This study was supported by grants from the Sichuan Provincial Health and Wellness Committee (Grant Number: 21PJ063) and the National Natural Science Foundation of China (Grant Number: 82001031).

All animal procedures in this study were reviewed and approved by the Ethical Committee of the West China School of Stomatology, Sichuan University (WCHSIRB-D-2020-114). Male C57BL/6 mice (5 weeks old) were raised in a humidity-controlled (53 &#177; 2%) and temperature-controlled (23 &#177; 2 &#176;C) facility and were on a 12 h light/dark cycle. See Table of Materials for details related to all materials, reagents, and instruments used in this protocol.
1. Freezing injury
<ol>
	<li>Anesthesia: Sedate the mice with isoflurane-soaked cotton balls and inject tiletamine-zolazepam intraperitoneally at a dose of 50 mg/kg. Pre- and post-operative analgesia was provided by intraperitoneal injection of meloxicam at a dosage of 1 mg/kg. Protect their eyes with vet ointment. A heating pad maintained at 38&#160;&#176;C was put under the surgical drape during the whole procedure to provide thermal support for the animal. Fix the mice with adhesive tape after laying them on their side on a surgical table (Figure 1). Assess the sufficiency of anesthesia by the loss of eyelid reflex, righting reflex, and pedal withdrawal reflex. 
	NOTE: When applying isofluorane anesthesia, hold the mouse in one hand and the isofluorane-soaked cotton ball in another. Make sure to keep the cotton ball at a distance from the animal to avoid discomfort and skin irritation.</li>
	<li>Preoperative hair removal: Use an electric razor to shave the hair on the calf and face of the mouse and apply the depilatory paste with a cotton swab onto the skin, covering the masseter muscle and the tibialis anterior muscle. After 1-2 min, wash away the paste with ethanol-based surgical wipe (Figure 2A and Figure 2G).&#160;The surgical sites were disinfected by three rounds of an iodine-based scrub and 75% ethanol.&#160;&#160; 
	NOTE: The electric razor alone was not sufficient for hair removal, especially in the masseter area. When applying the depilatory paste, it is important not to use excessive paste. Wipe the skin gently when washing away the paste to avoid skin burns.</li>
	<li>Surgical field exposure: Cover with surgical drapes prior to the surgical procedure. For the MAS muscle, palpate first along the inferior edge of the mandible. At 5 mm from the edge, make a 7 mm incision along the line connecting the oral commissure to the tragus (the dashed line indicates the location of the incision in Figure 2B).</li>
	<li>For the TA muscle, palpate first along the calf to locate the anterior edge of the tibia bone. Cut 2 mm posterior to the edge through the skin with a scalpel, starting from the knee and ending at the ankle (the dashed line indicates the location of the incision in Figure 2H). For each mouse, if the MAS and the TA muscle on the left side are injured, use the other muscle on the contralateral side as the control. 
	NOTE: This cut is very superficial to ensure that the muscles will not be injured during the incision. The incision on the face should not be too near to the tragus. Otherwise, the parotid gland will be exposed, leading to excessive bleeding during the following freezing process.</li>
	<li>Dry-ice freezing: Hold one piece of dry ice with precooled forceps along the long axes of the MAS and TA muscles (Figure 2C and Figure 2I, respectively). Place the dry ice directly onto the surface of the muscle for 5 s. Confirm that the muscle appears stiff and pale white immediately after removing the dry ice (Figure 2D and Figure 2J) but that it returns to its normal color and texture within 22-25 s, similar to the adjacent uninjured muscle (Figure 2E and Figure 2K). 
	NOTE: The forceps holding the dry ice should be precooled to avoid quick sublimation of the dry ice. A timer is mandatory in conducting the 5 s freezing injury and counting the 22-25 s recovery time of the muscle. Any muscle with recovery time shorter than 22 s or longer than 25 s should be abandoned. This step requires practice.</li>
	<li>Wound closure: After total recovery of the muscle, close the skin wound with at least three 7-0 sutures (Figure 2F and Figure 2L).</li>
	<li>Resuscitation: When the suture is complete, place the mice in the cage with thermal support and monitor them continuously until all righting reflexes have been regained.</li>
</ol>
2. Muscle collection
<ol>
	<li>Euthanize the mice with an overdose of isoflurane, followed by cervical dislocation. Harvest the MAS and TA muscles on both sides for histological analysis.</li>
	<li>MAS muscle dissection: Cut open the skin on the mouse&#39;s face and remove the parotid gland to expose the posterior part of the MAS muscle. Dissect MAS from its anterior attachment all the way to its posterior attachment on the mandible. The white dotted area indicates the exposed part of the MAS (Figure 3A). 
	NOTE: The MAS muscle is composed of deep and superficial compartments, both closely attached to the surface of the mandible. Be sure to press the scissors to the mandible during the dissection to ensure complete isolation of MAS. Be careful to distinguish the zygomatico-mandibularis from MAS by its boundaries between the zygomatic and the mandible.</li>
	<li>TA muscle dissection: Cut open the skin from the ankle to the knee and remove the fascia to expose the TA muscle. Use the tip of the micro-dissection forceps to isolate the tendon of the TA and slide it all the way up to the knee to break off the fibers attaching to the tibia. Then, cut the tendon on the ankle (Figure 3B). 
	NOTE: For TA muscle, there are a few muscle tendons around the mouse ankle; it would take practice to discern the tendon of TA, which is the largest and closest to the tibia.</li>
	<li>Precooling isopentane: Prepare two insulation barrels; fill the small barrel with isopentane and the large barrel with liquid nitrogen. Place the small barrel inside the big barrel 10 min prior to muscle dissection (Figure 3D).</li>
	<li>Embedding mold preparation: Make a cylindrical tin foil mold for each sample and fill them with optimum cutting temperature (OCT) compound.</li>
	<li>Fresh frozen: Submerge the muscle in OCT, holding it perpendicular to the OCT compound (Figure 3C). Use a needle holder to transfer the mold into the prechilled isopentane. Wait for 40 s for the OCT to become white and solid (Figure 3D). 
	NOTE: Make sure the position of the mold is not too low. Otherwise, the isopentane will mix with the OCT and impair the muscle-freezing process. The level of OCT in the mold should be comparable to the height of the muscle sample. Avoid an excessive amount of OCT to ensure quick and efficient freezing.</li>
	<li>Sample storage: Store fresh frozen muscle samples in clearly labeled 24-well plates. Section the samples immediately or store them at -80 &#176;C for long-term use.</li>
</ol>
3. Histological analysis
<ol>
	<li>Use a cryostat to cut 10 &#181;m thick sections from the muscle sample onto surface-treated slides.</li>
	<li>Place the slides in ice-cold acetone for 20 min and air dry them in the fume hood for 20 min.</li>
	<li>Wash the slides for 3 x 5 min with phosphate-buffered saline (PBS). 
	NOTE: From this step onward, the slides were kept in a humidity chamber to avoid drying.</li>
	<li>Sirius Red staining
	<ol>
		<li>Prepare Sirius Red working solution by dissolving 0.5 g of Sirius Red powder in 500 mL of saturated picric acid.</li>
		<li>Stain the slides with Sirius Red working solution for 1 h at room temperature.</li>
		<li>Wash the slides for 2 x 5 min with 0.5% acetic acid.</li>
		<li>After the staining process, samples should be dehydrated first in 95% ethanol for 15 s, followed by dehydration in anhydrous ethanol for 1 min. The final step involves mounting the slide with neutral balsam.</li>
	</ol>
	</li>
	<li>Immunohistological staining
	<ol>
		<li>Permeabilize the sections with 0.3% Triton in PBS for 20 min at room temperature, followed by a PBS wash.</li>
		<li>For Pax7 staining, block the sections with the reagent provided with the referenced kit; for laminin and Pdgfr&#945; staining, block with 5% bovine serum albumin (BSA) and 5% donkey serum in PBS for 1 h at room temperature.</li>
		<li>Incubate the sections with primary antibodies against Pax7, laminin, and Pdgfr&#945; overnight at 4 &#176;C, followed by washing with PBS for 3 x 5 min.</li>
		<li>Incubate the sections with fluorescent-dye conjugated secondary antibodies for 1 h at room temperature, followed by washing with PBS for 3 x 5 min.</li>
		<li>Incubate with 4&#39;,6-diamidino-2-phenylindole (DAPI) for 3 min at room&#160;temperature, followed by washing with PBS for 3 x 5 min.</li>
		<li>Place mounting medium on the section, place the coverslip on top, and use nail polish to seal the coverslip.</li>
	</ol>
	</li>
</ol>

