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

NOTE: The care and experimental manipulation of all mice in this paper was approved by the Cornell University Institutional Animal Care and Use Committee.

1. Setting up for Surgery

1. Sterilize all required tools using an autoclave or approved sterilization procedures. At a minimum, include a scalpel, scissors, vanna scissors, forceps, jeweler’s screwdriver, cotton swabs, a set of retractors, the implant (Figure 1A) and its delivery system (Figure 1D, E).
2. Wipe down all surfaces, including any stereoscope knobs, with 70% ethanol and/or suitable disinfectants.
3. Create a sterile field by overlaying the custom stereotax (Figure 1D) and surrounding benchtop with sterile drapes. A heating pad (preferably feedback controlled) should be placed on top of the stereotax and under the sterile drapes.

2. Preparing the Mouse for Surgery

1. Anesthetize a mouse under 5% isoflurane/oxygen in an induction chamber until breathing rate slows to approximately 1 breath/sec.
2. Transfer mouse to a staging area away from the sterile field, using a nose cone to continue isoflurane exposure. Reduce isoflurane to 2%. Apply eye salve to prevent corneal drying.
3. Administer glycopyrrolate (an anticholinergic drug) at 0.05 mg/100 g mouse intramuscularly into the hindlimbs using 29 – 30 G needles and 3/10 cc syringe, delivering approximately 50% of the injection volume per limb.
   NOTE: Glycopyrrolate serves to prevent fluid buildup in the lungs while the mouse is under anesthesia.
4. Using electrical clippers, shave a portion of the dorsal aspect of the mouse, approximately 3 cm long by 2 cm wide and centered on the thoracic arch. Clean any trimmed hair well away from the shaved area.
5. Administer the following injections subcutaneously and distal to the shaved patch using 29 – 30 G needles and 3/10 cc syringes: 1 ml/100 g mouse 5% glucose in saline (for hydration), 5 mg/kg mouse ketoprofen (an NSAID for analgesia), 0.2 mg/kg mouse dexamethasone (an antiinflammatory steroid to reduce inflammation).
6. Using cotton swabs, perform three alternating washes of iodine and 70% ethanol, waiting 1 min in between washes.
7. Administer 100 µl of 0.125 % bupivacaine (a peripheral nerve block) subcutaneously using 29 – 30 G needles and 3/10 cc syringe to the center of the shaved patch to reduce pain at the incision site.
8. Transfer mouse to the custom stereotax (Figure 1D), insert rectal thermometer, and wait approximately 10 min for bupivicaine to take effect.

3. Expose and Clean the Dorsal Laminae of 3 Adjacent Thoracic Vertebrae

1. Using the scalpel blade, create a small (5 mm or less) incision above the vertebrae of interest (Figure 2A).
2. Using either forceps or by inserting the tip of surgical scissors and spreading the handles, increase the size of the opening in the skin by blunt dissection.
3. Using a scalpel or surgical scissors, gently dissociate connective fascia between the skin and underlying muscle.
4. Use retractors to hold back the skin, allowing for as large a working area as possible (Figure 2B). Take care not to increase the load on the chest and thereby obstruct breathing.
5. Using a pair of forceps, grip the rostral-most vertebra that will be exposed through the overlying muscle. Using a scalpel, cut tissue away on either side of the dorsal process from all 3 vertebrae (Figure 2c).
6. Using any combination of a scalpel blade, cotton swabs, and/or bone scraper, remove all overlying tissue from the dorsal laminae (Figure 2D).
7. Using surgical scissors, carefully cut away tendons attached to the vertebrae on both lateral edges of the spinal column (Figure 2E).
8. Using the scalpel, gently remove as much tissue from the lateral edges of the vertebral column as possible in order to provide a clean clamping surface.
9. Gently debride the vertebrae of any remaining soft tissue using a bone scraper and/or cotton swab. Trim away any incongruous tissue elsewhere using surgical scissors (Figure 2F).
   NOTE: It is essential for both clamping and performing the dorsal laminectomy that the vertebrae are cleanly exposed. A cauterizer may be useful in controlling subsequent bleeding.
10. Shift retractors to hold back the tissue surrounding the vertebral column in addition to the skin (Figure 2F).
11. Attach the implant side bars to the prongs on the delivery posts and load these into the post-holders (as in Figure 1E).
12. Clamp the 3 vertebrae using the side bars, applying just enough pressure to hold securely (Figure 2G, H). Use forceps to help ensure a level clamping of the 3 vertebrae. Note that several attempts may be necessary. Ensure that the side bars are both level and parallel prior to performing the laminectomy.
13. Carefully clean and dry the dorsal surface of the clamped vertebrae using cotton swabs.

