Mechanical rejuvenation of senescent stem cells and aged bone via chromatin remodeling

Domain: physics
Published: 2026-01-13
Editorial outcome: minor_revision

Abstract
Bone aging compromises skeletal integrity and increases vulnerability to osteoporosis and other age-related disorders, underscoring the need for new therapeutic strategies. Although pharmacological and genetic approaches have been widely explored, how cellular mechanical remodeling contributes to bone aging remains unclear. Here, we find that senescent bone marrow stem cells show markedly reduced intracellular force and impaired mechanical behavior. Moderate mechanical stimulation in cell culture and in mice restores cellular force, increases chromatin accessibility at the FOXO1 locus, activates its expression, and reverses cellular senescence and bone aging. These mechanical interventions also improve physical performance in aged female mice and show a tendency to reduce systemic inflammation, whereas excessive force induces chromatin overextension and DNA damage, indicating the necessity of precise force control. In this work, we show that optimized mechanical stimulation provides a simple and effective strategy to counteract age-related bone deterioration and systemic inflammation, offering potential for clinical translation.

Introduction
Age-related bone loss, characterized by an imbalance between bone formation and resorption1,2, is a complex pathological process that leads to osteoporosis and an increased risk of fractures3–6. This disruption in bone homeostasis is exacerbated by cellular senescence, particularly in bone marrow mesenchymal stem/stromal cells (BMSCs), which exhibit diminished osteogenic potential and self-renewal capacity with aging7,8. As a result, bone remodeling is impaired, contributing to skeletal dysfunction and accelerating the aging process9–11. Despite growing interest in delaying bone aging, understanding the mechanisms underlying BMSC senescence remains critical for developing effective therapeutic strategies to counteract aging and its associated skeletal diseases.

Senescent BMSCs exhibit broad transcriptional reprogramming, particularly in genes governing proliferation, differentiation, and extracellular matrix organization12,13. These transcriptional alterations are tightly regulated by diverse biochemical signaling pathways, including metabolic14, transcriptional15, and epigenetic processes16. Dysregulated metabolic signaling, such as impaired mitochondrial function or altered NAD⁺/SIRT axis17,18, perturbs redox balance and energy homeostasis, thereby activating stress-responsive pathways that promote senescence. Concurrently, transcriptional regulators such as p53, FOXO, and NF-κB orchestrate the expression of senescence-associated genes, linking cellular stress to irreversible growth arrest and the senescence-associated secretory phenotype (SASP)15,19. At the chromatin level, epigenetic modifications—including histone acetylation, methylation, and DNA methylation changes—further consolidate the senescent state by repressing proliferation-related genes and altering chromatin accessibility16,20,21. Together, these biochemical processes converge to establish a stable and self-reinforcing senescence program. While these factors are well-documented, identifying straightforward, translatable interventions to reverse senescence presents a significant challenge.

In addition to these biochemical pathways, emerging evidence highlights a critical role for mechanical signaling in modulating transcriptional and epigenetic processes22. The intracellular traction forces, generated by non-muscle myosin II and counterbalanced by cell adhesive environment, can be directly transmitted into the cell nucleus to modulate chromatin structure and gene expression through the linker of nucleoskeleton and cytoskeleton (LINC) complex and lamin skeleton23. Furthermore, various aged cell types, including human dermal fibroblasts, display a marked reduction in chromatin accessibility, leading to impaired transcriptional activity and disrupted gene expression24–26. Given that cellular traction force regulates chromatin dynamics and transcriptional activity, it is plausible that it contributes to chromatin condensation and the associated loss of gene expression during cellular senescence. While it is known that cellular mechanotransduction can affect various cell behaviors27, such as cell shape28, differentiation29, and migration30, the precise role of mechanical signals in BMSC senescence remains poorly understood.

BMSCs are embedded in a dynamic microenvironment where they continuously perceive and respond to mechanical cues, affecting cellular traction force and directing cell behavior31. Aging significantly alters the physical properties of the bone microenvironment32,33, disrupting the mechanical signals transmitted to cells. These age-related changes suggest that senescent BMSCs may experience profound shifts in cellular mechanics and mechanotransduction. Exploring the interplay between mechanotransduction and cellular senescence is critical for developing novel strategies to intervene in the aging process and mitigate age-related skeletal disorders.

In this work, we show that aging reduces intracellular traction forces in BMSCs and aged bone tissue, a deficiency that can be reversed through appropriate mechanical stimulation to restore the cell mechanics. Mechanistically, the restoration of cellular traction force enhances chromatin accessibility, leading to the activation of FOXO1 expression. Importantly, FOXO1 knockdown abolished the mechanically rejuvenating effects, underscoring its critical role in mediating cellular responses to mechanical forces. Beyond bone recovery, mechanical interventions improved locomotor activity, alleviated physical frailty, and reduced systemic inflammation in aged mice. These findings highlight both local and systemic benefits of mechanical stimulation, offering a straightforward approach with significant translational potential for combating age-related tissue decline.

Results
Decreased cellular mechanics and mechanotransduction in aged BMSCs
To elucidate the relationship between cellular senescence and mechanotransduction, we isolated human bone marrow stromal cells (BMSCs) from 21 elderly donors (50–80 years) and 17 young donors (18–35 years) for in vitro analysis. The isolated BMSCs were cultured and characterized by flow cytometry, confirming typical MSC surface markers. Compared with young BMSCs, those from elderly donors displayed markedly reduced proliferative and osteogenic potential, indicating age-associated functional decline. In parallel, aged BMSCs exhibited increased p21⁺ and γH2A.X⁺ cells, enhanced SA-β-gal activity34, and upregulated expression of SASP factors (Supplementary Figs. 1–5).

To further delineate the alterations in cellular mechanics associated with aging, we applied a combination of traction force microscopy (TFM), Förster resonance energy transfer (FRET)-based actin tension probes35, and nanoindentation to quantify the mechanical integrity of BMSCs. In TFM assays, cells were cultured on deformable polyacrylamide gels embedded with fluorescent beads, allowing the traction stress generated by cell contraction to be calculated from bead displacement fields. Aged BMSCs produced substantially weaker traction forces than young cells (122 vs. 227 Pa; Fig. 1a), indicating a marked reduction in actomyosin-driven contractility. Consistent with this observation, FRET-based actin tension sensors—which report molecular-scale filament tension by detecting energy transfer between donor and acceptor fluorophores—revealed higher FRET efficiency in aged cells, signifying decreased actin filament tension (Fig. 1b). To complement these findings, nanoindentation analysis was performed to assess the stiffness of individual cells by measuring their resistance to localized deformation; the Young’s modulus of young BMSCs (≈790 Pa) was significantly higher than that of aged cells (≈470 Pa; Fig. 1c), reflecting a more robust cytoskeletal network in the young group. Confocal imaging further illustrated that actin stress fibers36 in aged cells were sparse, disorganized, and randomly oriented, whereas young BMSCs exhibited dense, well-aligned bundles (Fig. 1d). Together, these complementary assays consistently demonstrate that aged BMSCs exhibit markedly diminished traction forces, reduced cytoskeletal tension, lower stiffness, and disrupted actin organization, indicating a comprehensive decline in cellular mechanical competence with aging.

To determine whether this mechanical decline was accompanied by alterations in mechanotransduction, we focused on core components of the canonical cytoskeleton–nucleoskeleton–nucleus signaling axis. Specifically, we examined nonmuscle myosin IIa and its active phosphorylated form (P-myosin IIa at Ser1943), which directly drive traction force generation; Lamin A/C, which transmits mechanical stress from the cytoskeleton to the nucleus and maintains nuclear structural integrity; and the transcriptional coactivator YAP, whose nuclear localization serves as a readout of force-dependent mechanosignaling. Western blotting and immunofluorescence analyses revealed consistent reductions in total myosin IIa, P-myosin IIa, Lamin A/C, and nuclear YAP in aged BMSCs (Fig. 1e–f; Supplementary Fig. 6). These findings collectively indicate that the age-related decline in cellular mechanics is tightly linked to impairment of the cytoskeletal–nucleoskeletal mechanotransduction cascade, providing a mechanistic basis for the attenuated force responsiveness and functional deterioration observed in aged BMSCs.

Increased cellular traction force alleviates cellular senescence
Our study identified significant differences in mechanical properties between young and senescent cells, which led us to hypothesize that increased cellular traction force might contribute to functional restoration and alleviate senescence. The decreased mechanotransduction of senescent cells was then restored through mechanical stretching (5% strain and 0.02 Hz frequency). In addition, calyculin A (Supplementary Figs. 26–28 showed that calyculin A exhibited no cytotoxicity and maintained efficiency at the working concentration), the myosin light chain phosphatase inhibitor, was employed to promote the myosin II activity and cell traction force37. After confirming that both methods effectively increased cellular traction forces (Supplementary Fig. 7), we evaluated markers of cellular senescence. Enhanced traction forces resulted in a notable reduction in the expression of p16INK4a (Fig. 2a), p53, and p21WAF1/Cip1, as well as a decrease in the proportion of SA-β-gal-positive cells (Supplementary Figs. 9 and 10). Furthermore, enhancing cellular force in young BMSCs produced a modest reduction in senescence markers, indicating that aged cells are more responsive to mechanical cues than their younger counterparts (Supplementary Fig. 11). These findings provide strong evidence supporting the hypothesis that boosting mechanotransduction can mitigate cellular senescence-related markers.

Given that replicative depletion is a key feature of aging, we examined the impact of mechanotransduction on cell proliferation using EdU assays, Ki67 staining, and CCK8 assays (Fig. 2b and Supplementary Fig. 13). Our results showed that restoring traction force in senescent cells significantly improved proliferation. These findings highlight the critical role of mechanotransduction in aging, affecting both senescence markers and proliferative capacity.

Furthermore, to investigate the impact of the 3D microenvironment in bone marrow38 (soft tissue) on BMSCs, we employed gelatin methacrylate (GelMA) hydrogels and ECM-mimicking GelMA fiber networks (Young’s modulus of 1–4 kPa, Supplementary Fig. 14), both engineered to replicate bone marrow-like stiffness. Consistent with observations on 2D substrates, mechanotransduction activity, as indicated by Lamin A/C levels, remained higher in young cells compared to senescent cells in both 3D models (Fig. 2c, d). Moreover, enhanced mechanotransduction activity through calyculin A treatment significantly reduced p21 expression in senescent cells, highlighting the pivotal role of mechanotransduction in mitigating cellular senescence even within a 3D bone marrow-mimicking microenvironment.

On the other hand, we explored that weather-reduced mechanical stimuli could induce cellular senescence. To model reduced mechanical stimuli, young BMSCs were seeded on the soft hydrogels with a shear modulus of 1.5 kPa to adjust force balance at the cell adhesion interface, resulting in decreased intracellular traction39. Alternatively, blebbistatin (Supplementary Figs. 26–28 showed that blebbistatin exhibited no cytotoxicity and maintained efficiency at the working concentration), a selective myosin II inhibitor, was used to treat the young cells, which effectively eliminated intracellular traction forces40. Following validation of mechanotransduction disruption (Supplementary Fig. 8), we assessed levels of p16INK4a, p53, p21WAF1/Cip1 and SA-β-gal staining. Both mechanical and pharmacological interventions that attenuated cellular mechanotransduction induced age-associated phenotypes and promoted stem cell senescence (Fig. 2a and Supplementary Figs. 9 and 10). Diminishing cellular force in senescent cells further accelerated aging, albeit less markedly than in young cells, underscoring a reduced but still consequential sensitivity to mechanotransduction impairment (Supplementary Fig. 12). Moreover, EdU assays, Ki67 staining, and CCK8 assays demonstrated a significant reduction in proliferation capacity of young BMSCs subjected to diminished intracellular traction forces, reinforcing the conclusion that reduced mechanotransduction drives cellular senescence and impairs cellular proliferation (Fig. 2b and Supplementary Fig. 13). These effects were notably validated in the 3D GelMA hydrogels and ECM-mimicking fiber networks (Young’s modulus: 1-4 kPa), where reduced cellular forces induced by blebbistatin in young cells promoted senescence phenotypes (Fig. 2c, d).

Overall, cellular mechanotransduction serves as a pivotal regulator in the aging process.

Diminished mechanotransduction activity in aged bone tissues
Considering the decreased cellular traction force in aged BMSCs, we further dissected the mechanotransduction activity in aged tissues. Compared to 2-month-old mice, 20-month-old mice exhibited a significant decline in trabecular number, increased trabecular spacing, and decreased collagen I levels (Fig. 3a, b). These changes in bone microarchitecture severely compromise the niche supporting BMSCs, emphasizing the need to understand mechanotransduction pathways in aging.