Establishing an Orofacial Muscle Fibrosis Model in Mice via a Freezing Injury

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J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Outcome+of+patients+with+complete+unilateral+cleft+lip+and+palate:+20-year+follow-up+of+a+treatment+protocol.">Outcome of patients with complete unilateral cleft lip and palate: 20-year follow-up of a treatment protocol.</a> Plastic and Reconstructive Surgery. 143 (2), 359e-367e (2019).</li><li>Parsaei, Y., Chandler, L., Smetona, J. T., Lopez, J., Steinbacher, D. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aesthetic+repair+of+unilateral+cleft+lip+using+the+modified+inferior+triangle+and+adjunctive+techniques.">Aesthetic repair of unilateral cleft lip using the modified inferior triangle and adjunctive techniques.</a> Plastic and Reconstructive Surgery. 149 (1), 70e-73e (2022).</li><li>Schubert, F. R., Singh, A. J., Afoyalan, O., Kioussi, C., Dietrich, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=To+roll+the+eyes+and+snap+a+bite+-+function,+development+and+evolution+of+craniofacial+muscles.">To roll the eyes and snap a bite - function, development and evolution of craniofacial muscles.</a> Seminars In Cell &amp; Developmental Biology. 91, 31-44 (2019).</li><li>Vyas, B., Nandkishore, N., Sambasivan, R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Vertebrate+cranial+mesoderm:+developmental+trajectory+and+evolutionary+origin.">Vertebrate cranial mesoderm: developmental trajectory and evolutionary origin.</a> Cellular and Molecular Life Sciences: CMLS. 77 (10), 1933-1945 (2020).</li><li>Sambasivan, R., Kuratani, S., Tajbakhsh, S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=An+eye+on+the+head:+the+development+and+evolution+of+craniofacial+muscles.">An eye on the head: the development and evolution of craniofacial muscles.</a> Development. 138 (12), 2401-2415 (2011).</li><li>Cheng, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Head+muscle+fibro-adipogenic+progenitors+account+for+the+tilted+regeneration+towards+fibrosis.">Head muscle fibro-adipogenic progenitors account for the tilted regeneration towards fibrosis.</a> Biochemical and Biophysical Research Communications. 589, 131-138 (2022).</li><li>Pavlath, G. 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Part C, Methods. 25 (2), 59-70 (2019).</li><li>Lukjanenko, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Aging+disrupts+muscle+stem+cell+function+by+impairing+matricellular+WISP1+secretion+from+fibro-adipogenic+progenitors.">Aging disrupts muscle stem cell function by impairing matricellular WISP1 secretion from fibro-adipogenic progenitors.</a> Cell Stem Cell. 24 (3), 433-446.e7 (2019).</li><li>Dong, J., Dong, Y., Chen, Z., Mitch, W. 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The authors have no conflicts of interest to disclose.

