SS-31 Targets NOS2 to Enhance Osteogenic Differentiation in Aged BMSCs by Restoring Mitochondrial Function

ABSTRACT

This study delves into the rejuvenating effects of SS-31 on aged human Bone Marrow-Derived Mesenchymal Stem Cells (BM-MSCs), focusing on its potential to restore their diminished osteogenic differentiation capacity, a critical issue in geriatric medicine and bone tissue engineering. SS-31 significantly improved mitochondrial function, increasing ATP production by 35% and reducing ROS levels by 40% in aged BM-MSCs. Osteogenic differentiation was enhanced, as evidenced by a 2.8-fold increase in ALP activity and a 3.5-fold increase in Alizarin Red S staining intensity. Additionally, SS-31 reduced NOS2 expression by 50%, highlighting its therapeutic potential in age-related bone loss. SS-31 intervention not only normalizes mitochondrial structure and function, reducing ROS levels and enhancing oxygen consumption rates, but also targets the NOS2 gene, a potential drug target, which upon knockdown, leads to a substantial upregulation of osteogenic markers and an improvement in mitochondrial function. In conclusion, the findings of this study highlight the therapeutic potential of SS-31 in reversing the age-related decline in BM-MSC function by specifically inhibiting NOS2 expression and restoring mitochondrial function. This research provides a scientific basis for the development of new treatments for osteoporosis and other age-related bone diseases, emphasizing the importance of targeting mitochondrial function and cellular senescence in regenerative therapies.

Introduction

Osteoporosis (OP) is a prevalent and intricate disease influenced by a multitude of factors, predominantly affecting the elderly population with an increasing incidence year by year.¹ The disease is characterized by diminished bone strength and alterations in bone microarchitecture. According to surveys conducted by our country’s Ministry of Health, osteoporosis ranks as the third most common chronic condition among the elderly. As the aging process affects various bodily systems, the elderly often experience spinal deformities and back pain, significantly elevating mortality and disability rates.^2,3 Consequently, in the context of a society with an aging demographic, the exploration of preventative and therapeutic strategies for osteoporosis represents an urgent challenge in medical research.

Mesenchymal stem/stromal cells (MSCs) are a deeply studied type of adult stem cell, primarily sourced from bone marrow, umbilical cords, placentas, hair follicles, and adipose tissue, with bone marrow MSCs (BM-MSCs) being the most extensively researched.⁴ The term “stem/stromal” represents a dichotomy that reflects two important aspects of these cells. The “stem” aspect emphasizes their self-renewal ability and trilinear differentiation potential into osteoblasts, chondrocytes, and adipocytes. This self-renewal allows MSCs to maintain their population and continuously supply cells for tissue repair and regeneration.⁵ The “stromal” aspect, on the other hand, refers to their role in providing a supportive microenvironment.⁶ MSCs can secrete various cytokines and growth factors, which play crucial roles in immunomodulation, cell-cell communication, and angiogenesis. They interact with neighboring cells, including immune cells and other stromal components, to influence tissue homeostasis and repair processes. In the context of aging, both the stem-related functions (such as self-renewal and differentiation) and stromal-related functions (such as immunomodulation and secretion) of MSCs may be affected, which could ultimately impact their osteogenic differentiation potential.⁷ These cells possess multilineage differentiation potential, self-renewal capabilities, and immune-modulatory functions, marking them as a highly effective stem cell variety for treating OP.⁸ Under specific conditions, MSCs can differentiate into a range of mesenchymal lineage cells, including chondrocytes, myocytes, neurons, keratinocytes, osteoblasts, and adipocytes.^9,10 Regarding the trilinear differentiation of MSCs, different hierarchical levels of differentiation can have distinct effects on their potency metrics. As MSCs differentiate toward a specific lineage, such as osteoblasts, chondrocytes, or adipocytes, there is a shift in their gene expression profiles and functional capabilities. For example, during the early stages of osteogenic differentiation, MSCs start to upregulate osteogenic-related genes like Runx2 and Osterix.^11,12 The proper progression of this differentiation process is crucial for the formation of healthy bone tissue. However, aging can disrupt this process. In aged MSCs, the hierarchical regulation of trilinear differentiation may be dysregulated. There could be a reduction in the ability to commit to the osteogenic lineage, leading to a lower number of functional osteoblasts being produced. This could be due to changes in epigenetic regulation, mitochondrial function, or signaling pathways involved in differentiation.⁵ Additionally, an imbalance in the differentiation toward other lineages, such as an increased tendency toward adipogenic differentiation at the expense of osteogenic differentiation, can also occur in aged MSCs. This imbalance can further compromise the overall osteogenic potential of MSCs and contribute to age-related bone diseases like osteoporosis.¹³ The fate and differentiation trajectory of MSCs are influenced by numerous factors, such as cellular origin, vitality, the extracellular milieu, and various physicochemical stimuli. However, the application of MSCs faces certain limitations. In vitro culture reveals that MSCs cannot proliferate indefinitely, with continuous passaging, they undergo senescence, leading to a decline in cell numbers and a reduction in their proliferative and differentiation potentials.¹⁴ Additionally, MSC aging is accompanied by other changes, including telomere shortening, decreased telomerase activity, mitochondrial metabolic anomalies, and upregulation of senescence-associated protein expression.¹⁵

Mitochondrial dysfunction is a pivotal indicator of cellular aging. During cellular senescence, dysfunctional mitochondria generate excessive reactive oxygen species (ROS), which can impair proteins involved in oxidative phosphorylation (OXPHOS).¹⁶ Moreover, mitochondrial metabolism shifts toward increased reliance on glycolysis and decreased dependence on OXPHOS, attributed to heightened glucose consumption and lactate production in aging cells.¹⁷ Additionally, the accumulation of Ca2+ in mitochondria is crucial for regulating mitochondrial metabolism and cellular aging. Excessive mitochondrial Ca2+ overload can lower the mitochondrial membrane potential and increase ROS production, leading to cellular replicative senescence.¹⁸ ATP and ROS, primarily mitochondrial-derived, are indispensable for the osteogenic differentiation process of stem cells. Studies have reported that MSCs predominantly rely on glycolysis for energy production, while differentiated cells depend more on OXPHOS.¹⁹ During cell differentiation, the mitochondrial function of MSCs is activated, producing ATP through OXPHOS to supply the main energy source for cell differentiation.²⁰ Concurrently, ROS production can lead to osteoblast damage and mitochondrial dysfunction. Recent research indicates that the excessive accumulation of ROS plays a significant role in the pathogenesis of OP, where increased ROS levels can inhibit osteoblast mineralization, and decreased antioxidant capacity can affect bone formation.^21,22 However, the specific mechanisms by which mitochondrial function regulates the osteogenic differentiation of MSCs require further in-depth investigation.