To assess mechanotransduction activity, we analyzed the phosphorylation of focal adhesion kinase (p-FAK) at Tyr39741 and the nucleus YAP42, which are well-known mechanotransduction markers and are sensitive to intracellular traction force. Immunofluorescence or immunohistochemistry analysis revealed a notable downregulation of both p-FAK (Fig. 3c) and YAP (Fig. 3d) in bone marrow tissue from aged mice (20 months), in stark contrast to the tissue from younger mice (2 months). In addition, the nuclear lamina in aged mice displayed discontinuities and reduced lamin B levels, indicating compromised nuclear integrity and decreased nuclear tension (Fig. 3e). Aging markers, such as p16INK4a (Fig. 3f), P21 (Fig. 3g) and rH2A.X (Fig. 3h), were markedly elevated in older mice compared to younger mice.

These findings collectively highlight an age-associated decline in cellular mechanotransduction activity within the bone marrow, accompanied by compromised nuclear structure and increased senescence markers.

Mechanical stimulation modulates bone aging and remodeling in vivo
To investigate how mechanical cues regulate bone aging, we applied two complementary in vivo paradigms in both aged and young mice: vibrational loading (20 min/day for 30 days with rest days; force+) and tail suspension (30 consecutive days; force−) (Fig. 4a).

In aged mice, tail suspension did not further reduce trabecular or cortical bone mass (Supplementary Fig. 15), indicating that acute unloading has minimal impact once age-associated bone loss is established. We therefore focused on the effects of vibrational loading. Vibration markedly increased mechanotransduction in femoral trabecular bone, as indicated by elevated p-FAK levels, greater nuclear YAP localization, and improved nuclear envelope integrity reflected by higher Lamin B expression (Fig. 4b, c). These changes were accompanied by reductions in senescence-associated markers (p21, p16, and p53), demonstrated by immunohistochemistry and western blot (Fig. 4d and Supplementary Fig. 16). Bone marrow qPCR likewise showed downregulation of p21, p16, and p19 (Supplementary Fig. 16). Vibration also reduced inflammatory signaling43, with decreased expression of IL-1β, IL-6, CXCL1, CCL2, IL-1α, and TNF-α in bone marrow (Fig. 4e and Supplementary Fig. 17). Double immunofluorescence analysis of CD105 and TNF-α demonstrated a decreased proportion of inflammatory BMSCs following vibrational loading (Fig. 4f), accompanied by reduced expression of CD45 and IL-1β in trabecular bone (Supplementary Fig. 17). Histological staining and micro-CT analysis demonstrated recovery of trabecular architecture and bone mass following vibration (Fig. 4g–i), whereas cortical bone showed no detectable changes (Supplementary Fig. 18). Examination of osteoblast- and osteoclast-associated markers in bone marrow RNA and serum indicated that vibrational loading predominantly enhanced osteoblast activity, with marked increases in Col1a1, Bglap, and serum PINP, while osteoclast markers (Mmp9, Acp5) and CTX were less affected (Supplementary Fig. 19).

In young mice, vibrational loading did not induce detectable changes in trabecular or cortical bone mass (Supplementary Fig. 15), suggesting that younger bone is closer to a homeostatic state and less responsive to additional mechanical input. By contrast, tail suspension produced pronounced trabecular deterioration: mechanotransduction was impaired, as indicated by reduced p-FAK levels, loss of nuclear YAP, and decreased Lamin B expression (Fig. 4b, c), accompanied by upregulation of senescence-associated markers p16, p21, and p53, as shown by immunohistochemistry and Western blot (Fig. 4d and Supplementary Fig. 16). Bone marrow qPCR further confirmed increased expression of p21, p16, and p19 following unloading (Supplementary Fig. 16). Inflammatory cytokines, including IL-1β, IL-6, CXCL1, CCL2, IL-1α, and TNF-α, were significantly elevated in the bone marrow (Fig. 4e and Supplementary Fig. 17), and CD105/TNF-α double staining revealed an increased fraction of inflammatory BMSCs (Fig. 4f). Functionally, tail suspension caused marked trabecular bone loss, as assessed by histology and micro-CT (Fig. 4g–i), while cortical bone remained largely unaffected, as confirmed by micro-CT analysis (Supplementary Fig. 18). Osteoblast and osteoclast analyses confirmed that trabecular loss was primarily due to reduced osteoblast activity, rather than changes in osteoclast function (Supplementary Fig. 19).

Traction forces modulate chromatin accessibility at the FOXO1 locus
To investigate how intracellular traction force regulates aging, we focused on chromatin remodeling, a process linked to chromatin accessibility and gene expression44, and cellular functions. Chromatin remodeling is mediated by mechanotransduction, as previously reported45. Accordingly, we conducted a comprehensive analysis of chromatin accessibility in young and senescent cells. Confocal microscopy imaging of nuclear DNA distribution and subsequent calculation of chromatin condensation parameters (CCP) revealed distinct differences: senescent BMSCs exhibited a more heterogeneous DNA staining pattern compared to young cells, indicating increased chromatin condensation. When traction forces in senescent cells were enhanced using mechanical stretching or calyculin A treatment, DNA distribution became more homogeneous, reflecting decreased chromatin condensation. Conversely, reducing mechanical stimuli through soft hydrogels or blebbistatin treatment in young cells resulted in a DNA distribution resembling that of senescent cells (Fig. 5a, b).

We further assessed chromatin accessibility by analyzing histone modifications, which serve as key indicators of chromatin openness and DNA packaging. Chromatin accessibility is closely linked to histone acetylation (H3ac and H4ac), deacetylase 2 and 3 (HDAC2 and HDAC3), and demethylation of lysine 9 and/or 27 in histone H3 (H3K9me3, H3K27me3)46,47. Enhanced traction forces via calyculin A in senescent BMSCs significantly restored histone acetylation levels, whereas reduced traction forces in young cells triggered deacetylation of H3 and H4 (Supplementary Fig. 20). This pattern was corroborated by the expression levels of HDAC2 and HDAC3, which decreased in response to increased cellular forces (Supplementary Fig. 21). Additionally, the demethylation of H3K9me3, but not H3K27me3, was responsive to force modulation (Supplementary Fig. 20).

To further investigate the role of traction forces in chromatin accessibility, we performed ATAC-seq48 to analyze changes in the chromatin landscape. The analysis included senescent cells subjected to mechanical stretching (5% strain, 0.02 Hz), referred to as “senescent, Force (+),” and young MSCs exposed to reduced mechanical stimuli, referred to as “young, Force (-)”. Supplementary Table 1, Figs. 29–33 confirm the high quality of ATAC-seq libraries and reflect the effects of mechanical force modulation on chromatin accessibility. Chromatin accessibility at transcription start sites (TSSs) revealed a significant decrease in accessible peaks in the Force (−) group compared to the young group. In contrast, the Force (+) group exhibited higher TSS accessibility peaks compared to the senescent group, approaching levels observed in the young group (Fig. 5c). These findings align with CCP and histone modification data. KEGG enrichment analysis revealed that force-dependent changes in chromatin accessibility impacted key pathways involved in cellular senescence (Supplementary Fig. 22). To elucidate the mechanisms by which mechanical force regulates cellular senescence, we then concentrated on pathways associated with cellular senescence. Among these, FOXO signaling emerged as one of the most significantly impacted pathways (Fig. 5d), underscoring its pivotal role in the force-regulation of aging49. Chromatin landscape analysis of FOXO signaling genes (Fig. 5e), along with IGV visualization (Fig. 5f), specifically highlighted FOXO1 as a critical force-sensitive target. ATAC-seq data showed that chromatin accessibility at the TSS of FOXO1 was markedly reduced in the Force (−) group compared to the young group, consistent with the global decrease in TSS accessibility under reduced mechanical stimuli. In contrast, the Force (+) group exhibited increased TSS accessibility at FOXO1 compared to the senescent group, approaching levels observed in the young group (Fig. 5e, f). These trends align with the broader ATAC-seq findings, supporting the hypothesis that FOXO1 activation mediates the anti-senescence effects of traction forces.

Traction forces mediate cellular rejuvenation through FOXO1
To test whether intracellular forces regulate FOXO1 expression, we first examined its transcriptional levels under different mechanical conditions. Quantitative PCR revealed that senescent cells subjected to mechanical stretching showed a significant upregulation of FOXO1 expression, whereas young cells cultured on soft substrates (reduced force) displayed marked downregulation (Fig. 6a). We next assessed FOXO1 protein expression. Immunofluorescence staining demonstrated that traction force enhancement, either by mechanical stretching or calyculin A treatment, increased FOXO1 levels in senescent cells, whereas force reduction by blebbistatin treatment or soft substrates suppressed FOXO1 expression in young cells (Fig. 6b).

To determine whether FOXO1 is essential for force-mediated rejuvenation, we silenced FOXO1 in senescent cells (Fig. 6c) and evaluated senescence markers under mechanical stretching. Increased forces failed to reduce p21 and p16 expression upon FOXO1 knockdown (Fig. 6d), indicating that FOXO1 is indispensable for the rejuvenating effects of traction forces.

We then extended these observations in vivo. Aged mice subjected to vibration (Force+) exhibited significantly increased nuclear circularity compared with aged controls, whereas young mice under tail suspension (Force –) displayed reduced nuclear circularity (Fig. 6e). These findings reflect force-dependent alterations in nuclear mechanics and chromatin openness. We further assessed histone acetylation, a well-established marker of chromatin accessibility. Immunostaining for H4ac revealed increased levels in trabecular bone regions beneath the growth plate of aged mice exposed to vibration, whereas young mice subjected to tail suspension showed reduced H4ac expression (Fig. 6f, g).

Finally, to confirm FOXO1 dynamics in bone marrow stem cells, we performed CD105 and FOXO1 double immunohistochemistry. FOXO1 positivity was markedly upregulated in aged mice under vibration, but downregulated in young mice subjected to tail suspension (Fig. 6h, i), consistent with the in vitro findings.

In summary, these results demonstrate that intracellular traction forces modulate senescence through chromatin remodeling and FOXO1 activation.

Mechanical overstimulation accelerates cellular senescence
Moderate levels of mechanical stimulation significantly suppressed aging markers in BMSCs, as described above. We next asked whether intensifying the mechanical input would further augment these rejuvenating effects. To this end, cells were subjected to higher-frequency cyclic stretching (5% strain at 0.1 Hz; Fig. 7a) or to excessive strain amplitude (25% strain at 0.02 Hz; Fig. 7b). Contrary to expectation, these intensified regimens did not enhance rejuvenation. Instead, they provoked a senescent phenotype, as demonstrated by upregulation of senescence markers p16INK4a and p21WAF1/Cip1 (Fig. 7a–c). The pro-senescent effect of force overstimulation was further corroborated in senescent BMSCs exposed to continuous calyculin A for 72 h, both in ECM-mimicking 3D fiber networks (Fig. 7d) and on conventional tissue culture polystyrene (TCPS; Supplementary Figs. 23 and 24). These findings indicate that intracellular traction force is a double-edged sword in the aging process: while moderate levels restore youthful features, excessive force overwhelms cell tolerance, tipping the balance toward irreversible senescence.

Given the established role of DNA damage as a central driver of cellular aging, we next investigated whether force-induced senescence was associated with increased genomic instability. Comet assays revealed a pronounced rise in DNA damage under overstimulation, with significantly higher tail DNA content and extended tail moment length in senescent BMSCs subjected to high-intensity stretching, whereas moderate stimulation did not alter these parameters (Fig. 7e, f).

The force-induced DNA damage can be partially explained by the loss of steric effects resulting from highly condensed DNA, which was proven by a molecular model mimicking chromatin. The isolated DNA chains assembled onto protamine, mimicking the condensed chromatin structure within the cell nucleus, exhibit protection against fragmentation induced by ultraviolet irradiation. In contrast, freely dispersed DNA chains, which simulate the dissociation of DNA from histones under intracellular traction force, were fragmented into smaller pieces upon exposure to ultraviolet irradiation (Fig. 7g).

Optimal mechanical stimulation reduces bone aging and inflammation
To evaluate the dose-dependent effects of mechanical stimulation on bone recovery in aged mice, we performed vibrational loading over a 30-day period. Three experimental groups were established: intermittent vibration (force+, 20 min/day with alternating rest days), continual vibration (force++, 20 min/day without rest), and baseline controls (Fig. 8a). Intermittent vibration significantly enhanced trabecular bone formation, as evidenced by increased collagen deposition (Fig. 8b) and improved trabecular architecture observed via micro-CT (Fig. 8c). In contrast, continual vibration led to compromised trabecular structure and reduced collagen levels (Fig. 8b, c), indicating that excessive mechanical stimulation is detrimental to bone integrity.

