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. K., 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=Heterogeneity+among+muscle+precursor+cells+in+adult+skeletal+muscles+with+differing+regenerative+capacities.">Heterogeneity among muscle precursor cells in adult skeletal muscles with differing regenerative capacities.</a> Developmental Dynamics. 212 (4), 495-508 (1998).</li><li>Camacho-Alonso, F., 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=Regeneration+of+lingual+musculature+in+rats+using+myoblasts+over+porcine+bladder+acellular+matrix.">Regeneration of lingual musculature in rats using myoblasts over porcine bladder acellular matrix.</a> Oral Diseases. 27 (6), 1580-1589 (2021).</li><li>LeBoff, M. S., 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=The+clinician's+guide+to+prevention+and+treatment+of+osteoporosis.">The clinician's guide to prevention and treatment of osteoporosis.</a> Osteoporosis International. 33 (10), 2049-2102 (2022).</li><li>Julien, A., 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=Direct+contribution+of+skeletal+muscle+mesenchymal+progenitors+to+bone+repair.">Direct contribution of skeletal muscle mesenchymal progenitors to bone repair.</a> Nature Communications. 12 (1), 2860 (2021).</li><li>Mahdy, M. A. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Glycerol-induced+injury+as+a+new+model+of+muscle+regeneration.">Glycerol-induced injury as a new model of muscle regeneration.</a> Cell and Tissue Research. 374 (2), 233-241 (2018).</li><li>Guardiola, O., 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=Induction+of+acute+skeletal+muscle+regeneration+by+cardiotoxin+injection.">Induction of acute skeletal muscle regeneration by cardiotoxin injection.</a> Journal of Visualized Experiments: JoVE. (119), 54515 (2017).</li><li>Larouche, J. A., Wallace, E. C., Spence, B. D., Buras, E., Aguilar, C. A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Spatiotemporal+mapping+of+immune+and+stem+cell+dysregulation+after+volumetric+muscle+loss.">Spatiotemporal mapping of immune and stem cell dysregulation after volumetric muscle loss.</a> JCI Insight. 8 (7), e162835 (2023).</li><li>Anderson, S. E., 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=Determination+of+a+critical+size+threshold+for+volumetric+muscle+loss+in+the+mouse+quadriceps.">Determination of a critical size threshold for volumetric muscle loss in the mouse quadriceps.</a> Tissue Engineering. 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. E., Zhang, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=The+pathway+to+muscle+fibrosis+depends+on+myostatin+stimulating+the+differentiation+of+fibro/adipogenic+progenitor+cells+in+chronic+kidney+disease.">The pathway to muscle fibrosis depends on myostatin stimulating the differentiation of fibro/adipogenic progenitor cells in chronic kidney disease.</a> Kidney International. 91 (1), 119-128 (2017).</li><li>Lemos, D. R., 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=Nilotinib+reduces+muscle+fibrosis+in+chronic+muscle+injury+by+promoting+TNF-mediated+apoptosis+of+fibro/adipogenic+progenitors.">Nilotinib reduces muscle fibrosis in chronic muscle injury by promoting TNF-mediated apoptosis of fibro/adipogenic progenitors.</a> Nature Medicine. 21 (7), 786-794 (2015).</li><li>Joe, A. W. B., 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=Muscle+injury+activates+resident+fibro/adipogenic+progenitors+that+facilitate+myogenesis.">Muscle injury activates resident fibro/adipogenic progenitors that facilitate myogenesis.</a> Nature Cell Biology. 12 (2), 153-163 (2010).</li><li>Biferali, B., Proietti, D., Mozzetta, C., Madaro, L. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Fibro-adipogenic+progenitors+cross-talk+in+skeletal+muscle:+the+social+network.">Fibro-adipogenic progenitors cross-talk in skeletal muscle: the social network.</a> Frontiers In Physiology. 10, 1074 (2019).</li></ol>

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<...

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 border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<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&#160;</td>
			<td>&#160;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&#160;</td>
			<td>Pax7</td>
			<td>1:5</td>
		</tr>
		<tr>
			<td>Pdgfra</td>
			<td>R&#38;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 amon...

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 ...

