A Procedure for Implanting a Spinal Chamber for Longitudinal In Vivo Imaging of the Mouse Spinal Cord

In this video, we describe a procedure for implanting a chronic optical imaging chamber over the dorsal spinal cord of a live mouse. The chamber, surgical procedure, and chronic imaging are reviewed and demonstrated.

The overall goal of this procedure is to implant an imaging chamber over the dorsal spinal cord of a mouse to enable longitudinal optical imaging without the need for repeat surgical seizures. This is accomplished by first exposing three thoracic vertebrae and removing soft tissue above the spine. The second step is to clamp the exposed vertebrae using the side pieces of the chamber and perform a dorsal laminectomy.

Next, the top plate of the chamber is firmly attached over the exposed spinal cord and sealed using silicone elastomer and a cover glass. The final step is to seal the skin around the implant using adhesives and allow the animal to recover. Ultimately, multi photon fluorescence microscopy is used to image cellular behavior and tissue structure in the spinal cord with subcellular resolution over time.

The main advantage of our new methodology over existing methods, such as repeated surgical opening of the spinal cord, is the ability to have a nearly arbitrary number of imaging time points spread out over a long period of time without the complications associated with repeated surgery, such as potential infection, surgical trauma, or animal stress. We first had the idea for this method when we were considering the previous success of longitudinal imaging following spinal cord injury in the transparent zebra fish. We then decided to expand this paradigm to mammalian model.

Begin this procedure by shaving the thoracic arch area of a mouse anesthetized using isof fluorine. Then remove the trimmed hair. Next, administer the mixture of saline, ketoprofen, and dexamethasone to the mouse subcutaneously.

Using cotton swabs. Perform three alternating washes of iodine and 70%ethanol, and wait one minute in between washes. After that administer 100 microliters of 0.125%Bupivacaine to the center of the shaved patch subcutaneously to reduce pain at the incision site.

Then transfer the mouse to the custom stereo attacks. Insert the rectal thermometer and wait approximately 10 minutes for the bupivacaine to take effect under the microscope. Create a five millimeter incision above the vertebrae of interest with a scalpel blade.

After that, increase the size of the opening in the skin by blunt dissection. Next, gently dissociate the connective fascia between the skin and the underlying muscle. Hold the skin back for a larger working area with the retractors and to be careful not to increase any weight on the chest and obstruct breathing.

Then grip the rostral, most vertebra that will be exposed through the overlying muscle with a pair of forceps. Cut the tissue on both sides of the dorsal process away from three vertebrae. Subsequently, remove all the overlying tissue from the dorsal laminate.

Afterward, carefully cut away the tendons attached to the vertebrae on both lateral edges of the spinal column. Gently remove as much tissue from the lateral edges of the vertebra column as possible in order to provide a clean clamping surface. Then gently debride the vertebrae of any remaining soft tissue.

Using a bone scraper and cotton swab, trim away any incongruous tissue using the surgical scissors. Next, adjust the retractors to hold back the tissue surrounding the vertebra column. Together with the skin, attach the implant sidebars to the prongs on the delivery posts and place them into the post holders.

Next, clamp the three vertebrae. Using the sidebars by applying just enough pressure to hold them securely. Then use the forceps to help ensure a leveled clamping of the three vertebrae.

Make sure that the sidebars are both leveled and parallel prior to performing the laminectomy. After that, carefully clean and dry the dorsal surface of the clamped vertebrae using cotton swabs. In this procedure, insert the tips of the Vana scissors into the epidural space of the medial clamped vertebra and squeeze the handles lightly.

If the bone is dry, it should crack along the dorsal lamina. Repeat this procedure on the contralateral side to release the lamina while gripping the loose lamina Gently by the dorsal process with forceps, carefully cut away any connective tissue at the rostral and coddle ends of the lamina. Then thoroughly wash away any blood overlying the spinal cord with 0.9%saline or artificial cerebral spinal absorb excess fluid with cotton swabs.

After that, trim the lateral edges of the bone and proceed to the sidebars of the implant. Subsequently control the bleeding using cotton swabs and saline. In the event that a portion of the periosteum has detached and overlies the spinal cord, gently remove the tissue using cotton swabs and a 29 to 30 gauge needle.

Be careful not to injure the spinal cord and do not confuse the loose periosteum with a tightly wrapped dator. At the end. Seal the remaining exposed edges of the bone with dental, acrylic, and cyanoacrylate.

Take special care to keep the glue from getting onto the exposed spinal cord. Now, position the top plate on the animal carefully so that the four slots line up with the four holes on the sidebars and the opening overlies the exposed spinal cord. Using a jeweler screwdriver carefully start with the first screw, stabilizing the shaft of the screwdriver with another pair of forceps.

Do not tighten the screw at this stage, but insert it so that the first few threads are engaged. Repeat this process for the other three screws. With all four screws in place, tighten each screw alternately in the same small increments until all the screws are firmly secured.

Next, use the quick sill brand silicone elastomer to fill the space between the spinal cord and the glass window and apply it to the exposed spinal cord. Be sure to get rid of any silicone containing air bubbles from the mixing tip prior to its application. Then quickly place a five millimeter cover slip into the insert in the top plate.

Apply enough pressure to gradually squeeze the liquid elastomer to surround the spinal cord but not resulting in spinal cord compression. Afterward, cover the edges of the exuded elastomer with dental, acrylic and cyanoacrylate, and use the adhesives to adhere the elastomer to the surrounding bone as much as possible to prevent the elastomer from shifting over the surface of the spinal cord. Next, remove the retractors and pull the skin up around the edge of the implant.

Use the cyanoacrylate to adhere the skin to the edges of the implant. After that, insert the screws into the wings of the top plate. Subsequently seal the gaps in the screw slots of the top plate using dental acrylic before placing the animal back in the cage for recovery.

Before the imaging sessions, anesthetize the mouse again and attach the exposed screws of the implant to the threaded posts for stabilization. This figure shows that a chronic spinal chamber remains optically transparent over multiple weeks. These are white to light images of the spinal cord ranging from several minutes to three weeks post-operation.

The vascular landmarks are identifiable at all the time points. Little change is seen at one day post-surgery. Dense fibrous overgrowth is present in part of the window as early as three days and results in partial obfuscation of the imaging area.

Mild neovascularization is also present but does not obscure the original vasculature in white light or two PEF imaging. The chronic spinal chamber allows repeated multi photon imaging without the need for repeated surgeries Shown. Here are the images of a transgenic mouse expressing YFP in a subset of DRG axons in the dorsal fii of the spinal cord.

The vasculature was labeled by intravascular dye injection shown in red. The activity of microglia can also be tracked over time in thigh one YFP CX three CR one GFP double transgenic mice while axons and microglia remain visible. Contrast and resolution decline modestly over time.

Once mastered. This technique can be done in approximately one hour if it is performed properly After its development. This technique paved the way for us to explore the temporal and spatial heterogeneity of axon dieback after a spinal cord injury.

And mice.