Consistently, analysis of aging markers revealed that intermittent vibration suppressed expression of p16, p21, and p53, whereas continual vibration elevated these markers (Fig. 8d, e and Supplementary Fig. 25).

Inflammatory responses within trabecular bone followed a similar pattern. Intermittent vibration lowered IL-1β and CD45 expression (Fig. 8f), while continual vibration upregulated these markers. Bone marrow PCR analysis further supported these findings, with intermittent vibration reducing IL-1β, CXCL1, CCL2, and TNF-α expression, and continual vibration promoting their expression (Supplementary Fig. 25). DNA damage was elevated following continual vibration but remained unchanged under intermittent stimulation (Fig. 8g), highlighting the importance of force intensity in determining cellular outcomes. Together, these data indicate that judicious mechanical stimulation through intermittent vibration mitigates aging- and inflammation-associated markers while promoting trabecular bone recovery.

Beyond the skeletal effects, intermittent vibration conferred systemic benefits. Mice subjected to intermittent vibration demonstrated improved physical performance across grip strength, treadmill running and water maze tests. Serum analysis revealed reduced levels of IL-6 and IL-1β, as well as lower platelet counts (PLT), consistent with decreased systemic inflammation (Fig. 8h). Furthermore, assessment of key peripheral organs involved in immune modulation—liver and kidney—showed reduced expression of inflammatory markers CD68 and S100A8 following intermittent vibration (Fig. 8i, j), confirming broad anti-inflammatory effects.

Methods
BMSC isolation and culture
Bone marrow aspirates (~1–2 mL) were collected from the posterior superior iliac spine (PSIS) of healthy donors aged 18–35 years (young group) and 60–80 years (elderly group), all of Chinese descent, at West China Hospital of Sichuan University (n=38 donors). Donors were screened to exclude hematological disorders, and both male and female donors were included, with sex recorded at the time of collection (detailed donor information is provided in the Supplementary Information). Participants gave informed consent to the experiment and the publication of information. All human tissue samples were used exclusively for experimental analyses and were destroyed immediately upon completion of the assays. All procedures involving human participants were reviewed and approved by the Institutional Review Board of West China Hospital, Sichuan University (Approval No. WCHSIRB-D-20220290).

Bone marrow aspirates were processed to isolate mononuclear cells (MNCs) by density gradient centrifugation using Ficoll-Paque, removing red blood cells and other non-nucleated cells. The MNC fraction was plated in T-75 flasks and cultured in low-glucose DMEM supplemented with 10% fetal bovine serum (Gibco) at 37°C in a humidified atmosphere containing 5% CO2. After 48–72 h, non-adherent cells were removed by medium replacement, and adherent cells were expanded as bone marrow-derived stromal cells (BMSCs). Cells at passages 1–2 were used for all subsequent experiments.

This isolation procedure enriches for BMSCs, as evidenced by their adherence to plastic, fibroblast-like morphology, positive expression of MSC markers CD73 and CD146, negative expression of hematopoietic markers CD34 and CD45, and functional properties including colony-forming ability, proliferation, multilineage differentiation, and mechanosensitive responses.

BMSC treatment and mechanical manipulation
Chemical inhibitors. Blebbistatin (20 μM, B0560, Sigma) or calyculin A (0.1 nM, ab141784, Abcam) was added to the culture medium and maintained throughout the treatment period. Cell viability was assessed using live/dead assays according to the manufacturer’s instructions.

Mechanical stimulation. To enhance cellular traction force, cells were seeded in a stretching device (Cell & Force, Cell Tank, Hangzhou Surface & Force Technology Co., Ltd) with strains of 5% and 25% at frequencies of 0.02 and 0.1 Hz. To reduce cellular traction force, cells were cultured on soft hydrogels with a shear modulus of 1.5 kPa for 24 h.

Genetic manipulation. Electroporation (Celetrix Biotechnologies, China) was used to deliver siRNA targeting FOXO1 (sense: GAGCGUGCCCUACUUCAAG; antisense: CUUGAAGUAGGGCACGCUC). Plasmid DNA constructs were obtained from Hanbio Biotechnology (Shanghai, China).

Cellular mechanics and mechanotransduction in human BMSCs
Nanoindentation. Nano-indentation was used to measure Young’s modulus of young and senescent BMSC. The cells were pressed by a ball-shaped indenter (Piuma Chiaro, Optics11). The diameter of the nano-indentation indenter is 10.5 μm, the stiffness of the indenter is 0.05 N/m, and the indentation depth is 3 μm in this study. The modulus was determined by fitting force-indentation curves to known models according to the manufacturer’s instructions.

Traction force microscopy (TFM). TFM was used to detect cellular mechanotransduction according to our previous protocol56. The cells were seeded on deformable hydrogels embedded with fluorescent beads. The shifting of the beads, caused by cell adhesion-induced hydrogel deformation, could be recorded to calculate the traction force exerted by the cells. Firstly, RGD-decorated PEG hydrogels containing 0.5 μm fluorescent carboxylated polystyrene beads (Sigma) were fabricated and the micro-beads existed at the surface of the hydrogels. After hydrogels were soaked for 48 h, young and senescent BMSCs were seeded on the hydrogels. A confocal microscope (Leica) was exploited to take images of beads and the spreading area of cells, regarding the shifted position of beads induced by cell migration. Finally, both young and senescent cells were cleared up by dealing with 1% SDS and taken photos afterward to identify the original position of beads. Images were first aligned to correct for the experimental drift of the samples using the ImageJ plugin “Align slices in stack”. The displacement field in a spread cell region was subsequently calculated by the “particle image velocimetry” plugin in ImageJ. The obtained result was reconstructed using the “Fourier transform traction cytometry” plugin in order to generate the traction force field as a vector plot.

Quantification of fiber’s anisotropy
FibrilTool, a plugin of ImageJ, was introduced to quantify the orientation of fibrillar structures in the studies according to the protocol. After capturing the pictures of β-actin, we calculated the anisotropy of young and senescent cells, respectively, in order to compare the differences in cellular mechanical characteristics. Regarding the anisotropy score, the following convention was used, i.e., 0 for no order (purely isotropic arrays) and 1 for perfectly ordered (purely anisotropic arrays). Another plugin of ImageJ named OrientationJ was also used to measure the orientation of the fiber where the color representation reflected different orientations.

Actin tension sensor. Primary extracted young- and senescence-derived human BMSCs were spread on glass cell culture dishes and then cultured for 24 h in the absence of antibiotics. Opti-MEM (Gibco) was used to dilute plasmid Actin-cpstFRET-Actin (80643, Addgene), and X-tremeGENE HP DNA transfection reagent (Sigma) was added at 3 μL/μg to transfect cells for 48 h. Cells expressing β-actin-CFP/YFP-β-actin fusion protein at sufficiently high levels for easy visualization and imaging were selected for imaging. However, cells with excessively high expression levels were avoided. In addition, cells expressing the fusion protein through the whole cell, including the nucleus, were excluded, and only cells expressing the sensor in a discernible cytoplasmic skeleton were imaged.

At 0, 1, 2, 3, 4, 5, 6, 7, 8, 24 h after administration, cells were imaged using a confocal microscope with a ×20 objective (Leica). FRET results (530 nm) were captured with a 405 nm laser to excite β-actin-CFP. At least 20 cells were quantified for each condition. The cell images used for all values shown in the figure were acquired under the same imaging condition. Considering the different expression levels of cytoskeletal proteins in different cells, the YFP fluorescence intensity obtained by fixed 488 nm excitation was standardized to ensure the normal operation of the sensor and FRET assay. Moreover, the ratio of the fluorescence intensity of YFP (530 nm) excited at 405 nm and 488 nm (FRET/YFP) was used as a parameter to reflect the cytoskeletal tension.

Animal models and experimental procedures
Young and aged mice. Female C57BL/6J mice aged 2 months (young group) and 20 months (aged group) were obtained from Chengdu Dossy Experimental Animals Co., Ltd. To investigate the effects of mechanical interventions on bone development, bone aging, and mechanotransduction, both tail suspension and vibrational loading experiments were performed on mice from each age group. All animal procedures were approved by the Ethics Committee of West China Hospital of Stomatology, Sichuan University (approval number: WCHSIRB-D-2022-174), and were conducted in accordance with institutional and national guidelines for the care and use of laboratory animals.

Tail suspension model. Mice were subjected to hindlimb unloading using a tail suspension model for 30 consecutive days. Animals were suspended by taping the tail to a horizontal bar, allowing forelimb contact with the cage floor while keeping the hindlimbs elevated. Mice were housed individually in custom cages and provided with a complete nutritional gel diet during the suspension period. Tail suspension was applied to both young (2 months) and aged (20 months) mice to assess age-dependent responses.

Vibrational loading model. Mechanical stimulation was delivered using vibrational loading for 20 min per day over a 30-day period. Two vibration regimens were applied: (1) intermittent vibrational loading (one day of vibration followed by one day of rest, repeated for 30 days), and (2) daily continuous vibrational loading (20 min per day for 30 days). Vibrational loading was administered to both young and aged mice to evaluate the effects of increased mechanical stimulation across age groups. A baseline control group without mechanical intervention was included for each age.

MicroCT tomography evaluation. The femurs from mice with different treatments were scanned using micro-CT (SkyScan 1176 desktop X-ray Micro-CT system, Skyscan, Bruker) with 0.5 mm Al filter, image pixel size: 17.75 μm, voltage: 60 kV, electrical current: 400 μA, and exposure time: 1080 ms. The trabecular region of interest (ROI) in the micro-CT image was selected individually and manually to ensure the trabecular bones of the distal femoral metaphysis were included in the ROI as much as possible. The 3D reconstruction was performed using all the outlined slices by NRecon (Skyscan, Bruker) cone-beam reconstruction software. Three-dimensional microarchitecture parameters, including bone volume fraction (BV/TV), structure model index (SMI), trabecular number (Tb.N), trabecular thickness (Tb.Th), and trabecular bone pattern factor (TB.Pf), were calculated using CTAn (CTAn, Skyscan, Belgium).

Histological analyses. Histological analyses involved immunohistochemistry, immunofluorescence, hematoxylin and eosin (HE) staining, as well as Masson staining. The dissected femurs from mice with different treatments were fixed in 4% paraformaldehyde for 2 days and then decalcified in 12% EDTA for 1 week. Slides were subjected to sodium citrate buffer at 99°C for 20 min for antigen retrieval and then incubated with rabbit polyclonal anti-collagen I (1:800, Servicebio, Cat No: GB11022-3), rabbit anti-phospho-FAK (1:50, Thermo Fisher Scientific, Cat No: 700255), anti-Lamin B1 (1:100, Abcam, Cat No: ab16048), rabbit polyclonal anti-YAP (1:100, Genxspan, Cat No: GXP539020), rabbit polyclonal anti-p16 (1:100, Invitrogen, Cat No: MA5-17142), anti-p21 (1:100, Abcam, Cat No: ab188224), anti-CD45 (1:100, Abcam, Cat No: ab105550), anti-IL-1β (1:100, Santa cruz, Cat No: sc-52012), anti-H4AC (1:100, Abcam, Cat No: ab177790), rabbit polyclonal anti-FOXO1 (1:100, Cell signaling, 2880S), rabbit polyclonal anti-phospho-Histone H2A.X (ser139) (1:100, Cell Signaling Technology, Cat No: 9718), anti-DNA-RNA damage (1:100, Abcam, Cat No: ab62623). The anti-rabbit IgG antibody was used as secondary antibody (1:400, SeraCare, Cat No: 5220-0336). HE staining was used to assess the histological structure and morphology of the bone tissue. For Masson staining, the tissue was stained by Masson staining solution (Solarbio, China) according to the instruction. Additionally, liver and kidney tissues were stained using immunohistochemistry with anti-S100A8 (1:100, Abcam, Cat No: ab92331) and anti-CD68 (1:100, Abcam, Cat No: ab125212) antibodies to evaluate inflammatory markers. Immunohistochemistry and immunofluorescence slides were scanned in high resolution using the Pannoramic MIDI scanner (3DHISTECH, Hungary) for comprehensive analysis.

Physical test of animals
Grip-strength test. This grip-strength test was performed according to the previously described procedure57. Grip strength of the mouse’s limbs was recorded using a grip strength meter (SA 415, Jiangsu Science Bio-Tech Co., Ltd, Jiangsu, China). Before testing each mouse, the gauge was reset and stabilized to 0 g. The mouse was allowed to grasp a horizontally held metal grid with all four paws, and then it was gently pulled backward by the tail until it could no longer hold onto the grid. The peak pulling force, measured in grams, was automatically recorded on a digital force transducer. Each mouse underwent four trials after adequate rest, and the average value was used to represent the muscle grip strength of the individual mouse.

