BMP-2-driven osteo-organoid formation retains key osteogenic-support features and promotes bone repair following total-body irradiation

Fuwei Zhu; Luli Ji; Jing Wang; Changsheng Liu

doi:10.1016/j.bioactmat.2026.08.042

. 2026 Aug 27;67:774–793. doi: 10.1016/j.bioactmat.2026.08.042

BMP-2-driven osteo-organoid formation retains key osteogenic-support features and promotes bone repair following total-body irradiation

Fuwei Zhu ^a,^c, Luli Ji ^a,^c, Jing Wang ^a,^b,^c,^⁎, Changsheng Liu ^b,^c,^d,^⁎⁎

PMCID: PMC13544112 PMID: 42699681

Abstract

Irradiation stress causes persistent skeletal injury by disrupting marrow homeostasis, stromal function, and bone repair. However, the temporal remodeling of the native bone microenvironment and the capacity of a biomaterial-induced osteogenic environment to sustain regeneration under systemic irradiation remain unclear. Using 6 Gy total body irradiation (TBI), we characterized time-resolved changes in the native femur and evaluated BMP-2-loaded hyaluronic acid methacryloyl (BMP-2/HAMA)-induced osteo-organoids relative to their corresponding non-irradiated controls. TBI caused progressive femoral deterioration, including trabecular bone loss, persistent B-cell depletion, neutrophil-associated myeloid expansion, Type-H endothelial-cell decline, and accumulation of senescence-associated mesenchymal stromal cells (MSCs). Single-cell RNA sequencing further resolved mature B-cell loss and enrichment of inflammatory and mature neutrophil states. In osteo-organoids, irradiation delayed but did not abolish tissue maturation. By 6 weeks, irradiated osteo-organoids approached the tissue and cellular features of non-irradiated osteo-organoids at the mature 3-week stage, while the MSC-associated stromal compartment remained relatively stable. Osteo-organoid-derived MSCs exhibited less pronounced irradiation-associated dysfunction and retained clonogenic, osteogenic, and immunomodulatory capacities. In an irradiation-impaired femoral defect model, BMP-2/HAMA enhanced mineralized tissue formation, mechanical competence, and gait function. These findings support BMP-2/HAMA as a biomaterial-assisted strategy for functional bone regeneration under irradiation-impaired conditions.

Keywords: Irradiation stress, BMP-2, Osteogenic-support microenvironment, Osteo-organoid, Bone repair

Graphical abstract

Highlights

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TBI progressively disrupts the native femoral osteogenic-support microenvironment.
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Single-cell profiling reveals B-cell loss and neutrophil-state remodeling.
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Irradiation delays but does not abolish osteo-organoid maturation.
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Osteo-organoid MSCs retain clonogenic, osteogenic, and immunomodulatory functions.
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BMP-2/HAMA improves structural, mechanical, and gait outcomes in irradiated defects.

1. Introduction

Ionizing radiation can cause persistent skeletal injury, including deterioration of bone quality, disruption of marrow homeostasis, and impaired bone repair [[1], [2], [3]]. Although clinical irradiation varies substantially in exposure field, dose, and fractionation, experimental total body irradiation (TBI) provides a standardized model for investigating how systemic irradiation stress reshapes tissue microenvironments and regenerative capacity [[4], [5], [6]]. Irradiation not only directly affects osteoblast-lineage and osteoclast-lineage cells but also damages the vascular network, alters immune-cell composition and inflammatory signaling, and compromises marrow stromal and progenitor populations [[7], [8], [9], [10], [11], [12], [13], [14]]. Because bone formation and repair depend on coordinated interactions among these compartments [[15], [16], [17]], skeletal injury after irradiation cannot be fully understood from changes in bone-forming cells alone.

Here, we use the term osteogenic-support microenvironment to describe the local immune, vascular, and MSC-related conditions that collectively support bone formation, remodeling, and repair. Accordingly, we focused on three interconnected dimensions: immune-cell balance, bone-associated vascular support, and MSC abundance, senescence-associated status, and functional competence.

Despite increasing recognition that bone regeneration depends on coordinated immune, vascular, and stromal regulation, the temporal response of this multicompartment microenvironment to irradiation remains insufficiently understood. It is unclear whether its immune, vascular, and stromal modules undergo an acute disruption followed by recovery or whether they progressively deteriorate and impose sustained constraints on skeletal regeneration. Previous studies have frequently focused on individual cell populations, isolated molecular readouts, or single observation points, limiting an integrated understanding of how immune, vascular, and stromal remodeling evolves after irradiation and relates to impaired bone regeneration [[18], [19], [20]]. Defining these temporal changes is therefore important for understanding irradiation-associated disruption of the local bone microenvironment.

A related but distinct question is whether a biomaterial-defined local osteogenic environment can retain its tissue-forming and reparative capacities when the host is exposed to systemic irradiation stress. This question can be addressed using our previously established BMP-2/HAMA platform. Earlier studies demonstrated that implantation of cell-free BMP-2/HAMA scaffolds recruits host cells to form an in vivo osteo-organoid and that the BMP-2-driven tissue-forming process supports the acquisition of immunomodulatory function by osteo-organoid-derived MSCs [21,22]. Building on this foundation, we operationally define osteo-organoids in the present study as host-derived, ectopic bone-like tissues formed in vivo following subcutaneous implantation of cell-free BMP-2/HAMA scaffolds, distinguishing them from classical organoid systems that typically rely on the in vitro self-organization of stem or progenitor cells [23]. The porous HAMA matrix provides a local framework for host-cell recruitment, while BMP-2 initiates sequential fibrous and cartilaginous tissue formation, ossification, establishment of marrow-like regions, and later remodeling [22,24,25]. As an established cell-free platform with stage-associated tissue formation and MSC-related functions, this system enables us to determine whether biomaterial-induced bone-like tissue can sustain its maturation and cellular properties under systemic irradiation stress, with irradiated osteo-organoids evaluated against their corresponding stage-matched non-irradiated controls.

Here, we established a systemic irradiation-stress model using 6 Gy TBI and evaluated irradiation-induced changes in the native femur and BMP-2/HAMA-induced osteo-organoids in parallel, with each tissue assessed relative to its corresponding non-irradiated baseline. In the native femur, irradiation induced persistent B-cell loss, neutrophil expansion, depletion of Type-H endothelial cells, accumulation of senescence-associated MSCs, and progressive trabecular bone loss. Single-cell transcriptomic analysis further resolved mature B-cell depletion and enrichment of inflammatory and mature neutrophil states with transcriptional features related to oxidative stress, vascular regulation, and bone remodeling. In osteo-organoids, irradiation delayed but did not abolish tissue maturation, while osteo-organoid-derived MSCs retained comparatively greater clonogenic, differentiation, and immunomodulatory capacities than irradiated femur-derived BMSCs. BMP-2/HAMA further enhanced mineralized tissue formation, mechanical competence, and locomotor function in an irradiation-impaired femoral defect model. Together, these findings show that the native femoral microenvironment undergoes progressive disruption after irradiation, whereas biomaterial-induced bone-like tissue retains sustained maturation and repair-supportive cellular functions under the same systemic stress.

2. Results

2.1. TBI induces systemic injury and progressive femoral bone deterioration

To establish an irradiation-induced injury model, mice were subjected to 6 Gy total body irradiation (TBI), and systemic irradiation-associated changes were evaluated 4 weeks after exposure. Compared with age-matched controls, TBI-treated mice exhibited impaired hair regrowth and increased SA-β-gal staining in adipose tissue, indicating the development of senescence-associated tissue alterations (Fig. 1A–C). Peripheral blood analysis further revealed pronounced hematological abnormalities, including reductions in red blood cell and white blood cell counts, hematocrit, hemoglobin concentration, and lymphocyte proportion, accompanied by increased proportions of monocytes and neutrophils (Fig. 1D–J). Body weight declined during the early post-irradiation period and subsequently recovered. Survival monitoring over the 4-week observation period showed limited mortality, with 13 of 15 irradiated mice surviving to the predefined endpoint (Fig. 1K and L). Histological analyses of major organs revealed additional evidence of systemic tissue injury, including increased collagen deposition and p16 expression, together with altered HMGB1 staining in the liver, lung, and kidney (Supplementary Fig. 1).

Fig. 1 — Total body irradiation induces bone marrow inflammation and progressive femoral bone loss. (A) Representative gross images showing hair regrowth in mice subjected to total body irradiation (TBI) and non-irradiated controls (Ctrl). (B, C) Representative senescence-associated β-galactosidase (SA-β-gal) staining of adipose tissues and quantification of the relative SA-β-gal-positive staining intensity. Scale bar, 1 cm; n = 5. (D-J) Peripheral blood cell counts and hematological parameters, including red blood cell count (RBC; D), white blood cell count (WBC; E), hematocrit (HCT; F), hemoglobin concentration (HGB; G), lymphocyte percentage (LYMPH%; H), monocyte percentage (MONO%; I), and neutrophil percentage (NEUT%; J), in Ctrl and TBI mice; n = 6. (K) Changes in body weight after TBI, expressed as the percentage change relative to baseline; n = 6. (L) Survival curves of Ctrl and TBI mice during the 28-day observation period; 13 of 15 TBI mice survived. (M) Multiplex analysis of inflammatory and immune-related factors in femoral bone marrow supernatants from Ctrl and TBI mice, including IL-2, IL-3, IL-5, IL-6, IL-9, IL-10, IP-10, IL-12p70, IL-15, TNF-α, and IFN-γ; n = 6. (N) Representative hematoxylin and eosin (H&E)-stained sections of femurs from Ctrl mice and TBI mice at 1, 3, and 6 weeks after irradiation. Bt, bone trabeculae; Ca, cartilage; Ad, adipocytes. Scale bar, 100 μm. (O) Quantification of the adipocyte area within the femoral bone marrow; n = 6. (P) Representative two-dimensional and three-dimensional micro-computed tomography (micro-CT) images of distal femurs from Ctrl mice and TBI mice at 1, 3, and 6 weeks after irradiation. Scale bar, 1 mm. (Q-U) Micro-CT-based quantitative analyses of bone volume fraction (BV/TV; Q), trabecular thickness (Tb.Th; R), trabecular number (Tb.N; S), trabecular separation (Tb.Sp; T), and bone mineral density (BMD; U); n = 6. Data are presented as the mean ± SD. Statistical significance was determined using an unpaired two-tailed Student's t-test for comparisons between two groups and one-way ANOVA followed by Tukey's multiple-comparisons test for comparisons among multiple groups. For longitudinal body-weight measurements, two-way ANOVA followed by Sidak's multiple-comparisons test was used. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.

