Tricolor Transgenic Murine Model for Studying Growth Plate Injury

The research was performed in compliance with institutional guidelines. All animal procedures were approved by the University of Connecticut Health Center Institutional Animal Care and Use Committee (IACUC) prior to initiating the work. An outline of the protocol is described in the Figure 1 schematic.

Figure 1: Outline of the growth plate injury protocol in tricolor collagen reporter mice. Please click here to view a larger version of this figure.

1. Mouse breeding and preparation for surgery

1. Breed tricolor transgenic collagen reporter mice that express Col1a1-GFPTpz, Col2a1-CFP, and Col10a1-mCherry^{16,17,18,19,20,21} per standard breeding procedures to obtain pups for surgery that are 2 weeks old (± 1 day). Verify the triple transgene expression by genotyping a tail tip fragment under a fluorescence microscope. Only use the mice that are positive for all three colors to maximize the benefit of the reporter system during subsequent histological analysis.
   NOTE: This age of 2 weeks old (± 1 day) is selected because their bone growth plates are at a similar stage of development to the human adolescent²². This protocol applies to both sexes of mice.
2. On the day of surgery or one day before, uniquely identify each mouse used for surgery using an ear punch after disinfecting the ear and ear punch with alcohol antiseptic pads, weigh each mouse, and record the value.
3. Shave the entire right hind limb from the hip to the foot with an electric clipper while the mouse is under the effect of isoflurane anesthesia. To induce anesthesia in mice, use a mixture of isoflurane (2-3%) and 100% oxygen, administered at a flow rate of 1 L/min within a 1-2 L induction chamber. Remove the mouse and confirm that the anesthesia depth is sufficient with a toe pinch that should not cause the mouse to move.
   NOTE: This depth of anesthesia will last sufficiently long to conduct the shaving without requiring an anesthesia nose cone.
4. Employ X-ray imaging with a power setting of 26 kV (800 mA) to capture tibial images in live mice under isoflurane-induced anesthesia to record initial limb length.
   1. Prior to placing the mice in the X-ray cabinet, induce a profound state of anesthesia using a mixture of isoflurane (2-3%) and 100% oxygen, administered at a flow rate of 1 L/min within a 1-2 L induction chamber.
   2. Place three anesthetized mice at a time in parallel on their stomachs in the X-ray cabinet at a shelf height that allows for one image to be obtained that includes all three mice. Splay the legs so the tibial bones are not obscured under the mouse. For enhanced measurement accuracy, position a radiopaque scale near the mouse during imaging (Figure 2A).
5. Return the prepared mice to their housing cage with the mother mouse to await subsequent surgical procedures.

2. Preparation of surgical supplies and sterile work area

1. Sterilize the following items: 10-20 gauze pads, cotton-tipped applicator swabs, dental burs (0.5 mm diameter), dental handpieces, Graefe forceps, curved mosquito hemostatic forceps, curved fine scissors, needle holders, periodontal probes, and dental cleoid discoid carvers.
2. Collect additional necessary sterile supplies: an injectable suspension of buprenorphine, 20 G needles, 1 mL syringes, towel drapes, 5-0 undyed, braided, coated, vicryl sutures, povidone-iodine, phosphate-buffered saline, alcohol antiseptic pads, #15 scalpels, surgical gloves, environmental surface barrier tube sox, eye lubricant, 70% ethanol spray. Power on a glass bead sterilizer for additional sterilization of items during surgery and the electric heating pad under a clean mouse cage with sterilized bedding.
3. Clean and sterilize the fluorescence stereomicroscope stage, adjacent surfaces, and the surgical instrument stand using 70% ethanol. Cover these areas that will become the surgical workspace with sterile towel drapes.
4. Assemble the electronic high-speed dental drilling system. Connect the electronic foot controller to the control unit and cover the handpiece cord with a disinfected surface barrier tube sox. Attach the sterile 0.5 mm round dental bur. Turn on the controller and set it at a drive ratio of 1:1 and a max of 30,000 rpm.
5. Secure, with tape, the flexible isoflurane machine hose that terminates with the nose cone covered with a disinfected surface barrier tube sox on the fluorescence stereomicroscope stage.
6. Power on the fluorescence stereomicroscope and ancillary equipment and open the image acquisition software.