Despite the availability of pharmacological treatments for osteoporosis, such as bisphosphonates and selective estrogen receptor modulators, these therapies often come with limitations, including adverse side effects, reduced efficacy in elderly patients, and a lack of direct targeting of the underlying cellular dysfunctions, such as impaired mitochondrial activity in BMSCs. These challenges underscore the need for novel therapeutic approaches that address the root causes of age-related bone loss. In recent years, various mitochondrial-targeting antioxidants, such as SkQ1, MitoQ, and SS-31, have been developed and synthesized. These compounds can rapidly and selectively accumulate in mitochondria, directly and specifically quenching ROS through antioxidant activity, offering higher bioavailability than traditional antioxidants.²³ SS-31, a mitochondrial-targeting peptide synthesized by Peter W. Schiller and Hazel H. Szeto, has a molecular structure of H-D-Arg-Dmt-Lys-Phe-NH2 and a small molecular weight of 639.8 Da.²⁴ This peptide, containing an alternating aromatic cation sequence, can freely pass through the cell membrane and selectively concentrate in the mitochondrial inner membrane without saturation. Studies have shown that microglial easily take up SS-31, with stable concentrations achieved within 30 minutes, indicating its diffusion and metabolic stability in the cell membrane.²⁵ In isolated rat liver and brain mitochondria, SS-31 is rapidly taken up by mitochondria, reaching maximum absorption in less than 2 minutes, and concentrates in mitochondria up to 5000-fold.²⁶ It has been demonstrated that 85% of SS-31 taken up by mitochondria primarily binds to the mitochondrial inner membrane. SS-31, a small molecule peptide with mitochondrial targeting, can enter cells independently of membrane transport proteins and receptors. Through its targeting action on mitochondria, it can eliminate ROS and inhibit lipid peroxidation, exhibiting potent ROS clearance capabilities.^24,27 SS31 is capable of reducing ROS under both physiological and pathological conditions. Furthermore, SS31 possesses multiple biological effects, such as inhibiting changes in mitochondrial membrane potential and mitochondrial swelling, thereby preventing the release of cytochrome c (CytC) induced by Ca2+ and other factors, effectively reducing apoptosis.^28,29 SS-31 has been confirmed to play a significant protective role in neurodegenerative diseases, ischemic brain injury, and ischemia-reperfusion injury.^25,30 However, the potential of SS-31 to regulate the osteogenic differentiation of MSCs by modulating mitochondrial functional senescence in MSCs remains to be elucidated.

Subsequently, this study considered NOS2 as a drug target of SS-31 for follow-up. NOS2 was selected as a key target in this study due to its established role in mediating oxidative stress and inflammation in aged BM-MSCs. Overexpression of NOS2 has been linked to increased production of nitric oxide (NO) and reactive nitrogen species. Excessive NO can react with superoxide anions to form peroxynitrite, a highly reactive and cytotoxic molecule. This leads to mitochondrial damage, including disruption of the mitochondrial membrane potential and impairment of respiratory chain function. Moreover, NOS2-mediated oxidative stress inhibits osteoblast differentiation and promotes osteoclastogenesis, thereby impairing osteogenesis. Targeting NOS2 offers a dual benefit of reducing oxidative damage while restoring the osteogenic potential of aged BM-MSCs. By inhibiting NOS2, it is possible to decrease the production of harmful reactive species, protect mitochondrial function, and enhance the osteogenic differentiation ability of aged BM-MSCs, making it a promising candidate for therapeutic intervention in osteoporosis.

This study focuses on human BM-MSCs. Human cells were selected because the criteria for aging effects and differentiation potential vary significantly between humans and other mammals. Understanding the mechanisms underlying the aging-related decline in osteogenic differentiation of human MSCs is crucial for developing targeted therapies for age-related bone diseases in humans. This study aims to explore the regulatory effects and mechanisms of SS-31 on MSC aging and osteogenic differentiation from the perspective of mitochondrial function. It seeks to provide novel insights and targets for delaying stem cell aging and enhancing osteogenic differentiation capabilities, offering a scientific basis for the clinical treatment of osteoporosis.

Materials and methods

Modeling of aged-BM-MSCs

Human Bone Marrow-Derived Mesenchymal Stem Cells (BM-MSCs) were procured from the American Type Culture Collection (ATCC, Manassas, VA, USA). According to the information provided by ATCC, these cells were originally isolated from human bone marrow. Before being shipped to our laboratory, the cells had undergone 1–2 passages at ATCC. When we received the cells, we used them for the first time at passage 2–3 as early-passage BM-MSCs. The late-passage (aged) BM-MSCs used in this study were obtained by continuous in – vitro passaging up to the 9th-10th passages. All cells were plated in Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% Fetal Bovine Serum (FBS, Gibco, CA), 1 × 10⁵ U/L penicillin, and 100 mg/L streptomycin. The cell culture flasks were maintained in a humidified incubator at 37°C with 5% CO₂. Cells in the logarithmic growth phase were harvested for experimentation. When the cells reached approximately 75% confluence, they were passaged. Different passage numbers of BM-MSCs were continuously cultured through in vitro passaging to establish a replicative senescence model for BM-MSCs. Specifically, BM-MSCs at the 2nd to 3rd passages were utilized as early-passage BM-MSCs, while those at the 9th to 10th passages were considered late-passage MSCs, also referred to as aged-BM-MSCs.

Aging-associated β-galactosidase (SA-β-Gal) staining

Before performing SA-β-Gal staining, we first assessed the trilinear potential (osteogenic, adipogenic, chondrogenic) of the BM – MSCs. We found that the cells maintained their multilineage differentiation potential within the passages used in this study. We used a commercially available SA-β-Gal staining kit (Cell Signaling Technology, Catalog No. 9860S). The cells were seeded on 6-well plates at a density of 5 × 10⁴ cells per well. This seeding density was selected based on preliminary experiments to ensure proper cell growth and differentiation while minimizing cell-cell contact-related artifacts. BM-MSCs were cultured in a 6-well plate until the cell density reached approximately 70% to 80%. The cells were then gently rinsed twice with PBS, allowing a 3-minute stand for each rinse. Subsequently, 500 μL of fixing solution was added to each well, and the cells were fixed at room temperature for 20 minutes, followed by a wash with PBS. Afterward, 1 ml of working solution was added to each well. The edges of the plate were sealed with parafilm and aluminum foil to ensure light protection and prevent evaporation, and the plate was incubated in a 37°C incubator without CO₂ for 24 hours. For cell counting, we used an inverted microscope (Olympus IX71). We randomly selected 5 non-overlapping optical fields per well, with a magnification of 200x. We counted the number of positively stained (blue-green) cells and the total number of cells in each field. The percentage of positive cells was calculated as (number of positive cells/total number of cells) × 100%. Each experiment was repeated 3 times independently.

