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

Summary

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.

Abstract

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.

Introduction

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 atlases^2,3, 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 lambda^4,5, 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 injections^8,9.

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, mouse^8,9, 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.

Protocol

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.

1. 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).
2. Prepare autoclaved surgical instruments (Table of materials) and place them on a sterile surface.

2. Anesthesia induction and mouse preparation

1. 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.
2. Place the mouse (10-week-old male C57BL/6J) in the induction chamber.
3. Once breathing has slowed, open the induction chamber, and lift the mouse slightly. Use clippers to remove hair from head to shoulders.

3. Positioning of the mouse in the stereotaxic frame

1. 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.
2. Place the mouse in the stereotaxic frame using ear bars only.
   NOTE: Make sure the ear bars are even and the head is level.
3. Anteroflex the head of the mouse to a 90° 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° 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° between the plane of the frontal skull bone and the plane of the surface of the table as outlined in Figure 2A,C.
4. 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° 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.
5. Inject a single dose of 4 mg/kg Meloxicam slow-release (SR) subcutaneously (s.c.) at a volume of 2 µL/g body weight and place lubricant on the eyes.
6. 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.
7. Place a drape underneath the body.
8. Disinfect hands and put on sterile gloves.
9. Place a drape at the surgical site.

4. Surgery to access the cisterna magna

1. Make sure the mouse is appropriately anesthetized by pinching the toes or checking the corneal reflex.
2. Reduce isoflurane to maintenance levels (2.0).
3. 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.
4. 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.
5. 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.
6. Position the surgical microscope in place to better visualize the surgical field.
7. 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.
8. Reposition the retractors and adjust the tension by repositioning the hemostats, opening the view of the cisterna magna.
9. Use the blunt laminectomy forceps to separate the muscles further in the midline to get a good viewing window of the brainstem and cerebellum.
10. Repeat steps 4.7-4.9 as needed until the cerebellum and brainstem come in view below the dura.
11. 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.

5. Opening of the cisternal membrane

1. 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.
2. 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.
3. Once the dura is opened, drain excess cerebrospinal fluid with a sterile cue tip.

6. Identification of landmarks and zero point

1. 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.

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.

1. 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.
   1. For target distance >0.4 mm from caudal to the obex (Figure 1B, green) perform the following.
      1. 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.
      2. 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.
      3. 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.
   2. For target distance <0.4 mm from caudal to the obex (Figure 1B, orange) perform the following.
      1. 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.
      2. 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.
      3. 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.

8. Injection of the target

1. 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.
2. Repeat step 8.1. for multiple targets.

9. Closure of the surgical field

1. 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.
2. Close the skin with three nylon or polypropylene sutures (5-0 or 6-0).

10. Post-operative care

1. 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.
2. Monitor health status, weight, and sutures on post-operative days 1-3. Remove sutures on day 10 if not already removed.

Results

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...

Discussion

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

Materials

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