Rodent Model of Masseter Volumetric Muscle Loss for Studying Bioengineering Materials

Summary

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.

Abstract

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.

Introduction

Severe trauma, surgical excision, inflammation, and other acquired conditions can result in a degree of tissue loss that overwhelms the endogenous skeletal muscle repair mechanisms. Loss of resident cells and structures that promote the primary regenerative process can result in pathological remodeling and tissue fibrosis, resulting in long-term deficits of function and sensation and are referred to as volumetric muscle loss (VML)^1,2,3. The inflammatory response to VML injuries involves a well-documented and complex mechanism involving macrophages, cytokines, and myogenic cells that presents many theoretical targets in regenerative medicine⁴. While many in vitro studies have utilized these targets in animal models of extremity VML treatment, there is a lack of research on skeletal muscle regeneration in animal models of craniofacial VML^5,6,7.

Craniofacial tissue loss can result from conditions as described previously, and deficient craniofacial tissue can also occur in congenital conditions such as clefting, which in some cases involves a true volumetric deficiency of muscle tissue^8,9. Because muscles of the craniofacial region are important for function as well as aesthetic appearance, long-term effects of VML may have significant psychological affliction. Several aspects of craniofacial skeletal muscle are different from the somite-derived skeletal muscle found in extremities, including variations in gene expression, embryonic origin, satellite cell phenotype, satellite cell quantity, fiber composition, and architecture^10,11,12,13. These variations may result in VML injuries affecting craniofacial muscle differently than somite-derived muscle^14,15. To date, tissue engineering approaches shown to increase regeneration in animal models of extremity VML have not translated equivalently to craniofacial VML animal models¹⁶. This underscores the need to optimize in vivo approaches to animal craniofacial VML models.

While several in vivo studies of craniofacial VML have been conducted, the studies are small and the creation of a robust craniofacial muscle defect in animal models is challenging^8,13. Kim et al. reported the development of a mouse masseter VML model. However, this study only evaluated histology until 28 days following injury and had unclear power to detect differences in histologic outcomes between time points¹⁷. Rodriguez et al. reported the development of a sheep craniofacial VML model. However, they reported high variability within experimental groups, suggesting heterogeneity in the severity of the initial surgical injury¹⁶. Here, we report the protocol of our rat masseter VML model and demonstrate its utility in evaluating tissue-engineering approaches.

Protocol

This study was conducted in accordance with all applicable regulations, including adherence to the recommendations outlined in The Guide for the Care and Use of Laboratory Animals. The UCSF Institutional Animal Care and Use Program approved all animal procedures and postoperative care (IACUC protocol #AN195944-01).

1. VML surgery

1. Anesthetize male Sprague-Dawley rats at 12 weeks of age weighing 276-300 g in a sealed container using isofluorane general anesthesia at a rate of 1%-5% before being transferred to a sterile surgical field.
   1. Position the rodents on their left side such that the neck is extended in a neutral position and the nose fitted into the anesthetic nosecone, allowing exposure to the right side of the face.
   2. Taper isofluorane flow rate from induction to maintenance flow rate (approximately 2%).
2. Apply ophthalmic ointment to the eyes of the rodents.
3. Clear the operative site, bordered by a line between the right corner of the mouth, the base of the right ear superiorly, and the angle of the mandible inferiorly, of fur using depilatory cream.
   1. Disinfect the operative site several times in a circular motion using both ethanol and betadine scrub.
      NOTE: Sterile draping is not used in this protocol, given the size and location of access. The procedure is conducted using an aseptic technique in accordance with animal care guidelines.
4. Create a longitudinal incision 3-4 cm in length spanning from the inferior right whisker pad to the right ear.
   1. Elevate a skin flap, allowing visualization of the right masseter as well as the buccal and mandibular branches of the facial nerve (Figure 1A).
   2. Clear the skin from the underlying fascia using blunt separation and hold the skin edges in a retracted position using surgical clamps for optimal visualization of the underlying muscle.
5. Make a transverse incision 1-2 cm in length in the fascia overlying the anterior portion of the masseter. Use blunt separation to expand the space between the fascia and the masseter with care to maintain the overall integrity of the remaining fascia.
6. Using the fascial incision as a window to the superficial masseter, create a circular injury in the muscle measuring 5 mm across and 5 mm deep using a sterilized disposable punch biopsy. Ensure the defect is centered between the buccal and mandibular branches of the facial nerve, with care to avoid trauma to the nerves (Figure 1B).
7. Add the biomaterial to the circular masseter injury (Figure 1C).
8. Once the agent is suitably in place, suture the muscle-investing fascial window using a single 5-0 monofilament suture to avoid migration.
   NOTE: Solid-form agents are directly placed in the wound, and the tissue can be perturbed to promote saturation with blood, whereas liquid agents are added and allowed to gelate if necessary or immediately closed in place using fascia to avoid unwanted movement. In the experiment demonstrated here, an acrylated hyaluronic acid-based cryogel is administered.
9. Close the skin using either a simple interrupted or running subcuticular technique with 5-0 monofilament.
   1. Apply skin glue over the sutures to help prevent postoperative incision dehiscence.
10. Allow the rats to recover in a cage with an external heating source (heating pad beneath the cage or sterile hand warmers in the cage) and observe for 30 min postoperatively.
11. Dilute trimethoprim-sulfamethoxazole antibiotic (200 mg of trimethoprim and 40 mg of sulfamethoxazole in 5mL) in drinking water (5 mL/200 mL water) and administer it in rodents drinking water for 7 days postoperatively. Ensure perioperative analgesia is conducted according to institutional protocols.
    NOTE: In this study, 0.25% SC bupivacaine was given prior to surgical incision, 0.05 mg/kg SC buprenorphine was administered prior to recovery from anesthesia, and 2 mg/kg IP meloxicam was administered prior to recovery from anesthesia and again the following morning.