There are a variety of injury models for studying skeletal muscle regeneration, including the use of physical, chemical, and surgical stimuli10,11,12,13,14,15,16. Cardiotoxin and barium chloride are the two most widely used chemicals to initiate muscle regeneration10,11, but they result in transient fibrosis at ~7 dpi and complete regeneration after 3 weeks12,14. Local injection of glycerol is suitable for modeling muscle diseases with ectopic fat accumulation, such as Duchenne muscular dystrophy13. A surgical model of volumetric muscle loss could cause larger defects and thus lead to persistent fibrosis15,16. These models have been mainly tested in trunk and limb muscles, leaving orofacial muscle regeneration largely unexplored.
This study provides a rapid and reproducible method to study the characteristics of orofacial muscle regeneration. The whole freezing injury process of MAS and TA took only 20 min for each mouse. By carefully monitoring the freezing with a timer, the results could be made consistent in different batches of the experiment. The muscle harvest took 10 min for each mouse. The mice could tolerate simultaneous MAS and TA muscle freezing injury well. There was no impairment in locomotion. Malocclusion could be observed due to the improper application of pressure on the mandible, which could be overcome by pushing against the contralateral mandible.
In addition to staining with Sirius red and PDGFR&#945;, muscle fibrosis can be assessed by Masson&#39;s trichrome tissue staining and Col1 and &#945;-sma IHC. Masson trichrome staining is also used as a classic technique for staining collagen deposition. After staining, muscle fibers appear red, and collagen appears green or blue17. However, the steps of this method are relatively cumbersome. When tissue sections stained with Sirius Red are illuminated with a polarized light microscope, the fibrosis areas show significant birefringence. Moreover, different types of collagen exhibit different optical rotations and appear in different colors under the microscope18. In our preliminary research, we have included representative images8. Additionally, Col119 and &#945;-sma 17IHC staining have also been used to label fibrosis along with PDGFR&#945;, a membrane protein antibody that specifically labels FAPs.
There are several critical steps in this method. First, the freezing time (5 s) and recovery time (22-25 s) should be strictly monitored with a timer. A shorter freezing time will lead to complete regeneration in MAS, while a longer freezing time will be more likely to generate fibrosis in TA. More importantly, the degree of freezing could be carefully checked based on the recovery time. Muscles with a recovery time out of the 22-25 s range should not be incorporated into the study. Second, it is crucial to isolate the MAS muscle completely from the mandible. Muscle fibers along the inferior and posterior edges of the mandible fold back to the medial side. The parotid gland must be removed completely to fully expose the MAS and the scissors should be pressed to the surface of the mandible when dissecting. If poorly harvested, the cross-section of the MAS would appear incomplete, leaving out some regenerating myofibers or fibrotic areas. Third, the direction of muscle embedding should be perpendicular to the table to minimize the bias in later analysis of myofiber diameter. This is especially important for MAS because myofibers ran in different directions.
This protocol does have some limitations. First, this is a model of severe muscle injury because freezing will leave a &#34;dead zone&#34; in the injured muscle tissue17. All types of cells, including myofiber, MuSCs, FAPs, and endothelial cells, are damaged17. In contrast, chemical injuries, such as barium chloride injection, can preserve the MuSC viability18. Second, the age of the mice should be 4-5 weeks, thus limiting the interpretation and application of the data. Young mice at this age have a dominant content of neural-crest-derived FAPs in the MAS muscle19. Adult mice older than 8 weeks have a similar constitution of mesoderm-derived FAPs in MAS and TA muscles19, which may narrow their differences in muscle regeneration and fibrotic tissue formation.
The strength of this protocol is that it provides an easy and straightforward way to establish a model of orofacial muscle fibrosis. Subsequent morphological, histological, and molecular comparisons between the fibrotic MAS muscle and the fully regenerated TA muscle could be carried out to delineate the underlying mechanisms of regeneration. Following an injury to skeletal muscle, a range of cells participate in the process of regeneration. Recent research has identified FAPs as the primary cells responsible for creating the extracellular matrix in muscle tissue20. They also have the ability to prompt satellite cells to differentiate into muscle fibers21. These findings suggest that FAPs could be the key to understanding the limited myogenic capacity and excessive fibrosis present in orofacial muscles. While previous studies have focused on the impact of satellite cells on masseter muscle regeneration, our study highlights that FAP-mediated fibrogenesis is more pronounced during the regeneration of orofacial muscles. In contrast to TA FAPs, which rapidly decreased after an early peak, MAS FAPs continue to proliferate 14 days after injury8. Additionally, the PDGFR&#945;-positive area in MAS FAPs is significantly greater than that of TA. Our future research will delve deeper into the connection between orofacial muscle-oriented FAPs and fibrosis and develop more pertinent therapeutic approaches for patients with orofacial diseases.

Orofacial muscles are critical in daily physiological activities such as mastication, speech, respiration, and facial expression1. In congenital orofacial deformities, however, these muscles exhibit atrophic and fibrotic alterations, leading to impaired body health and social cognition2. Facial reconstructive surgery remains the first-line treatment, but up to 30-70% of postoperative patients still suffer from muscle loss and muscle dysfunction3,4 The failure of orofacial muscle regeneration has been attributed to intrinsic factors, which cannot be corrected by surgery alone. 
The emergence of orofacial muscles is an evolutionary novelty, accompanying the complex vertebrate head and chambered heart5,6. Unlike their somite-derived limb counterparts, orofacial muscles originate from the branchial arch7. These phylogenetic and ontogenetical characters may predispose them to distinct regenerative behaviors8. It has been reported that the masseter (MAS) muscle developed severe fibrosis at the time when the tibialis anterior (TA) muscle fully regenerated after exposure to the same extent of injury1,9. However, the underlying mechanism of regeneration remains poorly understood. 
In this study, a freezing injury model of the mice masseter muscle was established to facilitate the investigation into orofacial muscle regeneration. We chose 14 days after injury as the time point for assessing fibrosis phenotype as it was the earliest time point where discernible divergence was detectable between two muscles. Complete regeneration of the MAS after injury requires at least 40 weeks1. Consistently, this study revealed a remarkable deposition of collagen following freezing injury of MAS compared to the regular regeneration of the TA at 14 days post injury. With the help of this model, further mechanistic studies of muscle atrophy and fibrosis can be carried out, which will in turn help the development of potential therapeutic avenues to promote orofacial muscle regeneration after surgery.

