This work was supported by R01 NS079623, P01 HL149630, and P01 HL095491.

The author declares that the protocol follows the guidelines of the Institutional Animal Care and Use Committee at Beth Israel Deaconess Medical Center.
1. Preparation of surgical instruments and stereotaxic frame
NOTE: The surgery is performed under aseptic conditions. Sterility is maintained using the sterile tip technique.
<ol>
	<li>Install the stereotaxic arm with a micropipette or syringe filled with an injectable of choice (adeno-associated virus (AAV) or conventional tracer) on the stereotaxic frame and prepare the mouse adapter (Figure 2A).</li>
	<li>Prepare autoclaved surgical instruments (Table of materials) and place them on a sterile surface.</li>
</ol>
2. Anesthesia induction and mouse preparation
<ol>
	<li>Turn on O2 at 0.5 L/min and set the isoflurane vaporizer at 4.0, making sure that O2 flow is to the induction box. 
	CAUTION: Make sure that the isoflurane induction box is placed in a hood and that the isoflurane is scavenged away from the surgical site.</li>
	<li>Place the mouse (10-week-old male C57BL/6J) in the induction chamber.</li>
	<li>Once breathing has slowed, open the induction chamber, and lift the mouse slightly. Use clippers to remove hair from head to shoulders.</li>
</ol>
3. Positioning of the mouse in the stereotaxic frame
<ol>
	<li>Move the mouse to the stereotaxic frame and place the nose in a flexible nose cone. At this stage, make sure the O2 flow is now directed to the nose cone.</li>
	<li>Place the mouse in the stereotaxic frame using ear bars only. 
	NOTE: Make sure the ear bars are even and the head is level.</li>
	<li>Anteroflex the head of the mouse to a 90&#176; angle by manually guiding the nose. To secure this position, place a plastic barrier between the ear bar pillars of the mouse adapter, parallel to the pillars. The flat part of the skull serves as the reference, similar to the flat skull approach in conventional stereotaxic surgery. 
	NOTE: Do not over flex the head (i.e. beyond an angle of 90&#176; between the plane of the frontal skull bone and the plane of the surface of the table) as this impedes airflow through the upper airway. If airflow is impeded, reposition the mouse, making sure the body is supported underneath the trunk and a plastic card is set at 90&#176; between the plane of the frontal skull bone and the plane of the surface of the table as outlined in Figure 2A,C.</li>
	<li>Place the heating pad underneath the mouse, and then make sure that the neck and the rest of the body are positioned at the same level (i.e., at approximately 180&#176; or parallel to the table). The toolbox that holds the spring scissors can be used to lift the body to this position. 
	NOTE: This step is important as the caudal brainstem and upper cervical cord move depending on position, in contrast to more rostral parts of the CNS which are held in place by the skull.</li>
	<li>Inject a single dose of 4 mg/kg Meloxicam slow-release (SR) subcutaneously (s.c.) at a volume of 2 &#181;L/g body weight and place lubricant on the eyes.</li>
	<li>Clean the surgical incision site first with a 70% alcohol prep pad, then with a betadine prep pad, and then again with an alcohol prep pad and let dry.</li>
	<li>Place a drape underneath the body.</li>
	<li>Disinfect hands and put on sterile gloves.</li>
	<li>Place a drape at the surgical site.</li>
</ol>
4. Surgery to access the cisterna magna
<ol>
	<li>Make sure the mouse is appropriately anesthetized by pinching the toes or checking the corneal reflex.</li>
	<li>Reduce isoflurane to maintenance levels (2.0).</li>
	<li>Make a 1-1.2 cm incision with surgical blade #10 from the edge of the occipital bone toward the shoulders in one smooth movement.</li>
	<li>Make an incision in the midline raphe of the trapezius muscle. This exposes the paired longus capitis muscles. 
	NOTE: In mice, the trapezius muscle is a very thin, almost transparent muscle. Make sure to stay in the midline and do not cut into the underlying muscles as this will cause unnecessary bleeding.</li>
	<li>Place both retractor hooks in between the paired longus capitis muscles, one oriented to the left and the other to the right. The weight of the hemostats provides tension to the retractor hooks which can be modified by re-adjusting the position of the hemostats.</li>
	<li>Position the surgical microscope in place to better visualize the surgical field.</li>
	<li>Use the blunt laminectomy forceps to separate the left and right bellies of the paired longus capitis muscle, starting from the occiput, where the midline is readily visible. Guide the blunt forceps across the bone of the occiput in the midline down to where it meets the cisternal dura mater, and then continue across the dura mater to the atlas. 
	NOTE: There is no need to cut through the paired longus capitis muscles as nothing holds them together in the midline; doing so will cause unnecessary bleeding.</li>
	<li>Reposition the retractors and adjust the tension by repositioning the hemostats, opening the view of the cisterna magna.</li>
	<li>Use the blunt laminectomy forceps to separate the muscles further in the midline to get a good viewing window of the brainstem and cerebellum.</li>