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Research

JoVE Journal

Biology

671 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

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Human cancer and response to therapy is better represented in orthotopic animal models. This paper describes the development of an orthotopic mouse model of ovarian cancer, treatment of cancer via oral delivery of drugs, and monitoring of tumor cell behavior in response to drug treatment in real time using in vivo imaging system. In this orthotopic model, ovarian tumor cells expressing luciferase are applied topically by injecting them directly into the mouse bursa where each ovary is enclosed. Upon injection of D-luciferin, a substrate of firefly luciferase, luciferase-expressing cells generate bioluminescence signals. This signal is detected by the in vivo imaging system and allows for a non-invasive means of monitoring tumor growth, distribution, and regression in individual animals. Drug administration via oral gavage allows for a maximum dosing volume of 10 mL/kg body weight to be delivered directly to the stomach and closely resembles delivery of drugs in clinical treatments. Therefore, techniques described here, development of an orthotopic mouse model of ovarian cancer, oral delivery of drugs, and in vivo imaging, are useful for better understanding of human ovarian cancer and treatment and will improve targeting this disease.

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Orthotopic animal models of ovarian cancer replicate better human disease and therefore enhance our understanding of cancer progression and tumor response to therapy. A mouse model receives an intrabursal injection of luciferase-expressing ovarian tumor cells. Treatment is administered via oral gavage. Tumor growth is monitored by in vivo imaging system.

Human cancer and response to therapy is better represented in orthotopic animal models.  This paper describes the development of an orthotopic mouse model ...

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to replenish depleted endoplasmic reticulum (ER) or sarcoplasmic reticulum (SR) Ca2+ stores1,2. Since Ca2+ is a ubiquitous second messenger, it is not surprising to see that SOCE plays important roles in a variety of cellular processes, including proliferation, apoptosis, gene transcription and motility. Due to its wide occurrence in nearly all cell types, including epithelial cells and skeletal muscles, this pathway has received great interest3,4. However, the heterogeneity of SOCE characteristics in different cell types and the physiological function are still not clear5-7.
The functional channel properties of SOCE can be revealed by patch-clamp studies, whereas a large body of knowledge about this pathway has been gained by fluorescence-based intracellular Ca2+ measurements because of its convenience and feasibility for high-throughput screening. The objective of this report is to summarize a few fluorescence-based methods to measure the activation of SOCE in monolayer cells, suspended cells and muscle fibers5,8-10. The most commonly used of these fluorescence methods is to directly monitor the dynamics of intracellular Ca2+ using the ratio of F340nm and F380nm (510 nm for emission wavelength) of the ratiometric Ca2+ indicator Fura-2. To isolate the activity of unidirectional SOCE from intracellular Ca2+ release and Ca2+ extrusion, a Mn2+ quenching assay is frequently used. Mn2+ is known to be able to permeate into cells via SOCE while it is impervious to the surface membrane extrusion processes or to ER uptake by Ca2+ pumps due to its very high affinity with Fura-2. As a result, the quenching of Fura-2 fluorescence induced by the entry of extracellular Mn2+ into the cells represents a measurement of activity of SOCE9. Ratiometric measurement and the Mn+2 quenching assays can be performed on a cuvette-based spectrofluorometer in a cell population mode or in a microscope-based system to visualize single cells. The advantage of single cell measurements is that individual cells subjected to gene manipulations can be selected using GFP or RFP reporters, allowing studies in genetically modified or mutated cells. The spatiotemporal characteristics of SOCE in structurally specialized skeletal muscle can be achieved in skinned muscle fibers by simultaneously monitoring the fluorescence of two low affinity Ca2+ indicators targeted to specific compartments of the muscle fiber, such as Fluo-5N in the SR and Rhod-5N in the transverse tubules9,11,12.

The extent of store-operated Ca2+ entry (SOCE) can be monitored using fluorescent Ca2+ indicators. Mn2+ quenching of such indicators assays SOCE in cultured cells and skeletal muscle fibers. A technique allowing spatial and temporal resolution of SOCE by confocal imaging of mechanically skinned muscle fibers is also described.

