A Murine Distal Femoral Epiphysis Ischemia Model Reveals Spatiotemporal Stratification of Necrotic Bone Marrow Clearance and Associated Inflammatory Responses

A Murine Distal Femoral Epiphysis Ischemia Model Reveals Spatiotemporal Stratification of Necrotic Bone Marrow Clearance and Associated Inflammatory Responses | 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 Research Article A Murine Distal Femoral Epiphysis Ischemia Model Reveals Spatiotemporal Stratification of Necrotic Bone Marrow Clearance and Associated Inflammatory Responses Manjun Zhao, Zuyong Feng, Zhirui Hua, Shengping Tang, Jianhong Liu, and 5 more This is a preprint; it has not been peer reviewed by a journal. https://doi.org/ 10.21203/rs.3.rs-7583841/v1 This work is licensed under a CC BY 4.0 License Status: Published Journal Publication published 23 Nov, 2025 Read the published version in Journal of Orthopaedic Surgery and Research → Version 1 posted 15 You are reading this latest preprint version Abstract Background: Efficient clearance of necrotic marrow is essential for repair after epiphyseal ischemia, yet the tissue-level spatiotemporal evolution of this process and its immunoregulatory mechanisms remain incompletely defined. Methods : Using a refined juvenile ischemic osteonecrosis (JIO) model of the murine distal femoral epiphysis—with medial plus lateral approaches guided by mapped vascular entry points—we fully interrupted epiphyseal perfusion and harvested tissue at postoperative days 1–28. Coronal sections along the vascular entry axis underwent H&E/TUNEL and zonal IHC/IF (Ly6G, myeloperoxidase (MPO), F4/80, EMCN, iNOS, CD206). An in-vitro assay tested whether recombinant MPO(1 μg/mL) modulates LPS+IFN-γ–induced M1 polarization in RAW264.7 cells. Results : We consistently observed a necrosis-repair interface characterized by a centripetal tri-zonal progression: fibrotic margin →resorption front → necrotic core. Apoptotic marrow cells—including resident neutrophils—transitioned into secondary necrosis within the resorption zone. This zone demonstrated enrichment of EMCN⁺ microvessels alongside a reduction in hematoxylin-positive nuclear material, indicative of chromatin disposal supported by revascularization processes. At the fibrotic margin, F4/80⁺ macrophages engulfed MPO⁺ , and iNOS⁺ (M1) macrophages outnumbered CD206⁺ (M2). Furthermore, recombinant MPO enhanced M1 polarization of RAW264.7 cells in vitro. Conclusions : The JIO necrosis-repair interface adheres to a distinct spatiotemporal program in which apoptotic cells transition into secondary necrosis concurrently with revascularization efforts. Cells undergoing secondary necrosis are subsequently eliminated during the fibrovascular replacement processes. Notably, neutrophil-derived MPO spatially associates with M1-skewed macrophages in vivo and enhances M1 polarization in vitro; this suggests an immunoregulatory axis exists between neutrophils and macrophages at the interface. Strategies aimed at enhancing the clearance of secondary necrosis neutrophils and modulating this axis may facilitate inflammatory resolution and improve epiphyseal repair. epiphyseal ischemia Damage-associated molecular patterns (DAMPs) secondary necrosis neutrophil macrophage polarization myeloperoxidase revascularization mouse model Figures Figure 1 Figure 2 Figure 3 Figure 4 Figure 5 Introduction Legg–Calvé–Perthes disease (LCPD) is a pediatric avascular necrosis of the femoral head with substantial risk of long-term disability. The disease evolves from an ischemic necrosis phase into a repair phase characterized by resorption of necrotic bone, fibrovascular ingrowth, and new bone formation ( 1 – 3 ) . Interventional evidence from a piglet model shows that percutaneous three-needle lavage of the necrotic femoral head can accelerate repair ( 4 , 5 ) , supporting the concept that active removal of necrotic marrow contents can accelerate repair. However, mechanistic details of marrow clearance remain poorly defined, limiting the development of targeted therapies. Histopathology ( 2 , 6 ) and perfusion MRI ( 7 ) show that necrotic tissue abuts newly formed fibrovascular tissue, forming a distinct necrosis–repair interface that likely functions as a regulatory zone for clearing necrotic material. Because clinical specimens are scarce, the juvenile ischemic osteonecrosis (JIO) mouse model ( 8 ) —whose pathological trajectory parallels LCPD—has become a key tool for mechanistic study. Establishing and characterizing this interface in the JIO model is therefore essential to elucidate the cellular and molecular processes underlying marrow clearance. The direction of revascularization appears to follow native epiphyseal vascular entry points, as indicated by super-selective angiography ( 9 , 10 ) , perfusion MRI ( 11 , 12 ) , and large animal models ( 13 ) . In rats ( 14 ) , two of the four nutrient vessels enter via cartilage-free zones on the medial and lateral condyles (sites of synovial/ligament attachment) anatomically aligned with the coronal plane. We hypothesized that murine epiphyseal vasculature is similar and that coronal sectioning along the vascular entry axis would more clearly reveal the necrosis–repair interface, enabling accurate morphological analysis of marrow clearance. Neutrophils and macrophages, as professional phagocytes, are central to necrotic tissue disposal. Prior work in the JIO model showed macrophage phagocytosis of necrotic fat promotes anti-inflammatory polarization ( 15 ) . Yet the epiphysis also contains abundant marrow cells whose necrosis releases DAMPs (e.g., nucleic acids, HMGB1, S100), likely shaping inflammation and fibrosis ( 16 ) , differently from adipose-derived DAMPs. While macrophage handling of necrotic adipose has been described, the clearance of necrotic marrow cells themselves and their immune sequelae remain unclear. Furthermore, how neutrophil-derived enzymes such as MPO modulate macrophage polarization at the interface is unknown. Here, we anatomically localize vascular entry sites, reconstruct the necrosis–repair interface in coronal sections, delineate marrow cell death and clearance, and map vessels, neutrophils, and macrophages in space; we also test the effect of MPO on macrophage polarization in vitro. Materials and Methods Animals All procedures were approved by the Animal Care and Welfare Committee of Guangxi Medical University and complied with institutional guidelines (approval no.202401027). Male C57BL/6 mice were used (5-week-old). Six mice were allocated for distal femoral epiphyseal vascular anatomy, and 36 mice underwent JIO surgery of the right distal femoral epiphysis. The contralateral distal femoral epiphysis served as the intact normal control. Vascular anatomy of the distal femoral epiphysis Both knees of six mice were dissected to map the epiphyseal vascular entry points. Under a stereomicroscope (6.7–40×), a medial parapatellar approach was performed on mice anesthetized with 2.5% tribromoethanol (0.2 mL/10 g, i.p.) and provided with peri-operative analgesia (buprenorphine 0.05–0.1 mg/kg s.c., ± meloxicam 1–2 mg/kg s.c.). The medial approach facilitated exposure of the popliteal vessel (PV) and the medial proximal genicular vessel (MPGV). Following a medial capsulotomy, transection of the patellar ligament, and retraction of Hoffa’s fat pad allowed for visualization of the central genicular vessel (CGV). Additionally, the lateral proximal genicular vessel (LPGV) and lateral genicular vessel (LGV) were visualized on the lateral side of the epiphysis. All epiphyseal entry points were documented using a high-resolution camera. After documentation, animals were euthanized via pentobarbital overdose. Induction of juvenile ischemic osteonecrosis To achieve complete interruption of epiphyseal perfusion, all nutrient vessels were cauterized at their entry points into the epiphysis. Mice were anesthetized and received perioperative analgesia. They were positioned supine on a warming pad, and the right groin and hind limb were shaved. The operative field was prepared three times with povidone-iodine and 70% ethanol; ophthalmic ointment was applied to protect the eyes. Under a stereomicroscope (magnification ~ 6–40×), a medial parapatellar skin incision approximately 10–15 mm in length was made. The medial hamstrings were carefully dissected to expose and cauterize the epiphyseal branch of the popliteal vessel (PV). A medial parapatellar arthrotomy was subsequently performed to expose the medial proximal genicular (MPGV), which was cauterized prior to its entry into the epiphysis. Hoffa’s fat pad was retracted to reveal the central genicular vessel (CGV), which was also cauterized at its epiphyseal entry site. A separate lateral knee arthrotomy was then conducted to expose both the lateral proximal genicular vessel (LPGV) and lateral genicular vessel (LGV), which underwent similar cauterization at their respective epiphyseal entry points. Following these procedures, joint irrigation was performed, hemostasis confirmed, and closure of both capsule and muscles achieved using 8 − 0 monofilament nylon sutures; skin closure utilized 4 − 0 silk sutures. Postoperative analgesia consisted of buprenorphine 0.05–0.1 mg/kg s.c. every 12 h for 48 h and meloxicam 1–2 mg/kg s.c. every 24 h for 48–72 h. The left distal femoral epiphysis remained untouched as an intact normal control (refer to Fig. S1 for procedural steps). Tissue processing At postoperative days 1, 4, 7, 10, 14, and 28 (n = 6/time point), bilateral distal femora were harvested, fixed in 4% neutral-buffered formalin (24 h), decalcified in 14% EDTA (pH 7.4) for 10–14 days, embedded in paraffin, and sectioned coronally at 4 µm. Histology and TUNEL H&E staining was performed for morphology; TUNEL (Beyotime, C1098) identified apoptotic nuclei. Based on nuclear/tissue features, the interface was partitioned into: (i) Fibrotic zone (FZ) : Bone marrow space replaced by fibrovascular tissue with cells showing normal nuclear morphology. (ii) Resorption zone (RZ) : Bone marrow space filled with karyolytic cell remnants and residual cytoskeleton, with few intact nucleated cells. (iii) Necrotic zone (NZ) : Bone marrow cells with pyknotic or partially karyolytic nuclei and minimal inflammatory/fibroblastic infiltration. Immunohistochemistry (IHC) and immunofluorescence (IF) IHC. Only specimens with a clear fibrotic–resorption boundary were analyzed. After deparaffinization and sodium-citrate heat retrieval (10 mM citrate, 0.05% Tween-20, pH 6.0), sections were treated with 3% H₂O₂ and blocked with 10% goat serum (Solarbio, SL038). Primary antibodies (4°C, overnight): Ly6G (1:100, BD Pharmingen 551459), F4/80 (1:200, BioLegend 123101), endomucin/EMCN (1:200, Santa Cruz Biotechnology sc-65495), MPO (1:200, ZENBIO R25062), CD206 (1:2000, Proteintech 18704-1-AP), and iNOS (1:200, Invitrogen PA1-036). HRP-labeled secondary antibodies (Beyotime: A0208 anti-rabbit, A0192 anti-rat) and DAB chromogen were used, followed by hematoxylin counterstain. IF. Dual IF assessed F4/80–MPO colocalization near the interface. After antigen retrieval and blocking as above, primary antibodies were applied sequentially with matched fluorescent secondaries (1:100, Invitrogen A-11008; 1:500, Cell Signaling Technology 4417S) and DAPI nuclear stain. Images were acquired under unified exposure on fluorescence/confocal microscopy. Histomorphometry and statistics The contralateral distal femoral epiphysis served as the intact normal control for qualitative reference (Fig. 2 A). For semi-quantitative, within-animal comparisons on the same slides, the ipsilateral metaphysis was used as an internal control to minimize inter-animal and staining-batch variability. High-power fields (HPFs) were sampled from each zone on adjacent H&E, TUNEL, and IHC sections using fixed imaging settings. Quantitative endpoints included: fibrosis proportion (fibrovascular area/total marrow cavity), EMCN-positive area (% area/HPF), Ly6G- and MPO-positive cells per HPF, F4/80-positive area (% area/HPF), and hematoxylin-positive nuclear area fraction (after H–DAB color deconvolution). Regions of interest (ROIs) excluded cartilage, mineralized trabeculae, growth plate, and processing artifacts. Data are expressed as mean ± SEM. Normality was assessed by the Shapiro-Wilk test. For paired, within-animal comparisons, normally distributed data were analyzed with paired t-tests, and non-normal data with Wilcoxon signed-rank tests. Two-tailed p < 0.05 was considered statistically significant. RAW264.7 