The Establishment of Calvarial Suture-Bony Composite Defects in Rats: A Standardized Model for Suture-Regenerative Therapy Investigation

Summary

We describe the detailed surgical procedures of calvarial suture-bony composite defects in rats, alongside the investigations into the short-term and long-term prognoses of the model. We aim to construct a standardized model for developing suture-regenerative therapies.

Abstract

Large-scale calvarial defects often coincide with cranial suture disruption, leading to impairments in calvarial defect restoration and skull development (the latter occurs in the developing cranium). However, the lack of a standardized model hinders progress in investigating suture-regenerative therapies and poses challenges for conducting comparative analyses across distinct studies. To address this issue, the current protocol describes the detailed modeling process of calvarial suture-bony composite defects in rats.

The model was generated by drilling full-thickness rectangular holes measuring 4.5 mm × 2 mm across the coronal sutures. The rats were euthanized, and the cranium samples were harvested postoperatively at day 0, week 2, week 6, and week 12. µCT results from samples collected immediately post-surgery confirmed the successful establishment of the suture-bony composite defect, involving the removal of the coronal suture and the adjacent bone tissues.

Data from the 6^th and 12^th postoperative weeks demonstrated a natural healing tendency for the defect to close. Histological staining further validated this trend by showing increased mineralized fibers and new bone at the defect center. These findings indicate progressive suture fusion over time following calvarial defects, underscoring the significance of therapeutic interventions for suture regeneration. We anticipate that this protocol will facilitate the development of suture-regenerative therapies, offering fresh insights into the functional restoration of calvarial defects and reducing adverse outcomes associated with suture loss.

Introduction

Cranial sutures are dense fibrous connections between cranial bones, acting as joints to facilitate slight skull movement and providing a protective cushion for the brain under pressure¹. In recent years, increased research has highlighted the pivotal role of cranial sutures in skull development, craniofacial homeostasis, and inherent osteo-reparative potential^{2,3,4,5,6,7,8}. During periods of growth and development, cranial sutures act as the main growth centers in the skull⁴. New bone formation occurs at the osteogenic fronts on both sides of the sutures, while the cells within the sutures maintain an undifferentiated mesenchymal state, ensuring balanced skull expansion alongside brain growth¹. The loss of cranial sutures at this time results in a discrepancy between the growth of the skull and brain, leading to severe issues like brain injuries, hydrocephalus, increased intracranial pressure, and cognitive dysfunction^3,9.

Besides, cranial sutures play a crucial role in determining the prognosis of calvarial defects^5,7,10. The regenerative potential across the calvarial surface is unevenly distributed, with cranial sutures showing remarkably superior capabilities compared to non-suture regions^10,11. One study indicates that the speed of calvarial defect healing inversely correlates with the distance between the cranial suture and the injury site¹⁰. Specifically, the removal of coronal and sagittal sutures leads to the non-healing of parietal bone defects⁷, emphasizing the necessity of suture regeneration in calvarial defects. However, the focus of the current studies is predominantly on the restoration of cranial osseous structure while neglecting the regeneration of suture mesenchyme.

Regarding advancements in suture regeneration, promising outcomes have been observed with the transplantation of suture-containing bone flaps, mesenchymal stem cells (MSCs), and artificial biomaterials. When bone flaps with sutures were transplanted into calvarial defects, they successfully integrated and healed, in contrast to those without sutures displaying non-union and an inability to form periosteum, dura mater, or osteocytes⁵. Likewise, the implantation of MSCs derived from bone marrow into sagittal suture-bony composite defects facilitated the formation of suture-like gaps¹². Of note, a recent study highlighted the realization of suture regeneration with Gli1⁺ MSCs, allowing for intracranial pressure control, skull deformity correction, and enhanced neurocognitive function¹³. As regenerative medicine and biomedical engineering develop, researchers increasingly focus on tissue engineering biomaterials due to their adaptable and customizable characteristics¹⁴. Notably, polytetrafluoroethylene membranes have been proven to be effective in reconstructing cranial bone and suture mesenchyme simultaneously^15,16.

However, craniofacial research lacks established models for exploring suture mesenchymal regenerative therapies, unlike relatively mature models in repairing other tissues such as bone, skin, cartilage, and muscles¹⁷. The absence of a standardized model constrains the study of suture-regenerative therapies and makes it challenging to perform comparative analysis across different studies. Therefore, our study established a practicable and reproducible rat calvarial suture-bony composite defect. Through this method, we aim to develop appropriate clinical interventions for cranial suture reconstruction, offering novel perspectives on the functional repairment of calvarial defects and decreasing unfavorable outcomes resulting from suture loss.

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-2021-597). A total of 12 (3 rats at each of the four time points) Sprague-Dawley (SD) rats (male, 300 g, 8 weeks old) were obtained from a commercial source (see Table of Materials).