Given the high sensitivity of bone marrow and skeletal tissues to irradiation [26,27], we next assessed the inflammatory status of the femoral marrow microenvironment. Multiplex cytokine analysis revealed increased levels of several pro-inflammatory and immune-activating mediators, including IL-3, IL-6, IP-10, IL-12p70, IL-15, and IFN-γ, accompanied by a reduction in the anti-inflammatory cytokine IL-10 (Fig. 1M). These changes indicate that TBI shifts the femoral marrow toward a pro-inflammatory state characterized by enhanced inflammatory signaling and impaired anti-inflammatory regulation.

We then performed a longitudinal analysis at 1, 3, and 6 weeks after TBI to characterize the temporal progression of irradiation-induced femoral remodeling. These time points were selected to characterize progressive changes in the native femur and to provide a temporal framework aligned with the major stages of BMP-2/HAMA-induced osteo-organoid maturation. H&E staining revealed a time-dependent increase in adipocyte-like vacuoles within the marrow cavity, indicating progressive marrow adiposity (Fig. 1N and O). Consistently, SA-β-gal staining showed gradual accumulation of senescence-associated signals in femoral tissues from 1 to 6 weeks after TBI (Supplementary Fig. 2A and B).

OCN-positive areas progressively decreased over the same period, whereas TRAP-positive areas increased, indicating reduced osteogenic activity accompanied by enhanced osteoclast-associated remodeling (Supplementary Fig. 2C-E). In agreement with these histological findings, micro-computed tomography showed progressive deterioration of distal femoral trabecular architecture after irradiation (Fig. 1P). Quantitative analysis revealed reductions in bone volume fraction (BV/TV), trabecular thickness (Tb.Th), trabecular number (Tb.N), and bone mineral density (BMD), together with increased trabecular separation (Tb.Sp), with the greatest structural impairment observed at 6 weeks (Fig. 1Q–U). Collectively, these findings demonstrate that 6 Gy TBI induces systemic injury and marrow inflammation and drives progressive deterioration of the native femoral marrow and trabecular bone microenvironment.

2.2. TBI disrupts immune, vascular, and MSC-associated components of the femoral osteogenic-support microenvironment

Given the progressive loss of trabecular bone and remodeling of the marrow compartment after TBI, we next examined three predefined cellular modules of the femoral osteogenic-support microenvironment: immune homeostasis, vascular-osteogenic coupling, and MSC-associated stromal competence. Flow cytometry showed a progressive increase in myeloid cells and a corresponding decrease in lymphoid cells, resulting in a sustained elevation of the myeloid-to-lymphoid ratio (Fig. 2A–D). This immune shift was characterized by continuous B-cell loss and progressive Ly6G⁺ neutrophil expansion. Total T cells and CD8⁺ T cells reached their lowest levels at 3 weeks and then partially recovered, whereas NK cells and macrophages initially decreased at 1 week. NK cells remained below control levels after recovery, while macrophages subsequently increased and exceeded control levels by 6 weeks. CD4⁺ T cells and monocytes showed transient increases followed by later declines (Fig. 2E–M). CD19 and CD11b immunohistochemistry further supported the loss of B-cell-associated signals and enrichment of myeloid-cell-associated signals in irradiated femora (Supplementary Fig. 3A).

Fig. 2 — Total body irradiation induces progressive remodeling of the immune, vascular, and mesenchymal stromal cell compartments in the femoral bone marrow. (A) Representative flow cytometry plots showing the gating strategy and temporal changes in bone marrow immune-cell populations in femurs from non-irradiated control mice (Ctrl) and mice at 1, 3, and 6 weeks after total body irradiation (TBI-1W, TBI-3W, and TBI-6W, respectively). CD45⁺ cells were separated into myeloid and lymphoid populations. Lymphoid cells were further analyzed for B cells, total T cells, CD4⁺ T cells, CD8⁺ T cells, and natural killer (NK) cells, whereas myeloid cells were analyzed for monocytes, neutrophils, and macrophages. (B-M) Quantification of myeloid cells (B), lymphoid cells (C), the myeloid-to-lymphoid cell ratio (D), B cells (E), total T cells (F), CD4⁺ T cells (G), CD8⁺ T cells (H), the CD4⁺/CD8⁺ T-cell ratio (I), NK cells (J), neutrophils (K), monocytes (L), and macrophages (M). (N) Representative flow cytometry plots showing Type H endothelial cells, total endothelial cells, and arterial endothelial cells in the femoral bone marrow. Endothelial populations were analyzed within viable CD45^-Ter119^- cells. (O-Q) Quantification of Type H endothelial cells (O), total endothelial cells (P), and arterial endothelial cells (Q). (R) Representative flow cytometry plots showing bone marrow mesenchymal stromal cells (MSCs) within viable CD45⁻CD31^-Ter119^- cells and p16-positive cells within the MSC population. (S, T) Quantification of MSCs (S) and p16⁺ MSCs (T). Data are presented as the mean ± SD; n = 6 biologically independent mice per group. Statistical significance was determined by one-way ANOVA followed by Tukey's multiple-comparisons test. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.

We next examined the vascular compartment. Type-H endothelial cells are a specialized bone-associated vascular subtype linked to osteoprogenitor localization and angiogenesis-osteogenesis coupling [28,29]. TBI reduced total endothelial cells, arterial endothelial cells, and Type-H endothelial cells, with the latter progressively declining over time (Fig. 2N–Q). Consistently, CD31/EMCN immunofluorescence and VEGF ELISA indicated impaired vascular and angiogenic support after irradiation (Supplementary Fig. 3B and C).

Within the stromal compartment, the relative proportion of CD105⁺CD140a⁺ MSCs increased, while p16⁺ MSCs progressively accumulated after TBI (Fig. 2R–T). MSC absolute numbers decreased at 1 and 3 weeks but recovered by 6 weeks, indicating that numerical recovery was accompanied by increased senescence-associated remodeling (Supplementary Fig. 4).

Although total femoral cellularity was lowest at 1 week and subsequently recovered, absolute B-cell numbers continued to decrease, neutrophil numbers increased, and Type-H endothelial-cell numbers progressively declined (Supplementary Fig. 4). Collectively, these findings demonstrate that TBI disrupts the femoral osteogenic-support microenvironment through persistent B-cell depletion, neutrophil-dominant myeloid remodeling, loss of Type-H vascular support, and senescence-associated alteration of the MSC compartment.

2.3. Single-cell profiling reveals B-cell contraction and neutrophil-associated myeloid remodeling after TBI

Although flow cytometry identified sustained B-cell loss and neutrophil-associated myeloid expansion as dominant features of irradiation-induced immune remodeling, it could not resolve immune-cell heterogeneity or cell-state-specific transcriptional changes. We therefore performed longitudinal single-cell RNA sequencing (scRNA-seq) of femoral samples collected from the Femur-Ctrl, Femur-TBI-1W, Femur-TBI-3W, and Femur-TBI-6W groups. Condition-resolved t-SNE visualization was used to display the distribution of cells from each experimental group within the shared integrated embedding space (Supplementary Fig. 5A).

Integrated single-cell analysis identified major skeletal, stromal, vascular, hematopoietic, and immune cell populations in the femoral microenvironment, including B cells, T cells, NK cells, neutrophils, monocytes, macrophages, endothelial cells, mesenchymal stem/stromal cells, osteoblasts, osteoclasts, chondrocytes, and hematopoietic progenitor cells (Fig. 3A, B and Supplementary Fig. 5B). Cell-composition and differential abundance analyses identified B-cell depletion and neutrophil expansion as the most prominent cellular changes after TBI (Fig. 3C–E and Supplementary Fig. 5C and D). B-cell subclustering revealed a reduction in mature B cells after irradiation, providing a higher-resolution explanation for the persistent B-cell loss observed at the major-cell-type level (Supplementary Fig. 5E-H).

Fig. 3 — Single-cell transcriptomic profiling reveals B-cell contraction and transcriptional remodeling of neutrophil states after TBI. (A) Integrated t-SNE visualization of major skeletal, stromal, vascular, hematopoietic, and immune cell populations identified in femoral samples. (B) t-SNE plots showing the distribution of major cell populations in the Ctrl, TBI-1W, TBI-3W, and TBI-6W groups. (C) Relative proportions of major cell populations across different experimental groups. (D) Radar plot showing the relative abundance changes of major bone marrow cell populations after TBI. (E) Differential abundance analysis of cell populations between TBI and Ctrl groups, with dot size representing the absolute differential ratio and the vertical position indicating the direction and magnitude of change. (F) t-SNE visualization of neutrophil subpopulations identified from femoral bone marrow single-cell transcriptomic data in Ctrl and TBI groups. (G) Dot plot showing the expression patterns of representative marker genes associated with neutrophil subclusters, including antimicrobial, cycling, high-mitochondrial, immature, inflammatory, mature, and stress-associated neutrophil populations. Dot size indicates the percentage of cells expressing each gene, and color intensity represents the average expression level. (H) Differential abundance analysis of neutrophil subpopulations between Femur-TBI and Femur-Ctrl groups. (I, J) KEGG pathway enrichment analysis of genes upregulated in the Femur-TBI group relative to the Femur-Ctrl group in mature neutrophils and inflammatory neutrophils, respectively. Bar length represents the number of genes mapped to each pathway, and color indicates the adjusted P value. (K, L) Gene Ontology biological process enrichment analysis of differentially expressed genes in mature neutrophils and inflammatory neutrophils, respectively. Positive and negative signed -log10(q value) values indicate enrichment among genes upregulated and downregulated in Femur-TBI relative to Femur-Ctrl, respectively; the values shown beside the bars indicate the corresponding q values.

Given the pro-inflammatory milieu in the irradiated femoral marrow and the marked expansion of neutrophil-associated cells, we next examined the heterogeneity of this myeloid compartment. Subclustering identified antimicrobial, cycling, high-mitochondrial, immature, inflammatory, mature, and stress-associated neutrophil states (Fig. 3F and G). Among these subclusters, inflammatory neutrophils and mature neutrophils were enriched after TBI (Fig. 3H), indicating that the increase in neutrophils reflected selective expansion or accumulation of specific cellular states rather than a uniform increase across all neutrophil subsets.

We subsequently examined the transcriptional programs associated with the expanded mature and inflammatory neutrophil states. KEGG analysis of genes upregulated in mature neutrophils from Femur-TBI relative to Femur-Ctrl highlighted top-ranked pathways associated with inflammatory signaling and cell-matrix interactions, including TNF signaling, integrin signaling, ECM-receptor interaction, and focal adhesion (Fig. 3I). In inflammatory neutrophils, KEGG analysis of the upregulated genes highlighted osteoclast differentiation-related and inflammatory pathways (Fig. 3J).