4. Perform a Dorsal Laminectomy

1. Insert the tips of the vannas 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 length of the dorsal lamina. Repeat this procedure on the contralateral side to release the lamina.
2. Gently gripping the loose lamina by the dorsal process with forceps, carefully cut away any connective tissue at the rostral and caudal ends of the lamina and lift it away.
3. Using 0.9 % saline and/or artificial cerebral spinal fluid (ACSF), thoroughly wash away any blood overlying the spinal cord. Use cotton swabs to absorb excess fluid.
4. Using vannas scissors, carefully trim the lateral edges of the bone back to the side bars of the implant. Again, wash away and control bleeding using cotton swabs and saline/ACSF.
5. In the event that a portion of the periosteum has detached and overlies the spinal cord, gently remove this tissue using cotton swabs and/or a 29 – 30 G needle. Take care not to injure the spinal cord in this portion of the procedure. Do not confuse the loose periosteum with the tightly wrapped dura mater.
6. Carefully, seal the remaining exposed edges of the bone with one or both of dental acrylic and cyanoacrylate (Figure 2I). Take special care not to allow the glues to flow onto the exposed spinal cord.

5. Seal the Chamber

1. Position the top plate carefully so that the 4 slots line up with the 4 holes on the side bars and the opening overlies the exposed spinal cord (Figure 1B and 2J).
2. Using the jeweler’s screwdriver, carefully start 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 3 screws.
   NOTE: It is useful to use titanium or other non magnetizable forceps to help position the screws at this stage.
3. With all 4 screws in place, tighten each screw alternately by the same amount in small increments until screws are firmly secured (Figure 2K). Note that it is not unusual for the top plate to flex during this process. Take care not to tighten the screws too much, as the screw head can break and excessive downward pressure will dislodge the clamp and/or result in spinal cord injury.
4. Use a silicone elastomer to fill the space between the spinal cord and the glass window. Because it does not produce acetic acid during curing, use KwikSil brand elastomer and mixing tips. Apply a liberal portion of the silicone to the exposed spinal cord. Take care to get rid of any silicone containing air bubbles from the mixing tip prior to application to the spinal cord.
5. Working quickly, gently press a 5 mm #0 coverslip into the inset in the top plate. Apply enough pressure to gradually squeeze the liquid elastomer to surround the spinal cord, but not result in spinal cord compression (Figure 2L). Setting time for the elastomer is approximately 90 sec, so make sure the coverslip is applied rapidly.
6. In the event of a failed sealing, wait until the elastomer has set, gently remove it from the implant with forceps, and repeat 5.4 – 5.5.
7. Cover the edges of exuded elastomer with dental acrylic/cyanoacrylate and adhere to the surrounding bone as much as possible to prevent the elastomer from shifting over the surface of the spinal cord.
8. Remove retractors and pull the skin up around the edge of the implant. Using cyanoacrylate, adhere the skin to the edges of the implant (Figure 2M, N).
9. Insert #0-80 – 1/4” set screws into the wings of the top plate (Figure 1C and 2O).
10. Using dental acrylic, seal any gaps in the screw slots of the top plate (Figure 2P).

6. Postoperative Care

1. Remove animal from anesthesia and place in a large clean cage. Ensure that the cage is sufficiently large that the chamber will not snag on anything (such as the v-shaped dip for a water bottle and/or food) during the course of normal locomotion throughout the cage.
   NOTE: A standard rat cage is ideal.
2. Place the cage on a heated surface while the animal recovers. Do not return the mouse to the company of other animals until it has fully recovered.
3. Check animal behavior within 1 – 2 hr postoperation for signs of pain or distress and implant integrity.
   NOTE: Under normal conditions, the animal should be able to ambulate with modest perturbations to movement.
4. Continue to check animal every 8 – 12 hr for the first 72 hr, measuring weight and monitoring behavior. Check that the animals are active and mobile during the night. Wet food in the form of ground chow mixed with water or nutritional gels, chow pellets, and drinking water should be provided at ground level.  Additional hydration by subcutaneous or intraperiotoneal injections should be provided as necessary.
5. Administer 5 mg/kg mouse ketoprofen and 0.2 mg/kg mouse dexamethasone subcutaneously at 24 and 48 hr time points, postoperatively. Additionally, 0.005-0.010 mg/100 g mouse should be administered every 8 hr by subcutaneous injection for 72 hr .

7. In Vivo Imaging

1. Anesthetize a mouse under 5% isoflurane/oxygen in an induction chamber until breathing rate slows to approximately 1 breath/sec.
2. Transfer mouse to the custom stereotax (Figure 1D) with feedback controlled heating pad and insert rectal thermometer. Reduce isoflurane to 2%.
3. Administer glycopyrrolate 0.05 mg/100 g mouse intramuscularly into the hindlimbs using 29 – 30 G needles and 3/10 cc syringe, delivering approximately 50% of the injection volume per limb.
4. Administer 1 ml/100 g mouse 5% glucose (for hydration) subcutaneously in saline once per hr.
5. Insert the threaded chamber holders into the post-holders used for the surgery (Figure 1F). Adjust the height and twist the posts onto the set screws in the top plate.
6. With both holders attached, elevate the holders in the post-holders until the chest can freely expand upon inspiration.
   NOTE: Both the firm attachment of the holders and the free expansion of the chest significantly reduce motion artifacts due to breathing.
7. Perform imaging as desired.
8. When imaging is complete, lower posts, twist off chamber holders, remove rectal thermometer, and place mouse on a heated surface to recover. Do not return the mouse to the company of other animals until it has fully recovered.