Laminectomy and Spinal Cord Window Implantation in the Mouse

Summary

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

Abstract

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 — a tight junction protein.

Introduction

Transgenic animal models expressing fluorescent proteins, when combined with intravital microscopy, provide a powerful platform for addressing biology and pathophysiology. To apply these techniques to the spinal cord, specialized protocols are required to prepare the spinal cord for imaging. One such strategy is to conduct a laminectomy and spinal cord window implantation. The key features of an ideal laminectomy protocol for microscopy include preservation of native tissue structure and function, stability of the imaging field, quick processing time, and reproducibility of results. A particular challenge is to stabilize the imaging field against the motion induced by respiration and heartbeat. Multiple ex vivo and in vivo strategies have been reported to achieve these goals^1,2,3,4,5. Most in vivo methods involve clamping the sides of the spinal column^2,4 and is often followed by implanting a rigid metal apparatus^3,4 for stability during surgery and downstream imaging applications. Clamping the spinal column can potentially compromise blood flow and induce blood-brain barrier (BBB) protein remodeling.

The purpose of this method is to make the intact spinal cord available for optical imaging in the living mouse while minimizing the invasiveness of the protocol and improving outcomes. We describe a single laminectomy and cover-glass implantation procedure paired with a minimally invasive oval plastic 3D-printed backplate that still achieves robust mechanical stability. The backplate is directly adhered to the anterior and posterior vertebral spines with dental cement. The backplate is equipped with lateral extension arms with screw holes that rigidly attach to the microscope stage via a metal arm. This effectively anchors the intact anterior and posterior vertebra to the microscope stage, providing mechanical resistance to the motion artifact that would otherwise be introduced by respiration and heartbeat. The method has been optimized for laminectomy of a single vertebra at thoracic level 12, omitting the clamps utilized in alternative strategies for stability during in vivo imaging. The procedure is rapid, taking approximately 30 min per mouse.

This protocol can be used to study disease mechanisms of the BBB. The BBB is a dynamic microvascular system comprised of endothelial cells, vascular smooth muscle, pericytes, and astrocyte foot processes that provide a highly selective environment for the central nervous system (CNS). Representative data depict the application of this protocol in transgenic mice engineered to express enhanced green fluorescent protein (eGFP):Claudin-5, a BBB tight junction protein. The provided backplate printing files can also be customized for alternative applications.

Protocol

All experiments follow the University of Illinois, Chicago Institutional Animal Care and Use Committee protocols. This is a terminal procedure. 

1. Reagent Preparation

1. Prepare artificial cerebral spinal fluid (aCSF) to contain 125 mM NaCl, 5 mM KCl, 10 mM Glucose, 10 mM HEPES, 2 mM MgCl₂·6H₂O, 2 mM CaCl₂·2H₂O in ddH₂O. Sterile filter and freeze in individual-use aliquots. Warm aCSF in a water bath to 39 °C before use.
2. Warm low melting-point agarose (2%) in aCSF until fully dissolved in a water bath set to 65 °C. During the laminectomy, cool the melted agarose aliquot to 39 °C in a water bath, so that it can ready at close to physiologic temperature for step 5.2.
   Note: The agarose solution can be stored at -20 °C in single-use aliquots.
3. Prepare sterile 50 mg/mL carprofen in bacteriostatic water. Store at 4 °C.
4. Clean cover-glasses with 70% ethanol, three washes of ddH₂O, and store dry in a dust-free container.

2. Backplate 3D Printing

1. Use 3D CAD software is used to create a model to the dimensions shown in Figure 1. The interior is an ellipse widest at the bottom surface with respect to the printer and cut with a lofted cut to a smaller ellipse forming a lumen at the opposite surface. Two projecting arms with holes to accept screws extend laterally, for attachment to the backplate holding fork. From this 3D structure, create a triangulated 3D mesh file (.STL file).
   Note: See Figure 1B‒D and Supplementary Files 1 and 2.
2. Upload the triangulated 3D mesh file to a 3D printer.
3. Print backplates using a 0.4 mm hot-end nozzle and a 0.2 mm layer height. Select nozzle temperature of 205 °C, bed temperature of 45 °C, and printing speed of 45 mm/s.
4. Assess the resultant 3D printed backplates visually for structural integrity (Figure 1E); gross structural failure (absent lumen, collapsed wall) indicates printing defects (Figure 1F).

3. Surgical Preparation

1. Preheat the heating pad.
2. Load isoflurane into the delivery syringe while working in a chemical fume hood. Attach the delivery syringe to the isoflurane unit.
3. Select an 8‒12-week-old mouse. Weigh the animal. Induce anesthesia using 2% isoflurane in an induction chamber. Inject carprofen at 5 mg/kg subcutaneously.
4. Position the nose-cone and deliver isoflurane at 2% with a flow rate of 150 mL/min for maintenance of a surgical plane of anesthesia (Figure 2A‒E). Wrap the heating pad with a disposable absorbent pad for ease of cleanup.
5. Position an animal on the heating pad at the surgical station and install the nose cone. Lubricate the thermometer with petroleum jelly and insert it 5 mm into the rectum. Tape the thermometer probe to the tail for stability. Apply ophthalmic ointment to eyes of the mouse.
6. To maintain hydration, apply 200 µL of lactated Ringer’s solution by subcutaneous injection every 30 min until termination of the experiment.
7. Spray the dorsum with 70% ethanol, remove fur with clippers, and clean the site with povidone-iodine.