Treadmill exercise. The treadmill exercise was conducted as previously described58. A treadmill (SA101B, Jiangsu Science Bio-Tech Co., Ltd, China) with eight lanes (8.5 × 61 cm2 each) and individual moving belts was used to assess exercise endurance in mice. Each lane featured an electric shock grid (0.3 mA for 10 s) to motivate running. Mice underwent pre-training for 1 week at 5 m/min for 10 min, followed by 10 m/min for 30 min, with an acceleration of 70 m/min² on a flat incline. Sessions were conducted twice daily for 40 min with a minimum interval of 3 h. Subsequently, treadmill exercise was performed every morning for 3 days using the same protocol. Exercise tests continued until mice either touched the shock grid 150× or were unable to continue after a maximum shock duration of 10 s. Time to exhaustion, distance covered, and total shocks were recorded.

Morris water maze test. The Morris water maze was used to test spatial learning and memory. The Morris water maze test was performed in an opaque circular pool filled with water (diameter 1.5 m, height 0.6 m, 22−23°C). An 8 cm diameter escape platform was positioned in the second quadrant of the pool with its top 1 cm below the water surface. A video analysis system (SA201, Jiangsu Science Bio-Tech Co., Ltd, Jiangsu, China) was used to observe and record each mouse’s swimming pattern. All water maze training and testing were conducted during the animals’ active phase of the light cycle (07:00−19:00). Mice were released into the pool from four different starting positions in the pool and allowed to swim for a maximum of 60 s. If a mouse failed to find the platform within 60 s, it was gently guided to the platform and allowed to stay there for 30 s. Before the formal testing began, each animal received two swimming training sessions per day for 5 days, with the second session occurring at least 2 h after the first. During the training trials, the swimming speed, the escape latency and escape distance were recorded. After 5 days of training, mice that still cannot swim and are unable to find the platform within 60 s will be excluded from the study. The formal test was conducted on the 6th day after the training ended, with the platform removed from the pool. The number of crossings of the platform area, average swimming speed, total distance, and the time spent swimming in the second quadrant where the platform had been located were recorded. The swimming pattern of each mouse were recorded using a camera connected to a computer.

Enzyme-linked immunosorbent assay
After the behavioral tests, the mice were sacrificed, and serum was collected from the eyeballs for cytokine analysis. According to the manufacturer’s instructions, the quantitative ELISA kits (Mouse IL-1 beta Uncoated ELISA Kit, 88-7013, Thermo Fisher Scientific Inc, USA; Mouse IL-6 Uncoated ELISA Kit, 88-7064-88, Thermo Fisher Scientific Inc, USA) were used to measure the concentration of IL-1β and IL-6 in the serum. The procedures were done according to the instruction manual: (1) Firstly, coat each well of a Corning™ Costar™ 9018 ELISA plate with 100 μL/well of capture antibody diluted in Coating Buffer. Seal the plate and incubate overnight at 4°C. Block the wells with 200 μL of ELISA/ELISPOT Diluent (1×) and incubate at room temperature for 1 h. (2) Then, aspirate the wells and wash 3 times with >250 μL/well of Wash Buffer, allowing approximately 1 min of soaking for each wash. Blot the plate on absorbent paper to remove residual buffer. (3) 100 μL of standard, control buffer, and sample were added respectively into wells of the ELISA plate coated with antibody IL-1β or IL-6, then incubated overnight at 4°C for maximum sensitivity. (4) After that, the wells were washed 5 times. Add 100 μL/well of diluted Avidin-HRP. Seal the plate and incubate at room temperature for 30 min. (5) Washed again, add 100 μL/well of 1× TMB Solution and incubate at room temperature for 15 min in the dark. (6) Add 100 μL/well of Stop Solution. (7) Finally, read absorbance read plate at 450 nm within 30 min of stopping reaction. If wavelength substraction is available, substract the values of 570 nm from those of 450 nm and analyze data.

Western blotting
After euthanising the mice, femurs were quickly dissected, weighed and collected into Eppendorf tubes. Samples were snap-frozen on dry ice and stored at −80°C for subsequent analysis. Then tissues were lysed in radioimmunoprecipitation assay buffer (RIPA) supplemented with 1% protease inhibitor and 1% phosphatase inhibitor and homogenized using a homogenizer at 4°C. The mixture was spun down at 12,000×g for 5 min at 4°C to pellet the insoluble material. The supernatant was lysed through probe sonication on ice to release cytosolic proteins. Proteins were isolated by centrifugation at 12,000×g for 15 min at 4°C, resolved by SurePAGE 4–12% Bis-Tris gels, and transferred to PVDF membrane (0.45 μm). The membranes were blocked in block buffer for 1 h at room temperature and incubated with primary antibodies (anti-GAPDH antibody (1:5000, Servicebio, GB15004), anti-p53 antibody (1:1000, Cell Signaling, 2524) in primary antibody dilution buffer (Beyotime, China) overnight at 4°C. Blots were washed three times with PBST and incubated with secondary antibodies (goat anti-rabbit IgG secondary antibody HRP conjugated (1:10,000, Sianalway Antibody, L3012) and goat anti-mouse IgG secondary antibody HRP conjugated (1:10,000, Sianalway Antibody, L3032) at room temperature for 1 h. Blots were further washed three times with PBST for 5 min. Then blots were treated with an ECL plus detection kit and imaged using ChemiScope 6100 system.

RNA extraction and real-time PCR
Total RNA was isolated from cells using the FastPure Complex Tissue RNA Isolation Kit (RC113-01, Vazyme) following the manufacturer’s instructions. During purification, all centrifugation steps were performed at 12,000×g for 1 min at 4°C unless otherwise specified. RNA concentration and purity were assessed by spectrophotometry (NanoDrop One, Thermo Fisher Scientific). For complementary DNA (cDNA) synthesis, 1 μg of total RNA was reverse-transcribed using the HiScript III RT SuperMix for qPCR (R323-01, Vazyme) according to the manufacturer’s protocol. Quantitative PCR was performed using the Taq Pro Universal SYBR qPCR Master Mix (Q712-02, Vazyme) on a CFX96 Real-Time PCR Detection System (Bio-Rad). The thermal cycling conditions were: 95°C for 30 s, followed by 40 cycles of 95°C for 10 s and 60°C for 30 s. Melt curve analysis was carried out to ensure primer specificity. Gene expression was calculated and normalized to GAPDH as the internal reference. Primer sequences used in this study are listed in Supplementary Table 2.

Cellular functional assays
Immunofluorescence and confocal microscopy. Cells were fixed for 15 min at 37°C with 4% paraformaldehyde, permeabilized with 1% Triton-X100 (Sigma) for 15 min and blocked with 5% BSA diluted in PBS for 15 min at RT. BMSCs were then incubated, overnight at 4°C with the indicated primary antibodies in 1% BSA. The following antibodies were used: Myosin IIa (1:100, Cell Signaling Technology, Cat No: 3403), phospho-Myosin IIa (1:200, Cell Signaling Technology, Cat No: 14611), Lamin A/C (1:200, Cell Signaling Technology, Cat No: 4777), YAP (1:100, Cell Signaling Technology, Cat No: 14074), phospho-Histone H2A.X (ser139) (1:200, Cell Signaling Technology, Cat No: 9718), p16 (1:200, Invitrogen, Cat No: PA1-30670), p21 (1:200, Cell Signaling Technology, Cat No: 2947), P53 (1:100, Cell Signaling Technology, Cat No: 2524), HDAC2 (1:200, Santa Cruz Biotechnology, Cat SC-9959), HDAC3 (1:200, Cell Signaling Technology, Cat No: 3949), Histone H3ac (pan-acetyl) (1:200, Active motif, Cat No: 39040), Histone H4ac (pan-acetyl) (1:200, EMD Millipore, Cat No: 06-866), H3K9me3 (1:200, Active motif, Cat No: 39062), H3K27me3 (1:200, Active motif, Cat No: 39055), ki67 (1:200, Abcam, ab15580) and rabbit polyclonal anti-FOXO1 (1:100, Cell signaling, 2880S). Following this, cells were incubated for 2 h at 4°C with the appropriate secondary fluorescent antibodies diluted in 1% BSA, protected from light. At last, nuclei were stained with DAPI (1:1000, Solarbio, China). Images were collected under a confocal laser scanning microscope (CLSM, Olympus, Japan).

Western blot experiments. BMSCs were seeded in six-well plates and treated with different conditions for different times. Then the experiment was finished according to standard protocol. Protein concentration was determined using the BCA Protein Assay Kit (Bio-Rad, U.S.), and then 30 μg of total protein was run on 8–12% SDS-PAGE gel (Beyotime, China), and electro-transferred to polyvinylidene difluoride (PVDF) membranes (Millipore) for 2–3 h at 120 mA in transfer buffer. The following antibodies were used: Myosin IIa, phospho-Myosin IIa, Lamin A/C, and GAPDH. The concentration of these antibodies in this experiment was 1:1000. After three 10 min washes with TBST, membranes were incubated with the appropriate secondary antibody (1:5000) for 1 h at room temperature. Immunoreactive bands were visualized by Tanon-5200 Multi (Tanon luminous imaging image storage and management software) after staining by a western lightning chemiluminescence detection kit (Zen Bioscience, China).

Senescence-associated β-galactosidase (SA-β-Gal) Staining. Senescent cells were identified by the SA-β-gal staining kit (Solarbio, China) according to standard report. After incubating with the reagent about 8–12 h, senescent cells would be stained with aquamarine blue, whereas young cells were not able to be colored.

Cell proliferation assay. To investigate the effect of cellular mechanotransduction on BMSC proliferation, an EdU incorporation assay (Beyotime, China) was performed. After treating cells under different mechanical stimulation conditions, the assay was conducted according to the manufacturer’s instructions to assess DNA synthesis and proliferation activity.

ECM-mimicking gelatin methacrylate (GelMA) fiber networks. The networks of suspended GelMA fibers were fabricated by a combination of electrospinning and 3D printing technology. GelMA (grafting rate: 90%) was dissolved at 0.1−1 g/ml in hexafluoro isopropanol (HFIP, Aladdin) with 0.01% 2-hydroxy-4′-(2-hydroxyethoxy)−2-methylpropiophenone photoinitiator (Irgacure 2959, Aladdin). The electrospinning process was successfully conducted using a conventional roller electrospinning apparatus, which included a high-voltage power source, syringe pump, and grounded roller enclosed within an environmental chamber. Electrospinning was conducted with a flow rate of 0.5−1 h, voltage set at 12 kV, and a gap distance of 10 cm. The fibers are received at rotating roller speeds of 800 r.p.m. Samples were crosslinked for 5 min under ultraviolet light (100 mW cm−2) at 365 nm to stabilize the connection between the fiber and the receiving mold. The samples were then exposed to ultraviolet light for 5 min in an Irgacure 2959 ethanol solution (150 mg/ml) to achieve crosslinking. The fibers were collected on polylactic acid (PLA; Tough PLA, Ultimaker, Netherlands) 3D printed square array (side length 1 mm) functionalized with methacrylate to enhance fiber adhesion. Briefly, PLA square arrays are printed using an extruder 3D printer (Ultimaker S5, Ultimaker, Netherlands). PLA mold was immersed in 60 mg/mL hexamethylene diamine/isopropyl alcohol solution at 40°C for 2 h to produce PLA-NH2 mold. After that, the PLA-NH2 mold was placed in a 10% methacrylic anhydride/ethanol solution at 50°C for 2 h to obtain a methacrylated PLA mold.

The Young’s modulus of fiber network was measured by a bionanoindenter (Optics11, Piuma Chiaro, Netherlands). The probe with stiffness of 0.025 N/m (Young’s modulus range: 0.1–100 Kpa) was carefully selected, featuring a tip radius of the probe measuring 12.5 μm and applying a pressure depth of 10,000 nm. The obtained data were fitted with the Hertzian contact model and the degree of fitting was 80%.

Fabrication of GelMA hydrogels. The circular glass slides of 9 mm diameter treated with 3-(Trimethoxysilyl) propyl methacrylate (S111153, Aladdin) were sterilized: immersed in 75% alcohol for 30 min, soaked in penicillin-streptomycin solution for 2 h, and then washed three times with sterile PBS for 5 min each time. The modified glass slides were dried in an ultra-clean bench. The 3.5% (w/w) GelMA solution was prepared using GelMA with a grafting rate of 90% and sterilized through a 0.22 μm filter. The 0.25% (w/v) photoinitiator lithium phenyl-2,4,6-trimethylbenzoylphosphinate (LAP) was mixed with the GelMA solution. Young or senescent BMSCs were respectively resuspended in the GelMA solution at a concentration of 1.5×105 cells/ml to prepare the final hydrogel precursor. Next, 15 μl hydrogel precursor was evenly spread on the modified glass slides and crosslinked for 30 s under ultraviolet light (100 mW/cm) at 405 nm. After crosslinking, the cell-laden GelMA hydrogels were placed in complete culture medium and incubated in a 37°C incubator with 5% CO2. After cells extended in the hydrogels (normally 2 days), the young cells were treated with 20 μmol/ml blebbistatin for 24 h, while the senescent BMSCs were treated with 0.1 nmol/ml calyculin A for 24 h.

