Real-Time Imaging of CCL5-Induced Migration of Periosteal Skeletal Stem Cells in Mice

All mice were maintained in pathogen-free conditions, and all procedures were approved by Baylor College of Medicine’s Institutional Animal Care and Use Committee (IACUC).

1. Mouse preparation

1. Cross Mx1-Cre¹⁷ and Rosa26-loxP-stop-loxP-tdTomato (Tom)¹⁸ mice (purchased from the Jackson Laboratory) with αSMA-GFP mice (provided here by Drs. Ivo Kalajzic and Henry Kronenberg) to generate Mx1-Cre⁺Rosa26-Tomato⁺αSMA-GFP⁺ (Mx1/Tomato/αSMA-GFP) reporter mice (Figure 2A). 
2. For Mx1-Cre induction, inject mice with 25 mg/kg pIpC every other day for 10 days. 
3. Irradiate mice with 9.5 Gy 1 day prior to intravenous transplantation of 1 x 10⁶ whole bone marrow mononuclear cells from wild-type C57BL/6 mice (WT-BMT)¹⁹.  
4. After 6 to 8 weeks of recovery, subject mice to the in vivo imaging of periosteal cell migration protocol described below.
   NOTE: Irradiation and WT-BMT of Mx1-Cre mice are necessary to eliminate endogenously Mx1-labeled hematopoietic cells that may respond to an injury. Six to eight weeks post-irradiation, the host's hematopoietic cell count is at <5% and thus ready for imaging¹⁹.

2. Mouse restraint and imaging mount

1. Create a mouse restraint with a 50 mL conical tube (Figure 1A).
   1. Using pointed scissors, poke a hole through the bottom of the conical. Cut from the bottom hole to the top of the conical (the cut side will be the top of the restraint). 
   2. Make a cut approximately 4–5 cm long parallel to the previous cut and halfway down (~2 cm) from the top of the restraint. Remove this section of the conical and smooth the edges were necessary.
2. Construct a conical tube imaging mount using an adjustable angle plate, a 0.5” optical post, and a right-angle clamp for the 0.5” post, 0.1875” Hex (Figure 1B and Table of Materials).
   1. Adjust the angle plate to a 35° angle. On the left side of the angle plate, screw the optical post into the second hole from the back of the plate.
   2. On the right side of the adjustable plate, insert a spring-loaded 0.1875” Hex-locking 0.25” thumbscrew into the second hole from the front and last holes on the plate. The right-angle clamp will be used with the optical post to secure the conical mouse restraint in place.

3. Preoperative surgical procedure

1. Mouse anesthesia
   1. For pain management, subcutaneously inject the mouse with sustained release buprenorphine (1 mg/kg BW) 30–60 min prior to surgery.
   2. Anesthetize mice using rodent III CCM combination (combo III) anesthesia containing 37.5 mg/mL ketamine, 1.9 mg/mL xylazine, and 0.37 mg/mL acepromazine. The appropriate dosage for mice is 0.75–1.50 mL per 1 kg of body weight. The dose will last 30–45 min.
   3. Once the mouse is anesthetized, apply ophthalmic ointment to the eyes with a sterile cotton swab to prevent eye dehydration.
   4. For long-term imaging (>45 min), administer anesthesia again to maintain proper anesthetic depth as follows.
      1. Prepare a 1 mL syringe with a sufficient amount of combo III to maintain anesthesia for the planned imaging duration (0.75 mL/kg BW per 30 min of additional sedation). Attach a 25 G butterfly needle infusion set with 12” tubing to the syringe and push anesthetic through the tubing and needle.
      2. Insert the butterfly needle intraperitoneally, then tape the needle in place. 
         NOTE: This is how the additional combo III will be administered during imaging such that there is no need to move the animal or disrupt the plane of view.
   5. Provide thermal support with a heating pad kept at 37°C throughout the procedure. For prolonged imaging (>1hr) additional fluid support will be necessary.  Supplemental fluids can be provided I.P. with additional Combo III during imaging or 33 mL/kg saline (0.9 NaCl) subcutaneously.
2. Mouse restraint and preparation of the surgical site
   1. Use clippers to remove as much hair as possible from the top of the head. Apply depilatory cream to the shaved area to remove any remaining hair at the surgical site. 
   2. Ensure that the mouse is fully anesthetized by a lack of response to toe pinches. After 15 min, if the mouse is not fully anesthetized, administer a second smaller dose of combo III (<0.75 mL/kg BW).
   3. Place the mouse in the restraint tail-first, sliding it far enough backwards such that the nose slightly hangs over the edge of the conical.  
   4. Tape the front paws to the bottom of the conical tube using medical tape.
   5. Place conical tube with mouse on an imaging mount that is set at a 35° angle and secure using the right-angle clamp (Figure 1C).
   6. Once the mouse is secure in the imaging mount, clean the surgical site 3x with alternating scrubs of betadine scrub and 70% isopropyl alcohol. Again, check that the animal is fully sedated.
   7. Move the mouse to a clean absorbent pad and place a drape over the mouse to create a large sterile field.   