Figure 2: Key steps of the trilineage fluorescent reporter murine growth plate injury procedure. (A) Measuring tibial length by faxitron X-ray imaging using a radiopaque ruler placed alongside the mice in the X-ray cabinet. (B) Proper position of an anesthetized mouse for surgery under a fluorescence stereomicroscope. Proximal tibia indicated by a black arrow. (C) An example of an incision made to access the growth plate. (D) Fluorescence stereo microscopy illuminating the hypertrophic zone of the growth plate (white arrow). The adjacent proliferative zone is indicated by a yellow arrow. (E) Bright light illumination of the surgeon placing the 0.5 mm dental bur against the growth plate. (F) Precise placement of the dental bur is guided by fluorescence stereo microscopy into the hypertrophic zone. (G) An example of a Salter-Harris Type II-like growth plate defect (white arrow). Scale bars = 1 mm. Please click here to view a larger version of this figure.

3. Proximal tibia growth plate injury procedure

1. Anesthetize the mouse in the isoflurane anesthesia chamber, adjusting the isoflurane concentration to 2-3% and oxygen flow rate to 1 L/min. Await the induction of a profound anesthetic state, verified by a toe pinch test and the observation of respiratory patterns.
2. Administer half the prescribed dose of buprenorphine subcutaneously immediately following removal of the mouse from the chamber. The dosing of buprenorphine is as per the approved animal protocol.
3. Apply ocular lubricant to protect the mouse's eyes from drying out during surgery and transfer the mouse in a supine position to the isoflurane machine nose cone on the stereomicroscope stage (Figure 2B). Adjust the anesthesia flow through the nose cone to 2% and the oxygen flow rate to 1 L/min.
4. Disinfect the right hind limb, pelvic region, anterior aspect of the left hind limb, and tail sequentially with povidone-iodine followed by 70% ethanol.
5. Verify a continued stable depth of anesthesia through an additional toe pinch test and observation of respiratory patterns before surgery commencement.
6. Under bright light illumination, employ a #15 scalpel to create an incision in the skin just below the knee joint with an initial length of approximately 5 mm, to reveal the proximal end of the right tibia (Figure 2C). Use scissors to extend the cut if necessary. Keep the left contralateral tibia uninjured so that it serves as an internal uninjured control.
7. Perform a vertical blunt dissection through the overlying muscle at the proximal tibia using the back side of the #15 scalpel, removing the soft tissue for clear exposure of the growth plate.
8. Turn off the operating room lights and select the correct fluorescence channel to illuminate the desired region of the growth plate. Filter sets required to image each of the fluorescent collagen are Col 2 Cyan: ET436/20x (excitation), ET480/40 m (emission), Col 10 mCherry: ET577/20x (excitation), ET640/40 m (emission), Col 1 Topaz: ET500/20x (excitation), ET535/30 m (emission). Position the mouse to observe the proximal tibial growth plate (Figure 2D). Adjust the skin opening slightly more proximal and then, distal to ensure the tibial growth plate is in view and not the femur growth plate.
9. Create a Salter-Harris Type II-like lesion while looking through the microscope eyepiece by positioning the 0.5 mm dental drill bur parallel to the tibial axis in the middle of the hypertrophic growth plate zone, employing a lateral approach (Figure 2E,F). To reproducibly make a defect that mimics a Salter-Harris Type II lesion, keep the limb parallel to the work surface so that the bur entry pathway will not angle into the epiphysis. Apply pressure on the drill pedal to initiate bur rotation and gently depress the bur into the growth plate stopping before the defect goes any deeper than the bur diameter (Figure 2G).
   NOTE: A second pair of hands assisting in stabilizing the mouse limb is helpful.
10. Irrigate the lesion site with a drop of sterile PBS to remove any debris.
11. Confirm the defect's depth at 0.5 mm using a periodontal probe.