GO biological process analysis provided further insight into these transcriptional changes. In mature neutrophils, genes upregulated in Femur-TBI were enriched in processes related to the negative regulation of angiogenesis, blood vessel morphogenesis, and vascular development, together with cartilage development, collagen metabolism, chondrocyte differentiation, and endochondral bone morphogenesis (Fig. 3K). Downregulated genes were associated primarily with lymphocyte proliferation and adaptive immune responses. In inflammatory neutrophils, upregulated genes were enriched in reactive oxygen species metabolic processes, whereas downregulated genes were associated with B-cell receptor-related signaling annotations (Fig. 3L). Together, these enrichment patterns suggest that irradiation-associated mature and inflammatory neutrophil states exhibit transcriptional features related to inflammatory signaling, oxidative stress, vascular regulation, extracellular-matrix interactions, and bone remodeling within the femoral osteogenic-support microenvironment.

Ligand-receptor-based analysis further showed extensive predicted signaling among immune, stromal, vascular, and skeletal populations (Supplementary Fig. 5I). Source-centered networks suggested potential outgoing communication from inflammatory and mature neutrophils to MSCs, endothelial cells, osteoblasts, osteoclasts, and multiple immune populations (Supplementary Fig. 5J and K). These computational analyses suggest potential neutrophil-associated intercellular signaling within the irradiated femoral osteogenic-support microenvironment, although the underlying molecular pathways and functional consequences require further validation. Together, the single-cell data provide a cellular and transcriptional basis for the B-cell contraction and neutrophil-dominant myeloid remodeling observed after TBI.

2.4. BMP-2/HAMA-induced osteo-organoids show delayed but sustained maturation under irradiation stress

We next examined how systemic irradiation affected the stage-associated formation and maturation of BMP-2/HAMA-induced osteo-organoids. Osteo-organoids were generated by subcutaneous implantation of porous HAMA sponge scaffolds loaded with 30 μg BMP-2 into the dorsal region of mice [21]. We first characterized their formation and maturation under non-irradiated conditions. The constructs progressed from a fibrous proliferative phase containing cartilaginous regions at 1 week to active ossification at 2 weeks, followed by the formation of mature bone-like tissue with abundant marrow-like regions at 3 weeks and subsequent remodeling accompanied by adipose-like tissue accumulation at 6 weeks (Fig. 4A). Scaffold characterization showed that BMP-2-loaded HAMA retained an interconnected porous structure and comparable porosity and degradation behavior to PBS-loaded HAMA, while providing sustained BMP-2 release during the first week (Supplementary Fig. 6). To determine whether HAMA alone could initiate a comparable tissue-forming process, PBS-loaded HAMA scaffolds were implanted and evaluated at 1, 3, and 6 weeks under TBI. Macroscopic and micro-CT analyses showed no detectable mineralized or bone-like structure formation at any of these time points (Supplementary Fig. 7A and B), indicating that HAMA alone was insufficient to reproduce the osteo-organoid-forming trajectory observed with BMP-2/HAMA. Early BMP-Smad inhibition with LDN-193189 markedly impaired 3-week osteo-organoid formation, further supporting the requirement for early BMP signaling in this system (Supplementary Fig. 7C-I).

We then evaluated osteo-organoid maturation under TBI (Fig. 4B). Implantation of BMP-2/HAMA scaffolds did not produce detectable differences in the measured peripheral blood indices or femoral marrow cytokine profiles of TBI-exposed mice, indicating that no additional implantation-associated effect was detected for these measured systemic and femoral marrow endpoints (Supplementary Fig. 8). Macroscopic observation showed that non-irradiated osteo-organoids became highly vascularized and bone-like at 3 weeks and then entered a later post-maturation remodeling stage by 6 weeks. In contrast, TBI-exposed osteo-organoids showed delayed maturation at 3 weeks, whereas by 6 weeks they acquired more mature bone-like gross features (Supplementary Fig. 9).

Micro-CT analysis further confirmed irradiation-associated delayed maturation of the osteo-organoids. At 3 weeks, TBI markedly reduced mineralized tissue formation, as indicated by decreased BV/TV, Tb.Th, connectivity density, and bone mineral content compared with non-irradiated controls (Fig. 4C and D). By 6 weeks, however, TBI-exposed osteo-organoids showed increased mineralized structure compared with their 3-week irradiated counterparts and progressed toward the non-irradiated 3-week maturation state (Fig. 4C and D). Direct comparison between non-irradiated 3-week osteo-organoids and TBI-exposed 6-week osteo-organoids showed no significant differences in major micro-CT parameters, further supporting delayed rather than abolished mineralized tissue maturation after irradiation (Supplementary Fig. 10A-D).

Histological and immunohistochemical analyses yielded consistent findings. At 3 weeks, irradiated osteo-organoids had lower wet weights and reduced OCN- and TRAP-positive areas compared with non-irradiated osteo-organoids, indicating delayed acquisition of mature bone-like tissue features (Fig. 4E, F, H, I). SA-β-gal staining increased modestly during osteo-organoid maturation in both non-irradiated and irradiated groups. This pattern was consistent with stage-associated tissue remodeling and differed from the progressive accumulation of senescence-associated signals observed in the native femur (Fig. 4E–G). At 6 weeks, TBI-exposed osteo-organoids displayed OCN- and TRAP-positive tissue features comparable to those observed in non-irradiated 3-week osteo-organoids (Fig. 4J and K). Together, these findings demonstrate that systemic irradiation delays but does not abolish BMP-2/HAMA-induced osteo-organoid maturation.

2.5. TBI delays coordinated immune and vascular remodeling during osteo-organoid maturation

We next examined how the immune, vascular, and MSC-associated stromal modules of the osteogenic-support microenvironment changed during stage-associated osteo-organoid maturation. Under non-irradiated conditions, osteo-organoids showed a progressive decrease in myeloid-cell frequency and a progressive increase in lymphoid-cell frequency, resulting in a declining myeloid-to-lymphoid ratio over time (Fig. 5A–D). This remodeling pattern was accompanied by B-cell enrichment, increased CD8⁺ T-cell frequency, reduced NK-cell frequency, relatively stable neutrophil frequency, and gradual decreases in monocyte and macrophage populations (Fig. 5E–M), indicating a coordinated stage-associated immune remodeling pattern during osteo-organoid maturation.

Fig. 5 — Total body irradiation transiently disrupts and delays immune and vascular maturation within osteo-organoids. (A) Representative flow cytometry plots showing the gating strategy and temporal changes in immune-cell populations within osteo-organoids from non-irradiated control mice and TBI-treated mice at 1, 3, and 6 weeks. CD45⁺ cells were separated into myeloid and lymphoid populations. Lymphoid cells were further analyzed for B cells, total T cells, CD4⁺ T cells, CD8⁺ T cells, and natural killer (NK) cells, whereas myeloid cells were analyzed for neutrophils, monocytes, and macrophages. (B-M) Quantification of myeloid cells (B), lymphoid cells (C), the myeloid-to-lymphoid cell ratio (D), B cells (E), total T cells (F), CD4⁺ T cells (G), CD8⁺ T cells (H), the CD4⁺/CD8⁺ T-cell ratio (I), NK cells (J), neutrophils (K), monocytes (L), and macrophages (M). (N) Representative flow cytometry plots showing Type H endothelial cells, total endothelial cells, and arterial endothelial cells within osteo-organoids from Ctrl and TBI groups at 1, 3, and 6 weeks. Endothelial-cell populations were analyzed within viable CD45^-Ter119^- cells. (O-Q) Quantification of Type H endothelial cells (O), total endothelial cells (P), and arterial endothelial cells (Q). (R) Representative flow cytometry plots showing mesenchymal stromal cells (MSCs) within viable CD45⁻CD31^-Ter119^- cells and p16-positive cells within the MSC population. (S, T) Quantification of MSCs (S) and p16⁺ MSCs (T). Data are presented as the mean ± SD; n = 6 biologically independent samples per group. Statistical significance was determined by two-way ANOVA followed by Sidak's multiple-comparisons test. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.

TBI delayed this stage-associated immune transition. At 3 weeks, irradiated osteo-organoids exhibited a myeloid-skewed immune profile, characterized by reduced frequencies of lymphoid cells, B cells, and CD8⁺ T cells, together with increased frequencies of myeloid cells, neutrophils, monocytes, and macrophages relative to time-matched non-irradiated controls (Fig. 5B–M). CD11b and CD19 immunohistochemistry further supported the altered distribution of myeloid- and B-cell-associated signals within irradiated osteo-organoids (Supplementary Fig. 11A). By 6 weeks, the immune composition of irradiated osteo-organoids shifted toward that of non-irradiated osteo-organoids at the mature 3-week stage, although residual myeloid skewing remained (Supplementary Fig. 10E-P).

Vascular remodeling showed a similar delay. Under non-irradiated conditions, Type-H endothelial cells gradually decreased over time, consistent with the transition from active bone formation to later-stage remodeling. At 3 weeks, TBI reduced total endothelial cells, arterial endothelial cells, and Type-H endothelial cells relative to time-matched non-irradiated osteo-organoids (Fig. 5N–Q). CD31/EMCN immunofluorescence and VEGF ELISA further indicated impaired vascular and angiogenic support at this stage (Supplementary Fig. 11B and C). By 6 weeks, however, total, arterial, and Type-H endothelial-cell populations approached those observed in the non-irradiated 3-week mature reference state, with no significant differences detected between these groups (Supplementary Fig. 10Q-S).

In contrast, the MSC-associated stromal compartment remained comparatively stable. MSC frequency and the proportion of p16⁺ MSCs were similar between irradiated and time-matched non-irradiated osteo-organoids at all examined stages (Fig. 5R–T). To determine whether this comparatively preserved MSC-associated phenotype could be attributed to the HAMA scaffold itself, we additionally analyzed PBS-loaded HAMA implants at 3 weeks under TBI. HAMA/PBS implants contained only a small MSC population, and the recovered MSCs exhibited a higher proportion of p16⁺ cells than those in BMP-2/HAMA-induced osteo-organoids (Supplementary Fig. 12A-C), indicating that HAMA implantation alone did not reproduce the MSC-associated phenotype observed in BMP-2/HAMA-induced osteo-organoids. The relative preservation of the MSC-associated compartment was also evident when non-irradiated 3-week osteo-organoids were compared with irradiated 6-week osteo-organoids (Supplementary Fig. 10T and U). Absolute cell-number analysis further supported this delayed-convergence pattern. Total osteo-organoid cellularity was reduced at 3 weeks after TBI but increased by 6 weeks to a level comparable to that of the non-irradiated 3-week mature reference. At this stage, B-cell, Type-H endothelial-cell, and MSC numbers were also comparable between the two groups, despite residual neutrophil elevation (Supplementary Fig. 12D-L). Together, TBI delays stage-associated immune and vascular remodeling during osteo-organoid maturation, while the MSC-associated stromal compartment remains largely preserved. By 6 weeks, key cellular features of irradiated osteo-organoids approached those of non-irradiated osteo-organoids at the mature 3-week stage.