4. Surgical procedure

1. Opening incision
   1. Using fine-tipped forceps, pick up the skin immediately medial to the right ear. With microdissection scissors, cut the skin being held by the forceps. 
      NOTE: Initiating the incision using the method described above creates a rounded incision that is more suitable for suturing after the procedure.
   2. Make a transverse incision starting from the initial incision toward the left ear (<1 cm).  Make another incision, starting from the initial incision, towards the nose of the animal (~2–3 mm past the right eye).
   3. Separate the skin from the periosteum using the forceps and scissors to gently cut through the connective tissue separating the skin from calvaria periosteum. 
   4. Using a sterile cotton swab, apply some ophthalmic ointment to the top of the skin flap.  Gently pick up the skin flap and fold it over the left eye. The intersection of the sagittal and coronal sutures should be clearly exposed (Figure 1E).
      NOTE: The ophthalmic ointment is used to keep the skin flap in place.
   5. Flush the open surface with sterile PBS to clean the area of any blood and residual hair. 
      NOTE: If necessary, use fine-tipped forceps to remove hairs that remain after flushing, but be careful not to damage the periosteum.
2. Microfracture
   1. To generate microfractures on the calvaria, remove the plunger of a 29 G insulin syringe and cut the back end of the syringe, around the 300–400 uL marks.
      NOTE: This step makes it easier to maneuver under a dissection scope when producing the injury.
   2. Place the tip of a bevel (one to two needle widths) towards the nose from the coronal suture and one to two needle widths to the right of the sagittal suture (Figure 1E).
   3. Gently rotate the syringe clockwise, then counterclockwise several times.
      NOTE: In young mice, a very small amount of downward pressure is needed to create a circular microfracture. Be careful not to let the needle penetrate the brain, as this will cause bleeding and interfere with imaging.
   4. Use a 27 G needle to widen the microfracture to ~0.4 mm. If bone fragments remain in the microfracture, flush the area with PBS. If bone fragments still remain in the microfracture, use the 29 G needle to gently scoop them out. 
   5. Repeat steps 4.2.1–4.2.4 to create an injury on the left side of the sagittal suture.
      NOTE: At this point, there are two options: 1) immediately proceed to CCL5 treatment (section 4.3) and imaging (section 5); or 2) close the surgical site following the postoperative procedures in section 6. At 24 h later, anesthetize the mouse again, reopen the surgical sight, treat the injury with CCL5 (section 4.3), and perform intravital imaging (section 5) and postoperative care (section 6).
3. CCL5 treatment
   1. From a stock of RANTES (100 ng/mL), make a solution of 10 ng/mL using basement membrane matrix (Table of Materials). Keep this solution on ice to keep the matrix in liquid form.
   2. Treat each injury with 2 μL of CCL5 (10 ng/μL) by using a 2-20 μL pipette (this pipette tip has a larger diameter, which is more effective when using a viscous liquid). Allow the matrix to solidify.
   3. Once the matrix has solidified, gently cover the calvaria with the skin flap to ensure the tissue does not become dehydrated. Incubate for 1 h prior to imaging.

5. Intravital imaging

1. Microscope settings
   1. Turn on the multiphoton laser (femtosecond titanium-sapphire laser). Set the wavelength to 880 nm and adjust the power for second harmonic generation (SHG) imaging (440 nm) of the bone.
   2. Turn on the 488 nm and 561 nm solid state lasers for excitation of GFP- and tdTomato-expressing cells, respectively.
   3. Turn on PMT (photomultiplier tube) detectors for second harmonic generation (405–455 nm detection) and GFP (505–550 nm detection) signals. Turn on HyD 3 detector for tomato (590–620 nm detection) signal. 
2. Mounting
   1. Before moving the mouse to the scope, check the visual plane of the calvaria by gently pressing the back of the mouse’s head. If the plane is not level, adjust the position by gently rotating the mouse restraint to the left or the right. 
      NOTE: This is a critical step to ensure sufficient image quality.
   2. Apply sterile 2% methylcellulose in water (w/v) to cover the entire calvaria and skin flap and prevent drying of the tissue.
   3. Place the mouse on the XYZ-axis motorized microscope stage (Figure 1G). Using the epifluorescent light, align the mouse so that 1) it is facing to the microscope and 2) the intersection of the coronal and sagittal sutures is the centered reference point of the stage.
   4. Place the glass cover on the imaging area and double-check the alignment (Figure 1H).
3. Time-lapse imaging
   1. Use a low magnification lens (25x water immersion objective with NA = 0.95) to scan the calvaria. Acquire a reference image of the sagittal and coronal suture intersection.
   2. Using the SHG and fluorescent signals from the cells, locate each injury sight and record the XYZ coordinates as well as the distances from the sagittal and coronal sutures (Figure 2B).
   3. Pick a location on either the coronal or sagittal sutures for long-term imaging. Take a detailed Z-stack image of this area from reference during imaging. 
      NOTE: Since the sutures represent a known P-SSC niche, this is where the cells will be migrating from in response to the calvaria injury.
   4. Use the time-lapse software settings to record an image every 1 min for at least 1 h. In the time between snapshots, compare the current field of view with the initial field of view. If they are different, use the Z-stack image to determine in which direction the field of view needs to be adjusted.

6. Postoperative animal care

1. After imaging, rinse the exposed calvaria with sterile PBS to remove the methylcellulose. 
2. To close the incision, gently place the skin flap over the calvaria. Use sterile cotton swabs to remove residual ophthalmic ointment and dry the incision site.
3. Using monofilament nylon 5-0 size nonabsorbable sutures with an attached C-1 reverse cutting needle, close the surgical site. Use a sterile cotton swab to apply triple antibiotic ointment to the closed incision.
   NOTE: Scarring is reduced with high quality suturing techniques and minimum bleeding.
4. Place mouse in a clean cage on a warm surface and check every 15 min until sternal recumbency is achieved.
5. Administer a second dose of sustained release buprenorphine 72 h post-surgery.
6. Monitor daily for huddling behavior, ruffled fur, and/or lack of ambulation until sutures are removed (~1 week).