A Neurosphere Assay to Evaluate Endogenous Neural Stem Cell Activation in a Mouse Model of Minimal Spinal Cord Injury

This protocol was approved by the Animal Care Committee at the University of Toronto and is in accordance with the "Guide to the Care and Use of Experimental Animals" (2^nd Edition, Canadian Council on Animal Care, 2017).

1. Minimal Spinal Cord Injury Surgery

NOTE: Prior to surgery make sure that all surgical instruments and materials are sterilized by appropriate methods (Figure 1A).

1. Build a thoracic support arch by rolling up 4–5 squares of gauze and taping them together in the middle to obtain a fixed roll of gauze.
2. Place the mouse in the induction chamber and initiate anesthesia with 5% isoflurane. Confirm the absence of a toe pinch reflex before proceeding, as well as throughout the procedure. As the surgery can take upwards of 30 min to complete, optimize for the minimal isoflurane exposure (5 min at 5% and the remaining time at 2–3%).
3. Transfer the mouse to a nose cone attached to the anesthetic machine at 2–3% isoflurane maintenance dose. Use hair clippers to remove fur from the dorsum of the animal from mid-back up to the neck/ears to expose a wide rectangular area of skin for surgery.
4. Transfer the mouse to a stereotaxic instrument by placing its nose in the attached nose cone and stabilizing the skull with ear bars. Maintain isoflurane at 5% during placement of the ear bars and lower back to 2–3% once the mouse is secured in the stereotaxic device.
5. Administer preoperative analgesic medications intraperitoneally — Meloxicam (2.0 mg/kg). Generously apply eye lubrication onto the open mouse eyes to prevent corneal desiccation when under anesthesia.
6. Place the thoracic support arch underneath the mouse abdomen while straightening out the mouse body and spine by lightly pulling on the base of the tail. Use laboratory labeling tape to secure the tail and all extended limbs in a star-like position. Once secure, push the thoracic support arch rostrally, from the mouse abdomen towards the upper thorax — in order to prop up the thoracic spine (Figure 1B).
7. Prepare a sterile surgical field by disinfecting the fur-clipped skin area prepared in step 1.3 with 70% ethanol, followed by povidone-iodine. Repeat twice. Apply a sterile surgical drape to keep a wide sterile field.
8. Using a #10 scalpel blade, make a vertical incision parallel to the longitudinal axis of the animal from the mid-point of both shoulder blades to the curvature of the thoracic spine. Retract the skin to expose soft tissue and the spinal column contour (Figure 1C).
9. Identify the lower border of the suprascapular fat pad (this demarcates the T4/5 vertebral level). With the same #10 blade, carefully but with force, cut along both sides of the vertebral bone T5–T8/9 to detach the back-muscle tendons from the column.
10. Insert the teeth of retractors into the incision sites on either side of the spine. Adjust the exposure by expanding retractors to sufficiently elevate the spine without putting too much strain on the retracted muscle layers (Figure 1D).
11. Under the surgical microscope, carefully clean the residual muscle and other soft tissue overlying the spine to expose the vertebral bone (Figure 1E). Identify the vertebrae that will be removed by clasping the spinous process of the vertebrae with toothed forceps and moving it slightly up and down.
    NOTE: This should allow identification of intervertebral joints and expose the small openings (intervertebral foramina) underneath the vertebrae of interest.
12. Insert one head of the curved blunted scissors into either side of the exposed intervertebral foramen, caudal to the vertebral lamina to be excised out, and cut the connecting intervertebral joints bilaterally.
13. Lift the lamina upwards and cut off the upper attachment of the lamina to isolate and remove the bone.
    NOTE: This will expose the intact dural sac containing the spinal cord (Figure 1F). There might be excessive bleeding which can be controlled by placing precut 1 cm x 1 cm gauze onto the affected areas and/or wash with sterile PBS.
14. With the dorsal midline vein serving as a landmark, insert a 45° bent shaft of a 30 G needle tip (with needle bevel facing upwards) into the dorsolateral surface of the spinal cord (approximately 1 mm deep) at approximately 0.5 mm lateral to either side of the midline.
15. Move the needle ~2 mm from caudal to rostral (parallel to the midline) so that the entire length of the bevel of the needle is inserted into the cord. Remove the needle by retracing via the path of entry (Figure 1G).
    NOTE: There should be no bleeding but swelling of the spinal tissue may be seen at this step.
16. To close the wound, remove the retractor and suture the back muscles on either side of the injury together in the midline using a 6-0 absorbable suture. Use a 4-0 sterile silk suture to subsequently close the overlying skin.
17. Apply antibiotic ointment to the top of the superficial sutured skin to prevent postoperative infection using the tip of a cotton swab. Administer post-operative analgesic of 0.1 mg/kg of Buprenorphine subcutaneously along with fluids (1 mL of lactated Ringer's solution).
18. Turn off the isoflurane vaporizer and oxygen. Remove the mouse from the stereotaxic device and place in a clean cage with no bedding.
19. Place the mouse in the cage about 30 cm away from a heat lamp for recovery from anesthesia and monitor behavior following waking. The mouse should wake up within 5–10 min and have the hind-limb function with possible paresis and/or tail weakness upon waking.
20. Monitor mouse behavior over the next few days and provide proper post-operative care and analgesics (i.e., mash, lactated Ringer's solution fluid subcutaneous, 0.1 mg/kg buprenorphine 2x daily subcutaneously for 2–3 days, and proper weight monitoring) in accordance to local animal facility regulations and on an individual basis as recovery may vary.