2.6. Osteo-organoid-derived MSCs retain functional and immunomodulatory properties after irradiation

Although the MSC-associated compartment remained relatively stable in irradiated osteo-organoids, flow-cytometric analysis alone could not determine whether these cells retained their functional properties. To minimize the influence of prolonged secondary inflammation and tissue remodeling, MSCs were isolated from native femora and osteo-organoids at 1 week after TBI for comparative functional analyses.

MSCs isolated from native femora and BMP-2/HAMA-induced osteo-organoids were designated BMSCs and HMSCs, respectively; those isolated from TBI-exposed mice were further designated TBI-BMSCs and TBI-HMSCs. TBI markedly impaired BMSCs, as shown by reduced cell-cycle progression and Ki67 positivity, increased SA-β-gal staining, enhanced γH2AX signals, elevated p21 protein expression, and upregulated senescence- and inflammation-associated genes (Fig. 6A–K). Consistent with these transcriptional changes, ELISA of MSC-conditioned media showed markedly increased secretion of IL-6 and CCL2 by TBI-BMSCs, whereas both factors remained at low levels in BMSCs and TBI-HMSCs (Supplementary Fig. 13A and B). Overall, TBI-HMSCs exhibited substantially weaker damage-, senescence-, and inflammation-associated responses than TBI-BMSCs (Fig. 6A–K and Supplementary Fig. 13A and B). Direct in vitro irradiation further confirmed that osteo-organoid-derived MSCs were less susceptible to irradiation-induced apoptosis than femur-derived BMSCs (Supplementary Fig. 13C-E).

Fig. 6 — TBI-HMSCs exhibit reduced cellular damage and senescence while retaining key MSC functions. (A, B) Representative flow cytometric profiles of cell-cycle distribution and quantification of the proportions of cells in the G1 and S/G2 phases in Ctrl-BMSCs, TBI-BMSCs, and TBI-HMSCs; n = 5. (C, D) Representative flow cytometric analysis of Ki67 expression and quantification of Ki67-positive MSCs; n = 6. (E, F) Representative SA-β-gal staining and quantification of the relative number of SA-β-gal-positive cells; n = 6. Scale bar, 100 μm. (G, H) Representative immunofluorescence staining of γH2AX, F-actin, and nuclei, and quantification of the number of γH2AX foci per nucleus; n = 6. Scale bar, 25 μm. (I, J) Representative Western blot and densitometric analysis of p21 protein expression; n = 3. GAPDH was used as the loading control. (K) Quantitative real-time PCR analysis of senescence- and senescence-associated secretory phenotype-related genes, including P16, P21, P53, Il6, Ccl2, and Mmp13; n = 5 biologically independent samples, with three technical replicates per sample. (L, M) Representative colony-forming unit-fibroblast (CFU-F) assay and quantification of colony numbers; n = 5. Scale bar, 1 mm. (N-P) Representative Alizarin Red S and Oil Red O staining following osteogenic and adipogenic induction, respectively, and quantification of the positively stained areas; n = 6. Scale bars, 200 μm. (Q, R) Quantitative real-time PCR analysis of osteogenic genes (Runx2 and Sp7) and adipogenic genes (Pparg and Lpl); n = 5 biologically independent samples, with three technical replicates per sample. (S) Representative flow cytometric plots showing the polarization status of macrophages cultured in DMEM alone or cocultured with Ctrl-BMSCs, TBI-BMSCs, or TBI-HMSCs. CD86 and CD206 were used as markers of M1-and M2-like macrophages, respectively. (T, U) Quantification of M1-like CD86⁺CD206^- macrophages (T) and M2-like CD206⁺ macrophages (U); n = 6. (V) Quantitative real-time PCR analysis of macrophage polarization-related genes, including Inos, Il1b, Arg1, and Il10, after coculture with the indicated MSC populations; n = 5 biologically independent samples, with three technical replicates per sample. Data are presented as the mean ± SD. Statistical significance was determined by one-way ANOVA followed by Tukey's multiple-comparisons test. Panel J was performed using Kruskal-Wallis test followed by Dunn's multiple-comparisons test. ns, not significant; *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001.

Functional analyses showed that irradiated BMSCs exhibited reduced CFU-F formation, impaired osteogenic differentiation, and increased adipogenic differentiation. By contrast, TBI-HMSCs retained greater clonogenic and osteogenic capacities and showed a less pronounced adipogenic shift (Fig. 6L–R). We next assessed the immunomodulatory properties of these cells using macrophage co-culture assays. Compared with TBI-BMSCs, TBI-HMSCs induced a macrophage phenotype characterized by a lower proportion of CD86⁺CD206^- cells and a higher proportion of CD206-expressing cells, accompanied by decreased pro-inflammatory and increased anti-inflammatory gene expression (Fig. 6S–V and Supplementary Fig. 14). At the tissue level, immunofluorescence staining for iNOS and Arg-1 showed sustained enrichment of inflammatory macrophage-associated signals in irradiated femora, whereas osteo-organoids maintained a more balanced distribution of inflammatory and reparative macrophage-associated signals after TBI (Supplementary Fig. 15A-F). As an additional tissue-level readout of immunomodulatory signaling, PGE2 levels in the native femur peaked at 1 week after TBI and subsequently declined. In osteo-organoids, PGE2 levels gradually decreased with tissue maturation and did not differ significantly between the non-irradiated and TBI groups at the corresponding time points (Supplementary Fig. 15G and H). Together, these findings indicate that osteo-organoid-derived MSCs are less susceptible to irradiation-associated dysfunction and retain clonogenic, differentiation, and immunomodulatory capacities after irradiation.

2.7. BMP-2/HAMA promotes bone repair under irradiation-impaired conditions

Having characterized the irradiation response of osteo-organoids and their derived MSCs, we next evaluated the regenerative activity of BMP-2/HAMA in an irradiation-impaired femoral defect model. Micro-CT and H&E staining showed that TBI markedly impaired femoral defect repair, as evidenced by limited mineralized tissue formation and predominant fibrous tissue filling within the defect region (Fig. 7A). HAMA alone provided little regenerative benefit under TBI, whereas BMP-2/HAMA markedly enhanced mineralized tissue formation and promoted defect bridging (Fig. 7A). Quantitative micro-CT analysis confirmed that BMP-2/HAMA significantly increased BV/TV, Tb.N, Tb.Th, and BMD compared with TBI and TBI + HAMA groups (Fig. 7B–E).

Fig. 7 — BMP-2/HAMA hydrogel restores bone repair and functional recovery impaired by total body irradiation. (A) Representative micro-CT images and H&E staining of femoral defects in Ctrl, TBI, TBI + HAMA, and TBI + BMP-2/HAMA groups, showing bone regeneration under normal or TBI-impaired repair conditions. The red dashed circles indicate the defect regions in micro-CT images, and the black dashed circles indicate newly formed bone or residual scaffold regions in H&E-stained sections. Nb, newly formed bone; Ft, fibrous tissue; Sca, scaffold. Scale bar, 1 mm for micro-CT images; 200 μm for H&E. (B-E) Quantitative micro-CT analysis of regenerated bone, including BV/TV, Tb.N, Tb.Th, and BMD. (F) Representative image of biomechanical testing of regenerated femurs using a dynamic mechanical testing system. (G) Representative load-displacement curves of femurs from Sham, Ctrl, TBI, TBI + HAMA, and TBI + BMP-2/HAMA groups. (H, I) Quantitative analysis of maximum load (H) and fracture energy (I) obtained from biomechanical testing. (J) Representative gait patterns obtained by CatWalk gait analysis in Sham, Ctrl, TBI, TBI + HAMA, and TBI + BMP-2/HAMA groups. Different paw placements are indicated according to the CatWalk analysis system. (K) Representative paw-print images of hind limbs under intact and defect conditions. (L, M) Quantitative analysis of maximum paw intensity (L) and print area (M). Data are presented as the mean ± SD; n = 6 biologically independent animals per group. Statistical significance was determined by one-way ANOVA followed by Tukey's multiple-comparisons test. ns, not significant; *P < 0.05, *P < 0.01, ***P < 0.001, and ****P < 0.0001.

Three-point bending analysis further showed that irradiated femoral defects exhibited poor mechanical competence, with lower maximum load and fracture energy than non-irradiated defect controls (Fig. 7F–I). HAMA alone provided little improvement, whereas BMP-2/HAMA significantly increased both parameters, indicating enhanced structural integration and load-bearing capacity of the repaired femur. CatWalk gait analysis was used to assess gait-related function. Representative step-sequence patterns revealed disrupted alternating paw placement in TBI-treated defect mice, whereas BMP-2/HAMA treatment improved coordinated stepping patterns (Fig. 7J). Paw-print visualization and quantification showed reduced paw-contact intensity and print area after TBI, both of which were improved by BMP-2/HAMA treatment (Fig. 7K–M). Together, these findings indicate that BMP-2/HAMA enhances structural bone regeneration and improves the mechanical and locomotor outcomes of femoral defect repair under irradiation-impaired conditions.

To further characterize the local tissue state associated with bone repair, we examined immune, vascular, and MSC-related features within the defect region. BMP-2/HAMA-treated defects exhibited increased local CD19⁺ staining and CD31⁺EMCN⁺ Type-H vascular structures, together with a reduced proportion of p16⁺CD105⁺ stromal cells, compared with the irradiated untreated and HAMA-treated groups (Supplementary Fig. 16). These findings indicate that the improved structural and functional repair observed after BMP-2/HAMA treatment was accompanied by a more favorable local immune, vascular, and MSC-associated tissue state.