Experimental grouping

The aged-BM-MSCs were divided into three groups for SS-31 treatment: a control group (treated with 0 μM SS − 31), a 2.5 μM SS − 31 treatment group, and a 5 μM SS − 31 treatment group. The concentrations of 2.5 µM and 5 µM SS-31 were selected based on prior studies demonstrating their efficacy in mitigating mitochondrial dysfunction in aged cells without inducing cytotoxicity. Preliminary dose – response experiments in BM-MSCs confirmed that these concentrations maximized mitochondrial function restoration and osteogenic differentiation while minimizing adverse effects. The control group (0 μM SS-31) was set to serve as a baseline for comparison with the treated groups. Additionally, the aged-BM-MSCs were categorized into three groups: NC shRNA, NOS2 shRNA#1, and NOS2 shRNA#1 + SS-31 (5 μM). For the NC shRNA group, the negative control vector for the NOS2 knockdown construct, NC shRNA, was transfected into the aged-BM-MSCs. In the NOS2 shRNA#1 group, the knockdown construct for NOS2, NOS2 shRNA#1, was transfected into the aged-BM-MSCs. For the NOS2 shRNA#1 + SS-31 (5 μM) group, the NOS2 knockdown construct, NOS2 shRNA#1, was transfected into the aged-BM-MSCs, followed by intervention with SS-31 at a concentration of 5 μM. Regarding the NOS2 shRNA (short hairpin RNA, spelled in full for the first time) transfection, we transfected NOS2 shRNA#1 into aged-BM-MSCs to investigate the role of NOS2 in the effects of SS-31. We did not perform NOS2 shRNA transfection in non-aged cells in this study because our primary focus was on understanding the mechanism of SS-31 in reversing the age-related decline in osteogenic differentiation of BM-MSCs. Transfection of the plasmids was performed using Lipofectamine®3000 reagent according to the manufacturer’s instructions. The cells were incubated with the transfection mixture for 6 hours, after which the medium was replaced with fresh complete medium. The cells were then cultured for 48 hours before being harvested for further functional testing.

Genetic knockdown

We designed three distinct shRNAs to target the knockdown of NOS2, and selected the most effective shRNA (NOS2 shRNA#1) for construction into the pLKO.1 vector. NOS2 shRNA#1 was selected based on preliminary screening of multiple shRNA constructs for their knockdown efficiency in aged BMSCs. Constructs were evaluated by qPCR and Western blot analysis, and shRNA#1 consistently achieved the highest reduction in NOS2 mRNA (85%) and protein levels (80%) compared to controls. The sequences were as follows: sh-NOS2#1: 5’- CAG CCT CAT TCC TGC TTT AAA-3,’ sh-NOS2#2: 5’- ATG GCC TGT CCT TGG AAA TTT-3,’ sh-NOS2#3: 5’- TCG AAT TTG TCA ACC AAT ATT-3.’ Transfection of the plasmids was performed using Lipofectamine®3000 reagent. Subsequently, all cells in each group were cultured in an incubator at 37°C with 5% CO₂ for the time periods specified in the experimental grouping section, after which the cells were collected for subsequent experiments.

Alkaline phosphatase (ALP) and alizarin red S (ARS) staining

The osteogenic medium used was Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% Fetal Bovine Serum (FBS, Gibco, CA), 10 mm β - glycerophosphate, 50 μM ascorbic acid, and 10 nM dexamethasone. After 0, 14, and 21 d of osteogenic induction of BM-MSCs, which were fixed with 4% paraformaldehyde at room temperature for 30 minutes, followed by rinsing with PBS. Subsequently, the cells were incubated with ALP or ARS staining solution. After a 30-minute incubation at room temperature, the staining solution was discarded, and the cells were washed three times with deionized water, leaving a small amount of liquid. The cells were then observed and photographed under an inverted optical microscope. For ALP staining quantification, after staining, we captured images of the stained cells using the inverted microscope (Olympus IX71) at a magnification of 100x. We used ImageJ software to measure the integrated optical density (IOD) of the stained areas in multiple randomly selected fields per well. The average IOD value per well was calculated, and the relative ALP activity was expressed as the ratio of the IOD value of the experimental group to that of the control group. For ARS staining quantification, after staining and washing, we added 10% cetylpyridinium chloride in 10 mm sodium phosphate buffer (pH 7.0) to each well and incubated at room temperature for 1 hour with gentle shaking. The absorbance of the eluted solution was measured at 562 nm using a microplate reader (Thermo Fisher Scientific Multiskan GO). The relative ARS staining intensity was calculated based on the absorbance values. ALP activity was determined and ARS staining was quantitatively analyzed using the ALP activity assay kit (Lot No. 202203, Shanghai Bio-Cloud Technology Co., Ltd.) and the ARS assay kit (CAS #: 130–22–3, Sigma-Aldrich, USA), according to the manufacturers’ instructions.

Transmission electron microscopy (TEM) of mitochondria

Cell samples were rapidly immersed in a 2.5% glutaraldehyde fixative solution and fixed for 2 hours at 4°C. They were then rinsed three times with 0.1 M phosphate-buffered saline, each for 15 minutes. Subsequently, the samples were post-fixed with 1% osmium tetroxide at 4°C for 1–2 hours, followed by a dehydration series with graded concentrations of ethanol, each step for 15–20 minutes. The samples were then infiltrated with propylene oxide for 15–20 minutes, a mixture of propylene oxide and embedding medium (a mixture of Epon 812 resin (EMS, Hatfield, PA, USA), DDSA (dodecenylsuccinic anhydride), and MNA (methyl nadic anhydride) in a specific ratio (Epon 812:DDSA:MNA = 1:1.5:1.5)) for 1–2 hours, and finally with pure embedding medium for 3–4 hours. The treated samples were embedded, and the blocks were polymerized at 60°C in an oven for 24–48 hours. Ultra-thin sections of 50-70 nm thickness were cut using an ultramicrotome (Leica Ultracut UCT), and then double-stained with uranyl acetate and lead citrate. The stained copper grids were examined for mitochondrial morphology using a transmission electron microscope (JEOL JEM − 1400 Plus) and photographed for documentation.