macrophage assay with MPO RAW264.7 cells were maintained in DMEM supplemented with 10% FBS and 1% penicillin–streptomycin at 37°C in 5% CO₂, then seeded (6-well plates for RNA; glass coverslips in 24-well plates for IF) and allowed to adhere overnight. Four conditions were applied for 24 h: control (medium only); M1 induction (LPS 100 ng/mL + IFN-γ 20 ng/mL); M1 + MPO (same M1 induction with recombinant murine MPO 1 µg/mL added at the start); and MPO alone (1 µg/mL). For immunofluorescence, cells were rinsed with PBS, fixed in 4% paraformaldehyde (15 min), permeabilized with 0.1% Triton X-100 (10 min), blocked with 10% goat serum (30 min), incubated with anti-iNOS primary antibody overnight at 4°C, then with fluorescent secondary antibody (1:100, Invitrogen A-1101, 1 h, room temperature); nuclei were counterstained with DAPI. Images were acquired under identical exposure settings, and iNOS signal was quantified as mean fluorescence intensity from ≥ 3 non-overlapping fields per well and averaged per biological replicate. For qPCR, parallel wells were lysed in TRIzol for RNA extraction, reverse-transcribed to cDNA, and analyzed by SYBR Green qPCR for iNOS and TNFα, normalized to β-Actin and calculated using the 2^−ΔΔCt method. Results 1. Vascular anatomy of the distal femoral epiphysis in mice Detailed dissections in six mice mapped the epiphyseal entry points of the MPGV, CGV, LPGV, and LGV ( Fig. 1 ). The saphenous vessel gave rise to the popliteal trunk. The MPGV originated from the saphenous or proximal popliteal vessel, crossed the physis, and entered the medial epiphysis. The LPGV coursed along the lateral femur and supplied the lateral epiphysis through multiple branches that anastomosed with vessels in Hoffa’s fat pad. The LGV arose from the popliteal vessel and entered the lateral epiphysis. Extensive anastomoses among the MPGV, LPGV, and LGV were evident within Hoffa’s fat pad and near the lateral epiphysis. Because a medial arthrotomy alone did not fully expose the LGV, adding a lateral arthrotomy enabled direct visualization and complete cauterization of all epiphyseal vessels (Fig. S1). 2. Histological changes during revascularization Coronal sections showed progressive clearance of necrotic marrow with stepwise fibrovascular ingrowth (Fig. 2A–H). On day 1, marrow cells and osteocytes across the epiphysis displayed diffuse nuclear pyknosis with no fibrous infiltration. By day 4, karyolysis appeared in marrow cavities adjacent to the medial condylar surface, while the central epiphysis remained pyknotic. On day 7, fibroblasts and small vessels infiltrated the medial marrow cavity, and the karyolytic zone extended toward the center. By day 10, fibrovascular tissue had grown in from both condyles, leaving broad transitional areas of karyolysis; pyknotic nuclei persisted near the growth plate in the epiphyseal center. By day 14, the marrow cavity was almost completely replaced by fibrovascular tissue, with new bone forming along necrotic trabeculae. By day 28, regenerated marrow cells and abundant new bone were present within the cavity. ( Repaired region, see Fig.S2.) Fibrosis became detectable by day 4 and rapidly progressed from day 7 to day 10. By this time, the necrotic core, absorption zone, and fibrotic margin coexisted; consequently, by day 14, the necrotic bone marrow was nearly completely replaced. Serial sections corroborated this as a continuous progressive process: initially (or within the necrotic area), the nuclei underwent pyknosis, followed by the dissolution of pyknotic nuclei as microvessels and fibroblasts infiltrated the region. This invasion formed an absorption zone characterized by anucleate "ghost" cells or residual cytoskeleton. These remnants were gradually phagocytosed, leading to the replacement of the bone marrow cavity with fibrovascular tissue and completing the clearance of necrotic bone marrow. 3. Apoptosis and secondary necrosis of marrow cells and resident neutrophils High-power H&E and TUNEL staining revealed marked regional differences in cellular morphology (Fig. 3). Cells in the fibrotic zone and in the metaphysis appeared morphologically normal. In the necrotic zone, cells exhibited nuclear pyknosis, karyorrhexis, karyolysis, or reticular chromatin, while plasma membranes remained largely intact; in contrast, the resorption zone lacked discernible nuclear structures and contained abundant eosinophilic cytoskeletal remnants with only sparse infiltration by intact nucleated cells. TUNEL signals in the necrotic zone localized to pyknotic or fragmented nuclei, and apoptotic bodies were observed (Fig. 3), consistent with apoptosis. In the resorption zone, TUNEL-positive nuclei appeared swollen with loss of chromatin, indicating insufficient clearance of apoptotic cells and their progression to secondary necrosis (17, 18) . Resident bone marrow neutrophils showed a similar spatial pattern: Ly6G⁺ cells were morphologically normal in the metaphysis, displayed apoptotic nuclear changes in the necrotic zone, and exhibited near-complete nuclear lysis with membrane disruption in the resorption zone. Only scattered, morphologically normal neutrophils were seen in the fibrotic zone, and the number of Ly6G⁺ cells there was lower than in the resorption zone (Fig. 4A, B; p < 0.05 ). Although Ly6G⁺ counts did not differ markedly among the metaphysis, necrotic, and resorption zones, MPO⁺ cells declined significantly from the metaphysis to the necrotic and resorption zones (Fig. 4A, B; p < 0.05 ). These findings indicate that marrow stromal cells including resident neutrophils undergo apoptosis after ischemia, with many progressing to secondary necrosis when clearance is insufficient. Along the axis of fibrovascular ingrowth , nuclear morphology delineated regions enriched in apoptotic versus secondarily necrotic cells, suggesting that the secondary necrosis process during epiphyseal clearance is coupled to—and potentially regulated by—fibrovascularization . 4. Fibrovascular ingrowth and chromatin clearance EMCN immunostaining revealed abundant microvessels in the fibrotic, resorption, and metaphyseal zones, whereas the necrotic zone was avascular; morphologically, microvessels localized to the resorption side of the necrosis–resorption interface (Fig. 4A, C). Quantitatively, the EMCN-positive area in the necrotic zone was significantly lower than in the other zones, while the hematoxylin-positive nuclear area was significantly greater than in the resorption zone (Fig. 4B, F; p < 0.05). These findings suggest that microvascular ingrowth within the resorption zone may facilitate dissolution and clearance of nuclear material from apoptotic cells, thereby promoting their transition to secondary necrosis. 5. Macrophage involvement in clearing neutrophil remnants F4/80 immunohistochemistry showed a significantly larger positive area in the fibrotic zone than in the resorption zone (Fig. 4A–B). Ly6G⁺ and MPO⁺ cells were least abundant in the fibrotic zone. At high magnification, cells at the fibrotic–resorption interface were observed engulfing MPO-positive cytoskeletal remnants (Fig. 4D), and dual immunofluorescence confirmed F4/80–MPO colocalization (Fig. 4E). Together, these findings indicate that macrophages at the fibrotic margin phagocytose secondarily necrotic neutrophils , thereby promoting fibrovascular replacement of the marrow cavity. 6. Macrophage polarization at the fibrotic–resorption interface; MPO promotes M1 polarization in vitro IHC analysis revealed a predominance of iNOS⁺ (M1) macrophages over CD206⁺ (M2) macrophages at the fibrotic–resorption interface (Fig.5A,B;P < 0.05), indicating that M1 cells are frequently clustered in proximity to MPO-rich regions. In RAW264.7 cells, addition of recombinant MPO (1 μg/mL) during LPS+IFN-γ–driven M1 induction (vs. LPS+IFN-γ alone) increased iNOS immunofluorescence and upregulated iNOS and TNFα mRNA (qPCR; all P < 0.05) (Fig. 5C,D,E; P < 0.05). Discussion Vascular anatomy and model refinement The anatomical sites where vessels enter the epiphysis may determine the pattern of revascularization in necrotic epiphyses. Evidence from super-selective arterial perfusion imaging ( 9 , 10 ) , perfusion MRI ( 11 , 12 ) , and juvenile pig models ( 13 ) supports this view. In 2015, Kim et al. ( 8 ) used micro-CT to delineate the major arterial trunks supplying the murine distal femoral epiphysis, providing an anatomical basis for constructing the JIO mouse model. However, like other studies of murine hind-limb vasculature ( 19 ) that work did not clearly resolve epiphyseal entry points and thus could not elucidate the revascularization pattern of the necrotic murine epiphysis. In 2021, Ma et al. ( 14 ) mapped the nutrient vessels of the distal femoral epiphysis in a rat ischemic necrosis model and, on subsequent micro-CT during the repair phase, observed that neovessels preferentially entered through cartilage-free zones on the medial and lateral condyles. In the present study, we anatomically defined the epiphyseal entry points in mice, confirmed a murine–rat similarity in epiphyseal vascular anatomy, and identified a more extensive lateral vascular network with prominent anastomoses than previously reported. These findings provide additional anatomical context for subsequent studies of epiphyseal necrosis and repair. Mapping the epiphyseal vascular entry points clarified revascularization routes and justified adding a lateral approach to ensure complete interruption of perfusion. Coronal sectioning aligned with this entry axis consistently reconstructed the necrosis–repair interface and its centripetal advance; by days 7–10, a reproducible necrotic–resorption–fibrotic triad was evident. Building on this framework, temporal sequencing and zonal criteria delineated a distinct spatiotemporal stratification with a front that progresses from the periphery toward the center. The continuous, zone-specific shifts in nuclear and matrix features indicate that clearance is not uniform but proceeds in staged steps tightly coupled to vascular ingress or the local oxygen gradient and humoral factor. Patterns of necrosis and tissue clearance The mode of cell death dictates its clearance pathway. Although direct evidence from human Perthes disease tissue is lacking, porcine models indicate that apoptosis and oncosis (primary necrosis) coexist after epiphyseal ischemia ( 20 ) . Importantly, oncosis and secondary necrosis are morphologically similar at their terminal stages—cell swelling, loss of chromatin/nuclear staining, and “ghost cells”—and are therefore difficult to distinguish on static sections ( 21 ) . In JIO mouse ( 8 ) and rat models ( 14 ) , loss of marrow nuclear staining is commonly observed about one week after modeling; the diagnostic key, however, is not the end-stage morphology but whether an apoptotic phase precedes the anucleate state. In our spatiotemporal analyses, apoptotic cells were evident on postoperative day 1 and within the necrotic core, whereas abundant anucleate “ghost cells” emerged later, specifically within the resorption zone. We therefore infer that anucleate cells in the resorption zone primarily derive from uncleared apoptotic cells that progressed to secondary necrosis, rather than from primary oncosis. Temporally, marrow changes previously labeled as “oncosis” in JIO mice and juvenile pigs tend to occur at later time points, aligning more closely with the kinetics of secondary necrosis (late apoptotic lysis). Thus, a portion of the oncosis-like morphology reported in prior studies likely includes—or predominantly reflects—secondary necrosis of apoptotic cells. Mechanisms and cellular clearance of secondary necrosis Secondary necrosis represents a programmed lytic transition of uncleared apoptotic cells rather than passive decay. Mechanistically, caspase-3 cleavage of gasdermin E (GSDME/DFNA5) generates an N-terminal fragment that forms large pores in the plasma membrane, priming apoptotic cells for lysis—one of the key molecular switches from apoptosis to secondary necrosis ( 22 , 23 ) . Ninjurin-1 ( NINJ1 ) then executes plasma membrane rupture (PMR) as a shared terminal effector in late apoptosis, pyroptosis, and necrosis, enabling bulk release of cytosolic contents and DAMPs ( 24 , 25 ) . plasma membrane channel pannexin 1( PANX1 ) channels, beyond mediating “find-me” signals, can remain open to disturb ionic homeostasis and promote late cell lysis ( 26 ) . In parallel, chromatin is dismantled in an ordered fashion: (caspase-activated