Abstract
The treatment of osteoporosis and related bone defects remains challenging. This study identifies pyroptosis-driven inflammation as a key disruptor of bone homeostasis. To address this, we develop a magnesium-gelatin composite microsphere scaffold (GelMa/Mg/DMF MS) that exploit pyroptosis blockade and hydrogen-mediated inflammation regulation for osteoporosis treatment. This porous microsphere scaffold is implanted into bone defects to achieve the sustained release of hydrogen gas, magnesium ions (Mg2+), and dimethyl fumarate (DMF). DMF act by activating the nuclear factor erythroid-related factor 2 to prevent osteoblast pyroptosis, and combine with the antioxidant effects of hydrogen, effectively remodel the inflammatory microenvironment and create favorable conditions for the restoration of bone homeostasis. Mg2+ further expedite bone tissue repair. These results demonstrate that the GelMa/Mg/DMF MS effectively reverse inflammatory microenvironments both in vivo and in vitro, resulting in significant tissue repair. These results suggest the combination of hydrogen therapy and pyroptosis blockade as a potential therapeutic strategy.
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Introduction
Osteoporosis is a systemic skeletal disease characterized by reduced bone mass and microstructural deterioration1,2. The resulting comminuted fractures often lead to severe complications and significant bone defects, in which healing is particularly difficult to achieve under osteoporotic conditions3,4,5. With the aging of the global population, the incidence of osteoporosis and related bone defects is increasing annually, highlighting the pressing need for effective treatment options6,7. The traditional treatment of osteoporosis is conservative and includes oral calcium, vitamin D, and bisphosphonates, as well as subcutaneous injections of drugs, such as a monoclonal antibody targeting receptor activator of nuclear factor kappa-B ligand (RANKL, e.g., denosumab), a selective estrogen receptor modulator (e.g., raloxifene), and a parathyroid hormone analog (e.g., teriparatide)8,9. While these methods achieve clinical efficacy, they are also accompanied by several side effects, such as gastrointestinal reactions with alendronate, the risk of venous thromboembolism with raloxifene, and the risk of hypocalcemia and osteonecrosis of the jaw with teriparatide10,11,12. Recent research reports osteoporosis as a systemic inflammatory disease with an inflammatory microenvironment (IME) that increases osteoclast activity, thus impairing tissue repair and bone homeostasis13,14. This poses a major challenge to the efficacy of conventional treatments. Thus, targeting the IME to inhibit osteoclast function appears to be a promising approach for osteoporosis management15,16. While anti-inflammatory drugs are used clinically, they face challenges such as undesirable first-pass effects and delivery issues17. For example, ibuprofen, a non-steroidal anti-inflammatory drug, has a first-pass metabolism rate of approximately 60%, which reduces its bioavailability to around 80%18. Consequently, elevated dosages are necessary to attain the anticipated therapeutic outcomes, which can, in turn, increase the risk of gastrointestinal reactions, including ulcers. Therefore, exploring and developing more effective treatment strategies for osteoporosis remains crucial.
Hydrogen (H2) therapy is an emerging treatment modality that shows considerable promise in medicine19. H2 molecules have unique physical and chemical properties that allow them to exert localized anti-inflammatory effects by modulating the IME19,20,21,22. Moreover, using H2 as a therapeutic agent offers advantages such as high in situ synthesis efficiency, stability, rapid diffusion, and widespread distribution23,24. Additionally, as a gas, it can accelerate the release and diffusion of medications. However, since H2 therapy primarily acts through antioxidant mechanisms or by regulating inflammatory gene expression, it does not address the root causes of the IME, and the continuous production of inflammatory mediators may challenge the long-term efficacy of gas therapy. Thus, combining strategies that target initial inflammation sources using hydrogen therapy may provide a synergistic effect, effectively alleviating the IME in osteoporosis.
Pro-inflammatory forms of programmed cell death, such as ferroptosis, necroptosis, and pyroptosis, are recognized as crucial contributors to inflammation in various inflammatory diseases25,26,27. Inhibiting pathways related to inflammatory cell death has proven effective in reducing inflammation28,29. Pyroptosis, a highly inflammatory form of programmed cell death, can be triggered by various stimuli, including damage-associated molecular patterns, DNA damage, inflammatory cytokines, and related oxygen species (ROS)30,31. During pyroptosis, inflammatory cytokines such as interleukin-1β (IL-1β) and IL-18 are released into the local microenvironment, initiating an inflammatory cascade that severely hampers tissue repair32. In the presence of inflammation, osteoblast differentiation and tissue repair processes are diminished, while osteoclast progression is accelerated, thereby establishing a negative cycle that impedes the healing of bone tissue33,34. Thus, pyroptosis plays a pivotal role in promoting osteoporosis progression and inhibiting bone reconstruction. Based on these findings, we hypothesize that combining pyroptosis inhibition with hydrogen therapy could constitute a dual-pathway treatment approach that simultaneously inhibits inflammation sources and regulates the IME in osteoporosis.
Herein, we developed a magnesium-based polymer microsphere scaffold (MS) loaded with the Food and Drug Administration (FDA)-approved pyroptosis inhibitor dimethyl fumarate (DMF), aiming to achieve both hydrogen-mediated inflammation modulation and pyroptosis inhibition-induced inflammation blockade. The porous structure of the GelMa/Mg/DMF MS enhances fluid infiltration at the implantation site35. Internal magnesium spheres react with surrounding bodily fluids to generate anti-inflammatory hydrogen and reparative Mg2+, modulating the IME36,37. More importantly, the release and diffusion of H2 within the GelMa/Mg/DMF MS further accelerate the diffusion of the pyroptosis inhibitor DMF into the environment and inhibit osteoblast pyroptosis by inhibiting the NOD-like receptor thermal protein domain-associated protein 3/cysteinyl aspartate specific proteinase 1/gasdermin D (NLRP3/CASP1/GSDMD) signaling axis, thereby regulating the IME. The present study combined hydrogen therapy with pyroptosis blockade to treat osteoporosis, offering some potential future therapeutic strategy for the design and development of effective OP treatment strategies.
Results
Pyroptosis disrupts bone homeostasis in osteoporosis
A total of 32 patients with osteoporosis and 61 patients with normal bone mineral density (BMD) were included in the study to determine whether pyroptosis could disrupt bone homeostasis in osteoporosis. Patients with tumors, severe infections, or trauma were excluded. Table S1 summarizes the general information and bone metabolism indicators of these patients. Notably, no statistically significant difference was found in age between the two groups of patients. This is important because it helps to minimize the potential impact of age on inflammatory indicators. Osteoporosis was defined as a T-score on dual-energy X-ray absorptiometry (DXA) of less than −2.5, in accordance with methods established in clinical practice (Fig. 1A)3,38. Blood samples were collected, and the expression of several inflammatory factors was analyzed in serum using enzyme-linked immunosorbent assay (ELISA) (Fig. 1B), which revealed elevated levels of inflammatory factors, such as IL-1β and IL-18, in patients with osteoporosis compared with controls (Fig. S1A). These findings suggest that osteoporosis is not merely a localized alteration of the bone microstructure, but rather a systemic inflammatory disease. IL-1β and IL-18 are common inflammatory factors in the IME and serve as downstream signaling molecules in pyroptosis. Thus, their levels can indicate the extent of pyroptosis31. Further analysis revealed a negative correlation between BMD T-scores and the levels of these inflammatory factors, indicating that more severe osteoporosis was associated with higher IL-1β (R2 = 0.0696, p = 0.0106) and IL-18 levels (R2 = 0.1275, p = 0.0004) (Fig. 1C). Additionally, femur samples from patients with osteoporosis presented sparser and thinner trabeculae, increased trabecular separation, and activated osteoclasts, as identified by tartrate-resistant acid phosphatase (TRAP) staining (Fig. S1 B–C). Subsequent immunohistochemical staining revealed elevated levels of GSDMD, CASP1, and NLRP3 in patients with osteoporosis, providing strong evidence for the activation of the classical pyroptosis pathway in osteoporosis (Figs. 1D and S1D–F). Thus, a positive association between osteoporosis and pyroptosis could be inferred from the serological and histological observations. However, the specific cell type responsible for the promotion of pyroptosis remains unclear.