	<li>Repeat steps 4.7-4.9 as needed until the cerebellum and brainstem come in view below the dura.</li>
	<li>Using blunt laminectomy forceps, clear the dura of the small strands of connective tissue by moving the forceps from the midline in a lateral direction, until there is a clear view of the brainstem and to create more lateral space, as needed for the target.</li>
</ol>
5. Opening of the cisternal membrane
<ol>
	<li>Use the angled Dumont forceps (#4/45) to grab the dura, which extends from the occipital bone to the atlas. Grab the dura near the occipital bone and use the spring scissors to make a small opening (~0.5 to 1.5 mm) in the dura. 
	NOTE: At this rostral location, the space between the brainstem and the overlying dura is widest, providing ample room for safe manipulation of the dura.</li>
	<li>Use the spring scissors to lift the dura and open the dura further. The size of the window depends on the target. 
	NOTE: A larger window will be needed when making multiple longitudinal injections or bilateral injections; a small window will be sufficient when making single unilateral or midline injections.</li>
	<li>Once the dura is opened, drain excess cerebrospinal fluid with a sterile cue tip.</li>
</ol>
6. Identification of landmarks and zero point
<ol>
	<li>View the dorsal surface of the brainstem with detailed landmarks through the open dura. The obex, the point where the central canal opens into the IV ventricle, is the standard anterior-posterior and mediolateral zero point.</li>
</ol>
7. Target coordinates
NOTE: For various targets, we have included a list of standard coordinates with anterior posterior (AP) and mediolateral (ML) coordinates relative to zero-point bregma and cisterna magna coordinates with AP and ML coordinates relative to zero point obex to facilitate the transition between methodologies (Table 1). Dorsoventral (DV) coordinates are relative to the surface of the brain or cerebellum (standard approach) or the surface of the brainstem or upper cervical cord (cisterna magna approach) at the point of AP and ML entry. Planning should be done prior to surgery.
<ol>
	<li>Use the three sets of coordinates to determine the target: AP, ML, and DV. Due to the head position, the relative orientation of brainstem structures varies by location.
	<ol>
		<li>For target distance &#62;0.4 mm from caudal to the obex (Figure 1B, green) perform the following.
		<ol>
			<li>AP: Use any standard stereotaxic reference atlas (e.g., Paxinos and Franklin atlas2) or tissue series cut in the transverse plane to estimate the AP distance between obex and the target.</li>
			<li>ML: Use any standard stereotaxic reference atlas or tissue series cut in the transverse plane to estimate the ML distance between obex and the target.</li>
			<li>DV: Estimate coordinates relative to the surface of the brain or cerebellum at the AP and ML target point. Use any standard stereotaxic reference atlas or tissue series cut in the transverse plane to estimate the distance between the brainstem surface at the desired AP and ML coordinates and the target.</li>
		</ol>
		</li>
		<li>For target distance &#60;0.4 mm from caudal to the obex (Figure 1B, orange) perform the following.
		<ol>
			<li>AP: Adjust coordinates to account for anteroflexion of the brainstem. For ventral and rostral coordinates, the AP brainstem entry point will be more caudal relative to the target AP coordinate in the standard plane.</li>
			<li>ML: Derive target coordinates from a standard stereotaxic reference atlas or tissue series cut in the transverse plane. Coordinates will be relative to the visualized midline at the target AP level.</li>
			<li>DV: Estimate coordinates relative to the surface of the brainstem at the AP and ML target point. Adjust DV to account for anteroflexion of the brainstem. For ventral and rostral coordinates, the DV coordinates will be larger than the distance from the dorsal surface of the brainstem in the standard plane.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
8. Injection of the target
<ol>
	<li>Lower the pipette or syringe to the target using the stereotaxic arm and inject solution as for standard stereotaxic approaches. Leave in place for 1-5 min after injection, to avoid a needle track when using volumes between 3-50 nL. Then, lift the pipette or syringe using the stereotaxic arm.</li>
	<li>Repeat step 8.1. for multiple targets.</li>
</ol>
9. Closure of the surgical field
<ol>
	<li>Remove the hooks carefully from the surgical field. The paired longus capitis muscles will fall back into a neutral position, fully covering the cisterna magna. Do not close the trapezius muscle and dura mater in the midline as they are too fragile to hold sutures.</li>
	<li>Close the skin with three nylon or polypropylene sutures (5-0 or 6-0).</li>
</ol>
10. Post-operative care
<ol>
	<li>Turn off isoflurane and remove the mouse from the stereotaxic frame. Place the mouse in a clean cage on a heating pad and observe until awake and moving.</li>
	<li>Monitor health status, weight, and sutures on post-operative days 1-3. Remove sutures on day 10 if not already removed.</li>
</ol>