2. Masseter harvesting, freezing, and analysis

1. At predetermined time points, sacrifice the rats by inducing an overdose of a chemical agent (e.g., CO₂ or anesthetic) in the anesthetic induction chamber.
2. Confirm rodent sacrifice by conducting a bilateral thoracotomy using a single scalpel incision through 4^th rib ventrally to puncture the lungs (bilateral thoracotomy).
3. Reopen the surgical incision to allow visualization of the masseter, and remove the skin overlying the masseter for ease of access.
4. Dissect and remove the masseter from the mandible, beginning with separation at the mandibular angle. Ensure the dissection occurs along the bone surface to capture the entirety of the masseter thickness.
   1. Dissect anteriorly along the mandible, severing the attachment point of the tendon. Then, dissect along the zygomatic arch posteriorly (Figure 1D). At last, dissect the posterior attachment as there is an increased risk of bleeding.
5. Rinse the excised masseter in 1x PBS at room temperature (RT) and remove excessive moisture by thoroughly blotting the specimen with a paper towel.
6. Place the muscle in a cryomold and submerge it in optimal cutting temperature (OCT) embedding media, with the anterior end facing downwards and the posterior end facing upwards.
7. Add isopentane to a metal cup and cool it by submerging it in liquid nitrogen with care to prevent liquid nitrogen from entering it.
   1. Once the isopentane has reached optimal temperature (-140 to -149 °C) and thickened slightly, hold the cryomold containing OCT and the masseter partially submerged in the isopentane until the OCT has frozen through. Keep the frozen cryomold on dry ice until transferred to a freezer at -80 °C.
8. Cryosection frozen masseter samples with the orientation such that the anterior pole is sectioned first, moving posteriorly through the sample. Set the tissue block temperature to -20 °C and cryosection blade temperature to -15 °C. For every 200 µm, cut 4 slides (2 with 10 µm sections, 2 with 6 µm sections) before moving 200 µm further, cutting 4 additional slides and repeating throughout the sample.
9. Use the 10 µm sections for histology analysis (Masson's Trichrome, Hematoxylin and Eosin, Picrosirius Red, etc.) and the 6 µm sections for immunohistochemistry.
   NOTE: For histology, 6 µm sections may also be used.
10. Capture images using light or fluorescence microscopy.
11. Measure the cross-sectional areas of ~200 muscle fibers of masseter muscle using Image J software.

3. Immunohistochemical analysis

1. For immunohistochemical analysis of embryonic myosin heavy chain, fix slides in 4% PFA for 1 h at RT.
2. Wash the slides in PBS 3 times at RT, allowing each wash to sit for 10 min.
3. Permeabilize tissue by incubating the slides with 0.5% Triton X-100 at RT for 15 min.
4. Wash the slides in PBS 3 times at RT, allowing each wash to sit for 10 min.
5. Incubate with primary monoclonal antibody from host mice (1:200) in 2% normal goat serum (NGS) at 4 °C overnight.
6. Wash the slides in PBS-Tween 3 times at RT, allowing each wash to sit for 10 min.
7. Incubate with fluorophore-labeled anti-mouse secondary antibody from rabbit host (1:500) in 2% NGS at RT for 2 h.
8. Wash the slides in PBS-Tween 3 times at RT, allowing each wash to sit for 10 min.
9. Mount the slides using 3 drops of DAPI fluorescent mount before placing a coverslip.

Results

Outcomes for the evaluation of craniofacial VML and tissue regeneration using biomaterials include both quantitative and qualitative outcomes.

Figure 2 depicts an example of qualitative evaluation using the previously described model. The observation of de novo muscle fiber growth within our hydrogel is a qualitative positive outcome (Figure 2A) and suggests a biomaterial can provide sufficient architectural support and growt...

Discussion

There are several critical steps in the protocol where special attention is needed to achieve an optimal result. Step 1.4 describes the initial incision and blunt separation of the skin from the superficial masseter fascia. Blunt dissection should be done directly along the skin with scissors pointing away from the underlying muscle and fascia to prevent nicking and unintentionally creating a window through the fascia. The caudal aspect of the superficial masseter should be avoided to prevent unintentional injury to the ...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
F1.652 Myosin heavy chain (embryonic) monoclonal antibody	DSHB	F1.652
Goat anti-Mouse IgG2b Cross-Adsorbed Secondary Antibody, Alexa Fluor 647	Invitrogen	A-21242
Goat anti-Rabbit IgG (H+L) Highly Cross-Adsorbed Secondary Antibody, Alexa Fluor 488	Invitrogen	A-11034
Integra Standard Biopsy Punches, Disposable Standard biopsy punch; 5 mm, Diameter: 0.19 in., 0.5 cm	Integra	12460411
Mounting Medium with DAPI - Aqueous, Fluoroshield	Abcam	ab104139
Rabbit Anti-Mouse IgG H&L (Alexa Fluor 647) preadsorbed	Abcam	ab150127
Sulfamethoxazole/Trimethoprim Oral Suspension, Cherry Flavored, 473 mL	Med-Vet International	SKU: RXBAC-SUSP