A Pyroptosis disrupts bone homeostasis in patients with osteoporosis. (created with BioRender.com) B Inflammatory cytokine levels in the serum of patients. C Correlations between the serum levels of the cytokines interleukin-1β (IL-1β) and IL-18 and the T-score of bone mineral density (BMD) in patients. D Immunohistochemical staining of GSDMD, CASP1 and NLRP3 in patient bone tissue samples. Scale bars: 100 μm. E X-ray image of c57 mice (the white dashed box illustrated the microcomputed tomography (micro-CT) image of the lumbar spine). F The BMD of the C57 mouse lumbar spine and tibia was determined via dual-energy X-ray absorptiometry (DXA) method (n = 5 samples). G Changes in the level of inflammatory factors in c57 serum were detected via enzyme-linked immunosorbent assay (ELISA) (n = 3 samples). H Images of tibial sections after immunohistochemistry for GSDMD. Scale bars: 100 μm. I Immunofluorescence staining of OPN (green) and propidium iodide (PI, red) in the tibial. Scale bars: 100 μm. J Co-localization of OPN with PI was calculated via ImageJ. K Pyroptosis-related protein expression in bone marrow mesenchymal stromal cells (BMSCs) was detected by Western blotting. L Expression of GSDMD in BMSCs was detected by confocal laser scanning microscopy (CLSM). Scale bars: 50 μm. M Expression of NLRP3 in BMSCs was detected by CLSM. Scale bars: 50 μm. N Transmission electron microscopy (TEM) image of the BMSCs. Scale bars: 5 μm and 1 μm. O Quantitative analysis of the expression of NLRP3 and GSDMD in BMSCs was detected by CLSM (n = 5 independent experiments). P Alkaline phosphatase (ALP) staining of BMSCs after 14 days. Scale bars: 200 μm. Q Alizarin red S (ARS) staining of BMSCs after 21 days. Scale bars: 200 μm. R Tartrate-resistant acid phosphatase (TRAP) staining of bone marrow-derived macrophages (BMDMs) for 7 days. Scale bars: 50 μm. S Osteoclast ring of BMDMs was detected via CLSM. Scale bars: 100 μm. T Number of osteoclasts per field of view (n = 5 independent experiments). The data are presented as the means ± SD. Statistical significance was calculated by one-way analysis of variance (ANOVA) with Tukey’s post hoc test. Source data are provided as a Source Data file.
Bilateral ovariectomy (OVX) was performed on C57 mice to simulate estrogen-deficient osteoporosis to verify this hypothesis39. Eight weeks post-surgery, DXA revealed a significant reduction in BMD in the OVX group (Fig. 1E–F)40. In addition, a significant increase in serum levels of inflammatory factors such as IL-18 was observed in ovariectomized mice, suggesting an increase in inflammatory factors in vivo after ovariectomy, whereas these effects were not seen in the sham group (Figs. 1G and S1G). Histological analysis confirmed osteoporosis in the tibia, and hematoxylin and eosin (H&E) staining revealed reduced trabeculae and increased inflammatory cells (Fig. S1H). The application of Van Gieson (VG) stain facilitates the observation of red collagen and yellow mineralized bone matrix in bone tissue, thereby providing a comprehensive representation of the microscopic features of the bone tissue. The area of collagen deposition in the bone tissue was found to be decreased following OVX. Two months after ovariectomy, the mice exhibited sparse trabeculae, characterized by a decrease in trabecular area percentage and trabecular narrowing (Fig. S1I–J). Immunohistochemical analysis revealed GSDMD overexpression in the OVX group (Fig. 1H). Prior to euthanization, the C57 mice were received an intraperitoneal injection of propidium iodide (PI) to label dead cells, and osteopontin (OPN, a signature protein of osteoblasts) was immunofluorescently labeled (Fig. 1I)41. OPN and PI were co-localized after OVX, which suggests that osteoblasts undergo cell death in osteoporosis, likely through pyroptosis (Figs. 1J and S1K). Therefore, following the onset of osteoporosis, osteoblasts might undergo pyroptosis, which could not only result in bone loss but also trigger a systemic inflammatory response. Nevertheless, whether the osteogenic differentiation capacity of pyroptotic osteoblasts is affected and whether osteoclasts are involved in this process remain unclear.
We induced pyroptosis in bone marrow-derived mesenchymal stromal cells (BMSCs) using the well-known pyroptosis inducers lipopolysaccharides (LPS) and adenosine 5’-triphosphate (ATP) to investigate this phenomenon31,42. A comparison of the expression of pyroptosis-associated proteins by Western blotting (WB) analysis (Fig. S2A–B) and the levels of IL-18 and IL-1β in the cell supernatants by ELISA (Fig. S2C–D) revealed that 1 μg/mL LPS and 5 μM ATP significantly induced pyroptosis in the BMSCs. WB analysis demonstrated a stable increase in the number of N-terminal GSDMD fragments in BMSCs subsequent to pyroptosis, accompanied by considerable increase in the levels of CASP1 fragments and IL-18 (Figs. 1K and S3A). These findings suggest that GSDMD was cleaved by cleaved-CASP1 to generate N-GSDMD fragments, which possessed pore-forming activity. The GSDMD expression and its co-localization with the cell membrane marker Dil revealed that GSDMD aggregated on the cell membrane surface following activation (Fig. 1L). Light microscopy revealed that the BMSCs exhibited a rounded morphology and produced small bubbles following induction (Fig. 1M). Further detection of NLRP3 expression using confocal laser scanning microscopy (CLSM) indicated that NLRP3 expression in the BMSCs was also significantly elevated following pyroptosis. NLRP3, a member of the NOD-like receptor (NLR) subfamily, was identified as an upstream regulator of pyroptosis. NLRP3, in conjunction with the aptamer ASC protein PYCARD, forms a CASP1 activation complex and designates the NLRP3 inflammasome43,44. Subsequently, activated CASP1 induces the synthesis of pro-inflammatory cytokines and triggers the activation of GSDMD at the cellular membrane, which permits the release of cytokines into the extracellular space. Cell membrane rupture was observed in the pyroptosis-induced group using transmission electron microscopy (TEM), while this phenomenon was not detected in the control group (Fig. 1N). Thus, whether pyroptosis affected the osteogenic capacity of osteoblasts was investigated using alkaline phosphatase (ALP) and alizarin red S (ARS) staining. Pyroptosis impaired osteogenic differentiation in BMSCs, as indicated by reduced ALP and ARS staining (Figs. 1P, Q and S3B, C). The above results indicate that numerous inflammatory mediators were released following the onset of pyroptosis and that the pyroptosis-activated IME disrupts the ability of osteoblasts to repair tissue and promote osteogenic differentiation.
We co-cultured BMSCs and bone marrow-derived macrophages (BMDMs) in a Transwell system to explore whether this disruption affects bone homeostasis. TRAP staining revealed multinucleated giant cells, indicating osteoclast activation (Fig. 1R). An increase in the number of nuclei in these cells showed that osteoclast activity increased after osteoblast pyroptosis (Fig. S3D). Immunofluorescence staining revealed that activated osteoclasts formed a ring structure (Fig. 1S, T), with increased diameter correlated with increased osteoclast activity (Fig. S3E). Scanning electron microscopy (SEM) of bovine bone slices co-cultured with osteoclasts confirmed that pyroptosis in osteoblasts enhanced the bone-destructive capacity of osteoclasts, as indicated by larger eroded areas (Fig. S3F, G). Overall, osteoblast pyroptosis following osteoporosis not only impaired osteogenic differentiation but also promoted osteoclast activation, creating a vicious cycle that exacerbated osteoporosis. Therefore, novel strategies should be developed to break this cycle for effective osteoporosis management.
Fabrication and material characterization of the GelMa/Mg/DMF MS
In our study, the progression of osteoporosis has been demonstrated to be correlated with the level of pyroptosis in both clinical samples and animal models. Furthermore, pyroptosis has been shown to negatively impact tissue repair, and the pyroptosis-activated IME can induce osteoclasts, thereby accelerating the development of osteoporosis45. Thus, we designed a materials that not only modulated existing inflammation but also targeted the primary source of inflammation, thereby achieving a dual-pronged therapeutic approach for osteoporosis. Specifically, we successfully synthesized a GelMa/Mg/DMF MS by ultraviolet (UV) curing following initiator activation (Fig. S4A). This microsphere contained a gelatin framework and internal Mg spheres and DMF encapsulation. SEM images demonstrated that the simple GelMa MS exhibited a stable flower-like structure (Fig. 2A). Upon wrapping multiple Mg spheres, GelMa/Mg and GelMa/Mg/DMF MSs were larger in size than GelMa MS and maintained excellent spherical stability. Additionally, the surface exhibited a porous structure of uniform size, which could be advantageous for the controlled release of the contents. Energy dispersive spectroscopy confirmed stable Mg encapsulation (Fig. 2B, C). X-ray diffraction was used to characterize the crystalline structure (Fig. 2D), further suggesting the successful encapsulation of Mg inside the microspheres. The GelMa/Mg/DMF MS demonstrated efficient Mg ion release, with the rate dependent on the pH of the surrounding buffers. The degradation rate and Mg release were accelerated under acidic conditions (pH 6.4 and 5.5) compared with neutral conditions (pH = 7.4) (Fig. 2E). The structural integrity of the microspheres was maintained for up to 28 days, during which time most of the Mg ions were released (Fig. 2F). Gas release was still visible in the microspheres under optical microscopy after 28 days, suggesting a long-lasting release effect. In osteoporosis, osteoclast activation and the release of associated inflammatory factors tend to create an acidic bone metabolic microenvironment46. Thus, the acidic response of the GelMa/Mg/DMF MS may facilitate the release of Mg and DMF, which could enhance osteoblast activity and inhibit osteoclast activity. Furthermore, substantial DMF release was observed during this process (Fig. 2G). The released DMF was hypothesized to be capable of remodeling the IME in the osteoporotic state.