deoxyribonulease) CAD is activated by caspase-3–mediated (Inhibitor of Caspase-Activated DNase ) ICAD cleavage to initiate intracellular nucleosomal degradation ( 27 ) , while extracellular deoxyribonuclease I (DNase I)–family enzymes (e.g., DNase I, DNase γ) further process released nucleosomes; if clearance is inadequate, the accumulation of nucleosomal DNA and histones amplifies inflammation. Integrating prior work with our data, When revascularization reaches the necrosis–repair interface, it delivers humoral factors ( 28 ) (e.g., DNase I ( 29 ) , complement ( 30 ) and natural antibodies ( 31 ) ) and cellular effectors ( 32 ) ( neutrophils, macrophages) while restoring oxygen/ion flux and perfusion. These may inputs destabilize membrane homeostasis, amplify caspase-3/GSDME pore formation and PANX1 activity, and, via NINJ1, trigger PMR—converting “silent” apoptosis into secondary necrosis on the revascularized (resorption) side. Concurrently, DNases accelerate chromatin breakdown, macrophages/fibroblasts remove residual cytoskeleton, and microcirculation washes out soluble DAMPs and degradation products, shifting the milieu from inflammatory to reparative. This framework explains why secondary necrosis follows vessel ingrowth in time and localizes to the vascular side in space, yielding the observed centripetal triad of necrotic core → resorption front → fibrotic zone. Immune regulation of neutrophil secondary necrosis Our study shows that neutrophils in the resorption zone undergo secondary necrosis. Although secondarily necrotic neutrophils also lose plasma-membrane integrity, they are less pro-inflammatory than cells that undergo primary necrosis ( 33 ) , in part because the inflammatory activity of their released DAMPs is attenuated by apoptosis-associated degradative processes ( 34 ) . Fractionation of lysed neutrophils into membrane and soluble components indicates that the membrane fraction behaves like that of apoptotic cells—anti-inflammatory via phosphatidylserine (PS)—whereas proteases released during lysis are pro-inflammatory ( 35 ) . While some reports show anti-inflammatory effects of neutrophil α-defensins released from apoptotic/necrotic neutrophils ( 36 , 37 ) , other soluble factors can exacerbate inflammation: neutrophil elastase can cleave PS receptors and impair efferocytosis ( 38 ) , and secondarily necrotic neutrophils selectively release IL-16C and macrophage migration inhibitory factor(MIF) degradation to promote inflammatory responses ( 39 , 40 ) . In our study, whether MIF accounts for the relative paucity of macrophages in the resorption and necrotic zones remains unclear. Nevertheless, we observed MPO-rich fibrotic–resorption interfaces dominated by M1 (iNOS⁺) macrophages, and recombinant MPO promoted M1 polarization of RAW264.7 cells in vitro, supporting a pro-inflammatory role of MPO during necrotic marrow clearance. Whether MPO, IL-16C, and MIF can serve as inflammatory biomarkers of epiphyseal necrosis resolution warrants further investigation. Multiple lines of evidence implicate DAMP–PRR signaling in Perthes disease. Synovial studies show chronic synovitis with elevated IL-6 and increased HMGB1 that correlates with IL-6 ( 41 , 42 ) Necrotic bone activates macrophages via TLR4 ( 43 ) , while pharmacologic TLR4 inhibition accelerates epiphyseal repair ( 44 ) . In a juvenile pig model, minimally invasive lavage of necrotic marrow improved bone healing ( 5 ) . DAMPs from necrotic femoral heads suppress osteogenesis and promote mesenchymal-stem-cell fibrosis ( 16 ) , and macrophage clearance of necrotic fat drives anti-inflammatory polarization ( 15 ) . Yet how necrotic marrow cells—and the DAMPs they release—are cleared remains unclear. Our data reveal secondary necrosis of resident neutrophils at the necrosis–repair interface and suggest that targeting the removal of necrotic marrow cells, particularly secondarily necrotic neutrophils, may hasten inflammatory resolution and improve epiphyseal repair. Limitations Although our detailed anatomical model of the intraosseous epiphyseal plate (JIO) and the histopathological atlas of the necrosis-repair interface have advantages, this study still has several limitations. We only used male C57BL/6 mice, which may limit the generalizability of our findings to human LCPD. Most of the evidence was derived from static decalcified paraffin sections analyzed by semi-quantitative H&E/IHC/IF methods; in vivo imaging and functional readouts were not included in this study. Although the observed colocalization and regional associations are relevant, causal inferences still need to be established by genetic or pharmacological intervention of MPO, GSDME, NINJ1, complement components, deoxyribonuclease I (DNase I), and IL-16C/MIF, and direct assessment of phagocytic efficiency, DAMPs profiles, and cytokine kinetics. Finally, we did not evaluate biomechanical strength, three-dimensional microstructure, or therapeutic endpoints of intervention. Conclusions The necrosis–repair interface in the refined murine JIO model demonstrates that necrotic marrow clearance proceeds through a distinct, outward-to-inward tri-zonal program. Apoptotic marrow cells—including resident neutrophils—shift to secondary necrosis as revascularization advances, and the resulting remnants are eliminated during fibrovascular replacement. Neutrophil-derived MPO is spatially associated with M1-polarized macrophages in vivo and augments M1 polarization in vitro, suggesting an immunoregulatory neutrophil–macrophage axis at the interface. Strategies that enhance clearance of secondarily necrotic neutrophils and modulate this axis may accelerate inflammatory resolution and improve epiphyseal repair. Declarations Ethics approval and consent to participate The animal experiment was approved by the Animal Care & Welfare Committee of Guangxi Medical University (approval no.202401027; Nanning, China). Consent for publication This paper is approved by all authors for publication. Availability of data and materials The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request. Competing interests The authors declare no competing interests. Funding The authors declare that financial support was received for the research, authorship, and/or publication of this article. This work was supported by the Basic Research Capacity Enhancement Program for Young and Middle-aged Faculty at Universities and Colleges in Guangxi (Grant No. 2025KY0130); the Guangxi Natural Science Foundation—Joint Project on Regional High-Incidence Diseases Research (Grant No. 2023GXNSFAA026342); the Featured Innovation Team Cultivation Project of the First Affiliated Hospital of Guangxi Medical University (Grant No. YYZS2024002); the Youth Science and Technology Inspiration Star Program of the First Affiliated Hospital of Guangxi Medical University (Grant No. YYZS2022008); and the National Natural Science Foundation of China (Grant No. 82460422). Author contributions MZ designed the research and wrote the original draft. ZF, ZH, ST, JL and ZL performed the experiments. QH: Methodology, Project administration BL performed the analysis. XD and SL: Project administration, Supervision, Funding acquisition. All authors read and approved the final manuscript. The authors declare that all data were generated in-house, that no paper mill was used, and that no AI tool has been used for the generation of text or figures. Acknowledgements We thank the Animal Care & Welfare Core and institutional Histology/Imaging facilities for routine support. No professional writing services were used, and no materials were received as in-kind gifts; all experiments, analyses, and manuscript preparation were performed by the authors. All acknowledged units/cores have consented to be named. References Kim HK. Pathophysiology and new strategies for the treatment of Legg-Calvé-Perthes disease. J Bone Joint Surg Am. 2012;94(7):659-69. Jonsater S. Coxa plana; a histo-pathologic and arthrografic study. Acta Orthop Scand Suppl. 1953;12:5-98. Catterall A, Pringle J, Byers PD, Fulford GE, Kemp HB, Dolman CL, et al. A review of the morphology of Perthes' disease. J Bone Joint Surg Br. 1982;64(3):269-75. 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08:54:34","extension":"xml","order_by":29,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":116799,"visible":true,"origin":"","legend":"","description":"","filename":"6e9706484bbf405eac5aea99f69edda21structuring.xml","url":"https://assets-eu.researchsquare.com/files/rs-7583841/v1/f12267c96f316cb623ae84c9.xml"},{"id":91831678,"identity":"a2bb7cc7-95e5-44ca-8c23-b04e6547667c","added_by":"auto","created_at":"2025-09-22 09:10:34","extension":"html","order_by":30,"title":"","display":"","copyAsset":false,"role":"acdc-reference","size":130419,"visible":true,"origin":"","legend":"","description":"","filename":"earlyproof.html","url":"https://assets-eu.researchsquare.com/files/rs-7583841/v1/3331ec814af98a0c6389f668.html"},{"id":91829053,"identity":"aa3dea70-6341-4d35-81b1-5999751b2fe8","added_by":"auto","created_at":"2025-09-22 08:54:33","extension":"png","order_by":1,"title":"Figure 1","display":"","copyAsset":false,"role":"figure","size":388196,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eVascular anatomy of the distal femoral epiphysis in mice.\u003c/strong\u003e\u003cbr\u003e\n \u003cstrong\u003e(A)\u003c/strong\u003e Medial incision\u003cstrong\u003e with transection of the medial head of the gastrocnemius, \u003c/strong\u003eexposing the saphenous vessels (SV) and popliteal vessels (PV).\u003cstrong\u003e(B)\u003c/strong\u003e Medial capsulotomy showing the medial proximal genicular vessels (MPGV) crossing the physis.\u003cstrong\u003e(C)\u003c/strong\u003eBranch of the MPGV entering the medial epiphysis (black arrow).\u003cstrong\u003e(D)\u003c/strong\u003eTransection of the patellar ligament and retraction of Hoffa’s fat pad expose the central genicular vessel (CGV) as it enters the epiphysis. \u003cstrong\u003e(E)\u003c/strong\u003e With hip adduction, the lateral proximal genicular vessels (LPGV) and lateral genicular vessels (LGV) are seen entering the lateral epiphysis (black arrows) and interconnecting (white arrows).\u003cstrong\u003e(F)\u003c/strong\u003eAnastomoses among the MPGV, LPGV, and LGV within the fat pad (white arrows).\u003c/p\u003e","description":"","filename":"1.png","url":"https://assets-eu.researchsquare.com/files/rs-7583841/v1/aaf8fe9d253d321d1e011b08.png"},{"id":91829056,"identity":"b809e75b-a57c-408d-89a0-28b20cd6c1b4","added_by":"auto","created_at":"2025-09-22 08:54:33","extension":"png","order_by":2,"title":"Figure 2","display":"","copyAsset":false,"role":"figure","size":447906,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eHistological progression of necrotic marrow clearance and fibrovascular ingrowth in the distal femoral epiphysis after vascular occlusion.\u003c/strong\u003e\u003cbr\u003e\n \u003cstrong\u003e(A)\u003c/strong\u003e Contralateral (normal) epiphysis showing intact marrow cells (inset boxed).\u003cstrong\u003e(B)\u003c/strong\u003e Day 1: diffuse nuclear pyknosis in marrow cells.\u003cstrong\u003e(C)\u003c/strong\u003e Day 4: karyolysis in marrow adjacent to the medial condyle (\u003cstrong\u003ea\u003c/strong\u003e), with persistent pyknosis in the central epiphysis (\u003cstrong\u003eb\u003c/strong\u003e).\u003cstrong\u003e(D)\u003c/strong\u003e Day 7: fibroblasts infiltrate the medial marrow cavity containing karyolytic cells (\u003cstrong\u003ea\u003c/strong\u003e); residual pyknosis remains centrally (\u003cstrong\u003eb\u003c/strong\u003e).\u003cstrong\u003e(E)\u003c/strong\u003eDay 10: extensive fibrovascular ingrowth from the medial and lateral condyles (\u003cstrong\u003ea\u003c/strong\u003e), a central transitional zone with karyolytic cells (\u003cstrong\u003eb\u003c/strong\u003e), and persistent pyknotic nuclei near the growth plate (\u003cstrong\u003ec\u003c/strong\u003e).\u003cstrong\u003e(F)\u003c/strong\u003eDay 14: marrow cavity nearly completely replaced by fibrovascular tissue; new bone forms along necrotic trabeculae.\u003cstrong\u003e(G)\u003c/strong\u003e Day 28: regenerated marrow cells with abundant new bone.\u003cstrong\u003e(H)\u003c/strong\u003e Quantification of the fibrosis proportion over time.(H\u0026amp;E-stained coronal sections; Repaired region, see Fig.S2.)