<table><tbody><tr><td>1 mL syringe</td><td>Shifeng Medical Apparatus and Instrument (Chengdu, Sichuan, China)</td><td>1-ml syringe</td><td>/</td></tr><tr><td>Acetone</td><td>Chron Chemicals</td><td>Aceton</td><td>/</td></tr><tr><td>Adhesion microscope slides</td><td>Citotest Scientific</td><td>188105</td><td>/</td></tr><tr><td>Animal depilatory</td><td>Phygene Scientific</td><td>PH1877</td><td>/</td></tr><tr><td>BSA (bovine serum albumin)</td><td>Solarbio Life Sciences</td><td>A8010</td><td>/</td></tr><tr><td>DAPI</td><td>Solarbio Life Sciences</td><td>C0065</td><td>/</td></tr><tr><td>Donkey anti-goat Alexa Fluor 488</td><td>Abcam</td><td>ab150129</td><td>1:200</td></tr><tr><td>donkey serum</td><td>Solarbio Life Sciences</td><td>SL050</td><td>/</td></tr><tr><td>Dry Ice</td><td>Sinrro Technology (Chengdu, Sichuan, China)</td><td>rice-shaped dry ice</td><td>/</td></tr><tr><td>IFKine Red Donkey anti-rabbit</td><td>Abbkine Scientific Company</td><td>A24421</td><td>1:200</td></tr><tr><td>Insulation barrels (big)</td><td>Thermos</td><td>D600</td><td>/</td></tr><tr><td>Insulation barrels (small)</td><td>Polar Ware</td><td>250B</td><td>/</td></tr><tr><td>Isoflurane</td><td>RWD Life Technology Company (Shenzhen, Guangdong, China)</td><td>R510-22</td><td>/</td></tr><tr><td>Isopentane</td><td>MACKLIN</td><td>M813375</td><td>/</td></tr><tr><td>Laminin</td><td>Sigma-Aldrich</td><td>L9393</td><td>1:1000</td></tr><tr><td>Liquid nitrogen</td><td>Sinrro Technology (Chengdu, Sichuan, China)</td><td>/</td><td>/</td></tr><tr><td>M.O.M kit</td><td>Vector Laboratories</td><td>BMK-2202</td><td></td></tr><tr><td>Mice&amp;nbsp;</td><td>&amp;nbsp;Dashuo Biological Technology Company(Chengdu, Sichuan, China)</td><td>5 weeks old</td><td>/</td></tr><tr><td>mounting medium</td><td>Solarbio Life Sciences</td><td>S2100</td><td>/</td></tr><tr><td>Nertral balsam</td><td>Solarbio Life Sciences</td><td>G8590</td><td>/</td></tr><tr><td>Pax7</td><td>Developmental Studies Hybridoma Bank&amp;nbsp;</td><td>Pax7</td><td>1:5</td></tr><tr><td>Pdgfra</td><td>R&amp;amp;D systems</td><td>AF1062</td><td>1:40</td></tr><tr><td>Sirus Red Staining Kit</td><td>Solarbio Life Sciences</td><td>G1472</td><td>/</td></tr><tr><td>Surgical instruments (forceps, scissors, needle holder, scalpel, and suture)</td><td>Zhuoyue Medical Instrument (Suqian, Jiangsu, China)</td><td>/</td><td>/</td></tr><tr><td>Tissue-tek OCT</td><td>Sakura</td><td>4583</td><td>/</td></tr><tr><td>Triton</td><td>Shanghai Scigrace Biotech</td><td>ABIO-Biofroxx-0006A</td><td>/</td></tr><tr><td>Zoletil</td><td>Virbac</td><td>Zoletil 50</td><td>/</td></tr></tbody></table>

freezing injury in mouse masseter muscle to establish an orofacial muscle fibrosis model

Orofacial muscle constitutes a subset of skeletal muscle tissue, with a distinct evolutionary trajectory and development origin. Unlike the somite-derived limb muscles, the orofacial muscles originate from the branchial arches, with exclusive contributions from the cranial neural crest. A recent study has revealed that regeneration is also different in the orofacial muscle group. However, the underlying regulatory mechanism remains to be uncovered. Current skeletal muscle regeneration models mainly focus on the limb and trunk muscle. In this protocol, dry ice was used to induce freezing injury in the mouse masseter muscle and tibialis anterior muscle to create an orofacial muscle fibrosis model. The temporal dynamics of muscle satellite cells and fibro-adipogenic progenitors were different between the two muscles, leading to impaired myofiber regeneration and excessive extracellular matrix deposition. With the help of this model, a deeper investigation into muscle regeneration in the orofacial area could be carried out to develop therapeutic approaches for patients with orofacial diseases.