1. Presurgical preparation

1. Surgical items
   1. Prepare surgical instruments displayed in Figure 1A, including curved forceps, a disposable sterile scalpel, a periosteal elevator, an irrigation needle, cotton balls, a low-speed handpiece, a surgical motor, dental low-speed round burs (Figure 1B, 1.2 mm and 0.8 mm diameter, respectively), a needle holder, 3-0 monofilament sutures, and a straight scissor.
2. Sterilization and disinfection
   1. Sterilize the instruments by steam sterilization (125-135 °C, 20-25 min) in advance.
   2. Use ethanol to sterilize heat-sensitive medical equipment, such as electric shavers.
   3. Use sterile medical non-woven fabrics to cover the operation platform and disinfect the surrounding environment with 75% ethanol.
3. Anesthesia
   1. Prepare animals for surgery with a 1-week acclimation period.
   2. Inject the rats intraperitoneally with xylazine (10 mg/kg) and ketamine (100 mg/kg) 20 min before surgery for anesthesia or utilize any suitable anesthetic protocol to achieve general anesthesia.
      NOTE: The ketamine-xylazine anesthetic cocktail administered intraperitoneally in rats typically takes effect within 10-15 min, reaching peak anesthesia around 35-40 min after injection. To prevent hypothermia during anesthesia, rats are best placed on a heating pad.
   3. Administer a preoperative subcutaneous injection of carprofen at 5 mg/kg to provide analgesia.
   4. Use the "toe pinch method" to determine whether or not the rats are conscious and responsive.

2. Surgical process

1. Site preparation
   1. Place the rat in a prone position with its head naturally extending vertically, without requiring special devices or restraints to maintain this posture (Figure 2A).
   2. Apply veterinary ointment, i.e., Vaseline, to the rat's eyes to prevent corneal dryness.
   3. Remove the hair from the scalp between the nasal bridge and the cervical spine joint with an electric shaver (Figure 2B). Disinfect the surgical area in circular motions radiating from the center with 2% iodophor solution followed by 75% ethanol. Employ a surgical drape to cover the dorsum of the animal and the fur surrounding the incision.
2. Surgical site opening
   1. Starting from the midpoint of the nasal bone, make a 2 cm longitudinal skin incision with a disposable scalpel following the midline of the cranium (Figure 2C).
   2. Make a midline periosteal incision mirroring the initial point and extent of the skin layer with a scalpel (Figure 2D). Then, gently lift the periosteum on both sides of the incision with a periosteal elevator (Figure 2E1) to expose parietal bones, frontal bones, and coronal sutures (Figure 2E2).
      NOTE: When incising the periosteum, take care not to harm cranial sutures to prevent excessive bleeding when cutting. Adequate exposure of the coronal suture is of utmost importance for the following procedures.
   3. Rinse the wound with saline solution and dry the surgical area with cotton balls.
3. Suture-bony composite defect model establishment
   1. Set the surgical motor to 35,000 rpm by rotating the knob (Figure 1A, yellow arrow), then turn on the switch (Figure 1A, white arrow).
   2. Starting from any point on the coronal suture, apply vertical force using a 1.2 mm diameter round bur until a sense of breakthrough is felt.
      NOTE: Recommend the midpoint of the coronal suture (indicated by yellow arrows in Figure 2E2) as the starting point for penetration. When grinding, dental drills should be kept perpendicular to the cranial surface. Exercise caution not to continue drilling after penetrating the full thickness of the skull to prevent any further harm to the rats, including brain damage or cerebral hemorrhage.
   3. From the penetration point, move the bur laterally along the coronal suture to create an approximately 4 mm long positioning groove (Figure 2F1). Remove bone tissue with the bur on both sides of the positioning groove to initially form a rectangular full-thickness defect (Figure 2F2).
   4. Employ a 0.8 mm diameter round bur to refine details (Figure 2G1), involving the grinding of right angles and the smoothing of defect margins, ultimately achieving a standard rectangular defect measuring 2 mm in width and 4.5 mm in length (Figure 2G2).
      NOTE: To achieve complete removal of the coronal suture while preserving the sagittal and frontal sutures in 300 g SD rats, the maximum feasible defect length is approximately 4.5 mm. Considering the width of the coronal suture in the anterior-posterior direction (Supplementary Figure S1), the defect width was set as 2 mm.
   5. Create two defects across the left and right halves of the coronal suture for self-comparison.
   6. Maintain continuous irrigation of saline solution during the drilling procedure to safeguard against thermal injury to the cranium and brain. Meanwhile, use cotton balls to dry the operation area.
4. Sample dimension verification
   1. Regularly verify the length and width of the defects using a vernier caliper (Figure 2H1, H2) to ensure consistency across all samples.
5. Surgical site closure
   1. Close the skin with 3-0 monofilament sutures (Figure 2I).
6. Immediate postoperative in vivo micro-computed tomography (µCT)
   1. If feasible, conduct in vivo µCT scans on all rats immediately after surgery to confirm the success of the surgical procedure and monitor defect recovery trends for each individual.

