A rat model of the induced membrane technique using autologous iliac cancellous bone graft

A rat model of the induced membrane technique using autologous iliac cancellous bone graft | Research Square window.SnipcartSettings = { analytics: { enabled: false } }; (function() { var accessVector = localStorage.getItem('access_vector') || ''; window.dataLayer = window.dataLayer || []; if (accessVector) { window.dataLayer.push({ user: { profile: { profileInfo: { snid: accessVector } } } }); } })(); (function(w,d,s,l,i){w[l]=w[l]||[];w[l].push({'gtm.start':new Date().getTime(),event:'gtm.js'});var f=d.getElementsByTagName(s)[0],j=d.createElement(s),dl=l!='dataLayer'?'&l='+l:'';j.async=true;j.src='https://www.googletagmanager.com/gtm.js?id='+i+dl;f.parentNode.insertBefore(j,f);})(window,document,'script','dataLayer','GTM-K279D39R'); Browse Preprints In Review Journals COVID-19 Preprints AJE Video Bytes Research Tools Research Promotion AJE Professional Editing AJE Rubriq About Preprint Platform In Review Editorial Policies Our Team Advisory Board Help Center Sign In Submit a Preprint Cite Share Download PDF Article A rat model of the induced membrane technique using autologous iliac cancellous bone graft Kanu Shimokawa, Hidenori Matsubara, Tomo Hamada, Toshifumi Hikichi, and 2 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-8741751/v1 This work is licensed under a CC BY 4.0 License Status: Posted Version 1 posted You are reading this latest preprint version Abstract The induced membrane technique (IMT) is widely used for the treatment of segmental bone defects; however, small-animal models that faithfully reproduce clinical conditions remain limited. In particular, no rat IMT model using autologous iliac cancellous bone grafts (AICB) has been previously established. This study aimed to develop and characterize such a clinically relevant rat IMT model. A 5-mm segmental femoral defect was created in rats and stabilized with an external fixator. A polymethyl methacrylate spacer was inserted and replaced with bone graft two weeks later. Outcomes were compared among groups receiving no graft, artificial bone, AICB with IMT, and AICB without spacer placement. Bone regeneration was evaluated using computed tomography, histology, and real-time RT-PCR. AICB with IMT group achieved the highest bone union rate (66.7%) and demonstrated progressive increases in CT values and new bone formation. In the group, histological analyses demonstrated bone formation mediated by the induced membrane, accompanied by increased expression of osteogenic and angiogenic factors on real-time RT-PCR. This study establishes a reproducible rat IMT model using AICB, closely replicating the clinical IMT and providing an experimental platform for invnestigating IMT biology and therapeutic strategies for segmental bone defects. Biological sciences/Biotechnology Health sciences/Diseases Health sciences/Medical research Biological sciences/Stem cells Induced membrane technique Rat model autologous iliac cancellous bone graft Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Figure 6 Figure 7 Figure 8 Introduction Segmental bone defects arise from high-energy trauma, infection, and tumor resection, and remain a major clinical challenge in orthopedic surgery due to their complexity and the need for multiple surgical procedures, which significantly impair patients’ quality of life [ 1 , 2 ] . Among various reconstructive options, the induced membrane technique (IMT), also known as the Masquelet technique, has become increasingly adopted two-stage procedure for treating large bone defects and recalcitrant nonunions [ 3 , 4 ] . Compared with alternative strategies such as vascularized bone grafting or distraction osteogenesis [ 5 , 6 ] , IMT offers advantages in technical simplicity and is theoretically less influenced by defect size with respect to healing time [ 7 ] . Despite its widespread use, several aspects of IMT—including membrane biology, graft selection, and optimal surgical parameters—remain subjects of debate, in part because many of these factors are difficult to investigate in human clinical settings. Consequently, a variety of animal models have been developed to characterize induced membrane formation and to evaluate how procedural variables affect subsequent bone regeneration [ 8 – 10 ] . Autologous cancellous bone (ACB) remains the gold standard graft material for fracture reconstruction, delayed unions, and nonunions because of its inherent osteogenic, osteoinductive, and osteoconductive properties [ 11 ] . ACB contains osteoblasts, mesenchymal stem cells, growth factors, and an interconnected trabecular scaffold that facilitates vascular ingrowth and cellular infiltration, making it a reliable option for both acute and reconstructive trauma surgery [ 12 – 16 ] . In contrast, commonly used alternatives in small-animal research—including deproteinized bovine bone, heat-treated porcine bone, or freeze-dried allograft harvested from donor rats—serve primarily as osteoconductive scaffolds and lack the cellular and molecular components that confer the biological activity of fresh autograft [ 17 – 19 ] . These substitutes avoid the need for invasive bone harvesting but do not accurately replicate the clinical situation in which autograft is used. Moreover, models requiring donor animals increase cost and complexity. Although effective ACB models have been established in medium and large animals, there is a lack of small animal models that faithfully reproduce the clinical environment of autologous bone grafting [ 20 – 23 ] . Small animal models remain essential because they enable detailed investigation of tissue-level and molecular mechanisms, facilitate the development of new biomaterials, cellular therapies, and surgical strategies, and allow studies with larger sample sizes at lower cost. Our group previously established a rat model of autologous iliac crest bone harvesting and transplantation, demonstrating that reproducible autograft collection and successful graft incorporation can be achieved in a small-animal setting [ 24 ] . Building upon this validated platform, we sought to determine whether the IMT could similarly be performed using true autologous iliac crest grafts in rats—a clinically relevant but previously unreported approach. Therefore, the purpose of this study was to develop and characterize a rat IMT model using autologous iliac crest cancellous bone (AICB) grafts. This model aims to more closely replicate the clinical conditions of autograft-based IMT and to provide a robust experimental framework for future studies investigating graft biology, membrane function, and novel therapeutic strategies for segmental bone defects. Results Histological evaluation of femoral specimens harvested two weeks after the first surgery revealed the presence of a membrane-like structure surrounding the cavity previously occupied by the spacer (Fig. 3a). The membrane exhibited a characteristic bilayer architecture, consisting of an inner layer composed of dense fibrous tissue and an outer layer composed of loose connective tissue with abundant microvessels. These histological features were consistent with previously reported pathological characteristics of the induced membrane [ 10 ] (Fig. 3b,c). Representative longitudinal CT images of rats from Groups A–D at each time point are shown in Fig. 4 (Fig. 4a–c). The bone union rates at the graft site were 0% in Group A, 6.7% (1 rat) in Group B, 66.7% (10 rats) in Group C, and 20.0% (3 rats) in Group D. Group C demonstrated the highest bone union rate, with changes suggestive of new bone formation observed as early as two weeks after grafting, becoming more pronounced over time. Bone formation was predominantly observed on the medial side of the femur, whereas union on the lateral side tended to be delayed. In Groups A and B, bone formation at the defect site was minimal; the bone edges became rounded and regressed over time, and the synthetic bone graft in Group B showed a tendency toward resorption. Comparative CT analysis between Groups A and C demonstrated distinct temporal patterns of bone formation (Fig. 4d,e). In Group C, CT values increased progressively over time, and a significant overall temporal change was observed (p < 0.05). In Group A, CT values also showed a significant temporal change, characterized by an initial increase followed by slight decrease at later time points (p < 0.05). At all time points, CT values in Group C were significantly higher than those in Group A (p < 0.05). Similar temporal trends and significant intergroup differences were also observed in the proportion of newly formed bone, consistent with the findings for CT values. Histological sections stained with hematoxylin and eosin from Groups A–D are shown in Fig. 5. In Groups C and D, sagittal sections obtained two weeks after grafting confirmed that the transplanted cancellous bone was located within the bone defect. In Group C, the cancellous bone was progressively replaced by newly formed bone over time, and bridging of the defect by new bone was observed from four weeks onward. Notably, in Group C, infiltration of osteoblasts from the membrane surrounding the graft into the bone defect was observed in two to three cellular layers (Fig. 6a–f). This finding was evident from four to eight weeks after grafting, although the degree of osteoblast infiltration tended to decrease at eight weeks as bone maturation progressed. In Group D, although some replacement of the grafted cancellous bone by new bone was observed, the extent of bone formation was less pronounced than in Group C, no membrane structure was present on the outer surface, and osteoblast infiltration was limited. Representative histological images from Groups C and D at four weeks after grafting are shown in Fig. 7a and 7b. Quantitative analysis revealed that the CD31-positive total vascular area at the graft site was significantly greater in Group C than in Group D at two, and six weeks after grafting ( p < 0.05) (Fig. 7c). To evaluate the molecular effects of induced membrane–assisted bone grafting, real-time RT-PCR analysis was performed (Fig. 8). BMP-2, TGF-β, and VEGF were selected as representative markers of osteogenic differentiation, extracellular matrix production, and angiogenesis, respectively. In Group C, VEGF expression was significantly higher than in Group A at four weeks after grafting ( p < 0.05). Expression levels of BMP-2 and TGF-β at four weeks were significantly higher in Groups C and D than in Group A. Although expression levels of all three markers tended to be higher in Group C than in Group D, no statistically significant differences were observed between these two groups. Discussion/Conclusion The induced membrane technique (IMT) has been widely adopted as a treatment for segmental bone defects and nonunion. However, several aspects of this technique—including the optimal timing of grafting, selection of spacer materials, choice of graft material, and the potential role of adjunctive factors—remain subjects of ongoing debate in clinical practice. Accordingly, animal models used to investigate IMT should also be refined to better reflect clinical conditions before translation to human application. Among the various factors influencing the IMT, graft selection represents one of the most critical determinants of treatment success. Autograft, allograft, and synthetic bone substitutes are commonly used options. Previous clinical studies have reported IMT procedures using mixtures containing up to 64% allograft combined with autograft [ 27 ] . Nevertheless, a 3:1 ratio of autograft to allograft has traditionally been the most widely employed combination [ 28 – 31 ] , and autologous bone grafting remains the clinical gold standard. Despite this consensus, there is no uniform agreement regarding the optimal donor site for graft harvesting, either in clinical practice or in animal experiments. Harvesting graft material from the iliac crest is one of the most common clinical approaches. Gens et al. reported that, in a rat bone graft model, iliac crest–derived samples exhibited qualitatively higher alkaline phosphatase (ALP) levels than samples harvested from other skeletal sites, suggesting a higher concentration of osteoprogenitor cells [ 32 ] . Similarly, studies of human bone grafts have demonstrated that the mean concentration of osteoblastic progenitor cells is higher in iliac crest grafts than in tibial grafts [ 33 ] . In addition, from a functional perspective, harvesting bone from the iliac crest is expected to have less impact on postoperative locomotion than harvesting from the extremities. Another major advantage of the iliac