Spinal Cord Window Implantation in a Mouse for In Vivo Imaging

Abstract

Source: Pietruczyk, E. A., et al. Laminectomy and Spinal Cord Window Implantation in the Mouse. J. Vis. Exp. (2019)

This video demonstrates the implantation of a spinal cord window in an anesthetized mouse for in vivo imaging. The protocol stabilizes the exposed spinal cord using a borosilicate cover glass and a 3D-printed backplate. This method enables high-resolution two-photon microscopy to study blood-brain barrier dynamics and neurovascular interactions.

Protocol

All procedures involving animal models have been reviewed by the local institutional animal care committee and the JoVE veterinary review board.

1. 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 (artificial cerebrospinal fluid) 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 vertebrae 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 to the dental cement in the subsequent steps. The articular processes form a natural seat on which the backplate can rest stably (Figure 1). 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.

2. Imaging Preparation

1. Transfer the animal to 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, dichroic, and bandpass filters for the fluorophores present in the tissue6.

Results

Figure 1. Schematic depiction of the anatomic placement of the spinal cord window....

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
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	8009-03-08	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	#################	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