\u003c/p\u003e","description":"","filename":"2.png","url":"https://assets-eu.researchsquare.com/files/rs-7583841/v1/f7182c9b9d1d5879b4bb6633.png"},{"id":91829052,"identity":"5730d357-0c80-4285-9f9d-f6d62001c6b2","added_by":"auto","created_at":"2025-09-22 08:54:33","extension":"png","order_by":3,"title":"Figure 3","display":"","copyAsset":false,"role":"figure","size":642442,"visible":true,"origin":"","legend":"\u003cp\u003eH\u0026amp;E, TUNEL, and Ly6G IHC across fibrotic, resorption, necrotic, and metaphyseal zones (White arrows indicate apoptotic bodies).\u003c/p\u003e","description":"","filename":"3.png","url":"https://assets-eu.researchsquare.com/files/rs-7583841/v1/c8fa2f57a65667f7c0e85349.png"},{"id":91829060,"identity":"f2d2950e-5a81-42a7-bb1d-e348bc4a6f29","added_by":"auto","created_at":"2025-09-22 08:54:33","extension":"png","order_by":4,"title":"Figure 4","display":"","copyAsset":false,"role":"figure","size":714037,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eMicrovascular distribution and macrophage-mediated clearance of secondarily necrotic neutrophil remnants in the distal femoral epiphysis.\u003c/strong\u003e\u003cbr\u003e\n \u003cstrong\u003e(A)\u003c/strong\u003e Immunostaining for \u003cstrong\u003eEMCN, Ly6G, F4/80, \u003c/strong\u003eand \u003cstrong\u003eMPO\u003c/strong\u003e in the fibrotic, resorption, necrotic, and metaphyseal zones.\u003cbr\u003e\n \u003cstrong\u003e(B)\u003c/strong\u003e Quantitative analysis of \u003cstrong\u003eEMCN-positive area, Ly6G-positive cell counts\u003c/strong\u003e and\u003cstrong\u003e F4/80-positive area \u003c/strong\u003eacross the four zones.\u003cbr\u003e\n \u003cstrong\u003e(C)\u003c/strong\u003e \u003cstrong\u003eEMCN-positive microvessels\u003c/strong\u003e (white arrows) localize to the \u003cstrong\u003eresorption side\u003c/strong\u003e of the necrosis–resorption interface(upper left).\u003cbr\u003e\n \u003cstrong\u003e(D)\u003c/strong\u003e High-magnification images of the \u003cstrong\u003efibrotic–resorption interface\u003c/strong\u003e showing cells \u003cstrong\u003eengulfing MPO-positive cytoskeletal remnants\u003c/strong\u003e (black arrows).\u003cbr\u003e\n \u003cstrong\u003e(E)\u003c/strong\u003e \u003cstrong\u003eDual immunofluorescence\u003c/strong\u003e demonstrating \u003cstrong\u003ecolocalization of F4/80 and MPO\u003c/strong\u003e at the fibrotic–resorption interface (white arrows).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(F)\u003c/strong\u003e \u003cstrong\u003eMPO-positive cell counts\u003c/strong\u003e and Quantification of \u003cstrong\u003ehematoxylin-positive nuclear area\u003c/strong\u003e across the four zones.\u003c/p\u003e","description":"","filename":"4.png","url":"https://assets-eu.researchsquare.com/files/rs-7583841/v1/2b2a9fd4727010506cabbc0a.png"},{"id":91829057,"identity":"59c02877-aca5-498d-bf98-a054b850d257","added_by":"auto","created_at":"2025-09-22 08:54:33","extension":"png","order_by":5,"title":"Figure 5","display":"","copyAsset":false,"role":"figure","size":349447,"visible":true,"origin":"","legend":"\u003cp\u003e\u003cstrong\u003eMacrophage polarization at the fibrotic–resorption interface and MPO-driven promotion of M1 polarization in vitro.\u003c/strong\u003e\u003cbr\u003e\n \u003cstrong\u003e(A)\u003c/strong\u003e Immunohistochemistry for \u003cstrong\u003eiNOS\u003c/strong\u003e (M1 marker) and \u003cstrong\u003eCD206\u003c/strong\u003e (M2 marker) at the fibrotic–resorption interface of distal femoral epiphysis sections. \u003cstrong\u003eiNOS⁺\u003c/strong\u003e cells outnumber \u003cstrong\u003eCD206⁺\u003c/strong\u003ecells in this region.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e(B)\u003c/strong\u003eQuantification of \u003cstrong\u003eiNOS⁺\u003c/strong\u003e and \u003cstrong\u003eCD206⁺\u003c/strong\u003emacrophages per high-power field (HPF) at the fibrotic–resorption interface (mean ± SEM, \u003cstrong\u003en = 10\u003c/strong\u003e; *P \u0026lt; 0.05).\u003cbr\u003e\n \u003cstrong\u003e(C)\u003c/strong\u003e Immunofluorescence for \u003cstrong\u003eiNOS\u003c/strong\u003e in RAW264.7 cells after M1 induction with \u003cstrong\u003eLPS + IFN-γ\u003c/strong\u003e, with or without \u003cstrong\u003erecombinant MPO (1 μg/mL)\u003c/strong\u003e.\u003cbr\u003e\n \u003cstrong\u003e(D)\u003c/strong\u003e Quantification of \u003cstrong\u003eiNOS\u003c/strong\u003e mean immunofluorescence intensity with and without MPO.\u003cbr\u003e\n \u003cstrong\u003e(E)\u003c/strong\u003e qPCR showing significant upregulation of \u003cstrong\u003eiNOS \u003c/strong\u003eand\u003cstrong\u003eTNFα \u003c/strong\u003emRNA in MPO-treated, M1-polarized RAW264.7 cells compared with M1 induction alone (*P \u0026lt; 0.05, **P \u0026lt; 0.01, ***P \u0026lt; 0.001).\u003c/p\u003e","description":"","filename":"5.png","url":"https://assets-eu.researchsquare.com/files/rs-7583841/v1/a2c2673f61cf468b3dd22306.png"},{"id":96650896,"identity":"145125f2-2b29-419b-b863-1c71d58bd4c3","added_by":"auto","created_at":"2025-11-24 16:12:35","extension":"pdf","order_by":0,"title":"","display":"","copyAsset":false,"role":"manuscript-pdf","size":4263102,"visible":true,"origin":"","legend":"","description":"","filename":"manuscript.pdf","url":"https://assets-eu.researchsquare.com/files/rs-7583841/v1/d8502ec7-102f-4bf9-aeb0-e1a5713b516d.pdf"},{"id":91829055,"identity":"281e9fd6-2f74-4270-86f0-eee7a548a67b","added_by":"auto","created_at":"2025-09-22 08:54:33","extension":"docx","order_by":0,"title":"","display":"","copyAsset":false,"role":"supplement","size":1078604,"visible":true,"origin":"","legend":"","description":"","filename":"supplementarymaterials.docx","url":"https://assets-eu.researchsquare.com/files/rs-7583841/v1/712ccd48e87822074a0f3e53.docx"}],"financialInterests":"No competing interests reported.","formattedTitle":"A Murine Distal Femoral Epiphysis Ischemia Model Reveals Spatiotemporal Stratification of Necrotic Bone Marrow Clearance and Associated Inflammatory Responses","fulltext":[{"header":"Introduction","content":"\u003cp\u003eLegg–Calvé–Perthes disease (LCPD) is a pediatric avascular necrosis of the femoral head with substantial risk of long-term disability. The disease evolves from an ischemic necrosis phase into a repair phase characterized by resorption of necrotic bone, fibrovascular ingrowth, and new bone formation\u003csup\u003e(\u003cspan additionalcitationids=\"CR2\" citationid=\"CR1\" class=\"CitationRef\"\u003e1\u003c/span\u003e–\u003cspan citationid=\"CR3\" class=\"CitationRef\"\u003e3\u003c/span\u003e)\u003c/sup\u003e. Interventional evidence from a piglet model shows that percutaneous three-needle lavage of the necrotic femoral head can accelerate repair\u003csup\u003e(\u003cspan citationid=\"CR4\" class=\"CitationRef\"\u003e4\u003c/span\u003e, \u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e)\u003c/sup\u003e, supporting the concept that active removal of necrotic marrow contents can accelerate repair. However, mechanistic details of marrow clearance remain poorly defined, limiting the development of targeted therapies.\u003c/p\u003e\u003cp\u003eHistopathology\u003csup\u003e(\u003cspan citationid=\"CR2\" class=\"CitationRef\"\u003e2\u003c/span\u003e, \u003cspan citationid=\"CR6\" class=\"CitationRef\"\u003e6\u003c/span\u003e)\u003c/sup\u003e and perfusion MRI \u003csup\u003e(\u003cspan citationid=\"CR7\" class=\"CitationRef\"\u003e7\u003c/span\u003e)\u003c/sup\u003eshow that necrotic tissue abuts newly formed fibrovascular tissue, forming a distinct necrosis–repair interface that likely functions as a regulatory zone for clearing necrotic material. Because clinical specimens are scarce, the juvenile ischemic osteonecrosis (JIO) mouse model\u003csup\u003e(\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e)\u003c/sup\u003e—whose pathological trajectory parallels LCPD—has become a key tool for mechanistic study. Establishing and characterizing this interface in the JIO model is therefore essential to elucidate the cellular and molecular processes underlying marrow clearance.\u003c/p\u003e\u003cp\u003eThe direction of revascularization appears to follow native epiphyseal vascular entry points, as indicated by super-selective angiography\u003csup\u003e(\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e, \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e)\u003c/sup\u003e, perfusion MRI\u003csup\u003e(\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e, \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e)\u003c/sup\u003e, and large animal models\u003csup\u003e(\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e)\u003c/sup\u003e. In rats\u003csup\u003e(\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e)\u003c/sup\u003e, two of the four nutrient vessels enter via cartilage-free zones on the medial and lateral condyles (sites of synovial/ligament attachment) anatomically aligned with the coronal plane. We hypothesized that murine epiphyseal vasculature is similar and that coronal sectioning along the vascular entry axis would more clearly reveal the necrosis–repair interface, enabling accurate morphological analysis of marrow clearance.\u003c/p\u003e\u003cp\u003eNeutrophils and macrophages, as professional phagocytes, are central to necrotic tissue disposal. Prior work in the JIO model showed macrophage phagocytosis of necrotic fat promotes anti-inflammatory polarization\u003csup\u003e(\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e)\u003c/sup\u003e. Yet the epiphysis also contains abundant marrow cells whose necrosis releases DAMPs (e.g., nucleic acids, HMGB1, S100), likely shaping inflammation and fibrosis\u003csup\u003e(\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e)\u003c/sup\u003e, differently from adipose-derived DAMPs. While macrophage handling of necrotic adipose has been described, the clearance of necrotic marrow cells themselves and their immune sequelae remain unclear. Furthermore, how neutrophil-derived enzymes such as MPO modulate macrophage polarization at the interface is unknown. Here, we anatomically localize vascular entry sites, reconstruct the necrosis–repair interface in coronal sections, delineate marrow cell death and clearance, and map vessels, neutrophils, and macrophages in space; we also test the effect of MPO on macrophage polarization in vitro.\u003c/p\u003e\n\n\n\n\n\n\n\n\n\n\n\n\n\n"},{"header":"Materials and Methods","content":"\u003ch2\u003eAnimals\u003c/h2\u003e\u003cp\u003e All procedures were approved by the Animal Care and Welfare Committee of Guangxi Medical University and complied with institutional guidelines (approval no.202401027). Male C57BL/6 mice were used (5-week-old). Six mice were allocated for distal femoral epiphyseal vascular anatomy, and 36 mice underwent JIO surgery of the right distal femoral epiphysis. The contralateral distal femoral epiphysis served as the intact normal control.\u003c/p\u003e\u003ch3\u003eVascular anatomy of the distal femoral epiphysis\u003c/h3\u003e\u003cp\u003eBoth knees of six mice were dissected to map the epiphyseal vascular entry points. Under a stereomicroscope (6.7–40×), a medial parapatellar approach was performed on mice anesthetized with 2.5% tribromoethanol (0.2 mL/10 g, i.p.) and provided with peri-operative analgesia (buprenorphine 0.05–0.1 mg/kg s.c., ± meloxicam 1–2 mg/kg s.c.). The medial approach facilitated exposure of the popliteal vessel (PV) and the medial proximal genicular vessel (MPGV). Following a medial capsulotomy, transection of the patellar ligament, and retraction of Hoffa’s fat pad allowed for visualization of the central genicular vessel (CGV). Additionally, the lateral proximal genicular vessel (LPGV) and lateral genicular vessel (LGV) were visualized on the lateral side of the epiphysis. All epiphyseal entry points were documented using a high-resolution camera. After documentation, animals were euthanized via pentobarbital overdose.