4. Post injury procedures and closure

1. For time zero defect characterization, harvest the injured hind limb and control limb prior to tissue closure. Follow the tissue harvesting and preparation for subsequent microcomputed tomography (microCT) and cryo-histology imaging in section 6.
   NOTE: For experiments involving the implantation of a therapeutic substance in the growth plate defect, the defect itself can only accommodate a piece of biomaterial that is roughly spherical and 0.5 mm in diameter or 0.082 mm³ (0.082 µL) in volume.
2. Utilize forceps under microscopic visualization to insert the biomaterial or inject a liquid therapeutic into the defect within the growth plate. Use a larger volume of biomaterial or injected substance if the study design does not require the therapeutic to be limited to only the defect.
3. Carefully realign the skin edges, ensuring the implanted material, if any, remains securely within the defect site.
4. Employ interrupted suturing techniques with 5-0 polyglycolic acid sutures to seal the skin incision effectively.
5. Thoroughly cleanse the areas beyond the surgical region of povidone-iodine residue using swabs with sterile water, applying care to avoid contact with the wound and closely surrounding areas.
6. Remove the mouse from the isoflurane nose cone on the microscope stage to a recovery cage, positioning it laterally on fresh bedding over a heating pad.
7. Closely observe the mouse's respiratory rate for approximately 5 min, administering the remaining buprenorphine dose for analgesia just as the isoflurane anesthesia's effects diminish, indicated by an increase in respiratory rate, but before the onset of mobility. Maintain the mouse in an isolated recovery cage until it demonstrates full ambulatory capabilities, thereafter, reuniting it with its mother and siblings.
8. Conduct daily inspections for the initial 48 h post operation, followed by weekly assessments up to the point of euthanasia. Focus on signs of infection, locomotion efficacy, accessibility to food, and the integrity of sutures.
9. Wean the mice at the standard time (i.e., 3 weeks of age).

5. Limb length measurements

1. Based on the time points determined by the experimental hypothesis and design, anesthetize the mice with isoflurane and conduct X-ray imaging of the full length of both the injured and uninjured tibiae as shown in Figure 2A and described in section 1.4. Suitable landmarks for making consistent limb length measurements include the top of the tibial epiphysis and the distal end of the tibia at the tibiotalar joint as shown in Figure 3.
   NOTE: Imaging is used to assess limb length and limb length discrepancies due to the injury, as well as the development of the bony bridge within the growth plate tissues at specific post-operative intervals.

6. Tissue dissection, fixation, microCT imaging, and embedding

1. Euthanize the mouse via CO₂ asphyxiation preferably using an automated CO₂ induction system with a confirmation of death by an alternate method such as cervical dislocation.
2. Isolate both intact hind limbs and remove skin and muscle from the bone and knee capsular area in preparation for histological and microCT analysis.
   1. Before putting the hind limbs into the fixative, excise the patella carefully by cutting it off using micro-dissecting scissors to facilitate fixative penetration into the knee cavity. Employ a 29 G insulin syringe to thoroughly distribute cold 10% buffered formalin within all areas of the knee cavity. Severe the diaphyseal region of the femur and tibia to improve fixative access to the marrow space. Maintain the joint tissue in a fully extended position within the fixative for 24-36 h at 4 °C by tying it with gauze to a thin dowel.
      CAUTION: Formalin is toxic and should be handled in a fume hood while wearing appropriate personal protective equipment.
3. After 24-48 h of formalin fixation at 4 °C, conduct high-resolution X-ray microCT of contralateral and injured samples by microCT imaging after transferring samples to PBS, for assessment of bony bridge development. Use a voxel size of 6.0 µm, sample times of 330,000 ms, and energy settings of 55,000 V with an intensity of 145 µA.
4. After microCT imaging and an additional 24 h of formalin fixation at 4 °C, rinse the samples in 1X PBS for 3 x 5 min and then, immerse samples in 10% sucrose in 1X PBS for 1 h, 20% sucrose in 1X PBS for 1 h, and 30% sucrose in 1X PBS overnight. Use a 29 G insulin syringe filled with sucrose solution to ensure the sucrose solution permeates all knee cavity regions. Transfer to a -80 °C freezer if not embedding after the overnight sucrose immersion to ensure retention of the GFP reporter and tissue enzymatic activity.
5. Remove any residual muscle tissue from the joint region before embedding. Equilibrate the sample with a cryo-embedding medium overnight, facilitating medium penetration into the knee cavity.
6. View the microCT images to determine where the defect and bony bridge are located prior to embedding. Apply a thin layer of the cryo-embedding medium into a cryomold and place the dissected and still connected tibia/femur/joint such that the region of interest faces 90° to the surface of the embedding medium.
   NOTE: This orientation allows the sectioning to capture the lateral view of the laterally created defect. If the bony bridge is near the edge of the defect, then this orientation will also avoid the possibility of missing the region of interest during the first thicker sections taken through the tissue.
7. Position the cryomold on dry ice until the cryo-embedding medium solidifies, securing the specimen. Continue to fill the cryomold with the medium while keeping it on dry ice.
8. After securing the specimen, immerse the cryomold in 2-methyl-butane chilled with dry ice until fully frozen. Post freezing, remove excess 2-methyl-butane, wrap the cryomolds in cellophane, and store at -20 °C or -80 °C.
   NOTE: For prolonged storage beyond 1-2 months in a cryo-embedding medium, -80 °C is recommended to prevent drying out. Tape-assisted cryosectioning and adhesion of the sections to the glass slides are described previously²³.