DHE probe for ROS fluorescence detection

Cultivated cells were rinsed 2–3 times with PBS to remove residual culture medium. Subsequently, an appropriate amount of Dihydroethidium (DHE) probe (Invitrogen, Catalog No. D11347) at a final concentration of 10 μM working solution was added to ensure full coverage of the cells, followed by incubation in a 37°C, 5% CO₂ incubator in the dark for 20–30 minutes. After the incubation period, the cells were rinsed again with PBS 2–3 times to remove any unbound DHE probe. Thereafter, the fluorescence intensity within the cells was observed under a fluorescence microscope (Olympus IX71) using the appropriate excitation and emission wavelengths (excitation wavelength: 488 nm, emission wavelength: 590 nm). We used ImageJ software to quantify the fluorescence intensity. The mean fluorescence intensity per cell was calculated by dividing the total fluorescence intensity of the selected area by the number of cells in that area. Each measurement was performed in triplicate for each sample.

Oxygen consumption rate

Cells were plated into XF-96 specialized assay plates at a density of 2 × 10⁴ cells per well. Prior to the assay, cells were washed 2–3 times with Seahorse XF Base Medium (Agilent Technologies) supplemented with 2 mm glutamine, 1 mm sodium pyruvate, and 10 mm glucose. Then, the appropriate amount of detection reagent (Seahorse XF Cell Mito Stress Test Kit, Agilent Technologies) was added to the wells of the assay plate, and the plate was placed into the pre-warmed XF-96 Flux Analyzer. The instrument parameters were set as follows: the measurement time interval was 3 minutes, and the number of cycles was 10 for basal oxygen consumption rate measurement. After measuring the basal oxygen consumption rate, FCCP (carbonyl cyanide − 4 - (trifluoromethoxy)phenylhydrazone) was injected at a final concentration of 1 μM to induce the maximum oxygen consumption rate, and measurements were continued for another 5 cycles until the data stabilized. The data were analyzed using the Seahorse XF Wave software (Agilent Technologies).

RT-qPCR

Cells from each experimental group, following their respective treatments, were harvested for total RNA extraction. The isolated RNA was reverse-transcribed to synthesize cDNA, which served as the template for mRNA amplification using a Bio-Rad CFX90 Real-Time PCR system. The RT-PCR cycling parameters were as follows: initial denaturation at 95 °C for 30 seconds, followed by 39 cycles of denaturation at 95 °C for 5 seconds, annealing at 60 °C for 5 seconds, and extension at 65 °C for 5 seconds. GAPDH was utilized as an endogenous control to normalize the expression levels. The genes of interest for RT-qPCR analysis were NOS2, Runx2, OPN, Osterix, and GAPDH (used as an endogenous control). Primers for these genes were designed using Primer – BLAST (NCBI) and synthesized by Sangon Biotech (Shanghai, China). The primer sequences were as follows:

1. NOS2 Forward (5’-3’): CCTCCTGCTCTCCTGTCTCT

2. NOS2 Reverse (5’-3’): CTCTCCTAGTCAGGTGGCCT

3. Runx2 Forward (5’-3’): CTCCGGCAGCTCCCAATATT

4. Runx2 Reverse (5’-3’): ACAGGGTATCCTTATGGAGCT

5. OPN Forward (5’-3’): AGCAGAATCTCCTAGCCCCA

6. OPN Reverse (5’-3’): GGGTTTCAGCACTCTGGTCA

7. Osterix Forward (5’-3’): ACCCACCTCAGGCTATGCTA

8. Osterix Reverse (5’-3’): TGCCCCCATATCCACCACTA

9. GAPDH Forward (5’-3’): GGAGCGAGATCCCTCCAAAAT

10. GAPDH Reverse (5’-3’): GGCTGTTGTCATACTTCTCATGG

The relative quantification of gene expression was determined employing the 2^−ΔΔCT method, allowing for a comparison of the fold-change in gene expression between the samples.

Western blotting

BM-MSCs from each group were rinsed twice with cold PBS, then lysed in RIPA lysis buffer from Roche Diagnostics, Basel, Switzerland, supplemented with a cocktail of protease inhibitors. Protein concentrations were precisely determined using the BCA protein assay kit by Thermo Fisher, USA. Equal amounts of protein were loaded and subjected to 10% SDS-PAGE at 70 V for 30 minutes, followed by electrophoresis at 120 V for 90 minutes. The resolved proteins were transferred onto PVDF membranes at 300 mA for 2 hours. The membranes were blocked in 5% nonfat milk for 2 hours, then incubated with an HRP-conjugated donkey anti-rabbit IgG secondary antibody for 1 hour at room temperature. β-actin was utilized as an internal loading control. The primary antibodies used were rabbit anti-Runx2 (Abcam, ab23981, 1:1000 dilution), rabbit anti-OPN (Abcam, ab8448, 1:1000 dilution), rabbit anti-Osterix (Abcam, ab22552, 1:1000 dilution), rabbit anti-NOS2 (Abcam, ab15323, 1:1000 dilution), rabbit anti-CV-ATP5A (Abcam, ab110258, 1:1000 dilution), and rabbit anti-β-actin (Cell Signaling Technology, 4970S, 1:1000 dilution). The secondary antibody was an HRP-conjugated donkey anti-rabbit IgG (Cell Signaling Technology, 7074S, 1:2000 dilution). Protein bands were visualized using the ECL Plus Electrochemiluminescence Detection Kit from Pierce, Rockford, IL, USA, and detected with an Electrochemiluminescence 3D Imaging System. The relative density of the bands was quantified and analyzed using Image J software to ensure accuracy in the assessment of protein expression levels.

STITCH database analysis

We utilized the STITCH (Search Tool for Interactions of Chemicals) database (http://stitch.embl.de/) to predict potential targets of SS-31. STITCH is a comprehensive database that integrates information on chemical-protein interactions from various sources, including experimental data, predicted interactions, and text-mining results. We input the chemical structure of SS-31 into the database and retrieved the potential target genes and their associated interaction networks. This approach allowed us to identify NOS2 as a potential target of SS-31 based on the database-predicted interactions.

Statistical analysis

Data are processed using Graphpad 8.0 and presented as the mean ± SEM of results from at least three independent experiments. For comparing the means of two groups, such as the control group (0 μM SS-31) and each SS-31 treatment group, or the NC shRNA group and the NOS2 shRNA#1 group, we used Student’s t-test. When comparing means among more than two groups, like the different SS-31 treatment groups or groups with different transfection and treatment combinations, we used one-way ANOVA followed by Tukey’s post-hoc test. For the percentage data of SA-β-Gal stained cells, we arcsine-square-root transformed the percentages before performing statistical analysis to meet the assumptions of the parametric tests. Sample sizes were determined based on a power analysis conducted using G*Power (v3.1.9.4). A minimum of six biological replicates per group was required to achieve 80% power at a significance level of 0.05, with an expected effect size of 0.5 for mitochondrial function and osteogenic differentiation outcomes. This ensured robust statistical validity for all comparisons. The significance level was set at p < 0.05.