ATAC-seq methodology for chromatin accessibility analysis
Cell Sample Preparation. Bone marrow aspirates were obtained from patients aged 60–80 years (old group) and 18–30 years (young group). Bone marrow-derived mesenchymal stem cells (BMSCs) were cultured by plating 1 mL of bone marrow into T-75 flasks containing growth medium supplemented with 10% fetal bovine serum (Gibco). BMSCs at passages 1–2 were used for all downstream experiments.

To enhance cellular traction forces, senescent BMSCs were seeded onto a stretching device (Cell & Force, Cell Tank, Hangzhou Surface & Force Technology Co., Ltd) and subjected to cyclic mechanical stimulation at 5% strain and 0.02 Hz for 2 h. Conversely, to attenuate traction forces, young BMSCs were cultured on soft hydrogels with a shear modulus of 1.5 kPa for 24 h.

ATAC-seq library preparation and sequencing. ATAC-seq libraries were generated using the Hyperactive ATAC-Seq Library Prep Kit for Illumina (Vazyme, TD711) in strict adherence to the manufacturer’s protocol. Importantly, intact whole cells were utilized rather than isolated nuclei to preserve the physiological relevance of cellular traction forces, as nuclear isolation disrupts these forces and may alter the chromatin landscape. This approach was critical for accurately examining the effects of mechanical forces on chromatin accessibility.

For each sample, 50,000 cells were used. Cells were washed twice with ice-cold 1× TW buffer (500×g, 4°C), resuspended in precooled lysis buffer, and incubated on ice for 5 min. After centrifugation at 500×g for 5 min, the supernatant was removed, and the cell pellet was resuspended in a precooled Tn5 transposome/transposition reaction mix. The reaction mix was gently vortexed to ensure uniform distribution and incubated at 37°C for 30 min to facilitate tagmentation. The reaction was terminated by adding a stop buffer, and the fragmented DNA was purified using VAHTS DNA Clean Beads, including two washes with 80% ethanol. The DNA was finally eluted in 26 μL of nuclease-free water.

Library amplification was performed under the following conditions: initial denaturation at 72°C for 3 min and 95°C for 3 min, followed by 14 PCR cycles (98°C for 10 s, 60°C for 5 s, 72°C for 1 min), with a final extension at 72°C for 1 min and a hold at 12°C. The amplified libraries were further purified with VAHTS DNA Clean Beads, washed twice with 80% ethanol, and eluted in 22 μL of nuclease-free water. Sequencing was conducted on an Illumina NovaSeq X (150-bp paired-end reads), generating approximately 8 Gb of data per library. Read quality was evaluated using FastQC (v0.11.9), and sequencing services were provided by Berry Genomics Co., Ltd (China).

ATAC-seq analysis. ATAC-seq analysis was performed by Touty.io (Shanghai, China). Raw sequencing reads were aligned to the human genome reference sequence (GRCh38) using the BWA-MEM algorithm. To ensure data quality, duplicate reads were removed and reads with a MAPQ score below 30 were excluded. Additionally, genomic regions listed in the ENCODE blacklisted regions were omitted from the analysis.

Peak calling was performed using Genrich (v0.6.1) with the following parameters: PCR duplicates were removed (-r), signals from mitochondrial and Y chromosomes were excluded (-e MT, Y), and ATAC-specific settings were applied (-j). Quantification of identified peaks was performed using BAMscale (v1.0), and the resulting data were normalized to TPM (transcripts per million). BigWig tracks were generated using deepTools (v3.5.3) with a bin size of 10 bp and normalized to TPM to facilitate visualization of ATAC-seq signals in regions of interest.

Differential peak analysis was conducted in R (v4.1.3) using the DESeq2 package (v1.38.0). Statistical significance was determined using a threshold of |log2FC|>0.5 and p-value < 0.05. Pairwise comparisons were denoted as “A-vs-B,” where “A” represents the experimental group and “B” represents the control group.

DNA protection experiments
DNA sample preparation. T4-DNA, 165.65 kbp was purchased from Nippon Gene, Tokyo, and used without further purification. YOYO-1 fluorescence staining dye was purchased from Invitrogen, Carlsbad, CA. T4-DNA was stained with YOYO-1 with an intercalation ratio of 100 base-pairs per dye molecule. Samples were prepared by dialysing solutions of T4-DNA against 10 mM Tris/HCl (1xT) buffer in micro-dialysers. The Tris is adjusted with HCl to pH 8.5, i.e., 2.9 mM Tris Cl and 7.1 mM Tris. The ionic strength of the buffer was calculated with the Davies equation for estimating the activity coefficients of the ions and a dissociation constant pK=8.08 for Tris. Protamine chloride from salmon was purchased from Sigma-Aldrich and dissolved in the Tris/HCl buffer. For the pre-equilibrated samples, solutions of protamine and DNA were subsequently mixed in equal volumes and incubated for 24 h at 277 K. The final DNA concentration is 3 mg/L. The buffer used for unpacking of DNA inside the channels was of the same composition, but with 100 mM of added NaCl. No anti-photobleaching agent was used, since it might interfere with protein-DNA binding.

Fluorescence and dark-field imaging: Bare and YOYO-1-stained DNA molecules dispersed in the relevant buffer were imaged in millimeter-sized droplets from bulk solution. For experiments conducted using FM, at least 10-run series, each of which is 5–10 s in length at 100 fps, was recorded. To prevent photocleavage, the imaging time is determined corresponding to experimental conditions. In DM, ~10 run series of 30 s length were recorded for each experimental condition. Each data point in this study was obtained by tracking and analyzing more than 200 single DNA molecules. The fluorescence of the stained DNA molecules was visualized with an Olympus IX71 inverted fluorescence microscope equipped with a 100 W mercury lamp, a UV filter set, and a 100-oil immersion objective. The exposure time of the YOYO-1 fluorescence was controlled using a UV light shutter. Images were collected with a charge-coupled device camera (Olympus DP70). For dark-field imaging, DNA molecules were imaged with a Nikon Eclipse 50i microscope with a CytoViva illuminator and a 100× oil immersion objective. Images were collected with a CCD camera (JVC TK-C921EG) connected via an analog-to-digital video converter (ADVC55, Canopus) to a computer.

RNA extraction and real-time PCR
Total RNA was extracted and purified using the FastPure Complex Tissue/Cell Total RNA Isolation Kit (RC113-01, Vazyme) according to the manufacturer’s protocol. Reverse transcription to complementary DNA (cDNA) was performed with the HiScript III RT SuperMix for qPCR (R323-01, Vazyme). Quantitative real-time PCR (qPCR) was conducted using Taq Pro Universal SYBR qPCR Master Mix (Q712-02, Vazyme) on a CFX96 Real-Time System (Bio-Rad). Gene expression levels were normalized to GAPDH expression. Primer sequences used in this study are listed in Supplementary Table 2.

Statistical analysis
All data are presented as mean±SD. Statistical analyses were performed using Prism 10.2.3 (GraphPad Software). Two-tailed Student’s t-tests were used for comparisons between two groups, and one-way ANOVA followed by appropriate post hoc tests was applied for comparisons involving more than two groups. P value < 0.05 was considered statistically significant. Exact P values and sample sizes (n) are provided in the corresponding figure and figure legends.

Reporting summary
Further information on research design is available in the Nature Portfolio Reporting Summary linked to this article.

Data availability
The ATAC-seq datasets generated in this study have been deposited in the NCBI Gene Expression Omnibus under accession number GSE287556. All other data supporting the findings of this study, including source data for all figures and supplementary information, are provided with this paper. Source data are provided with this paper.