crest is the ability to obtain a relatively large amount of cancellous bone. This point is particularly important in small animals such as rats, in which long bones consist predominantly of cortical bone and contain only minimal cancellous components. Although rat models of autologous bone grafting have been previously reported, most have relied on cortical bone as the graft material [ 34 – 36 ] . While autologous cortical grafts provide high mechanical strength and immediate structural support, they exhibit limited osteoinductive potential compared with cancellous bone [ 37 – 39 ] . Consequently, cancellous bone is generally more effective for promoting bone union and is preferentially used in clinical bone grafting. Despite these advantages, a systematic review analyzing 47 studies investigating IMT in animal models found that only seven studies harvested autologous grafts from the iliac crest, all of which involved rabbits or sheep; no rat iliac crest bone graft models were included [ 1 ] . This lack of rat models likely reflects the absence of established techniques for safely and reliably harvesting iliac bone from small animals. Many previously reported rat IMT models have therefore relied on allografts or synthetic bone substitutes [ 1 , 8 – 10 , 40 ] , and bone grafting using caudal vertebrae has been particularly common. However, the rat tail plays an important role in thermoregulation and temperature sensation, and tail amputation in live animals is not recommended, raising animal welfare concerns [ 32 , 41 ] . Moreover, caudal vertebrae are composed primarily of cortical bone, with only minimal cancellous bone available for harvesting. For these reasons, the iliac crest appears to offer clear advantages over the tail as a donor site for bone grafting. To our knowledge, the present study is the first to establish a rat IMT model using autologous iliac cancellous bone grafting. Using this approach, an adequate volume of cancellous bone was safely harvested, and the surgical procedures closely replicated the clinical IMT. Histological evaluation following spacer removal confirmed the presence of a characteristic bilayer membrane structure consistent with previously described features of the induced membrane, indicating successful membrane formation. In the comparative analysis, bone union was not achieved in Group A (no graft) or Group B (artificial bone graft), whereas successful bone union was observed in 67% of rats in Group C (AICB with IMT). These findings indicate that cancellous bone grafting plays a favorable role in promoting bone healing. Furthermore, the higher bone union rate in Group C compared with Group D (without induced membrane) suggests that the presence of an induced membrane provides an additional biological advantage beyond AICB alone. Collectively, these results demonstrate that the IMT using AICB yields the most consistent and stable bone union outcomes among the models tested, supporting its validity as a reproducible experimental platform. CT evaluation revealed that in some cases, bony continuity was established as early as two weeks after grafting. Quantitative analysis further demonstrated that in Group C, both mean CT values and the proportion of newly formed bone increased progressively over time and were significantly higher than those observed in Group A. In contrast, these parameters showed a slight decline at eight weeks in Group A, which may reflect changes associated with nonunion. Bone union tended to occur earlier on the medial side of the femur, a finding that may reflect not only differences in surrounding soft tissue volume but also the incision of the lateral membrane during spacer removal. Histological analyses provided further evidence of osteoblast infiltration from the induced membrane into the bone defect. This influx of osteoblasts was most prominent at four weeks after grafting and gradually decreased by eight weeks as bone maturation progressed. These observations support the concept that the induced membrane actively contributes to osteogenesis. In contrast, no membrane-like structure was histologically identified in Group D, in which AICB grafting was performed without spacer placement. Quantitative analyses demonstrated that angiogenesis, as assessed by immunohistochemistry, was significantly greater in Group C, and real-time RT-PCR revealed that VEGF expression was significantly upregulated only in Group C compared with Group A. Thus, although angiogenic activity was clearly enhanced in the IMT-treated Group C, no marked differences in BMP-2 or TGF-β expression were observed between Groups C and D. This finding may be attributable to the biological effects inherent to cancellous bone grafting itself. Nevertheless, the substantial difference in union rates between the two groups suggests that the physical effects associated with membrane formation, such as prevention of graft dispersion and protection against external contamination, may have also played a role. Indeed, many of the rats that failed to achieve bone union exhibited, to some extent, signs of infection. Taken together, these findings suggest that induced membrane formation using the IMT provides a more favorable biological and mechanical environment for bone healing than cancellous bone grafting alone. Several limitations of this model should be acknowledged. First, the volume of cancellous bone that can be harvested from the iliac crest is limited. Hamada et al. reported a mean harvested cancellous bone volume of 73.8 ± 5.5 mm³ in a rat autologous cancellous bone graft model [ 24 ] , corresponding to a graft length of approximately 5.8 mm with a 4-mm diameter or 10 mm with a 3-mm diameter. Therefore, for large bone defects exceeding 10 mm, the use of AICB alone may be insufficient in rats of this size. Second, in the present study, the duration of spacer placement was set at two weeks. Although this interval was slightly shorter than that reported in many previous studies [ 8 , 9 , 40 ] , histological evaluation of the membrane immediately before grafting confirmed a characteristic bilayer structure consistent with prior literature. Moreover, multiple findings suggested that this membrane exerted a favorable effect on subsequent bone healing, indicating that a two-week waiting period was sufficient in this model. Nevertheless, the optimal duration of spacer placement remains to be determined, and it is possible that the selected interval was not the most favorable. Further studies directly comparing different waiting periods are warranted to clarify the optimal timing for grafting in IMT. Third, external fixation was used for stabilization in this study. This choice was made because IMT is often applied clinically to traumatic bone defects or cases complicated by infection, in which external fixation is frequently preferred over internal fixation. However, maintaining strict postoperative immobilization in rats is challenging, and loosening of fixation pins due to contact with the cage or pin tract infections may have contributed to nonunion in some cases. Although IMT models using internal fixation with plates have been reported [ 7 , 42 ] , the use of external fixation in small animals may increase the susceptibility to fixation-related complications, which should be considered when interpreting the results. In conclusion, we successfully established a reproducible rat IMT model using autologous iliac cancellous bone grafting. This model closely replicates the clinical IMT and provides a valuable experimental platform for investigating the biological mechanisms underlying IMT and for developing novel therapeutic strategies for the treatment of segmental bone defects. Methods Animal experiments: All animal experiments were approved by the Animal Experimentation Committee of Kanazawa University (Approval No. AP-204176). All procedures were performed in accordance with the institutional guidelines, the ARRIVE guidelines (PLoS Biol 8(6): e1000412, 2010), and the relevant regulations. Anesthesia and euthanasia protocols adhered to the American Veterinary Medical Association (AVMA) Guidelines for the Euthanasia of Animals (2020). The study does not involve endangered or CITES-listed species. Healthy male Sprague–Dawley rats (12 weeks old, 350–400 g) were obtained for this study. The animals were housed individually under specific pathogen-free conditions with a 12-h light/dark cycle and given free access to food and water. All rats were acclimatized to the laboratory environment for one week prior to experimentation. For all procedures, anesthesia was induced by intraperitoneal injection of medetomidine (0.15 mg/kg), midazolam (2 mg/kg), and butorphanol (2.5 mg/kg). At the end of the study, the rats were euthanized by intraperitoneal administration of pentobarbital (450 mg/kg). IMT procedure: All rats underwent two surgical procedures. Figure 1 illustrates the overall design of the first-stage procedure. The femoral defect model was adapted from previously reported methods in which nonunion was consistently observed without fixation implants [ 25 , 26 ] . Each rat was placed in the prone position on the operating table. A longitudinal lateral skin incision was made over the right femur, and the quadriceps and hamstring muscles were split and retracted (Fig. 1a,b). After pre-drilling with a 1.4-mm Kirschner wire, an external fixator (Meira Co., Nagoya, Japan) was mounted to the femur using four self-tapping pins (1.6 mm in diameter; Japan Medicalnext Co., Osaka, Japan) (Fig. 1c). Two osteotomies were created with a bone saw between the second and third pins to generate a 5-mm segmental bone defect (Fig. 1d). A polymethyl methacrylate (PMMA) spacer was inserted into each defect (Fig. 1e). The muscle layer, subcutaneous tissue, and skin were closed with simple interrupted sutures. To investigate bone regeneration, all rats underwent a second surgery two weeks after the initial procedure. The rats were positioned identically to the first surgery, and the previous lateral skin incision was reopened. After confirming the presence of membrane-like tissue (induced membrane) overlying the spacer (Fig. 1f), the membrane was incised with a scalpel, and the spacer was removed (Fig. 1g). The membrane cavity between the bone ends was then filled according to one of the following three conditions: no graft material (Group A), a β-TCP bone substitute block (Group B), or AICB (Group C) (Fig. 1h, 2e). The incised membrane was sutured, and the wound was closed in layers. Additionally, to determine whether placement of a cement spacer is essential for IMT, a separate group was created in which the bone defect was generated during the first surgery but closed without inserting a spacer. During the second surgery, these rats received AICB grafting in the same manner as Group C (Group D). Autologous iliac bone graft model: AICB harvesting was performed according to the procedure described by Hamada et al. [ 24 ] . A longitudinal midline skin incision of approximately 4 cm was made over the highest point of the bilateral iliac crests (Fig. 2a). The gluteus maximus was elevated from the dorsal midline, and the dorsomedial and dorsolateral fibers of the coccygeus muscle were separated to expose the iliac crest (Fig. 2b,c). Care was taken to avoid injury to deep neurovascular structures while performing the osteotomy, and each iliac crest was harvested bilaterally. Using a scalpel, cancellous bone was separated from the cortical shell, and the morselized cancellous bone was packed into a cylindrical mold and compressed to form a 5-mm-thick block (Fig. 2d). In group C, the AICB block was transplanted into the femoral bone defect. (Fig. 2e). The submuscular and gluteal layers were closed with simple interrupted sutures, followed by closure of the skin. Imaging Evaluation: Computed tomography (CT) imaging was performed using a Latheta LCT-200 system (Hitachi Aloka Medical, Tokyo, Japan). Scans were obtained immediately after grafting and at 2, 4, 6, and 8 weeks postoperatively to evaluate longitudinal bone formation and the presence of bone union. Fifteen rats per group were assessed to determine the bone union rate. Coronal CT images were used to define the femoral axis. To exclude pre-existing cortical bone from the analysis, bone formation was evaluated only within the central defect region, defined as a 5-mm long-axis × 4-mm short-axis region of interest. CT values were calibrated using water (CTw = 0) and air (CTa = − 1000) as reference standards, and regions with CT values exceeding + 1000 were extracted as newly formed bone. Image analysis was performed using Synapse VINCENT software (Fujifilm, Tokyo, Japan). For Groups