HE and Sirius Red staining (Figure 4 and Supplemental Figure S1) revealed complete muscle regeneration of TA in this freezing-injury model. In contrast, MAS exhibited impaired myofiber regeneration and excessive extracellular matrix deposition. The histology of intact MAS and TA muscle is shown in Figure 4A,B, where myofibers are in alignment and the fibrotic area only appeared in the interstitial space and among different bundles. While the TA muscle remained largely the same muscle structure as its intact control at 14 days post injury (Figure 4G-J), the structure of the MAS muscle was severely disturbed (Figure 4C-F). Fibrotic areas appeared in the place of nearly all the muscle fibers in the anterior, middle, and posterior cross-sections of the MAS muscle (Figure 4C-F).
Immunohistological staining of Pax7 (Supplemental Figure S2) and Pdgfr&#945; facilitated further investigation into the muscle satellite cells (MuSCs) and fibro-adipogenic progenitors (FAPs), respectively. In the MAS muscle, at 14 dpi, densely populated nuclei appeared but without proportionally increased MuSCs (Figure 5A,B). A large number of centrally nucleated myofibers (shown by the blue arrowhead) were detected in TA at 14 dpi (Figure 5H), while the contour of myofibers was barely noticeable in MAS in the injured area (Figure 5E,F). Instead, the infiltration of Pdgfr&#945;-positive FAPs and newly formed small-diameter myofibers (pointed by white arrows) were observed (Figure 5F).
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/65847/65847fig01.jpg" /> 
Figure 1: An overview of the surgical setup. (A) The mouse was anesthetized and fixed on the operation table. Hair was removed from the face and leg on the left side. A timer was used to monitor the muscle freezing and recovery time. (B) Dry ice, with a diameter of 3.5 mm and a height of 6-12 mm, was prepared in a glass beaker for freezing. The forceps were prechilled in dry ice for later use. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65847/65847fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/65847/65847fig02v2.jpg" /> 
Figure 2: The intraoperative record of MAS and TA freezing injury. (A) Hair removal for MAS freezing. (B) Open the skin to expose the MAS muscle. (C) Freezing with dry ice along the long axis of MAS. (D) The appearance of MAS immediately after 5 s of freezing. (E) The appearance of MAS after 22-25 s of recovery. (F) Facial wound closure. (G) Hair removal for TA freezing. (H) Open the skin to expose the TA muscle. (I) Freezing using dry ice along the long axis of TA. (J) The appearance of TA right after 5 s of freezing. (K) The appearance of TA after 22-25 s of recovery. (L) Leg wound closure. Abbreviations: MAS = masseter; TA = tibialis anterior.&#160;Dashed lines in Figures 2B and 2H&#160; indicate the location of the incisions.&#160;<a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65847/65847fig02largev2.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" class="xfigimg" src="/files/ftp_upload/65847/65847fig03.jpg" /> 
Figure 3: MAS and TA muscle harvest. (A) The appearance of MAS muscle at 14 days post freezing injury. The red dashed oval indicates MAS; the white dashed triangle shows the exposure of the MAS muscle. (B) The appearance of TA muscle at 14 days post freezing injury. The red dashed oval indicates TA. (C) Submerge the muscle sample in OCT compound in a perpendicular manner. (D) Transfer the muscle sample to nitrogen-cooled isopentane. Abbreviations: MAS = masseter; TA = tibialis anterior; OCT = optimum cutting temperature. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65847/65847fig03large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 4" class="xfigimg" src="/files/ftp_upload/65847/65847fig04.jpg" /> 
Figure 4: Histological analysis of MAS and TA muscles. Sirius red staining of intact (A-E) Intact muscle section of MAS and anterior, middle, and posterior sections of MAS at 14 dpi. (F-J) Intact muscle section of TA and anterior, middle, and posterior section of TA at 14 dpi. Scale bar = 100 &#181;m. Abbreviations: MAS = masseter; TA = tibialis anterior; dpi = days post injury. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65847/65847fig04large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 5" class="xfigimg" src="/files/ftp_upload/65847/65847fig05.jpg" /> 
Figure 5: Immunofluorescent analysis of MAS and TA muscles. Immunostaining of Pax7 and DAPI (blue) in (A) intact MAS, (B) 14 dpi MAS, (C) intact TA, and (D) 14 dpi TA. Immunostaining of laminin, Pax7, and DAPI (blue) in (E) intact MAS, (F) 14 dpi MAS, (G) intact TA, and (H) 14 dpi TA. Scale bar = 20 &#181;m. Abbreviations: MAS = masseter; TA = tibialis anterior; dpi = days post injury; DAPI = 4&#39;,6-diamidino-2-phenylindole. The white arrows indicate the representative small diameter fiber of the MAS. The blue dotted area and arrows indicate the representative regenerating myofibers and central nuclei of the TA.&#160;<a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65847/65847fig05large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
Supplemental Figure S1: Persistent fibrosis during muscle regeneration. (A) Schematic illustration of TA and MAS muscle regeneration induced by freezing injury. (B-G) Representative images of hematoxylin and eosin staining in TA and MAS muscle cross-sections at different time points post injury. Scale bars = 100 &#181;m. This figure is reproduced from Cheng et al.8. Abbreviations: MAS = masseter; TA = tibialis anterior; dpi = days post injury. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65847/65847_Figure S1.zip" target="_blank">Please click here to download this File.</a>
Supplemental Figure S2: Impairment of myogenesis process. (A) Immunofluorescence images of Pax7 (green) and DAPI (blue) in TA and MAS muscle. (B) Quantification of Pax7 positive/DAPI nuclei percentage. Scale bar = 100 &#181;m * Indicates significantly different from the intact muscle control. # Indicates significantly different from the other muscle at the same time point. **p &#60; 0.01, ##p &#60; 0.01. This figure is reproduced from Cheng et al.8. Abbreviations: MAS = masseter; TA = tibialis anterior; dpi = days post injury; DAPI = 4&#39;,6-diamidino-2-phenylindole. <a href="https://www-jove-com.bdigitaluss.remotexs.co/files/ftp_upload/65847/65847_Figure S2.zip" target="_blank">Please click here to download this File.</a>

Watch this Scientific Journal Video about Freezing Injury in Mouse Masseter Muscle to Establish an Orofacial Muscle Fibrosis Model at JoVE.com

Author Spotlight: Exploring Orofacial Muscle Regeneration &#8211; Insights and Innovations

The goal of this protocol is to establish an orofacial muscle fibrosis model. Comparison of the histology between mice masseter and tibialis anterior muscle after freezing injury confirmed masseter muscle fibrosis. This model will facilitate further investigation into the mechanism underlying orofacial muscle fibrosis.

