Intravital Imaging of Axonal Interactions with Microglia and Macrophages in a Mouse Dorsal Column Crush Injury

NOTE: All animals should be housed and utilized in accordance with the animal care and use committee (IACUC) at the institution. All procedures described here are approved by the Case Western Reserve University IACUC.

1. Transgenic Animals

1. Use commercially available fluorescent reporter mice to label axons (Thy-1 YFP H)⁵⁰ and microglia / monocytes (CX3CR1^{GFP/GFP 51}; see list of materials). These mice should be intercrossed and maintained as individual breeding colonies according to institutional animal care policies.

2. Bone Marrow Chimeras

1. Identify animals that are at least 8 weeks of age for use as recipient animals.
2. Place host animals in a circular irradiation holding cage designed to hold the mouse in a position to ensure even, whole body irradiation.
3. Set timer on a cesium (Cs-137) irradiator to administer a whole-body radiation dosage of 800-1,000 Rads to the animals according to manufacturer’s instructions (examples shown here are 980 Rads).
4. Return animals to their cages to rest for a minimum of 4 hr.
5. Choose donor animals of the appropriate genotype (see Figure 2A) as the source of marrow progenitor cells.
   NOTE: The appropriate donors are animals expressing different lineage-specific fluorescent reporters from the host to identify different cell populations, or one of the same genotype as controls.
6. Fill one Petri dish with 70% alcohol, and a second dish with 10 ml Roswell Park Memorial Institute (RPMI) media and a 40 µm cell strainer soaked in the media. Fill a 15 ml conical tube with RPMI media. Keep the dishes and tube on ice.
7. Sacrifice the donor animals by CO₂ asphyxiation according to institutional guidelines. Clean the skin and fur by soaking the entire animal with 70% ethanol until all fur is wet.
8. Remove the skin from lower half of animal by cutting around the mid-section and pulling skin down over the hind legs to fully expose the leg muscles.
9. Remove the tibia by anterior overextension at the knee joint to dislocate the bone, then using sterile forceps and scissors, peel and cut muscles away from both ends of the tibia. Place the bone in the Petri dish with alcohol for up to 30 sec then transfer to the cell strainer inside the Petri dish containing the media.
10. Dislocate the hip joint and peel away the muscles attached to the femur, placed the femur briefly in alcohol and then place it in the same cell strainer inside the dish.
11. In a tissue culture hood, use sterile scissors to cut off both ends of the bones and insert a 21 gauge needle on a 1 ml syringe into the end of the bone. Flush the marrow into the 15 ml conical with media. Repeat the procedures for the remaining bones.
12. Using the back of the plunger from a 3 ml syringe, crush bone ends in the cell strainer on the petri dish to extract more cells.
13. Place the filter on top of a 50 ml conical tube and transfer media with aspirated cells from the 15 ml conical tube. Use the back of the plunger to crush the marrow in order to obtain a single cell suspension. Wash the 15 ml conical tube, bone fragments and the cell strainer with 30 ml of media into a 50 ml conical tube.
14. Centrifuge the cells for 5 min at 500 x g to pellet cells.
15. Lyse red blood cells with 2 ml of Ammonium-Chloride-Potassium (ACK) lysis buffer for 2 min. Quench the lysis buffer with 10 ml of media containing 10% fetal bovine serum. Centrifuge at 500 x g for 5-10 min to pellet.
16. Wash cells once in 10 ml of RPMI media and twice in saline, centrifuging at 500 x g for 5-10 min to pellet for each wash.
17. Resuspend cells to a final concentration of 3 x 10⁶ cells in 200 µl of saline.
18. Transfer 3 x 10⁶ marrow cells via tail vein injection into irradiated recipient mice.
19. Place acidified water (pH 3.0) in recipient cage and allow mice to recover.
    NOTE: Mice that fail the transplantation will die within 10-14 days after the procedure.
20. After 8 weeks, verify marrow reconstitution by examining peripheral blood immune cells using flow cytometry (Figure 2B-C).

3. Dorsal Column Crush Injury

NOTE: Choose appropriate chimera animals or double transgenic animals for surgery based on the experiment desired. Animals with either labeled resident or bone marrow derived cells were created in previous steps.