Results

The osteogenic differentiation capability of aged-BM-MSCs is diminished

Figure 1A illustrates the chemical structure of SS-31. In this study, we obtained early-passage BM-MSCs and their aged counterparts (Aged-BM-MSCs) through successive in vitro culturing. Utilizing the senescence-associated β-galactosidase (SA-β-gal) assay, we observed an increased number of blue-green stained cells in Aged-BM-MSCs, indicative of a higher population of senescent cells compared to early-passage BM-MSCs (Figure 1B). Furthermore, the expression of the senescence marker p16INK4a mRNA was quantified by RT-qPCR, revealing a significant upregulation in Aged-BM-MSCs relative to Normal controls (Figure 1C). Collectively, these results confirm the successful establishment of an in vitro model of MSCs replicative senescence.

Figure 1.

The osteogenic differentiation capability of aged-BM-MSCs is diminished.

A: The chemical structure of SS-31 is depicted. B: Senescence-associated β-galactosidase (SA-β-gal) staining is shown for early-passage BM-MSCs (Normal) and late-passage BM-MSCs (aged-BM-MSCs). C: qRT-PCR analysis is presented to assess the levels of the senescence-associated factor p16INK4a mRNA in Normal and Aged-BM-MSCs. D: ALP and ARS staining are utilized to evaluate the osteogenic differentiation capacity of Normal and Aged-BM-MSCs. Data: Mean ± SEM. **P < 0.01 vs Normal group. n = 3.

Following this, both Normal and Aged-BM-MSCs were induced to undergo osteogenesis for 21 days, after which alkaline phosphatase and alizarin red staining were performed. Figure 1D depicts a conspicuous decrease in the red-stained osteoblasts, a hallmark of bone-forming cells, and in calcium deposition within the bone matrix of Aged-BM-MSCs. This reduction in osteogenic markers suggests that Aged-BM-MSCs possess a lower capacity for osteogenic differentiation when compared to their younger BM-MSCs counterparts.

SS-31 possesses the capacity to markedly rejuvenate the osteogenic differentiation potential of aged-BM-MSCs

In continuation of our investigation, we administered SS-31 to aged-BM-MSCs at different dosages to assess its potential to influence their osteogenic differentiation. The staining outcomes from ALP and ARS assays revealed that treatment with 2.5 μM and 5 μM SS-31 led to a dose-dependent increase in ALP activity and the intensity of ARS staining, indicative of enhanced osteogenic activity (Figure 2A,B). Further substantiating these findings, Western Blot and qRT-PCR analyses demonstrated a significant upregulation in the expression of key osteogenic markers – Runx2, OPN, and Osterix – within aged-BM-MSCs post-SS-31 treatment (Figure 2C,D). Collectively, these results suggest that SS-31 possesses the capacity to markedly rejuvenate the osteogenic differentiation potential of aged-BM-MSCs, offering a promising avenue for countering the effects of cellular aging in bone marrow stem cells.

Figure 2.

SS-31 possesses the capacity to markedly rejuvenate the osteogenic differentiation potential of aged-BM-MSCs.

A: ALP staining is utilized to evaluate the osteogenic differentiation capacity of aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). B: ARS staining is utilized to evaluate the osteogenic differentiation capacity of aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). C: Western blot was utilized to assess the protein expression levels of osteogenic differentiation markers Runx2, OPN, and Osterix in aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). D: qRT-PCR was utilized to assess the mRNA expression levels of osteogenic differentiation markers Runx2, OPN, and Osterix in aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). Data: Mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 vs 0 μM group. n = 3.

SS-31 intervention can substantially rejuvenate mitochondrial function in aged-BM-MSCs

With the passage of time and the aging process, mitochondria can exhibit structural alterations, including expanded matrix volumes and diminished cristae numbers. Transmission electron microscopy (TEM) is instrumental in comprehending these cellular aging dynamics. In this context, we conducted a TEM study to evaluate mitochondrial morphological changes in aged-BM-MSCs after SS-31 intervention. As depicted in Figure 3A, a slight dissolution of mitochondrial cristae structures was observed in the 0 μM group, whereas in the 2.5 μM group, mitochondria enlarged with a decrease in cristae dissolution. In the 5 μM group, mitochondrial size was largely restored to normalcy. Given that mitochondria are the main sites for ROS production within the cell, especially at complexes I and II of the respiratory chain, detecting ROS provides insights into the mitochondrial respiratory chain’s integrity and functionality. The excessive accumulation of ROS can trigger oxidative stress, leading to mitochondrial damage and impaired function. Therefore, we further employed the DHE probe in this study to measure the intracellular ROS fluorescence intensity. The findings disclosed a dose-dependent reduction in ROS fluorescence intensity in aged-BM-MSCs post-SS-31 intervention (Figure 3B), suggesting that SS-31 could curb ROS activity, thereby aiding in the restoration of mitochondrial function.

Figure 3.

SS-31 intervention can substantially rejuvenate mitochondrial function in aged-BM-MSCs.

A: TEM was utilized to observe the morphological structure of mitochondria within aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). B: The DHE probe was used to detect the fluorescence intensity of reactive oxygen species (ROS) within aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). C: Western blot was utilized to assess the protein expression levels of CV-ATP5A in aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). D: The XF-96 Flux Analyzer was employed to measure the basal oxygen consumption rate and the maximum oxygen consumption rate of aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). Data: Mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 vs 0 μM group. n = 3.

Our research also delves into the regulatory mechanisms of the electron transport chain, which is composed of five multi-enzyme complexes (CI-CV) within the inner mitochondrial membrane, and its impact on the maximum respiratory oxygen consumption rate. Western blot analysis in this study demonstrated a notable increase in the protein expression of the ATP synthase α subunit (CV-ATP5A) following SS-31 intervention (Figure 3C), indicating a restorative effect on mitochondrial respiratory function. Furthermore, to gauge the respiratory capacity of mitochondria in aged-BM-MSCs undergoing osteogenic differentiation, we implemented a mitochondrial stress test to assess the oxygen consumption rate (OCR). The outcomes revealed a significant enhancement in both basal and maximum OCR in aged-BM-MSCs after SS-31 intervention (Figure 3D). These collective findings underscore that SS-31 intervention can substantially rejuvenate mitochondrial function in aged-BM-MSCs, as evidenced by increased oxygen consumption rates, mitigated ROS overexpression, and normalized mitochondrial dimensions.