A Scanning electron microscopy (SEM) images of the different samples. Scale bars: 20 μm. B Element mapping images of the GelMa/Mg/DMF MS. Scale bars: 20 μm. C Energy dispersive spectroscopy (EDS) spectrum of the GelMa/Mg/DMF MS. D X-ray diffraction (XRD) patterns of the different samples. E Mg ion release of GelMa/Mg/DMF MS detected by inductively coupled plasma-optical emission spectrometry (ICP) (n = 3 independent experiments). F Microsphere morphology of GelMa/Mg/DMF MS under light microscopy at different times of degradation (schematic diagram is shown in the upper right corner, created with BioRender.com). Scale bars: 20 μm. G Detection of dimethyl fumarate (DMF) release via UV-vis spectroscopy (n = 5 independent experiments). H Detection of hydrogen release from materials by reduction of methylene blue. I Detection of the total antioxidant capacity of GelMa/Mg/DMF MS via the 3, 3’, 5, 5’-tetramethylbenzidine (TMB) method. The data are presented as the means ± SD. Source data are provided as a Source Data file.
In addition, hydrogen gas released from the GelMa/Mg/DMF MS was detected using methylene blue (MB) assays, with platinum catalysis facilitating a redox reaction that transformed blue oxidized MB to colorless reduced MB in fewer than 3 min (Fig. 2H)47,48. The oxide tetramethylbenzidine (oxTMB) and 2,2’-azino-bis (3-ethylbenzothiazoline-6-sulfonic acid) (ABTS+) assays, which are commonly used for antioxidant detection49, revealed significant changes in specific peaks in UV-vis spectroscopy, indicating their ability to scavenge ROS (Figs. 2I and S4B). The findings suggest a reduction in absorbance with the introduction of the Mg-containing MS (GelMa/Mg and GelMa/Mg/DMF MSs), indicating that the materials possess robust antioxidant capability, depending on both time and concentration (Fig. S4C–E). The GelMa/Mg/DMF MS was capable of scavenging ABTS+ radicals at a fast rate through continuous H2 generation, even at a lower concentration, which resulted in a decreased absorbance. These findings indicated that the GelMa/Mg/DMF MS was capable of producing a specific quantity of H2 at the initial stage of bone repair, which exerted beneficial local anti-inflammatory and antioxidant effects. Consequently, the targeted modulation of the IME generated by pyroptotic osteoblasts in osteoporosis was successfully achieved using the GelMa/Mg/DMF MS. The GelMa/Mg/DMF MS effectively targeted the early inflammatory response by releasing H2, and DMF blocked the malignant cycle of osteoblasts and osteoclasts and ultimately reshaped bone homeostasis in osteoporosis.
Modulation of the IME by GelMa/Mg/DMF MSs
Co-culturing BMSCs with GelMa/Mg/DMF MSs and subsequent live/dead cell staining revealed minimal red fluorescence, indicating high cytocompatibility (Figs. 3A and S5A). The GelMa/Mg/DMF MSs also exhibited favorable biocompatibility with Raw 264.7 cells, which was further confirmed by methylthiazolyldiphenyl-tetrazolium bromide (MTT) assays at different time points (Fig. 3B). Flow cytometry analysis revealed macrophage polarization changes in Raw 264.7 cells (Fig. 3C). The express of the cell marker CD86 was increased in M1-type macrophages treated with LPS (3.55 ± 0.10 vs. 6.24 ± 0.16%), whereas cells treated with GelMa/Mg/DMF MSs were resistant to M1 polarization induced by LPS (4.76 ± 0.11%). In contrast, the level of the cellular marker CD206 was increased in M2-type macrophages after pretreatment with GelMa/Mg/DMF MSs compared to the LPS-treated group (33.2 ± 1.47% vs. 0.23 ± 0.02%) (Fig. S5B). The shift in the macrophage phenotype from the M1-type to the M2-type was attributed to the attenuation of the LPS-induced inflammatory response by hydrogen synergized with DMF after pretreatment with GelMa/Mg/DMF MSs. ROS are highly reactive molecules that play pivotal roles in the pathogenesis and treatment of inflammatory diseases19,50. Dichlorodihydrofluorescein diacetate (DCF) is a valuable instrument for the immediate assessment of the intracellular redox status, with the degree of DCF fluorescence reflecting the degree of ROS. CLSM revealed that LPS treatment elevated DCF levels in Raw 264.7 cells. However, the introduction of Mg-containing MSs significantly reduced the increase in DCF levels, likely due to the release of H2 gas, which possessed ROS-scavenging properties (Fig. 3D, E). Flow cytometry further revealed the ameliorative effect of GelMa/Mg/DMF MSs on the induction of inflammation by LPS (Figs. 3H and S5C). Notably, inducible nitric oxide synthase (iNOS) expression increased with LPS treatment but decreased with GelMa/Mg/DMF MS treatment (Fig. 3F and J). Conversely, arginase-1 (Arg1) expression decreased following LPS treatment but increased after GelMa/Mg/DMF MS treatment (Fig. 3G and 3J). Phalloidin staining highlighted the characteristic protrusions of M2-type macrophages. Together, these results revealed that GelMa/Mg/DMF MSs exhibited ROS scavenging effects by releasing H2 and promoted macrophage polarization (Fig. 3K). GelMa/Mg MSs have been shown to exert certain modulatory effects on the early inflammatory response, potentially by releasing hydrogen. Accordingly, GelMa/Mg/DMF MSs were introduced with the objective of further enhancing the local anti-inflammatory and macrophage polarization effects, regulating and remodeling the IME, and ultimately modulating bone homeostasis in patients with osteoporosis.
A Live/dead assay of BMSCs and Raw 264.7 cells for 72 h. Scale bars: 200 μm. B Methylthiazolyldiphenyl-tetrazolium bromide (MTT) assay of BMSCs and Raw 264.7 cells (n = 5 independent experiments). C Detection of macrophage polarization levels via flow cytometry. D Quantitative analysis of the expression of dichlorodihydrofluorescein diacetate (DCF) in Raw 264.7 cells was detected by CLSM (n = 5 independent experiments). E DCF expression in Raw 264.7 cells was detected via CLSM. Scale bars: 10 μm. F Detection of DCF in Raw 264.7 cells by flow cytometry. Scale bars: 10 μm. G The expression of inducible nitric oxide synthase (iNOS) in Raw 264.7 cells was detected via CLSM. Scale bars: 10 μm. H Arginase-1 (Arg1) expression in Raw 264.7 cells was detected via CLSM. I Quantitative analysis of the expression of iNOS in Raw 264.7 cells was detected by CLSM (n = 5 independent experiments). J Quantitative analysis of the expression of Arg1 in Raw 264.7 cells was detected by CLSM (n = 5 independent experiments). K The GelMa/Mg/DMF MSs promote macrophage polarization levels and ROS scavenging and thereby orchestrate the IME. (created with BioRender.com) The data are presented as the means ± SD. Statistical significance was calculated by one-way ANOVA with Tukey’s post hoc test. Source data are provided as a Source Data file.