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The authors have nothing to disclose.

Standard stereotaxic surgery commonly relies on skull landmarks to calculate the coordinates of target sites in the CNS1. Target sites are then accessed via burr holes that are drilled through the skull1. This method is not ideal for the caudal brainstem as target sites are located distant from the skull landmarks in the anteroposterior and dorsoventral planes2 and as the anatomy of the skull and overlying muscles make access challenging<sup...

Standard stereotaxic surgery to target brain sites in mice1 commonly involves fixation of the skull using a set of ear bars and a mouth bar. Coordinates are then estimated based on reference atlases2,3, and skull landmarks, namely, bregma (the point where the sutures of the frontal and parietal bones come together) or lambda (the point where the sutures of the parietal and occipital bones come together; Figure 1A,B). Through a burr hole into the skull above the estimated target, the target region can then be reached, either for delivery of microinjections or instrumentation with cannulas or optic fibers. Due to variation in the anatomy of these sutures and errors in the localization of bregma or lambda4,5, the position of zero points in relation to the brain varies from animal to animal. While small errors in targeting, that result from this variability, are not a problem for large or nearby targets, their impact is greater for smaller areas of interest that are remote from the zero points in the anteroposterior or dorsoventral planes and/or when studying animals of varying size due to age, strain and/or sex. There are several additional challenges that are unique for the medulla oblongata and the upper cervical cord. First, small changes in anteroposterior coordinates are associated with significant changes in dorsoventral coordinates relative to the dura, due to the position and shape of the cerebellum (Figure 1Bi)2,6,7. Second, the upper cervical cord is not contained within the skull2. Third, the slanting position of the occipital bone and overlying layer of neck muscles2 makes the standard stereotaxic approach even more challenging for structures located near the transition between the brainstem and spinal cord (Figure 1Bi). Finally, many targets of interest in the caudal brainstem and cervical cord are small2, requiring precise and reproducible injections8,9.
An alternative approach through the cisterna magna circumvents these problems. The cisterna magna is a large space that extends from the occipital bone to the atlas (Figure 1A, i.e., the second vertebral bone)10. It is filled with cerebrospinal fluid and covered by dura mater10. This space between the occipital bone and the atlas opens when anteroflexing the head. It can be accessed by navigating in between the overlying paired bellies of the longus capitis muscle, exposing the dorsal surface of the caudal brainstem. Regions of interest can then be targeted based upon the landmarks of these regions themselves if they are located near the dorsal surface; or by using the obex, the point where the central canal opens into the IV ventricle, as a zero point for coordinates to reach deeper structures. This approach has been successfully used in a variety of species, including the rat11, cat12, mouse8,9, and non-human primate13 to target the ventral respiratory group, medullary medial reticular formation, the nucleus of the solitary tract, area postrema, or hypoglossal nucleus. However, this approach is not widely utilized as it requires knowledge of anatomy, a specialized toolkit, and more advanced surgical skills compared to the standard stereotaxic approach.
Here we describe a step-by-step surgical approach to reach the brainstem and upper cervical cord via the cisterna magna, visualize landmarks, set the zero point (Figure 2), and estimate and optimize target coordinates for stereotaxic delivery of microinjections into the discrete brainstem and spinal cord regions of interest (Figure 3). We then discuss the advantages and disadvantages related to this approach.