Store operated Ca2+ entry (SOCE), earlier termed capacitative Ca2+ entry, is a tightly regulated mechanism for influx of extracellular Ca2+ into cells to ...

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, fatigability, and recovery after fatigue can be obtained to assess specific aspects of the excitation-contraction coupling (ECC) process such as excitability, contractile machinery and Ca2+ handling ability. This method removes the nerve and blood supply and focuses on the isolated skeletal muscle itself. We routinely use this method to identify genetic components that alter the contractile property of skeletal muscle though modulating Ca2+ signaling pathways. Here, we describe a newly identified skeletal muscle phenotype, i.e., mechanic alternans, as an example of the various and rich information that can be obtained using the in vitro muscle contractility assay. Combination of this assay with single cell assays, genetic approaches and biochemistry assays can provide important insights into the mechanisms of ECC in skeletal muscle.

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

We describe a method to directly measure muscle force, muscle power, contractile kinetics and fatigability of isolated skeletal muscles in an in vitro system using field stimulation. Valuable information on Ca2+ handling properties and contractile machinery of the muscle can be obtained using different stimulating protocols.

 
Described here is a method to measure contractility of isolated skeletal muscles. Parameters such as muscle force, muscle power, contractile kinetics, ...

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections (bupivacaine, cardiotoxin or notexin), muscle transplantations (denervation-devascularization induced regeneration), intensive exercise, but also murine muscular dystrophy models such as the mdx mouse (for a review of these approaches see 1). In zebrafish, genetic approaches include mutants that exhibit muscular dystrophy phenotypes (such as runzel2 or sapje3) and antisense oligonucleotide morpholinos that block the expression of dystrophy-associated genes4. Besides, chemical approaches are also possible, e.g. with Galanthamine, a chemical compound inhibiting acetylcholinesterase, thereby resulting in hypercontraction, which eventually leads to muscular dystrophy5. However, genetic and pharmacological approaches generally affect all muscles within an individual, whereas the extent of physically inflicted injuries are more easily controlled spatially and temporally1. Localized physical injury allows the assessment of contralateral muscle as an internal control. Indeed, we recently used laser-mediated cell ablation to study skeletal muscle regeneration in the zebrafish embryo6, while another group recently reported the use of a two-photon laser (822 nm) to damage very locally the plasma membrane of individual embryonic zebrafish muscle cells7.
Here, we report a method for using the micropoint laser (Andor Technology) for skeletal muscle cell injury in the zebrafish embryo. The micropoint laser is a high energy laser which is suitable for targeted cell ablation at a wavelength of 435 nm. The laser is connected to a microscope (in our setup, an optical microscope from Zeiss) in such a way that the microscope can be used at the same time for focusing the laser light onto the sample and for visualizing the effects of the wounding (brightfield or fluorescence). The parameters for controlling laser pulses include wavelength, intensity, and number of pulses.
Due to its transparency and external embryonic development, the zebrafish embryo is highly amenable for both laser-induced injury and for studying the subsequent recovery. Between 1 and 2 days post-fertilization, somitic skeletal muscle cells progressively undergo maturation from anterior to posterior due to the progression of somitogenesis from the trunk to the tail8, 9. At these stages, embryos spontaneously twitch and initiate swimming. The zebrafish has recently been recognized as an important vertebrate model organism for the study of tissue regeneration, as many types of tissues (cardiac, neuronal, vascular etc.) can be regenerated after injury in the adult zebrafish10, 11.

The method presented here comprises the precise injury of live zebrafish embryos with high-energy laser pulses and the subsequent analysis of these injuries and their recovery with time. We also show how genetically labeled single or groups of skeletal muscle cells can be tracked during and after laser light induced damage.

Various experimental approaches have been used in mouse to induce muscle injury with the aim to study muscle regeneration, including myotoxin injections ...