4. Laminectomy

1. Position the animal between the ear bars; these maintain the head position of the mouse with respect to the nose-cone.
2. Confirm that animal is deeply anesthetized as assessed by lack of interdigital pinch reflex and steady respiratory pattern.
3. Make a 1.5 cm rostral-caudal incision at midline over the lower thoracic/upper lumbar region using #11 blade (Figure 2). Separate the skin by grasping it with blunt toothed forceps and/or gloved fingers. Use forceps to separate and peel back any remaining transparent connective tissue underneath the skin. The superficial musculature should now be exposed; displace this with a foam surgical spear.
4. Use foam surgical spears (or a curette) to clear away the remaining, deeper musculature of the target vertebra (thoracic 12). To create a seat for the backplate, also clear away muscle from the posterior aspect of thoracic 11, and the anterior aspect of thoracic 13. Control any bleeding by applying gentle pressure with a surgical spear or use a minimal pulse with a cautery gun. Continue to remove the remaining muscle away from the tendons using forceps
5. Once muscles are removed, carefully detach the tendons by cutting with forceps. There should be plenty of space for visualizing and manipulating the cord when this step is complete. Check that the dura matter of the inter-vertebral space, the semi-transparent laminar bone, the central superficial blood vessel underneath the bone, and anterior radiating artery are now clearly visible.
6. Wet the region with warm aCSF. Use the microdrill to repetitively thin the laminar bone using straight strokes parallel to the long axis of the spinal cord (Figure 2, Figure 3). If desired, utilize a gliding stage to rotate the surgical platform for enhanced ergonomic comfort (for example, a right-handed operator may rotate the surgical platform counterclockwise for the drilling step).
   Note: The gliding stage used is constructed of an upper aluminum plate that slides ±15 mm with respect to the fixed base plate.
7. Gently grasp the superficial spinous process with forceps and lift the vertebra; the bone should lift away easily. If there is resistance, repeat bone thinning with the drill and if necessary use iris scissors, being careful to aim the scissor tips upward to avoid damaging the tissue.
   Note: In order to maintain the dura intact, it is essential not to tug on the bone.
8. Use #4 forceps to clear away any bone shards. Use a surgical spear to apply gentle steady pressure to control any bleeding. Rinse tissue with warm aCSF. Do not allow the tissue to dry out.

5. Cover-glass Implantation

1. Gently apply a 3 mm borosilicate cover-glass to the exposed cord.
2. Ensure agarose is cooled to 39 °C. Using a small spatula, apply warm 2% agarose/aCSF to the edge of the cover-glass and allow capillary action to draw it under the surface.
   Note: At temperatures below 39 °C, the agarose may start to gel. If this occurs, rewarm using a water bath or microwave. Some operators prefer to first apply one drop of agarose and lay the cover-glass on top.
3. Apply tissue adhesive to the exposed bony articular processes of the intact adjacent vertebra at thoracic level 11 vertebral spine and thoracic level 13 vertebral spine. Apply additional tissue adhesive in a ring around the laminectomy site, over the adjacent tendon and transverse process.
   Note: Tissue adhesive is required for proper adherence of the dental cement in the subsequent steps. The articular processes form a natural seat on which the backplate can rest stably (Figure 3). Adhesion to the articular processes will form the strongest points of attachment.
4. Mix dental cement with accelerant in a porcelain mixing tray. Utilize a small spatula to transfer dental cement onto the tissue adhesive layer. Use dental cement to adhere the backplate to the surgical field, centered over the window. Allow 10 minutes for the dental cement to cure.
   Note: The firm adhesion of the backplate to the anterior and posterior articular processes provides the fundamental structural stability of the implant.
5. Use additional dental cement to fill in the interior base of the backplate, and the underside of the backplate : tissue interface.
   Note: The extra application of dental cement improves adherence and reduces risk of leakage of microscope objective immersion fluid (saline) out the bottom of the backplate.
6. Advance the forked backplate holder to the appropriate position over the window. Secure the backplate into the backplate holder with screws.
   Note: This protocol utilized a custom-machined backplate holder (Figure 2G‒H).
7. Apply saline to the backplate to test for leakage. If any fluid leaks, dry the area and apply more dental cement.