2. Neurosphere Assay Dissection

NOTE: Follow proper sterile tissue culture procedures throughout.

1. Enzymatic Solution Preparation
   1. Add 32 mL of Earle's Balanced Salt Solution (EBSS) to the albumin ovomucoid inhibitor containing glass bottle and gently vortex until dissolved.
   2. Add 5 mL of EBSS to a pre-portioned papain glass vial and place in a 37 °C water bath to dissolve. The solution will become clear (Solution 1).
   3. Take 500 µL of EBSS and add to the pre-portioned DNase I glass vial to obtain a concentration of 1 mg/mL (Solution 2). Mix very gently by tilting the vial back and forth and add 250 µL of this mixture to Solution 1.
      NOTE: All solutions made (Solution 1 and 2) are enough for processing of two pieces of the tissue. If processing more pieces of tissue, accommodate with more solution prep.
2. Tissue Preparation and Dissection
   1. Place a laboratory wipe soaked in isoflurane into the mouse cage and allow 2 – 3 min for vapors to completely anesthetize mouse. Following knockout, remove the mouse from the cage and confirm the absence of a toe-pinch reflex.
   2. Perform cervical dislocation with a blunt metal object (i.e., metal cage card) to euthanize the animal. Spray the euthanized animal from its neck to mid back generously with 70% ethanol.
   3. At the base of the skull, use scissors to cut off the head and discard into a bio bag. Make a midline incision in the skin along the entire length of the back to expose the muscle and vertebral bone contour (Figure 2A).
   4. Lift the medial aspect of each scapula (i.e., shoulder blade — identified by overlying fat pad) and use scissors to cut away soft tissue (between scapulae and vertebrae). Separate fully to expose the underlying vertebral column.
   5. Following the curvature of the spinal column, insert the same pair of scissors into the upper thoracic aperture and isolate the vertebral column by cutting away attached ribs, muscle, and internal organs (Figure 2B).
   6. Insert scissors into the caudal vertebral foramen/opening and cut the intervertebral joints on either side of the laminae (dorsal surface). Take particular care to not damage the intact cord. Continue this process until the desired level and/or length of the cord is exposed.
   7. Carefully cut the spinal nerve(s) extending laterally from the cord prior to transferring the spinal cord tissue into a Petri dish with regular artificial cerebrospinal fluid. Keep the sample on ice (Figure 2C).
   8. Under a dissection microscope, cut the spinal cord tissue to include ~2 mm rostral and caudal to the injury site. Use two pairs of forceps to gently separate the cord into 2 halves longitudinally along the dorsal and ventral fissures (Figure 2C').
   9. With microscissors, cut out the injured dorsolateral white matter. Place the piece into a 15 mL conical tube.
   10. Holding one half of the spinal cord down with forceps, tease and remove white matter using another pair of fine forceps along the rostrocaudal extent of the spinal cord to isolate the desired periventricular region. Repeat for the other half of the spinal cord.
   11. Place pieces of periventricular tissue into a separate 15 mL conical tubes. Use microscissors and gently mince the tissue against the walls of the conical tubes.
       NOTE: Tissue Dissociation Modifications are from Papain Dissociation System Protocol²³ and previously described tissue preparation procedures²⁴.
