Spinal Cord Lateral Hemisection and Asymmetric Behavioral Assessments in Adult Rats

All surgical and animal handling procedures were performed as approved under the Guide for the Care and Use of Laboratory Animals (National Research Council) and the Guidelines of the Indiana University School of Medicine Institutional Animal Care and Use Committee.

1. General Consideration

1. Use adult female Sprague-Dawley (SD) rats (weighing 200 g, n=12) for this study. Habituate animals to all testing environments and collect baseline data for all behavior tests one week prior to the surgical procedure.
2. Perform the behavioral assessments by two observers who are blinded to the experimental groups.

2. Animal Preparation

1. Clean the surgical table with 70% ethanol. Place a pre-warmed heating pad on the surgical table. Cover the surgical area with a sterile surgical drape. Place the sterile gauze, cotton swabs, and autoclaved surgical tools on the surface of the surgical drape.
2. Turn on a microbead sterilizer for the intersurgery sterilization of surgical tools.
   NOTE: An example of the tools used in this experiment is shown in Figure 1.
3. Anesthetize the rat with an intraperitoneal (i.p.) injection of ketamine (87.7 mg/kg) and xylazine (12.3 mg/kg). Ensure that the proper plane of anesthesia is reached by no response to the toe pinch stimulus. Apply vet ointment to the eyes of the animal to prevent corneal drying during surgery.
4. Remove the hair overlying the thoracic vertebrae by shaving (Figure 2A). Remove the shaved fur with a vacuum equipped with a HEPA filter.
5. Clean the surgical area with three alternating scrubs of iodine-based scrub and ethanol.
6. Cover the animal with a sterile drape with a fenestration over the proposed incision site (Figure 2B). Note; In video, the surgical drape has been omitted for demonstration purposes. 

3. Spinal Hemisection

1. Touch the 13^th rib which is the lowest rib in the rat and a floating rib that does not connect to the sternum. Follow the 13^th rib dorsally to identify its connection with the T13 vertebra and then count up to identify the T10 vertebra.
2. Use a scalpel blade (#15, Figure 1) to perform a 3 – 4 cm midline skin incision on the back overlying the 8-11^th vertebral spinous processes.
3. Under a surgical microscope, bluntly dissect and separate the paraspinal muscles laterally from the spinous processes towards the facets of the T9 and T10 vertebrae on both sides using the same scalpel blade.
   NOTE: This approach will nicely tease the tissue apart without causing hemorrhage.
4. Stabilize the spine using a modified stabilizing holder. Make a slit on both sides of the lateral vertebral bone. Slide the stainless steel arms underneath the exposed transverse process facets and tighten the screws to secure stability.
5. Use a retractor to retract muscles from the surgical area (Figure 2B) and to expose the T8-11 vertebral laminae and spinous processes (Figure 2C).
   NOTE: There is a large gap between the T8 and T9 spinous processes, which are landmarks for identifying T9 (Figure 2C, dorsal view). From the lateral view, the spinous process of the T9 vertebra points caudally, the T10 spinous process points dorsally, and the T11 spinous process points rostrally; so, the 3 spinous processes form a pyramid and the T10 spinous process forms the peak (Figure 2D, lateral view).
6. Perform a dorsal laminectomy on the T9 vertebra using a rongeur. Snip away the T9 spinous process and remove a small portion of the lamina left to the midline (Figure 3A, dashed line), and the entire right portion of the lamina as laterally as possible (Figure 3A, dashed line). For laminectomy, insert the rongeur gently under the lamina and snip a small piece of bone at a time until a desired region of laminectomy is completed (Figure 3B and Figure 3C).
7. Under a surgical microscope, identify the dorsal midline of the spinal cord (Figure 3C). Insert a needle (30 G) vertically through the midline into the spinal cord with the beveled side facing the right side (Figure 4A).
   NOTE: The needle should penetrate the entire spinal cord to reach the ventral wall of the vertebral canal.
8. Stop any bleeding with a small piece of sterile gelfoam.
9. Insert one tip of an iridectomy/microsurgical scissors through the midline needle track, and the other tip along the lateral surface of the right hemi-cord, and then make a complete cut on the right hemi-cord with the scissors (Figure 4B).
   NOTE: Use a sharp micro-scissors for the spinal cord cut to minimize compression lesion to the spinal cord during the cutting.
10. Use the lateral edge of the same needle as a knife to cut through the lesion gap to confirm a complete right hemisection. Verify the completeness of the right hemisection by visualizing the bottom of the vertebral canal with the surgical microscope (Figure 4C, cross-sectional view; Figure 4D, lateral view; Figure 4E, dorsal view).
11. Place a small piece of gelatin sponge over the lesion site (Figure 4F). Use cement mixture and build a narrow bridge over the sponge and the spinous processes of T8 and T10 (Figure 4G, H).
    NOTE: The purpose of using a cement bridge is two-fold: 1) it separates the scar developed at the injury site from the rest of the tissue, and 2) it makes easier to dissect out the spinal cord segment after animal sacrifice.
12. Suture the muscle and skin layers separately with 4-0 silk thread.
13. Inject 0.9% sterile saline subcutaneously to maintain hydration. Inject an analgesic agent Buprenorphine (0.05-2.0mg/kg S) 8-12 h/day subcutaneously for 2 days. Press urinary bladder 2-3 times daily for the first week and 1-2 times in the following weeks till spontaneous bladder voiding returns.

