF and oxygenate with a mixture of 95% O₂ and 5% CO₂ (also called "carbogen"). 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.
   5. 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.
   6. Remove excess fluids as needed by vacuum suction filtration system.
   7. 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.
   8. 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.
2. 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.
   1. 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.
   2. 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 "flaps" 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).
   3. 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).
3. Perform dorsal laminectomy.
   1. 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.
   2. 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).
4. Perform ventral laminectomy.
   1. 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).
   2. 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).
   3. 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 Figure 3C).
   4. 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).
   5. 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.
   6. 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' origin at the spinal cord as possible; avoid stretching the roots.
   7. 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.
   8. 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.
   9. 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).
   10. 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).
5. Remove dura mater.
   NOTE: The isolated brainstem-spinal cord can be used for en bloc in vitro recording as has been previously described^3,7. 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.
   1. 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.
   2. 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.
   3. 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.
   4. 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.
   5. Use a fine insect pin to tease away any remaining dura and remove remaining blood vessels in close proximity to cranial and cervical rootlets.

4. Slice Protocol

1. After removing the dura, place the brainstem in the center of the paraffin platform on the plastic block (hereafter referred to as the "cutting block"). 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.
2. 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.
3. Make an initial slice to remove the uneven, extraneous tissue on the rostral-most end. This initial cut can be 200 - 300 µm 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.
4. Continue cutting rostral-to-caudal until the IX rootlets are near the surface of the cut face of the brainstem.
5. 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 "wedge" shape to the cut slice and cut slices of approximately 100 - 200 µm 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).
6. Cut a 300 - 500 µm slice of brainstem from these rostral neuroanatomical markers to capture the pBC and associated transmission circuitry.
   NOTE: An "ideal" 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.

5. Recording Procedures

1. 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 clamping^10,11.

Results

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 preBötzinger Complex, premotor neurons projecting to the hypoglossal motor ...

Discussion

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 others^17,18,19, will allow reproducible preparation of robust and viable tissue that is widely adap...

Materials

Name	Company	Catalog Number	Comments
NaCl	Fisher Scientific	S271-500
KCl	Sigma Aldrich	P5405-1KG
NaHCO3	Fisher Scientific	BP328-1
NaH2PO4 •H2O	Sigma Aldrch	S9638-25G
CaCl2•2H2O	Sigma Aldrich	C7902-500G
MgSO4•7H2O	Sigma Aldrich	M7774-500G
D-Glucose	Sigma Aldrich	G8270-1KG
Cold-Light source Halogen lamp 150 W	AmScope	H2L50-AY
Dissection Microscope	Leica	M-60
Vibratome 1000 Plus	Vibratome	W3 69-0353
Magnetic Base	Kanetic	MB-B-DG6C
Isoflurane, USP	Patterson Veterinary	NDC 14043-704-06
Sword Classic Double Edge Blades	Wilkinson	97573
Histoclear	Sigma-Aldrich	H2779
Dumont #5 Fine Forceps	Fine Science Tools	11254-20
Dumont #5/45 Forcep	Fine Science Tools	11251-35
Scalpel Blades #10	Fine Science Tools	10010-00
Scalpel Handel #3	Fine Science Tools	10003-12
Spring Scissors Straight	Fine Science Tools	15024-10
Narrow Pattern Forcep Serrated/straight	Fine Science Tools	11002-12
Castroviejo Micro Dissecting Spring Scissors; Straight	Roboz	RS-5650
Vannas Scissors 3" Curved	Roboz	RS-5621
Insect pins, 0	Fine Science Tools/8840604	26000-35
Insect pins, 0, SS	Fine Science Tools	26001-35
Insect pins, 00	Fine Science Tools	26000-30
Insect pins, 00, SS	Fine Science Tools	26001-30
Insect pins, 000	Fine Science Tools	26000-25
Insect pins, 000, SS	Fine Science Tools	26001-25
Minutien pins, 0.10 mm	Fine Science Tools	26002-10
Minutien pins, 0.15 mm	Fine Science Tools	26002-15
Minutien pins, 0.2 mm	Fine Science Tools	26002-20
Fisher Tissue prep Parafin	fisher	T56-5
Graphite	fisher	G67-500
Delrin Plastic	Grainger	3HMT2
18 Gauge Hypodermic Needle	BD	305195