Freezing Injury in Mouse Masseter Muscle to Establish an Orofacial Muscle Fibrosis Model

Orofacial muscle constitutes a subset of skeletal muscle tissue, with a distinct evolutionary trajectory and development origin. Unlike the somite-derived ...

freezing-injury-mouse-masseter-muscle-to-establish-an-orofacial

Universidad San Sebastian

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Biology

737 Views.  Sichuan University. The goal of this protocol is to establish an orofacial muscle fibrosis model. Comparison of the histology between mice masseter and tibialis anterior muscle after freezing injury confirmed masseter muscle fibrosis. This model will facilitate further investigation into the mechanism underlying orofacial muscle fibrosis.

Video: Author Spotlight: Exploring Orofacial Muscle Regeneration – Insights and Innovations

Tibial Nerve Transection - A Standardized Model for Denervation-induced Skeletal Muscle Atrophy in Mice

The tibial nerve transection model is a well-tolerated, validated, and reproducible model of denervation-induced skeletal muscle atrophy in rodents. Although originally developed and used extensively in the rat due to its larger size, the tibial nerve in mice is big enough that it can be easily manipulated with either crush or transection, leaving the peroneal and sural nerve branches of the sciatic nerve intact and thereby preserving their target muscles. Thus, this model offers the advantages of inducing less morbidity and impediment of ambulation than the sciatic nerve transection model and also allows investigators to study the physiologic, cellular and molecular biologic mechanisms regulating the process of muscle atrophy in genetically engineered mice. The tibial nerve supplies the gastrocnemius, soleus and plantaris muscles, so its transection permits the study of denervated skeletal muscle composed of fast twitch type II fibers and/or slow twitch type I fibers. Here we demonstrate the tibial nerve transection model in the C57Black6 mouse. We assess the atrophy of the gastrocnemius muscle, as a representative muscle, at 1, 2, and 4 weeks post-denervation by measuring muscle weights and fiber type specific cross-sectional area on paraffin-embedded histologic sections immunostained for fast twitch myosin.

The tibial nerve transection model is a well-tolerated, validated, and reproducible model of skeletal muscle atrophy. The model surgical protocol is described and demonstrated in C57Black6 mice. &#160;

The tibial nerve transection model is a well-tolerated, validated, and reproducible model of denervation-induced skeletal muscle atrophy in rodents. ...

Medicine

Modeling Stroke in Mice: Permanent Coagulation of the Distal Middle Cerebral Artery

Stroke is the third most common cause of death and a main cause of acquired adult disability in developed countries. Only very limited therapeutical options are available for a small proportion of stroke patients in the acute phase. Current research is intensively searching for novel therapeutic strategies and is increasingly focusing on the sub-acute and chronic phase after stroke because more patients might be eligible for therapeutic interventions in a prolonged time window. These delayed mechanisms include important pathophysiological pathways such as post-stroke inflammation, angiogenesis, neuronal plasticity and regeneration. In order to analyze these mechanisms and to subsequently evaluate novel drug targets, experimental stroke models with clinical relevance, low mortality and high reproducibility are sought after. Moreover, mice are the smallest mammals in which a focal stroke lesion can be induced and for which a broad spectrum of transgenic models are available. Therefore, we describe here the mouse model of transcranial, permanent coagulation of the middle cerebral artery via electrocoagulation distal of the lenticulostriatal arteries, the so-called &#8220;coagulation model&#8221;. The resulting infarct in this model is located mainly in the cortex; the relative infarct volume in relation to brain size corresponds to the majority of human strokes. Moreover, the model fulfills the above-mentioned criteria of reproducibility and low mortality. In this video we demonstrate the surgical methods of stroke induction in the &#8220;coagulation model&#8221; and report histological and functional analysis tools.

Various murine models of middle cerebral artery occlusion (MCAO) are widely used in experimental brain research. Here, we demonstrate the model of transcranial permanent distal MCAO which produces consistent cortical infarction of a size corresponding to damage imposed by the majority of human ischemic strokes.

Stroke is the third most common cause of death and a main cause of acquired adult disability in developed countries. Only very limited therapeutical ...

Facial Nerve Axotomy in Mice: A Model to Study Motoneuron Response to Injury

The goal of this surgical protocol is to expose the facial nerve, which innervates the facial musculature, at its exit from the stylomastoid foramen and either cut or crush it to induce peripheral nerve injury. Advantages of this surgery are its simplicity, high reproducibility, and the lack of effect on vital functions or mobility from the subsequent facial paralysis, thus resulting in a relatively mild surgical outcome compared to other nerve injury models. A major advantage of using a cranial nerve injury model is that the motoneurons reside in a relatively homogenous population in the facial motor nucleus in the pons, simplifying the study of the motoneuron cell bodies. Because of the symmetrical nature of facial nerve innervation and the lack of crosstalk between the facial motor nuclei, the operation can be performed unilaterally with the unaxotomized side serving as a paired internal control. A variety of analyses can be performed postoperatively to assess the physiologic response, details of which are beyond the scope of this article. For example, recovery of muscle function can serve as a behavioral marker for reinnervation, or the motoneurons can be quantified to measure cell survival. Additionally, the motoneurons can be accurately captured using laser microdissection for molecular analysis. Because the facial nerve axotomy is minimally invasive and well tolerated, it can be utilized on a wide variety of genetically modified mice. Also, this surgery model can be used to analyze the effectiveness of peripheral nerve injury treatments. Facial nerve injury provides a means for investigating not only motoneurons, but also the responses of the central and peripheral glial microenvironment, immune system, and target musculature. The facial nerve injury model is a widely accepted peripheral nerve injury model that serves as a powerful tool for studying nerve injury and regeneration.

We present a surgical protocol detailing how to perform a cut or crush axotomy on the facial nerve in the mouse. The facial nerve axotomy can be employed to study the physiological response to nerve injury and test therapeutic techniques.

The goal of this surgical protocol is to expose the facial nerve, which innervates the facial musculature, at its exit from the stylomastoid foramen and ...