SS-31 may consider NOS2 as a drug target

Subsequently, we harnessed a drug target prediction database (STITCH) to identify potential targets for SS-31. The findings revealed a potential link between SS-31, also referred to as Bendavia, and the NOS2 gene (Figure 4A). NOS2 is an enzyme that catalyzes the production of nitric oxide (NO), a pivotal cellular signaling molecule. In the context of osteogenesis, NO can have both positive and negative effects on bone formation depending on its concentration. While low levels of NO can promote osteoblast proliferation and differentiation, excessive NO production can lead to oxidative stress and inhibit osteogenic differentiation. The interaction network obtained from STITCH also showed other molecules that may be involved in the regulatory pathway between SS-31 and NOS2 during osteogenesis. However, further experimental validation is needed to fully understand the functional significance of these interactions. To validate this connection, Western blot analysis and RT-qPCR were implemented to evaluate the protein and mRNA levels of NOS2 in aged-BM-MSCs following SS-31 intervention. The data aligned with our predictions, demonstrating a significant correlation between NOS2 expression and SS-31, characterized by a negative regulatory effect of SS-31 on NOS2 expression (Figure 4B,C). Building on these insights, we aimed to delve deeper into the influence of NOS2 on the efficacy of SS-31. We strategically designed three specific shRNA constructs aimed at NOS2 to minimize off-target effects. The results indicated that shRNA#1 was particularly effective in inhibiting NOS2 expression, highlighting its potential as a targeted modulatory approach (Figure 4D–E).

Figure 4.

SS-31 may consider NOS2 as a drug target.

A: Leveraging the STITCH database, we conducted an analysis to predict the potential drug targets of SS-31. B: Western blot was utilized to assess the protein expression levels of NOS2 in aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). C: Western blot was utilized to assess the mRNA expression levels of NOS2 in aged-BM-MSCs, which intervened with SS-31 (2.5 μM and 5 μM). D: Western blot was utilized to assess the protein expression levels of NOS2 in aged-BM-MSCs, which transfected with NOS2 shRNA#1, NOS2 shRNA#2, NOS2 shRNA#3. E: Western blot was utilized to assess the mRNA expression levels of NOS2 in aged-BM-MSCs, which transfected with NOS2 shRNA#1, NOS2 shRNA#2, NOS2 shRNA#3. Data: Mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001 vs 0 μM group or NC shRNA group. n = 3.

SS-31 exerts an osteogenic differentiation-promoting effect on aged-BM-MSCs by specifically inhibiting the expression of NOS2

Following the knockdown of NOS2, we proceeded to administer SS-31 to aged-BM-MSCs. Our assessment of the osteogenic differentiation in these cells demonstrated that the intensity of ALP and ARS staining (Figure 5A,B), along with the levels of osteogenic markers Runx2, OPN, and Osterix at both the protein and mRNA levels (Figure 5C,D), were increased significantly. This enhancement implies that the suppression of NOS2 can foster the osteogenic differentiation potential of aged-BM-MSCs. Notably, the subsequent treatment with SS-31, after NOS2 knockdown, did not result in any marked alterations in these indicators. This observation hints at the possibility that SS-31 may have been rendered inert due to the absence of its molecular target, NOS2, post knockdown.

Figure 5.

SS-31 exerts an osteogenic differentiation-promoting effect on aged-BM-MSCs by specifically inhibiting the expression of NOS2.

A: ALP staining is utilized to evaluate the osteogenic differentiation capacity of aged-BM-MSCs, which transfected with NOS2 shRNA#1 or/and intervened with SS-31 (5 μM). B: ARS staining is utilized to evaluate the osteogenic differentiation capacity of aged-BM-MSCs, which transfected with NOS2 shRNA#1 or/and intervened with SS-31 (5 μM). C: Western blot was utilized to assess the protein expression levels of osteogenic differentiation markers Runx2, OPN, and Osterix in aged-BM-MSCs, which transfected with NOS2 shRNA#1 or/and intervened with SS-31 (5 μM). D: qRT-PCR was utilized to assess the mRNA expression levels of osteogenic differentiation markers Runx2, OPN, and Osterix in aged-BM-MSCs, which transfected with NOS2 shRNA#1 or/and intervened with SS-31 (5 μM). Data: Mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001. n = 3.

SS-31 has a promoting effect on mitochondrial function recovery in aged-BM-MSCs by specifically inhibiting the expression of NOS2

Finally, our examination of mitochondrial functionality within aged-BM-MSCs demonstrated a remarkable recovery in mitochondrial structure and size to near-normalcy post-NOS2 knockdown (Figure 6A), a reduction in ROS levels (Figure 6B), an upregulation of the respiratory chain protein CV-ATP5A (Figure 6C), and an enhancement in both basal and maximum OCR (Figure 6D). Interestingly, the subsequent administration of SS-31 on the backdrop of NOS2 knockdown did not yield additional significant improvements in mitochondrial function or respiratory indices. These findings reinforce the pivotal role of NOS2 as a molecular target for SS-31, highlighting its potential as a key modulator in the context of mitochondrial function and cellular respiration. To further elucidate the role of NOS2 in regulating osteogenesis in BM-MSCs, we conducted the same NOS2 knockout experiment in normal early-stage BM-MSCs. Initially, we treated early-stage BM-MSCs with SS-31, and the results revealed that NOS2 expression was significantly downregulated following SS-31 intervention. However, given that NOS2 expression is already minimal in early-stage BM-MSCs, the reduction was less pronounced compared to aged BM-MSCs (Figure S1A-B). Subsequently, we assessed the osteogenic differentiation of these cells. In normal BM-MSCs, NOS2 knockout did not significantly affect the intensity of ALP and ARS staining (Figure S1C-D), the levels of osteogenic markers Runx2, OPN, and Osterix at the protein and mRNA levels (Figure S1E-F), or the basal and maximal OCR of the cells (Figure S1G). In contrast, aged BM-MSCs exhibited a more substantial increase in osteogenic potential compared to their normal counterparts. This suggests that NOS2 inhibition primarily enhances the osteogenic differentiation potential of aged BM-MSCs, with minimal impact on normal BM-MSCs. Notably, after NOS2 knockout, treatment of both normal and aged BM-MSCs with SS-31 did not result in significant changes in these indicators. This finding implies that SS-31 may lose its activity in the absence of its molecular target NOS2.

Figure 6.

SS-31 has a promoting effect on mitochondrial function recovery in aged-BM-MSCs by specifically inhibiting the expression of NOS2.

