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

Summary

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.

Abstract

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.

Introduction

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 dysfunction^3,4 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 heart^5,6. 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 injury^1,9. 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.

Protocol

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 ± 2%) and temperature-controlled (23 ± 2 °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

1. 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 °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.
2. 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). The surgical sites were disinfected by three rounds of an iodine-based scrub and 75% ethanol.  
   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.
3. 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).
4. 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.
5. 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.
6. Wound closure: After total recovery of the muscle, close the skin wound with at least three 7-0 sutures (Figure 2F and Figure 2L).
7. 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.

2. Muscle collection

1. Euthanize the mice with an overdose of isoflurane, followed by cervical dislocation. Harvest the MAS and TA muscles on both sides for histological analysis.
2. MAS muscle dissection: Cut open the skin on the mouse'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.
3. 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.
4. 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).
5. Embedding mold preparation: Make a cylindrical tin foil mold for each sample and fill them with optimum cutting temperature (OCT) compound.
6. 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.
7. Sample storage: Store fresh frozen muscle samples in clearly labeled 24-well plates. Section the samples immediately or store them at -80 °C for long-term use.

3. Histological analysis

1. Use a cryostat to cut 10 µm thick sections from the muscle sample onto surface-treated slides.
2. Place the slides in ice-cold acetone for 20 min and air dry them in the fume hood for 20 min.
3. 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.
4. Sirius Red staining
   1. Prepare Sirius Red working solution by dissolving 0.5 g of Sirius Red powder in 500 mL of saturated picric acid.
   2. Stain the slides with Sirius Red working solution for 1 h at room temperature.
   3. Wash the slides for 2 x 5 min with 0.5% acetic acid.
   4. 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.
5. Immunohistological staining
   1. Permeabilize the sections with 0.3% Triton in PBS for 20 min at room temperature, followed by a PBS wash.
   2. For Pax7 staining, block the sections with the reagent provided with the referenced kit; for laminin and Pdgfrα staining, block with 5% bovine serum albumin (BSA) and 5% donkey serum in PBS for 1 h at room temperature.
   3. Incubate the sections with primary antibodies against Pax7, laminin, and Pdgfrα overnight at 4 °C, followed by washing with PBS for 3 x 5 min.
   4. 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.
   5. Incubate with 4',6-diamidino-2-phenylindole (DAPI) for 3 min at room temperature, followed by washing with PBS for 3 x 5 min.
   6. Place mounting medium on the section, place the coverslip on top, and use nail polish to seal the coverslip.

Results

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...

Discussion

There are a variety of injury models for studying skeletal muscle regeneration, including the use of physical, chemical, and surgical stimuli^{10,11,12,13,14,15,16}. Cardiotoxin and barium chloride are the two most widely used chemicals to initiate muscle regeneration¹⁰<...

Materials

List of materials used in this article

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