Evaluating Biomaterials in Craniofacial VML Injury Using a Rat Model

Rodent Model of Masseter Volumetric Muscle Loss for Studying Bioengineering Materials

Craniofacial volumetric muscle loss (VML) injuries can occur as a result of severe trauma, surgical excision, inflammation, and congenital or other acquired conditions. Treatment of craniofacial VML involves surgical, functional muscle transfer. However, these procedures are unable to restore normal function, sensation, or expression, and more commonly, these conditions go untreated. Very little research has been conducted on skeletal muscle regeneration in animal models of craniofacial VML. This manuscript describes a rat model for the study of craniofacial VML injury and a protocol for the histological evaluation of biomaterials in the treatment of these injuries. Liquid hydrogel and freeze-dried scaffolds are applied at the time of surgical VML creation, and masseters are excised at terminal time points up to 12 weeks with high retention rates and negligible complications. Hematoxylin and eosin (HE), Masson's Trichrome, and immunohistochemical analysis are used to evaluate parameters of skeletal muscle regeneration as well as biocompatibility and immunomodulation. While we demonstrate the study of a hyaluronic-acid-based hydrogel, this model provides a means for evaluating subsequent iterations of materials in VML injuries.

Here, we describe a protocol for the surgical creation of a volumetric muscle loss (VML) injury in the rat masseter, providing a reproducible and accessible model for the study of craniofacial muscle injuries and their treatment using biomaterials such as the novel hydrogel.

Craniofacial volumetric muscle loss (VML) injuries can occur as a result of severe trauma, surgical excision, inflammation, and congenital or other ...

Bioengineering

Modeling Stroke in Mice: Focal Cortical Lesions by Photothrombosis

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, there remain limited therapeutic options for stroke patients. Therefore, more research is needed for pathophysiological pathways such as post-stroke inflammation, angiogenesis, neuronal plasticity, and regeneration. Given the inability of in vitro models to reproduce the complexity of the brain, experimental stroke models are essential for the analysis and subsequent evaluation of novel drug targets for these mechanisms. In addition, detailed standardized models for all procedures are urgently needed to overcome the so-called replication crisis. As an effort within the ImmunoStroke research consortium, a standardized photothrombotic mouse model using an intraperitoneal injection of Rose Bengal and the illumination of the intact skull with a 561 nm laser is described. This model allows the performance of stroke in mice with allocation to any cortical region of the brain without invasive surgery; thus, enabling the study of stroke in various areas of the brain. In this video, the surgical methods of stroke induction in the photothrombotic model along with&#160;histological analysis are demonstrated.

Described here is the photothrombotic stroke model, where a stroke is produced through the intact skull by inducing permanent microvascular occlusion using laser illumination after administration of a photosensitive dye.

Stroke is a leading cause of death and acquired adult disability in developed countries. Despite extensive investigation for novel therapeutic strategies, ...

Neuroscience

Force Assessment of Rat Masseter Muscle After VML Injury

In Vivo Functional Assessment of Rat Masseter Muscle Following Surgical Creation of a Volumetric Muscle Loss (VML) Injury

Volumetric Muscle Loss (VML) is prevalent in civilian and military populations and represents a debilitating skeletal muscle injury surpassing the body&#39;s natural regenerative capacity. These injuries disrupt not only muscle fibers but also nerves, blood vessels, and the extracellular matrix, overwhelming the regenerative capacity of skeletal muscle and leading to severe fibrosis and permanent deficiencies in muscle structure and function. Current clinical management has many limitations, and thus, research is ongoing to develop more effective therapeutic approaches. Notably, however, much of the preclinical emphasis on VML injuries has focused on limb and trunk muscles, with limited investigation into craniofacial muscles. Differences in developmental biology and regenerative capacity between craniofacial and limb/trunk muscles may provide crucial insights that drive more injury-specific VML treatment options. Moreover, evaluation of functional recovery is critical to establishing therapeutic efficacy. In this regard, in vivo testing of muscle contraction with percutaneous nerve stimulation is a minimally invasive method that allows for repeated functional assessment over the course of a study - in the same animal. In light of these considerations, this paper describes a method for the in vivo assessment of muscle function in the rat masseter muscle before and after a VML injury. This protocol is the first published instance to detail the creation and functional evaluation of a biologically relevant craniofacial VML injury in the rat.

Volumetric muscle loss (VML) injuries exceed endogenous regenerative ability, resulting in permanent functional deficits. Current VML research primarily focuses on limb and trunk muscles. To extend mechanistic studies of VML to craniofacial muscles, this article describes an in vivo method for longitudinal assessment of masseter muscle function pre- and post-VML injury.

Volumetric Muscle Loss (VML) is prevalent in civilian and military populations and represents a debilitating skeletal muscle injury surpassing the ...

A Model for Encephalomyosynangiosis Treatment after Middle Cerebral Artery Occlusion-Induced Stroke in Mice

There is no effective treatment available for most patients suffering with ischemic stroke, making development of novel therapeutics imperative. The brain's ability to self-heal after ischemic stroke is limited by inadequate blood supply in the impacted area. Encephalomyosynangiosis (EMS) is a neurosurgical procedure that achieves angiogenesis in patients with moyamoya disease. It involves craniotomy with placement of a vascular temporalis muscle graft on the ischemic brain surface. EMS has never been studied in the setting of acute ischemic stroke in mice. The hypothesis driving this study is that EMS enhances cerebral angiogenesis at the cortical surface surrounding the muscle graft. The protocol shown here describes the procedure and provides initial data supporting the feasibility and efficacy of the EMS approach. In this protocol, after 60 min of transient middle cerebral artery occlusion (MCAo), mice were randomized to either MCAo or MCAo + EMS treatment. The EMS was performed 3-4 h after occlusion. The mice were sacrificed 7 or 21 days after MCAo or MCAo + EMS treatment. Temporalis graft viability was measured using nicotinamide adenine dinucleotide reduced-tetrazolium reductase assay. A mouse angiogenesis array quantified angiogenic and neuromodulating protein expression. Immunohistochemistry was used to visualize graft bonding with brain cortex and change in vessel density. The preliminary data here suggest that grafted muscle remained viable 21 days after EMS. Immunostaining showed successful graft implantation and increase in vessel density near the muscle graft, indicating increased angiogenesis. Data show that EMS increases fibroblast growth factor (FGF) and decreases osteopontin levels after stroke. Additionally, EMS after stroke did not increase mortality suggesting that protocol is safe and reliable. This novel procedure is effective and well-tolerated and has the potential to provide information of novel interventions for enhanced angiogenesis after acute ischemic stroke.