Modulation of the pyroptosis-induced IME enhances osteoblast differentiation
Despite the favorable anti-inflammatory effects of H2 generated by Mg spheres, as demonstrated in previous experiments, H2 did not significantly mitigate osteoblast pyroptosis. TEM showed that rupture of the cell membrane was still observed following GelMa/Mg MS treatment (Fig. 4A, black arrow), leading to the release of inflammatory factors and the activation of downstream inflammatory cascades. We targeted the pore-forming protein N-GSDMD with DMF to prevent cell membrane rupture after treatment with GelMa/Mg/DMF MSs to address this (Fig. 4B). WB analysis revealed elevated GSDMD levels following LPS and ATP treatment. However, GelMa/Mg/DMF MS treatment significantly decreased N-GSDMD, NLRP3, and IL-18 levels (Fig. S6A). CLSM revealed GSDMD protein expression, with high localization at the cell membrane (Dil staining) after LPS/ATP treatment, which was likely inhibited by GelMa/Mg/DMF MSs (Fig. 4C). Semi-quantitative analysis further confirmed that GelMa/Mg/DMF MSs reduced GSDMD expression compared with GelMa/Mg MSs alone (Fig. 4D). Nuclear factor erythroid-related factor 2 (NRF2) is a master transcriptional regulator of many genes involved in the adaptive response to oxidative stress51,52. Increased oxidative stress is also a key factor associated with the aging process, and compelling evidence suggests that NRF2 not only decreases with age, but also reduces secretory phenotypes associated with cellular senescence and aging52,53. This phenomenon has been demonstrated in many models of inflammatory and autoimmune diseases, so it is not surprising that NRF2 has been recognized as a viable drug target for the treatment of a wide range of diseases. NRF2 is the direct target of microsphere-assisted release of DMF, a cysteine-responsive compound that targets specific cysteine residues on the KEAP1 protein in the NRF2 pathway, in particular KEAP1 Cys151. Upon binding to KEAP1, DMF covalently modifies KEAP1, thereby inhibiting the KEAP1-CUL3 E3 ubiquitin ligase, and deubiquitylating NRF26. NRF2 is deubiquitinated and degraded, accumulates in the cell and is transported to the nucleus. In the nucleus, NRF2 dimerizes with small MAF proteins and binds to the antioxidant response element (ARE), which drives the expression of its target genes (e.g., antioxidant enzymes and phase II detoxification enzymes, such as NQO1, GCLC, and HO-1) to enhance cellular antioxidant defenses and maintain cellular redox54. The present study demonstrated that NRF2 is a direct target of microsphere-assisted release of DMF, and WB confirmed that DMF activates the NRF2/HO-1 pathway (Fig. S6 B, C). This finding suggests that anti-inflammatory pathway activation through the NRF2 /HO-1 pathway may be a key pathway in osteoporosis. The modulation of the IME was assessed by analyzing inflammatory factor levels in the cell supernatants. The results demonstrated the suppression of pro-inflammatory cytokines (IL-1β, IL-18, interferon (IFN)-γ, and tumor necrosis factor-alpha (TNF)-α) and an increase in anti-inflammatory mediators (transforming growth factor (TGF)-β and IL-10) (Figs. 4E and S6D), indicating that GelMa/Mg/DMF MSs effectively blocked the pyroptosis of BMSCs, thereby alleviating the IME caused by pyroptosis.
A TEM image of BMSCs after different treatments. Scale bars: 5 μm. B The expression of pyroptosis-related proteins in BMSCs was detected by Western blotting. C The expression of GSDMD in BMSCs was detected via CLSM. Scale bars: 50 μm. D Quantitative analysis of the expression of GSDMD in BMSCs was detected by CLSM (n = 5 independent experiments). E Changes in the levels of inflammatory factors within cell supernatants were detected via ELISA (n = 5 independent experiments). F ALP staining of BMSCs after 7 days and 14 days. Scale bars: 200 μm. G ARS staining of BMSCs after 14 days and 21 days. Scale bars: 200 μm. H The expression of osteogenic differentiation-related proteins in BMSCs was detected by Western blotting. I The GelMa/Mg/DMF MSs promoted osteogenic differentiation by targeting the N-GSDMD to inhibit pyroptosis in BMSCs and thereby orchestrated the IME. (created with BioRender.com) The data are presented as the means ± SD. Statistical significance was calculated by one-way ANOVA with Tukey’s post hoc test. Source data are provided as a Source Data file.
Further investigation of the impact on osteogenic differentiation revealed that the GelMa/Mg/DMF MSs significantly promoted osteoblast differentiation, as evidenced by ALP and ARS staining (Fig. 4F, G). This effect increased over time, likely due to the sustained release of Mg2+, which enhanced osteogenic differentiation and bone remodeling. ALP activity corroborated this phenomenon (Fig. S6E). Additionally, osteogenic-related protein expression was analyzed by WB. The expression level of OPN, which is closely related to bone formation and development, was increased in Mg-containing MSs compared to the control group (Figs. 4H and S6F)41,55. Similarly, Mg-containing MSs increased the protein expression of runt-related transcription factor 2 (Runx2), SP7 (Osx), and osteocalcin (OCN) in BMSCs, key indicators of bone formation and late bone mineralization56,57. It could be inferred that MSs have the potential to accelerate bone formation and mineralization by producing low doses of Mg2+, thereby accelerating osteogenic differentiation. Taken together, GelMa/Mg/DMF MSs demonstrated superior efficacy in inhibiting pyroptosis and regulating the IME. They played a crucial role in both the upstream and downstream processes of inflammation, potentially mitigating the inflammatory milieu associated with osteoporosis (Fig. 4I). Therefore, the GelMa/Mg/DMF MSs promoted osteogenic differentiation by releasing Mg2+, thereby facilitating bone formation and remodeling in inflammatory states.
Regulation of pyroptosis-induced IME facilitates the phosphorylation of ERK in osteoclasts
Clinical data analysis revealed that osteoblasts underwent pyroptosis in patients with osteoporosis. A Transwell co-culture system of BMSCs and BMDMs was used to investigate the role of GelMa/Mg/DMF MSs in this process to further verify the impact of BMSC pyroptosis on BMDMs, (Fig. 5A). TRAP staining revealed that BMSC pyroptosis facilitated the differentiation of BMDMs into osteoclasts (Figs. 5B and S6A). Quantitative analysis confirmed that GelMa/Mg/DMF MSs significantly inhibited osteoclast formation (Fig. 5C). Conversely, the functionality of the co-cultured osteoclasts was validated by examining resorption traces on bovine bone fragments using SEM (Fig. 5D). Quantitative analysis of the resorbed areas on bovine bone slices revealed that pyroptosis promoted BMDM activity, which was effectively blocked by GelMa/Mg/DMF MSs (Fig. 5E). The pH of the co-culture medium was measured, revealing that BMDM activation released inflammatory substances, creating an acidic environment that was partially neutralized by GelMa/Mg/DMF MSs (Fig. 5F). The parietal membrane, a specialized plasma membrane located at the base of the cell, is in direct contact with the bone matrix, and its surface forms a “ruffled border” that is the immediate site where bone resorption occurs. Osteoclasts secrete hydrogen ions (H+) to the bone surface at the ruffled border, thereby lowering the local pH to 4.5-5.0 and facilitating the dissolution of bone mineral, which is typically confined to the bone resorption cavity below the sealing zone. In cases of osteoporotic bone defects, the sealing zone is destabilized by anchoring due to abnormal activation of osteoclasts, leading to hyperfunction and rapid dissolution of the bone matrix. This results in weakening of the sealing zone barrier and loss of local control. The V-type H⁺-ATPase proton pump located on the parietal membrane of hyperactivated osteoclasts is subject to continuous overloading, resulting in the secretion of substantial quantities of H⁺ into the bone surface58,59. Concurrently, H⁺ leakage into the surrounding bone microenvironment occurs, leading to a decline in the pH of the tissue fluid60,61,62. This process, in turn, facilitates the release of Mg2+ from the microspheres. The Mg2+ released by GelMa/Mg/DMF MSs not only stimulated osteoblast activity but also generated alkaline substances that regulated the IME, thereby inhibiting osteoclast activity63,64.
A The GelMa/Mg/DMF MSs coordinate bone homeostasis by inhibiting the pyroptosis of BMSCs, thereby suppressing the activity of BMDMs. (created with BioRender.com) B TRAP staining of BMDMs after 7 days. Scale bars: 50 μm. C Quantification of the number of nuclei per osteoclast (n = 5 independent experiments). D SEM images of BMDMs on the surface of bovine bone slices after 7 days of incubation. Scale bars: 20 μm. E Quantitative analysis of the surface resorbed area of SEM images of bovine bone slices (n = 5 independent experiments). F pH levels of the co-culture medium (n = 5 independent experiments). G The gene ontology (GO) enrichment analysis of differentially expressed genes (DEGs) in BMDMs. H Gene set enrichment analysis of DEGs. I Protein expression in BMDMs was detected via Western blotting analysis. J Osteoclast ring formation and p-ERK expression in BMDMs were detected via CLSM. Scale bars: 100 μm. K GelMa/Mg/DMF MSs inhibit pyroptosis-induced IME promoting phosphorylation of ERK in osteoclasts. (created with BioRender.com) The data are presented as the means ± SD. Statistical significance was calculated by one-way ANOVA with Tukey’s post hoc test. Source data are provided as a Source Data file.
RNA sequencing was performed on co-cultured BMDMs, with the LPS group serving as the control, to further explore the mechanism of BMDM activation. The volcano plot and differential gene heatmap revealed that a greater number of differential genes were generated after GelMa/Mg/DMF MS treatment (Fig. S7B–C). The extracellular regulated protein kinase (ERK)-related pathway was enriched according to the gene ontology (GO) enrichment analysis (Fig. 5G). Further analysis by gene set enrichment analysis (GSEA) confirmed that GelMa/Mg/DMF MSs had an inhibitory effect on the activation of the ERK pathway (Fig. 5H). WB analysis revealed that pyroptotic BMSCs activated ERK phosphorylation in BMDMs, thereby promoting activator protein 1 (AP1) expression, whereas GelMa/Mg/DMF MSs inhibited this process (Figs. 5I and S7D). Once activated by phosphorylation, ERK is a pivotal protein in this pathway that is activated by phosphorylation and translocates from the cytoplasm to the nucleus65. It also mediates the transcriptional activation of Elk-1 and ATF and is involved in a multitude of biological responses, including cell proliferation and differentiation, as well as the maintenance of cell morphology66.