\u003c/p\u003e\u003ch3\u003eInduction of juvenile ischemic osteonecrosis\u003c/h3\u003e\u003cp\u003eTo achieve complete interruption of epiphyseal perfusion, all nutrient vessels were cauterized at their entry points into the epiphysis. Mice were anesthetized and received perioperative analgesia. They were positioned supine on a warming pad, and the right groin and hind limb were shaved. The operative field was prepared three times with povidone-iodine and 70% ethanol; ophthalmic ointment was applied to protect the eyes. Under a stereomicroscope (magnification ~ 6–40×), a medial parapatellar skin incision approximately 10–15 mm in length was made. The medial hamstrings were carefully dissected to expose and cauterize the epiphyseal branch of the popliteal vessel (PV). A medial parapatellar arthrotomy was subsequently performed to expose the medial proximal genicular (MPGV), which was cauterized prior to its entry into the epiphysis. Hoffa’s fat pad was retracted to reveal the central genicular vessel (CGV), which was also cauterized at its epiphyseal entry site. A separate lateral knee arthrotomy was then conducted to expose both the lateral proximal genicular vessel (LPGV) and lateral genicular vessel (LGV), which underwent similar cauterization at their respective epiphyseal entry points. Following these procedures, joint irrigation was performed, hemostasis confirmed, and closure of both capsule and muscles achieved using 8 − 0 monofilament nylon sutures; skin closure utilized 4 − 0 silk sutures. Postoperative analgesia consisted of buprenorphine 0.05–0.1 mg/kg s.c. every 12 h for 48 h and meloxicam 1–2 mg/kg s.c. every 24 h for 48–72 h. The left distal femoral epiphysis remained untouched as an intact normal control (refer to Fig. \u003cspan refid=\"MOESM1\" class=\"InternalRef\"\u003eS1\u003c/span\u003e for procedural steps).\u003c/p\u003e\u003ch3\u003eTissue processing\u003c/h3\u003e\u003cp\u003eAt postoperative days 1, 4, 7, 10, 14, and 28 (n = 6/time point), bilateral distal femora were harvested, fixed in 4% neutral-buffered formalin (24 h), decalcified in 14% EDTA (pH 7.4) for 10–14 days, embedded in paraffin, and sectioned coronally at 4 µm.\u003c/p\u003e\u003ch3\u003eHistology and TUNEL\u003c/h3\u003e\u003cp\u003eH\u0026amp;E staining was performed for morphology; TUNEL (Beyotime, C1098) identified apoptotic nuclei. Based on nuclear/tissue features, the interface was partitioned into: (i) \u003cb\u003eFibrotic zone (FZ)\u003c/b\u003e: Bone marrow space replaced by fibrovascular tissue with cells showing normal nuclear morphology. (ii) \u003cb\u003eResorption zone (RZ)\u003c/b\u003e: Bone marrow space filled with karyolytic cell remnants and residual cytoskeleton, with few intact nucleated cells. (iii) \u003cb\u003eNecrotic zone (NZ)\u003c/b\u003e: Bone marrow cells with pyknotic or partially karyolytic nuclei and minimal inflammatory/fibroblastic infiltration.\u003c/p\u003e\u003ch2\u003eImmunohistochemistry (IHC) and immunofluorescence (IF)\u003c/h2\u003e\u003cp\u003eIHC. Only specimens with a clear fibrotic–resorption boundary were analyzed. After deparaffinization and sodium-citrate heat retrieval (10 mM citrate, 0.05% Tween-20, pH 6.0), sections were treated with 3% H₂O₂ and blocked with 10% goat serum (Solarbio, SL038). Primary antibodies (4°C, overnight): Ly6G (1:100, BD Pharmingen 551459), F4/80 (1:200, BioLegend 123101), endomucin/EMCN (1:200, Santa Cruz Biotechnology sc-65495), MPO (1:200, ZENBIO R25062), CD206 (1:2000, Proteintech 18704-1-AP), and iNOS (1:200, Invitrogen PA1-036). HRP-labeled secondary antibodies (Beyotime: A0208 anti-rabbit, A0192 anti-rat) and DAB chromogen were used, followed by hematoxylin counterstain.\u003c/p\u003e\u003cp\u003eIF. Dual IF assessed F4/80–MPO colocalization near the interface. After antigen retrieval and blocking as above, primary antibodies were applied sequentially with matched fluorescent secondaries (1:100, Invitrogen A-11008; 1:500, Cell Signaling Technology 4417S) and DAPI nuclear stain. Images were acquired under unified exposure on fluorescence/confocal microscopy.\u003c/p\u003e\u003ch3\u003eHistomorphometry and statistics\u003c/h3\u003e\u003cp\u003eThe contralateral distal femoral epiphysis served as the intact normal control for qualitative reference (Fig.\u0026nbsp;\u003cspan refid=\"Fig2\" class=\"InternalRef\"\u003e2\u003c/span\u003eA). For semi-quantitative, within-animal comparisons on the same slides, the ipsilateral metaphysis was used as an internal control to minimize inter-animal and staining-batch variability. High-power fields (HPFs) were sampled from each zone on adjacent H\u0026amp;E, TUNEL, and IHC sections using fixed imaging settings. Quantitative endpoints included: fibrosis proportion (fibrovascular area/total marrow cavity), EMCN-positive area (% area/HPF), Ly6G- and MPO-positive cells per HPF, F4/80-positive area (% area/HPF), and hematoxylin-positive nuclear area fraction (after H–DAB color deconvolution). Regions of interest (ROIs) excluded cartilage, mineralized trabeculae, growth plate, and processing artifacts. Data are expressed as mean ± SEM. Normality was assessed by the Shapiro-Wilk test. For paired, within-animal comparisons, normally distributed data were analyzed with paired t-tests, and non-normal data with Wilcoxon signed-rank tests. Two-tailed p \u0026lt; 0.05 was considered statistically significant.\u003c/p\u003e\u003ch3\u003eRAW264.7 macrophage assay with MPO\u003c/h3\u003e\u003cp\u003eRAW264.7 cells were maintained in DMEM supplemented with 10% FBS and 1% penicillin–streptomycin at 37°C in 5% CO₂, then seeded (6-well plates for RNA; glass coverslips in 24-well plates for IF) and allowed to adhere overnight. Four conditions were applied for 24 h: control (medium only); M1 induction (LPS 100 ng/mL + IFN-γ 20 ng/mL); M1 + MPO (same M1 induction with recombinant murine MPO 1 µg/mL added at the start); and MPO alone (1 µg/mL). For immunofluorescence, cells were rinsed with PBS, fixed in 4% paraformaldehyde (15 min), permeabilized with 0.1% Triton X-100 (10 min), blocked with 10% goat serum (30 min), incubated with anti-iNOS primary antibody overnight at 4°C, then with fluorescent secondary antibody (1:100, Invitrogen A-1101, 1 h, room temperature); nuclei were counterstained with DAPI. Images were acquired under identical exposure settings, and iNOS signal was quantified as mean fluorescence intensity from ≥ 3 non-overlapping fields per well and averaged per biological replicate. For qPCR, parallel wells were lysed in TRIzol for RNA extraction, reverse-transcribed to cDNA, and analyzed by SYBR Green qPCR for iNOS and TNFα, normalized to β-Actin and calculated using the 2^−ΔΔCt method.\u003c/p\u003e"},{"header":"Results","content":"\u003cp\u003e\u003cstrong\u003e1. Vascular anatomy of the distal femoral epiphysis in mice\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eDetailed dissections in six mice mapped the epiphyseal entry points of the MPGV, CGV, LPGV, and LGV (\u003cstrong\u003eFig. 1\u003c/strong\u003e). The saphenous vessel gave rise to the popliteal trunk. The MPGV originated from the saphenous or proximal popliteal vessel, crossed the physis, and entered the medial epiphysis. The LPGV coursed along the lateral femur and supplied the lateral epiphysis through multiple branches that anastomosed with vessels in Hoffa\u0026rsquo;s fat pad. The LGV arose from the popliteal vessel and entered the lateral epiphysis. Extensive anastomoses among the MPGV, LPGV, and LGV were evident within Hoffa\u0026rsquo;s fat pad and near the lateral epiphysis. Because a medial arthrotomy alone did not fully expose the LGV, adding a lateral arthrotomy enabled direct visualization and complete cauterization of all epiphyseal vessels (Fig. S1).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e2. Histological changes during revascularization\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eCoronal sections showed progressive clearance of necrotic marrow with stepwise fibrovascular ingrowth (Fig. 2A\u0026ndash;H). On day 1, marrow cells and osteocytes across the epiphysis displayed diffuse nuclear pyknosis with no fibrous infiltration. By day 4, karyolysis appeared in marrow cavities adjacent to the medial condylar surface, while the central epiphysis remained pyknotic. On day 7, fibroblasts and small vessels infiltrated the medial marrow cavity, and the karyolytic zone extended toward the center. By day 10, fibrovascular tissue had grown in from both condyles, leaving broad transitional areas of karyolysis; pyknotic nuclei persisted near the growth plate in the epiphyseal center. By day 14, the marrow cavity was almost completely replaced by fibrovascular tissue, with new bone forming along necrotic trabeculae. By day 28, regenerated marrow cells and abundant new bone were present within the cavity. \u003cem\u003e(\u003c/em\u003eRepaired region, see Fig.S2.)\u003c/p\u003e\n\u003cp\u003eFibrosis became detectable by day 4 and rapidly progressed from day 7 to day 10. By this time, the necrotic core, absorption zone, and fibrotic margin coexisted; consequently, by day 14, the necrotic bone marrow was nearly completely replaced. Serial sections corroborated this as a continuous progressive process: initially (or within the necrotic area), the nuclei underwent pyknosis, followed by the dissolution of pyknotic nuclei as microvessels and fibroblasts infiltrated the region. This invasion formed an absorption zone characterized by anucleate \u0026quot;ghost\u0026quot; cells or residual cytoskeleton. These remnants were gradually phagocytosed, leading to the replacement of the bone marrow cavity with fibrovascular tissue and completing the clearance of necrotic bone marrow.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e3. Apoptosis and secondary necrosis of marrow cells and resident neutrophils\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eHigh-power H\u0026amp;E and TUNEL staining revealed marked regional differences in cellular morphology (Fig. 3). Cells in the fibrotic zone and in the metaphysis appeared morphologically normal. In the necrotic zone, cells exhibited nuclear pyknosis, karyorrhexis, karyolysis, or reticular chromatin, while plasma membranes remained largely intact; in contrast, the resorption zone lacked discernible nuclear structures and contained abundant eosinophilic cytoskeletal remnants with only sparse infiltration by intact nucleated cells. TUNEL signals in the necrotic zone localized to pyknotic or fragmented nuclei, and apoptotic bodies were observed (Fig. 3), consistent with apoptosis. In the resorption zone, TUNEL-positive nuclei appeared swollen with loss of chromatin, indicating insufficient clearance of apoptotic cells and their progression to secondary necrosis\u003csup\u003e(17, 18)\u003c/sup\u003e\u003cstrong\u003e.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eResident bone marrow neutrophils showed a similar spatial pattern: \u003cstrong\u003eLy6G⁺\u003c/strong\u003e cells were morphologically normal in the metaphysis, displayed \u003cstrong\u003eapoptotic nuclear changes\u003c/strong\u003e\u003cstrong\u003e\u0026nbsp;\u003c/strong\u003ein the necrotic zone, and exhibited \u003cstrong\u003enear-complete nuclear lysis with membrane disruption\u003c/strong\u003e in the resorption zone. Only scattered, morphologically normal neutrophils were seen in the fibrotic zone, and the number of Ly6G⁺ cells there was lower than in the resorption zone (Fig. 4A, B; \u003cstrong\u003ep \u0026lt; 0.05\u003c/strong\u003e). Although Ly6G⁺ counts did not differ markedly among the metaphysis, necrotic, and resorption zones, \u003cstrong\u003eMPO⁺ cells declined significantly\u003c/strong\u003e from the metaphysis to the necrotic and resorption zones (Fig. 4A, B; \u003cstrong\u003ep \u0026lt; 0.05\u003c/strong\u003e). These findings indicate that marrow stromal cells including resident neutrophils undergo \u003cstrong\u003eapoptosis\u003c/strong\u003e after ischemia, with many progressing to \u003cstrong\u003esecondary necrosis\u003c/strong\u003e when clearance is insufficient. Along the axis of \u003cstrong\u003efibrovascular ingrowth\u003c/strong\u003e, nuclear morphology delineated regions enriched in apoptotic versus secondarily necrotic cells, suggesting that the secondary necrosis process during epiphyseal clearance is \u003cstrong\u003ecoupled to\u0026mdash;and potentially regulated by\u0026mdash;fibrovascularization\u003c/strong\u003e\u003cstrong\u003e.\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e4. Fibrovascular ingrowth and chromatin clearance\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eEMCN immunostaining revealed abundant microvessels in the fibrotic, resorption, and metaphyseal zones, whereas the necrotic zone was avascular; morphologically, microvessels localized to the resorption side of the necrosis\u0026ndash;resorption interface (Fig. 4A, C). Quantitatively, the EMCN-positive area in the necrotic zone was significantly lower than in the other zones, while the hematoxylin-positive nuclear area was significantly greater than in the resorption zone (Fig. 4B, F; p \u0026lt; 0.05). These findings suggest that microvascular ingrowth within the resorption zone may facilitate dissolution and clearance of nuclear material from apoptotic cells, thereby promoting their transition to secondary necrosis.