<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Alcohol pad</td>
			<td>Med-Vet International</td>
			<td>SKU: MDS090735Z</td>
			<td>skin preparation for the prevention of surgical site infection</td>
		</tr>
		<tr>
			<td>Angled forceps, Dumont #5/45</td>
			<td>FST</td>
			<td>11251-35</td>
			<td>only to grab dura</td>
		</tr>
		<tr>
			<td>Betadine pad</td>
			<td>Med-Vet International</td>
			<td>SKU:PVP-PAD</td>
			<td>skin preparation for the prevention of surgical site infection</td>
		</tr>
		<tr>
			<td>Cholera toxin subunit-b, Alexa Fluor 488/594 conjugate</td>
			<td>Thermo Fisher Scientific</td>
			<td>488: C34775, 594: C22842</td>
			<td>Fluorescent tracer</td>
		</tr>
		<tr>
			<td>Clippers</td>
			<td>Wahl</td>
			<td>Model MC3, 28915-10</td>
			<td>for shaving fur at surgical site</td>
		</tr>
		<tr>
			<td>Electrode holder with corner clamp</td>
			<td>Kopf</td>
			<td>1770</td>
			<td>to hold glass pipette</td>
		</tr>
		<tr>
			<td>Flowmeter</td>
			<td>Gilmont instruments</td>
			<td>model # 65 MM</td>
			<td>to regulate flow of isoflurane and oxygen to mouse on the surgical plane</td>
		</tr>
		<tr>
			<td>Fluorescent microspheres, polystyrene</td>
			<td>Thermo Fisher Scientific</td>
			<td>F13080</td>
			<td>Fluorescent tracer</td>
		</tr>
		<tr>
			<td>Heating pad</td>
			<td>Stoelting</td>
			<td>53800M</td>
			<td>thermoregulation</td>
		</tr>
		<tr>
			<td>Induction chamber with port hook up kit</td>
			<td>Midmark Inc</td>
			<td>93805107 92800131</td>
			<td>chamber providing initial anasthesia</td>
		</tr>
		<tr>
			<td>Insulin Syringe</td>
			<td>Exelint International</td>
			<td>26028</td>
			<td>to administer saline and analgesic</td>
		</tr>
		<tr>
			<td>Isoflurane</td>
			<td>Med-Vet International</td>
			<td>SKU:RXISO-250</td>
			<td>inhalant anesthetic</td>
		</tr>
		<tr>
			<td>Isoflurane Matrix VIP 3000 vaporizer</td>
			<td>Midmark Inc</td>
			<td>91305430</td>
			<td>apparatus for inhalant anesthetic delivery</td>
		</tr>
		<tr>
			<td>Laminectomy forceps, Dumont #2</td>
			<td>FST</td>
			<td>11223-20</td>
			<td>only to clean dura</td>
		</tr>
		<tr>
			<td>Medical air, compressed</td>
			<td>Linde</td>
			<td>UN 1002</td>
			<td>used with stimulator &#38; PicoPump for providing air for precision solution injection</td>
		</tr>
		<tr>
			<td>Meloxicam SR</td>
			<td>Zoo Pharm LLC</td>
			<td>Lot # MSR2-211201</td>
			<td>analgesic</td>
		</tr>
		<tr>
			<td>Microhematocrit borosilicate glass pre calibrated capillary tube</td>
			<td>Globe Scientific Inc</td>
			<td>51628</td>
			<td>for transfection of material to designated co-ordinates</td>
		</tr>
		<tr>
			<td>Mouse adaptor</td>
			<td>Stoelting</td>
			<td>0051625</td>
			<td>&#160;adapting rat stereotaxic frame for mouse surgery</td>
		</tr>
		<tr>
			<td>Needle holder, Student Halsted- Mosquito Hemostats</td>
			<td>FST</td>
			<td>91308-12</td>
			<td>for suturing</td>
		</tr>
		<tr>
			<td>Oxygen regulator</td>
			<td>Life Support Products</td>
			<td>S/N 909328, lot 092109</td>
			<td>regulate oxygen levels from oxygen tank</td>
		</tr>
		<tr>
