A Mouse Model of Ankle-Subtalar Complex Joint Instability

The successful establishment of the ASCJ inability mouse model extends a simple ankle inability animal model, and provides a new concepts for the study of the clinical foot diseases. We have established an animal model that's more in line with the actual clinical situation, which can be used for further research on the diagnosis and the treatment of the foot's diseases. One week before the experiment, subject the mice to balance beam and gait training, going from one end of a balance beam or U-shaped pipe to the other, without stopping.

After anesthetizing and removing the hair on the ankle joint of the mouse's right hind limbs, disinfect the exposed skin with an iodine swab. Transfer the mouse to the microsurgery animal operating room using a sterile surgical pad. For the transverse cervical and anterior talofibular ligament groups, make a seven millimeter oblique downward longitudinal incision with a scalpel on the skin above the right ankle joint.

And probe with microscopic straight forceps in the front of the ankle joint. Ensuring the exposure of the anterior talofibular ligament at the lower border of the talus body and the fibula, cut it gently with a scalpel. Separate the long and short fibular tendons, and the extensor digitorum longus tendons.

Expose the cervical ligament, and cut it with a scalpel. For the transverse cervical ligament plus deltoid ligament group, make an eight-millimeter vertical incision on the medial skin of the right ankle joint, and bluntly separate the DL from the medial malleolus to cut it. Then cut the cervical ligament as demonstrated earlier.

For the sham group, perform sham surgery on the right ankle joint. After flushing the incision with sterile normal saline, suture it with 5-0 surgical nylon thread, followed by disinfecting the sutured incision with an iodine swab. Once the angle joint swelling has reduced at two weeks after surgery, exercise the mice in the mouse rotary fatigue machine for one hour every day.

For the balance beam test, clamp a circular wooden beam inclined at 15 degrees at one end with a photography tripod clip, and place the other end on a work surface connected to a closed cassette. Perform balance beam training one week before surgery to ensure that the mice smoothly move from one end of the beam to the other. Consider a mouse to have passed the test when it passes through the beam twice in 60 seconds without pausing.

After each trial, spray the balance beam with 75%alcohol to present the residual odor of the previous mouse from affecting the next mouse. Perform the balance beam test and record the average time of each mouse passing the balance beam twice in a row. And the number of times the right hind foot slips off the beam, as dependent variables.

For footprint analysis, place a U-shaped plastic channel on the experimental table, and connect one end of the plastic channel to a closed cassette. Place a plain pigment paper flat in the channel. Then hold the mouse with both hands, and paint the front and rear feet evenly with non-toxic red and green pigment.

Now, place the painted mouse at one end of the channel, and allow it to move to the other end of the cassette. Take the pigmented paper with the footprints, mark it, and allow it to dry in a ventilated and shaded space. Spray the U-shaped plastic channel with 75%alcohol after each mouse had passed, to prevent the previous mouse's residual odor from affecting the next mouse.

Select three consecutive clear mouse footprints on each paper. Using a ruler, measure the step length of the footprint of the mouse's right foot, with the step width between the left and right footprints. Use microscopic tweezers and scissors to remove excess soft tissue around the ankle specimens.

And then, place the specimens in a centrifuge tube containing a 10%EDTA decalcification solution, and mark the different groups. Next, place the centrifuge tube on a shaking table for decalcification. Change the decalcification solution once per day, and determine the decalcification of the specimens.

After one month of decalcification, dehydrate the specimens with gradient alcohol, and then use N-butanol for eight hours for clearing purposes. Finally, immerse the cleared specimens in paraffin in a coronal position for embedding. Before sectioning, transfer the specimens from the four degree Celsius refrigerator to a minus 20 degree Celsius freezer for about 10 minutes, to facilitate the complete planes cutting.

Fix the specimens on the microtome and section them in a thickness of six micrometers. At the same time, use a microscope to observe whether the samples have been cut at the expected level for easy penetration of a syringe needle into the bone tissue. After cutting two or three complete paraffin sections in a row, transfer them to the tablet machine, set at 40 degrees Celsius, for full expansion.

Then remove the paraffin sections with a glass slide. Finally, mark the slides with groups and numbers. To observe the intact ASCJ joint structure under a microscope.

Place the sections in an incubator set at 60 degrees Celsius for 40 to 50 minutes. Then de-wax the sections with xylene thrice, as described in the manuscript. Place the de-waxed sections in 100%ethanol twice for three minutes, followed by 90%and 80%ethanol for five minutes each.

Then wash the sections with double distilled water for five minutes. After soaking in Hematoxylin for one minute, wash the sections with double distilled water until they become colorless. Soak the sections in 1%acid ethanol differentiation solution for 30 seconds, and wash them with double-distilled water.

Once the sections are stained with 1%ammonia solution, and washed with double-distilled water, stain the samples with eosin staining solution for one minute. And then put them in 95%and 100%ethanol in succession for one minute each. Finally, treat the sections with xylene four for one minute.

Air dry the sections and cover them with a cover slip, after pasting a drop of neutral resin onto the specimens on the slides. Then take images with an upright fluorescence microscope in bright field at five and 20X. Soak the hematoxylin and ammonia-stained sections in 0.05%fast green for two minutes.

Followed by soaking the sections in 1%acetic acid solution for 30 seconds, and 0.1%safranin for five minutes. Then process the slides with ethanol and xylene as demonstrated earlier. Take images with an upright fluorescence microscope in bright field at 5X and 20X.

After three days, eight weeks, and 12 weeks of surgery, the severed ligament groups took significantly longer to pass through the balance beam than the sham group. Before surgery, the right hind foot step length was comparable among the three mice groups. However, 12 weeks after surgery, the step length was shorter in the ligament cut group than in the sham group.

Before surgery, the thermal nociception response time of the mice's feet during activity is similar among the three groups of mice. After surgery, the mice in the ligament cut group displayed longer response times compared to the sham group. The ASCJ of the right hind foot in severed ligament groups showed rough surfaces, osteophyte accumulation, degenerative changes.

And significantly higher bone volume fraction than the sham group. The cartilage layer of the ASCJ in the ligament cut group showed discontinuity, and decreased number of chondrocytes. Also, the Malkin and Osteoarthritis Research Society International scores of the mice with ligament amputation were significantly higher than the sham group.

The type two collagen in the sham group's ASCJ articular cartilage layer of the right hind foot was more uniform, and higher than that of the two groups of mice with severed ligaments. This study can be used for the diagnosis and the treatments for ankle instability, and provide an experimental reference for the clinical treatments of this disease.