Evaluation of Muscle Function of the Extensor Digitorum Longus Muscle Ex vivo and Tibialis Anterior Muscle In situ in Mice

Body movements are mainly provided by mechanical function of skeletal muscle. Skeletal muscle is composed of numerous bundles of myofibers that are sheathed by intramuscular connective tissues. Each myofiber contains many myofibrils that run longitudinally along the length of the myofiber. Myofibrils are the contractile apparatus of muscle and they are composed of repeated contractile units known as sarcomeres. A sarcomere unit contains actin and myosin filaments that are spaced by the Z discs and titin protein. Mechanical function of skeletal muscle is defined by the contractile and passive properties of muscle. The contractile properties are used to characterize the amount of force generated during muscle contraction, time of force generation and time of muscle relaxation. Any factor that affects muscle contraction (such as interaction between actin and myosin filaments, homeostasis of calcium, ATP/ADP ratio, etc.) influences the contractile properties. The passive properties refer to the elastic and viscous properties (stiffness and viscosity) of the muscle in the absence of contraction. These properties are determined by the extracellular and the intracellular structural components (such as titin) and connective tissues (mainly collagen) 1-2. The contractile and passive properties are two inseparable aspects of muscle function. For example, elbow flexion is accomplished by contraction of muscles in the anterior compartment of the upper arm and passive stretch of muscles in the posterior compartment of the upper arm. To truly understand muscle function, both contractile and passive properties should be studied.
The contractile and/or passive mechanical properties of muscle are often compromised in muscle diseases. A good example is Duchenne muscular dystrophy (DMD), a severe muscle wasting disease caused by dystrophin deficiency 3. Dystrophin is a cytoskeletal protein that stabilizes the muscle cell membrane (sarcolemma) during muscle contraction 4. In the absence of dystrophin, the sarcolemma is damaged by the shearing force generated during force transmission. This membrane tearing initiates a chain reaction which leads to muscle cell death and loss of contractile machinery. As a consequence, muscle force is reduced and dead myofibers are replaced by fibrotic tissues 5. This later change increases muscle stiffness 6. Accurate measurement of these changes provides important guide to evaluate disease progression and to determine therapeutic efficacy of novel gene/cell/pharmacological interventions. Here, we present two methods to evaluate both contractile and passive mechanical properties of the extensor digitorum longus (EDL) muscle and the contractile properties of the tibialis anterior (TA) muscle.

Evaluation of Muscle Function of the Extensor Digitorum Longus Muscle Ex vivo and Tibialis Anterior Muscle In situ in Mice

Changes in limb muscle contractile and passive mechanical properties are important biomarkers for muscle diseases. This manuscript describes physiological assays to measure these properties in the murine extensor digitorum longus and tibialis anterior muscles.

Body  movements are mainly provided by mechanical function of skeletal muscle. Skeletal  muscle is composed of numerous bundles of myofibers that are ...

Generation of Myospheres From hESCs by Epigenetic Reprogramming

Generation of a homogeneous and abundant population of skeletal muscle cells from human embryonic stem cells (hESCs) is a requirement for cell-based therapies and for a "disease in a dish" model of human neuromuscular diseases. Major hurdles, such as low abundance and heterogeneity of the population of interest, as well as a lack of protocols for the formation of three-dimensional contractile structures, have limited the applications of stem cells for neuromuscular disorders. We have designed a protocol that overcomes these limits by ectopic introduction of defined factors in hESCs - the muscle determination factor MyoD and SWI/SNF chromatin remodeling complex component BAF60C - that are able to reprogram hESCs into skeletal muscle cells. Here we describe the protocol established to generate hESC-derived myoblasts and promote their clustering into tridimensional miniaturized structures (myospheres) that functionally mimic miniaturized skeletal muscles7.

Here, we describe a protocol based on epigenetic reprogramming of human embryonic stem cells (hESCs) toward generating a homogeneous population of skeletal muscle progenitors that under permissive culture conditions form three-dimensional clusters of contractile myofibers (myospheres), which recapitulate biological features of human skeletal muscles.

Generation of a homogeneous and abundant population of skeletal muscle cells from human embryonic stem cells (hESCs) is a requirement for cell-based ...