   12. Add 2.5 mL of new Solution 1 (step 2.1.3) to the tissue sample and place on a rocker at 37 °C for 30 min.
   13. Gently triturate the tissue in solution with a 1,000 µL pipette tip. Centrifuge the cloudy cell suspension at 300 x g for 5 min at RT.
   14. Prepare Solution 3 with 2.7 mL of Earle's Balanced Salt Solution, 300 µL of ovomucoid inhibitor solution (step 2.1.1) and 150 µL of the DNase I solution (step 2.1.3).
   15. Discard the supernatant from step (step 2.2.13) and resuspend in 1.5 mL of Solution 3 from the previous step (step 2.2.14).
   16. Layer this new cell suspension (step 2.2.15) on top of 5 mL of ovoimucoid inhibitor solution (step 2.1.1) in a new 15 mL conical tube to create a discontinuous density gradient. Centrifuge at 300 x g for 5 min at RT.
   17. Discard supernatant and resuspend in 2 mL of neurobasal media. Centrifuge at 300 x g for 3 min at RT.
   18. Discard supernatant and resuspend in 1 mL of neurobasal media. Filter the suspension using a 20 µm cell strainer followed by 4 mL of media for a total volume of 5 mL.
3. Plating
   1. Prepare culture media for neurosphere growth by supplementing neurobasal media with epidermal growth factor (20 ng/mL) / fibroblast growth factor (20 ng/mL)/ heparin (2 µg/mL) [for definitive NSCs] or leukemina inhibitory factor (10 ng/mL) [for primitive NSCs]. Add 10 mL of either media into a 25 mL tissue culture flask.
   2. Use a hemocytometer and Trypan blue dye to calculate the number of cells in suspension from (step 2.2.18)²⁵. Once calculated, add cells to tissue culture flasks prepared in (step 2.3.1) to a clonal density of 10 cells/µL and place tissue culture flask with added cells into the incubator for 24 h (37 °C, 95% humidity, 5% CO₂).
      NOTE: An example of the counting technique is as follows: Take 10 µL of the cell suspension and mix with 10 µL of Trypan blue dye. Add 10 µL of this mixture into a hemocytometer. Under a 10X light microscope, count the numbers of live cells and sum all cells counted within 4 of the 4 x 4 quadrants. Use the formula: 100,000/[(X cells/4) x 2 x 10 x 1,000] to give the volume of cell suspension to be plated in each 25 mL tissue culture flask to ensure clonal density (10 cells/µL).
   3. After 24 h, gently swirl the flask and pour contents into a 15 mL conical tube. Centrifuge at 300 x g for 5 min at RT.
   4. Discard the supernatant and re-suspend the cells in 1–2 mL of fresh neurobasal media.
   5. Repeat step 2.3.2 to count the number of cells in suspension and plate cells at clonal density in a 24-well tissue culture plate containing 500 µL each of their respective mitogen supplemented media (EFH for definitive NSC growth and LIF for primitive NSC growth).
      NOTE: Use the adapted formula (5,000/[X cells/4) x 2 x 10 x 1,000) to calculate the volume of the cell suspension to be plated in each well containing 500 µL of media.
   6. Place plates containing cells in the incubator (37 °C, 95% humidity, 5% CO₂) to grow for 7 days.
4. Counting
   1. Following 7 days in culture, remove plates and view under a light microscope. Quantify neurospheres by counting spherical colonies that are >80 µm in diameter for dNSCs cultures and >50 µm diameter for pNSCs cultures.
      NOTE: If desired, neurospheres can be further passaged²⁶ or differentiated. If cells are no longer needed, dispose by adding accelerated hydrogen peroxide to the wells (approx. 1 mL) for 20 min so that the color of the medium turns yellow. Then discard.