A and C, rats at each time point (N = 5 per group) were further analyzed to calculate the mean CT value within the region of interest and the proportion of newly formed bone. Temporal changes and intergroup comparisons were subsequently evaluated. Histological Evaluation: At each designated time point, harvested femora were fixed in 10% neutral-buffered formalin with surrounding soft tissues and the external fixator left in situ. The specimens were then dehydrated through a graded ethanol series (70%, 80%, 90%, and 100%). After decalcification in a 10% sodium citrate–formic acid solution, the external fixators were removed. The specimens were embedded in paraffin, sectioned in the sagittal plane, and stained with hematoxylin and eosin (H&E). Histological evaluation was performed using a BZ-9000 optical microscope (Keyence, Osaka, Japan). Specimens were collected at 2, 4, 6, and 8 weeks after grafting in all groups to assess histological bone formation and its relationship with the induced membrane. In addition, specimens from Groups C and D were collected immediately after grafting to evaluate the presence or absence of the induced membrane prior to transplantation. In Group C, membrane formation consistent with previous reports was anticipated, whereas no membrane formation was expected in Group D. Immunohistochemistry: To assess angiogenesis at the graft site, immunohistochemical staining for endothelial cells was performed using an anti-CD31 antibody (Abcam, Cambridge, UK; ab28364; 1:50 dilution). Tissue sections were incubated at room temperature for 15 minutes in Liberate Antibody Binding Solution (LAB solution; Polysciences, Philadelphia, PA, USA), followed by blocking with Protein Block Serum-Free (Dako, Glostrup, Denmark) and peroxidase-blocking solution containing hydrogen peroxide (Dako, Glostrup, Denmark) for 10 minutes each. Sections were then incubated overnight at 4°C with the primary anti-CD31 antibody diluted in Tris-HCl buffer containing stabilizing proteins and 0.015 mol/L sodium azide (Dako Antibody Diluent; Dako, Glostrup, Denmark). After incubation with a secondary antibody (Dako REAL EnVision/HRP, Rabbit/Mouse; Dako, Glostrup, Denmark) for 30 minutes at room temperature, immunoreactivity was visualized using DAB chromogen (Dako REAL DAB+ Chromogen; Dako, Glostrup, Denmark). The reaction was terminated by rinsing the slides with deionized water once an optimal signal-to-noise ratio was achieved. To specifically investigate angiogenesis associated with induced membrane formation, analyses were performed in Groups C and D. Vascular area at the graft site was measured at 2, 4, 6, and 8 weeks after grafting. In each group, five randomly selected high-power fields (×40) from five nonconsecutive sections were quantified. Measurement of vascular area and image analysis were performed using a BZ-X800 all-in-one fluorescence microscope with time-lapse functionality (Keyence, Osaka, Japan). Real-Time RT-PCR Analysis: Total RNA was extracted from graft-site tissues using the NucleoSpin RNA II kit (Takara Bio, Otsu, Japan). To evaluate the effects of bone grafting and induced membrane formation on angiogenesis-related gene expression, analyses were conducted using samples from Groups A, C, and D. Harvested tissues were mechanically homogenized using a syringe (n = 5 per group at each time point). RNA concentration and purity were assessed using a UV–visible spectrophotometer (NanoDrop Lite; Thermo Scientific, Waltham, MA, USA). For real-time RT-PCR, 6 µg of total RNA was reverse-transcribed into complementary DNA (cDNA) using the RevertAid First Strand cDNA Synthesis Kit (Thermo Scientific, Waltham, MA, USA) and a thermal cycler (T100™ Thermal Cycler; Bio-Rad, Hercules, CA, USA). Quantitative PCR was performed using SYBR Green PCR Master Mix (Applied Biosystems, Foster City, CA, USA) on an ABI Prism 7900 system (Applied Biosystems, Foster City, CA, USA), in accordance with the manufacturer’s instructions. Primers for rat glyceraldehyde-3-phosphate dehydrogenase (GAPDH), bone morphogenetic protein-2 (BMP-2), vascular endothelial growth factor (VEGF), and transforming growth factor-β1 (TGF-β1) were purchased from Hokkaido System Science Co., Ltd. (Japan), and their sequences are listed in Table 1. The amplification parameters were an initial 95˚C incubation step for 15 minutes, followed by 20 amplification cycles of 94˚C for 15 seconds, 60˚C for 30 seconds, and 72˚C for 30 seconds. The reactions ended with a 72˚C extension step for 7 minutes, followed by storage at 4˚C overnight. The expression levels of each target gene were calculated relative to the level of GAPDH for each sample. Statistical Analysis: Data are presented as mean ± standard deviation (SD). Intergroup comparisons of CT values, CD31-positive vascular area, and relative gene expression levels were performed using the Mann–Whitney U test,, as the data did not meet assumptions of normality and the sample size in each group was small (n = 5) and nonparametric analysis was considered more appropriate. Temporal changes in CT values were evaluated using the Kruskal–Wallis test to assess overall trends over time. A p-value of < 0.05 was considered statistically significant. Declarations Funding Declaration: This work was supported by the National Mutual Insurance Federation of Agricultural Cooperatives (ZENKYOREN) in fiscal year 2021, the Shibuya Science Culture and Sports Foundation in fiscal year 2020, and by Japan Society for the Promotion of Science (KAKENHI) Grant-in-Aid for Scientific Research (C) in fiscal year from 2021 to 2023. Author Contribution K.S., H.M., and T.H. conceptualized and designed the study. K.S. primarily performed the experiments. Data interpretation and discussion were conducted by all authors, including T.H. and Y.N. K.S. performed data curation, formal analysis, and drafted the original manuscript. H.M. and S.D. supervised the study. K.S. acquired the funding. All authors reviewed and approved the final manuscript. Acknowledgements: Not applicable Data Availability The datasets generated and/or analyzed during the current study are available from the corresponding author on reasonable request. Additional Information (including a Competing Interests Statement): The authors declare that they have no competing interests related to this work. References Sun, H. et al. The induced membrane technique in animal models: a systematic review. OTA. Int. 10; 5(1 Suppl): e176 (2022). Svedbom, A. et al. Osteoporosis in the European Union: a compendium of country-specific reports. Arch. Osteoporos. 8 (1), 137 (2013). Pelissier, P. et al. Induced membranes secrete growth factors including vascular and osteoinductive factors and could stimulate bone regeneration. J. Orthop. Res. 22 (1), 73–79 (2004). Taylor, B. C., French, B. G., Fowler, T. T., Russell, J. & Poka, A. Induced membrane technique for reconstruction to manage bone loss. J. Am. Acad. Orthop. Surg. 20 (3), 142–150 (2012). Glynou, S. P., Georgiannakis, A., Ardolino, D., Craxford, S. & Vris, A. Vascularised Fibula Transfer for Post-traumatic Critical Tibial Bone Defects: A Systematic Review. Strategies Trauma. Limb Reconstr. 20 (1), 37–46 (2025). Tsuchiya, H. & Tomita, K. Distraction osteogenesis for treatment of bone loss in the lower extremity. J. Orthop. Sci. 8 (1), 116–124 (2003). Mauffrey, C., Barlow, B. T. & Smith, W. Management of segmental bone defects. J. Am. Acad. Orthop. Surg. 23 , 143–153 (2015). McBride-Gagyi, S. et al. Altering spacer material affects bone regeneration in the Masquelet technique in a rat femoral defect. J. Orthop. Res. 23866. 10.1002/jor (2018). Gouron, R. et al. Osteoclasts and their precursors are present in the induced-membrane during bone reconstruction using the Masquelet technique. J. Tissue Eng. Regen Med. 11 (2), 382–389 (2017). Tang, Q., Tong, M., Zheng, G., Shang, P. & Liu, H. Masquelet's induced membrane promotes the osteogenic differentiation of bone marrow mesenchymal stem cells by activating the Smad and MAPK pathways. Am. J. Transl Res. 10 (4), 1211–1219 (2018). Giannoudis, P. V., Dinopoulos, H. & Tsiridis, E. Bone substitutes: an update. Injury 36 Suppl. 3: S20–S27 (2005). Cassidy, C. et al. Norian SRS cement compared with conventional fixation in distal radial fractures. A randomized study. J. Bone Joint Surg. Am. 85 (11), 2127–2137 (2003). Finkemeier, C. G. Bone-grafting and bone-graft substitutes. J. Bone Joint Surg. Am. 84 (3), 454–464 (2002). Kelly, C. M. et al. The use of a surgical grade calcium sulfate as a bone graft substitute: Results of a multicenter trial. Clin. Orthop. Relat. Res. 382 , 42–50 (2001). Khan, S. N. et al. The biology of bone grafting. J. Am. Acad. Orthop. Surg. 13 (1), 77–86 (2005). Ray, R. D. Vascularization of bone grafts and implants. Clin. Orthop. Relat. Res. 87 , 43–48 (1972). Fitch, R., Kerwin, S., Sinibaldi, K. R. & Newman, H. E. Bone autografts and allografts in dogs. Comp. Vet. Cont. Ed. 19 (5), 558–575 (1997). McLaughlin, R. M. & Roush, J. K. Autogenous cancellous and corticocancellous bone grafting. Vet. Med. 93 , 1071–1074 (1998). Orth, M. et al. BMP-2-coated mineral coated microparticles improve bone repair in atrophic non-unions. Eur. Cell. Mater. 33 , 1–12 (2017). Athanasiou, V. T. et al. Histological comparison of autograft, allograft-DBM, xenograft, and synthetic grafts in a trabecular bone defect: An experimental study in rabbits. Med. Sci. Monit. 16 (1), BR24–31 (2010). Fox, S. M. Cancellous bone grafting in the dog—An overview. J. Am. Anim. Hosp. Assoc. 20 , 840–848 (1984). Klaue, K. et al. Bone regeneration in long-bone defects: Tissue compartmentalisation? In vivo study on bone defects in sheep. Injury 40 (Suppl 4), S95–102 (2009). Rosenthal, A. H. & Buchman, S. R. Volume maintenance of inlay bone grafts in the craniofacial skeleton. Plast. Reconstr. Surg. 112 (3), 802–811 (2003). Hamada, T., Matsubara, H., Hikichi, T., Shimokawa, K. & Tsuchiya, H. Rat model of an autologous cancellous bone graft. Sci. Rep. 11 (1), 18001 (2021). Glatt, V., Evans, C. H. & Matthys, R. Design, characterisation and in vivo testing of a new, adjustable stiffness, external fixator for the rat femur. Eur. Cell. Mater. 23 , 289–298 (2012). Harada, N. et al. Bone regeneration in a massive rat femur defect through endochondral ossification achieved with chondrogenically differentiated MSCs in a degradable scaffold. Biomaterials 35 (27), 7800–7810 (2014). Pesciallo, C. A., Garabano, G., Dainotto, T. & Ernst, G. Masquelet technique in post-traumatic infected femoral and tibial segmental bone defects. Union and reoperation rates with high proportions (up to 64%) of allograft in the second stage. Injury 52 (11), 3471–3477 (2021). Masquelet, A. C. & Begue, T. The concept of induced membrane for reconstruction of long bone defects. Orthop. Clin. North. Am. 41 (1), 27–37 (2010). Masquelet, A., Kanakaris, N. K., Obert, L., Stafford, P. & Giannoudis, P. V. Bone repair using the Masquelet technique. J. Bone Joint Surg. Am. 101 (11), 1024–1036 (2019). Karger, C., Kishi, T., Schneider, L., Fitoussi, F. & Masquelet, A. C. Treatment of posttraumatic bone defects by the induced membrane technique. Orthop. Traumatol. Surg. Res. 98 (1), 97–102 (2012). Baldwin, P. et al. Autograft, allograft, and bone graft substitutes: Clinical evidence and indications for use in the setting of orthopaedic trauma surgery. J. Orthop. Trauma. 33 (4), 203–213 (2019). Gens, L. et al. Surgical technique and comparison of autologous cancellous bone grafts from various donor sites in rats. J. Orthop. Res. 41 (4), 834–844 (2023). Hyer, C. F. et al. Quantitative assessment of the yield of osteoblastic connective tissue progenitors in bone marrow aspirate from the iliac crest, tibia, and calcaneus. J. Bone Joint Surg. 95 (14), 1312–1316 (2013). Bosemark, P., Isaksson, H., McDonald, M. M., Little, D. G. & Tägil, M. Augmentation of autologous bone graft by a combination of bone morphogenic protein and bisphosphonate increased both callus volume and strength. Acta Orthop. 