1. Prepare and autoclave surgical tools, including rongeurs, Dumont #4 forceps, iris scissors, forceps for holding tissues, needle drivers and wound clips.
2. Prepare medications to administer to the mice for pain control. Bupernex and Marcaine are commonly used for pain control. Use appropriate medications as required and approved by the Institutional Animal Care and Use Committee (IACUC).
3. Induce anesthesia with 4% isoflurane and maintain with 1-2% isoflurane to achieve a breathing rate of 60-100 breaths per min. Apply eye lubricant to the animal’s eyes to prevent dryness under anesthesia and confirm lack of response by foot pinch. Maintain the temperature using a heating pad and monitor the temperature using a rectal probe.
4. Shave the back of the mouse with an electric shaver over the targeted spinal cord level, and then swab the area three times with povidone-iodine followed by twice with 70% alcohol. Place the animal on a sterile drape, and cover the animal with another sterile drape with a hole cut in an appropriate location to visualize the surgical site. Maintain all tools on sterile drapes throughout the surgery.
5. Cut the skin along the midline at the desired spinal level with iris scissors to expose the muscle on the back of the animal.
6. Under a dissecting microscope, cut the back muscles, including the trapezius and the ligaments of the latissimus dorsi along the midline where they attach to the dorsal spinal processes, and expose the vertebral column.
7. Remove the rotator muscles between the dorsal and transverse processes of the vertebra to reveal the lamina of the specific vertebra to be removed (T11 in this case). Identify T11 by the insertion of the last non-floating rib onto this process.
8. Insert tips of rongeurs into the space above and below the process to be removed and lift slightly to make sure they are securely in place around the vertebral lamina but not too deep to injure the spine. Close rongeurs in one movement to remove the lamina.
9. Enlarge the laminectomy as needed by using the rongeurs to remove small parts of the remaining vertebra.
10. Measure 1 mm from the tips of a #4 forceps by holding them against a ruler and marking with a permanent marker. Measure a 1 mm in separation between the two tines by holding the forceps against the ruler and fix the maximum width of the tips to 1 mm by sliding a rubber forceps ring into place over the shaft of the forceps.
11. Create a small hole by inserting a 30 gauge needle through the dura at the two insertion points for the tines of the forceps to access the spinal cord.
12. Insert the forceps tines at a 90° angle to the dura through the holes to the depth of 1 mm marked on the sides of the forceps, making sure the mark is above the dura. Pinch close the forceps and hold for 10 sec. Release, and repeat forceps squeeze two more times for a total of three holds. Remove forceps from the spinal cord.
13. Soak a piece of absorbable gelatin compressed sponge in saline and place over the injury site.
14. Using a running stitch, suture the muscle in place over the laminectomy and soaked gelatin sponge with #4 synthetic non-absorbable nylon sutures.
15. Clip the skin in place using 5 mm wound clips.
16. Inject 10 µl Marcaine (1.0 mg/kg) in 490 µl of saline subcutaneously around the incision site and 5 µl Bupernex (0.1 mg/kg) intramuscularly into the gluteus muscles.
17. Allow animal to recover on a warm pad and monitor for any difficulties in movement or signs of paralysis in the hind limbs. Do not leave animal unattended or in the presence of other animals until fully recovered from anesthesia. Do not use any animals that display observable motor deficits.
    NOTE: Although this crush lesion can be performed at any level along the spinal cord, practically, T10–L4 pose fewer technical challenges for imaging with respect to breathing and stabilizing body movement using clamps. Clamp placement must be modified around the rib cage to image at thoracic levels.

4. Intravital Imaging

1. Prepare and autoclave surgical tools, including Dumont #4 forceps, iris scissors, forceps for holding tissue, needle drivers and wound clips.
2. Anesthetize the mouse as before with isoflurane. Induce anesthesia at 4-5% and maintain at 1-2% as needed to maintain a breath rate of 60-100 breaths per minute and a lack of response to foot pinch. Keep the mouse heated, monitor breath rate when not imaging and continuously monitor animal temperature with a rectal probe.
3. Clean the skin around the wound clips with Betadine and then alcohol and remove the wound clips according to manufacturer instructions. Once wound clips are removed, remove hair with Nair if needed and clean the site again with povidone-iodine and 70% alcohol.
4. Re-open the skin with scissors. Remove any remaining sutures from muscle and pull muscle apart with forceps, cutting with scissors if needed.
5. Under a dissecting microscope, create pockets for the spinal cord clamps by cutting the muscle with scissors at the side of the transverse processes on the vertebra one vertebral level above and below the laminectomy.
6. Place clamps around each of these vertebra and tighten. Adjust as needed to allow room for microscope objective to reach the spinal cord. Make sure that the spinal cord is parallel to the imaging platform, and maintain tissue stability for imaging. If breathing artifact is seen, readjust clamps accordingly to minimize motion in the imaging field.
7. Cover the clamps with parafilm.
8. Remove the absorbable gelatin compressed sponge from the spinal cord with forceps, delicately picking off as many pieces as possible, also removing as much scar tissue from over the meninges as possible. Cover exposed spinal cord with saline and new sponge if needed.
   NOTE: If any bleeding occurs from the surrounding muscle, either staunch with sponge or cauterize as needed. However, take care not to touch the spinal cord with the cautery. Cover the spinal cord with either a piece of dampened tissue or gelatin compressed sponge when cauterizing.
9. Create a well to hold the immersion fluid by first sealing the skin and tissue with Vetbond, and then using dental acrylic to build up a well between the two clamps of the spinal cord holder and the sides of the animal. Wait until the acrylic cures.
10. Fill the well with artificial cerebral spinal fluid (aCSF) and check again for any bleeding around the surgery site.
11. Optionally at this point, inject fluorescent vessel dyes intravenously by tail vein injection to highlight the blood vessels during imaging.
    NOTE: Fluorescent dextran above 70 kDa is often used, although quantum dots and fluorescently labeled lectins also serve as useful vessel dyes. 50 µl of 150,000 WM TRITC-dextran is typically injected intravenously.
12. To obtain physiologically relevant immune cell motility, the temperature of the mouse must be maintained at 37 °C. Move the mouse to a temperature controlled environment under the microscope for imaging and monitor the animal for correct anesthesia level by breath rate and foot pinch. Monitor the animal frequently during imaging to determine depth of anesthesia, aiming to achieve a respiratory rate of 60 – 100 breaths per minute.
13. Locate the imaging site through the microscope eyepiece.
14. Adjust the 2-photon laser scanning microscope to take a z-stack image every 30-60 sec to obtain dynamic imaging data.
15. Once imaging session is completed remove the animal from the heated box.
16. Loosen the spinal clamps and remove the mouse from the clamps. Pull the acrylic well away from the skin while removing the clamps.
17. If the mouse is to be imaged again, place saline soaked gelatin compressed sponge over the spinal cord. Close the muscle and skin over the surgery site. Do not leave the animal unattended or in the presence of other animals while recovering from anesthesia. If the animal shows any signs of neurologic deficit after recovery, the animal should be euthanized. Otherwise, euthanize the mouse in a CO₂ chamber before it emerges from anesthesia according to IACUC-approved euthanasia protocol.
18. Analyze fluorescent images using image analysis software by following the manufacturer’s instructions.