6. Imaging Preparation

1. Transfer the animal on the surgical platform to the optical table.
   Note: Our surgical platform, backplate holder, and isoflurane nose-cone holder can be transported between surgical and two-photon imaging stations as one unit, while applying continuous isoflurane anesthesia (Figure 2D,H). Similar units can be assembled from holding forks, beams, and supportive pillar posts obtainable from commercial sources (e.g. ThorLabs). For intravital microscopy, there should be at least 11 inches of clearance between the microscope objective and the optical table to accommodate the height of the surgical platform.
2. Affix the surgical platform to the optical table using a stainless-steel mounting post and counterbored clamping fork.
3. Apply fresh saline into the well of the backplate. Lower a water-immersion lens into the well.
4. Use transmitted or epifluorescence light to identify the area of interest and focus. Switch to laser scanning mode and perform in vivo imaging according to the appropriate two-photon laser excitation wavelength, dichroics and bandpass filters for the fluorophores present in the tissue⁶.

Results

Implanted glass windows and intravital two-photon microscopy provides a useful tool for assessing dynamic changes in CNS proteins. The functional integrity of the BBB is influenced by the expression, subcellular localization, and turnover rates of tight junction proteins⁷. Previous studies have demonstrated that tight junction proteins undergo rapid and dynamic remodeling at steady state⁸. The currently described laminectomy and glass window...

Discussion

The method described here allows for stable imaging of the spinal cord in mice through a glass window. This method has been applied to assess BBB remodeling in transgenic eGFP:Claudin5+/- mice that express a fluorescent BBB tight junction protein, but it could be applied equally well for studies of any fluorescent proteins or cells in the spinal cord.

Multiple methods for laminectomy and spinal cord stabilization have been developed. All protocols address stabilizing the spinal cord during ima...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
3D printer	Raise3D	Pro2	For printing backplates
PLA 3D printing filament	Inland	PLA+-175-B	Black plastic 3D printing material
3D CAD software	Dassault Systemes	Solidworks software	used to design 3D shapes
3D printer software	Raise3D	Ideamaker software	software used to interface with the 3D printer
3D printed oval backplate	custom		Stabilizing imaging field
Surgical dissecting microscope	Leica	M205 C	Equipped with Leica FusionOptics, Planapo 0.63x M-series objective, and gliding stage
Microscope camera	Leica	MC170	HD color camera for visualizing surgical field
Gliding stage	Leica	10446301	The gliding stage is constructed of two metal plates. The base plate is fixed. The upper plate slides on greased interface to allow rotational and linear movement.
Surgical station and stabilization fork	Whale Manufactoring	custom	Laminectomy
SomnoSuite low-flow isoflurane delivery unit	Kent Scientific	SS-01	Surgical anesthesia administration with integrated digitial vaporizer
Stainless steel 1.5 inch mounting post	ThorLabs	P50/M	For mounting surgical station onto optical table for two-photon imaging
Counterbored Clamping Fork for 1.5" mounting Post	ThorLabs	PF175	For stabilizing surgical station mount onto optical table for two-photon imaging
Ideal bone microdrill	Harvard apparatus	72-6065	Thinning bone for laminectomy
Water bath	Fisher Scientific	15-462-10	Warming saline
Cautery gun	FST	18010-00	Cauterizing minor bleeds
Heating pad	Benchmark	BF11222	1.9” x 4.5” silicone heater with 20” Teflon leads, 10W, 5V
K type thermocoupled rectal probe	Physitemp	RET3	Measuring mouse body temperature
petroleum jelly	Sigma	8009-03-8	Lubricating rectal probe
Feedback-regulated thermal controller	custom	NA	Commercially available alternatives include the Physitemp TCAT series
PVA Surgical eye spears	Beaver-visitec international	40400-8	Absorbing blood
Electric trimmer	Wahl	41590-0438	Trimming mouse fur
Blade, #11	FST	14002-14	Surgical tool
Forceps, #5	FST	11254-20	Surgical tool
Forceps, #4	FST	14002-14	Surgical tool
Titatnium toothed forceps	WPI	555047FT	Surgical tool
Titanium Iris scissors	WPI	555562S	Surgical tool
Vetbond tissue adhesive	3M	084-1469SB	Preparing tissue surface for dental acrylic
Ceramic mixing tray	Jack Richeson	420716	Mixing dental acrylic agent with accelerant
Orthojet dental acrylic	Lang Dental	1520BLK, 1503BLK	Permanently bonding backplate to tissue
Small round cover glass, #1 thickness, 3 mm	Harvard apparatus	64-0720	optical window
NaCl	Fisher Scientific	7647-14-5	For aCSF
KCl	Fisher Scientific	7447-40-7	For aCSF
Glucose	Fisher Scientific	50-99-7	For aCSF
HEPES	Sigma	7365-45-9	For aCSF
MgCl2·6H2O	Fisher Scientific	7791-18-6	For aCSF
CaCl2·2H2O	Fisher Scientific	10035-04-8	For aCSF
Carprofen	Rimadyl	QM01AE91	Analgesia
Bacteriostatic water	Henry Schein	2587428	Diluent for carprofen
Isoflurane	Henry Schein	11695-6776-2	Anesthesia
Lactated ringer solution	Baxter	0338-0117-04	Hydration for mouse
Agarose High EEO	Sigma	A9793	gel point 34-37 degrees C
Opthalmic lubricating ointment	Akwa Tears	68788-0697	Prevent corneal drying
MOM Two-Photon Microscope	Sutter