84 (1), 106–111 (2013). Gouron, R. et al. Osteoclasts and their precursors are present in the induced-membrane during bone reconstruction using the Masquelet technique. J. Tissue Eng. Regen Med. 11 (2), 382–389 (2017). Murakami, H., Nakasa, T., Ishikawa, M., Adachi, N. & Ochi, M. Autologous bone grafts with MSCs or FGF-2 accelerate bone union in large bone defects. J. Orthop. Surg. Res. 11 (1), 105 (2016). Dell, P. C., Burchardt, H. & Glowczewskie, F. P. A roentgenographic, biomechanical, and histological evaluation of vascularized and non-vascularized segmental fibular canine autografts. J. Bone Joint Surg. Am. 67 (1), 105–112 (1985). Enneking, W. F., Burchardt, H., Puhl, J. J. & Piotrowski, G. Physical and biological aspects of repair in dog cortical-bone transplants. J. Bone Joint Surg. Am. 57 (2), 237–252 (1975). Doi, K., Tominaga, S. & Shibata, T. Bone grafts with microvascular anastomosis of vascular pedicles: An experimental study in dogs. J. Bone Joint Surg. Am. 59 (6), 806–815 (1977). Bosemark, P., Perdikouri, C., Pelkonen, M., Isaksson, H. & Tagil, M. The masquelet induced membrane technique with BMP and a synthetic scaffold can heal a rat femoral critical size defect. J. Orthop. Res. 33 (4), 488–495 (2015). Yehuda, S. d-Amphetamine and the sensory role of a rat's tail in thermoregulation or what the rat's tail tells the rat's brain. Behav. Biol. 13 (2), 233–238 (1975). Ma, Y. F. et al. Calcium sulfate induced versus PMMA-induced membrane in a critical-sized femoral defect in a rat model. Sci. Rep. 8 (1), 637 (2018). Additional Declarations No competing interests reported. Cite Share Download PDF Status: Posted Version 1 posted You are reading this latest preprint version Research Square lets you share your work early, gain feedback from the community, and start making changes to your manuscript prior to peer review in a journal. As a division of Research Square Company, we’re committed to making research communication faster, fairer, and more useful. We do this by developing innovative software and high quality services for the global research community. Our growing team is made up of researchers and industry professionals working together to solve the most critical problems facing scientific publishing. Also discoverable on Platform About Our Team In Review Editorial Policies Advisory Board Help Center Resources Author Services Accessibility API Access RSS feed Manage Cookie Preferences © Research Square 2026 | ISSN 2693-5015 (online) Privacy Policy Terms of Service Do Not Sell My Personal Information {"props":{"pageProps":{"initialData":{"identity":"rs-8741751","acceptedTermsAndConditions":true,"allowDirectSubmit":true,"archivedVersions":[],"articleType":"Article","associatedPublications":[],"authors":[{"id":591449142,"identity":"79b06381-ee1a-45b0-92a4-e460da116b5d","order_by":0,"name":"Kanu Shimokawa","email":"","orcid":"","institution":"Kanazawa University Hospital","correspondingAuthor":false,"prefix":"","firstName":"Kanu","middleName":"","lastName":"Shimokawa","suffix":""},{"id":591449143,"identity":"ddd8db7a-7500-43fc-a06c-0c14d2d51fdf","order_by":1,"name":"Hidenori Matsubara","email":"data:image/png;base64,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","orcid":"","institution":"Kanazawa University Hospital","correspondingAuthor":true,"prefix":"","firstName":"Hidenori","middleName":"","lastName":"Matsubara","suffix":""},{"id":591449144,"identity":"db1c810e-9a02-4d59-b74b-861c9e679c46","order_by":2,"name":"Tomo Hamada","email":"","orcid":"","institution":"Kanazawa University Hospital","correspondingAuthor":false,"prefix":"","firstName":"Tomo","middleName":"","lastName":"Hamada","suffix":""},{"id":591449145,"identity":"599f1224-f330-40b3-b4b4-45ba0ca94bdc","order_by":3,"name":"Toshifumi Hikichi","email":"","orcid":"","institution":"Kanazawa University Hospital","correspondingAuthor":false,"prefix":"","firstName":"Toshifumi","middleName":"","lastName":"Hikichi","suffix":""},{"id":591449146,"identity":"5f21c61e-365a-449f-957b-5309c4de1928","order_by":4,"name":"Yusuke Nakazawa","email":"","orcid":"","institution":"Kanazawa University Hospital","correspondingAuthor":false,"prefix":"","firstName":"Yusuke","middleName":"","lastName":"Nakazawa","suffix":""},{"id":591449147,"identity":"e1e5aec6-21c5-463d-90c2-d41d03c11b46","order_by":5,"name":"Satoru Demura","email":"","orcid":"","institution":"Kanazawa University Hospital","correspondingAuthor":false,"prefix":"","firstName":"Satoru","middleName":"","lastName":"Demura","suffix":""}],"badges":[],"createdAt":"2026-01-30 13:39:30","currentVersionCode":1,"declarations":"","doi":"10.21203/rs.3.rs-8741751/v1","doiUrl":"https://doi.org/10.21203/rs.3.rs-8741751/v1","draftVersion":[],"editorialEvents":[],"editorialNote":"","failedWorkflow":false,"files":[{"id":103014829,"identity":"800f5d2a-56ce-4c3c-96a9-ffdbe5dcd558","added_by":"auto","created_at":"2026-02-19 16:15:58","extension":"jpg","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":229871,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eOverview of the fsurgery:\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e(a-b) A skin incision was made over the right femur.\u003c/p\u003e\n\u003cp\u003e(c) An external fixator was fixed in the femur.\u003c/p\u003e\n\u003cp\u003e(d-e) 5-mm segmental defect was created and a polymethyl methacrylate spacer was placed in each defect. *In Group D, the defect site was closed without placement of a spacer.\u003c/p\u003e\n\u003cp\u003e(f) A membrane has formed on the surface of the spacer, and Incise the membrane and remove the spacer.\u003c/p\u003e\n\u003cp\u003e(g) After spacer removal. In Group A, the wound is closed without grafting.\u003c/p\u003e\n\u003cp\u003e(h) In Group B, artificial bone is transplanted.\u003c/p\u003e","description":"","filename":"Figure1.jpg","url":"https://assets-eu.researchsquare.com/files/rs-8741751/v1/7b631925841bc91b4559165a.jpg"},{"id":103050360,"identity":"b436d6c4-bef8-4164-914d-c19f9fa35fd8","added_by":"auto","created_at":"2026-02-20 07:49:37","extension":"jpg","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":179253,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eOverview of the iliac crest bone graft harvesting:\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e(a) A skin incision was made along the dorsal midline region.\u003c/p\u003e\n\u003cp\u003e(b) Dissect the gluteus maximus muscle from the dorsal midline.\u003c/p\u003e\n\u003cp\u003e(c) Separate the dorsomedial and dorsolateral fibers of the coccygeus muscle to expose the iliac crest.\u003c/p\u003e\n\u003cp\u003e(d) Bilateral iliac crests and the crushed cancellous bone\u003c/p\u003e\n\u003cp\u003e(e) Autologous cancellous bone is transplanted after removal of the PMMA spacer.\u003c/p\u003e","description":"","filename":"Figure2.jpg","url":"https://assets-eu.researchsquare.com/files/rs-8741751/v1/873954adb82aedf979b3721f.jpg"},{"id":103049994,"identity":"0c61bffe-6611-4bf7-a90b-f5c63ddeffca","added_by":"auto","created_at":"2026-02-20 07:47:35","extension":"jpg","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":166104,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eHistological features of the induced membrane after spacer removal:\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e(a) Histological section of the femur immediately after spacer removal (magnification ×2). The outlined area indicates the induced membrane tissue.\u003c/p\u003e\n\u003cp\u003e(b) High-magnification view (×20) of the membrane showing a characteristic bilayer structure composed of an outer layer rich in vascular components and an inner layer consisting primarily of dense fibrous tissue.\u003c/p\u003e\n\u003cp\u003e(c) Higher-magnification image (×40) focusing on the outer layer of the membrane.\u003c/p\u003e","description":"","filename":"Figure3.jpg","url":"https://assets-eu.researchsquare.com/files/rs-8741751/v1/9d301bba6c97c42f8e1647b1.jpg"},{"id":103014836,"identity":"401ec8b7-c708-496f-9672-093e790abe31","added_by":"auto","created_at":"2026-02-19 16:15:59","extension":"jpg","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":192247,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eComputed tomography evaluation of bone formation:\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e(a) Serial computed tomography (CT) images of a representative rat from Group C (autologous iliac cancellous bone graft) at consecutive time points.\u003c/p\u003e\n\u003cp\u003e(b) Representative X-ray CT images of rats in Group C at 8 weeks post-bone grafting.\u003c/p\u003e\n\u003cp\u003e(c) Serial CT images of representative rats from Groups A, B, C, and D at consecutive time points.\u003c/p\u003e\n\u003cp\u003e(d) Mean CT values at the graft site in Groups A (without graft) and C (cancellous bone graft) at each time point. Data are presented as mean ± standard deviation (SD).\u003c/p\u003e\n\u003cp\u003e(e) Proportion of newly formed bone (CT value \u0026gt; +1000) within the region of interest at each time point.\u003c/p\u003e\n\u003cp\u003e*Significant differences between Groups A and C (p \u0026lt; 0.05).\u003c/p\u003e","description":"","filename":"Figure4.jpg","url":"https://assets-eu.researchsquare.com/files/rs-8741751/v1/647bb055780e974e088c433b.jpg"},{"id":103014844,"identity":"24091b00-4868-482d-a8f7-d163bbfe2bf2","added_by":"auto","created_at":"2026-02-19 16:16:01","extension":"jpg","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":260298,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eHistological evaluation of longitudinal femoral sections:\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e(a) Longitudinal sections of rat femora following formalin fixation and decalcification.\u003c/p\u003e\n\u003cp\u003e(b) Representative hematoxylin and eosin (H\u0026amp;E)–stained histological sections corresponding to (a), shown at low magnification (magnification ×2).\u003c/p\u003e\n\u003cp\u003e(c) The grafted region (outlined by a box) was evaluated. Representative images from Groups A, B, C, and D at each time point are presented in a tabulated format (magnification ×4).\u003c/p\u003e","description":"","filename":"Figure5.jpg","url":"https://assets-eu.researchsquare.com/files/rs-8741751/v1/8f8458815b49b072d0a86f5e.jpg"},{"id":103014827,"identity":"bf7a2932-3122-4626-ae76-9ddddaa8b44a","added_by":"auto","created_at":"2026-02-19 16:15:58","extension":"jpg","order_by":6,"title":"Figure 6","display":"","copyAsset":false,"role":"figure","size":286377,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eHistological findings at the grafted site in Group C:\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e(a, b) At 4 weeks after grafting, osteoblasts infiltrating from the induced membrane toward the bone defect are observed, forming two to three cellular layers (arrows).\u003c/p\u003e\n\u003cp\u003e(c, d) At 6 weeks after grafting, progressive bone sclerosis is evident, and continuous osteoblast infiltration from the membrane is observed.\u003c/p\u003e\n\u003cp\u003e(e, f) At 8 weeks after grafting, bone maturation is apparent, accompanied by a decreasing trend in osteoblast infiltration from the induced membrane.\u003c/p\u003e\n\u003cp\u003eOriginal magnifications: (a, c, e) ×4; (b, d, f) ×20.\u003c/p\u003e","description":"","filename":"Figure6.jpg","url":"https://assets-eu.researchsquare.com/files/rs-8741751/v1/8c056d09f891acce6d2792f1.jpg"},{"id":103014828,"identity":"c76b969f-79f7-4a38-b1d2-d25fa359bc5a","added_by":"auto","created_at":"2026-02-19 16:15:58","extension":"jpg","order_by":7,"title":"Figure 7","display":"","copyAsset":false,"role":"figure","size":112147,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eCD31 immunohistochemical analysis:\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e(a–d) Representative immunohistochemical images of CD31 staining, a marker for vascular endothelial cells, in Group C and Group D.\u003c/p\u003e\n\u003cp\u003e(a, b) Group C at low magnification (×4) and high magnification (×40), respectively.\u003c/p\u003e\n\u003cp\u003e(c, d) Group D at low magnification (×4) and high magnification (×40), respectively.\u003c/p\u003e\n\u003cp\u003eThe areas enclosed by solid squares in (a) and (c) are shown at higher magnification in (b) and (d), respectively. The dotted square indicates the grafted region.\u003c/p\u003e\n\u003cp\u003e(e) Quantitative analysis of the total area of CD31-positive vessels at each time point. Data are presented as mean ± standard deviation (SD).\u003c/p\u003e\n\u003cp\u003e*Significant differences between Groups C and D (p \u0026lt; 0.05).\u003c/p\u003e","description":"","filename":"Figure7.jpg","url":"https://assets-eu.researchsquare.com/files/rs-8741751/v1/02db2399e216f16a8b7b105c.jpg"},{"id":103014832,"identity":"7744d029-c5b1-4503-b255-4934ebaf7d36","added_by":"auto","created_at":"2026-02-19 16:15:59","extension":"jpg","order_by":8,"title":"Figure 8","display":"","copyAsset":false,"role":"figure","size":63125,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eGene expression analysis by real-time RT-PCR:\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eRelative expression levels of BMP-2, TGF-β, and VEGF among the different groups at different time points of implantation determined using real-time RT-PCR.\u003c/p\u003e\n\u003cp\u003eThe data represent the mean value.\u003c/p\u003e\n\u003cp\u003e*Significant differences between the groups (p \u0026lt; 0.05).