3. Discussion

In this study, we evaluated the responses of the native femur and BMP-2/HAMA-induced osteo-organoids to systemic irradiation, with each tissue assessed relative to its corresponding non-irradiated baseline. TBI progressively disrupted the native femoral osteogenic-support microenvironment, as reflected by persistent immune imbalance, Type-H endothelial-cell loss, and MSC senescence, accompanied by marrow adiposity and trabecular deterioration. In osteo-organoids, irradiation caused a marked delay in stage-associated maturation rather than irreversible arrest. Mineralization continued, immune and vascular features subsequently approached the non-irradiated mature state, and MSC abundance and senescence-associated status remained comparatively preserved. Consistent with this sustained tissue-forming trajectory, osteo-organoid-derived MSCs retained clonogenic, osteogenic, and immunomodulatory capacities, while BMP-2/HAMA improved mineralized tissue formation, mechanical integrity, and gait-related outcomes in irradiation-impaired femoral defects. Together, these findings suggest that the skeletal consequences of irradiation are shaped not only by systemic stress but also by the local tissue state and the ongoing tissue-forming process within that environment.

Previous studies have shown that irradiation impairs bone repair by damaging osteogenic cells, disrupting marrow stromal populations, injuring the vasculature, and altering immune responses [[7], [8], [9], [10],12,13,18,27]. The importance of such multicellular organization is increasingly recognized in bone marrow organoid models, which aim to recapitulate three-dimensional interactions among hematopoietic, vascular, and stromal compartments [30]. Our longitudinal findings extend this view by showing that these changes occur as coordinated remodeling of the native femoral osteogenic-support microenvironment rather than as isolated abnormalities. Persistent mature B-cell loss and neutrophil-dominant myeloid remodeling were accompanied by progressive Type-H endothelial-cell depletion and accumulation of senescence-associated MSCs. Single-cell analysis further showed enrichment of inflammatory and mature neutrophil states. Their transcriptional profiles were associated with ROS metabolism, TNF and cell-matrix signaling, negative regulation of vascular formation, and osteoclast differentiation-related pathways. These signatures provide potential links between immune remodeling, vascular decline, and unbalanced bone turnover, although enrichment and ligand-receptor analyses remain hypothesis-generating and do not establish direct causality.

The irradiation response of osteo-organoids should be interpreted in the context of their distinct tissue state. The adult femur is a mature skeletal organ maintained through continuous homeostatic remodeling [31,32], whereas osteo-organoids are actively forming ectopic bone-like tissues involving host-cell recruitment, ossification, vascularization, marrow-like region formation, and later remodeling [33,34]. More broadly, emerging bone-organoid platforms encompass diverse engineering strategies that integrate cellular and biomaterial cues to generate bone-like tissues with increasing structural and functional complexity [35]. Therefore, the present study does not imply direct equivalence between the two tissues, but evaluates their irradiation-response trajectories relative to their own baselines. The marked mineralization and vascular impairment observed in osteo-organoids at 3 weeks showed that they remained susceptible to TBI. Nevertheless, their continued maturation and the approach of several structural and cellular features toward the non-irradiated 3-week mature reference by 6 weeks indicate delay rather than irreversible arrest. This finding is consistent with reports that biomaterial-induced juvenile ossicles can retain regenerative activity under adverse host conditions, including the ability of rhBMP-2/gelatin-induced ossicles to promote bone repair in aged mice [36], and extends this concept to systemic irradiation stress.

The continued maturation of osteo-organoids may be related to the local tissue-forming process established by BMP-2/HAMA. Our previous studies showed that cell-free BMP-2/HAMA scaffolds recruit host cells to generate in vivo osteo-organoids and that the BMP-2-driven tissue-forming process supports the acquisition of immunomodulatory properties by osteo-organoid-derived MSCs [21,22]. Hydrogel-based biomaterials can influence bone regeneration by modulating local cellular and signaling microenvironments [37]. HAMA provides a porous matrix for local cell recruitment and BMP-2 retention, while BMP-2 initiates osteogenic differentiation and coordinated tissue remodeling. HAMA/PBS controls failed to form detectable mineralized tissue under TBI and, at 3 weeks, showed lower MSC abundance with increased p16 positivity. Early BMP-Smad inhibition further impaired osteo-organoid formation, supporting a role for canonical BMP signaling during initiation. These findings support a role for BMP-2-driven osteoinduction in establishing the tissue-forming process, rather than a direct radioprotective effect of BMP-2. Its osteoinductive activity may remain sufficient to sustain tissue formation after an early irradiation-induced delay. The weaker damage- and senescence-associated responses of TBI-HMSCs, together with their comparatively preserved clonogenic, osteogenic, and macrophage-modulatory functions, were temporally associated with this sustained trajectory. However, the present data do not establish that MSC preservation is necessary or sufficient for the later maturation of irradiated osteo-organoids. Tissue PGE2 levels appeared to vary primarily with osteo-organoid maturation rather than irradiation status, and their cellular source remains unresolved. Importantly, in the orthotopic defect model, the limited effect of HAMA alone indicates that provision of a scaffold was insufficient to overcome irradiation-impaired repair. The improvements in mechanical integrity and limb use following BMP-2/HAMA implantation therefore provide functional evidence that BMP-2/HAMA promoted structurally integrated repair rather than mineral deposition alone. This finding places the present approach within a broader direction of regenerative biomaterial strategies aimed at coupling bone formation with structural and functional reconstruction, including emerging biomaterial-based organoid-like constructs for bone-defect repair [38,39]. More broadly, organoid-based and organoid-derived therapeutic strategies are being explored for skeletal regeneration [40], although their translational efficacy and safety require further validation.

Several limitations should be acknowledged. First, 6 Gy TBI provides a reproducible systemic irradiation-stress model but does not reproduce the localized and fractionated exposure patterns used clinically. Under localized or fractionated irradiation, the magnitude and temporal course of osteo-organoid maturation delay, as well as associated immune, vascular, and MSC-related changes, may differ. Validation in clinically relevant irradiation models is therefore required. Second, although coordinated immune, vascular, and MSC-related changes were identified, the individual causal contributions of these components remain unresolved. GO, KEGG, and ligand-receptor analyses are predictive, and early BMP-Smad inhibition establishes a requirement for osteo-organoid initiation but not the mechanisms governing its later response to irradiation. Longitudinal scRNA-seq was also not performed in osteo-organoids, limiting molecular resolution of irradiation-associated remodeling in this tissue. Third, the origins, recruitment kinetics, survival, and fate of host-derived stromal cells within osteo-organoids remain unresolved and will require lineage-tracing and fate-mapping studies. Therefore, the comparatively preserved MSC-associated phenotype under TBI cannot yet be attributed to intrinsic stromal radioresistance rather of a defined stromal population than differences in cell recruitment, retention, or survival. Finally, longer follow-up is needed to determine the stability and remodeling of BMP-2/HAMA-induced bone-like tissues and to evaluate potential risks associated with BMP-2 delivery, including off-target ossification.

4. Conclusions

When evaluated relative to their respective non-irradiated baselines, the native femur and BMP-2/HAMA-induced osteo-organoids exhibited distinct response trajectories to systemic irradiation stress. TBI progressively disrupted the native femoral osteogenic-support microenvironment through persistent immune imbalance, Type-H endothelial-cell decline, and MSC senescence, accompanied by progressive trabecular bone loss. By contrast, irradiation delayed but did not terminate osteo-organoid maturation; several immune and vascular features subsequently approached the non-irradiated mature reference, while MSC abundance and senescence-associated status remained comparatively preserved. Osteo-organoid-derived MSCs showed weaker irradiation-associated dysfunction and retained clonogenic, osteogenic, and immunomodulatory capacities. In an irradiation-impaired femoral defect model, BMP-2/HAMA enhanced mineralized tissue formation, mechanical integrity, and gait-related outcomes, accompanied by more favorable local immune, vascular, and stromal features. Together, these findings support BMP-2/HAMA as a biomaterial-assisted platform for sustaining bone-like tissue formation and promoting functional bone repair under irradiation-impaired conditions.

5. Materials and methods

5.1. Animals and ethics statement

Male C57BL/6 mice aged 6-8 weeks were purchased from Shanghai JieSiJie Laboratory Animal Co., Ltd. Mice were maintained under specific pathogen-free conditions with a 12 h light/12 h dark cycle and ad libitum access to food and water. All animal procedures were approved by the Institutional Animal Care and Use Committee of East China University of Science and Technology (Approval No. ECUST-2024-022) and were performed in accordance with relevant institutional and national guidelines for animal welfare. Animals were randomly assigned to experimental groups. For longitudinal analyses, mice were euthanized at the indicated terminal time points; therefore, samples collected at 1, 3, and 6 weeks represented independent animal cohorts rather than repeated measurements from the same animals.

5.2. Total body irradiation model

Mice received single-dose total body irradiation (TBI) at 6 Gy with a dose rate of 1.28 Gy/min. Irradiation was performed at the National Facility for Protein Science Shanghai using an X-ray small-animal irradiator (RADSOURCE, RS2000). Body weight and survival were monitored after irradiation. Peripheral blood, femurs, and BMP-2/HAMA osteo-organoids were collected at the indicated time points for downstream analyses. For longitudinal femoral and osteo-organoid studies, samples were collected at 1, 3, and 6 weeks after irradiation unless otherwise specified.

5.3. Synthesis and fabrication of HAMA scaffolds

Sodium hyaluronic acid (HA; Bloomage Biotechnology Co., Ltd.) was used to synthesize hyaluronic acid methacrylate (HAMA) as previously described [41]. Briefly, HA was dissolved in deionized water under ice cooling, and methacrylic anhydride was added dropwise while maintaining the pH at 8.5-9.5 for 4 h. The reaction mixture was stirred overnight, dialyzed against deionized water for 7 days, frozen at −80 °C, and lyophilized to obtain HAMA.

To fabricate porous scaffolds, HAMA was dissolved in ultrapure water containing 0.1 wt/vol% lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP; StemEasy Biotech) to a final concentration of 3 wt/vol%. The precursor solution was transferred into molds and photocrosslinked under UV irradiation for 30 s. The hydrogels were then freeze-dried to generate porous HAMA scaffolds with dimensions of 5 mm × 5 mm × 5 mm. The BMP-2 dose was selected based on our previously optimized BMP-2/HAMA osteo-organoid platform [21,22]. Each sterile scaffold was loaded with 30 μL of recombinant human BMP-2 solution (1.00 mg/mL; rhBMP-2; Shanghai Rebone Biomaterials Co., Ltd.), corresponding to 30 μg BMP-2 per scaffold. The solution was allowed to fully absorb into the scaffold, after which the BMP-2/HAMA scaffolds were lyophilized and stored at −20 °C until implantation.