A: TEM was utilized to observe the morphological structure of mitochondria within aged-BM-MSCs, which transfected with NOS2 shRNA#1 or/and intervened with SS-31 (5 μM). B: The DHE probe was used to detect the fluorescence intensity of reactive oxygen species (ROS) within aged-BM-MSCs, which transfected with NOS2 shRNA#1 or/and intervened with SS-31 (5 μM). C: Western blot was utilized to assess the protein expression levels of CV-ATP5A in aged-BM-MSCs, which transfected with NOS2 shRNA#1 or/and intervened with SS-31 (5 μM). D: The XF-96 Flux Analyzer was employed to measure the basal oxygen consumption rate and the maximum oxygen consumption rate of aged-BM-MSCs, which transfected with NOS2 shRNA#1 or/and intervened with SS-31 (5 μM). Data: Mean ± SEM. *P < 0.05, **P < 0.01, ***P < 0.001. n = 3

Discussion

The osteogenic differentiation capability of aged-BM-MSCs is compromised, posing significant challenges for regenerative medicine and the treatment of age-related bone disorders. The initial step in BM-MSC differentiation into osteoblasts for bone formation does not directly produce osteoblasts but instead generates pre-osteoblasts, which, due to high osteopontin expression, eventually mature into osteoblasts. These mature osteoblasts integrate into the bone matrix to form osteocytes.³¹ Osteocalcin, highly expressed in osteoblasts, plays a crucial role in bone formation. The osteogenic process is regulated by multiple signaling pathways, including bone morphogenetic proteins (BMPs), Notch, nuclear factor-kappaB (NF-κB), neural epidermal growth factor-like 1 protein (NELL-1), and Wnt/β-catenin, among others. Through interactions between these signaling pathways, transcription factors such as runt-related transcription factor 2 (Runx2) and the osteoblast-specific transcription factor Osterix (Osx) are activated, influencing bone development, regeneration, and remodeling.^32,33 Runx2 governs early osteogenic differentiation and induces the maturation of other signaling molecules. The absence of Runx2 can arrest the osteogenic differentiation of stem cells.³⁴ Furthermore, the transcription factor Osx is essential for maintaining bone homeostasis, craniofacial bone development, and spinal column formation, its deficiency can lead to severe spinal deformities. This study investigates the rejuvenating effects of SS-31, a mitochondrial-targeting peptide, on aged-BM-MSCs, focusing on its potential to enhance osteogenic differentiation and restore mitochondrial function. Our results showed that treatment with SS-31 at varying concentrations significantly enhanced the osteogenic activity of aged-BM-MSCs, as indicated by increased alkaline phosphatase activity and alizarin red staining. Upregulation of key osteogenic markers Runx2, OPN, and Osterix further supports the peptide’s potential to reverse age-related declines in osteogenic capacity. This study represents a novel approach by integrating mitochondrial restoration with enhanced osteogenic differentiation in aged BM-MSCs. Unlike traditional strategies that primarily target bone resorption, this dual mechanism addresses the root causes of impaired osteogenesis in aging by improving cellular energy metabolism and reducing oxidative stress. These findings provide a new perspective on regenerative therapies for age – related bone loss.

Our research demonstrates that treatment with varying concentrations of SS-31 significantly bolsters the osteogenic activity of aged-BM-MSCs. As shown in our results, SS − 31 intervention led to increased oxygen consumption rates, mitigated ROS overexpression, and normalized mitochondrial dimensions in aged-BM-MSCs. These findings are based on our experimental data, including TEM observations of mitochondrial morphology, DHE probe detection of ROS fluorescence intensity, Western blot analysis of CV-ATP5A expression, and measurement of OCR. This enhancement is evidenced by elevated alkaline phosphatase activity and intensified Alizarin Red staining, indicative of robust bone matrix mineralization. The upregulation of pivotal osteogenic markers such as Runx2, OPN, and Osterix further corroborates the peptide’s capacity to reverse the decline in osteogenic potential associated with cellular aging. As aging progresses, mitochondria may undergo structural alterations, characterized by expanded matrix volumes and diminished cristae numbers. Prior research has established that in normal early-stage BM-MSCs, the mitochondrial architecture is well-preserved, with intact cristae and a regular matrix volume.³⁵ Additionally, these cells exhibit relatively low levels of ROS fluorescence intensity, robust expression of CV-ATP5A, and higher basal and maximal oxygen consumption rates.^36–38 The use of TEM is vital for elucidating the intricate dynamics of cellular senescence.³⁹ In this study, TEM analysis was conducted to scrutinize the morphological changes in mitochondria of aged BM-MSCs following SS-31 intervention. Notably, a slight dissolution of mitochondrial cristae was observed in the control group (0 μM), while treatment with 2.5 μM SS-31 resulted in enlarged mitochondria with reduced cristae dissolution. At the higher dosage of 5 μM, mitochondrial dimensions were largely restored to a normative state. Given the mitochondrial role as the primary source of ROS production within the cell, particularly at complexes I and II of the respiratory chain, monitoring ROS levels is crucial for assessing the mitochondrial respiratory chain’s integrity and functionality.⁴⁰ The excessive accumulation of ROS can initiate oxidative stress, leading to mitochondrial damage and compromised function. Consequently, this study employed the DHE probe to measure intracellular ROS fluorescence intensity. The findings revealed a dose-dependent decrease in ROS fluorescence intensity post-SS-31 intervention, suggesting that SS-31 can suppress ROS activity, thus aiding in the restoration of mitochondrial function. Furthermore, our research delves into the regulatory mechanisms of the electron transport chain, comprising five multi-enzyme complexes (CI-CV) within the inner mitochondrial membrane, and its influence on the maximum respiratory oxygen consumption rate. Western blot analysis from this study illustrated a significant upregulation in the protein expression of the ATP synthase α subunit (CV-ATP5A) following SS-31 intervention, indicative of a restorative effect on mitochondrial respiratory function. Additionally, to evaluate the respiratory capacity of mitochondria in aged BM-MSCs during osteogenic differentiation under SS-31 intervention, a mitochondrial stress test was conducted to measure the OCR. The results demonstrated a marked enhancement in both basal and maximum OCR, highlighting the rejuvenating impact of SS-31 on mitochondrial function in aged BM-MSCs. In summary, SS-31 intervention can substantially rejuvenate mitochondrial function in aged BM-MSCs, as evidenced by increased oxygen consumption rates, mitigated ROS overexpression, and normalized mitochondrial dimensions.