The protocol aims to provide methods for encephalomyosynangiosis-grafting of a vascular temporalis muscle flap on the pial surface of ischemic brain tissue-for the treatment of non-moyamoya acute ischemic stroke. The approach's efficacy in increasing angiogenesis is evaluated using a transient middle cerebral artery occlusion model in mice.

There is no effective treatment available for most patients suffering with ischemic stroke, making development of novel therapeutics imperative. The ...

Electroporation and Nerve Injury in Mouse Levator Auris Longus Muscle

Combined In Vivo Electroporation and Short&#45;Term Reinnervation of the Cranial Levator Auris Longus Skeletal Muscle

The neuromuscular junction (NMJ) is the peripheral synapse controlling the contraction of skeletal muscle fibers to allow the coordinated movement of many organisms. At the NMJ, a presynaptic motor axon terminal contacts a muscle postsynaptic domain and is covered by terminal Schwann cells. The integrity and function of the NMJ is compromised under several conditions, including aging, neuromuscular diseases, and traumatic injuries. To analyze the potential contribution of muscle-derived proteins to NMJ maintenance and regeneration, an in vivo gene transfer strategy has been combined with the denervation of the cranial levator auris longus (LAL) muscle after mechanical nerve injury. Previous findings showed that the forced expression of control fluorescent proteins does not alter NMJ organization or neurotransmission. This procedure aims to describe a detailed method of in vivo electroporation of the LAL muscle followed by transection or crushing of the specific branch of the facial nerve innervating cranial muscles, leading to NMJ denervation and reinnervation, respectively. The combination of these experimental strategies in the convenient LAL muscle constitutes an efficient method to study the potential contribution of muscle protein overexpression or silencing in the context of short-term NMJ reinnervation.

Author Spotlight: Regenerative Roles of Muscle Proteins at Neuromuscular Junction Post-Nerve Injury

Here, we present a protocol that combines in vivo electroporation and denervation of the cranial levator auris longus (LAL) muscle. This procedure enables the study of the potential role of muscle-derived proteins in the regeneration of the neuromuscular synapse.

The neuromuscular junction (NMJ) is the peripheral synapse controlling the contraction of skeletal muscle fibers to allow the coordinated movement of many ...

The Establishment of a Murine Mandibular Molar Extraction Socket Healing Model

This study introduces the development of a molar extraction model in the murine mandible to provide a practicable model for studying alveolar bone regeneration and intramembranous ossification. C57/J6 mice were used to extract the mandibular first molar to establish this model. They were euthanized, and the bilateral mandibles harvested, at 1 week and 4 weeks post-surgery, respectively. Subsequent serial stereoscopic harvest, histological assessment, and immunofluorescence staining were performed to demonstrate successful surgery. Immediately after surgery, the stereoscopic images displayed an empty extraction socket. The hematoxylin and eosin (H&#38;E) at 1 week and Masson staining at 4 weeks post-surgery showed that the area of the original root was partially and fully filled with bone trabeculae, respectively. The immunofluorescence staining showed that, compared with the homeostasis side, the Sp7 expression increased at 1 week post-surgery, suggesting vigorous osteogenesis in the alveolar fossa. All these results demonstrated a practicable murine tooth extraction socket healing model. Upcoming studies revealing the mechanisms of jawbone defect healing or socket healing could adopt this method.

This protocol demonstrates step-by-step details of how to extract the mandibular first molar in the mouse. It provides an alternative method for researchers focusing on jawbone healing and regeneration.

This study introduces the development of a molar extraction model in the murine mandible to provide a practicable model for studying alveolar bone ...

Standardized Mandibular Fracture Protocol for Bone Repair in Mice

Less-Invasive Technique for Non-stabilized Mandibular Fracture in Mouse Models

Non-stabilized fractures can be made at mandibular sites in mice, thus making it possible to analyze bone repair using an endochondral ossification mode. To most accurately reflect this process in vivo, it is necessary to have a standardized protocol to avoid excessive bone loss and soft tissue damage, particularly at the mandibular site, an anatomical site characterized by minimal access. To our knowledge, we describe for the first time a less-invasive protocol of non-stabilized mandibular fracture in mice. Adult mice are anesthetized with isoflurane and receive a preoperative dose of buprenorphine by subcutaneous injection. A submandibular approach is performed, with a skin incision made along the inferior border of the mandible. The masseter muscle is elevated in a subperiosteal plane along the mandibular ramus with a periosteal elevator. A vertical and complete fracture of the ramus is performed from the basilar border to the coronoid notch (between condylar and coronoid processes), using a piezoelectric bone surgery device under saline serum irrigation at the basilar edge and scissors to complete the fracture of the mandibular ramus. The skin approach is closed by silk sutures. This detailed procedure for non-stabilized mandibular fractures offers an efficient procedure allowing minimal bone loss and soft tissue damage. This technique ensures successful and consistent results to accurately reflect the process of endochondral bone repair in mouse models.

This protocol presents a comprehensive and efficient method for analyzing mandibular bone repair. We describe a reproducible technique for non-stabilized mandibular fracture in a mouse model, allowing analysis of the process of endochondral bone repair with minimal tissue damage and bone loss.

Non-stabilized fractures can be made at mandibular sites in mice, thus making it possible to analyze bone repair using an endochondral ossification mode. ...