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e5.\u0026nbsp;\u003c/strong\u003e\u003cstrong\u003eMacrophage involvement in clearing neutrophil remnants\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eF4/80 immunohistochemistry showed a significantly larger positive area in the \u003cstrong\u003efibrotic\u003c/strong\u003e\u003cstrong\u003e\u0026nbsp;\u003c/strong\u003ezone than in the \u003cstrong\u003eresorption\u003c/strong\u003e zone (Fig. 4A\u0026ndash;B). Ly6G⁺ and \u003cstrong\u003eMPO⁺\u003c/strong\u003e cells were least abundant in the fibrotic zone. At high magnification, cells at the \u003cstrong\u003efibrotic\u0026ndash;resorption interface\u003c/strong\u003e were observed engulfing \u003cstrong\u003eMPO-positive cytoskeletal remnants\u003c/strong\u003e (Fig. 4D), and \u003cstrong\u003edual immunofluorescence\u003c/strong\u003e confirmed \u003cstrong\u003eF4/80\u0026ndash;MPO colocalization\u003c/strong\u003e (Fig. 4E). Together, these findings indicate that \u003cstrong\u003emacrophages at the fibrotic margin phagocytose secondarily necrotic neutrophils\u003c/strong\u003e\u003cstrong\u003e,\u003c/strong\u003e thereby promoting \u003cstrong\u003efibrovascular replacement\u003c/strong\u003e of the marrow cavity.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003e6. Macrophage polarization at the fibrotic\u0026ndash;resorption interface; MPO promotes M1 polarization in vitro\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eIHC analysis revealed a predominance of iNOS⁺ (M1) macrophages over CD206⁺ (M2) macrophages at the fibrotic\u0026ndash;resorption interface (Fig.5A,B;P \u0026lt; 0.05), indicating that M1 cells are frequently clustered in proximity to MPO-rich regions. In RAW264.7 cells, addition of recombinant MPO (1 \u0026mu;g/mL) during LPS+IFN-\u0026gamma;\u0026ndash;driven M1 induction (vs. LPS+IFN-\u0026gamma; alone) increased iNOS immunofluorescence and upregulated iNOS and TNF\u0026alpha; mRNA (qPCR; all P \u0026lt; 0.05) (Fig. 5C,D,E; P \u0026lt; 0.05).\u003c/p\u003e"},{"header":"Discussion","content":"\u003cdiv id=\"Sec13\" class=\"Section2\"\u003e\u003ch2\u003eVascular anatomy and model refinement\u003c/h2\u003e\u003cp\u003eThe anatomical sites where vessels enter the epiphysis may determine the pattern of revascularization in necrotic epiphyses. Evidence from super-selective arterial perfusion imaging\u003csup\u003e(\u003cspan citationid=\"CR9\" class=\"CitationRef\"\u003e9\u003c/span\u003e, \u003cspan citationid=\"CR10\" class=\"CitationRef\"\u003e10\u003c/span\u003e)\u003c/sup\u003e, perfusion MRI\u003csup\u003e(\u003cspan citationid=\"CR11\" class=\"CitationRef\"\u003e11\u003c/span\u003e, \u003cspan citationid=\"CR12\" class=\"CitationRef\"\u003e12\u003c/span\u003e)\u003c/sup\u003e, and juvenile pig models \u003csup\u003e(\u003cspan citationid=\"CR13\" class=\"CitationRef\"\u003e13\u003c/span\u003e)\u003c/sup\u003e supports this view. In 2015, Kim et al. \u003csup\u003e(\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e)\u003c/sup\u003e used micro-CT to delineate the major arterial trunks supplying the murine distal femoral epiphysis, providing an anatomical basis for constructing the JIO mouse model. However, like other studies of murine hind-limb vasculature\u003csup\u003e(\u003cspan citationid=\"CR19\" class=\"CitationRef\"\u003e19\u003c/span\u003e)\u003c/sup\u003e that work did not clearly resolve epiphyseal entry points and thus could not elucidate the revascularization pattern of the necrotic murine epiphysis. In 2021, Ma et al. \u003csup\u003e(\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e)\u003c/sup\u003e mapped the nutrient vessels of the distal femoral epiphysis in a rat ischemic necrosis model and, on subsequent micro-CT during the repair phase, observed that neovessels preferentially entered through cartilage-free zones on the medial and lateral condyles. In the present study, we anatomically defined the epiphyseal entry points in mice, confirmed a murine\u0026ndash;rat similarity in epiphyseal vascular anatomy, and identified a more extensive lateral vascular network with prominent anastomoses than previously reported. These findings provide additional anatomical context for subsequent studies of epiphyseal necrosis and repair.\u003c/p\u003e\u003cp\u003eMapping the epiphyseal vascular entry points clarified revascularization routes and justified adding a lateral approach to ensure complete interruption of perfusion. Coronal sectioning aligned with this entry axis consistently reconstructed the necrosis\u0026ndash;repair interface and its centripetal advance; by days 7\u0026ndash;10, a reproducible necrotic\u0026ndash;resorption\u0026ndash;fibrotic triad was evident. Building on this framework, temporal sequencing and zonal criteria delineated a distinct spatiotemporal stratification with a front that progresses from the periphery toward the center. The continuous, zone-specific shifts in nuclear and matrix features indicate that clearance is not uniform but proceeds in staged steps tightly coupled to vascular ingress or the local oxygen gradient and humoral factor.\u003c/p\u003e\u003c/div\u003e\u003cdiv id=\"Sec14\" class=\"Section2\"\u003e\u003ch2\u003ePatterns of necrosis and tissue clearance\u003c/h2\u003e\u003cp\u003eThe mode of cell death dictates its clearance pathway. Although direct evidence from human Perthes disease tissue is lacking, porcine models indicate that apoptosis and oncosis (primary necrosis) coexist after epiphyseal ischemia\u003csup\u003e(\u003cspan citationid=\"CR20\" class=\"CitationRef\"\u003e20\u003c/span\u003e)\u003c/sup\u003e. Importantly, oncosis and secondary necrosis are morphologically similar at their terminal stages\u0026mdash;cell swelling, loss of chromatin/nuclear staining, and \u0026ldquo;ghost cells\u0026rdquo;\u0026mdash;and are therefore difficult to distinguish on static sections\u003csup\u003e(\u003cspan citationid=\"CR21\" class=\"CitationRef\"\u003e21\u003c/span\u003e)\u003c/sup\u003e. In JIO mouse\u003csup\u003e(\u003cspan citationid=\"CR8\" class=\"CitationRef\"\u003e8\u003c/span\u003e)\u003c/sup\u003e and rat models\u003csup\u003e(\u003cspan citationid=\"CR14\" class=\"CitationRef\"\u003e14\u003c/span\u003e)\u003c/sup\u003e, loss of marrow nuclear staining is commonly observed about one week after modeling; the diagnostic key, however, is not the end-stage morphology but whether an apoptotic phase precedes the anucleate state.\u003c/p\u003e\u003cp\u003eIn our spatiotemporal analyses, apoptotic cells were evident on postoperative day 1 and within the necrotic core, whereas abundant anucleate \u0026ldquo;ghost cells\u0026rdquo; emerged later, specifically within the resorption zone. We therefore infer that anucleate cells in the resorption zone primarily derive from uncleared apoptotic cells that progressed to secondary necrosis, rather than from primary oncosis. Temporally, marrow changes previously labeled as \u0026ldquo;oncosis\u0026rdquo; in JIO mice and juvenile pigs tend to occur at later time points, aligning more closely with the kinetics of secondary necrosis (late apoptotic lysis). Thus, a portion of the oncosis-like morphology reported in prior studies likely includes\u0026mdash;or predominantly reflects\u0026mdash;secondary necrosis of apoptotic cells.\u003c/p\u003e\u003c/div\u003e\u003cdiv id=\"Sec15\" class=\"Section2\"\u003e\u003ch2\u003eMechanisms and cellular clearance of secondary necrosis\u003c/h2\u003e\u003cp\u003eSecondary necrosis represents a programmed lytic transition of uncleared apoptotic cells rather than passive decay. Mechanistically, caspase-3 cleavage of gasdermin E \u003cb\u003e(GSDME/DFNA5)\u003c/b\u003e generates an N-terminal fragment that forms large pores in the plasma membrane, priming apoptotic cells for lysis\u0026mdash;one of the key molecular switches from apoptosis to secondary necrosis\u003csup\u003e(\u003cspan citationid=\"CR22\" class=\"CitationRef\"\u003e22\u003c/span\u003e, \u003cspan citationid=\"CR23\" class=\"CitationRef\"\u003e23\u003c/span\u003e)\u003c/sup\u003e. Ninjurin-1 (\u003cb\u003eNINJ1\u003c/b\u003e) then executes plasma membrane rupture \u003cb\u003e(PMR)\u003c/b\u003e as a shared terminal effector in late apoptosis, pyroptosis, and necrosis, enabling bulk release of cytosolic contents and DAMPs \u003csup\u003e(\u003cspan citationid=\"CR24\" class=\"CitationRef\"\u003e24\u003c/span\u003e, \u003cspan citationid=\"CR25\" class=\"CitationRef\"\u003e25\u003c/span\u003e)\u003c/sup\u003e. plasma membrane channel pannexin 1(\u003cb\u003ePANX1\u003c/b\u003e) channels, beyond mediating \u0026ldquo;find-me\u0026rdquo; signals, can remain open to disturb ionic homeostasis and promote late cell lysis \u003csup\u003e(\u003cspan citationid=\"CR26\" class=\"CitationRef\"\u003e26\u003c/span\u003e)\u003c/sup\u003e. In parallel, chromatin is dismantled in an ordered fashion: (caspase-activated deoxyribonulease) \u003cb\u003eCAD\u003c/b\u003e is activated by caspase-3\u0026ndash;mediated (Inhibitor of Caspase-Activated DNase )\u003cb\u003eICAD\u003c/b\u003e cleavage to initiate intracellular nucleosomal degradation\u003csup\u003e(\u003cspan citationid=\"CR27\" class=\"CitationRef\"\u003e27\u003c/span\u003e)\u003c/sup\u003e, while extracellular deoxyribonuclease I \u003cb\u003e(DNase I)\u0026ndash;family enzymes\u003c/b\u003e (e.g., DNase I, DNase γ) further process released nucleosomes; if clearance is inadequate, the accumulation of nucleosomal DNA and histones amplifies inflammation.\u003c/p\u003e\u003cp\u003eIntegrating prior work with our data, When revascularization reaches the necrosis\u0026ndash;repair interface, it delivers humoral factors\u003csup\u003e(\u003cspan citationid=\"CR28\" class=\"CitationRef\"\u003e28\u003c/span\u003e)\u003c/sup\u003e (e.g., DNase I \u003csup\u003e(\u003cspan citationid=\"CR29\" class=\"CitationRef\"\u003e29\u003c/span\u003e)\u003c/sup\u003e, complement\u003csup\u003e(\u003cspan citationid=\"CR30\" class=\"CitationRef\"\u003e30\u003c/span\u003e)\u003c/sup\u003eand natural antibodies\u003csup\u003e(\u003cspan citationid=\"CR31\" class=\"CitationRef\"\u003e31\u003c/span\u003e)\u003c/sup\u003e) and cellular effectors\u003csup\u003e\u003cb\u003e(\u003cspan citationid=\"CR32\" class=\"CitationRef\"\u003e32\u003c/span\u003e)\u003c/b\u003e\u003c/sup\u003e ( neutrophils, macrophages) while restoring oxygen/ion flux and perfusion. These may inputs destabilize membrane homeostasis, amplify caspase-3/GSDME pore formation and PANX1 activity, and, via NINJ1, trigger PMR\u0026mdash;converting \u0026ldquo;silent\u0026rdquo; apoptosis into secondary necrosis on the revascularized (resorption) side. Concurrently, DNases accelerate chromatin breakdown, macrophages/fibroblasts remove residual cytoskeleton, and microcirculation washes out soluble DAMPs and degradation products, shifting the milieu from inflammatory to reparative. This framework explains why secondary necrosis follows vessel ingrowth in time and localizes to the vascular side in space, yielding the observed centripetal triad of necrotic core \u0026rarr; resorption front \u0026rarr; fibrotic zone.\u003c/p\u003e\u003c/div\u003e\u003cdiv id=\"Sec16\" class=\"Section2\"\u003e\u003ch2\u003eImmune regulation of neutrophil secondary necrosis\u003c/h2\u003e\u003cp\u003eOur study shows that neutrophils in the resorption zone undergo secondary necrosis. Although secondarily necrotic neutrophils also lose plasma-membrane integrity, they are less pro-inflammatory than cells that undergo primary necrosis \u003csup\u003e(\u003cspan citationid=\"CR33\" class=\"CitationRef\"\u003e33\u003c/span\u003e)\u003c/sup\u003e, in part because the inflammatory activity of their released DAMPs is attenuated by apoptosis-associated degradative processes \u003csup\u003e(\u003cspan citationid=\"CR34\" class=\"CitationRef\"\u003e34\u003c/span\u003e)\u003c/sup\u003e. Fractionation of lysed neutrophils into membrane and soluble components indicates that the membrane fraction behaves like that of apoptotic cells\u0026mdash;anti-inflammatory via phosphatidylserine (PS)\u0026mdash;whereas proteases released during lysis are pro-inflammatory \u003csup\u003e(\u003cspan citationid=\"CR35\" class=\"CitationRef\"\u003e35\u003c/span\u003e)\u003c/sup\u003e. While some reports show anti-inflammatory effects of neutrophil α-defensins released from apoptotic/necrotic neutrophils\u003csup\u003e(\u003cspan citationid=\"CR36\" class=\"CitationRef\"\u003e36\u003c/span\u003e, \u003cspan citationid=\"CR37\" class=\"CitationRef\"\u003e37\u003c/span\u003e)\u003c/sup\u003e, other soluble factors can exacerbate inflammation: neutrophil elastase can cleave PS receptors and impair efferocytosis \u003csup\u003e(\u003cspan citationid=\"CR38\" class=\"CitationRef\"\u003e38\u003c/span\u003e)\u003c/sup\u003e, and secondarily necrotic neutrophils selectively release IL-16C and macrophage migration inhibitory factor(MIF) degradation to promote inflammatory responses\u003csup\u003e(\u003cspan citationid=\"CR39\" class=\"CitationRef\"\u003e39\u003c/span\u003e, \u003cspan citationid=\"CR40\" class=\"CitationRef\"\u003e40\u003c/span\u003e)\u003c/sup\u003e. In our study, whether MIF accounts for the relative paucity of macrophages in the resorption and necrotic zones remains unclear. Nevertheless, we observed MPO-rich fibrotic\u0026ndash;resorption interfaces dominated by M1 (iNOS⁺) macrophages, and recombinant MPO promoted M1 polarization of RAW264.7 cells in vitro, supporting a pro-inflammatory role of MPO during necrotic marrow clearance. Whether MPO, IL-16C, and MIF can serve as inflammatory biomarkers of epiphyseal necrosis resolution warrants further investigation.