5.4. Characterization of HAMA scaffolds and BMP-2 release

The porous morphology of freeze-dried HAMA scaffolds was observed by scanning electron microscopy. Scaffold porosity was measured using a liquid-displacement pycnometer method with n-hexane as the displacement liquid. Briefly, freeze-dried HAMA scaffolds were weighed to obtain the dry scaffold mass (W_S). A pycnometer was first filled with n-hexane to the calibration mark and weighed (W₁). The scaffold was then immersed in n-hexane in the pycnometer, and the system was maintained until no visible air bubbles remained. The pycnometer was refilled with n-hexane to the calibration mark and weighed again (W₂). The n-hexane-saturated scaffold was then removed, and the pycnometer containing the remaining n-hexane was weighed (W₃). Scaffold porosity was calculated as: Porosity (%) = [(W₂ − W₃ − W_S)/(W₁− W3)] × 100.

The degradation behavior of HAMA scaffolds was evaluated in hyaluronidase-containing simulated body fluid at 37 °C. Briefly, PBS-loaded HAMA scaffolds and BMP-2-loaded HAMA scaffolds were weighed to obtain the initial dry mass (W₀) and then immersed in simulated body fluid containing hyaluronidase (100 U/mL). At predetermined time points (0, 1, 3, 5, 7, 10, and 14 days), scaffolds were collected, gently rinsed with deionized water, frozen, lyophilized, and weighed to obtain the remaining dry mass (W_t). The remaining mass ratio was calculated as: Remaining mass (%) = (W_t/W₀) × 100.

For in vitro BMP-2 release analysis, BMP-2/HAMA scaffolds were incubated in PBS at 37 °C. At each time point, part of the supernatant was collected and replaced with fresh medium. BMP-2 concentration in the supernatant was measured using a Human BMP-2 ELISA kit (Boster Biological Technology, Wuhan, China; EK0311) according to the manufacturer's instructions. Cumulative BMP-2 release was calculated relative to the total BMP-2 loaded in each scaffold.

5.5. Subcutaneous implantation and osteo-organoid induction

For survival surgery, mice were anesthetized with 4% isoflurane for induction and maintained under 1.5-2.0% isoflurane delivered through a nose cone during surgery. The depth of anesthesia was monitored according to the respiratory pattern and absence of the pedal withdrawal reflex, and the isoflurane concentration was adjusted as necessary. Mice were placed on a warming pad to maintain body temperature. Meloxicam was administered subcutaneously at 5 mg/kg every 24 h for 72 h after surgery for postoperative analgesia. Dorsal hair was removed, and the surgical site was disinfected with iodophor. All procedures were performed under aseptic conditions using sterile drapes and sterilized surgical instruments. A small dorsal skin incision was made to create a subcutaneous pocket, and one BMP-2/HAMA scaffold was implanted into each mouse. The incision was closed using 4-0 sutures. Mice were maintained on a warming pad until recovery from anesthesia and were monitored daily for general condition and wound healing.

For the TBI groups, mice received 6 Gy total body irradiation on the day of implantation. BMP-2/HAMA constructs were harvested at 1, 3, or 6 weeks after implantation for downstream analyses, including gross examination, micro-CT, histology, immunostaining, flow cytometry and local mediator assays.

5.6. Early BMP-Smad inhibition

To examine the requirement for early canonical BMP-Smad signaling in BMP-2/HAMA-induced bone-like tissue formation, mice received LDN-193189 dihydrochloride (MedChemExpress, HY-12071), a BMP type I receptor inhibitor [42]. LDN-193189 powder was first dissolved in DMSO to prepare a stock solution and then diluted with sterile PBS to obtain the working solution immediately before injection. LDN-193189 was administered intraperitoneally at 3 mg/kg every 12 h from day 0 to day 5 after BMP-2/HAMA scaffold implantation. Vehicle-treated mice received the same volume of DMSO/PBS solution according to the same dosing schedule. Control mice received the corresponding vehicle. Osteo-organoids were harvested at 3 weeks after implantation for micro-CT.

5.7. Sample collection and processing

Mice were euthanized at the indicated time points by overdose of sodium pentobarbital, followed by cervical dislocation as a secondary method to ensure death before tissue collection. For systemic phenotyping, peripheral blood was collected before tissue harvest. Liver, lung, kidney, and adipose tissue samples were collected where indicated. Liver, lung, and kidney tissues were fixed in 4% paraformaldehyde, dehydrated, paraffin-embedded, and sectioned at 7 μm. Fresh adipose tissue was processed for SA-β-gal staining.

Femurs were harvested at 1, 3, or 6 weeks after TBI. Samples designated for micro-CT were fixed in 4% paraformaldehyde and stored in PBS before scanning. For paraffin histology, femurs were decalcified in 10% EDTA (pH 7.4) at 4 °C with solution changes every 3 days, followed by paraffin embedding and sectioning. For cryosection-based assays, samples were processed as described below.

Osteo-organoids were retrieved at 1, 3, or 6 weeks after implantation. Gross morphology and wet weight were recorded. Samples for micro-CT were fixed in 4% paraformaldehyde. Samples for histology were decalcified in 10% EDTA at 4 °C with solution changes every 3 days and embedded in paraffin. Fresh samples were processed for flow cytometry, single-cell RNA sequencing, or biochemical assays as described below.

5.8. Peripheral blood analysis and bone marrow multiplex cytokine assay

Peripheral blood was collected from mice at the indicated time points after TBI. Complete blood count analysis was performed using an automated hematology analyzer (Mindray, China) according to the manufacturer's instructions. Parameters including white blood cell count, red blood cell count, hemoglobin, platelets, lymphocytes, neutrophils, and monocytes were recorded.

For bone marrow cytokine analysis, femoral bone marrow was flushed from each femur with 500 μL of PBS. The bone marrow suspension was centrifuged at 3000 g for 10 min at 4 °C, and the supernatant was collected for cytokine measurement. Cytokine levels were quantified using an ABPlex Mouse Cytokine Assay Kit (ABclonal, China) according to the manufacturer's instructions.

5.9. ELISA quantification of tissue and MSC-secreted factors

VEGF and prostaglandin E2 (PGE2) were quantified in femoral and osteo-organoid tissue homogenates. Briefly, tissues were weighed and homogenized in ice-cold PBS containing protease inhibitors. Homogenates were centrifuged at 12,000 rpm for 10 min at 4 °C, and the supernatants were collected. VEGF and PGE2 levels were measured using ELISA kits (VEGF: Boster Biological Technology, Cat. No. EK0541; PGE2: ELK Biotechnology, Cat. No. ELK10928) according to the manufacturers’ instructions. Mediator levels were normalized to tissue wet weight and expressed as pg/mg tissue.

For analysis of MSC-secreted factors, BMSCs and HMSCs were seeded in 6-well plates at 3 × 10⁵ cells per well and cultured in 1 mL of low-serum culture medium containing 2% FBS. After 48 h, conditioned media were collected and centrifuged at 3000 × g for 15 min to remove suspended cells and cellular debris. IL-6 and CCL2 concentrations in the resulting supernatants were measured using ELISA kits (IL-6: Boster Biological Technology, Cat. No. EK0411; CCL2: Boster Biological Technology, Cat. No. EK0568) according to the manufacturer's instructions. All MSC groups were analyzed using the same initial cell number, culture volume, serum concentration, and conditioned-medium collection period.

5.10. Histological staining

For hematoxylin and eosin staining, paraffin sections were deparaffinized, rehydrated, and subjected to standard H&E staining. Safranin O/Fast Green staining was performed using a bone tissue Safranin O/Fast Green staining kit (Servicebio, G1053, Wuhan, China) according to the manufacturer's instructions. Sirius Red staining was performed to evaluate collagen deposition using a commercial kit (Servicebio, G1078) according to the manufacturer's instructions. TRAP staining was performed on paraffin sections of femurs and osteo-organoids using a commercial kit (Servicebio, G1050) according to the manufacturer's instructions. Nuclei were counterstained with Mayer's hematoxylin. Images were acquired using a Leica DMi8 microscope. Quantification was performed in ImageJ using predefined regions of interest and identical thresholding criteria across groups.

5.11. SA-β-gal staining and quantification

Senescence-associated β-galactosidase staining was performed using a commercial kit (Beyotime, C0602) according to the manufacturer's protocol. For bone tissues, fresh samples were fixed in 4% paraformaldehyde for 4 h at 4 °C, decalcified in EDTA for 7 days, cryoprotected in 20% sucrose and 2% polyvinylpyrrolidone overnight, embedded in OCT, and sectioned at 10 μm using a freezing microtome (RWD Life Science, FS800). Sections were incubated with freshly prepared SA-β-gal staining solution at 37 °C overnight. SA-β-gal-positive area was quantified relative to the total analyzed tissue area in comparable anatomical regions.

For adipose tissue, freshly collected fat was fixed in 4% paraformaldehyde for 15 min, rinsed with PBS, and incubated in SA-β-gal staining solution at 37 °C overnight. Samples were subsequently immersed in PBS before imaging.

For adherent cultured cells, cells were fixed and stained using the same kit. SA-β-gal positivity in cultured cells was quantified as the percentage of SA-β-gal-positive cells among total cells. All quantification was performed using standardized image acquisition and predefined analysis criteria.

5.12. Immunohistochemistry and immunofluorescence staining

Immunohistochemistry and immunofluorescence staining were performed on paraffin or frozen sections according to standard protocols. For immunohistochemistry, sections were stained with antibodies against OCN, p16, HMGB1, CD11b, or CD19, followed by HRP-conjugated secondary antibodies and DAB development. Nuclei were counterstained with hematoxylin. For immunofluorescence staining, sections were incubated with antibodies against F4/80 together with iNOS or Arg-1 to evaluate macrophage polarization-associated features, or with antibodies against CD31 and EMCN to evaluate vascular features, with an antibody against CD19 to assess local B-cell-associated staining, or with antibodies against CD105 and p16 to evaluate senescence-associated stromal cells. Fluorescent secondary antibodies were applied, and nuclei were counterstained with DAPI. Images were acquired using light or fluorescence microscopy. IHC signals were quantified as positively stained area in randomly selected fields, whereas CD19⁺ staining, F4/80⁺iNOS⁺, F4/80⁺Arg-1⁺, and CD31⁺EMCN⁺ structures, and p16⁺CD105⁺ stromal cells were quantified from immunofluorescence images.