Peer review rounds
[{"round": 1, "reviewer_comments": {"reviewer_1": "In the manuscript entitled \"Mechanical Rejuvenation of Senescent Stem Cells and Aged Bone via Chromatin Remodeling\", the authors reported a marked reduction in intracellular traction force and cellular mechanics in senescent bone marrow mesenchymal stem cells (BMSCs). They demonstrated that the moderate mechanical stimuli effectively restored cellular force, enhanced chromatin accessibility at the FOXO1 locus, activated its expression, and reversed cellular senescence and bone aging. This study unveils that optimizing mechanical stimulation provides a promising, straightforward approach to counteract age-related bone degeneration and systemic inflammation, offering significant potential for clinical translation. The presented findings are new and interesting, and reviewer has the following comments.\n1) In Fig.1E results, the authors performed Western blot (WB) analysis on several key proteins involved in mechanotransduction. However, given the multitude of cellular mechanotransduction pathways, the authors should clarify whether this investigation focuses on a predefined, specific pathway or represents a broader screening of multiple associated pathways. This clarification would significantly enhance comprehension of the research scope and improve the accuracy of the study.\n2) In Fig.2 staining results, the authors used different senescence markers (p16 in some panels, p21 in others) across various experimental models. While both p16 and p21 are established markers of cellular senescence, they are implicated in distinct mechanistic pathways. The selective use of these markers may lead to confusion. It is recommended that the authors provide a justification for this experimental design choice to clarify whether it reflects a targeted investigation of specific senescence pathways or a practical consideration based on assay optimization.\n3) In lines 242~246, there appears to be an error in the figure citations. The text references \"Figure 1E~H\" but based on the context and the described content, these panels correspond more appropriately to “Figure 3E~H”. It is likely a typographical error, and the authors are advised to correct the citation to ensure accuracy and proper alignment with the presented data.\n4) In Figure 4F, the Western Blot results for p53, the internal control GAPDH bands appear excessively faint, suggesting potential issues such as insufficient protein loading or suboptimal detection conditions. It is strongly recommended that the authors repeat it with standardized protein quantification and equal loading validation to ensure credible results.\n5) In Figure 7D staining results, the scale bars appear inconsistent. The nuclei in the p21 fluorescence channel are conspicuously larger than those in other channels. please check it.", "reviewer_2": "This study by Lie et al. explores how mechanical stimulation influences the same pathways as senescence or aging. The authors examine both human bone marrow mesenchymal cells and mouse models. The study provides valuable insights, identifying several mechanistic pathways that are regulated in both senescence and mechanical stimulation. The regulation of FOXO1 is particularly exciting.\nHowever, there are several missing elements, such as special controls, that need to be addressed along with other uncertain aspects. A more in-depth investigation of the same models in both older and younger samples is necessary, as well as mechanistic and written analyses, to enhance the translational relevance of the study.\nMajor comments\n1. First, I don’t understand why the authors have only treated the old cells and mice with mechanical stimulation, and only the young with limited mechanical stimulation. I believe it’s an important control, even if negative, to also examine, for example, young mice with mechanotransduction. This needs to be performed, or there should be a very good reason why it isn’t. The whole story for me falls apart on this because I don’t see proper controls, and it’s possible that limited loading is beneficial for the bone and shares the exact mechanism as senescence.\n2. The isolation of BMSCs needs to be explained better. How can you be certain that these cells are the ones you are working with?\n3. Why are all technical replicates displayed in Figures 1 and 2? Should you not just calculate the mean of each technical replicate and then present the acutely different individuals in the Figure? Or it then doesn’t reach significance, which seems hardly unlikely when there are so many individuals, 21+17.\n4. Aging is linked to inflammation. Why not examine specific local markers in BM or cortical bone? For example, RNA might not be as visible, but it needs further investigation for Figures 4 and 8.\n5. BMSCs are mainly found in the BM of the cortical bone. Why did the authors not look at cortical bone? This needs to be included, and differences among various bones should be discussed.\n6. A part about limitations needs to be addressed and included, since there are several limitations within this study.\nMinor comment\n1. You need to explain in a better way how the biochemical shares the same pathways with senescence. I have a problem understanding this when reading the introduction. Also include more references about this connection as well as other things in the introduction. For example, on line 39, This disruption…. this especially refers to BMSC needs references.\n2. You need to do something with figure 2, as it is problematic to understand which illustration is shown in each figure.\n3. Where am I looking in Figures 2D and E? I could not find the information, neither in the results, the figure legend, nor in the materials and methods. The femur is lifted, but not in which part of the femur, the cortical, or the trabecular. Needs to be lifted in MM and either in the results or the figure legend.\n4. In addition, the manuscript is acceptable, but it lacks several details in the results, and I will need to review the MM extensively. Therefore, for clarity, the manuscript would benefit from including more explanation also in the results section.", "reviewer_3": "To the best of my knowledge, this is the first study to clearly delineate how extracellular mechanical cues are transmitted to the chromatin, thereby regulating chromatin accessibility and ultimately mesenchymal stem cell (MSC) aging. Building on this mechanistic discovery, the authors further developed a vibration-based intervention that effectively counteracts bone aging in a simple and translatable manner, highlighting its therapeutic potential. Overall, the work is carefully executed, and the results are convincing. I have only a few minor points that could further improve clarity and presentation:\n1. The Methods section could be reordered to align with the sequence in which the results are presented, which would improve readability and facilitate comprehension.\n2. The description of BMSC isolation is generally clear, and the identification methods are well presented. For completeness, it would be helpful if the authors could specify the anatomical site from which the BMSCs were obtained, as this detail is important for reproducibility and interpretation.\n3. In Figure 7D, the scale bars appear to be inconsistent. For example, the nuclei in the p21 fluorescence channel look noticeably larger than those in the other channels. Double-checking the scaling to ensure consistency across the images would further improve clarity.\n4. A point that could further strengthen the manuscript is the central role of FOXO1. While the data convincingly demonstrate its force-dependent activation, adding more discussion on the potential mechanisms by which FOXO1 regulates cellular senescence would enhance the manuscript.\n5．This study complements Yue et al. (Cell, 2025), who identified a population of force-sensitive cells, and Liu et al. (Cell, 2025), who reported exercise-mediated anti-aging effects. By elucidating the intracellular molecular mechanisms linking mechanical force to aging, the present work provides a valuable counterpart to these recent advances. I recommend that the authors cite both studies and discuss how their findings integrate with these results, thereby placing their work in the broader context of mechanobiology and bone aging."}, "author_response": "Point-to-point response\nReviewer: 1\nReviewer comment:\nIn the manuscript entitled \"Mechanical Rejuvenation of Senescent Stem Cells and\nAged Bone via Chromatin Remodeling\", the authors reported a marked reduction\nin intracellular traction force and cellular mechanics in senescent bone marrow\nmesenchymal stem cells (BMSCs). They demonstrated that the moderate\nmechanical stimuli effectively restored cellular force, enhanced chromatin\naccessibility at the FOXO1 locus, activated its expression, and reversed cellular\nsenescence and bone aging. This study unveils that optimizing mechanical\nstimulation provides a promising, straightforward approach to counteract age-\nrelated bone degeneration and systemic inflammation, offering significant\npotential for clinical translation. The presented findings are new and interesting,\nand reviewer has the following comments.\nResponse: We thank the reviewer for the constructive comments and positive\nrecognition of our work.\n(1) In Fig.1E results, the authors performed Western blot (WB) analysis on several\nkey proteins involved in mechanotransduction. However, given the multitude of\ncellular mechanotransduction pathways, the authors should clarify whether this\ninvestigation focuses on a predefined, specific pathway or represents a broader\nscreening of multiple associated pathways. This clarification would significantly\nenhance comprehension of the research scope and improve the accuracy of the\nstudy.\nResponse: We thank the reviewer for this insightful comment. Our study specifically\nfocused on a well-established and canonical mechanotransduction axis linking\ncytoskeletal contractility to nuclear mechanosignaling, rather than performing a broad,\nunbiased screening of all mechanotransduction pathways. In particular, we examined\nrepresentative components across the key structural and signaling nodes of this cascade:\n1. Nonmuscle Myosin IIa and its phosphorylated form (Ser1943), which\ngenerate intracellular traction forces and serve as indicators of actomyosin\ncontractility;\n2. Lamin A/C, the principal nucleoskeletal component that transmits cytoskeletal\ntension to the nuclear envelope and maintains nuclear mechanical integrity; and\n3. Yes-associated protein (YAP), a force-sensitive transcriptional coactivator,\nwhose nuclear localization provides a downstream readout of mechanical input.\nThis axi, actomyosin cytoskeleton - nucleoskeleton (Lamin A/C) - nuclear\nmechanosignaling (YAP), constitutes a central route by which physical forces are\nconverted into biochemical responses in stromal cells. We have updated the main text\nto clarify that our Western blot analysis was designed to interrogate this classical\nmechanotransduction pathway, rather than serving as a comprehensive screen of all\npossible mechanotransduction networks.\n(2)In Fig.2 staining results, the authors used different senescence markers (p16 in\nsome panels, p21 in others) across various experimental models. While both p16\nand p21 are established markers of cellular senescence, they are implicated in\ndistinct mechanistic pathways. The selective use of these markers may lead to\nconfusion. It is recommended that the authors provide a justification for this\nexperimental design choice to clarify whether it reflects a targeted investigation of\nspecific senescence pathways or a practical consideration based on assay\noptimization.\nResponse: We thank the reviewer for this valuable comment. We fully agree that p16\nand p21 are regulated through distinct mechanistic pathways, p16 primarily reflecting\nRb pathway activation, while p21 is more closely linked to p53-mediated responses.\nIn our study, the use of different markers across experimental models was not intended\nto suggest reliance on a single pathway, but rather to achieve a comprehensive and\ncross-validated assessment of cellular senescence.\nSpecifically, we systematically examined multiple senescence indicators,\nincluding p16, p21, p53, and SA-β-gal, to capture the multifaceted nature of the\nsenescence phenotype. These complementary markers collectively provide a more\nreliable evaluation than any single indicator alone, given the context-dependent\nvariability of individual pathways. The corresponding results for these additional\nmarkers are presented in the Supplementary Fig 9-13, ensuring that our conclusions are\nsupported by consistent and convergent evidence across independent senescence\npathways.\n(3) In lines 242~246, there appears to be an error in the figure citations. The text\nreferences \"Figure 1E~H\" but based on the context and the described content,\nthese panels correspond more appropriately to “Figure 3E~H”. It is likely a\ntypographical error, and the authors are advised to correct the citation to ensure\naccuracy and proper alignment with the presented data.\nResponse: We are grateful to the reviewer for carefully identifying this oversight. The\ncitation was indeed incorrect; the correct reference should be to Figure 3E–H rather\nthan Figure 1E–H. This error has been corrected in the revised manuscript to ensure\naccuracy and consistency throughout the text.\n(4) In Figure 4F, the Western Blot results for p53, the internal control GAPDH\nbands appear excessively faint, suggesting potential issues such as insufficient\nprotein loading or suboptimal detection conditions. It is strongly recommended\nthat the authors repeat it with standardized protein quantification and equal\nloading validation to ensure credible results.\nResponse: We thank the reviewer for this valuable comment. We fully agree that the\nGAPDH bands in Figure 4F appeared faint, likely due to suboptimal exposure during\nthe initial Western blot detection. To address this concern, we have repeated the\nexperiment with standardized protein quantification and validated equal loading across\nall lanes. The updated results are now provided in Supplementary Section 7, Figure 16,\nwhich confirms the reliability of the original findings.\nFig. S16.Senescence-associated markers in trabecular bone and bone marrow.\n(a) Western blot analysis of p53 expression in trabecular bone from aged mice, aged\nmice with vibration, young mice, and young mice with tail suspension. p53 protein\nlevels were decreased in aged mice after vibration and increased in young mice after\ntail suspension (n=3 per group). (b) qPCR analysis of senescence-associated genes p16,\np19, and p21 in bone marrow from the same groups (n=3 per group). Expression levels\ndecreased after vibration in aged mice and increased after tail suspension in young mice.\n(5) In Figure 7D staining results, the scale bars appear inconsistent. The nuclei in\nthe p21 fluorescence channel are conspicuously larger than those in other channels.\nplease check it.\nResponse: We thank the reviewer for carefully noting this issue. We have confirmed\nthat the original description of the scale bars in Figure 7D was incorrect. The figure\nlegend has now been revised to accurately state: “Scale bar, 50 μm for a, b, e, f, g; 10\nμm for d.”\nReviewer: 2\nReviewer comment:\nThis study by Lie et al. explores how mechanical stimulation influences the same\npathways as senescence or aging. The authors examine both human bone marrow\nmesenchymal cells and mouse models. The study provides valuable insights,\nidentifying several mechanistic pathways that are regulated in both senescence\nand mechanical stimulation. The regulation of FOXO1 is particularly exciting.\nHowever, there are several missing elements, such as special controls, that need to\nbe addressed along with other uncertain aspects. A more in-depth investigation of\nthe same models in both older and younger samples is necessary, as well as\nmechanistic and written analyses, to enhance the translational relevance of the\nstudy.\nResponse: We sincerely thank the reviewer for the thoughtful suggestions.