Concurrently, the activation of BMDMs resulted in the formation of multinucleated giant cells, which subsequently constituted a ring of osteoclasts. The CLSM analysis indicated that pyroptotic BMSCs affected BMDMs to form more multinucleated giant cells than the control group. Conversely, GelMa/Mg/DMF MSs inhibited the formation of these cells (Fig. 5J). The diameter of the multinucleated giant cells was significantly increased in the pyroptotic BMSCs groups and was approximately three times larger than that of the control group. This larger diameter was indicative of functional activation. In contrast, no increase in diameter was observed following treatment with GelMa/Mg/DMF MSs (Fig. S7E). The expression of p-ERK in multinucleated giant cells was also identified using CLSM. Consistent with WB results, GelMa/Mg/DMF MSs inhibited the expression of p-ERK in multinucleated giant cells (Fig. S7F).
Initially, the GelMa/Mg/DMF MSs indirectly inhibited BMDM activity by mitigating BMSC pyroptosis. The subsequent direct application of GelMa/Mg/DMF MSs to BMDMs, as observed through TRAP staining, inhibited BMDM function, likely due to the antioxidant properties of H2 generated by the MSs (Fig. S7G–H). Consistent with the above results, the resorbed area of bovine bone fragments was reduced upon treatment with GelMa/Mg/DMF MSs (Fig. S7I–J). Overall, the findings demonstrated that GelMa/Mg/DMF MSs effectively suppressed ERK phosphorylation, thereby promoting AP1 expression within osteoclasts by inhibiting pyroptotic BMSCs and neutralizing inflammatory conditions, and they offer potential therapeutic benefits to overcome the vicious inflammatory cycle in osteoporosis (Fig. 5K).
GelMa/Mg/DMF MSs enhance osteoporosis treatment in vivo
The above in vitro study demonstrated that GelMa/Mg/DMF MSs inhibited osteoclast pyroptosis and remodeled the IME. The potential regenerative effects on tissue repair can be elucidated through the modeling of bone defects in osteoporotic mice39. Eight weeks post-ovariectomy, female C57 mice developed osteoporosis due to estrogen deficiency (Fig. S8A). Cranial defects were created and filled with microsphere scaffolds from different groups (Fig. S8B). Cranial defects reconstruction was observed at two distinct time points (1 and 8 weeks) following the suturing of the defects in a layer-by-layer fashion (Fig. S8C). Microcomputed tomography (micro-CT) analysis revealed that GelMa/Mg/DMF MSs effectively facilitated cranial defect repair, with significant healing observed at 8 weeks (Fig. 6A). The BMD and bone volume-to-total volume (BV/TV) metrics indicated superior osteogenic efficiency in the GelMa/Mg/DMF MS group compared with the controls (Fig. 6B, C). While a substantial body of literature exists on the promotion of osteogenesis by ions such as Ca and Mg, research on the role of ions in the context of osteoporosis and inflammatory disease states is limited63,67. In pathological states such as osteoporosis, the body’s inflammatory indexes are activated, which, in turn, reduces the efficiency of tissue reconstruction. The GelMa/Mg/DMF MS treatment group exhibited delayed osteoporosis progression at the site of cranial bone defects, as evidenced by less trabecular loss and improved distribution. However, considering that osteoporosis is a systemic disease, we further evaluated the impact of GelMa/Mg/DMF MSs on systemic bone mass (Fig. 6D, E). This involved observing microstructural changes in the representative spine and tibia using micro-CT (Fig. S9A, B). The reduction in BMD and the increase in trabecular separation in representative tibiae and spines following the onset of osteoporosis were further quantified using micro-CT analysis (Fig. 6F, G). The utilization of micro-CT technology revealed a disparity in the initial bone density between the GelMa/Mg/DMF MS and GelMa/Mg MS groups. This finding suggests that the early DMF release has the potential to impede pyroptosis, facilitate local bone remodeling of the bone, and enhance osteoblast activity, thereby contributing to an increase in bone density. However, no statistical difference was observed between the two groups in terms of BMD over time. Nonetheless, a significant discrepancy was detected with respect to bone volume fraction, suggesting the possibility that GelMa/Mg/DMF MSs could augment bone volume through the induction of rebalancing bone homeostasis. A quantitative analysis of bone trabeculae further confirmed that GelMa/Mg/DMF MSs increased the number of trabeculae and thus promoted the restructuring of bone homeostasis (Fig. 6H, I).
A Coronal and three-dimensional reconstructed images eight weeks after surgery for cranial defects viewed by micro-CT. Quantitative results of BMD B and bone volume-to-total volume (BV/TV) C in the cranial defect of C57 mice determined by micro-CT (n = 5 samples). D Cross-sectional image of the tibial observed via micro-CT. E Cross-sectional image of a representative spine viewed via micro-CT. Quantitative results of BMD in the tibial F and spine G of C57 mice determined by micro-CT (n = 5 samples). Quantitative results of the number of trabeculae (Tb.N) H and the degree of separation of trabeculae (Tb.Sp) I in the cranial defect of C57 mice determined by micro-CT (n = 5 samples). J Fluorescence microscope observation of a typical image of calcein and alizarin red staining. Scale bars: 200 μm. K Quantification of the mineralization rate (MAR) (n = 5 samples). L Compression experimental diagram. (created with BioRender.com) M Stress-strain curve of the femur. N Quantitative analysis of the modulus of elasticity of the femur of C57 mice (n = 5 samples). The data are presented as the means ± SD. Statistical significance was calculated by one-way ANOVA with Tukey’s post hoc test. Source data are provided as a Source Data file.
However, treatment with GelMa/Mg/DMF MSs mitigated the bone loss associated with disrupted estrogen metabolism and increased the whole-body bone mass and trabecular number while reducing the degree of trabecular separation (Fig. S9C, D). The rationale behind this phenomenon might be that the GelMa/Mg/DMF MSs in the injured area not only generated H2 locally to exert anti-inflammatory effects but also released Mg2+ and DMF into the blood circulation, thereby inhibiting pyroptosis and exerting anti-inflammatory effects at a systemic level. This process ultimately reversed the inflammatory environment induced by osteoporosis and further promoted systemic bone remodeling. Calcium AM and alizarin red double-labeling intraperitoneal injection before euthanization allowed for the measurement of the mineralization rate (MAR), which revealed that GelMa/Mg/DMF MSs accelerated tissue repair and reconstruction (Fig. 6J, K). The structural changes that occur in bone tissue as a result of osteoporosis led to a reduction in bone strength and an increased risk of fracture (Fig. 6L). Therefore, mechanical testing indicated that GelMa/Mg/DMF MSs enhanced the femur’s elastic modulus, aiding in the restoration of the mechanical properties of bone tissue by regulating bone homeostasis (Fig. 6M, N).
The histological analysis of cranial defects by H&E staining revealed reduced local inflammation and a more compact bone structure in the GelMa/Mg/DMF MS group at 1 week and 8 weeks post-surgery (Figs. 7A and S10A). The cranial breaks in the different groups exhibited substantial differences, which were a manifestation of osteoporosis. This was attributed to the metabolic estrogen imbalance. However, in the GelMa/Mg/DMF MS group, the structure of the fractured skull was more compact, and the gap was smaller, indicating osteoclast inhibition. The collagen fibers in the mesh of the bone defect site were identified using Masson’s staining. The GelMa/Mg/DMF MS group exhibited enhanced bridging of the bone defect compared with the control group, accompanied by a distinct reddish hue, indicative of heightened maturity of the regenerated bone (Fig. 7B). Additionally, TRAP staining revealed activated osteoclasts in the control group’s cranial breaks, whereas the GelMa/Mg/DMF MS group exhibited a notable reduction in osteoclasts (Figs. 7C and S10B, black arrows). The osteoblasts were further labeled by immunohistochemical staining at the site of the injury (Figs. 7D and S10C). The quantification of OCN-positive cells revealed that GelMa/Mg/DMF MSs facilitated osteogenic activity via Mg2+ release (Fig. 7E). Moreover, immunohistochemical staining of GSDMD revealed that GelMa/Mg/DMF MSs significantly inhibited pyroptosis in cranial defects compared with the other groups (Fig. 7F, G). As demonstrated by the findings of the preceding experiments, the osteogenic differentiation capability of osteoblasts decreased following the onset of pyroptosis. The investigation further revealed that GelMa/Mg/DMF MSs could impede the occurrence of local pyroptosis at the site of injury, thereby prompting osteoblasts to act as mediators in the restoration of the defective area.