\u003c/p\u003e","description":"","filename":"Figure8.jpg","url":"https://assets-eu.researchsquare.com/files/rs-8741751/v1/194ce4d8aa07288dde1bcc48.jpg"},{"id":104714777,"identity":"09226367-bbb4-4c5f-a0a5-833f73af4def","added_by":"auto","created_at":"2026-03-16 10:58:19","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":2167033,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-8741751/v1/77bd807b-ddbe-4cb2-ba7e-f63c24bccd0c.pdf"}],"financialInterests":"No competing interests reported.","formattedTitle":"A rat model of the induced membrane technique using autologous iliac cancellous bone graft","fulltext":[{"header":"Introduction","content":"\u003cp\u003eSegmental bone defects arise from high-energy trauma, infection, and tumor resection, and remain a major clinical challenge in orthopedic surgery due to their complexity and the need for multiple surgical procedures, which significantly impair patients\u0026rsquo; quality of life\u003csup\u003e[\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e, \u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e]\u003c/sup\u003e. Among various reconstructive options, the induced membrane technique (IMT), also known as the Masquelet technique, has become increasingly adopted two-stage procedure for treating large bone defects and recalcitrant nonunions\u003csup\u003e[\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e, \u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e]\u003c/sup\u003e. Compared with alternative strategies such as vascularized bone grafting or distraction osteogenesis\u003csup\u003e[\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e, \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e]\u003c/sup\u003e, IMT offers advantages in technical simplicity and is theoretically less influenced by defect size with respect to healing time\u003csup\u003e[\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e]\u003c/sup\u003e. Despite its widespread use, several aspects of IMT\u0026mdash;including membrane biology, graft selection, and optimal surgical parameters\u0026mdash;remain subjects of debate, in part because many of these factors are difficult to investigate in human clinical settings. Consequently, a variety of animal models have been developed to characterize induced membrane formation and to evaluate how procedural variables affect subsequent bone regeneration\u003csup\u003e[\u003cspan additionalcitationids=\"CR9\" citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e]\u003c/sup\u003e.\u003c/p\u003e \u003cp\u003eAutologous cancellous bone (ACB) remains the gold standard graft material for fracture reconstruction, delayed unions, and nonunions because of its inherent osteogenic, osteoinductive, and osteoconductive properties\u003csup\u003e[\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e]\u003c/sup\u003e. ACB contains osteoblasts, mesenchymal stem cells, growth factors, and an interconnected trabecular scaffold that facilitates vascular ingrowth and cellular infiltration, making it a reliable option for both acute and reconstructive trauma surgery\u003csup\u003e[\u003cspan additionalcitationids=\"CR13 CR14 CR15\" citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e]\u003c/sup\u003e. In contrast, commonly used alternatives in small-animal research\u0026mdash;including deproteinized bovine bone, heat-treated porcine bone, or freeze-dried allograft harvested from donor rats\u0026mdash;serve primarily as osteoconductive scaffolds and lack the cellular and molecular components that confer the biological activity of fresh autograft\u003csup\u003e[\u003cspan additionalcitationids=\"CR18\" citationid=\"CR17\" class=\"CitationRef\"\u003e17\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e]\u003c/sup\u003e. These substitutes avoid the need for invasive bone harvesting but do not accurately replicate the clinical situation in which autograft is used. Moreover, models requiring donor animals increase cost and complexity.\u003c/p\u003e \u003cp\u003eAlthough effective ACB models have been established in medium and large animals, there is a lack of small animal models that faithfully reproduce the clinical environment of autologous bone grafting\u003csup\u003e[\u003cspan additionalcitationids=\"CR21 CR22\" citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e]\u003c/sup\u003e. Small animal models remain essential because they enable detailed investigation of tissue-level and molecular mechanisms, facilitate the development of new biomaterials, cellular therapies, and surgical strategies, and allow studies with larger sample sizes at lower cost. Our group previously established a rat model of autologous iliac crest bone harvesting and transplantation, demonstrating that reproducible autograft collection and successful graft incorporation can be achieved in a small-animal setting\u003csup\u003e[\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e]\u003c/sup\u003e. Building upon this validated platform, we sought to determine whether the IMT could similarly be performed using true autologous iliac crest grafts in rats\u0026mdash;a clinically relevant but previously unreported approach.\u003c/p\u003e \u003cp\u003eTherefore, the purpose of this study was to develop and characterize a rat IMT model using autologous iliac crest cancellous bone (AICB) grafts. This model aims to more closely replicate the clinical conditions of autograft-based IMT and to provide a robust experimental framework for future studies investigating graft biology, membrane function, and novel therapeutic strategies for segmental bone defects.\u003c/p\u003e"},{"header":"Results","content":"\u003cp\u003eHistological evaluation of femoral specimens harvested two weeks after the first surgery revealed the presence of a membrane-like structure surrounding the cavity previously occupied by the spacer (Fig.\u0026nbsp;3a). The membrane exhibited a characteristic bilayer architecture, consisting of an inner layer composed of dense fibrous tissue and an outer layer composed of loose connective tissue with abundant microvessels. These histological features were consistent with previously reported pathological characteristics of the induced membrane\u003csup\u003e[\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e]\u003c/sup\u003e (Fig.\u0026nbsp;3b,c).\u003c/p\u003e \u003cp\u003eRepresentative longitudinal CT images of rats from Groups A\u0026ndash;D at each time point are shown in Fig.\u0026nbsp;4 (Fig.\u0026nbsp;4a\u0026ndash;c). The bone union rates at the graft site were 0% in Group A, 6.7% (1 rat) in Group B, 66.7% (10 rats) in Group C, and 20.0% (3 rats) in Group D. Group C demonstrated the highest bone union rate, with changes suggestive of new bone formation observed as early as two weeks after grafting, becoming more pronounced over time. Bone formation was predominantly observed on the medial side of the femur, whereas union on the lateral side tended to be delayed. In Groups A and B, bone formation at the defect site was minimal; the bone edges became rounded and regressed over time, and the synthetic bone graft in Group B showed a tendency toward resorption.\u003c/p\u003e \u003cp\u003eComparative CT analysis between Groups A and C demonstrated distinct temporal patterns of bone formation (Fig.\u0026nbsp;4d,e). In Group C, CT values increased progressively over time, and a significant overall temporal change was observed (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05). In Group A, CT values also showed a significant temporal change, characterized by an initial increase followed by slight decrease at later time points (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05). At all time points, CT values in Group C were significantly higher than those in Group A (p\u0026thinsp;\u0026lt;\u0026thinsp;0.05). Similar temporal trends and significant intergroup differences were also observed in the proportion of newly formed bone, consistent with the findings for CT values.\u003c/p\u003e \u003cp\u003eHistological sections stained with hematoxylin and eosin from Groups A\u0026ndash;D are shown in Fig.\u0026nbsp;5. In Groups C and D, sagittal sections obtained two weeks after grafting confirmed that the transplanted cancellous bone was located within the bone defect. In Group C, the cancellous bone was progressively replaced by newly formed bone over time, and bridging of the defect by new bone was observed from four weeks onward. Notably, in Group C, infiltration of osteoblasts from the membrane surrounding the graft into the bone defect was observed in two to three cellular layers (Fig.\u0026nbsp;6a\u0026ndash;f). This finding was evident from four to eight weeks after grafting, although the degree of osteoblast infiltration tended to decrease at eight weeks as bone maturation progressed. In Group D, although some replacement of the grafted cancellous bone by new bone was observed, the extent of bone formation was less pronounced than in Group C, no membrane structure was present on the outer surface, and osteoblast infiltration was limited.\u003c/p\u003e \u003cp\u003eRepresentative histological images from Groups C and D at four weeks after grafting are shown in Fig.\u0026nbsp;7a and 7b. Quantitative analysis revealed that the CD31-positive total vascular area at the graft site was significantly greater in Group C than in Group D at two, and six weeks after grafting (\u003cem\u003ep\u003c/em\u003e\u0026thinsp;\u0026lt;\u0026thinsp;0.05) (Fig.\u0026nbsp;7c).\u003c/p\u003e \u003cp\u003eTo evaluate the molecular effects of induced membrane\u0026ndash;assisted bone grafting, real-time RT-PCR analysis was performed (Fig.\u0026nbsp;8). BMP-2, TGF-β, and VEGF were selected as representative markers of osteogenic differentiation, extracellular matrix production, and angiogenesis, respectively. In Group C, VEGF expression was significantly higher than in Group A at four weeks after grafting (\u003cem\u003ep\u003c/em\u003e\u0026thinsp;\u0026lt;\u0026thinsp;0.05). Expression levels of BMP-2 and TGF-β at four weeks were significantly higher in Groups C and D than in Group A. Although expression levels of all three markers tended to be higher in Group C than in Group D, no statistically significant differences were observed between these two groups.\u003c/p\u003e"},{"header":"Discussion/Conclusion","content":" \u003cdiv id=\"Sec3\" class=\"Section2\"\u003e \u003cp\u003eThe induced membrane technique (IMT) has been widely adopted as a treatment for segmental bone defects and nonunion. However, several aspects of this technique\u0026mdash;including the optimal timing of grafting, selection of spacer materials, choice of graft material, and the potential role of adjunctive factors\u0026mdash;remain subjects of ongoing debate in clinical practice. Accordingly, animal models used to investigate IMT should also be refined to better reflect clinical conditions before translation to human application. Among the various factors influencing the IMT, graft selection represents one of the most critical determinants of treatment success. Autograft, allograft, and synthetic bone substitutes are commonly used options. Previous clinical studies have reported IMT procedures using mixtures containing up to 64% allograft combined with autograft\u003csup\u003e[\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e]\u003c/sup\u003e. Nevertheless, a 3:1 ratio of autograft to allograft has traditionally been the most widely employed combination\u003csup\u003e[\u003cspan additionalcitationids=\"CR29 CR30\" citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e]\u003c/sup\u003e, and autologous bone grafting remains the clinical gold standard. Despite this consensus, there is no uniform agreement regarding the optimal donor site for graft harvesting, either in clinical practice or in animal experiments.