			<td>Oxygen tank, compressed</td>
			<td>Linde</td>
			<td>USP UN 1072</td>
			<td>provided along with isoflurane anasthesia</td>
		</tr>
		<tr>
			<td>Plastic card</td>
			<td>not applicable</td>
			<td>not applicable</td>
			<td>any firm plastic card, cut to fit the stereotactic frame (e.g. ID card)</td>
		</tr>
		<tr>
			<td>Pneumatic PicoPump ( or similar)</td>
			<td>World Precision Instruments (WPI)</td>
			<td>SYS-PV820</td>
			<td>For precision solution injection</td>
		</tr>
		<tr>
			<td>Saline, sterile</td>
			<td>Mountainside Medical Equipment</td>
			<td>H04888-10</td>
			<td>to replace body fluids lost during surgery</td>
		</tr>
		<tr>
			<td>Scalpel handle, #3</td>
			<td>FST</td>
			<td>10003-12</td>
			<td>to hold scalpel</td>
		</tr>
		<tr>
			<td>Scissors, Wagner</td>
			<td>FST</td>
			<td>14070-12</td>
			<td>to cut polypropylene suture</td>
		</tr>
		<tr>
			<td>Spring scissors, Vannas 2.5mm with accompanying box</td>
			<td>FST</td>
			<td>15002-08</td>
			<td>scissors only to open dura, box to elevate body</td>
		</tr>
		<tr>
			<td>Stereotactic micromanipulator</td>
			<td>Kopf</td>
			<td>1760-61</td>
			<td>attached to electrode holder to adjust position based on co-ordinates</td>
		</tr>
		<tr>
			<td>Stereotactic &#39;U&#39; frame assembly and intracellular base plate</td>
			<td>Kopf</td>
			<td>1730-B, 1711</td>
			<td>frame for surgery</td>
		</tr>
		<tr>
			<td>Sterile cotton tipped applicators</td>
			<td>Puritan</td>
			<td>25-806 10WC</td>
			<td>absorbing blood from surgical field</td>
		</tr>
		<tr>
			<td>Sterile non-fenestrated drapes</td>
			<td>Henry Schein</td>
			<td>9004686</td>
			<td>for sterile surgical field</td>
		</tr>
		<tr>
			<td>Sterile opthalmic ointment</td>
			<td>Puralube</td>
			<td>P1490</td>
			<td>ocular lubricant</td>
		</tr>
		<tr>
			<td>Stimulator &#38; Tubing</td>
			<td>Grass Medical Instruments</td>
			<td>S44</td>
			<td>to provide controlled presurred air for precision solution injection</td>
		</tr>
		<tr>
			<td>Surgical Blade #10</td>
			<td>Med-Vet International</td>
			<td>SKU: 10SS</td>
			<td>for skin incision</td>
		</tr>
		<tr>
			<td>Surgical forceps, Extra fine Graefe</td>
			<td>FST</td>
			<td>11153-10</td>
			<td>to hold skin</td>
		</tr>
		<tr>
			<td>Surgical gloves</td>
			<td>Med-Vet International</td>
			<td>MSG2280Z</td>
			<td>for asceptic surgery</td>
		</tr>
		<tr>
			<td>Surgical microscope</td>
			<td>Leica</td>
			<td>Model M320/ F12</td>
			<td>for 5X-40X magnification of surgical site</td>
		</tr>
		<tr>
			<td>Suture 5-0 polypropylene</td>
			<td>Oasis</td>
			<td>MV-8661</td>
			<td>to close the skin</td>
		</tr>
		<tr>
			<td>Tegaderm</td>
			<td>3M</td>
			<td>3M ID 70200749250</td>
			<td>provides sterile barrier</td>
		</tr>
		<tr>
			<td>Universal Clamp and stand post</td>
			<td>Kopf</td>
			<td>1725</td>
			<td>attached to stereotactic U frame and intracellular base plate</td>
		</tr>
		<tr>
			<td>Wound hook with hartman hemostats</td>
			<td>FST</td>
			<td>18200-09, 13003-10</td>
			<td>to separate muscles and provide surgical window</td>
		</tr>
	</tbody>
</table>