Coronal images of the cranial defect after H&E staining A and Masson staining B 8 weeks after surgery. Scale bars: 100 μm. C Coronal images of the cranial defect after TRAP staining 8 weeks after surgery for the cranial defect. Scale bars: 100 μm and 20 μm. Coronal images of the cranial defect after immunohistochemistry for OCN D and GSDMD F 8 weeks after surgery for the cranial defect. Scale bars: 100 μm and 20 μm. Quantitative analysis of the percentage area of immunohistochemical staining for OCN E and GSDMD G 8 weeks after surgery for the cranial defect. (n = 5 samples). H Detection of inflammatory cytokines in the C57 serum at one week post-operation (n = 5 samples). I Images of cranial defects after immunofluorescence staining for Arg1+ (red) and iNOS+ (green). Scale bars: 100 μm and 20 μm. J Images of scalp tissue surrounding cranial defects after immunofluorescence staining for Arg1+ (red) and iNOS+ (green). Scale bars: 500 μm. K GelMa/Mg/DMF MSs inhibits pyroptosis and remodels the IME in osteoporosis. (created with BioRender.com) The data are presented as the means ± SD (n = 5 samples). Statistical significance was calculated by one-way ANOVA with Tukey’s post hoc test. Source data are provided as a Source Data file.
Moreover, ELISA analysis of the serum samples revealed reduced the levels of inflammatory cytokines (IL-1β, IL-18, IFN-γ, and TNF-α) and increased the levels of anti-inflammatory cytokines (IL-10 and TGF-β) in the GelMa/Mg/DMF MS group than the control group, corroborating the cellular findings (Figs. 7H and S10D). Immunofluorescence staining indicated that GelMa/Mg/DMF MSs modulated macrophage polarization toward the anti-inflammatory M2 phenotype. All these results suggest that the GelMa/Mg/DMF MSs regulated the local IME, thereby facilitating the repair and reconstruction of bone tissue in an inflammatory setting. Given the pivotal role of macrophages in the IME, we further examined their distribution at the injury site using immunofluorescence staining. This involved iNOS labeling for M1-type macrophages and Arg1 labeling for M2-type macrophages (Fig. 7I). Consistent with the cellular experiments results, GelMa/Mg/DMF MSs promoted macrophage polarization toward the M2 phenotype (Fig. S10E). One week post-surgery, M1-type macrophages were initially predominant at the injury site. However, the GelMa/Mg/DMF MSs modulated the IME, suppressed M1-type macrophages and promoted M2-type macrophage polarization.
Micro-CT analysis suggested that GelMa/Mg/DMF MSs exerted a beneficial effect on osteoporotic bone tissue throughout the body. Further examination of tibial tissue sections from C57 mice revealed a slight increase in the number and density of bone trabeculae in the GelMa/Mg/DMF MS group compared with those in the control groups (Fig. S11A). TRAP staining confirmed a decrease in the number of osteoclasts in the GelMa/Mg/DMF MS group (Fig. S11B). Moreover, immunohistochemical staining revealed a significant increase in OCN-positive cells, indicating enhanced osteogenic activity (Fig. S11C). The histological examination of localized scalp tissue near the cranial injury revealed the infiltration of inflammatory cells prompted by the injury (Fig. S11D). Immunofluorescence staining revealed a predominance of M1-type macrophages post-injury. However, GelMa/Mg/DMF MSs modulated this response, promoting the polarization of macrophages toward the anti-inflammatory M2 phenotype (Figs. 7J and S11E).
Finally, the in vivo biosafety of the GelMa/Mg/DMF MSs was evaluated. H&E staining of major organs (including the heart, liver, spleen, lungs, and kidneys) was performed to further assess tissue toxicity. The results demonstrated that GelMa/Mg/DMF MSs were highly histocompatible (Fig. S12A). Furthermore, the levels of alanine aminotransferase (ALT), aspartate aminotransferase (AST), ALP, total bilirubin (TBIL), creatinine (CRE), and blood urea nitrogen (BUN) were comparable between the control group and the GelMa/Mg/DMF MS group (Fig. S12B), indicating that the degradation of magnesium-ionized microspheres did not result in any observable systemic toxicity.
Overall, the incorporation of gel microspheres at the defect site modulated macrophage polarization and attenuated local inflammation at an early stage. Conversely, the inhibition of osteoclast activation by preventing osteoblast pyroptosis and remodeling of the local IME resulted in long-term tissue regeneration and repair (Fig. 7K). Therefore, GelMa/Mg/DMF MSs facilitated tissue repair and reconstruction in the context of osteoporosis, thus offering a promising approach to address bone loss.
Discussion
The treatment of osteoporosis and related bone defects has long presented a considerable challenge to healthcare professionals. The present study utilized a comprehensive analytical approach, encompassing the examination of samples obtained from individuals diagnosed with osteoporosis. The analysis yielded compelling evidence that the inflammatory microenvironment associated with pyroptosis exerts a pivotal role in disrupting bone homeostasis. In light of the findings, a magnesium-gelatin composite microsphere scaffold was developed, leveraging pyroptosis blockade and hydrogen-mediated inflammation regulation for the treatment of osteoporosis. The implantation of the porous microsphere scaffold was undertaken in order to achieve the sustained release of hydrogen gas, Mg2+ and DMF. The DMF operates by inhibiting the NLRP3/CASP1/GSDMD signaling pathway to prevent osteoblast pyroptosis. In combination with the antioxidant effects of hydrogen, this effectively remodels the inflammatory microenvironment and creates favorable conditions for the restoration of bone homeostasis. The results of this study demonstrated that GelMa/Mg/DMF MSs effectively reversed the inflammatory microenvironment both in vivo and in vitro, resulting in significant tissue repair.
Our study has some shortcomings, the first being that our retrospective approach to collecting serum from controls and patients with osteoporosis precluded the establishment of a robust causal relationship, a necessity that can only be addressed through prospective studies. Moreover, samples obtained from a single center are susceptible to regional limitations. Second, patients diagnosed with osteoporosis tend to have low levels of vitamin D. Thus, it was not possible to avoid the potential bias of vitamin D on markers of inflammation in patients’ serum, and further investigation with a greater number of clinical samples is needed to establish a more robust relationship between osteoporosis and complications.
In summary, the pyroptosis-blocking complex microsphere scaffold, GelMa/Mg/DMF MS, was successfully constructed for treating osteoporosis. Preliminary results indicated that the microspheres respond to changes in pH and IME through their porous structure, thereby achieving sustained and stable release of Mg2+ and DMF. The internal magnesium spheres generated H2 gas for effective anti-inflammatory effects, which synergized with DMF to promote macrophage polarization toward an anti-inflammatory M2 phenotype. GelMa/Mg/DMF MSs targeted GSDMD and impeded the pore-forming activity of N-GSDMD, thereby inhibiting osteoblast pyroptosis and diminishing the activation of the p-ERK/AP1 pathway in osteoclasts triggered by pyroptotic osteoblasts. GelMa/Mg/DMF MSs appears to disrupt the pyroptosis-induced IME, effectively breaking the vicious cycle in osteoporosis (Fig. 8).
(created with BioRender.com).
Methods
Analysis of clinical samples
Patient specimens were collected in accordance with the ethical standards approved by the Medical Ethics Committee of The First Affiliated Hospital of Soochow University (Jiangsu, China, Ethic approval number: 2024499). Participants gave a written informed consent for the experiment. Bone density was determined by DXA method through X-ray examination. Patient serum was collected and analyzed via ELISA to determine the presence of a range of cytokines, including IL-1β, IL-2, IL-4, IL-5, IL-6, IL-8, IL-10, IL-12P70, IL-17A, IL-18, IFN-α2, IFN-γ, and TNF-α (Dakewe).
Synthesis of the GelMa/Mg/DMF microsphere-scaffold (MSs)
The preformed gelatin was dissolved in water at 60 °C. Lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP, Sigma, USA) was added to the dissolved gelatin, which was then ultrasonically dispersed and promptly combined with magnesium spheres and DMF (T0492, TargetMol, USA). This mixture was introduced in a controlled fashion to the pre-formed mineral oil, which was agitated at a rate of 200 rpm. Following a two-minute stirring and mixing period, the product was subjected to ultraviolet (UV) curing with a handheld UV lamp for an additional three minutes. Centrifugation at 6800 x g and washing with water were then performed three times to obtain the resulting GelMa/Mg/DMF MSs.
Characterization
The morphology of GelMa/Mg/DMF MS was characterized by SEM (G500, Zeiss). The crystal structure of GelMa/Mg/DMF MS was measured by XRD (Panalytical Empyrean). The absolute concentration of Mg ions was measured by inductively coupled plasma-optical emission spectroscopy (ICP-OES, Avio 200). The absorption spectra were obtained by a UV-vis-NIR spectrophotometer (Thermo50 UV-vis-NIR spectrophotometer, Thermo Scientific). The detection of hydrogen release from a material is achieved through the use of MB. Detect the removal of ROS using oxTMB and ABTS+ as analytical reagents.