\u003c/p\u003e\u003cp\u003eMultiple lines of evidence implicate DAMP\u0026ndash;PRR signaling in Perthes disease. Synovial studies show chronic synovitis with elevated IL-6 and increased HMGB1 that correlates with IL-6\u003csup\u003e(\u003cspan citationid=\"CR41\" class=\"CitationRef\"\u003e41\u003c/span\u003e, \u003cspan citationid=\"CR42\" class=\"CitationRef\"\u003e42\u003c/span\u003e)\u003c/sup\u003e Necrotic bone activates macrophages via TLR4 \u003csup\u003e(\u003cspan citationid=\"CR43\" class=\"CitationRef\"\u003e43\u003c/span\u003e)\u003c/sup\u003e, while pharmacologic TLR4 inhibition accelerates epiphyseal repair \u003csup\u003e(\u003cspan citationid=\"CR44\" class=\"CitationRef\"\u003e44\u003c/span\u003e)\u003c/sup\u003e. In a juvenile pig model, minimally invasive lavage of necrotic marrow improved bone healing \u003csup\u003e(\u003cspan citationid=\"CR5\" class=\"CitationRef\"\u003e5\u003c/span\u003e)\u003c/sup\u003e. DAMPs from necrotic femoral heads suppress osteogenesis and promote mesenchymal-stem-cell fibrosis\u003csup\u003e(\u003cspan citationid=\"CR16\" class=\"CitationRef\"\u003e16\u003c/span\u003e)\u003c/sup\u003e, and macrophage clearance of necrotic fat drives anti-inflammatory polarization \u003csup\u003e(\u003cspan citationid=\"CR15\" class=\"CitationRef\"\u003e15\u003c/span\u003e)\u003c/sup\u003e. Yet how necrotic marrow cells\u0026mdash;and the DAMPs they release\u0026mdash;are cleared remains unclear. Our data reveal secondary necrosis of resident neutrophils at the necrosis\u0026ndash;repair interface and suggest that targeting the removal of necrotic marrow cells, particularly secondarily necrotic neutrophils, may hasten inflammatory resolution and improve epiphyseal repair.\u003c/p\u003e\u003c/div\u003e\u003cdiv id=\"Sec17\" class=\"Section2\"\u003e\u003ch2\u003eLimitations\u003c/h2\u003e\u003cp\u003eAlthough our detailed anatomical model of the intraosseous epiphyseal plate (JIO) and the histopathological atlas of the necrosis-repair interface have advantages, this study still has several limitations. We only used male C57BL/6 mice, which may limit the generalizability of our findings to human LCPD. Most of the evidence was derived from static decalcified paraffin sections analyzed by semi-quantitative H\u0026amp;E/IHC/IF methods; in vivo imaging and functional readouts were not included in this study. Although the observed colocalization and regional associations are relevant, causal inferences still need to be established by genetic or pharmacological intervention of MPO, GSDME, NINJ1, complement components, deoxyribonuclease I (DNase I), and IL-16C/MIF, and direct assessment of phagocytic efficiency, DAMPs profiles, and cytokine kinetics. Finally, we did not evaluate biomechanical strength, three-dimensional microstructure, or therapeutic endpoints of intervention.\u003c/p\u003e\u003c/div\u003e"},{"header":"Conclusions","content":"\u003cp\u003eThe necrosis\u0026ndash;repair interface in the refined murine JIO model demonstrates that necrotic marrow clearance proceeds through a distinct, outward-to-inward tri-zonal program. Apoptotic marrow cells\u0026mdash;including resident neutrophils\u0026mdash;shift to secondary necrosis as revascularization advances, and the resulting remnants are eliminated during fibrovascular replacement. Neutrophil-derived MPO is spatially associated with M1-polarized macrophages in vivo and augments M1 polarization in vitro, suggesting an immunoregulatory neutrophil\u0026ndash;macrophage axis at the interface. Strategies that enhance clearance of secondarily necrotic neutrophils and modulate this axis may accelerate inflammatory resolution and improve epiphyseal repair.\u003c/p\u003e"},{"header":"Declarations","content":"\u003cp\u003e\u003cstrong\u003eEthics approval and consent to participate\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe animal experiment was approved by the Animal Care \u0026amp; Welfare Committee of Guangxi Medical University (approval no.202401027; Nanning, China).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eConsent for publication\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThis paper is approved by all authors for publication.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAvailability of data and materials\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eCompeting interests\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare no competing interests.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eFunding\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eThe authors declare that financial support was received for the research, authorship, and/or publication of this article. This work was supported by the Basic Research Capacity Enhancement Program for Young and Middle-aged Faculty at Universities and Colleges in Guangxi (Grant No. 2025KY0130); the Guangxi Natural Science Foundation—Joint Project on Regional High-Incidence Diseases Research (Grant No. 2023GXNSFAA026342); the Featured Innovation Team Cultivation Project of the First Affiliated Hospital of Guangxi Medical University (Grant No. YYZS2024002); the Youth Science and Technology Inspiration Star Program of the First Affiliated Hospital of Guangxi Medical University (Grant No. YYZS2022008); and the National Natural Science Foundation of China (Grant No. 82460422).\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAuthor contributions\u0026nbsp;\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eMZ designed the research and wrote the original draft.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eZF, ZH, ST, JL and ZL performed the experiments.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eQH: Methodology, Project administration\u003c/p\u003e\n\u003cp\u003eBL performed the analysis.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eXD and SL: Project administration, Supervision, Funding acquisition.\u003c/p\u003e\n\u003cp\u003eAll authors read and approved the final manuscript.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003eThe authors declare that all data were generated in-house, that no paper mill was used, and that no AI tool has been used for the generation of text or figures.\u0026nbsp;\u003c/p\u003e\n\u003cp\u003e\u003cstrong\u003eAcknowledgements\u003c/strong\u003e\u003c/p\u003e\n\u003cp\u003eWe thank the Animal Care \u0026amp; Welfare Core and institutional Histology/Imaging facilities for routine support. No professional writing services were used, and no materials were received as in-kind gifts; all experiments, analyses, and manuscript preparation were performed by the authors. All acknowledged units/cores have consented to be named.\u003c/p\u003e"},{"header":"References","content":"\u003col\u003e\n \u003cli\u003eKim HK. Pathophysiology and new strategies for the treatment of Legg-Calv\u0026eacute;-Perthes disease. J Bone Joint Surg Am. 2012;94(7):659-69.\u003c/li\u003e\n \u003cli\u003eJonsater S. Coxa plana; a histo-pathologic and arthrografic study. Acta Orthop Scand Suppl. 1953;12:5-98.\u003c/li\u003e\n \u003cli\u003eCatterall A, Pringle J, Byers PD, Fulford GE, Kemp HB, Dolman CL, et al. A review of the morphology of Perthes\u0026apos; disease. J Bone Joint Surg Br. 1982;64(3):269-75.\u003c/li\u003e\n \u003cli\u003eAlves do Monte F, Sung Park M, Gokani V, Singhal M, Ma C, Aruwajoye OO, et al. Development of a novel minimally invasive technique to washout necrotic bone marrow content from epiphyseal bone: A preliminary cadaveric bone study. Orthop Traumatol Surg Res. 2020;106(4):709-15.\u003c/li\u003e\n \u003cli\u003eKim HKW, Park MS, Alves do Monte F, Gokani V, Aruwajoye OO, Ren Y. Minimally Invasive Necrotic Bone Washing Improves Bone Healing After Femoral Head Ischemic Osteonecrosis: An Experimental Investigation in Immature Pigs. J Bone Joint Surg Am. 2021;103(13):1193-202.\u003c/li\u003e\n \u003cli\u003eDolman CL, Bell HM. The pathology of Legg-Calv\u0026eacute;-Perthes disease. A case report. J Bone Joint Surg Am. 1973;55(1):184-8.\u003c/li\u003e\n \u003cli\u003eKim HK, Burgess J, Thoveson A, Gudmundsson P, Dempsey M, Jo CH. Assessment of Femoral Head Revascularization in Legg-Calv\u0026eacute;-Perthes Disease Using Serial Perfusion MRI. J Bone Joint Surg Am. 2016;98(22):1897-904.\u003c/li\u003e\n \u003cli\u003eKamiya N, Yamaguchi R, Aruwajoye O, Adapala NS, Kim HK. Development of a mouse model of ischemic osteonecrosis. Clin Orthop Relat Res. 2015;473(4):1486-98.\u003c/li\u003e\n \u003cli\u003eAtsumi T, Yamano K, Muraki M, Yoshihara S, Kajihara T. The blood supply of the lateral epiphyseal arteries in Perthes\u0026apos; disease. J Bone Joint Surg Br. 2000;82(3):392-8.\u003c/li\u003e\n \u003cli\u003eAtsumi T, Yoshihara S, Hiranuma Y. Revascularization of the artery of the ligamentum teres in Perthes disease. Clin Orthop Relat Res. 2001(386):210-7.\u003c/li\u003e\n \u003cli\u003eMorris WZ, Valencia AA, McGuire MF, Kim HKW. The Role of the Artery of Ligamentum Teres in Revascularization in Legg-Calve-Perthes Disease. J Pediatr Orthop. 2022;42(4):175-8.\u003c/li\u003e\n \u003cli\u003eLamer S, Dorgeret S, Khairouni A, Mazda K, Brillet PY, Bacheville E, et al. Femoral head vascularisation in Legg-Calv\u0026eacute;-Perthes disease: comparison of dynamic gadolinium-enhanced subtraction MRI with bone scintigraphy. Pediatr Radiol. 2002;32(8):580-5.\u003c/li\u003e\n \u003cli\u003eKim HK, Bian H, Aya-ay J, Garces A, Morgan EF, Gilbert SR. Hypoxia and HIF-1alpha expression in the epiphyseal cartilage following ischemic injury to the immature femoral head. Bone. 2009;45(2):280-8.\u003c/li\u003e\n \u003cli\u003eMa C, Andre G, Edwards D, Kim HKW. A rat model of ischemic osteonecrosis for investigating local therapeutics using biomaterials. Acta Biomater. 2021;132:260-71.\u003c/li\u003e\n \u003cli\u003eDeng Z, Kim HKW, Hernandez PA, Ren Y. Fat Phagocytosis Promotes Anti-Inflammatory Responses of Macrophages in a Mouse Model of Osteonecrosis. Cells. 2024;13(14).\u003c/li\u003e\n \u003cli\u003eDeng Z, Ren Y, Park MS, Kim HKW. Damage associated molecular patterns in necrotic femoral head inhibit osteogenesis and promote fibrogenesis of mesenchymal stem cells. Bone. 2022;154:116215.\u003c/li\u003e\n \u003cli\u003eSilva MT, do Vale A, dos Santos NM. Secondary necrosis in multicellular animals: an outcome of apoptosis with pathogenic implications. Apoptosis. 2008;13(4):463-82.\u003c/li\u003e\n \u003cli\u003eSilva MT. Secondary necrosis: the natural outcome of the complete apoptotic program. FEBS Lett. 2010;584(22):4491-9.\u003c/li\u003e\n \u003cli\u003eKochi T, Imai Y, Takeda A, Watanabe Y, Mori S, Tachi M, et al. Characterization of the arterial anatomy of the murine hindlimb: functional role in the design and understanding of ischemia models. PLoS One. 2013;8(12):e84047.\u003c/li\u003e\n \u003cli\u003eKothapalli R, Aya-ay JP, Bian H, Garces A, Kim HK. Ischaemic injury to femoral head induces apoptotic and oncotic cell death. Pathology. 2007;39(2):241-6.\u003c/li\u003e\n \u003cli\u003eWeerasinghe P, Buja LM. Oncosis: an important non-apoptotic mode of cell death. Exp Mol Pathol. 2012;93(3):302-8.\u003c/li\u003e\n \u003cli\u003eRogers C, Fernandes-Alnemri T, Mayes L, Alnemri D, Cingolani G, Alnemri ES. Cleavage of DFNA5 by caspase-3 during apoptosis mediates progression to secondary necrotic/pyroptotic cell death. Nat Commun. 2017;8:14128.