stereotaxic surgical approach to microinject the caudal brainstem and upper cervical spinal cord via the cisterna magna in mice

Stereotaxic surgery to target brain sites in mice is commonly guided by skull landmarks. Access is then obtained via burr holes drilled through the skull. This standard approach can be challenging for targets in the caudal brainstem and upper cervical cord due to specific anatomical challenges as these sites are remote from skull landmarks, leading to imprecision. Here we outline an alternative stereotaxic approach via the cisterna magna that has been used to target discrete regions of interest in the caudal brainstem and upper cervical cord. The cisterna magna extends from the occipital bone to the atlas (i.e., the second vertebral bone), is filled with cerebrospinal fluid, and is covered by dura mater. This approach provides a reproducible route of access to select central nervous system (CNS) structures that are otherwise hard to reach due to anatomical barriers. Furthermore, it allows for direct visualization of brainstem landmarks in close proximity to the target sites, increasing accuracy when delivering small injection volumes to restricted regions of interest in the caudal brainstem and upper cervical cord. Finally, this approach provides an opportunity to avoid the cerebellum, which can be important for motor and sensorimotor studies.

The cisterna magna approach makes it possible to target caudal brainstem and upper cervical cord structures that are otherwise hard to reach via standard stereotaxic approaches or are prone to inconsistent targeting. The surgery to reach the cisterna magna requires incisions of the skin, a thin layer of trapezius muscle, and opening of the dura mater and is therefore well tolerated by mice. It is especially efficient and less invasive when targeting multiple (longitudinally dispersed or bilateral) sites, as it d...

Watch this Scientific Journal Video about Stereotaxic Surgical Approach to Microinject the Caudal Brainstem and Upper Cervical Spinal Cord via the Cisterna Magna in Mice at JoVE.com

Stereotaxic Surgical Approach to Microinject the Caudal Brainstem and Upper Cervical Spinal Cord via the Cisterna Magna in Mice

Stereotaxic surgery to target brain sites in mice commonly involves access through the skull bones and is guided by skull landmarks. Here we outline an alternative stereotaxic approach to target the caudal brainstem and upper cervical spinal cord via the cisterna magna that relies on direct visualization of brainstem landmarks.

Stereotaxic Surgical Approach to Microinject the Caudal Brainstem and Upper Cervical Spinal Cord via the Cisterna Magna in Mice

Stereotaxic surgery to target brain sites in mice is commonly guided by skull landmarks. Access is then obtained via burr holes drilled through the skull. ...

stereotaxic-surgical-approach-to-microinject-caudal-brainstem-upper

Universidad de Cartagena UDEC

Research

JoVE Journal

Neuroscience

9.3K Views.  Harvard Medical School. Stereotaxic surgery to target brain sites in mice commonly involves access through the skull bones and is guided by skull landmarks. Here we outline an alternative stereotaxic approach to target the caudal brainstem and upper cervical spinal cord via the cisterna magna that relies on direct visualization of brainstem landmarks.

Video: Stereotaxic Surgical Approach to Microinject the Caudal Brainstem and Upper Cervical Spinal Cord via the Cisterna Magna in Mice

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

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

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

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

Stereotaxic Injection of a Viral Vector for Conditional Gene Manipulation in the Mouse Spinal Cord

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central nervous system. We demonstrate a stereotaxic injection technique that allows targeted gene expression or silencing in the dorsal horn of the mouse spinal cord. The surgical procedure is brief. It requires laminectomy of a single vertebra, providing for quick recovery of the animal and unimpaired motility of the spine. Controlled injection of a small vector suspension volume at low speed and use of a microsyringe with beveled glass cannula minimize the tissue lesion. The local immune response to the vector depends on the intrinsic properties of the virus employed; in our experience, it is minor and short-lived when a recombinant adeno-associated virus is used. A reporter gene such as enhanced green fluorescent protein facilitates monitoring spatial distribution of the vector, and the efficacy and cellular specificity of the transfection.

Viral vectors allow for targeted gene manipulation. We demonstrate a method for conditional gene expression or ablation in the mouse spinal cord, using stereotaxic injection of a viral vector into the dorsal horn, a prominent site of synaptic contact between primary somatosensory afferents and neurons of the central nervous system.