3. Postsurgical care

1. According to animal care protocols, administer established analgesics as necessary after surgery, for instance, carprofen (5 mg/kg, subcutaneous use).
2. Transfer the rats to a constant heating pad (37 °C) for postoperative recovery.
3. Once fully conscious, relocate the rats to their housing cage containing clean bedding.
   NOTE: Continuously monitor the rats post-surgery. Do not leave them unattended until they can maintain sternal recumbency. Keep operated rats isolated from others until fully recovered.
4. Conduct analgesia management and postoperative monitoring for 24 h, followed by daily assessments throughout the first week post surgery. Monitor the rats at least 1-2x per week thereafter.

4. Sample collection and data analysis

1. Sample preparation
   1. Collect cranium specimens at postoperative day 0, week 2, week 6, and week 12. Euthanize the rats using CO₂ inhalation.
   2. Fix the samples in 4% paraformaldehyde at 4 °C for 24 h before further analysis.
2. µCT evaluation
   1. Perform µCT scans on cranial bones of postoperative day 0, week 6, week 12 with the following scan parameters: X-ray tube potential, 70 kVp; X-ray intensity, 0.2 mA; filter, AL 0.5 mm; integration time, 1 x 300 ms; and voxel size, 10 µm.
   2. Obtain 3D reconstruction and cross-sectional images with image processing software (see Table of Materials).
   3. Measure residual defect volume and perform statistical analysis with corresponding software (see Table of Materials).
3. Histological staining
   1. Decalcify the cranial bones in 12% (w/v) ethylene diamine tetraacetic acid solution (pH = 7.2) at 4 °C for 6 weeks.
      NOTE: Decalcify the samples with solution changes every 3 days. Utilize a shaker to expedite the process. Completion is indicated when a 25 G needle easily penetrates the sample.
   2. Proceed with dehydration, paraffin embedding, and preparation of 6 µm sections using standard protocols¹⁸.
   3. Conduct histological analysis using hematoxylin and eosin (H&E) and Masson's trichrome staining following kit protocol¹⁸.

Results

In this study, the rat calvarial suture-bony composite defect was established by drilling a 4.5 mm x 2 mm rectangular hole across the coronal suture. The surgical schematic illustration and the research flow chart are depicted in Figure 3. The 3D image and the cross-sectional view of postoperative 0-day samples, namely samples collected immediately after surgery, confirmed the successful creation of a full-thickness calvarial defect, involving the complete removal of the coronal suture as we...

Discussion

Conventional calvarial defect models, whether involving cranial sutures or not, primarily concentrate on the repair of hard tissue, often neglecting the vital regeneration of suture mesenchyme^19,20. In suture regeneration research, prior models, like those by Mardas et al.^15,16, utilizing a trephine bur to create a 5 mm circular defect across the sagittal suture of rats, resulted in substantial hard tissu...

Materials

Name	Company	Catalog Number	Comments
4% paraformaldehyde	Biosharp	BL539A
2% Iodophor solution	Chengdu Jinshan Chemical Reagent Co., Ltd.	None
75% Ethanol	Chengdu Jinshan Chemical Reagent Co., Ltd.	None
Cotton balls	Haishi Hainuo Group Co., Ltd.	None
Cotton swabs	Lakong Medical Devices Co.,	None
Curved forceps	Chengdu Shifeng Co., Ltd.	None
Dataviewer and Ctan software for residual defect volume assessments	Bruker	None
Dental low-speed round burs	Dreybird Medical Equipment Co., Ltd.	RA3-012 RA1-008
Disposable sterile scalpel	Hangzhou Huawei Medical Supplies Co., Ltd.	None
Disposable syringes (22 G)	Chengdu Shifeng Co., Ltd.	SB1-089(IX)
Electric shaver	JASE	BM320210
Ethylene Diamine Tetraacetic Acid (EDTA)	BioFroxx	1340GR500
Hematoxylin and Eosin Stain Kit	Biosharp	BL700B
Irrigation needle (23 G)	Sichuan New Century Medical Polymer Products Co., Ltd.	None
Low-speed handpiece	Guangzhou Dental Guard Technology Co., Ltd.	None
Masson’s Trichrome Stain Kit	Solarbio	G1340
Medical non-woven fabrics	Henan Yadu Industrial Co., Ltd.	None
Micro-computed tomography (µCT)	Scanco Medical AG	µCT45
Mimics 20.0 for cross-sectional images	Materialise	None
Needle holders	Chengdu Shifeng Co., Ltd.	None
Periosteal elevator	Chengdu Shifeng Co., Ltd.	None
Saline solution	Sichuan Kelun Pharmaceutical Co., Ltd.	None
Scanco medical visualizer software for 3D image reconstruction	Scanco Medical AG	None
SPSS Statistics 20.0 for statistical analysis	IBM	None
Sprague-Dawley rats	Byrness Weil Biotech Ltd	None
Straight Scissors	Chengdu Shifeng Co., Ltd.	None
Surgical Motor	MARATHON	N3-140232
Surgical sutures (3-0 monofilament)	Hangzhou Huawei Medical Supplies Co., Ltd.	None