\u003c/p\u003e \u003cp\u003eHarvesting graft material from the iliac crest is one of the most common clinical approaches. Gens et al. reported that, in a rat bone graft model, iliac crest\u0026ndash;derived samples exhibited qualitatively higher alkaline phosphatase (ALP) levels than samples harvested from other skeletal sites, suggesting a higher concentration of osteoprogenitor cells\u003csup\u003e[\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e]\u003c/sup\u003e. Similarly, studies of human bone grafts have demonstrated that the mean concentration of osteoblastic progenitor cells is higher in iliac crest grafts than in tibial grafts\u003csup\u003e[\u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e]\u003c/sup\u003e. In addition, from a functional perspective, harvesting bone from the iliac crest is expected to have less impact on postoperative locomotion than harvesting from the extremities. Another major advantage of the iliac crest is the ability to obtain a relatively large amount of cancellous bone. This point is particularly important in small animals such as rats, in which long bones consist predominantly of cortical bone and contain only minimal cancellous components. Although rat models of autologous bone grafting have been previously reported, most have relied on cortical bone as the graft material\u003csup\u003e[\u003cspan additionalcitationids=\"CR35\" citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e]\u003c/sup\u003e. While autologous cortical grafts provide high mechanical strength and immediate structural support, they exhibit limited osteoinductive potential compared with cancellous bone\u003csup\u003e[\u003cspan additionalcitationids=\"CR38\" citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e]\u003c/sup\u003e. Consequently, cancellous bone is generally more effective for promoting bone union and is preferentially used in clinical bone grafting.\u003c/p\u003e \u003cp\u003eDespite these advantages, a systematic review analyzing 47 studies investigating IMT in animal models found that only seven studies harvested autologous grafts from the iliac crest, all of which involved rabbits or sheep; no rat iliac crest bone graft models were included\u003csup\u003e[\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e]\u003c/sup\u003e. This lack of rat models likely reflects the absence of established techniques for safely and reliably harvesting iliac bone from small animals. Many previously reported rat IMT models have therefore relied on allografts or synthetic bone substitutes\u003csup\u003e[\u003cspan citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e, \u003cspan additionalcitationids=\"CR9\" citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e\u0026ndash;\u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e, \u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e]\u003c/sup\u003e, and bone grafting using caudal vertebrae has been particularly common. However, the rat tail plays an important role in thermoregulation and temperature sensation, and tail amputation in live animals is not recommended, raising animal welfare concerns\u003csup\u003e[\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e, \u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e41\u003c/span\u003e]\u003c/sup\u003e. Moreover, caudal vertebrae are composed primarily of cortical bone, with only minimal cancellous bone available for harvesting. For these reasons, the iliac crest appears to offer clear advantages over the tail as a donor site for bone grafting.\u003c/p\u003e \u003cp\u003eTo our knowledge, the present study is the first to establish a rat IMT model using autologous iliac cancellous bone grafting. Using this approach, an adequate volume of cancellous bone was safely harvested, and the surgical procedures closely replicated the clinical IMT. Histological evaluation following spacer removal confirmed the presence of a characteristic bilayer membrane structure consistent with previously described features of the induced membrane, indicating successful membrane formation. In the comparative analysis, bone union was not achieved in Group A (no graft) or Group B (artificial bone graft), whereas successful bone union was observed in 67% of rats in Group C (AICB with IMT). These findings indicate that cancellous bone grafting plays a favorable role in promoting bone healing. Furthermore, the higher bone union rate in Group C compared with Group D (without induced membrane) suggests that the presence of an induced membrane provides an additional biological advantage beyond AICB alone. Collectively, these results demonstrate that the IMT using AICB yields the most consistent and stable bone union outcomes among the models tested, supporting its validity as a reproducible experimental platform.\u003c/p\u003e \u003cp\u003eCT evaluation revealed that in some cases, bony continuity was established as early as two weeks after grafting. Quantitative analysis further demonstrated that in Group C, both mean CT values and the proportion of newly formed bone increased progressively over time and were significantly higher than those observed in Group A. In contrast, these parameters showed a slight decline at eight weeks in Group A, which may reflect changes associated with nonunion. Bone union tended to occur earlier on the medial side of the femur, a finding that may reflect not only differences in surrounding soft tissue volume but also the incision of the lateral membrane during spacer removal.\u003c/p\u003e \u003cp\u003eHistological analyses provided further evidence of osteoblast infiltration from the induced membrane into the bone defect. This influx of osteoblasts was most prominent at four weeks after grafting and gradually decreased by eight weeks as bone maturation progressed. These observations support the concept that the induced membrane actively contributes to osteogenesis. In contrast, no membrane-like structure was histologically identified in Group D, in which AICB grafting was performed without spacer placement. Quantitative analyses demonstrated that angiogenesis, as assessed by immunohistochemistry, was significantly greater in Group C, and real-time RT-PCR revealed that VEGF expression was significantly upregulated only in Group C compared with Group A. Thus, although angiogenic activity was clearly enhanced in the IMT-treated Group C, no marked differences in BMP-2 or TGF-β expression were observed between Groups C and D. This finding may be attributable to the biological effects inherent to cancellous bone grafting itself. Nevertheless, the substantial difference in union rates between the two groups suggests that the physical effects associated with membrane formation, such as prevention of graft dispersion and protection against external contamination, may have also played a role. Indeed, many of the rats that failed to achieve bone union exhibited, to some extent, signs of infection. Taken together, these findings suggest that induced membrane formation using the IMT provides a more favorable biological and mechanical environment for bone healing than cancellous bone grafting alone.\u003c/p\u003e \u003cp\u003eSeveral limitations of this model should be acknowledged. First, the volume of cancellous bone that can be harvested from the iliac crest is limited. Hamada et al. reported a mean harvested cancellous bone volume of 73.8\u0026thinsp;\u0026plusmn;\u0026thinsp;5.5 mm\u0026sup3; in a rat autologous cancellous bone graft model\u003csup\u003e[\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e]\u003c/sup\u003e, corresponding to a graft length of approximately 5.8 mm with a 4-mm diameter or 10 mm with a 3-mm diameter. Therefore, for large bone defects exceeding 10 mm, the use of AICB alone may be insufficient in rats of this size. Second, in the present study, the duration of spacer placement was set at two weeks. Although this interval was slightly shorter than that reported in many previous studies\u003csup\u003e[\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e, \u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e, \u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e]\u003c/sup\u003e, histological evaluation of the membrane immediately before grafting confirmed a characteristic bilayer structure consistent with prior literature. Moreover, multiple findings suggested that this membrane exerted a favorable effect on subsequent bone healing, indicating that a two-week waiting period was sufficient in this model. Nevertheless, the optimal duration of spacer placement remains to be determined, and it is possible that the selected interval was not the most favorable. Further studies directly comparing different waiting periods are warranted to clarify the optimal timing for grafting in IMT. Third, external fixation was used for stabilization in this study. This choice was made because IMT is often applied clinically to traumatic bone defects or cases complicated by infection, in which external fixation is frequently preferred over internal fixation. However, maintaining strict postoperative immobilization in rats is challenging, and loosening of fixation pins due to contact with the cage or pin tract infections may have contributed to nonunion in some cases. Although IMT models using internal fixation with plates have been reported\u003csup\u003e[\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e, \u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e]\u003c/sup\u003e, the use of external fixation in small animals may increase the susceptibility to fixation-related complications, which should be considered when interpreting the results.\u003c/p\u003e \u003cp\u003eIn conclusion, we successfully established a reproducible rat IMT model using autologous iliac cancellous bone grafting. This model closely replicates the clinical IMT and provides a valuable experimental platform for investigating the biological mechanisms underlying IMT and for developing novel therapeutic strategies for the treatment of segmental bone defects.\u003c/p\u003e \u003c/div\u003e"},{"header":"Methods","content":"\u003cp\u003eAnimal experiments:\u003c/p\u003e \u003cp\u003e All animal experiments were approved by the Animal Experimentation Committee of Kanazawa University (Approval No. AP-204176). All procedures were performed in accordance with the institutional guidelines, the ARRIVE guidelines (PLoS Biol 8(6): e1000412, 2010), and the relevant regulations. Anesthesia and euthanasia protocols adhered to the American Veterinary Medical Association (AVMA) Guidelines for the Euthanasia of Animals (2020). The study does not involve endangered or CITES-listed species.\u003c/p\u003e \u003cp\u003eHealthy male Sprague\u0026ndash;Dawley rats (12 weeks old, 350\u0026ndash;400 g) were obtained for this study. The animals were housed individually under specific pathogen-free conditions with a 12-h light/dark cycle and given free access to food and water. All rats were acclimatized to the laboratory environment for one week prior to experimentation. For all procedures, anesthesia was induced by intraperitoneal injection of medetomidine (0.15 mg/kg), midazolam (2 mg/kg), and butorphanol (2.5 mg/kg). At the end of the study, the rats were euthanized by intraperitoneal administration of pentobarbital (450 mg/kg).\u003c/p\u003e \u003cp\u003eIMT procedure:\u003c/p\u003e \u003cp\u003eAll rats underwent two surgical procedures. Figure\u0026nbsp;1 illustrates the overall design of the first-stage procedure. The femoral defect model was adapted from previously reported methods in which nonunion was consistently observed without fixation implants\u003csup\u003e[\u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e, \u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e]\u003c/sup\u003e. Each rat was placed in the prone position on the operating table. A longitudinal lateral skin incision was made over the right femur, and the quadriceps and hamstring muscles were split and retracted (Fig.\u0026nbsp;1a,b). After pre-drilling with a 1.4-mm Kirschner wire, an external fixator (Meira Co., Nagoya, Japan) was mounted to the femur using four self-tapping pins (1.6 mm in diameter; Japan Medicalnext Co., Osaka, Japan) (Fig.\u0026nbsp;1c). Two osteotomies were created with a bone saw between the second and third pins to generate a 5-mm segmental bone defect (Fig.\u0026nbsp;1d). A polymethyl methacrylate (PMMA) spacer was inserted into each defect (Fig.\u0026nbsp;1e). The muscle layer, subcutaneous tissue, and skin were closed with simple interrupted sutures.\u003c/p\u003e \u003cp\u003eTo investigate bone regeneration, all rats underwent a second surgery two weeks after the initial procedure. The rats were positioned identically to the first surgery, and the previous lateral skin incision was reopened. After confirming the presence of membrane-like tissue (induced membrane) overlying the spacer (Fig.\u0026nbsp;1f), the membrane was incised with a scalpel, and the spacer was removed (Fig.\u0026nbsp;1g). The membrane cavity between the bone ends was then filled according to one of the following three conditions: no graft material (Group A), a β-TCP bone substitute block (Group B), or AICB (Group C) (Fig.\u0026nbsp;1h, 2e). The incised membrane was sutured, and the wound was closed in layers.\u003c/p\u003e \u003cp\u003eAdditionally, to determine whether placement of a cement spacer is essential for IMT, a separate group was created in which the bone defect was generated during the first surgery but closed without inserting a spacer. During the second surgery, these rats received AICB grafting in the same manner as Group C (Group D).