Intraparenchymal injection of a viral vector enables conditional gene manipulation in distinct populations of neurons or particular regions of the central ...

Spinal Cord Transection in the Larval Zebrafish

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

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

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

Diffusion Imaging in the Rat Cervical Spinal Cord

Magnetic resonance imaging (MRI) is the state of the art approach for assessing the status of the spinal cord noninvasively, and can be used as a diagnostic and prognostic tool in cases of disease or injury. Diffusion weighted imaging (DWI), is sensitive to the thermal motion of water molecules and allows for inferences of tissue microstructure. This report describes a protocol to acquire and analyze DWI of the rat cervical spinal cord on a small-bore animal system. It demonstrates an imaging setup for the live anesthetized animal and recommends a DWI acquisition protocol for high-quality imaging, which includes stabilization of the cord and control of respiratory motion. Measurements with diffusion weighting along different directions and magnitudes (b-values) are used. Finally, several mathematical models of the resulting signal are used to derive maps of the diffusion processes within the spinal cord tissue that provide insight into the normal cord and can be used to monitor injury or disease processes noninvasively.

The goal of this protocol is to obtain high-quality diffusion weighted magnetic resonance imaging (DWI) of the rat spinal cord for noninvasive characterization of tissue microstructure. This protocol describes optimizations of the MRI sequence, radiofrequency coil, and analysis methods to enable DWI images free from artifacts.

Magnetic resonance imaging (MRI) is the state of the art approach for assessing the status of the spinal cord noninvasively, and can be used as a ...

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

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

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

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

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

Cannula Implantation into the Cisterna Magna of Rodents

Cisterna magna cannulation (CMc) is a straightforward procedure that enables direct access to the cerebrospinal fluid (CSF) without operative damage to the skull or the brain parenchyma. In anesthetized rodents, the exposure of the dura mater by blunt dissection of the neck muscles allows the insertion of a cannula into the cisterna magna (CM). The cannula, composed either by a fine beveled needle or borosilicate capillary, is attached via a polyethylene (PE) tube to a syringe. Using a syringe pump, molecules can then be injected at controlled rates directly into the CM, which is continuous with the subarachnoid space. From the subarachnoid space, we can trace CSF fluxes by convective flow into the perivascular space around penetrating arterioles, where solute exchange with the interstitial fluid (ISF) occurs. CMc can be performed for acute injections immediately following the surgery, or for chronic implantation, with later injection in anesthetized or awake, freely moving rodents. Quantitation of tracer distribution in the brain parenchyma can be performed by epifluorescence, 2-photon microscopy, and magnetic resonance imaging (MRI), depending on the physico-chemical properties of the injected molecules. Thus, CMc in conjunction with various imaging techniques offers a powerful tool for assessment of the glymphatic system and CSF dynamics and function. Furthermore, CMc can be utilized as a conduit for fast, brain-wide delivery of signaling molecules and metabolic substrates that could not otherwise cross the blood brain barrier (BBB).

Here we describe a protocol to perform cisterna magna cannulation (CMc), a minimally invasive way to deliver tracers, substrates and signaling molecules into the cerebrospinal fluid (CSF). Combined with different imaging modalities, CMc enables glymphatic system and CSF dynamics assessment, as well as brain-wide delivery of various compounds.

Cisterna magna cannulation (CMc) is a straightforward procedure that enables direct access to the cerebrospinal fluid (CSF) without operative damage to ...

Laminectomy and Spinal Cord Window Implantation in the Mouse

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated digital vaporizer is utilized to achieve a stable plane of anesthesia at a low-flow rate of isoflurane. A single vertebral spine is removed, and a commercially available cover-glass is overlaid on a thin agarose bed. A 3D-printed plastic backplate is then affixed to the adjacent vertebral spines using tissue adhesive and dental cement. A stabilization platform is used to reduce motion artifact from respiration and heartbeat. This rapid and clamp-free method is well-suited for acute multi-photon fluorescence microscopy. Representative data are included for an application of this technique to two-photon microscopy of the spinal cord vasculature in transgenic mice expressing eGFP:Claudin-5 &#8212; a tight junction protein.

