Bone loss or the absence of bone formation is a persistent clinical issue affecting both humans and other species. Various etiologies of bone loss or impaired formation occur in clinical contexts such as severe trauma, tumor resection, congenital disorders, infections, and systemic disease (Hardy and Cooper, 2009; Roddy et al., 2018; Aalami et al., 2004). When confronted with bone defects, reconstruction of bone is essential to restore skeletal function and integrity. To investigate the clinical applications of bone regenerative therapies, preclinical experimental models have been widely utilized to provide a “proof of principle” (Vajgel et al., 2014). The critical size defect (CSD) model has become an invaluable tool in studying bone regeneration and evaluating the limits of spontaneous healing. A critical size defect is defined as the smallest osseous lesion that will fail to heal spontaneously within the lifespan of the organism, thus necessitating reintervention for successful bone regeneration and healing. Schmitz and Hollinger introduced the CSD concept, emphasizing defect size as a key factor in healing (Schmitz and Hollinger, 1986). The critical size defect model is intended to standardize the testing of bone repair methods and materials that could be used as either alternatives or supplements in existing auto- and allograft materials. By utilizing the CSD model, researchers can provide a broad understanding of the underlying mechanisms of bone formation. Although this model is highly used and has proven to be highly effective in pre-clinical research studies, the concept of the Critical Size Defect model is subject to discrepancies due to variation across studies. Differences in species, age, defect size, and surgical technique affect study outcomes and complicate standardization. Animal models ranging from mice to non-human primates demonstrate differing intrinsic healing capacities. These variations offer valuable insight but hinder efforts to define a consistent “critical” size. Younger animals, in particular, show stronger regenerative abilities than older ones (Vajgel et al., 2014; Cooper et al., 2010) (Table 1).

A diverse range of animal models, from guinea pigs and mice to rats and even non-human primates, has been employed to investigate the Critical Size Defect (CSD) model (Cooper et al., 2010; Yoon et al., 2007; Spicer et al., 2012; Delgado-Ruiz et al., 2015; Marei et al., 2018; Yu et al., 2008; Voss et al., 2022; Moest et al., 2020) (Table 2). Each species showcases distinct intrinsic healing capabilities, contributing unique insights to the study of bone regeneration. Moreover, the varied sizes of these animal models enable researchers to explore a spectrum of defect sizes, enriching the breadth of experimental investigations. Additionally, the age of the organisms plays a pivotal role in shaping study outcomes. Studies consistently demonstrate that younger organisms exhibit a more robust capacity for healing, often showcasing more complete regenerative responses compared to their older counterparts (Aalami et al., 2004).

Another key factor influencing the experimental outcome in critical size defect (CSD) models is the surgical techniques employed. While several techniques have been described in the previous literature for creating the calvarial defect model, with slight variations in execution. However, options may be constrained by the small size, thin bone structure, and anatomical characteristics of the rat calvaria (Cooper et al., 2010). Commonly reported methods include trepanning, drilling, and punching. The trepanning method involves using a trephine bur of a specific diameter to create the defect, offering high consistency and reproducibility—attributes essential for comparative studies (Pereira et al., 2011). The drilling method utilizes a dental or orthopedic drill to achieve the desired defect size, but is highly operator-dependent, often resulting in variability in defect dimensions. The punch method, similar to the trepanning technique, employs a biopsy punch and has also demonstrated consistency and reproducibility (Kasuya et al., 2018).

A consistent priority across all surgical approaches is the preservation of surrounding tissues during the procedure. This is crucial to enable a more accurate assessment of the bone’s intrinsic healing potential, free from extrinsic or iatrogenic interference. Of particular importance is the dura mater—a dense membrane located directly beneath the calvarial bone in rats. As the outermost layer of the meninges, it not only protects the central nervous system but also contributes significantly to the regenerative and developmental processes of the calvaria. Despite its wide usage, inconsistencies persist regarding the exact parameters that constitute a CSD. Variability in species, age, defect size, surgical technique, and healing timelines challenge the establishment of universal standards (Cooper et al., 2010; Spicer et al., 2012; Senos and Hankenson, 2020). For instance, younger animals demonstrate superior regenerative capacity compared to adults, and surgical techniques vary in reproducibility depending on the method used (trephine vs. drill vs. punch) (Hudieb et al., 2021).

In rodent models, particularly rats, critical size calvarial defects are most frequently studied. Traditional practices recognize 5 mm and 8 mm defects as “critical,” yet limited data exists for intermediate or smaller defect sizes (Cooper et al., 2010; Spicer et al., 2012; Senos and Hankenson, 2020). Furthermore, defect size is often reported based on the nominal trephine bur diameter rather than the actual postoperative measurement, contributing further to inter-study variability.

This study aims to clarify and propose standardization for the rat calvarial CSD model by:

• Precisely defining critical defect size based on postoperative measurements.

Through this comprehensive evaluation, we aim to strengthen the translational relevance of rat calvarial CSD models in bone tissue engineering applications.