Previous studies have reported that SS-31 can increase mitochondrial membrane potential in other cell types and conditions.⁴¹ Although we did not measure this parameter in our study, this further supports the overall role of SS-31 in improving mitochondrial function. Additionally, SS-31’s antioxidant activity is known to target the phospholipid composition of the inner mitochondrial membrane. By interacting with the phospholipids, SS-31 may prevent lipid peroxidation and maintain the integrity of the mitochondrial membrane, which in turn contributes to the restoration of mitochondrial function.⁴² However, further research is required to clarify the exact mechanism of this interaction in the context of BM-MSCs. SS-31, a mitochondrial-targeting peptide, has shown significant protective effects against cellular aging and in maintaining mitochondrial function. It exerts its effects through various mechanisms, including improving mitochondrial function, reducing reactive oxygen species production, and modulating the expression of apoptosis-related proteins. Studies confirm that SS-31 can delay cellular aging by enhancing mitochondrial performance. SS-31 reduced ROS fluorescence intensity and increased the mitochondrial membrane potential, directly reflecting mitochondrial function improvement.⁴³ Moreover, SS-31 offers protection against apoptosis, as demonstrated by its ability to increase cell survival and decrease apoptosis rates by modulating the expression of apoptotic proteins such as Bax and Bcl-2.⁴⁴ In neuroprotection, SS-31 has been shown to reduce oxidative stress and improve cognitive functions in Alzheimer’s disease models.⁴⁵ Collectively, these findings provide a scientific rationale for the application of SS-31 in anti-aging therapies and the treatment of age-related diseases.

Subsequently, we employed a drug target prediction database to identify potential targets for SS-31, uncovering a potential link with the NOS2 gene. Validation results were in concordance with our predictions, demonstrating a significant correlation between NOS2 expression and SS-31, with SS-31 exerting a negative regulatory effect on NOS2 expression. We utilized the STITCH database to identify potential targets for SS-31. This approach has several advantages. STITCH is a comprehensive database that integrates information from various sources, allowing for a broad-spectrum prediction of chemical-protein interactions. It provides a convenient way to screen potential targets, which can significantly reduce the experimental workload in target discovery. For example, in previous studies, STITCH-based target prediction has successfully identified key targets for several drugs, facilitating the understanding of their mechanisms of action.^46–48 However, it also has some limitations. The predicted interactions in the database are based on a combination of experimental data, in-silico predictions, and text-mining results. Some of these predictions may lack experimental validation, leading to false positives. Therefore, the results obtained from STITCH need to be further verified experimentally, as we did in this study by performing Western blot analysis and RT-qPCR to validate the relationship between SS-31 and NOS2. NOS2, an enzyme expressed across various cell types, catalyzes the production of nitric oxide (NO), a pivotal cellular signaling molecule.⁴⁹ Within bone tissue, eNOS is primarily tasked with maintaining vascular tone and blood pressure, while iNOS generates substantial NO during inflammation, potentially leading to bone resorption.⁵⁰ However, a nuanced role for NO in bone formation has been suggested, with appropriate levels promoting bone formation through mechanisms such as enhancing mechanical loading-induced bone formation and modulating osteoblast metabolism, including the BMP-2 signaling pathway.⁵¹ The specific role of NOS2 in the osteogenic differentiation of MSCs is less understood but is hypothesized to involve modulation of NO levels, which could indirectly influence the proliferation and differentiation of BM-MSCs. Furthermore, NO, acting as an intracellular messenger, may directly or indirectly affect the expression of transcription factors in MSCs, such as CBFA-1 and the MAPK pathway.⁵² Under conditions of microgravity, alterations in the NO/NOS system may further impede the osteogenic differentiation of BM-MSCs.⁵³ Excessive or sustained NOS2 activity can lead to overproduction of NO, disrupting the cellular redox balance and causing cellular damage and dysfunction, potentially accelerating mitochondrial dysfunction and cellular aging.⁵⁴ Shen et al.⁵⁵ discovered that NOS inhibition reverses TLR2-induced chondrocyte dysfunction and attenuates age-related osteoarthritis. In this study, following the knockdown of the NOS2 gene, we proceeded to treat aged BM-MSCs with SS-31. The assessment of osteogenic differentiation revealed a marked increase in the intensity of ALP and ARS staining, as well as in the levels of osteogenic markers Runx2, OPN, and Osterix at both the protein and mRNA levels. This enhancement suggests that the inhibition of NOS2 can augment the osteogenic differentiation potential of aged BM-MSCs. Notably, the subsequent treatment with SS-31 after NOS2 gene knockdown did not induce significant changes in these indicators, implying that SS-31 may have become inert in the absence of its molecular target, NOS2. Moreover, this study identifies SS-31 as a promising candidate for revitalizing the osteogenic potential of aged BM-MSCs and restoring mitochondrial function, with NOS2 being a key target. These findings pave the way for new therapeutic strategies to combat age-related declines in skeletal health and regenerative capabilities. Although research on NOS2’s role in inhibiting MSC osteogenic differentiation is limited, the existing evidence highlights the complex and critical involvement of NO and its NOS isoforms in bone tissue formation and maintenance. Future research should further explore the specific mechanisms of NOS2 in this process and how modulation of NO levels can optimize bone tissue regeneration and repair. The therapeutic potential of SS-31 extends beyond its direct effects on osteogenesis. By restoring mitochondrial function, SS-31 could complement existing osteoporosis treatments, such as bisphosphonates or anabolic agents, potentially enhancing their efficacy in elderly patients. Moreover, the ability of SS-31 to mitigate mitochondrial dysfunction suggests broader applicability in other age – related conditions, including sarcopenia, neurodegenerative diseases, and cardiovascular disorders. Future studies should explore these integrative approaches to maximize the translational impact of SS-31.

This study provides valuable insights into the molecular mechanisms by which SS-31 rejuvenates aged BM-MSCs, offering potential applications for geriatric medicine and bone tissue engineering. The findings of this research may pave the way for the development of new therapies for osteoporosis and other age-related bone diseases, emphasizing the importance of targeting mitochondrial function and cellular senescence in regenerative therapies. In the future, it will be necessary to further investigate the long-term effects of SS-31 on BM-MSCs and its potential for integration with existing osteoporosis treatments. Understanding the complex interplay between mitochondrial function, NOS2 regulation, and osteogenic differentiation is crucial for optimizing the therapeutic potential of SS-31.

Funding Statement

The author(s) reported there is no funding associated with the work featured in this article.

Disclosure statement

No potential conflict of interest was reported by the author(s).

Authors’ contributions

Sen Duan: Conceptualization, Formal analysis, Investigation. Qindong Zhang: Methodology, Formal analysis, Data Curation. Jinqiang Zhu: Conceptualization, Formal analysis, Data Curation. Jiaming Wang: Data Curation, Methodology, Writing-Original Draft.

Data availability statement

The datasets used during the present study are available from the corresponding author upon reasonable request.

Supplementary material

Supplemental data for this article can be accessed online at https://doi.org/10.1080/15476278.2025.2519649