\n(1) First, I don’t understand why the authors have only treated the old cells and\nmice with mechanical stimulation, and only the young with limited mechanical\nstimulation. I believe it’s an important control, even if negative, to also examine,\nfor example, young mice with mechanotransduction. This needs to be performed,\nor there should be a very good reason why it isn’t. The whole story for me falls\napart on this because I don’t see proper controls, and it’s possible that limited\nloading is beneficial for the bone and shares the exact mechanism as senescence.\nResponse: We thank the reviewer for emphasizing the importance of including control\nconditions for mechanical interventions. In our main study, we focused on increasing\nmechanical load in aged mice and reducing load in young mice to probe age-dependent\neffects, consistent with our prior in vitro observations of diminished\nmechanotransduction in aged BMSCs.\nTo address the reviewer’s concern, we performed the reverse interventions, aged\nmice subjected to tail suspension (aged_ force−) and young mice receiving vibrational\nloading (young_ force+), and these data are now included in Supplementary Section 7,\nFig. S15. Quantitative micro-CT analyses of trabecular bone (BV/TV, Tb.N, BMD,\nTb.Th, Tb.Sp, Tb.Pf) and cortical bone (Ct.Th, Ct.Ar/Tt.Ar, Ct.Ar) revealed negligible\nchanges relative to age-matched controls. These results confirm that reverse\ninterventions do not significantly alter bone structure, supporting the rationale for our\nprimary experimental design.\nTogether, these data validate our focus on aged mice with vibrational loading and\nyoung mice with tail suspension, demonstrating that the observed effects are age-\ndependent and not confounded by mechanical stimulation alone.\nFig. S15. Micro-CT analysis of trabecular and cortical bone in young and aged\nmice under additional mechanical control conditions. (a) Representative μCT\nreconstructions of distal femoral trabecular bone (up) and mid-diaphyseal cortical bone\n(down). Images show aged controls, aged mice subjected to tail suspension (30\nconsecutive days; force-), young controls, and young mice subjected to vibrational\nloading (20 min/day for 30 days, alternating with rest days; force+). Images are\nrepresentative of n = 5–7 mice per group. Tail suspension in aged mice did not produce\novert deterioration in trabecular or cortical bone compared with aged controls. Similarly,\nvibrational loading in young mice yielded minimal structural changes relative to young\ncontrols. (b) Quantification of trabecular bone parameters in the distal femur, including\nbone volume fraction (BV/TV), trabecular number (Tb.N), bone mineral density\n(BMD), trabecular thickness (Tb.Th), trabecular separation (Tb.Sp), and trabecular\npattern factor (Tb.Pf). Consistent with representative images, force- in aged mice did\nnot exacerbate trabecular bone loss, and force+ in young mice had only marginal effects.\n(c) Quantification of cortical bone parameters at the femoral mid-diaphysis:\ncortical thickness (Ct.Th), cortical area fraction (Ct.Ar/Tt.Ar), and cortical area (Ct.Ar).\nCortical bone architecture in aged mice was not further compromised by tail suspension\n(force-), and vibrational loading (force+) in young mice produced negligible effects.\n(2) The isolation of BMSCs needs to be explained better. How can you be certain\nthat these cells are the ones you are working with?\nResponse:\nWe appreciate the reviewer’s important concern regarding the identification of BMSCs.\nBone marrow aspirates were obtained from the posterior superior iliac spine, which is\na clinically recognized site for bone marrow harvest. Mononuclear cells were isolated\nby density gradient centrifugation, and adherent cells were expanded under standard\nMSC culture conditions. To confirm their identity, we performed flow cytometric\nanalysis showing positive expression of CD73 and CD146, and negative expression of\nhematopoietic/endothelial markers CD45 and CD34. In addition, functional validation\nwas carried out. The isolated cells displayed colony-forming unit, robust proliferation\nas demonstrated by EdU incorporation and crystal violet staining, and multilineage\ndifferentiation into osteoblasts and adipocytes. Given the focus of our study, we also\nconfirmed age-related functional alterations by SA-β-gal staining, p21 immunostaining,\nand γ-H2AX detection (Supplementary Section 2, Fig. 2-5). Together, these phenotypic,\nfunctional, and senescence-related assays provide strong evidence that the cells used in\nthis study are bona fide bone marrow-derived mesenchymal stem/stromal cells\n(BMSCs).\n(3) Why are all technical replicates displayed in Figures 1 and 2? Should you not\njust calculate the mean of each technical replicate and then present the acutely\ndifferent individuals in the Figure? Or it then doesn’t reach significance, which\nseems hardly unlikely when there are so many individuals, 21+17.\nResponse: We thank the reviewer for raising the concern regarding individual donor\nvariability. As many of our assays required a relatively large number of cells, it was not\nfeasible to obtain sufficient cells from a single donor, therefore, in the previous\nexperiments, cells within the same group were pooled from multiple donors of similar\nage. In the previous version, we had already examined P-myosin IIa levels in BMSCs\nfrom 10 young donors (18–35 years) and 10 elderly donors (50–80 years), and observed\nconsistent age-dependent differences in force-related signaling across donors\n(Supplementary Section1; Section 3, Fig. 6a).\nFollowing the reviewer’s important suggestion, we have now performed a more\nsystematic set of independent biological replicate experiments using BMSCs isolated\nseparately from these 10 young and 10 elderly donors. For each donor, multiple cells\n(≥50) were analyzed, and the mean value was treated as one biological replicate. To\nfurther strengthen the analysis, Lamin A/C was included as an additional marker of\nmechanical force. Consistent age-dependent differences were observed across all\ndonors, confirming that these molecular phenotypes are driven by donor age rather than\nindividual variability (Supplementary Section 3, Fig. 6b).\nFurthermore, mechanosensitive functional assays were also independently repeated\nfor biological replicates: elderly donor-derived BMSCs subjected to cyclic stretch\n(increased force) exhibited suppressed P16 expression, whereas young donor-derived\nBMSCs cultured on soft hydrogels (reduced force) showed increased P16 expression\n(each data point represents one donor, n = 10 per group) (Supplementary Section 3, Fig.\n6c). These results confirm reproducible age-dependent mechanosensitive responses.\nAfter establishing consistent age-related phenotypes, cells from donors within each\nage group were pooled for experiments requiring large cell numbers (Fig.1-2). Pooling\nwas only performed after confirming that age-dependent differences were consistent\nacross individual donors, ensuring that all downstream observations reflect age-\nassociated mechanobiological changes rather than donor-specific variability.\nFig. S6. Donor variation in mechanotransduction and force responsiveness of\nhuman BMSCs. (a) Immunofluorescence (IF) quantification of phosphorylated\nMyosin IIa (P-myosin IIa). (b) IF quantification of Lamin A/C. Elderly donor-derived\nBMSCs exhibited markedly lower P-myosin IIa and Lamin A/C levels compared with\nyoung donor-derived cells. (c) Assessment of cellular senescence by P16\nimmunofluorescence under altered mechanical conditions: elderly donor-derived\nBMSCs subjected to cyclic stretch (enhanced force) displayed reduced P16 expression,\nwhereas young donor-derived BMSCs cultured on soft hydrogels (reduced force)\nshowed increased P16 expression. Each data point represents the mean IF intensity from\none donor (n = 10 per group).\n(4) Aging is linked to inflammation. Why not examine specific local markers in\nBM or cortical bone? For example, RNA might not be as visible, but it needs\nfurther investigation for Figures 4 and 8.\nResponse: We thank the reviewer for raising the point regarding local inflammatory\nmarkers in bone marrow (BM) and trabecular/cortical bone. To address this, we\nperformed both transcriptional and histological analyses in our study. In Figure 4e,\nqPCR analysis of BM from aged mice revealed that vibrational loading decreased\nexpression of IL-1β, IL-6, CXCL1, and CCL2, whereas tail suspension in young mice\nincreased these cytokines. Complementary immunofluorescence analysis of trabecular\nbone (Fig. 4f) showed that TNF-α⁺, CD105⁺ cells were reduced after vibration in aged\nmice and elevated after tail suspension in young mice (n = 5 per group).\nSupplementary data further support these findings. In Supplementary Section 7, Fig.\nS17, BM qPCR analysis demonstrated decreased IL-1α and TNF-α following vibration\nin aged mice and increased expression after tail suspension in young mice, while\nimmunohistochemistry of trabecular bone confirmed corresponding changes in CD45\nand IL-1β staining.\nMoreover, Supplementary Section 10, Fig. S24 extends these observations by\nshowing that intermittent vibration in aged mice reduced senescence markers (p16, p21)\nand inflammatory cytokines (IL-1β, CXCL1, CCL2, TNF-α), whereas continual\nvibration promoted their expression in bone marrow.\nTogether, these data demonstrate that mechanical interventions modulate local\ninflammatory and senescence-associated pathways within BM and bone tissue,\nproviding direct evidence that tissue-resident cells respond to mechanical cues in an\nage- and context-dependent manner.\nFig. 4 | Mechanical Stimulation Modulates Bone Aging, Inflammation, and\nRemodeling in vivo.\n(e) qPCR analysis of inflammatory cytokines IL-1β, IL-6, CXCL1, and CCL2 in bone\nmarrow. Expression decreased after vibration in aged mice and increased after tail\nsuspension in young mice. (f) Representative double immunofluorescence images of\nCD105 and TNF-α in trabecular bone. Quantification of TNF-α⁺ CD105⁺ cells shows a\ndecrease after vibration in aged mice and an increase after tail suspension in young mice\n(n=5)\nFig. S17. Inflammatory markers in bone marrow and trabecular bone.\n(a) qPCR analysis of inflammatory cytokines IL-1α and TNF-α in bone marrow from\naged mice, aged mice with vibration, young mice, and young mice with tail suspension\n(n=5 per group). In aged mice, vibration reduced IL-1α and TNF-α expression\ncompared with aged controls. In young mice, tail suspension increased IL-1α and TNF-\nα levels. (b) Immunohistochemistry images of CD45 and IL-1β in trabecular bone from\nthe same groups (n=5 per group). In aged mice, vibration decreased CD45 and IL-1β\nstaining. In young mice, tail suspension increased CD45 and IL-1β staining. Scale bars:\n20 μm.\nFig. S24. qPCR analysis of senescence- and inflammation-associated gene\nexpression in bone marrow following vibrational loading.\n(a) Relative mRNA expression of senescence markers (p16 and p21) in bone marrow\nfrom aged mice subjected to intermittent or continual vibration treatment for 30 days,\nas assessed by qPCR. Intermittent vibration reduced expression of senescence markers,\nwhereas continual vibration increased their levels. (b) Relative mRNA expression of\ninflammatory cytokines and chemokines (IL-1β, CXCL1, CCL2, and TNF-α) in bone\nmarrow from the same groups. Intermittent vibration reduced inflammatory gene\nexpression, whereas continual vibration promoted their expression.\n(5) BMSCs are mainly found in the BM of the cortical bone. Why did the authors\nnot look at cortical bone? This needs to be included, and differences among\nvarious bones should be discussed.\nResponse: We thank the reviewer for the comment regarding the assessment of cortical\nbone, as BMSCs are predominantly localized within the bone marrow of cortical bone.\nTo address this, we performed micro-CT analysis of the femoral midshaft to evaluate\ncortical bone architecture (Supplementary Section 7, Fig. S18). Representative μCT\nreconstructions and quantitative analysis demonstrated that, in aged mice, vibrational\nloading had minimal impact on cortical thickness (Ct.Th), cortical area fraction\n(Ct.Ar/Tt.Ar), total cross-sectional area (Tt.Ar), cortical area (Ct.Ar), and marrow area\n(Ma.Ar) compared with age-matched controls. Similarly, in young mice, tail suspension\ndid not significantly alter any of these cortical parameters relative to young controls (n\n= 5 per group).\nThese results indicate that, under the current experimental conditions, cortical bone\nstructure is largely preserved and responds less robustly to short-term mechanical\ninterventions than trabecular bone. This finding complements our analyses of trabecular\nbone and bone marrow, providing a more comprehensive understanding of bone tissue\nresponses to mechanical cues across different bone compartments.\nFig. S18. Cortical bone architecture in the femoral midshaft assessed by micro-CT.\n(a) Representative μCT reconstructions of cortical bone at the femoral mid-diaphysis\nfrom aged controls, aged mice subjected to vibrational loading (20 min/day for 30 days,\nalternating with rest days), young controls, and young mice subjected to tail suspension\n(30 consecutive days) (n = 5 per group). (b) Quantitative analysis of cortical bone\nparameters. For aged mice, vibrational loading had negligible effects on cortical\nthickness (Ct.Th), cortical area fraction (Ct.Ar/Tt.Ar), total cross-sectional area (Tt.Ar),\ncortical area (Ct.Ar), and marrow area (Ma.Ar) compared with aged controls. For young\nmice, tail suspension did not significantly alter any of these cortical parameters relative\nto young controls (n = 5 per group).\n(6) A part about limitations needs to be addressed and included, since there are\nseveral limitations within this study.\nResponse: We have addressed the reviewer’s comment by including a discussion of\nthe study’s limitations and future directions in the Discussion section (see revised\nmanuscript).\nLimitations and future directions: Despite these advances, several limitations remain.\nIn vivo measurement of cellular forces remains technically challenging, with existing\nmethods offering limited spatiotemporal resolution; developing high-precision, high-\nresolution techniques will be essential for future studies. While FOXO1 is identified as\na key mediator linking mechanical cues to cellular aging, the downstream epigenetic\nand transcriptional mechanisms remain incompletely understood. The functional\nrelevance of FOXO1-bound target genes to the aging process warrants further\nsystematic investigation. Finally, although this study focused on BMSCs and bone\ntissue, determining whether targeted mechanical stimulation can similarly rejuvenate\nother cell populations or tissues will broaden the translational potential of this approach.\nMinor comment\n(1) You need to explain in a better way how the biochemical shares the same\npathways with senescence. I have a problem understanding this when reading the\nintroduction. Also include more references about this connection as well as other\nthings in the introduction. For example, on line 39, This disruption…. this\nespecially refers to BMSC needs references.\nResponse: We thank the reviewer for raising this important point. We have revised the\nIntroduction to clarify how biochemical signaling pathways converge on BMSC\nsenescence. Specifically, we now describe that senescent BMSCs undergo broad\ntranscriptional reprogramming, particularly in genes governing proliferation,\ndifferentiation, and extracellular matrix organization. These changes are tightly\nregulated by multiple biochemical processes: metabolic dysregulation (e.g., impaired\nmitochondrial function, altered NAD⁺/SIRT axis) perturbs redox balance and energy\nhomeostasis, activating stress-responsive pathways; transcriptional regulators such as\np53, FOXO, and NF-κB orchestrate the expression of senescence-associated genes,\nlinking cellular stress to growth arrest and the senescence-associated secretory\nphenotype (SASP); and epigenetic modifications, including histone acetylation,\nmethylation, and DNA methylation, consolidate the senescent state by repressing\nproliferation-related genes and altering chromatin accessibility. Collectively, these\nbiochemical processes converge to establish a stable, self-reinforcing senescence\nprogram. This revision emphasizes that different biochemical signals act through\ninterconnected pathways to drive BMSC aging, providing a clearer mechanistic context\nfor the study.\n(2) You need to do something with figure 2, as it is problematic to understand\nwhich illustration is shown in each figure.