Buprenorphine (Ethiqa, catalog number 1217793), Lidocaine, 1 wt/vol% with 1:100,000 epinephrine (Henry Schein, catalog number 1047099), Bupivacaine HCL 0.5%. (Covertus, catalog number 054893), Sterile normal saline (Henry Schein, catalog number 6985812), Formalin (Fisher Scientific, catalog number SF100). Isoflurane (Henry Schein, catalog number 1084262), Phosphate buffered saline (Gibco, catalog number 21600010) ethanol, 70% (VWR, catalog number 71001–654), Oxygen, USP Grade (Matheson Tri-Gas), Carbon dioxide, USP Grade (Matheson Tri-Gas).

Animals: All animals survived until 8 weeks. There were no injuries to the dura while creating the defect. The wound area showed no signs of inflammation or adverse reactions after surgery.

Micro CT: Micro-CT was employed to obtain 3D reconstruction images for characterizing the quality and quantity of bone healing and newly formed bone (Bouxsein et al., 2010). The qualitative evaluation of spontaneous bone healing ability and comparison among the groups demonstrated that bone healing began at the edges of the defect (Figures 4A–C). However, over the course of 8 weeks, the defect was not completely healed, indicating that it was of a critical size. Figure 4B shows the trans-axial view (2-D) of bone healing at Day 0 and Week 8, showing that no considerable bone healing occurred until Week 8. Day 0 to Week. Figure 4C shows the quantitative data for bone volume (mm³), percent bone volume, trabecular thickness (Tb Th), Trabecular space (Tb Sp), and trabecular number (Tb N). The quantitative data on bone volume and percent volume showed that there was no significant increase from day 0 to week 1. Bone healing began at week 4 and continued until week 8. Most of the healing occurred around the perimeter, and the central area remained empty. The percentage of bone healing ranged from 0.1 to a maximum of 7% among all the rats, except for one defect, where the maximum healing occurred at 11.49% (data not shown). Trabecular thickness and trabecular number significantly increased (P < 0.05) compared to day 0 or day 7 to 4 weeks and 8 weeks, while trabecular space showed a slight decrease. However, there was no significant change throughout the time. The trabecular thickness increased by over 2.5-fold at 4- and 8-week post-surgery compared to day 0, while the trabecular number showed approximately a fourfold increase in the 4-week and 8-week groups compared to day 0.

Histology: Histological data provide two-dimensional measurements (Dempster et al., 2013). We collected data at the end of the study period. The rats were euthanized at the end of the study period (8 weeks). The histological sections were stained with H&E. The defect showed deposition of connective tissue consisting of fibroblasts and loose collagen fibers. Areas of necrotic bone without osteocytes were seen in some samples. Except in two defects where there was little bone formation observed, no areas of newly forming bone were identified (Figures 5A,B). The areas where small bone formation occurred were towards the periphery, which is in agreement with our micro-CT results. Overall results showed that there was no functional bone formation at the end of 8 8-week period, indicating that the defect made was of critical size.

This study presents a rat model for critical-size calvarial defects, standardized to improve reproducibility and translational potential in bone tissue engineering research. The goal of this study was to provide a critical size defect model that will be useful for bone regenerative therapies. To ensure compatibility with future phases of this research involving xenogeneic cell transplantation, we selected athymic rats as our animal model. Athymic (nude) rats lack a thymus and are therefore immunodeficient, making them incapable of mounting a T-cell–mediated immune response. This unique immunological profile allows for the engraftment and survival of human-derived stem cells. This continuity in model selection improves the translational relevance of our findings and facilitates comparison between scaffold-only and stem cell-augmented healing outcomes (Macchiarini et al., 2008; Bugelski and Martin, 2012).

Although we followed standard surgical techniques, keeping animal welfare in mind, we made some improvisations on specific surgical techniques. A 2 cm midline incision was made along the calvarium, undermining to expose coronal sutures and parietal plates. The incision length was selected to allow sufficient tissue retraction without interfering with the cutting edge of the trephine burr. We selected not to use the L-shaped incision as previously reported due to its association with increased inflammation and a higher risk of tissue damage associated with longer incisions.

Another key modification in our surgical technique was to create a standardized surgical field and minimize endogenous sources of osteogenesis. To achieve this, the periosteum was carefully dissected away from the calvarium prior to defect creation. The periosteum is a highly vascularized membrane rich in mesenchymal stem cells, known to contribute significantly to calvarial bone regeneration through both direct osteoprogenitor activity and the secretion of pro-angiogenic and osteoinductive factors, such as vascular endothelial growth factor (VEGF) and bone morphogenetic proteins (BMPs) (Colnot, 2009; Bouletreau et al., 2002). Previous studies have shown that spontaneous healing in critical-sized defect models can be influenced by periosteal preservation, thereby introducing variability that may confound the interpretation of experimental interventions (Histing et al., 2011; Schindeler et al., 2008). By removing the periosteum, we aimed to eliminate this source of variability and improve internal consistency across experimental groups.