Cell culture
BMSCs and BMDMs were obtained by flushing the femur and tibia of 6-weeks-old C57 mice. C57 mice were obtained from Changzhou Cavens Model Animal Co., Ltd. RAW 264.7 was obtained from the Cell Bank, Shanghai Institutes for Biological Sciences, Chinese Academy of Sciences. BMSCs and BMDMs were cultured in α-MEM supplemented with 10% fetal bovine serum (FBS, BI, USA) and 100 mg/mL penicillin-streptomycin (Sigma, USA)68,69. The BMSCs and BMDMs were co-cultured via a transwell system. The BMSCs were seeded in the upper chamber, and the BMDMs were grown in the lower chamber. The incorporation of Rankl (50 ng/ml, O35235, lifetein, USA) and M-CSF (30 ng/ml, C756, Novoprotein, Shanghai, China) into the BMDMs medium enhanced the induction process70,71,72. The second and third generations of BMSCs were incubated at 37 °C in saturated humidity and 5% CO₂ to allow for optimal growth. To assess the response of BMSCs and BMDMs, all microsphere samples were sterilized overnight in UV light.
For live/dead dual-staining in vitro, BMSCs and Raw 264.7 cells were incubated with GelMa/Mg/DMF (10 μg/mL) for 24 h, followed by staining with calcein AM and propidium iodide for 30 min according to the recommended protocol (K2247, APExBIO, Houston, USA). All the images were acquired via confocal laser scanning microscopy (CLSM, LSM800, Zeiss).
For in vitro cell viability, the MTT assay (M8180, Beijing Solarbio Science & Technology Co., Ltd.) was performed to determine the relative cell viability after different treatments.
Osteogenesis-related experiments
For osteogenic differentiation, BMSCs were cultured in α-MEM and subsequently transferred to standard osteogenic induction medium (OIM). The composition of OIM was as follows: 0.05 mM vitamin C (Sigma, USA), 10 mM β-glycerophosphate (Sigma, USA), and 1 × 108 M dexamethasone (Sigma, USA)67,73. The medium was replaced every two days. Once the cell density reached approximately 70-80%, the α-MEM was replaced with the OIM to induce the differentiation of the BMSCs for a period of 7/14/21 days. Subsequently, the differentiation of BMSCs was evaluated through ALP staining, ALP activity measurement, and ARS staining39,74. Following a 7/14/21-day culture period, an ALP assay kit (Beyotime, China) was employed to quantify ALP activity. The results were normalized to total cellular protein, with a medium lacking the specified material serving as a control. Following a period of 7/14/21 days, the cell monolayer was washed twice with PBS and treated with 10% neutrophil formalin for 15 mins. Following two rinses in PBS, the samples were subjected to staining in accordance with the protocols of the ALP and ARS assay kits and observed under a light microscope (Olympus, IX71, Japan).
To verify the oteogenesis-related protein expression in vitro, BMSCs were incubated with GelMa/Mg/DMF (10 μg/mL) for different time and then collected and suspended in 4 °C cell lysis buffer. Proteins were quantified using the BCA Protein Assay Kit (Beijing Bose Biotechnology Co., Ltd., AKPR017). The antibodies were as follows: OPN antibody (A21084, Abclonal, Dilution: 1:1000), OCN antibody (A20800, Abclonal, Dilution: 1:1000), Runx2 antibody (82636-2-RR, ProteinTech, Dilution: 1:1000), SP7 antibody (ab209484, Abcam, Dilution: 1:1000).
Osteoclast-related experiments
Following a seven-day incubation period, the activity of the co-cultured BMDMs was detected via Trap staining (Beyotime, China), in accordance with the protocol outlined in the assay kit. The area of absorption on bovine bone slices was observed through a seven-day co-culture of bovine bone slices and BMDMs via SEM and quantified via ImageJ75. The size of the osteoclast rings and the p-ERK expression were observed via immunofluorescence. The BMDMs were initially blocked with 5% FBS and subsequently incubated with the primary antibody (ProteinTech, China) for a period of three hours. Following this, the cells were washed three times with PBS, after which the secondary antibody (Alexa Fluor 488, Abcam, USA) was applied for one hour. The size of the osteoclast ring diameters was then measured via ImageJ.
To verify the osteoclast-related protein expression in vitro, the co-cultured BMDMs were collected and suspended in 4 °C cell lysis buffer for western blotting according to the recommended protocol (SurePAGE, M00654, GenScript). The antibodies were as follows: ERK1/2 antibody (R013501, Epizyme, Shanghai, China, Dilution: 1:1000), Phospho-ERK1/2 (Tyr222/Tyr205) antibody (M010247, Epizyme, Shanghai, China, Dilution: 1:1000), AP1 antibody (24909-1-AP, ProteinTech, Dilution: 1:1000).
A total of 1 μg of RNA from BMDMs was employed for the subsequent library preparation (AC0103, Shandong Sparkjade Biotechnology Co., Ltd.). Poly (A) mRNA isolation was conducted using Oligo (dT) beads. The fragmentation of the mRNA was conducted using divalent cations and a high temperature. Priming was conducted using random primers. First-strand and second-strand cDNA were synthesized. The purified double-stranded cDNA was subsequently subjected to repair at both ends and addition of an adenine residue at the 3’ end, followed by a T-A ligation to add adapters to both ends. Subsequently, the adaptor-ligated DNA was subjected to size selection using DNA Clean Beads. Subsequently, each sample was subjected to polymerase chain reaction (PCR) amplification using P5 and P7 primers, and the resulting PCR products were subsequently validated. The libraries with distinct indexes were multiplexed and sequenced on an Illumina HiSeq, Illumina Novaseq, or MGI2000 instrument using a 2×150 paired-end configuration, in accordance with the manufacturer’s instructions.
The differential expression analysis was conducted using the DESeq2 Bioconductor package, which is based on a negative binomial distribution. The estimates of dispersion and logarithmic fold changes incorporate data-driven prior distributions. The P-value of genes was set at ≤0.05 to identify those that were differentially expressed. GOSeq (v1.34.1) was employed to identify Gene Ontology terms that annotate a list of enriched genes with a Padj less than or equal to 0.05. Furthermore, the R package topGO was employed for the construction of a directed acyclic graph (DAG). GSEA analysis was performed with GOBP_NEGATIVE_REGULATION_OF_ERK1_AND_ERK2_CASCADE using the cluster Profiler package.
Cellular pyroptosis related experiments
To verify the relative protein expression associated with pyroptosis in vitro, BMSCs were incubated with GelMa/Mg/DMF (10 μg/mL) for different time and then collected and suspended in 4 °C cell lysis buffer for western blotting according to the recommended protocol (SurePAGE, M00654, GenScript). The antibodies were as follows: GSDMD antibody (ab219800, Abcam, Dilution: 1:1000), Caspase-1 antibody (ab179515, Abcam, Dilution: 1:1000), NLRP3 antibody (ab263899, Abcam, Dilution: 1:1000), NRF2 antibody (AF0639, Affinity, Dilution: 1:1000) and HO-1 antibody (R014764, Epizyme, Shanghai, China, Dilution: 1:1000).
To verify the microstructure of cellular pyroptosis in vitro, BMSCs were incubated with GelMa/Mg/DMF (50 μg/mL) for 12 h, followed by incubation with LPS (1 μg/ml, L4391, Sigma) for 4 hours and ATP (5 μM, A2383, Sigma) for 6 h. The cells were then collected and fixed in electron microscopy fixative, and transmission electron microscopy (TEM, tecnai F20) was performed according to the recommended protocol.
To test GSDMD and NLRP3 expression, BMSCs cells were incubated with GelMa/Mg/DMF for 12 h, followed by incubation with LPS (1 μg/ml, L4391, Sigma) for 4 h and ATP (5 μM, A2383, Sigma) for 6 h. The collected cells were first blocked with 5% FBS and then incubated with the primary antibody (Abcam, USA) for 3 h. After washing with PBS three times, the cells were then incubated with secondary antibody (Alexa Fluor 488, Abcam, USA) for 1 h and incubated with Dil (C1036, Beyotime, China) according to the recommended protocol. All the images were acquired via CLSM. The other groups included the control, GelMa, and GelMa/Mg groups.
Inflammation-related experiments
Raw 264.7 cells were incubated with GelMa/Mg/DMF (50 μg/mL) for 48 h, followed by incubation with LPS (1 μg/ml) for 24 hours. Following treatment, macrophages were gently detached, washed with ice-cold PBS, and resuspended for staining. Cells were first incubated with a viability dye (dead cell staining buffer) to exclude non-viable cells, followed by Fcγ receptor blockade (BD Pharmingen, USA; 10 min) to minimize nonspecific antibody binding. Subsequently, cells were stained with FITC-conjugated anti-mouse CD11b and Percp-cy5.5-conjugated F4/80 antibodies (BioLegend, USA; 30 min, 4 °C) to identify macrophages. For intracellular marker analysis, cells were fixed and permeabilized using fixation/permeabilization buffer (eBioscience™, room temperature, 30 min), washed twice with permeabilization buffer (eBioscience™, 5 min each), and then stained with antibodies against CD86-PE (M1 marker) and CD206-APC (M2 marker) (30 min, room temperature).