\nResponse: We thank the reviewer for pointing out the ambiguity in Figure 2. We have\nrevised the figure to clearly indicate which panels correspond to each experimental\ncondition and marker. The updated version improves clarity and ensures that the data\nare easily interpretable. The revised figure is shown below.\nFig. 2 | Cellular mechanotransduction regulates cell senescence and cell\nproliferation. (a) Senescent and young BMSCs were subjected to mechanical\ninterventions to alter intracellular traction forces. In senescent cells, traction force was\nincreased using either mechanical stretching (5% strain, 0.02 Hz) or calyculin A\ntreatment (myosin light chain phosphatase inhibitor), resulting in a significant reduction\nof p16INK4a expression. In young cells, traction force was reduced using either soft\nhydrogels (1.5 kPa) or blebbistatin treatment (selective myosin II inhibitor), which led\nto elevated p16INK4a expression (n = 50, three technical replicates). (b) Proliferation\nanalysis using Ki67 immunostaining and EdU incorporation. Senescent BMSCs were\nsubjected to enhanced traction force via mechanical stretching, while young BMSCs\nwere exposed to reduced traction force on soft hydrogels (1.5 kPa). Representative\nimages and quantification (n = 6–8 images per condition, three technical replicates)\nare shown. (c-d) Mechanotransduction and senescence analysis in 3D bone marrow-\nmimicking microenvironments. Senescent and young BMSCs were cultured in (c)\nGelMA hydrogels or (d) ECM-mimicking 3D fiber networks (Young’s modulus: 1–4\nkPa). Cellular traction forces were pharmacologically modulated: senescent BMSCs\nwere treated with calyculin A to enhance force, whereas young BMSCs were treated\nwith blebbistatin to reduce force. Immunofluorescence analysis of p21 and Lamin A/C\nwas performed to assess cellular senescence and mechanotransduction activity,\nrespectively (n = 35 cells per condition, three technical replicates). Data are presented\nas mean ± SD. Scale bar, 10 μm for p21 in e, f, 50 μm for others.\n(3) Where am I looking in Figures 2D and E? I could not find the information,\nneither in the results, the figure legend, nor in the materials and methods. The\nfemur is lifted, but not in which part of the femur, the cortical, or the trabecular.\nNeeds to be lifted in MM and either in the results or the figure legend.\nResponse: We thank the reviewer for this comment. We have clarified Figures 2D and\n2E to ensure that the data being presented are unambiguous. The relevant information\nregarding what is shown in each panel is now explicitly described in the Results section,\nthe figure legend, and the Materials and Methods, ensuring full clarity and\nreproducibility.\n(4) In addition, the manuscript is acceptable, but it lacks several details in the\nresults, and I will need to review the MM extensively. Therefore, for clarity, the\nmanuscript would benefit from including more explanation also in the results\nsection.\nResponse: We thank the reviewer for this suggestion. We have revised the Results\nsection to include additional explanations and clarifications, enhancing the clarity and\ninterpretability of the data throughout the manuscript.\nReviewer: 3\nReviewer comment:\nTo the best of my knowledge, this is the first study to clearly delineate how\nextracellular mechanical cues are transmitted to the chromatin, thereby\nregulating chromatin accessibility and ultimately mesenchymal stem cell (MSC)\naging. Building on this mechanistic discovery, the authors further developed a\nvibration-based intervention that effectively counteracts bone aging in a simple\nand translatable manner, highlighting its therapeutic potential. Overall, the work\nis carefully executed, and the results are convincing. I have only a few minor points\nthat could further improve clarity and presentation:\nResponse: We sincerely thank the reviewer for the encouraging comments.\n(1) The Methods section could be reordered to align with the sequence in which\nthe results are presented, which would improve readability and facilitate\ncomprehension.\nResponse: We appreciate the reviewer’s constructive suggestion. In the revised\nmanuscript, we have reorganized the Methods section to follow the same order as the\ncorresponding Results section. This restructuring ensures a more coherent flow and\nimproves readability, allowing readers to more easily connect each methodological\ndescription with the related experimental findings.\n(2) The description of BMSC isolation is generally clear, and the identification\nmethods are well presented. For completeness, it would be helpful if the authors\ncould specify the anatomical site from which the BMSCs were obtained, as this\ndetail is important for reproducibility and interpretation.\nResponse: We thank the reviewer for this helpful suggestion. In the revised manuscript,\nwe have added this information to the Methods section. Specifically, we now state that\nhuman bone marrow aspirates were collected from the posterior superior iliac spine.\nThis clarification ensures reproducibility and provides important anatomical context for\nthe isolation of BMSCs.\n(3) In Figure 7D, the scale bars appear to be inconsistent. For example, the nuclei\nin the p21 fluorescence channel look noticeably larger than those in the other\nchannels. Double-checking the scaling to ensure consistency across the images\nwould further improve clarity.\nResponse: Thank you for bringing this to our attention. We carefully re-examined\nFigure 7D and confirmed that the apparent difference was due to an error in the\noriginally indicated scale bars rather than an actual discrepancy in image magnification.\nThe figure has been corrected, and the legend now accurately reads: “Scale bar, 50 μm\nfor a, b, e, f, g; 10 μm for d.” This correction ensures consistency and clarity across\nall panels.\n(4) A point that could further strengthen the manuscript is the central role of\nFOXO1. While the data convincingly demonstrate its force-dependent activation,\nadding more discussion on the potential mechanisms by which FOXO1 regulates\ncellular senescence would enhance the manuscript.\nResponse: We thank the reviewer for this constructive suggestion. In response, we have\nexpanded the Discussion to elaborate on the mechanistic role of FOXO1 in regulating\ncellular senescence. Specifically, we now highlight that FOXO1, a transcription factor\nwith established roles in oxidative stress resilience, DNA repair, apoptosis, and\nmetabolic regulation, functions as a central effector translating mechanical cues into\nanti-senescent gene expression programs. This addition emphasizes how FOXO1\nintegrates mechanical inputs with cellular stress-response and longevity pathways,\nthereby reinforcing its pivotal role in maintaining genomic stability and delaying\nsenescence. The revised discussion provides a more comprehensive mechanistic\ncontext for FOXO1’s contribution to force-dependent regulation of aging.\n(5) This study complements Yue et al. (Cell, 2025), who identified a population of\nforce-sensitive cells, and Liu et al. (Cell, 2025), who reported exercise-mediated\nanti-aging effects. By elucidating the intracellular molecular mechanisms linking\nmechanical force to aging, the present work provides a valuable counterpart to\nthese recent advances. I recommend that the authors cite both studies and discuss\nhow their findings integrate with these results, thereby placing their work in the\nbroader context of mechanobiology and bone aging.\nResponse: We appreciate the reviewer’s valuable suggestion. In accordance with the\ncomment, we have revised the Discussion to incorporate and contextualize the recent\nfindings of Liu et al. (Cell, 2025) and Yue et al. (Cell, 2025) within the broader\nframework of our study. Specifically, we now state that:\n“The physiological relevance of mechanotransduction in systemic aging is further\nsupported by recent clinical and experimental evidence. Liu et al. (Cell, 2025) showed\nthat exercise elevates systemic anti-senescent and anti-inflammatory factors across\nmultiple tissues, rescuing age-associated phenotypes, while Yue et al. (Cell, 2025)\nidentified a population of force-sensitive progenitor cells that mediate cartilage\nrejuvenation and suppress osteoarthritis progression. Together, these studies\nsubstantiate that appropriately applied mechanical cues confer anti-aging benefits at\nboth the cellular and tissue levels. Nonetheless, interindividual variability in senescence\nstatus and mechanosensitivity presents challenges for standardizing mechanical\ninterventions, emphasizing the importance of precision mechanotherapy for\ntranslational applications.”\nPoint-to-point response\nReviewer 2\nComment:\nFirstly, I would like to emphasize the great work the authors have done in the\nrevision process.\n(1) Regarding comment 1. Removing the force in aged mice, as well as inducing\nforce in young mice, has no effect. This is great, now shown in the supplement\nFigure. But I don't understand why you have compared these young and aged mice\nto each other. Possible to compare aged-young with the same force, but otherwise\nremove this comparison and possibly put in two figures.\nResponse：We thank the reviewer for this insightful comment and appreciate the careful\nattention to this issue. We agree that aged and young mice should not be directly\ncompared under different mechanical conditions, as these control experiments were\ndesigned solely to evaluate the effects of reduced force in aged mice and increased force\nin young mice within each respective age group. In accordance with the reviewer’s\nsuggestion, we have reorganized the Supplementary Figures to present the data\nseparately:\n Panels a–b now include only trabecular and cortical bone analyses from aged\ncontrol and aged_ force– mice.\n Panels c–d now include the corresponding analyses from young control and\nyoung_ force+ mice.\nThis revised structure removes any unintended cross-age comparisons and ensures\nthat each age group is evaluated exclusively relative to its own mechanical condition.\nWe believe this adjustment substantially improves clarity and fully addresses the\nreviewer’s concern.\nFig. S15. Micro-CT analysis of trabecular and cortical bone in aged and young\nmice under additional mechanical control conditions.\n(a) Representative μCT reconstructions of distal femoral trabecular bone from four\ngroups: aged control, aged tail suspension (force–), young control, and young\nvibrational loading (force+). (b) Quantitative analysis of trabecular microarchitecture\nin the four groups, including bone volume fraction (BV/TV), trabecular number (Tb.N),\nbone mineral density (BMD), trabecular thickness (Tb.Th), trabecular separation\n(Tb.Sp), and trabecular pattern factor (Tb.Pf). Tail suspension did not further\nexacerbate trabecular deterioration in aged mice, whereas vibrational loading induced\nminimal changes in young mice. (c) Representative μCT reconstructions of cortical\nbone at the femoral mid-diaphysis from the same four groups. (d) Quantitative analysis\nof cortical parameters (cortical thickness, Ct.Th; cortical area fraction, Ct.Ar/Tt.Ar;\ncortical area, Ct.Ar) in the four groups. Cortical bone architecture remained largely\nunchanged under either mechanical unloading in aged mice or vibrational stimulation\nin young mice. n = 5 biologically independent mice per group, Two-sided Student’s t-\ntest with adjustment for multiple comparisons. Exact P values and sample sizes (n) are\nprovided in the figure. All raw data and uncropped scans are provided in the Source\nData file.\n(2) I have a hard time discerning what has actually been changed in the\nmanuscript and what is merely explained to me. Please ensure that my questions\nand concerns are not only for my benefit, but also that they are addressed in the\nmanuscript so that the reader can gain a deeper understanding.\nResponse：We sincerely apologize for the lack of clarity in our previous response. In\nthe previous and current revised manuscripts, all the points you raised were carefully\nincorporated. We have clarified the relevant sections to ensure that readers can directly\nfollow the experimental rationale, results, and mechanistic insights. These revisions aim\nto improve the overall readability and logical flow of the manuscript, so that the content\nis fully self-contained and accessible without referring to the response letter.\n(3) I only have one new question, which is whether the effects are mainly\ndependent on bone building (osteoblasts) or decreasing bone resorption\n(osteoclasts). This could be checked, just as the immune parameters in the RNA or\nserum of the mice, using PINP or CTX. Perhaps no difference is available since\nthe effect began a long time ago, and now there's a new balance. However, I believe\nthis is beneficial for the study, particularly in identifying which part is secondarily\naffected in Figure 4.\nResponse：We thank the reviewer for this important question and for the helpful\nsuggestion regarding the assessment of bone formation versus resorption. To address\nthis, we performed additional analyses: bone marrow RNA was used to measure\nosteoblast-related genes (Col1a1, Bglap) and osteoclast-related genes (Mmp9, Acp5),\nand serum was collected to quantify bone turnover markers (PINP for bone formation\nand CTX for bone resorption).\nOur results indicate that mechanical manipulation affects both osteoblast and\nosteoclast markers. However, the changes are more pronounced in osteoblast-related\ngenes and in the serum PINP levels, suggesting that the observed effects are primarily\nmediated through modulation of osteoblast activity, with secondary effects on\nosteoclast function. These findings provide additional mechanistic insight into the\neffects illustrated in Figure 4 and strengthen the conclusion that mechanical forces\npredominantly influence bone formation.\nFigure S19. Mechanical loading predominantly affects osteoblast activity in aged\nand young mice.\n(a) qPCR analysis of bone marrow gene expression for osteoblast markers (Col1a1,\nBglap) and osteoclast markers (Mmp9, Acp5) in aged mice with increased mechanical\nloading and young mice with reduced loading (n = 3 biologically independent mice per\ngroup). Mechanical manipulation impacts both osteoblast and osteoclast markers, with\na stronger effect on osteoblast-associated genes. (b) Serum bone turnover markers PINP\n(formation) and CTX (resorption) in the same groups (n = 5 biologically independent\nmice per group). Consistent with gene expression, mechanical loading primarily\nmodulates bone formation. One-way ANOVA followed by Tukey’s post hoc test (two-\nsided) was used, and adjustment for multiple comparisons was applied. Exact P values\nand sample sizes (n) are provided. All raw data supporting the findings of this study are\nprovided in the Source Data file."}, {"round": 2, "reviewer_comments": {"reviewer_1": "the revision has addressed the reviewer's comments. reviewer has no further comment", "reviewer_2": "Firstly, I would like to emphasize the great work the authors have done in the revision process.\nRegarding comment 1. Removing the force in aged mice, as well as inducing force in young mice, has no effect. This is great, now shown in the supplement Figure. But I don't understand why you have compared these young and aged mice to each other. Possible to compare ageed-young with the same force, but otherwise remove this comparison and possibly put in two figures.\nI have a hard time discerning what has actually been changed in the manuscript and what is merely explained to me. Please ensure that my questions and concerns are not only for my benefit, but also that they are addressed in the manuscript so that the reader can gain a deeper understanding.\nI only have one new question, which is whether the effects are mainly dependent on bone building (osteoblasts) or decreasing bone resorption (osteoclasts). This could be checked, just as the immune parameters in the RNA or serum of the mice, using PINP or CTX. Perhaps no difference is available since the effect began a long time ago, and now there's a new balance. However, I believe this is beneficial for the study, particularly in identifying which part is secondarily affected in Figure 4.", "reviewer_3": "The authors have satisfactorily addressed all of my concerns. I recommend acceptance."}, "author_response": ""}]

Decision letter