This protocol describes implantation of a glass window onto the spinal cord of a mouse to facilitate visualization by intravital microscopy.

This protocol describes a method for spinal cord laminectomy and glass window implantation for in vivo imaging of the mouse spinal cord. An integrated ...

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.

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 ...

A Revised Surgical Approach to Induce Endolymphatic Hydrops in the Guinea Pig

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) remain unclear. In order to adequately study the attributes of endolymphatic hydrops, such as the origins of low-frequency hearing loss, a reliable model is needed. The guinea pig is a&#160;good model because it hears in the low-frequency regions that are putatively affected by endolymphatic hydrops. Previous research has demonstrated that endolymphatic hydrops can be induced surgically via intradural or extradural approaches that involve drilling on the endolymphatic duct and sac. However, whether it was possible to create an endolymphatic hydrops model using an extradural approach that avoided dangerous drilling on the endolymphatic duct and sac was unknown. The objective of this study was to demonstrate a revised extradural approach to induce experimental endolymphatic hydrops at 30 days post-operatively by obliterating the endolymphatic sac and injuring the endolymphatic duct with a fine pick. The sample size consisted of seven guinea pigs. Functional measurements of hearing were made and temporal bones were subsequently harvested for histologic analysis. The approach had a success rate of 86% in achieving endolymphatic hydrops. The risk of cerebrospinal fluid leak was minimal. No perioperative deaths or injuries to the posterior semicircular canal occurred in the sample. The presented method demonstrates a safe and reliable way to induce endolymphatic hydrops at a relatively quick time point of 30 days. The clinical implications are that the presented method provides a reliable model to further explore the origins of low-frequency hearing loss that can be associated endolymphatic hydrops.

This article demonstrates an extradural approach to obliterate the guinea pig endolymphatic sac and injure the endolymphatic duct with a fine pick in order to induce experimental endolymphatic hydrops.

Endolymphatic hydrops is an enlargement of scala media that is most often associated with Meniere&#39;s disease, though the pathophysiologic mechanism(s) ...

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to intracranial delivery, intra-arterial administration of NSCs is less invasive and produces a more diffuse distribution of NSCs within the brain parenchyma. Further, intra-arterial delivery allows the first-pass effect in the brain circulation, lessening the potential for trapping of cells in peripheral organs, such as liver and spleen, a complication associated with peripheral injections. Here, we detail the methodology, in both mice and rats, for delivery of NSCs through the common carotid artery (mouse) or external carotid artery (rat) to the ipsilateral hemisphere after an ischemic stroke. Using GFP-labeled NSCs, we illustrate the widespread distribution achieved throughout the rodent ipsilateral hemisphere at 1 d, 1 week and 4 weeks after postischemic delivery, with a higher density in or near the ischemic injury site. In addition to long-term survival, we show evidence of differentiation of GFP-labeled cells at 4 weeks. The intra-arterial delivery approach described here for NSCs can also be used for administration of therapeutic compounds, and thus has broad applicability to varied CNS injury and disease models across multiple species.

A method for delivering neural stem cells, adaptable for injecting solutions or suspensions, through the common carotid artery (mouse) or external carotid artery (rat) after ischemic stroke is reported. Injected cells are distributed broadly throughout the brain parenchyma and can be detected up to 30 d after delivery.

Neural stem cell (NSC) therapy is an emerging innovative treatment for stroke, traumatic brain injury and neurodegenerative disorders. As compared to ...

Stereotaxic Surgical Approach to Microinject the Caudal Brainstem and Upper Cervical Spinal Cord via the Cisterna Magna in Mice

Summary

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Chapters in this video

Spinal Cord Electrophysiology II: Extracellular Suction Electrode Fabrication

Stereotaxic Injection of a Viral Vector for Conditional Gene Manipulation in the Mouse Spinal Cord

Spinal Cord Transection in the Larval Zebrafish

Diffusion Imaging in the Rat Cervical Spinal Cord

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

Cannula Implantation into the Cisterna Magna of Rodents

Laminectomy and Spinal Cord Window Implantation in the Mouse

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

A Revised Surgical Approach to Induce Endolymphatic Hydrops in the Guinea Pig

Intra-Arterial Delivery of Neural Stem Cells to the Rat and Mouse Brain: Application to Cerebral Ischemia