Raw 264.7 cells that underwent treatment were subjected to a 10 μM concentration of DCF (Beyotime, China) staining, which is a fluorescent probe used to detect reactive oxygen species (ROS). The staining was performed for 30 minutes in an environment that was free from light. Subsequently, the cells were observed via CLSM and CYTEK NORTHERN LIGHTS (NL-CLC V16-B14-R8).
To test iNOS and Arg1 expression, Raw 264.7 cells were incubated with GelMa/Mg/DMF for 12 h. The collected cells were first blocked with 5% FBS (JYK-FBS-303, INNER MONGOLIA JIN YUAN KANG BIOTECHNOLOGY CO., LTD) and then incubated with primary antibody (16001-1-AP, 22226-1-AP, ProteinTech, China) for 3 h. After washing with PBS three times, the cells were then incubated with secondary antibody (Alexa Fluor 488, Abcam, USA) for 1 h and incubated with ActinRed 555 (R37112, Thermo Fisher) according to the recommended protocol. All the images were acquired via CLSM. The other groups included the control, GelMa, and GelMa/Mg groups.
The cell supernatants were collected and analyzed by ELISA for the presence of a range of cytokines, including IL-1β, IL-18, IFN-γ, TNF-α, transforming growth factor-β (TGF-β) and IL-10 (AMOY LUNCHANGSHUO BIOTECH, CO., LTD).
Construction of the OVX and cranial defect model
The experiment was conducted using six-week-old female C57 mice, which were procured from Changzhou Cavens Model Animal Co. Ltd. and subsequently housed under pathogen-free conditions. The mice were maintained under a 14-hour light/10-hour dark cycle, with temperatures ranging from 18 to 23 °C and humidity levels maintained between 40 and 60%. The mice were housed in microisolator cages to ensure the containment of pathogens. The experiment protocol was approved by the Animal Committee of Medical Ethics Committee of Soochow University (Jiangsu, China, Ethic approval number: 202310A0867). All of the reagents used in the experiment were non-toxic and non-radioactive. The animal anesthetic used in this experiment was isoflurane, administered via inhalation. General analgesia (buprenorphine, mundipharma, China) was administered subcutaneously immediately before surgery at a dose of 0.05 mg per kg. Local anesthesia was also applied subcutaneously under the scalp. Eye gel was applied to prevent the eyes from drying out during surgery. After the experiment, the mice were sacrificed using CO2 gas.
For the OVX model, following anesthesia in 8-week C57 mice, a dorsal double incision was made to expose both ovaries successively, the ovarian arteries were ligated and both ovaries were resected. The incisions were closed layer by layer76,77,78.
Eight weeks following the ovariectomy, the C57 mice were randomly divided into four groups (n = 5 per group) and treated as follows: (1) control, PBS; (2) filled with GelMa microspheres; (3) filled with GelMa/Mg microspheres; and (4) filled with GelMa/Mg/DMF microspheres. The C57 mice were anaesthetized, and the skull was exposed while the mice were in the prone position. A two-millimeter round bone defect was created in the middle of the skull on each side using a ring drill. The defects were then filled with microspheres from each group and the incision was closed layer by layer after hemostasis.
After the mouse model of cranial defects was established, alizarin red S (30 mg/kg, Maclean’s, China) and calcineurin (20 mg/kg, Maclean’s, China) were injected intraperitoneally 1 and 2 weeks before sampling, respectively67. Bone density was determined by DXA method through X-ray examination (iNSiGHT VET DXA).
At 1 and 8 weeks post-surgery, the mice were sacrificed by inhaled CO2 at the end of treatment, and the skull, tibia and spine were removed with great care for subsequent experiments. Additionally, the scalp tissue in close proximity to the skull was excised, and the heart, liver, spleen, lungs, and kidneys were removed for biosafety evaluation. Serum was collected and analyzed for biochemical analysis and for the presence of a range of cytokines, including IL-1β, IL-18, IFN-γ, TNF-α, TGF-β and IL-10.
Morphological analysis
For micro-CT analysis, the cranial tibia and spine of mice were fixed in 10% formalin for a period of 24 h. The reconstruction of bone tissue was evaluated using micro-CT (SkyScan1174, Bruker, Belgium; voltage 50 kV, current 800 μA, pitch 2, filter 0.11 mm Cu)35. The postoperative bone formation was evaluated through the measurement of BMD, BV/TV, the number of trabeculae (Tb.N), and the degree of separation of trabeculae (Tb.Sp).
Mechanical performance assay
The hardness of uncalcified femurs was measured in compression mode with a universal material testing machine (24TM-50, Instron, USA). The stress-strain curve was obtained by compressing the mouse femur.
Histological analysis
For immunohistochemical staining, the skulls and tibias of the mice euthanized at 8 weeks were fixed in 4% paraformaldehyde for at least 48 h and then soaked in 10% ethylene diamine tetraacetic acid (EDTA) for 14 days79. The tissue slices were prepared via paraffin embedding of decalcified tissue and stained with H&E, V&G, Masson, and TRAP to observe the effects on bone repair and the mitigation of systemic osteoporosis. Following a one-week decalcification period, the bone tissue was subjected to immunohistochemical staining for GSDMD, OPN, and OCN. Furthermore, immunofluorescence staining (iNOS+ and Arg1 + ) was also performed on the skull and adjacent scalp tissue. Tibia samples were dehydrated and embedded in paraffin. After embedding in methyl (China Biotechnology Co., Ltd.) until solidification, 300 μm thick sections were allocated for hard tissue staining. The rate of calcium salt deposition was calculated by determining the average width/interval days of the fluorescent double-labeled bands to derive the MAR.
Statistics & Reproducibility
All quantitative experiments were performed in triplicate unless otherwise indicated. All the statistical analysis was performed by GraphPad Prism 10.0. The data are presented as the mean ± standard deviation (SD). All experiments were repeated once with similar results unless otherwise stated in the Figure or Supplementary Figure legends. Significant differences between two groups were determined via two-tailed unpaired t-test. One-way analysis of variance (ANOVA) was employed for the significant differences among multiple comparisons. The sample size was selected with the objective of detecting a minimum effect size of 1.5, with a power of at least 80% and a type I error rate of 0.05 for each comparison. Statistically speaking, a P value less than 0.05 was deemed to be statistically significant.
Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.
Data availability
The RNA-seq data generated in this study have been deposited in the NCBI (National Center for Biotechnology Information) database under accession code PRJNA 1295853. GSEA analysis was performed under Mouse Gene Set (https://www.gsea-msigdb.org/gsea/msigdb/mouse/geneset/GOBP_NEGATIVE_REGULATION_OF_ERK1_AND_ERK2_CASCADE). All data supporting the research results described in this article are available in the main text of the article, the supplementary information file, and the source data file. Source data are provided with this paper.
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Acknowledgements
This article was partially supported by the National Research Programs of China (2022YFB3804600, granted to Liang Cheng), National Key Research and Development Program of China (2022YFC2009705, granted to Jun Zou), National Natural Science Foundation of China (52472288, granted to Liang Cheng), (82172506, granted to Jun Zou), (82030068, granted to Huilin Yang), (52203362, granted to Fei Gong), Collaborative Innovation Center of Suzhou Nano Science and Technology (Granted to Liang Cheng), the 111 Project, and Joint International Research Laboratory of Carbon-Based Functional Materials and Devices, Suzhou Key Laboratory of Nanotechnology and Biomedicine, and Key Laboratory of Structural Deformities in Children of Suzhou (SZS2022018, granted to Liang Cheng), the Priority Academic Program Development of Jiangsu Higher Education Institutions (PAPD, granted to Huilin Yang). Jiangsu Funding Program for Excellent Postdoctoral Talent (2024ZB491, granted to Shunyi Lu), China Postdoctoral Science Foundation (2024M752333, granted to Shunyi Lu). AI-assisted technology was used to refine language and ensure clarity in the preparation of this manuscript. All content was reviewed and approved by the authors.
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H. Yang, L. Cheng, J. Zou, Z.L. Zhang and J. Ge conceived and designed the study, analyzed the data, and provided funding. S. Lu, Z. Song and F. Gong characterized the GelMa/Mg/DMF MSs both in vitro and in vivo and performed the data analysis. S. Lu, P. Yang, G. Hou, Z.M. Zhang and Y. Yang carried out the cytological experiments. S. Lu, J. Cao, P. Yang, Z. Han, Y. He and Y. Teng established the in vivo OVX model, acquired medical imaging, and collected the samples. All authors wrote and approved the publication of the manuscript.
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Lu, S., Cao, J., Song, Z. et al. Pyroptosis-responsive microspheres modulate the inflammatory microenvironment to retard osteoporosis in female mice. Nat Commun 16, 8127 (2025). https://doi.org/10.1038/s41467-025-63456-5
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DOI: https://doi.org/10.1038/s41467-025-63456-5