7. Sequential imaging, staining, and reimaging

1. Implement a sequential procedure of imaging, staining, and reimaging to facilitate the detection and colocalization of multiple biological signals within the same tissue section as described previously²³. Specifically for the growth plate sections from the tricolor collagen reporter mice, the following steps are recommended. This ordered approach ensures comprehensive visualization of both molecular signals and structural details.
   1. Initially, capture endogenous fluorescent signals for the three collagens in the first round of imaging.
   2. Apply calcein blue staining and image for mineralized tissue²³, followed by the tartrate-resistant acid phosphatase (TRAP) enzymatic activity staining and imaging²³.
      NOTE: While the TRAP enzymatic activity staining results are not displayed here, this staining step is performed as it is essential for the removal of mineral before the Safranin O/Fast Green staining results presented in the representative results section.
   3. Conduct 4',6-diamidino-2-phenylindole (DAPI) nuclear staining as described below and then image.
      1. Place slides that have undergone prior imaging into a Coplin jar containing 1x PBS. Leave them submerged until the coverslips detach from the slides. Once detached, remove and thoroughly dry the coverslips.
      2. Apply DAPI counterstaining solution to the coverslips, utilizing a 1:1,000 dilution of DAPI in a mixture of 50% glycerol and 1x PBS. Proceed with the imaging process post application.
   4. Conclude with the application of Safranin O/Fast Green stain to accentuate tissue architecture and then image.
      NOTE: Perform this as the concluding step to correlate chromogenic images with fluorescent signals.
      1. Prepare Weigert's Iron Hematoxylin solutions. Prepare Solution A by dissolving 1 g of hematoxylin in 100 mL of 95% ethanol and Solution B by combining 4 mL of 29% Ferric chloride solution, 95 mL of deionized water, and 1 mL of concentrated HCl. Mix equal parts of Solution A and B to create Weigert's working hematoxylin solution, stable for approximately 4 weeks.
      2. Submerge slides in deionized water for 2 x 2 min to hydrate.
      3. Apply Weigert's working hematoxylin solution for 5 min.
      4. Wash slides in tap water for 5 min, then briefly in deionized water for 1 min.
      5. Stain with 0.2% Fast Green solution (0.2 g of Fast Green FCF in 100 mL of deionized water) for 2 min.
      6. Briefly rinse in 1% acetic acid for 1 min.
      7. Stain with 0.1% Safranin O solution (0.1 g of Safranin O in 100 mL of deionized water) for 1 min.
      8. Rinse in deionized water until achieving a visually balanced color, approximately 5 min.
      9. Mount slides in 30% glycerol in deionized water (avoid PBS) and proceed to imaging immediately to prevent color diffusion from the tissue. To accentuate the resting zone cells of the growth plate conduct image with a Cy5 filter: ET640/30x (EX), ET690/50 m (EM).