\u003c/p\u003e \u003cp\u003eAutologous iliac bone graft model:\u003c/p\u003e \u003cp\u003eAICB harvesting was performed according to the procedure described by Hamada et al.\u003csup\u003e[\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e]\u003c/sup\u003e. A longitudinal midline skin incision of approximately 4 cm was made over the highest point of the bilateral iliac crests (Fig.\u0026nbsp;2a). The gluteus maximus was elevated from the dorsal midline, and the dorsomedial and dorsolateral fibers of the coccygeus muscle were separated to expose the iliac crest (Fig.\u0026nbsp;2b,c). Care was taken to avoid injury to deep neurovascular structures while performing the osteotomy, and each iliac crest was harvested bilaterally. Using a scalpel, cancellous bone was separated from the cortical shell, and the morselized cancellous bone was packed into a cylindrical mold and compressed to form a 5-mm-thick block (Fig.\u0026nbsp;2d). In group C, the AICB block was transplanted into the femoral bone defect. (Fig.\u0026nbsp;2e). The submuscular and gluteal layers were closed with simple interrupted sutures, followed by closure of the skin.\u003c/p\u003e \u003cp\u003eImaging Evaluation:\u003c/p\u003e \u003cp\u003eComputed tomography (CT) imaging was performed using a Latheta LCT-200 system (Hitachi Aloka Medical, Tokyo, Japan). Scans were obtained immediately after grafting and at 2, 4, 6, and 8 weeks postoperatively to evaluate longitudinal bone formation and the presence of bone union. Fifteen rats per group were assessed to determine the bone union rate. Coronal CT images were used to define the femoral axis. To exclude pre-existing cortical bone from the analysis, bone formation was evaluated only within the central defect region, defined as a 5-mm long-axis \u0026times; 4-mm short-axis region of interest. CT values were calibrated using water (CTw\u0026thinsp;=\u0026thinsp;0) and air (CTa\u0026thinsp;=\u0026thinsp;\u0026minus;\u0026thinsp;1000) as reference standards, and regions with CT values exceeding\u0026thinsp;+\u0026thinsp;1000 were extracted as newly formed bone. Image analysis was performed using Synapse VINCENT software (Fujifilm, Tokyo, Japan). For Groups A and C, rats at each time point (N\u0026thinsp;=\u0026thinsp;5 per group) were further analyzed to calculate the mean CT value within the region of interest and the proportion of newly formed bone. Temporal changes and intergroup comparisons were subsequently evaluated.\u003c/p\u003e \u003cp\u003eHistological Evaluation:\u003c/p\u003e \u003cp\u003eAt each designated time point, harvested femora were fixed in 10% neutral-buffered formalin with surrounding soft tissues and the external fixator left in situ. The specimens were then dehydrated through a graded ethanol series (70%, 80%, 90%, and 100%). After decalcification in a 10% sodium citrate\u0026ndash;formic acid solution, the external fixators were removed. The specimens were embedded in paraffin, sectioned in the sagittal plane, and stained with hematoxylin and eosin (H\u0026amp;E). Histological evaluation was performed using a BZ-9000 optical microscope (Keyence, Osaka, Japan).\u003c/p\u003e \u003cp\u003eSpecimens were collected at 2, 4, 6, and 8 weeks after grafting in all groups to assess histological bone formation and its relationship with the induced membrane. In addition, specimens from Groups C and D were collected immediately after grafting to evaluate the presence or absence of the induced membrane prior to transplantation. In Group C, membrane formation consistent with previous reports was anticipated, whereas no membrane formation was expected in Group D.\u003c/p\u003e \u003cp\u003eImmunohistochemistry:\u003c/p\u003e \u003cp\u003eTo assess angiogenesis at the graft site, immunohistochemical staining for endothelial cells was performed using an anti-CD31 antibody (Abcam, Cambridge, UK; ab28364; 1:50 dilution). Tissue sections were incubated at room temperature for 15 minutes in Liberate Antibody Binding Solution (LAB solution; Polysciences, Philadelphia, PA, USA), followed by blocking with Protein Block Serum-Free (Dako, Glostrup, Denmark) and peroxidase-blocking solution containing hydrogen peroxide (Dako, Glostrup, Denmark) for 10 minutes each.\u003c/p\u003e \u003cp\u003eSections were then incubated overnight at 4\u0026deg;C with the primary anti-CD31 antibody diluted in Tris-HCl buffer containing stabilizing proteins and 0.015 mol/L sodium azide (Dako Antibody Diluent; Dako, Glostrup, Denmark). After incubation with a secondary antibody (Dako REAL EnVision/HRP, Rabbit/Mouse; Dako, Glostrup, Denmark) for 30 minutes at room temperature, immunoreactivity was visualized using DAB chromogen (Dako REAL DAB+ Chromogen; Dako, Glostrup, Denmark). The reaction was terminated by rinsing the slides with deionized water once an optimal signal-to-noise ratio was achieved.\u003c/p\u003e \u003cp\u003eTo specifically investigate angiogenesis associated with induced membrane formation, analyses were performed in Groups C and D. Vascular area at the graft site was measured at 2, 4, 6, and 8 weeks after grafting. In each group, five randomly selected high-power fields (\u0026times;40) from five nonconsecutive sections were quantified. Measurement of vascular area and image analysis were performed using a BZ-X800 all-in-one fluorescence microscope with time-lapse functionality (Keyence, Osaka, Japan).\u003c/p\u003e \u003cp\u003eReal-Time RT-PCR Analysis:\u003c/p\u003e \u003cp\u003eTotal RNA was extracted from graft-site tissues using the NucleoSpin RNA II kit (Takara Bio, Otsu, Japan). To evaluate the effects of bone grafting and induced membrane formation on angiogenesis-related gene expression, analyses were conducted using samples from Groups A, C, and D. Harvested tissues were mechanically homogenized using a syringe (n\u0026thinsp;=\u0026thinsp;5 per group at each time point). RNA concentration and purity were assessed using a UV\u0026ndash;visible spectrophotometer (NanoDrop Lite; Thermo Scientific, Waltham, MA, USA).\u003c/p\u003e \u003cp\u003eFor real-time RT-PCR, 6 \u0026micro;g of total RNA was reverse-transcribed into complementary DNA (cDNA) using the RevertAid First Strand cDNA Synthesis Kit (Thermo Scientific, Waltham, MA, USA) and a thermal cycler (T100\u0026trade; Thermal Cycler; Bio-Rad, Hercules, CA, USA). Quantitative PCR was performed using SYBR Green PCR Master Mix (Applied Biosystems, Foster City, CA, USA) on an ABI Prism 7900 system (Applied Biosystems, Foster City, CA, USA), in accordance with the manufacturer\u0026rsquo;s instructions.\u003c/p\u003e \u003cp\u003ePrimers for rat glyceraldehyde-3-phosphate dehydrogenase (GAPDH), bone morphogenetic protein-2 (BMP-2), vascular endothelial growth factor (VEGF), and transforming growth factor-β1 (TGF-β1) were purchased from Hokkaido System Science Co., Ltd. (Japan), and their sequences are listed in Table\u0026nbsp;1. The amplification parameters were an initial 95˚C incubation step for 15 minutes, followed by 20 amplification cycles of 94˚C for 15 seconds, 60˚C for 30 seconds, and 72˚C for 30 seconds. The reactions ended with a 72˚C extension step for 7 minutes, followed by storage at 4˚C overnight. The expression levels of each target gene were calculated relative to the level of GAPDH for each sample.\u003c/p\u003e \u003cdiv id=\"Sec5\" class=\"Section2\"\u003e \u003ch2\u003eStatistical Analysis:\u003c/h2\u003e \u003cp\u003eData are presented as mean\u0026thinsp;\u0026plusmn;\u0026thinsp;standard deviation (SD). Intergroup comparisons of CT values, CD31-positive vascular area, and relative gene expression levels were performed using the Mann\u0026ndash;Whitney U test,, as the data did not meet assumptions of normality and the sample size in each group was small (n\u0026thinsp;=\u0026thinsp;5) and nonparametric analysis was considered more appropriate. Temporal changes in CT values were evaluated using the Kruskal\u0026ndash;Wallis test to assess overall trends over time. A p-value of \u0026lt;\u0026thinsp;0.05 was considered statistically significant.\u003c/p\u003e \u003c/div\u003e"},{"header":"Declarations","content":"\u003ch2\u003eFunding Declaration:\u003c/h2\u003e \u003cp\u003eThis work was supported by the National Mutual Insurance Federation of Agricultural Cooperatives (ZENKYOREN) in fiscal year 2021, the Shibuya Science Culture and Sports Foundation in fiscal year 2020, and by Japan Society for the Promotion of Science (KAKENHI) Grant-in-Aid for Scientific Research (C) in fiscal year from 2021 to 2023.\u003c/p\u003e\u003ch2\u003eAuthor Contribution\u003c/h2\u003e\u003cp\u003eK.S., H.M., and T.H. conceptualized and designed the study. K.S. primarily performed the experiments. Data interpretation and discussion were conducted by all authors, including T.H. and Y.N. K.S. performed data curation, formal analysis, and drafted the original manuscript. H.M. and S.D. supervised the study. K.S. acquired the funding. All authors reviewed and approved the final manuscript.\u003c/p\u003e\u003ch2\u003eAcknowledgements:\u003c/h2\u003e \u003cp\u003eNot applicable\u003c/p\u003e\u003ch2\u003eData Availability\u003c/h2\u003e\u003cp\u003eThe datasets generated and/or analyzed during the current study are available from the corresponding author on reasonable request.\u003c/p\u003e\u003cp\u003eAdditional Information (including a Competing Interests Statement):\u003c/p\u003e\n\u003cp\u003eThe authors declare that they have no competing interests related to this work.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\u003cli\u003e\u003cspan\u003eSun, H. et al. The induced membrane technique in animal models: a systematic review. OTA. Int. 10; 5(1 Suppl): e176 (2022).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eSvedbom, A. et al. Osteoporosis in the European Union: a compendium of country-specific reports. \u003cem\u003eArch. Osteoporos.\u003c/em\u003e \u003cb\u003e8\u003c/b\u003e (1), 137 (2013).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePelissier, P. et al. Induced membranes secrete growth factors including vascular and osteoinductive factors and could stimulate bone regeneration. \u003cem\u003eJ. Orthop. Res.\u003c/em\u003e \u003cb\u003e22\u003c/b\u003e (1), 73\u0026ndash;79 (2004).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eTaylor, B. C., French, B. G., Fowler, T. T., Russell, J. \u0026amp; Poka, A. Induced membrane technique for reconstruction to manage bone loss. \u003cem\u003eJ. Am. Acad. Orthop. Surg.\u003c/em\u003e \u003cb\u003e20\u003c/b\u003e (3), 142\u0026ndash;150 (2012).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGlynou, S. P., Georgiannakis, A., Ardolino, D., Craxford, S. \u0026amp; Vris, A. Vascularised Fibula Transfer for Post-traumatic Critical Tibial Bone Defects: A Systematic Review. \u003cem\u003eStrategies Trauma. Limb Reconstr.\u003c/em\u003e \u003cb\u003e20\u003c/b\u003e (1), 37\u0026ndash;46 (2025).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eTsuchiya, H. \u0026amp; Tomita, K. Distraction osteogenesis for treatment of bone loss in the lower extremity. \u003cem\u003eJ. Orthop. Sci.\u003c/em\u003e \u003cb\u003e8\u003c/b\u003e (1), 116\u0026ndash;124 (2003).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMauffrey, C., Barlow, B. T. \u0026amp; Smith, W. Management of segmental bone defects. \u003cem\u003eJ. Am. Acad. Orthop. Surg.\u003c/em\u003e \u003cb\u003e23\u003c/b\u003e, 143\u0026ndash;153 (2015).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMcBride-Gagyi, S. et al. Altering spacer material affects bone regeneration in the Masquelet technique in a rat femoral defect. \u003cem\u003eJ. Orthop. Res.\u003c/em\u003e 23866. \u003cspan class=\"ExternalRef\"\u003e\u003cspan class=\"RefSource\"\u003e10.1002/jor\u003c/span\u003e\u003cspan address=\"10.1002/jor\" targettype=\"DOI\" class=\"RefTarget\"\u003e\u003c/span\u003e\u003c/span\u003e (2018).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGouron, R. et al. Osteoclasts and their precursors are present in the induced-membrane during bone reconstruction using the Masquelet technique. \u003cem\u003eJ. Tissue Eng. Regen Med.\u003c/em\u003e \u003cb\u003e11\u003c/b\u003e (2), 382\u0026ndash;389 (2017).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eTang, Q., Tong, M., Zheng, G., Shang, P. \u0026amp; Liu, H. Masquelet's induced membrane promotes the osteogenic differentiation of bone marrow mesenchymal stem cells by activating the Smad and MAPK pathways. \u003cem\u003eAm. J. Transl Res.\u003c/em\u003e \u003cb\u003e10\u003c/b\u003e (4), 1211\u0026ndash;1219 (2018).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGiannoudis, P. V., Dinopoulos, H. \u0026amp; Tsiridis, E. Bone substitutes: an update. \u003cem\u003eInjury 36 Suppl.