Tissue Triage and Freezing for Models of Skeletal Muscle Disease

Skeletal muscle is a unique tissue because of its structure and function, which requires specific protocols for tissue collection to obtain optimal results from functional, cellular, molecular, and pathological evaluations. Due to the subtlety of some pathological abnormalities seen in congenital muscle disorders and the potential for fixation to interfere with the recognition of these features, pathological evaluation of frozen muscle is preferable to fixed muscle when evaluating skeletal muscle for congenital muscle disease. Additionally, the potential to produce severe freezing artifacts in muscle requires specific precautions when freezing skeletal muscle for histological examination that are not commonly used when freezing other tissues. This manuscript describes a protocol for rapid freezing of skeletal muscle using isopentane (2-methylbutane) cooled with liquid nitrogen to preserve optimal skeletal muscle morphology. This procedure is also effective for freezing tissue intended for genetic or protein expression studies. Furthermore, we have integrated our freezing protocol into a broader procedure that also describes preferred methods for the short term triage of tissue for (1) single fiber functional studies and (2) myoblast cell culture, with a focus on the minimum effort necessary to collect tissue and transport it to specialized research or reference labs to complete these studies. Overall, this manuscript provides an outline of how fresh tissue can be effectively distributed for a variety of phenotypic studies and thereby provides standard operating procedures (SOPs) for pathological studies related to congenital muscle disease.

The analysis of skeletal muscle tissues to determine structural, functional, and biochemical properties is greatly facilitated by appropriate preparation. This protocol describes appropriate methods to prepare skeletal muscle tissue for a broad range of phenotyping studies.

Skeletal muscle is a unique tissue because of its structure and function, which requires specific protocols for tissue collection to obtain optimal ...

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age are significant public health burdens. We currently understand little about the genetic regulation of muscle health with disease or age. The nematode C. elegans is an established model for understanding the genomic regulation of biological processes of interest. This worm&#8217;s body wall muscles display a large degree of homology with the muscles of higher metazoan species. Since C. elegans is a transparent organism, the localization of GFP to mitochondria and sarcomeres allows visualization of these structures in vivo. Similarly, feeding animals cationic dyes, which accumulate based on the existence of a mitochondrial membrane potential, allows the assessment of mitochondrial function in vivo. These methods, as well as assessment of muscle protein homeostasis, are combined with assessment of whole animal muscle function, in the form of movement assays, to allow correlation of sub-cellular defects with functional measures of muscle performance. Thus, C. elegans provides a powerful platform with which to assess the impact of mutations, gene knockdown, and/or chemical compounds upon muscle structure and function. Lastly, as GFP, cationic dyes, and movement assays are assessed non-invasively, prospective studies of muscle structure and function can be conducted across the whole life course and this at present cannot be easily investigated in vivo in any other organism.

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Skeletal muscle is essential for locomotion and is the bodies&#8217; main protein store. Muscle health measurements within C. elegans are described. Prospective changes to muscle structure and function are assessed using localized GFP and cationic dyes.

Muscle is a dynamic tissue that responds to changes in nutrition, exercise, and disease state. The loss of muscle mass and function with disease and age ...

Murine Aortic Crush Injury: An Efficient In Vivo Model of Smooth Muscle Cell Proliferation and Endothelial Function

Arterial reconstruction, whether angioplasty or bypass surgery, involves iatrogenic trauma causing endothelial disruption and vascular smooth muscle cell (VSMC) proliferation. Common murine models study small vessels such as the carotid and femoral arteries. Herein we describe an in vivo system in which both VSMC proliferation and endothelial barrier function can be simultaneously assessed in a large vessel. We studied the infrarenal aortic response to injury in C57BL/6 mice. The aorta was injured from the left renal vein to the aortic bifurcation by 30 transmural crushes of 5-seconds duration with a cotton-tipped applicator. Morphological changes were assessed with conventional histology. Aorta wall thickness was measured from the luminal surface to the adventitia. EdU integration and counter staining with DAPI and alpha-actin was used to demonstrate VSMC proliferation. Activation of ERK1/2, a known moderator of intimal hyperplasia formation, was determined by Western Blot analysis. The effect of inflammation was determined by immunohistochemistry for B-cells, T-cells, and macrophages. En face sections of endothelium were visualized with scanning electron microscopy (SEM). Endothelial barrier function was determined with Evans Blue staining. Transmural injury resulted in aortic wall thickening. This injury induced VSMC proliferation, most prominently at 3 days after injury, and early activation of ERK1/2 and decreased p27kip1 expression. Injury did not result in increased B-cells, T-cells, or macrophages infiltration in the vessel wall. Injury caused partial endothelial cell denudation and loss of cell-cell contact. Injury resulted in a significant loss of endothelial barrier function, which returned to baseline after seven days. The murine transmural blunt aortic injury model provides an efficient system to simultaneously study both VSMC proliferation and endothelial barrier function in a large vessel.