\u003c/li\u003e\n \u003cli\u003eDe Schutter E, Ramon J, Pfeuty B, De Tender C, Stremersch S, Raemdonck K, et al. Plasma membrane perforation by GSDME during apoptosis-driven secondary necrosis. Cell Mol Life Sci. 2021;79(1):19.\u003c/li\u003e\n \u003cli\u003eDondelinger Y, Priem D, Huyghe J, Delanghe T, Vandenabeele P, Bertrand MJM. NINJ1 is activated by cell swelling to regulate plasma membrane permeabilization during regulated necrosis. Cell Death Dis. 2023;14(11):755.\u003c/li\u003e\n \u003cli\u003eKayagaki N, Kornfeld OS, Lee BL, Stowe IB, O\u0026apos;Rourke K, Li Q, et al. NINJ1 mediates plasma membrane rupture during lytic cell death. Nature. 2021;591(7848):131-6.\u003c/li\u003e\n \u003cli\u003eChekeni FB, Elliott MR, Sandilos JK, Walk SF, Kinchen JM, Lazarowski ER, et al. Pannexin 1 channels mediate \u0026apos;find-me\u0026apos; signal release and membrane permeability during apoptosis. Nature. 2010;467(7317):863-7.\u003c/li\u003e\n \u003cli\u003ePorter AG, J\u0026auml;nicke RU. Emerging roles of caspase-3 in apoptosis. Cell Death Differ. 1999;6(2):99-104.\u003c/li\u003e\n \u003cli\u003eSchuermans S, Kestens C, Marques PE. Systemic mechanisms of necrotic cell debris clearance. Cell Death Dis. 2024;15(8):557.\u003c/li\u003e\n \u003cli\u003eSchuermans S, Quanico J, Kestens C, Vandendriessche S, Slowikowski E, Crijns ML, et al. Degradation rather than disassembly of necrotic debris is essential to enhance recovery after acute liver injury. Cell Mol Life Sci. 2025;82(1):190.\u003c/li\u003e\n \u003cli\u003eVandendriessche S, Mattos MS, Bialek EL, Schuermans S, Proost P, Marques PE. Complement activation drives the phagocytosis of necrotic cell debris and resolution of liver injury. Front Immunol. 2024;15:1512470.\u003c/li\u003e\n \u003cli\u003eMattos MS, Vandendriessche S, Schuermans S, Feyaerts L, H\u0026ouml;velmeyer N, Waisman A, et al. Natural antibodies are required for clearance of necrotic cells and recovery from acute liver injury. JHEP Rep. 2024;6(4):101013.\u003c/li\u003e\n \u003cli\u003ePhipps MC, Huang Y, Yamaguchi R, Kamiya N, Adapala NS, Tang L, et al. In vivo monitoring of activated macrophages and neutrophils in response to ischemic osteonecrosis in a mouse model. J Orthop Res. 2016;34(2):307-13.\u003c/li\u003e\n \u003cli\u003eLoh W, Vermeren S. Anti-Inflammatory Neutrophil Functions in the Resolution of Inflammation and Tissue Repair. Cells. 2022;11(24).\u003c/li\u003e\n \u003cli\u003eSachet M, Liang YY, Oehler R. The immune response to secondary necrotic cells. Apoptosis. 2017;22(10):1189-204.\u003c/li\u003e\n \u003cli\u003eFadok VA, Bratton DL, Guthrie L, Henson PM. Differential effects of apoptotic versus lysed cells on macrophage production of cytokines: role of proteases. J Immunol. 2001;166(11):6847-54.\u003c/li\u003e\n \u003cli\u003eMiles K, Clarke DJ, Lu W, Sibinska Z, Beaumont PE, Davidson DJ, et al. Dying and necrotic neutrophils are anti-inflammatory secondary to the release of alpha-defensins. J Immunol. 2009;183(3):2122-32.\u003c/li\u003e\n \u003cli\u003eBrook M, Tomlinson GH, Miles K, Smith RW, Rossi AG, Hiemstra PS, et al. Neutrophil-derived alpha defensins control inflammation by inhibiting macrophage mRNA translation. Proc Natl Acad Sci U S A. 2016;113(16):4350-5.\u003c/li\u003e\n \u003cli\u003eVandivier RW, Fadok VA, Hoffmann PR, Bratton DL, Penvari C, Brown KK, et al. Elastase-mediated phosphatidylserine receptor cleavage impairs apoptotic cell clearance in cystic fibrosis and bronchiectasis. J Clin Invest. 2002;109(5):661-70.\u003c/li\u003e\n \u003cli\u003eRoth S, Agthe M, Eickhoff S, M\u0026ouml;ller S, Karsten CM, Borregaard N, et al. Secondary necrotic neutrophils release interleukin-16C and macrophage migration inhibitory factor from stores in the cytosol. Cell Death Discov. 2015;1:15056.\u003c/li\u003e\n \u003cli\u003eRoth S, Solbach W, Laskay T. IL-16 and MIF: messengers beyond neutrophil cell death. Cell Death Dis. 2016;7(1):e2049.\u003c/li\u003e\n \u003cli\u003eKamiya N, Yamaguchi R, Adapala NS, Chen E, Neal D, Jack O, et al. Legg-Calv\u0026eacute;-Perthes disease produces chronic hip synovitis and elevation of interleukin-6 in the synovial fluid. J Bone Miner Res. 2015;30(6):1009-13.\u003c/li\u003e\n \u003cli\u003eKamiya N, Kim HK. Elevation of Proinflammatory Cytokine HMGB1 in the Synovial Fluid of Patients With Legg-Calv\u0026eacute;-Perthes Disease and Correlation With IL-6. JBMR Plus. 2021;5(2):e10429.\u003c/li\u003e\n \u003cli\u003eAdapala NS, Yamaguchi R, Phipps M, Aruwajoye O, Kim HKW. Necrotic Bone Stimulates Proinflammatory Responses in Macrophages through the Activation of Toll-Like Receptor 4. Am J Pathol. 2016;186(11):2987-99.\u003c/li\u003e\n \u003cli\u003eYu R, Ma C, Li G, Xu J, Feng D, Lan X. Inhibition of Toll-Like Receptor 4 Signaling Pathway Accelerates the Repair of Avascular Necrosis of Femoral Epiphysis through Regulating Macrophage Polarization in Perthes Disease. Tissue Eng Regen Med. 2023;20(3):489-501.\u003c/li\u003e\n\u003c/ol\u003e"}],"fulltextSource":"","fullText":"","funders":[],"hasAdminPriorityOnWorkflow":false,"hasManuscriptDocX":true,"hasOptedInToPreprint":true,"hasPassedJournalQc":"","hasAnyPriority":false,"hideJournal":false,"highlight":"","institution":"","isAcceptedByJournal":true,"isAuthorSuppliedPdf":false,"isDeskRejected":"","isHiddenFromSearch":false,"isInQc":false,"isInWorkflow":false,"isPdf":false,"isPdfUpToDate":true,"isWithdrawnOrRetracted":false,"journal":{"display":true,"email":"[email protected]","identity":"journal-of-orthopaedic-surgery-and-research","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"josr","sideBox":"Learn more about [Journal of Orthopaedic Surgery and Research](http://josr-online.biomedcentral.com)","snPcode":"13018","submissionUrl":"https://submission.nature.com/new-submission/13018/3","title":"Journal of Orthopaedic Surgery and Research","twitterHandle":"@MSKmedBMC","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true},"keywords":"epiphyseal ischemia, Damage-associated molecular patterns (DAMPs), secondary necrosis, neutrophil, macrophage polarization, myeloperoxidase, revascularization, mouse model","lastPublishedDoi":"10.21203/rs.3.rs-7583841/v1","lastPublishedDoiUrl":"https://doi.org/10.21203/rs.3.rs-7583841/v1","license":{"name":"CC BY 4.0","url":"https://creativecommons.org/licenses/by/4.0/"},"manuscriptAbstract":"\u003cp\u003e\u003cstrong\u003eBackground:\u003c/strong\u003e Efficient clearance of necrotic marrow is essential for repair after epiphyseal ischemia, yet the tissue-level spatiotemporal evolution of this process and its immunoregulatory mechanisms remain incompletely defined. \u003cbr\u003e\n \u003cstrong\u003eMethods\u003c/strong\u003e: Using a refined juvenile ischemic osteonecrosis (JIO) model of the murine distal femoral epiphysis—with medial plus lateral approaches guided by mapped vascular entry points—we fully interrupted epiphyseal perfusion and harvested tissue at postoperative days 1–28. Coronal sections along the vascular entry axis underwent H\u0026amp;E/TUNEL and zonal IHC/IF (Ly6G, myeloperoxidase (MPO), F4/80, EMCN, iNOS, CD206). An in-vitro assay tested whether recombinant MPO(1 μg/mL) modulates LPS+IFN-γ–induced M1 polarization in RAW264.7 cells.\u003cbr\u003e\n \u003cstrong\u003eResults\u003c/strong\u003e: We consistently observed a necrosis-repair interface characterized by a centripetal tri-zonal progression: fibrotic margin →resorption front → necrotic core. Apoptotic marrow cells—including resident neutrophils—transitioned into secondary necrosis within the resorption zone. This zone demonstrated enrichment of EMCN⁺ microvessels alongside a reduction in hematoxylin-positive nuclear material, indicative of chromatin disposal supported by revascularization processes. At the fibrotic margin, F4/80⁺ macrophages engulfed MPO⁺ , and iNOS⁺ (M1) macrophages outnumbered CD206⁺ (M2). Furthermore, recombinant MPO enhanced M1 polarization of RAW264.7 cells in vitro. \u003cbr\u003e\n \u003cstrong\u003eConclusions\u003c/strong\u003e: The JIO necrosis-repair interface adheres to a distinct spatiotemporal program in which apoptotic cells transition into secondary necrosis concurrently with revascularization efforts. Cells undergoing secondary necrosis are subsequently eliminated during the fibrovascular replacement processes. Notably, neutrophil-derived MPO spatially associates with M1-skewed macrophages in vivo and enhances M1 polarization in vitro; this suggests an immunoregulatory axis exists between neutrophils and macrophages at the interface. Strategies aimed at enhancing the clearance of secondary necrosis neutrophils and modulating this axis may facilitate inflammatory resolution and improve epiphyseal repair.\u003c/p\u003e","manuscriptTitle":"A Murine Distal Femoral Epiphysis Ischemia Model Reveals Spatiotemporal Stratification of Necrotic Bone Marrow Clearance and Associated Inflammatory Responses","msid":"","msnumber":"","nonDraftVersions":[{"code":1,"date":"2025-09-22 08:54:28","doi":"10.21203/rs.3.rs-7583841/v1","editorialEvents":[{"type":"communityComments","content":0},{"type":"decision","content":"Revision requested","date":"2025-10-24T05:16:33+00:00","index":"","fulltext":""},{"type":"reviewerAgreed","content":"56960936748751252206581075296769810139","date":"2025-10-16T11:50:26+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-10-13T14:48:54+00:00","index":"hide","fulltext":""},{"type":"editorInvitedReview","content":"","date":"2025-10-13T05:14:47+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"184366508225410424521717564728502011332","date":"2025-10-13T01:43:19+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"91789077356679471970402943181242468129","date":"2025-10-10T14:33:02+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"202827012487901019112774741188995454357","date":"2025-10-10T10:22:18+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"167896230682930338687409153467976648450","date":"2025-10-10T08:48:02+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"298598353933703655827060365038791685666","date":"2025-09-18T17:20:21+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"82952668186920541704392485628926785058","date":"2025-09-16T22:03:54+00:00","index":"hide","fulltext":""},{"type":"reviewerAgreed","content":"144847523346980127561250502404577740976","date":"2025-09-15T18:16:13+00:00","index":"hide","fulltext":""},{"type":"reviewersInvited","content":"","date":"2025-09-13T09:45:00+00:00","index":"","fulltext":""},{"type":"editorAssigned","content":"","date":"2025-09-13T07:30:00+00:00","index":"","fulltext":""},{"type":"checksComplete","content":"","date":"2025-09-12T23:44:57+00:00","index":"","fulltext":""},{"type":"submitted","content":"Journal of Orthopaedic Surgery and Research","date":"2025-09-10T14:16:16+00:00","index":"","fulltext":""}],"status":"published","journal":{"display":true,"email":"[email protected]","identity":"journal-of-orthopaedic-surgery-and-research","isNatureJournal":false,"hasQc":true,"allowDirectSubmit":false,"externalIdentity":"josr","sideBox":"Learn more about [Journal of Orthopaedic Surgery and Research](http://josr-online.biomedcentral.com)","snPcode":"13018","submissionUrl":"https://submission.nature.com/new-submission/13018/3","title":"Journal of Orthopaedic Surgery and Research","twitterHandle":"@MSKmedBMC","acdcEnabled":true,"dfaEnabled":true,"editorialSystem":"em","reportingPortfolio":"BMC/SO AJ","inReviewEnabled":true,"inReviewRevisionsEnabled":true}}],"origin":"","ownerIdentity":"faed072f-85c0-42c5-bef4-f2a0632c8ca6","owner":[],"postedDate":"September 22nd, 2025","published":true,"recentEditorialEvents":[],"rejectedJournal":[],"revision":"","amendment":"","status":"published-in-journal","subjectAreas":[],"tags":[],"updatedAt":"2025-11-24T16:08:22+00:00","versionOfRecord":{"articleIdentity":"rs-7583841","link":"https://doi.org/10.1186/s13018-025-06523-3","journal":{"identity":"journal-of-orthopaedic-surgery-and-research","isVorOnly":false,"title":"Journal of Orthopaedic Surgery and Research"},"publishedOn":"2025-11-23 15:59:16","publishedOnDateReadable":"November 23rd, 2025"},"versionCreatedAt":"2025-09-22 08:54:28","video":"","vorDoi":"10.1186/s13018-025-06523-3","vorDoiUrl":"https://doi.org/10.1186/s13018-025-06523-3","workflowStages":[]},"version":"v1","identity":"rs-7583841","journalConfig":"researchsquare"},"__N_SSP":true},"page":"/article/[identity]/[[...version]]","query":{"redirect":"/article/rs-7583841","identity":"rs-7583841","version":["v1"]},"buildId":"8U1c8b4HqxoKbykW_rLl7","isFallback":false,"isExperimentalCompile":false,"dynamicIds":[84888],"gssp":true,"scriptLoader":[]}