The rodent critical size defect (CSD) model is well-established in bone regeneration research, with adequate data characterizing the healing pattern in defects that exceed defined thresholds. In particular, standardized models frequently define CSDs as 8 mm defects in rat calvaria and 5 mm in mouse calvaria, as these sizes consistently fail to heal spontaneously within the animal’s lifetime (Schmitz and Hollinger, 1986). While numerous studies have explored these defect sizes, there is a notable lack of evidence and consensus regarding the healing response in sub-critical defects—those smaller than the established CSD thresholds. Although many publications reference the concept of a “critical size,” the literature lacks a universally accepted percentage-based threshold for defining criticality. This is primarily due to variations in experimental design, animal models, anatomical sites, and healing assessment methodologies across studies, which complicate direct comparisons and the establishment of standardized criteria. The goal of this article is to provide a standardization of the CSD model in the context of defects, while emphasizing clinical relevance. Thus, we have chosen to investigate the time course of spontaneous bone healing and evaluate whether the defect size that we made was critical. In our study, we considered day 0 as our baseline (Control) and evaluated bone healing.

While many studies define defect size based on the bur used, we measured each defect postoperatively with precision calipers. This revealed that a head diameter of 3.75 mm (inner diameter) bur consistently produced 4.5 mm (with 4.75 mm outer diameter) diameter defects, likely due to bur geometry, drilling pressure, and microfracture at the margins. Vajgel et al. highlighted these inconsistencies, noting that while a 5 mm defect is often considered appropriate for the CSD model in rodents, further studies are necessary to establish standardized criteria (Vajgel et al., 2014). Accurate measurement is crucial in determining whether a defect meets the criteria for critical size. Table 3 represents the studies which has mentioned the bur size.

In contrast to studies that report defect size based on trephine bur diameter, we utilized a 3.75 mm round bur to create defects that were consistently measured postoperatively as 4.5 mm in diameter using precision calipers. We confirmed the diameter of the defect utilizing the ROI in micro-CT. The discrepancy between bur diameter and final defect size is attributable to several factors, including bur geometry, drilling angle, pressure during surgery, and minor bone fragmentation at the edges of the defect (Hidalgo and Pusic., 2002; Schmitz and Hollinger, 1986). For this reason, all measurements reported in this study reflect the actual post-surgical defect dimensions rather than the nominal bur size. This distinction is crucial in determining what qualifies as a critical-sized defect and in comparing results across studies. Due to the geometry of trephine burs, an 8 mm trephine bur will make a defect larger than 8 mm. Previous studies reported the size of the bur and not the post-operative defect size (Spicer et al., 2012). Bur size doesn’t accurately represent the true size of the defects, which could create significant discrepancies in the study results. Such inconsistencies have been underreported in the literature, despite their potential impact on the reproducibility and interpretation of bone healing outcomes in cranial defect models.

Micro-CT analysis revealed that bone formation was not uniform throughout the defect, more towards the periphery not in the center. Our results revealed that there was no spontaneous healing until 8 weeks, indicating that the defect we made was a critical-size defect. The percent bone formation has not exceeded beyond—7%, except in one defect (11.49%). A 2023 study by Qabbani et al. found an average healing rate of 10% at the control defect site with no intervention (Al Qabbani et al., 2023). This study used 24 Sprague-Dawley rats with a similar 8-week period, however, a 5 mm defect.

In contrast with the micro-CT data, Histological analysis at 8 weeks post-surgery revealed no evidence of bone formation. Histologically, we observed the presence of dense fibrous tissue interspersed with fibroblastic cells, with a little or no mineralization. This discrepancy can be attributed to the presence of small and scattered mineral deposits that are sufficient to generate a micro-CT signal but are lost during the decalcification process used for histological examination. Study limitations also include a relatively small sample size and the restriction of histological assessment to a single time point—8 weeks post-surgery.

The recent advances in the scientific field, offering NAMs (new approach methods) such as organoids and organ-on-a-chip technologies, hold significant promise as alternatives to traditional animal surgical methods (Zhou et al., 2025a). Artificial intelligence (AI)-based technology, a game-changer in drug discovery, has ushered in human-relevant research that has entered clinical trials (Zhou et al., 2025b; Liu et al., 2025). Organoids and Organ-on-Chip (OoCs) each offer unique advantages, such as cellular fidelity and ease of implantation (Liu et al., 2025). While organoids provide several benefits, such as cellular fidelity and ease of implantation, they differ from natural organs in that they lack vascularization and standardization. Especially in bone regenerative therapies, vascularization plays a crucial role. Furthermore, animal model-based testing of monoclonal antibodies and biologics has been particularly challenging as species-specific targets and immunogenicity limit the predictive power of animal models. In this scenario, animal models remain essential to providing a systemic environment of a whole organism.

Thus, our overall results support the assertion that 4.5 mm calvarial defects in athymic rats fail to heal spontaneously over 8 weeks. Minimal bone fill and lack of bridging on histology confirm this. These findings align with and build upon earlier reports, underscoring the importance of both measurement standardization and periosteum removal in CSD models.

Thus, we developed the critical size calvarial defect model. This study demonstrates the successful creation of critical-size calvarial defects in the athymic rat model, which will be favorable for testing the vascularized cell-scaffold system. These results support the validity of our model for evaluating bone regeneration strategies and biomaterial performance in future preclinical studies.