Murine Aortic Crush Injury: An Efficient In Vivo Model of Smooth Muscle Cell Proliferation and Endothelial Function

Restenosis following cardiovascular procedures (bypass surgery, angioplasty, or stenting) is a significant problem reducing the durability of these procedures. An ideal therapy would inhibit smooth muscle cell (VSMC) proliferation while promoting regeneration of the endothelium. We describe a model for simultaneous assessment of VSMC proliferation and endothelial function in vivo.

Arterial reconstruction, whether angioplasty or bypass surgery, involves iatrogenic trauma causing endothelial disruption and vascular smooth muscle cell ...

A Multi-hole Cryovial Eliminates Freezing Artifacts when Muscle Tissues are Directly Immersed in Liquid Nitrogen

Studies on skeletal muscle physiology face the technical challenge of appropriately processing the specimens to obtain sections with clearly visible cytoplasmic compartments. Another hurdle is the tight apposition of myofibers to the surrounding tissues. Because the process of tissue fixation and paraffin embedding leads to the shrinkage of muscle fibers, freezing is an optimal means of hardening muscle tissue for sectioning. However, a commonly encountered issue, the formation of ice crystals, occurs during the preparation of frozen sections because of the high water content of muscle. The protocol presented here first describes a simple and efficient method for properly freezing muscle tissues by immersing them in liquid nitrogen. The problem with using liquid nitrogen alone is that it causes the formation of a nitrogen gas barrier next to the tissue, which acts as an insulator and inhibits the cooling of the tissues. To avoid this "vapor blanket" effect, a new cryovial was designed to increase the speed of liquid flow around the tissue surface. This was achieved by punching a total of 14 inlet holes in the wall of the vial. According to bubble dynamics, a higher rate of liquid flow results in smaller bubbles and fewer chances to form a gas barrier. When liquid nitrogen flows into the cryovial through the inlet holes, the flow velocity around the tissue is fast enough to eliminate the gas barrier. Compared to the method of freezing muscle tissues using pre-chilled isopentane, this protocol is simpler and more efficient and can be used to freeze muscle in a throughput manner. Furthermore, this method is optimal for institutions that do not have access to isopentane, which is extremely flammable at room temperature.

This protocol describes a procedure to freeze muscle tissues by plunging them directly into liquid nitrogen. This protocol also highlights a new cryovial that can avoid the &#34;blanket effect&#34; of nitrogen gas when liquid nitrogen contacts the tissue surface of a specimen.

Studies on skeletal muscle physiology face the technical challenge of appropriately processing the specimens to obtain sections with clearly visible ...

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

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Chapters in this video

In vivo Imaging and Therapeutic Treatments in an Orthotopic Mouse Model of Ovarian Cancer

Fluorescence-based Measurement of Store-operated Calcium Entry in Live Cells: from Cultured Cancer Cell to Skeletal Muscle Fiber

Ex Vivo Assessment of Contractility, Fatigability and Alternans in Isolated Skeletal Muscles

Laser-inflicted Injury of Zebrafish Embryonic Skeletal Muscle

Evaluation of Muscle Function of the Extensor Digitorum Longus Muscle Ex vivo and Tibialis Anterior Muscle In situ in Mice

Generation of Myospheres From hESCs by Epigenetic Reprogramming

Tissue Triage and Freezing for Models of Skeletal Muscle Disease

Methods to Assess Subcellular Compartments of Muscle in C. elegans

Murine Aortic Crush Injury: An Efficient In Vivo Model of Smooth Muscle Cell Proliferation and Endothelial Function

A Multi-hole Cryovial Eliminates Freezing Artifacts when Muscle Tissues are Directly Immersed in Liquid Nitrogen