4. Postoperative Animal Care

1. Return the animal to its single-housed home cage. Provide moist rodent chow or gel on the bottom of the cage to aid in animal’s ability to eat/hydrate. Place a heating pad below the cage during post-surgery recovery. Make sure that the heating pad only covers half of the cage bottom to avoid overheating.
2. Inject 0.9% sterile saline subcutaneously to maintain hydration. Inject an analgesic agent Buprenorphine (0.05-2.0mg/kg S) 8-12 h/day subcutaneously for 2 days. Press urinary bladder 2-3 times daily for the first week and 1-2 times in the following weeks till spontaneous bladder voiding returns.

5. Assess the Unilateral Hemisection Stepping (UHS)

NOTE: The unilateral hemisection stepping (UHS) test is a direct measure of the ability of SCI animals to utilize their ipsilesional hindlimb in the open field. As mentioned in 1.1, the animals were acclimated to an open field environment (diameter 42 inches) ¹⁵ twice a day for 7 days. Two observers blinded to the animal groups perform the test. UHS score at both baseline (7 days prior to the T9 HX) and time points after injury will be collected. The steps for assessment are described as followed.

1. Place the animal into an open field environment and examine the animal’s locomotion for 4 min.
   NOTE: During testing, encouragement may be given to the animal to move actively.
2. With the form provided in Table 1, assign a value of 1 for Yes and 0 for No for each behavior category and then sum the total value to give a final UHS score of 0 to 8.
   NOTE: According to Table 1 0: no observable movement of the hindlimb; 1 – 4: isolated movements of 3 hindlimb joints (hip, knee, and ankle); 5: sweeping with no weight support; 6: placing with no weight support; 7: placing with weight support; and 8: stepping with weight support.
3. Collect UHS scores at both baseline (7 days prior to the T9 HX) and time points after injury.
   NOTE: The scores will be assessed at various time points after the T9 HX.

6. Coupling

1. Analyze the CPL (gait coupling) with a video recording an animal walking on a narrow runway device or a simple open-field.
2. In the coupling section of Table 1, assign a score of 0 for “No”, 1 for “Irregular/clumsy”, and 2 for “Normal” for each CPL category.
   NOTE: The coupling (CPL) test is to evaluate the coordination of alternating movements of limbs, including homologous CPL (front-front/rear-rear limbs, Figure 5A), diagonal CPL (front left-rear right or front right-rear left limbs, Figure 5B), and homolateral CPL (front-rear limbs on the same side, Figure 5C). Following a T9 HX, the deficits of the hindlimb on the ipsilesional side become visible resulting in the alternation of the homologous CPL (Figure 5D), diagonal CPL (Figure 5E), and homolateral CPL (Figure 5F).

7. Contact Placing

NOTE: The hindlimb contact placing test is used to evaluate the motor integration of hindlimb responses to proprioceptive stimuli ¹⁶. Proprioception is considered to be intact if the animal steps up with its hindlimb onto the surface after the hindlimb is pulled down below the surface.

1. Hold the animal in a vertical position so that both hindlimbs are available for the placing response.
2. Brush the dorsal surface of a hindlimb forward lightly towards the edge of a surface (e.g., animal working bench).
3. Observe the foot placement onto the surface and assign a hindlimb contact placing score. 0: no placing; 1: placing.
   NOTE: The dorsal surface receives stimulation and the foot will subsequently extend and place the foot on the surface if the reflex is intact. The assessment form is also in Table 1.

8. Grid Walking

NOTE: The grid walking test assesses spontaneous motor deficits and limb movements involved in precise stepping, coordination, and accurate paw placement.

1. Place a rat on an elevated plastic-coated wire mesh grid (36 × 38 cm with 3 cm² openings) and allow it to walk freely across the platform for 30 steps.
2. Count the total number of footsteps and the number of foot missteps for each limb. Video recordings are made to confirm the counting.
   NOTE: Two blinded observers assess paw placement of forelimbs and hindlimbs as animals walk.
3. Assign the grid walking scores for each hindlimb as follows – 0: missteps greater than 15; 1: missteps less than or equal to 15; 2: missteps less than or equal to 10; and 3: missteps less than or equal to 5.
   NOTE: The scoring assessment is based on Table 1. The cutoffs scores are used as measures of severity of motor deficits.

9. Perfusion and Tissue Processing

1. After appropriate anesthesia similar in step 2.3, perfuse the animals carefully following the transcardial perfusion protocol ¹⁷.
2. Dissect and collect the spinal cord samples and post-fix them in 4% PFA overnight.
3. The samples can then be transferred to 30% sucrose solution.
4. Cut the spinal cord in cross sections and stain the selected sections with an axon marker SMI-31 and astrocytic marker glial fibrillary acidic protein (GFAP) according to standard procedures¹⁸.