\u003c/em\u003e 3: S20\u0026ndash;S27 (2005).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eCassidy, C. et al. Norian SRS cement compared with conventional fixation in distal radial fractures. A randomized study. \u003cem\u003eJ. Bone Joint Surg. Am.\u003c/em\u003e \u003cb\u003e85\u003c/b\u003e (11), 2127\u0026ndash;2137 (2003).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFinkemeier, C. G. Bone-grafting and bone-graft substitutes. \u003cem\u003eJ. Bone Joint Surg. Am.\u003c/em\u003e \u003cb\u003e84\u003c/b\u003e (3), 454\u0026ndash;464 (2002).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKelly, C. M. et al. The use of a surgical grade calcium sulfate as a bone graft substitute: Results of a multicenter trial. \u003cem\u003eClin. Orthop. Relat. Res.\u003c/em\u003e \u003cb\u003e382\u003c/b\u003e, 42\u0026ndash;50 (2001).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKhan, S. N. et al. The biology of bone grafting. \u003cem\u003eJ. Am. Acad. Orthop. Surg.\u003c/em\u003e \u003cb\u003e13\u003c/b\u003e (1), 77\u0026ndash;86 (2005).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRay, R. D. Vascularization of bone grafts and implants. \u003cem\u003eClin. Orthop. Relat. Res.\u003c/em\u003e \u003cb\u003e87\u003c/b\u003e, 43\u0026ndash;48 (1972).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFitch, R., Kerwin, S., Sinibaldi, K. R. \u0026amp; Newman, H. E. Bone autografts and allografts in dogs. \u003cem\u003eComp. Vet. Cont. Ed.\u003c/em\u003e \u003cb\u003e19\u003c/b\u003e (5), 558\u0026ndash;575 (1997).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMcLaughlin, R. M. \u0026amp; Roush, J. K. Autogenous cancellous and corticocancellous bone grafting. \u003cem\u003eVet. Med.\u003c/em\u003e \u003cb\u003e93\u003c/b\u003e, 1071\u0026ndash;1074 (1998).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eOrth, M. et al. BMP-2-coated mineral coated microparticles improve bone repair in atrophic non-unions. \u003cem\u003eEur. Cell. Mater.\u003c/em\u003e \u003cb\u003e33\u003c/b\u003e, 1\u0026ndash;12 (2017).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eAthanasiou, V. T. et al. Histological comparison of autograft, allograft-DBM, xenograft, and synthetic grafts in a trabecular bone defect: An experimental study in rabbits. \u003cem\u003eMed. Sci. Monit.\u003c/em\u003e \u003cb\u003e16\u003c/b\u003e (1), BR24\u0026ndash;31 (2010).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eFox, S. M. Cancellous bone grafting in the dog\u0026mdash;An overview. \u003cem\u003eJ. Am. Anim. Hosp. Assoc.\u003c/em\u003e \u003cb\u003e20\u003c/b\u003e, 840\u0026ndash;848 (1984).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKlaue, K. et al. Bone regeneration in long-bone defects: Tissue compartmentalisation? In vivo study on bone defects in sheep. \u003cem\u003eInjury\u003c/em\u003e \u003cb\u003e40\u003c/b\u003e (Suppl 4), S95\u0026ndash;102 (2009).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eRosenthal, A. H. \u0026amp; Buchman, S. R. Volume maintenance of inlay bone grafts in the craniofacial skeleton. \u003cem\u003ePlast. Reconstr. Surg.\u003c/em\u003e \u003cb\u003e112\u003c/b\u003e (3), 802\u0026ndash;811 (2003).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHamada, T., Matsubara, H., Hikichi, T., Shimokawa, K. \u0026amp; Tsuchiya, H. Rat model of an autologous cancellous bone graft. \u003cem\u003eSci. Rep.\u003c/em\u003e \u003cb\u003e11\u003c/b\u003e (1), 18001 (2021).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGlatt, V., Evans, C. H. \u0026amp; Matthys, R. Design, characterisation and in vivo testing of a new, adjustable stiffness, external fixator for the rat femur. \u003cem\u003eEur. Cell. Mater.\u003c/em\u003e \u003cb\u003e23\u003c/b\u003e, 289\u0026ndash;298 (2012).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHarada, N. et al. Bone regeneration in a massive rat femur defect through endochondral ossification achieved with chondrogenically differentiated MSCs in a degradable scaffold. \u003cem\u003eBiomaterials\u003c/em\u003e \u003cb\u003e35\u003c/b\u003e (27), 7800\u0026ndash;7810 (2014).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003ePesciallo, C. A., Garabano, G., Dainotto, T. \u0026amp; Ernst, G. Masquelet technique in post-traumatic infected femoral and tibial segmental bone defects. Union and reoperation rates with high proportions (up to 64%) of allograft in the second stage. \u003cem\u003eInjury\u003c/em\u003e \u003cb\u003e52\u003c/b\u003e (11), 3471\u0026ndash;3477 (2021).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMasquelet, A. C. \u0026amp; Begue, T. The concept of induced membrane for reconstruction of long bone defects. \u003cem\u003eOrthop. Clin. North. Am.\u003c/em\u003e \u003cb\u003e41\u003c/b\u003e (1), 27\u0026ndash;37 (2010).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMasquelet, A., Kanakaris, N. K., Obert, L., Stafford, P. \u0026amp; Giannoudis, P. V. Bone repair using the Masquelet technique. \u003cem\u003eJ. Bone Joint Surg. Am.\u003c/em\u003e \u003cb\u003e101\u003c/b\u003e (11), 1024\u0026ndash;1036 (2019).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eKarger, C., Kishi, T., Schneider, L., Fitoussi, F. \u0026amp; Masquelet, A. C. Treatment of posttraumatic bone defects by the induced membrane technique. \u003cem\u003eOrthop. Traumatol. Surg. Res.\u003c/em\u003e \u003cb\u003e98\u003c/b\u003e (1), 97\u0026ndash;102 (2012).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBaldwin, P. et al. Autograft, allograft, and bone graft substitutes: Clinical evidence and indications for use in the setting of orthopaedic trauma surgery. \u003cem\u003eJ. Orthop. Trauma.\u003c/em\u003e \u003cb\u003e33\u003c/b\u003e (4), 203\u0026ndash;213 (2019).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGens, L. et al. Surgical technique and comparison of autologous cancellous bone grafts from various donor sites in rats. \u003cem\u003eJ. Orthop. Res.\u003c/em\u003e \u003cb\u003e41\u003c/b\u003e (4), 834\u0026ndash;844 (2023).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eHyer, C. F. et al. Quantitative assessment of the yield of osteoblastic connective tissue progenitors in bone marrow aspirate from the iliac crest, tibia, and calcaneus. \u003cem\u003eJ. Bone Joint Surg.\u003c/em\u003e \u003cb\u003e95\u003c/b\u003e (14), 1312\u0026ndash;1316 (2013).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBosemark, P., Isaksson, H., McDonald, M. M., Little, D. G. \u0026amp; T\u0026auml;gil, M. Augmentation of autologous bone graft by a combination of bone morphogenic protein and bisphosphonate increased both callus volume and strength. \u003cem\u003eActa Orthop.\u003c/em\u003e \u003cb\u003e84\u003c/b\u003e (1), 106\u0026ndash;111 (2013).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eGouron, R. et al. Osteoclasts and their precursors are present in the induced-membrane during bone reconstruction using the Masquelet technique. \u003cem\u003eJ. Tissue Eng. Regen Med.\u003c/em\u003e \u003cb\u003e11\u003c/b\u003e (2), 382\u0026ndash;389 (2017).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMurakami, H., Nakasa, T., Ishikawa, M., Adachi, N. \u0026amp; Ochi, M. Autologous bone grafts with MSCs or FGF-2 accelerate bone union in large bone defects. \u003cem\u003eJ. Orthop. Surg. Res.\u003c/em\u003e \u003cb\u003e11\u003c/b\u003e (1), 105 (2016).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eDell, P. C., Burchardt, H. \u0026amp; Glowczewskie, F. P. A roentgenographic, biomechanical, and histological evaluation of vascularized and non-vascularized segmental fibular canine autografts. \u003cem\u003eJ. Bone Joint Surg. Am.\u003c/em\u003e \u003cb\u003e67\u003c/b\u003e (1), 105\u0026ndash;112 (1985).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eEnneking, W. F., Burchardt, H., Puhl, J. J. \u0026amp; Piotrowski, G. Physical and biological aspects of repair in dog cortical-bone transplants. \u003cem\u003eJ. Bone Joint Surg. Am.\u003c/em\u003e \u003cb\u003e57\u003c/b\u003e (2), 237\u0026ndash;252 (1975).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eDoi, K., Tominaga, S. \u0026amp; Shibata, T. Bone grafts with microvascular anastomosis of vascular pedicles: An experimental study in dogs. \u003cem\u003eJ. Bone Joint Surg. Am.\u003c/em\u003e \u003cb\u003e59\u003c/b\u003e (6), 806\u0026ndash;815 (1977).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eBosemark, P., Perdikouri, C., Pelkonen, M., Isaksson, H. \u0026amp; Tagil, M. The masquelet induced membrane technique with BMP and a synthetic scaffold can heal a rat femoral critical size defect. \u003cem\u003eJ. Orthop. Res.\u003c/em\u003e \u003cb\u003e33\u003c/b\u003e (4), 488\u0026ndash;495 (2015).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eYehuda, S. d-Amphetamine and the sensory role of a rat's tail in thermoregulation or what the rat's tail tells the rat's brain. \u003cem\u003eBehav. Biol.\u003c/em\u003e \u003cb\u003e13\u003c/b\u003e (2), 233\u0026ndash;238 (1975).\u003c/span\u003e\u003c/li\u003e \u003cli\u003e\u003cspan\u003eMa, Y. F. et al. Calcium sulfate induced versus PMMA-induced membrane in a critical-sized femoral defect in a rat model. \u003cem\u003eSci. Rep.\u003c/em\u003e \u003cb\u003e8\u003c/b\u003e (1), 637 (2018).\u003c/span\u003e\u003c/li\u003e\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":true,"highlight":"","institution":"","isAcceptedByJournal":false,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true},"keywords":"Induced membrane technique, Rat model, autologous iliac cancellous bone graft","lastPublishedDoi":"10.21203/rs.3.rs-8741751/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-8741751/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003eThe induced membrane technique (IMT) is widely used for the treatment of segmental bone defects; however, small-animal models that faithfully reproduce clinical conditions remain limited. In particular, no rat IMT model using autologous iliac cancellous bone grafts (AICB) has been previously established. This study aimed to develop and characterize such a clinically relevant rat IMT model.\u003c/p\u003e \u003cp\u003eA 5-mm segmental femoral defect was created in rats and stabilized with an external fixator. A polymethyl methacrylate spacer was inserted and replaced with bone graft two weeks later. Outcomes were compared among groups receiving no graft, artificial bone, AICB with IMT, and AICB without spacer placement. Bone regeneration was evaluated using computed tomography, histology, and real-time RT-PCR.\u003c/p\u003e \u003cp\u003eAICB with IMT group achieved the highest bone union rate (66.7%) and demonstrated progressive increases in CT values and new bone formation. In the group, histological analyses demonstrated bone formation mediated by the induced membrane, accompanied by increased expression of osteogenic and angiogenic factors on real-time RT-PCR.\u003c/p\u003e \u003cp\u003eThis study establishes a reproducible rat IMT model using AICB, closely replicating the clinical IMT and providing an experimental platform for invnestigating IMT biology and therapeutic strategies for segmental bone defects.\u003c/p\u003e","manuscriptTitle":"A rat model of the induced membrane technique using autologous iliac cancellous bone graft","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2026-02-19 16:15:49","doi":"10.21203/rs.3.rs-8741751/v1","editorialEvents":[{"type":"communityComments","content":0}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"researchsquare","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":true,"externalIdentity":"","sideBox":"","snPcode":"","submissionUrl":"/submission","title":"Research Square","twitterHandle":"researchsquare","acdcEnabled":true,"dfaEnabled":false,"editorialSystem":"","reportingPortfolio":"","inReviewEnabled":false,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"977bbebe-b287-4491-8ff2-fca867e9c3e9","owner":[],"postedDate":"February 19th, 2026","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"posted","subjectAreas":[{"id":62940974,"name":"Biological sciences/Biotechnology"},{"id":62940975,"name":"Health sciences/Diseases"},{"id":62940976,"name":"Health sciences/Medical research"},{"id":62940977,"name":"Biological sciences/Stem cells"}],"tags":[],"updatedAt":"2026-03-16T10:57:16+00:00","versionOfRecord":[],"versionCreatedAt":"2026-02-19 16:15:49","video":"","vorDoi":"","vorDoiUrl":"","workflowStages":[]},"version":"v1","identity":"rs-8741751","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-8741751","identity":"rs-8741751","version":["v1"]},"buildId":"XKTyCvWXoU3ODBz1xrDgd","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}