For γH2AX staining, adherent MSCs were seeded in glass-bottom confocal dishes and cultured under the indicated conditions until attachment. Cells were fixed with 4% paraformaldehyde, permeabilized, blocked, and incubated with an anti-γH2AX antibody overnight at 4 °C, followed by fluorescent secondary antibody incubation and DAPI counterstaining. For quantitative analysis, DAPI staining was used to identify and segment individual nuclei. Discrete γH2AX-positive foci within each nucleus were counted in randomly selected fields, and DNA damage was expressed as the mean number of γH2AX foci per nucleus. At least 50 nuclei from 5 randomly selected fields were analyzed for each biological replicate.

No-primary-antibody controls were used to assess nonspecific background staining in immunohistochemistry and immunofluorescence analyses. In these controls, the primary antibody was omitted, and samples were incubated only with the corresponding secondary antibody. These controls were processed in parallel with experimental samples under identical staining and imaging conditions, and quantification thresholds were set based on the observed background signals.

5.13. Micro-CT scanning and analysis

Samples were scanned using a micro-computed tomography system (SkyScan 1076, Bruker). Three-dimensional reconstruction was performed using VGStudio MAX software (Volume Graphics), and quantitative morphometric analysis was conducted using Scanco Medical software (Scanco Medical AG). Femoral and osteo-organoid datasets were analyzed independently, and statistical comparisons were performed only within the corresponding tissue context.

For femoral trabecular analysis, the region of interest was defined as a 2.0-mm metaphyseal region beginning 1.0 mm distal to the growth plate and extending toward the diaphysis. The growth plate and cortical shell were excluded from the analysis. A fixed threshold of 1100 was applied to all femoral samples. Parameters including BV/TV, Tb.N, Tb.Th, Tb.Sp, and BMD were calculated.

For osteo-organoids, the region of interest was defined by manually delineating the outer boundary of the entire construct on serial transverse sections, independently of mineralization intensity. Mineralized tissue within this construct-level ROI was segmented using a fixed grayscale threshold of 900, which was applied uniformly to all groups. Thus, total volume represented the entire osteo-organoid construct, whereas bone volume represented threshold-defined mineralized voxels within the construct. HAMA exhibited attenuation below the mineralization threshold and was therefore not included in the segmented mineralized volume or mineral-content measurements. Trabecular morphometric parameters were calculated from the segmented mineralized network within the predefined construct-level ROI. Representative ROI images are presented in Supplementary Fig. 17.

5.14. Preparation of single-cell suspensions for flow cytometry

For MSC analysis, femoral and osteo-organoid samples were processed using mechanical disruption followed by enzymatic digestion [43]. Femurs and tibiae were dissected, cut into small fragments, and processed in HBSS containing 2% FBS. Osteo-organoids were minced in multiple directions with curved-blade ophthalmic scissors. Samples from both tissue contexts were digested in HBSS containing collagenase (3 mg/mL; Gibco, 17018029), neutral protease (4 mg/mL; Roche, 4942078001), and DNase I (200 U/mL; Thermo Fisher Scientific, 89836) at 37 °C for 15 min. Digestion was stopped with Mg²⁺/Ca²⁺-free HBSS containing 2% FBS and 2 μM EDTA. Cells were collected by centrifugation, resuspended, and filtered through a 70 μm cell strainer.

For immune and endothelial profiling, femoral bone marrow cells were obtained by flushing the femoral marrow cavity with ice-cold PBS containing 2% FBS using a 26-G needle, followed by filtration through a 70-μm cell strainer. Osteo-organoids were mechanically dissociated in ice-cold PBS containing 2% FBS and similarly filtered through a 70-μm cell strainer. The resulting cell suspensions were treated with ACK red blood cell lysis buffer, and DNase I was added to minimize cell clumping.

5.15. Flow cytometry

Single-cell suspensions were stained with a near-infrared viability dye (Thermo Fisher Scientific, L10119) at 4 °C for 30 min, blocked with anti-mouse CD16/32 antibody, and incubated with surface-antibody cocktails at 4 °C for 30 min. For intracellular or nuclear markers such as p16, staining was performed using the True-Nuclear™ Transcription Factor Buffer Set (BioLegend, 424401) according to the manufacturer's protocol.

MSCs were defined as viable singlets that were CD45^-Ter119⁻CD11b⁻CD31⁻CD105⁺CD140a⁺. Endothelial cells were identified within the CD45^-Ter119⁻CD31⁺ population, and Type-H endothelial cells were analyzed using CD31 and Endomucin markers. Immune subsets were identified within the viable CD45⁺ population using lineage-specific markers.

Data were acquired on a CytoFLEX flow cytometer (Beckman Coulter) or an SFLO full-spectrum flow cytometer (Conpomed, China) and analyzed using FlowJo software. The complete gating strategy, fluorescence-minus-one controls, compensation matrices, and viability/doublet/debris exclusion steps are provided in Supplementary Figs. 18-20. Detailed antibody information is provided in Table S2.

5.16. Single-cell RNA sequencing and analysis

Single-cell RNA sequencing library construction and deep sequencing were performed by BGI Tech Solutions Co., Ltd./BGI Genomics (Shenzhen, China). Briefly, single-cell suspensions were first assessed by microscopy after 0.4% trypan blue or AOPI staining to evaluate cell viability, aggregation, background debris, and sample quality. Qualified single-cell suspensions were subjected to library preparation using the DNBelab C4 single-cell RNA-seq platform according to the manufacturer's protocol. This platform is based on droplet microfluidics and bead-based mRNA capture. Single cells, oil, and capture beads were loaded onto the DNBelab C4 system to generate water-in-oil droplets, in which cells were lysed and mRNA molecules were captured by barcoded beads. Reverse transcription was then performed to generate cDNA, followed by emulsion breaking, cDNA amplification, fragmentation, end repair, A-tailing, adaptor ligation, PCR amplification, and library purification. Libraries were quality-checked, circularized to generate single-stranded circular DNA molecules, and converted into DNA nanoballs by rolling-circle amplification. Sequencing was performed on the DNBSEQ platform using combinatorial probe-anchor synthesis technology.

Raw sequencing data from each sample were processed using DNBelab C4scRNA (v1.0.1) to generate raw gene expression matrices. Downstream analysis was performed using Seurat (v3.2.0). Cells were retained for analysis if they contained more than 200 detected genes and less than 15% mitochondrial gene expression. Potential doublets were identified using the DoubletFinder package, and only cells classified as singlets were retained for downstream analysis. After quality control, samples were merged, and batch effects were corrected before integrated analysis.

The integrated gene expression matrix was normalized, and the top 2000 highly variable genes were selected for principal component analysis. The first 15 principal components were used for downstream dimensionality reduction and clustering. tSNE and UMAP were used to visualize cellular heterogeneity. Marker genes for each cluster were identified using the FindAllMarkers function in Seurat with logFC > 0.25, min.pct > 0.1, and adjusted P value ≤ 0.05. Cell types were annotated using SCSA based on the identified marker genes.

5.17. MSC culture, CFU-F assay, and multilineage differentiation

After isolation, cells were seeded in α-MEM supplemented with 20% FBS, 1% sodium pyruvate, 1% penicillin-streptomycin, and 10 μM ROCK inhibitor. After 12 h, non-adherent cells were removed by washing with PBS, and adherent MSCs were maintained in fresh culture medium. MSCs at passage P2 were used for downstream assays.

For CFU-F assays, 500 purified MSCs were seeded per well in six-well plates and cultured for 8 days at 37 °C in a humidified incubator containing 5% O₂. Cells were fixed in 4% paraformaldehyde for 30 min and stained with 0.1% crystal violet (Servicebio, G1014). Colonies were imaged using an APX100 all-in-one fluorescence imaging system (Olympus). Colony-forming efficiency was expressed as the number of colonies per 500 MSCs initially seeded.

For osteogenic and adipogenic differentiation assays, 3 × 10⁵ MSCs were seeded per well in six-well plates coated with 0.1% gelatin. Osteogenic and adipogenic induction was performed using OriCell™ Mesenchymal Stem Cell Osteogenic Differentiation Medium (Cyagen, MUXMX-90021) and Adipogenic Differentiation Medium (Cyagen, MUXMX-90031), respectively, according to the manufacturer's instructions. Osteogenic differentiation was evaluated by Alizarin Red S staining, and adipogenic differentiation was evaluated by Oil Red O staining. Quantification was performed using identical acquisition and analysis settings across groups.

5.18. In vitro irradiation of MSCs

MSCs isolated from femora and osteo-organoids were cultured under identical conditions and subjected to in vitro irradiation at 6 Gy with a dose rate of 1.28 Gy/min using an RS2000 X-ray irradiator. Cells were collected at 48h after irradiation for apoptosis analyses.

5.19. Isolation and polarization of bone marrow-derived macrophages

Bone marrow cells were obtained by flushing femurs from 8 to 12-week-old C57BL/6 mice with DMEM supplemented with 10% FBS, 10 ng/mL recombinant murine M-CSF (PeproTech, 315-02), and 1% penicillin-streptomycin. Cells were seeded in 100 mm culture dishes and maintained at 37 °C in a humidified incubator. After 14 h, non-adherent cells were collected and centrifuged at 300 × g for 10 min to obtain bone marrow-derived monocytes. M0 macrophages were generated by culturing monocytes in DMEM containing 10% FBS and 30 ng/mL recombinant murine M-CSF for 3 days. For M1-like polarization, M0 macrophages were cultured in DMEM containing 10% FBS, 30 ng/mL recombinant murine IFN-γ, and 100 ng/mL LPS for 24 h.

5.20. Macrophage-MSC co-culture

Macrophage-MSC co-cultures were established using a Transwell system at a macrophage-to-MSC ratio of 3:1. Macrophages were seeded in the lower chamber of 12-well plates at 3 × 10⁵ cells per well in co-culture medium consisting of α-MEM:DMEM = 1:1 supplemented with 10% FBS and 1% penicillin-streptomycin. After overnight adherence, MSCs were seeded into 0.4 μm pore-size polyester Transwell inserts (Corning) at 1 × 10⁵ cells per insert and placed above the macrophage layer. Co-cultures were maintained for 48 h. At the endpoint, macrophages and MSCs were harvested separately for flow cytometry and qPCR.

For flow cytometry, Macrophage phenotypes were assessed based on CD86 and CD206 expression. CD86⁺CD206⁻ macrophages were quantified as the CD86 single-positive population, whereas total CD206⁺ macrophages included both CD86⁻CD206⁺ and CD86⁺CD206⁺ populations. Because a substantial fraction of CD206-expressing cells co-expressed CD86, these populations were not interpreted as mutually exclusive M1-and M2-like macrophage states.

5.21. Western blotting

For western blotting, total protein was extracted from cells using ice-cold radioimmunoprecipitation assay (RIPA) buffer containing phenylmethylsulfonyl fluoride (PMSF). Cell lysates were collected, mixed with loading buffer, and boiled for 10 min. Equal amounts of protein were separated by 8% SDS-PAGE and transferred onto polyvinylidene fluoride membranes (Millipore). The membranes were blocked with QuickBlock™ Western Blocking Buffer (Beyotime Biotechnology, Shanghai, China) at room temperature and incubated with the indicated primary antibodies overnight at 4 °C. After washing with TBST, the membranes were incubated with HRP-conjugated secondary antibodies. GAPDH was used as a loading control. Protein bands were visualized using a chemiluminescence imaging system (Tanon, Shanghai, China). Band intensity was quantified using ImageJ. Antibody information is provided in Table S2.

5.22. Quantitative real-time PCR

Total RNA was extracted using TRIzol reagent and reverse-transcribed using the PrimeScript RT reagent kit (Takara, RR047A) according to the manufacturer's instructions. qPCR was performed on a CFX96 Touch Real-Time PCR Detection System (Bio-Rad, USA) using TB Green Premix Ex TaqII (Takara, RR420A) according to the manufacturer's instructions. The cycling program consisted of 95 °C for 30 s, followed by 40 cycles of 95 °C for 5 s and 60 °C for 30 s. Mouse Gapdh was used as the endogenous reference gene. Each biological sample was analyzed in three technical replicates, and the mean Ct value was used for subsequent calculation. Relative gene expression was calculated using the 2^−ΔΔCt method. Primer sequences and primer-efficiency information are provided in Table S1. Melting curve and titration curve analyses are presented in Supplementary Fig. 21.

5.23. Irradiation-impaired bone-defect model and BMP-2/HAMA treatment

To evaluate functional bone repair under irradiation stress, a distal femoral bone defect model was established in TBI-exposed mice. Mice were assigned to the following groups: Defect, TBI + Defect, TBI + HAMA, and TBI + BMP-2/HAMA. As described above, mice in the irradiation groups were subjected to 6 Gy total body irradiation. At 24 h after TBI, mice were anesthetized with isoflurane, and the distal femur was exposed through a small lateral incision under aseptic conditions. A standardized monocortical circular defect with a diameter of 1.2 mm was created using a dental drill without penetrating the opposite cortex. The indicated scaffolds were implanted into the defect site, and the wound was closed in layers.

For postoperative analgesia, meloxicam was administered subcutaneously at 5 mg/kg every 24 h for 72 h after surgery. Mice were maintained on a warming pad until recovery from anesthesia and then returned to their cages. Bone repair was evaluated at the indicated time points after surgery by micro-CT, histological analysis, mechanical testing, and gait analysis.

No animals were excluded from the femoral defect experiment because of postoperative complications, and the final sample sizes are reported in the corresponding figure legends.

5.24. Biomechanical testing

At 4 weeks after surgery, femora were harvested for biomechanical testing. Three-point bending tests were performed using a dynamic biomaterial mechanical testing system (ElectroForce 3230 Series III, Waters, USA). Each femur was placed on two lower supports with the defect region positioned centrally between the supports, and load was applied vertically to the femoral shaft until failure. Maximum load and fracture energy were calculated from the load-displacement curves.

5.25. Gait analysis

Gait performance was evaluated at 2 weeks after bone defect surgery using a CatWalk gait analysis system (Noldus Information Technology). Mice were allowed to walk freely across the illuminated glass walkway, and valid runs were recorded according to the manufacturer's instructions. Gait parameters, including regularity index, maximum contact intensity, and paw print area, were analyzed to assess limb use and functional recovery.

5.26. Blinding and randomization

Animals were randomly assigned to experimental groups. For image-based analyses, micro-CT ROI delineation, histomorphometric quantification, immunostaining quantification, and flow-cytometry gating review, samples were coded before analysis, and investigators were blinded to group allocation until quantification was completed.

5.27. Statistical analysis

Statistical analysis was performed using GraphPad Prism 10.0.3. Data are presented as mean ± SD unless otherwise stated. Normality was assessed using the Shapiro-Wilk test. For comparisons between two independent groups, an unpaired two-tailed Student's t-test was used when data were normally distributed; otherwise, the Mann-Whitney U test was used. For p21 Western blot densitometric analysis, three independent biological replicates were included per group, and comparisons among Ctrl, TBI-BMSC, and TBI-HMSC groups were performed using the Kruskal-Wallis test followed by Dunn's multiple-comparisons test. For comparisons among more than two groups with one experimental factor, one-way ANOVA followed by Tukey's multiple-comparisons test was used. For analyses involving two experimental factors, such as irradiation condition and time, two-way ANOVA followed by Sidak's multiple-comparisons test was used. Longitudinal 1-, 3-, and 6-week datasets were derived from independent terminal cohorts and were therefore analyzed as independent samples rather than repeated measures. A P value below 0.05 was considered statistically significant. Significance levels are indicated as *P < 0.05, **P < 0.01, ***P < 0.001, and ****P < 0.0001. In all graphs, n indicates the number of independent biological replicates, animals, or independent osteo-organoids, as specified in the corresponding figure legends.

CRediT authorship contribution statement

Fuwei Zhu: Writing – original draft, Visualization, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Luli Ji: Writing – original draft, Methodology, Investigation, Formal analysis, Data curation, Conceptualization. Jing Wang: Writing – review & editing, Supervision, Funding acquisition. Changsheng Liu: Writing – review & editing, Supervision, Funding acquisition.

Ethics approval and consent to participate

All animal procedures were approved by the Animal Care and Use Committee of the East China University of Science and Technology (Approval No. ECUST-2024-022).

Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Acknowledgments

Our x-ray irradiation work was performed at the National Center for Protein Science (Shanghai) with the aid of C. Zheng and H. Chen. We express our sincere gratitude to Shanghai Tuling Biotechnology Co., Ltd for their invaluable assistance with data analysis. We also extend our heartfelt thanks to Yuanhao Zhou,Xiaochao Zhao and Jianpeng Zhang for their professional support and insightful contributions throughout the data analysis process. This research was supported by the Key Program of the National Natural Science Foundation of China (No. 32230059), the Excellence Research Group Program of National Natural Science Foundation of China (No. T2288102), the National Natural Science Foundation of China (No. 32471406), the Foundation of Frontiers Science Center for Materiobiology and Dynamic Chemistry (JKVD1211002), and the Young Scientists Fund of the National Natural Science Foundation of China (No. 32401128).

Footnotes

Peer review under the responsibility of editorial board of Bioactive Materials.

^{Appendix A}

Supplementary data to this article can be found online at https://doi.org/10.1016/j.bioactmat.2026.08.042.

Contributor Information

Jing Wang, Email: wangjing08@ecust.edu.cn.

Changsheng Liu, Email: liucs@ecust.edu.cn.

Appendix A. Supplementary data

The following is the Supplementary data to this article:

Multimedia component 1

mmc1.docx^{(14.3MB, docx)}

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[bib1] 1.Williams H.J., Davies A.M. The effect of X-rays on bone: a pictorial review. Eur. Radiol. 2006;16:619–633. doi: 10.1007/s00330-005-0010-7. [DOI] [PubMed] [Google Scholar]

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[bib3] 3.Baxter N.N., Habermann E.B., Tepper J.E., Durham S.B., Virnig B.A. Risk of pelvic fractures in older women following pelvic irradiation. JAMA. 2005;294:2587–2593. doi: 10.1001/jama.294.20.2587. [DOI] [PubMed] [Google Scholar]

[bib4] 4.Zhou B.O., Yu H., Yue R., Zhao Z., Rios J.J., Naveiras O., Morrison S.J. Bone marrow adipocytes promote the regeneration of stem cells and haematopoiesis by secreting SCF. Nat. Cell Biol. 2017;19:891–903. doi: 10.1038/ncb3570. [DOI] [PMC free article] [PubMed] [Google Scholar]

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PERMALINK

BMP-2-driven osteo-organoid formation retains key osteogenic-support features and promotes bone repair following total-body irradiation

Fuwei Zhu

Luli Ji

Jing Wang

Changsheng Liu

Abstract

Graphical abstract

Highlights

1. Introduction

2. Results

2.1. TBI induces systemic injury and progressive femoral bone deterioration

Fig. 1.

2.2. TBI disrupts immune, vascular, and MSC-associated components of the femoral osteogenic-support microenvironment

Fig. 2.

2.3. Single-cell profiling reveals B-cell contraction and neutrophil-associated myeloid remodeling after TBI

Fig. 3.

2.4. BMP-2/HAMA-induced osteo-organoids show delayed but sustained maturation under irradiation stress

Fig. 4.

2.5. TBI delays coordinated immune and vascular remodeling during osteo-organoid maturation

Fig. 5.

2.6. Osteo-organoid-derived MSCs retain functional and immunomodulatory properties after irradiation

Fig. 6.

2.7. BMP-2/HAMA promotes bone repair under irradiation-impaired conditions

Fig. 7.

3. Discussion

4. Conclusions

5. Materials and methods

5.1. Animals and ethics statement

5.2. Total body irradiation model

5.3. Synthesis and fabrication of HAMA scaffolds

5.4. Characterization of HAMA scaffolds and BMP-2 release

5.5. Subcutaneous implantation and osteo-organoid induction

5.6. Early BMP-Smad inhibition

5.7. Sample collection and processing

5.8. Peripheral blood analysis and bone marrow multiplex cytokine assay

5.9. ELISA quantification of tissue and MSC-secreted factors

5.10. Histological staining

5.11. SA-β-gal staining and quantification

5.12. Immunohistochemistry and immunofluorescence staining

5.13. Micro-CT scanning and analysis

5.14. Preparation of single-cell suspensions for flow cytometry

5.15. Flow cytometry

5.16. Single-cell RNA sequencing and analysis

5.17. MSC culture, CFU-F assay, and multilineage differentiation

5.18. In vitro irradiation of MSCs

5.19. Isolation and polarization of bone marrow-derived macrophages

5.20. Macrophage-MSC co-culture

5.21. Western blotting

5.22. Quantitative real-time PCR

5.23. Irradiation-impaired bone-defect model and BMP-2/HAMA treatment

5.24. Biomechanical testing

5.25. Gait analysis

5.26. Blinding and randomization

5.27. Statistical analysis

CRediT authorship contribution statement

Ethics approval and consent to participate

Declaration of competing interest

Acknowledgments

Footnotes

Contributor Information

Appendix A. Supplementary data

References

Associated Data

Supplementary Materials

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